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Library of Congress Cataloging-in-Publication Data

Insect reproduction / Simon R. Leather and Jim Hardie, editors.

p. cm.
Includes bibliographical references (p. ) and index.
ISBN 0-8493-6695-X (alk. paper)
1. Insects--Reproduction. I. Leather, S. R. (Simon R.)
II. Hardie, Jim.
QL495.I4985 1995
595.7’016--dc20 95- 16294

A Library of Congress record exists under LC control number: 95016294

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ISBN 13: 978-1-315-89450-8 (hbk)

ISBN 13: 978-1-351-07360-8 (ebk)

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 INTRODUCTION
This book, consisting of ten review chapters contributed by leading workers in their
respective fields, from around the world, covers the whole subject of insect reproduction. It
begins with the basic physiological questions of insect reproduction, moves on to discuss the
new advances seen in the fields of behavioral and ecological mechanisms, and culminates by
examining the recent work on evolutionary biology and its application in the field.
Each chapter, although including a brief review of the basic seminal work, focuses mainly
on the advances made within the last ten years and highlights those areas in which the
respective authors see the greatest scope for further important advances. By allowing each
author full rein to explore their chapter subject using their particular "hobby horse," it has been
possible to make this not just a book of review chapters, but one in which exciting new ideas
have been raised.
This book should be of general interest to all entomologists, whether they are in pure or
applied fields, and should also be an important asset to any teaching program where entomol-
ogy is taught at the undergraduate and post-graduate level.

SRL, JH.
 THE EDITORS
Dr. Simon R. Leather is presently Lecturer in Applied Ecology and Pest Management at
Imperial College, London. He obtained his B.Sc. from the University of Leeds, England in
1977 with first class honors in Agricultural Zoology. After receiving his Ph.D. in 1980 from
the University of East Anglia in Norwich, he embarked on further research in aphid ecology.
Then followed a ten year spell with the British Forestry Commission where he worked in the
Research Division, primarily on the population biology of forest pests with particular refer-
ence to their reproductive behavior. He started his current position in 1992. Dr. Leather is a
Fellow of the Royal Entomological Society, a Member of the British Ecological Society, a
Member of the Flora and Fauna Preservation Society, and a Member of the Institute of
Biology. He sits on the Council of the Royal Entomological Society and edits their journal
Antenna.
Dr. Jim Hardie is presently a Principal Research Fellow at Imperial College, London. He
obtained a Ph.D. degree from the University of Birmingham, England in 1975 and a D.Sc.
from London University in 1989. He has worked in the field of aphid physiology for more than
twenty years and is regarded as one of the leading figures in this area. He is a Fellow of the
Institute of Biology, the Royal Entomological Society, and the Royal Microscopical Society.
 CONTRIBUTORS
Roger L. Blackman, B.Sc., Ph.D. Klaus H. Hoffmann, Prof. Dr.
Department of Entomology Department of Animal Ecology I
The Natural History Museum University of Bayreuth
London, England Bayreuth, Germany

Simon R. Leather, B.Sc., Ph.D.

Carol L. Boggs, Ph.D. Department of Biology
Center for Conservation Biology Imperial College of Science,
Department of Biological Sciences Technology, and Medicine
Stanford University University of London
Stanford, California Silwood Park
and Rocky Mountain Biological Ascot, England
Laboratory
Crested Butte, Colorado Athol McLachlan, Ph.D., D.Sc.
Department of Agriculture and
A. F. G. Dixon, B.Sc., D. Phil. Environmental Sciences
School of Biological Sciences University of Newcastle-upon-Tyne
University of East Anglia Newcastle-upon-Tyne, England
Norwich, England
Rachel Neems, B.Sc., Ph.D.
Department of Genetics
Cedric Gillott, B.Sc., Ph.D., D.Sc. University of Leeds
Department of Biology Leeds, England
University of Saskatchewan
Saskatoon, Canada Richard Wall, B.Sc., Ph.D.
School of Biological Sciences
Jim Hardie, B.Tech., Ph.D., D.Sc. University of Bristol
Department of Biology Bristol, England
Imperial College of Science,
Technology, and Medicine Christer Wiklund, Ph.D.
University of London Department of Zoology
Silwood Park University of Stockholm
Ascot, England Stockholm, Sweden
 TABLE OF CONTENTS
Chapter 1
Oogenesis and the Female Reproductive System .................................................................. 1
Klaus H. Hoffmann

Chapter 2
Insect Male Mating Systems .................................................................................................
33
Cedric Gillott

Chapter 3
Sex Determination in Insects ................................................................................................
57
Roger L. Blackman

Chapter 4
Hormones and Reproduction ................................................................................................95
Jim Hardie

Chapter 5
Fatal Attraction: The Disruption of Mating and Fertilization for Insect Control .............109
Richard Wall

Chapter 6
Parthenogenesis in Insects with Particular Reference to the
Ecological Aspects of Cyclical Parthenogenesis in Aphids ...............................................
131
A. F. G. Dixon

Chapter 7
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects ............ 143
Simon R. Leather

Chapter 8
Protandry and Mate Acquisition .........................................................................................
175
Christer Wiklund

Chapter 9
Swarm-Based Mating Systems ...........................................................................................199
Athol McLachlan and Rachel Neems

Chapter 10
Male Nuptial Gifts: Phenotypic Consequences and Evolutionary Implications................2 15
Carol L. Boggs

Index ....................................................................................................................................
243
 DEDICATION
This book is dedicated to our families in recognition of the support given during the
somewhat lengthy process that ensued once we had embarked upon this task. So, thank you
Gill, Fern, John, James, Thomas, and Matthew from Simon, and thank you Ros, Sally, Nicola,
and Robert from Jim.
 Chapter 1

OOGENESIS AND THE FEMALE REPRODUCTIVE SYSTEM

.
Klaus H Hoffmann

CONTENTS
I . Introduction ................................................................................................................... 1

I1. Morphology of the Female Reproductive System ........................................................2

A . External Genitalia ...................................................................................................
4
B . Structure of the Ovary ............................................................................................
4
C . Structure of the Female Accessory Reproductive System .....................................6
1. Spermatheca and Spermathecal Accessory Glands ........................................... 7
2 . Colleterial Glands and Mesodermal Accesso~yGlands .................................... 8
3 . Milk Glands ........................................................................................................
9
D . Endocrine Control of Differentiation of Accessory Glands and Ducts ............... 10

I11. Origin and Formation of the Germ Cells ...................................................................10

IV . Oogenesis .................................................................................................................... 11
A. Early Events in Oogenesis .................................................................................... 11
1. Oocyte Differentiation ..................................................................................... 14
2 . Endocrine Control of Early Oogenesis............................................................ 15
3 . Follicle Cell Differentiation ............................................................................. 15
4 . Trophic Function of Nurse Cells ..................................................................... 17
B . Previtellogenesis ................................................................................................... 19
C. Vitellogenesis ........................................................................................................19
1. Vitellogenin and Vitellin Chemistry ................................................................19
2. Vitellogenin Genes ...........................................................................................21
3 . Vitellogenin Synthesis ..................................................................................... 21
4 . Vitellogenin Secretion ...................................................................................... 22
5. Uptake of Vitellogenin by the Ovary ..............................................................23
D. Chorionization ....................................................................................................... 25
1. The Vitelline Membrane .................................................................................. 26
2. Chorion Formation ........................................................................................... 27

V. Ovulation and Oviposition ..........................................................................................

28

V1. Concluding Remarks ................................................................................................... 29

References ............................................................................................................................. 29

.
I INTRODUCTION
Insect reproduction results from a succession of interdependent steps which are often quite
different in nature and take place at various stages of the insect life cycle. The main reproduc-
tive events in females are sex determination. gonial mitoses and meioses. differentiation of the

.
0-8493-6695-X/95/$0.00+$.M
6 1995 by CRC PRSS Inc.
2 Insect Reproduction

reproductive organs, previtellogenesis and vitellogenesis, accessory gland functioning, sexual

behavior, mating, ovulation, and oviposition. It is the function of the female reproductive tract
to produce the eggs and to deposit them at an appropriate time and in an appropriate place.
In addition, the female reproductive tract must receive the spermatozoa from the male and
transport them to the spermatheca where they are stored until they are used to fertilize the eggs
as they are oviposited.
Insect reproduction strictly depends on environmental factors. Factors which may affect
reproduction include temperature, humidity, photoperiod, nutritive conditions, and a suitable
egg-laying substrate.
Regulation of insect reproduction involves numerous sensory receptors, nerve transmis-
sion, and integration in the brain, which regulates the synthesis of the two groups of insect
developmental hormones, the juvenile hormones and the ecdysteroids, and produces its own
neurohormones.' The mechanisms which regulate each reproductive step may vary with insect
species (see Chapter by Hardie).
Present knowledge of oogenesis has progressed variously, depending on the event consid-
ered. The best-known field is that of vitellogenesis and its endocrine regulation, while the
early events of ovarian development and its "fine-tuning" control by hormones are less well
understood.
Sexual reproduction is the general rule in insects, although many exceptions and modifi-
cations are observed (see Chapter by Dixon). In the more highly evolved social insects
(Hymenoptera), reproduction is limited to a small number of individuals, often one queen and
a small number of males (drones).

11. MORPHOLOGY OF THE FEMALE REPRODUCTIVE SYSTEM

The central site of egg production is the ovary. Ovaries are usually located dorsolateral to
the gut and each comprises a number of tubular ovarioles ensheathed by a network of
connective tissue (Figure 1). Each ovariole is composed of somatic and germ cell tissues. The
number of ovarioles per ovary varies from one (e.g., in some viviparous aphids and in dung
beetles) to about 3000 (in some higher termite queen^).^ Three basic types of ovary organi-
zation are found in insects (Figure 2). The panoistic ovarian, probably the most primitive one,
is found in the oldest families of insects, such as in Archaeognatha, Zygentoma, Odonata,
Plecoptera, and in most orthopteroid insects, but also in some Megaloptera (Corydalidae) and
most Siphonaptera. In the panoistic ovary, specialized nutritive cells are absent and most of
the informational resources of the oocyte are provided by the synthetic activity of the oocyte
nucleus itself. The polytrophic meroistic ovary is found in most endopterygotes, and in
Dermaptera, Psocoptera, and in Phthiraptera. In polytrophic ovaries, a number of nurse cells
are enclosed in each follicle along with an oocyte. In the Hemiptera, polyphage Coleoptera,
Megaloptera (Sialidae)/Raphidioptera, and also in the most "primitive" winged insects, the
Ephemer~ptera,~ telotrophic meroistic ovaries are found. In this ovarian type, a syncytium of
nutritive cells is connected with each oocyte by means of a trophic cord. In the vast majority
of insect species, the nutrive, or yolk, contribution is supplied largely by the fat body, but in
some cases the follicle cells can also serve as additional source of yolk. Accumulation of yolk
(vitellogenesis; Section 1V.C) normally occurs only in the terminal oocyte, that is, the oocyte
closest to the oviduct. Another follicle cell function is the formation of the protective layers
of the egg. These include the vitelline membrane and the chorion (Section 1V.D).
Besides the ovary, oviduct-associated secretory cells can have important contributions to
egg production (Section 1I.C). The most significant of such structures are the spermathecal
accessory glands and the female accessory glands, such as the colleterial glands. Last,
secretory functions of the vagina may play a role in egg production. Like other terrestrial
Oogenesis and the Female Reproductive System

$ p * r m o l h ~ c o l dond
common oviduct

FIGURE l. Female reproductive system: diagram of common type found in many insects. (From Gillott, C.,
Enromology, Plenum, New York, 1980, chap. 19. With permission.)

nutc

fc

PANOISTIC POLYTROPHIC TELOTROPHIC

FIGURE 2. Schematic diagrams showing the types of ovarioles. Meroistic ovaries may be organized in the
telotrophic and the polytrophic way, respe~tively.~.~~ch = chorion; fc = follicle cell; gm = germarium; nc =nurse cells;
nutc = nutritive cord; oo = oocyte; tf = terminal filament. (From Gillott,C., Enromology, Plenum, New York, 1980,
chap. 19. With permission.)
 Insect Reproduction

terao VIII. IX. and X

gonostyle

aonoooohvsis

FIGURE 3. The primitive structure of the pterygote ovipositor in the thysanuran Lepisma. (From Gillott, C . ,
Entomology, Plenum, New York, 1980, chap. 19. With permission.)

animals, insects have had to solve the problem of bringing together sperm and egg in the
absence of surrounding water (internal fertilization, see Chapter by Wall). Almost all insects
store the spermatozoa received from the male in a specialized organ, the spermatheca, until
they are used to fertilize the mature eggs.

A. EXTERNAL GENITALIA
The morphology of the organs specialized for copulation and oviposition is highly varied.
In the mayflies, the oviducts open directly to two genital pores behind the seventh abdominal
segment. In most insects, the appendages of the genital segments (eighth and ninth abdominal
segment) form an ovipositor. In the apterygotes and some of the winged insects, the ovipositor
is a simple opening both for copulation and for the deposition of eggs. The primitive structure
of the pterygote ovipositor can be seen already in the thysanuran Lepisma (Figure 3).4 Among
Pterygota, an ovipositor is found in Notoptera (Grylloblattodea),Dictyoptera, Ensifera, Caelifera,
and Hymenoptera, some Odonata and most Hemiptera, Thysanoptera, and Psocoptera. The
structure and elaborateness of the ovipositor is determined by the site of egg deposition. The
ovipositor of Hymenoptera may be considerably modified for boring, piercing, sawing, and
stinging. In the stinging Hymenoptera, such as bees, the eggs are released at the base of the
ovipositor, and the ovipositor is modified by the addition of poison glands and reservoirs that
evacuate the venom through the hollow sting. The ovipositor of Drosophila has sharpened
ends that penetrate the surface of fruit, while the ovipositors of some of the predatory wasps
are long (up to 15 cm in length) and sharp to penetrate the body of the insect prey.5

B. STRUCTURE OF THE OVARY

Insects have become particularly adept in manufacturing large numbers of oocytes within
the ovary. The fruit fly, Drosophila melanogaster, can produce, during a 10-week reproduc-
tion period, a quantity of eggs equivalent to 30 times her body eight.^ Her newly formed
oocytes can undergo a 100,000-fold increase in volume within 3 days. Such reproductive feats
are possible because of certain evolutionary adaptations: (1) Oocytes have developed methods
for incorporating massive quantities of female-specific proteins (vitellogenins) which are
transported in the hernolymph from the fat body to the ovary. (2) Mechanisms have evolved
for loading unfertilized eggs with the ribosomes, RNA, and long-lived mRNA that are
required in early embryogenesis. (3) The ovarioles are supplied by tracheae of the aeriferous
type, with a great diversity in the methods by which oxygen is delivered to the individual
Oogenesis and the Female Reproductive System

FIGURE 4. Panoistic ovariole of a cricket, Glyllus bimaculatus de Geer, with a mature egg in the terminal oocyte.
Photograph courtesy of K. H. Hoffmann, Bayreuth.

Panoistic ovarioles can be developed by blocking germ cell cluster divisions totally, as is
found in most "primitive" insects, or after germ cell cluster formation by final cleavage of
cystocytes, all of which develop as oocytes as found in stone flies or thripss In the panoistic
ovary, each of the ovarioles is composed of a terminal filament, the germarium, a series of
oocytes at the previtellogenic phase of development, one or more oocytes in the process of
vitellogenesis, and last, the mature egg (Figure 2A; Figure 4). The terminal filament is made
up of a group of flattened cells, surrounded by a basement lamina and an ovarian sheath, both
of the latter surrounding the entire ovariole. The oogonia are located in the most anterior
region of the germarium, followed by a zone of oocytes in the early stages of meiosis. At the
posterior end of the germarium, the oocytes are beginning to be surrounded by a monolayer
of follicle cells. The size increase of oocytes at the previtellogenic stage is accomplished by
an expansion of the cytoplasmic volume. In many cases, a multilayered pad of interfollicular
tissue is located between successive oocytes? In many insects with panoistic ovaries, vitello-
genesis commences in the penultimate oocyte only after ovulation at the terminal one (e.g.,
in Locusta migratoria and Schistocerca gregaria [Caelifera]),or in the case of ovoviviparous
cockroaches,after the loss of the egg case. The inhibitory effect is mediated by the interfollicular
cells which pass an inhibitory substance from the anterior to the more posterior oocyte. In
contrast, secretion from such cells located proximal to the oocyte stimulates vitellogenesis. In
other species (e.g., in Periplaneta americana [Blattodea] and Melanoplus sanguinipes
[Caelifera]), two or more oocytes may be vitellogenic at the same time, although at different
stages of the yolk deposition cycle. All the above-mentioned insects produce eggs in batches.
In the stick insect Clitumnus extadentatus, the different ovarioles can mature asynchronously,
and the female lays a few eggs per day for several weeks. The number of ovarioles in panoistic
ovaries can range from 4-3000 (5 in Acrididae, 15-30 in Tettigonioidea, 150-170 in Gryllidae,
about 3000 in Isoptera queens).
A terminal filament and a germarium are also found in polytrophic meroistic ovaries
(Figure 2B). In this case, the anterior region of the germarium contains one or more stem-line
oogonia and a number of daughter cells or cystoblasts. The cystoblasts divide to give a cluster
of cells remaining connected by structures called "ring canals" or "intercellular bridges." The
innovation of the polytrophic ovary is the differentiation of only one oocyte, which generates
from one, central cell of the cluster, whereas all other siblings are transformed into nurse cells.
6 Insect Reproduction

In many cases, clusters follow the 2"-rule ( l oocyte + 2"- 1 nurse cells), in which n represents
the number of cell cycles. Identical characters in polytrophic meroistic ovaries among hemi-
metabolous and holometabolous insects indicate a "basic type" of common rigi in.^ In Droso-
phila, where the number of cells in the cluster is 16, the clustering of the cells, as well as the
formation of the ring canals, is mediated by structures called fusomes. The fusomes contain
a random array of membranous vesicles and fibrils,'0." but no mitochondria and few ribo-
somes and microtubules. In the central region of the germarium, prefollicular cells grow
around the oocyte-nurse cell cluster, while in the posterior section of the germarium, typical
egg chambers are detectable; which means the oocyte-nurse cell complex is completely
surrounded by a unilayer of follicle cells. Previtellogenesis includes the enlargement of the
oocyte, an increase in the number of follicle cells, and the polyploidization of the follicle cells
and nurse cells. An epithelial sheath surrounds each ovariole and consists of a thin, acellular,
inner membrane; a median cellular network of muscle and tracheal cells; and an outer
epithelia1 membrane. The number of ovarioles in polytrophic ovaries can be highly variable
(usually 4 in the Lepidoptera, 10-30 in Drosophila, and 70-100 in Musca or Lucilia [Diptera]).
Commonly, a single oocyte per ovariole will become vitellogenic at one time.
Telotrophic ovaries differ from polytrophic ovaries by retention of all nurse cells in an
anterior trophic chamber and by changing oocyte-nurse cell determination. This type of ovary
developed independently three times (in Hemiptera, RaphidiopteraJMegaloptera [Sialidae],
and in polyphagous Coleoptera) from polytrophic ancestors and once directly from panoistic
ancestors in mayflie~.~ Despite fundamental differences between the subtypes of telotrophic
ovarioles, they share some common characters. As the oocytes move towards the region of
follicle formation, they become surrounded by prefollicular cells. The connection to the
tropharium is retained via a cytoplasmic strand, the nurse strand or trophic cord. The young
oocytes subsequently enlarge by incorporation of nurse cell material, transported through the
nutritive cords (previtellogenesis). The vitellogenic growth phase begins when yolk spheres
are observed to accumulate in the oocytes. The nutritive cords collapse during vitellogenesis.
The follicular cells surround each growing oocyte to form a monolayered epithelium, and this
tissue will secrete a vitellin membrane and the chorion. An interesting variant of the telotrophic
ovary is found in the polyphagous Coleoptera. The number of ovarioles remains more or less
constant in each species, but varies between species from 1 (some Scarabaeinae) to about 1000
in blister beetles (Meloe pro~carabaeus).~ In Creophilus maxillosus (Staphylinidae), the
differentiation of nurse cells and oocytes occurs within linear chains of sibling cells (linear
chain model). Only the most basal member of the sibling cluster develops into an oocyte; the
others differentiate into nurse cell^.^.^

C. STRUCTURE OF THE FEMALE ACCESSORY REPRODUCTIVE SYSTEM

The bottom of each ovariole forms a small duct or pedicel (Figure 1). The pedicels of each
group unite to form a calyx, and each calyx opens into a lateral oviduct. Usually the oviducts
of the two sides join to form a single median oviduct. The common oviduct is ectodermal in
origin and typically it is heavily invested with muscles. The lateral oviducts can be of
mesodermal or ectodermal origin. The presence or absence of a cuticular intima indicates the
origin. In Oncopeltusfasciatus (Hemiptera), the lateral oviducts undergo a drastic larval-adult
transformation during the last (fifth) larval stadium.12The long and thin larval oviducts shorten
and become very wide. This transformation is ecdysteroid dependent in a dose-related manner
and only takes place in the absence of juvenile hormone. The morphological transformation
is accompanied by dramatic cytological changes. Whereas the cells of the anterior part of the
oviduct commence with a strong secretory activity, the cells of the caudal part form a bizarre
pattern of cell projections which deposit the cuticle. The secretory material of the ductal
system may act as a lubricant for egg passage, as protective oothecal coverings, or as glues
to attach eggs to various substrates or to hold batches of eggs t ~ g e t h e rThe
. ~ distal end of the
common oviduct is called the gonopore, which serves for the discharge of eggs. In the
Oogenesis and the Female Reproductive System 7

Dermaptera, the gonopore is the external opening, located on the posterior part of the seventh
abdominal segment. In most insects, however, the gonopore opens into a genital chamber. The
opening to the outside is the vulva. The genital chamber can be of variable complexity and
is often associated with an ovipositor. Within some orders, an invagination of the primitive
genital chamber forms a distinct intermediate structure, the vagina, between the external vulva
and the gonopore. Generally, the vagina is not secretory and consists of a single layer of
epithelia1 cells, covered by a cuticular intima and surrounded by muscle. In many species, the
genital chamber has become modified to form a bursa copulatrix. An important function for
this organ is to receive spermatophores or seminal fluid. In Lepidoptera, the bursa remains in
the eighth abdominal segment, whereas the eggs are deposited through a separate opening, the
oviporus, on segment IX. In this case, the bursa is connected with the vagina by the seminal
duct. In other insects, however, there is no separation between the copulatory and egg-laying
apertures, and both of them open as the vulva on segment IX. In the bursa of some butterflies
(e.g., genus Danaus), tooth-like dentata are present and may function in tearing open the
spermatophore. Some secretory activity may also be associated with the bursa copulatrix,
since empty spermatophores are digested within the bursa of some insects. In ovoviviparous
and viviparous species, a brood pouch or uterus forms as an expansion of the vagina wall.
Two ectodermal glands (ectademia) are connected with the genital chamber or vagina. One
is the spermatheca in which spermatophoresare stored and that has a gland attached to its duct.
The other is a paired structure, the accessory glands or colleterial glands, with functions
associated with egg deposition.

1. Spermatheca and Spermathecal Accessory Glands

The morphology and arrangement, as well as the number of spermathecae, vary from
species to species. In most insects, the spermatheca is a single organ, spherical or ovoid in
form. In some insects, such as cockroaches and mosquitoes, secretory cells are associated with
the walls of the storage portion of the spermatheca, whereas in others, such as honeybees and
many beetles, the distal end of the spermatheca is specialized for secretion and is referred to
as the spermathecal accessory gland. In the genus Rhodnius (Heteroptera), the spermathecae
are a pair of blind tubules which open into the common oviduct near its junction with the
lateral oviducts.
In spite of the morphological diversity, the cellular elements of the secretory part of the
spermatheca are rather constant. A generalized diagram of a spermathecal secretory cell is
given in Figure 5. The cells form a cup-shaped cavity at their apical surface, and the membrane
in this region is thrown into numerous microvilli. The central cavity communicates with the
spermathecal lumen via a cuticle-lined ductule. In other forms, however, several ductules may
open at a single pore. The cuticle lining the ductule and the spermathecal lumen is produced
by separate duct cells which are interposed between the secretory cells and the lumen of the
spermatheca. Ultrastructural studies showed that the secretory cells have a phenotype associ-
ated with cells which are specialized for export protein synthesis?
The secretion produced by the gland cells is a mucoprotein or mucopolysaccharideand may
be used by the spermatozoa for an energy source.13 Removal of the spermathecal accessory
gland from females of Anthonomus grandis (Coleoptera) results in a gradual loss of motility
among spermatozoa in the spermatheca. The epithelium of the spermatheca, particularly in the
storage part, exhibits characteristics of ion-transporting epithelia and thus may be responsible
for providing an ionic milieu within the lumen of the spermatheca different from that in the
hemolymph. In some insects, the spermathecae may be sufficiently permeable to allow the
passage of various male secretory products.
Information on the control of spermatheca differentiation is rather limited. In the beetle
Tenebrio molitor, the differentiation of the spermatheca occurs in the pupal stage prior to the
eclosion to the adult. The differentiation process can be divided into three phases: ( l ) cellular
proliferation, (2) cellular morphogenesis, and (3) cuticulogenesis. From in vitro culture
 Insect Reproduction

FIGURE 5. Diagrammatic representation of a spermathecal secretory cell and its spatial relationship to the duct cell
and basement membrane. CC, central cavity; CU, cuticle; DC, duct or canal cell; DU, ductule; ER, rough endoplasmic
reticulum; LU, lumen; MV, microvilli; N. nucleus; SC, secretory cell. (From Kaulenas, M. S., Insect Accessory
Reproductive Structures. Function, Structure, and Development. Springer-Verlag, Berlin, 1992. With permission.)

experiments, it was concluded that cuticulogenesis is under control of 20-hydroxyecdysone;

the hormone is necessary to initiate cuticle deposition.14 With regard to the control of the
acquisition of definitive levels of differentiated functions by the spermatheca, some informa-
tion is available for a number of Orthopterans. In Chorthippus curtipennis (Caelifera,Acridinae),
allatectomy leads to degeneration of the spermatheca, while in Gomphocerus rufus (Caelifera,
Acridinae), allatectomy resulted in the inability of females to dissolve transferred spermato-
phores, suggesting a failure in the production of spermathecal proteolytic enzymes? The
results suggest that juvenile hormone is necessary to initiate and possibly maintain differentiative
secretory function in these grasshoppers. On the other hand, allatectomy in L. migratoria
(Caelifera, Locustinae) does not alter the histology of the spermatheca. In Rhodnius prolixus
(Heteroptera), removal of the neurosecretory cells in the pars intercerebralis results in a failure
of the secretion to appear in the lumen of the gland as a result of the absence of a myotropin
from the neurosecretory cells. The myotropin acts on the muscles of the spermatheca to
squeeze the secretion from the reservoir beneath the intima into the lumen.13

2. Colleterial Glands and Mesodermal Accessory Glands

Many insects produce protective coatings for the newly laid eggs. The Lepidoptera coat
individual eggs with a proteinaceous glue that hardens on contact with air and attaches the
eggs onto appropriate substrates. Accessory glands in the genus Musca (Diptera) contribute
in aiding in the fertilization process by providing secretions which assist in liberating sperm
acrosomal contents.15 Aquatic insects often produce egg cases of a gelled substance, but the
most complex of these structures may be the tough ootheca of cockroaches and mantids. In
mantids, the glands produce a polymerized protein foam. In cockroaches, several layers of
tanned protein form a complicated egg chamber with elaborate respiratory apparatus and
release valves. The sources of these protective devices are the colleterial glands which branch
off the vagina or genital chamber. In most cases, the colleterial glands are a pair of organs
composed of a number of multibranched tubules and are formed from invagination of the
epidermis (Figure 1). The morphology of the colleterial glands has been well documented.
Five cell types compose the left colleterial gland; four types of glandular cells are homologous
Oogenesis and the Female Reproductive System 9

with the dermal glands of the integument, and a chitinogenic cell type is homologous with the
epidermal cell of the integument. The latter cell type is found interspaced between the
glandular cells and secretes the protective intima which lines the lumen of the tubule. Each
of the gland cell types is distinct from the other, and is located in separate regions of the tubule.
Type 1 cells are found in the most posterior region of the tubule and are in a presecretory stage.
Type 2 gland cells are found anterior to type 1 cells and lack a well-defined rough endoplasmic
reticulum (ER), but are packed with mitochondria. The morphology of the type 3 cells
resembles that of both type 2 and type 4, and appears to be a transient intermediate form. The
type 4 cells dominate the anterior ends of the tubules and are packed with rough ER and with
mitochondria.16
The colleterial gland of P. americana has been studied most extensively?J3 The left side
tubules (type 4 cells) secrete the proteins (oothecins) which make up the structure of the
ootheca, together with a polyphenol oxidase (type 2 cells) as well as the precursor of the
tanning agent, the 4-0-P-glucoside of protocatechine acid. The right colleterial gland secretes
the enzyme P-glucosidase. The opening for the duct from the right gland is more anterior than
the opening for the duct from the left gland. At the time of ovulation (see Section VI), the
chorionated oocytes migrate down the oviduct to the genital atrium for fertilization. After
fertilization, the eggs are transported more posteriorly and pass the duct of the right gland
where the contents of the gland (P-glucosidase) are secreted onto the fertilized egg. Thereafter,
as the egg moves more posteriorly, it encounters the secretions of the left colleterial gland. The
subsequent mixing of the secretions from both glands results in the hardened ootheca.I6
Apparently, juvenile hormone affects the synthesis of oothecins in left colleterial
gland^?.'^.'^.^^ Analyses of the Periplaneta oothecin cDNA sequences and of the resultant
predicted amino acid sequences have confirmed the existence of I I major glycine-rich
oothecins which represent six size classes with molecular weights of 14.5, 15.5, 17-18.5,
23.5-26, 28.0, and 37-39 kDa, respectively.18The oothecin sequences have numerous simi-
larities to silkrnoth chorion proteins. In mantids, the chemistry of the ootheca proteins shows
some unique features. The glucosides identified in five mantid species are 3-0-P-glucosides
of N-P-alanyldopamine and N-(N-malonyl-P-alany1)dopamine.The light color of the ootheca
and the occurrence of phenolic compounds modified at the P position of the parent compounds
suggest that P-sclerotization occurs in mantid oothecae.
In S. gregaria and other Orthopterans, the foamy ootheca is produced, at least in part, by
mesodermal accessory glands (mesadenes) which consist of convoluted blind tubules opening
into the proximal end of the lateral oviducts. In spite of the great structural differences between
glands responsible for oothecal formation in cockroaches and locusts, similar mechanisms
may operate to harden the ootheca.

3. Milk Glands
In tsetse flies, members of the genus Glossina, the accessory glands of the female are
transformed into milk glands and supply a secretion upon which the developing larva feeds
(adenotrophic vivipary). In these flies, the female ovulates a single egg into a uterus, where
it hatches. The growth of the larva is rapid under such circumstances. In Glossina austeni for
example, development from an egg to a fully grown larva weighing some 30 mg requires only
9-10 days. The fully mature larva is then "larviposited." The milk gland in Glossina is a
branched tubular structure ramifying throughout the abdomen and emptying into the uterus via
a single muscular collecting duct which contains two channels.I3 The tubules consist of a
single layer of secretory cells similar to those in other accessory glands. The tubules undergo
cyclic changes in diameter, largely as a result of changes in the volume of secretory reservoirs.
The secretory reservoir is an extracellular structure, formed by a cup-shaped invagination of
the apical membrane of each of the secretory cells. In G. austeni, the tubular diameter reaches
a peak of about 100 pm 3 days before each larviposition, and a diameter of about 30 pm at
10 Insect Reproduction

each larviposition, followed by a resynthesis of the "synthetic machinery." Removal of the

corpora allata greatly reduces the production of milk in tsetse flies.
Milk glands also occur in some cockroaches. In the viviparous cockroach Diploptera
punctata, nutrient is supplied to the developing embryos, which increase in weight by a factor
of about 50 before they hatch. The developing embryos receive this nutrition by ingesting a
fluid secretion ("milk") in the brood pouch. The brood pouch is a part of the genital chamber.
The intima of this cuticle-lined chamber is penetrated by pores, each of which is the opening
of a ductule leading from a secretory cell. In Diploptera, juvenile hormone is necessary to
allow the decline of milk gland synthesis at the termination of pregnancy.lg

D. ENDOCRINE CONTROL OF DIFFERENTIATION

OF ACCESSORY GLANDS AND DUCTS
The dependency on ecdysone and 20-hydroxyecdysone of the organogenesis of accessory
glands and genital ducts has been demonstrated in several insect species.'-20The development
of the colleterial glands of P. americana, which occurs at the end of the last larval instar,
requires ecdysone. In young females of L. migratoria and S. gregaria, implantation of
additional corpora allata accelerated the development of both the oviducts and colleterial
glands. In T.molitor, Ephestia kiihniella and Samia cynthia, L. migratoria, and 0.fasciatus,
duct differentiation only occurs in the presence of ecdysone in vivo and in vitro. However,
other tissues of epidermic origin, such as the spermatheca, escape mitotic stimulation by
ecdysone.
Besides the effects exerted by hormones upon the reproductive organs, humoral relation-
ships between gonads and ducts have been demonstrated. In Drosophila, female ducts were
sometimes found to be attached to the gonads, thus causing degeneration. In the bugs
Dysdercus fasciatus and Triatoma infestans, atrophy of one ovary was observed together with
the regression of the upper part of the corresponding lateral oviduct, and the experimental
section of the oviduct had the same effect in both species.'

111. ORIGIN AND FORMATION OF THE GERM CELLS

Both sperm and eggs are derived from primordial germ cells set aside very early in the
development of the embryo. Among the orthopteroid insects, in species where the germ cells
appear early in development, such as in the house cricket Acheta domesticus and the grass-
hopper Melanoplus differentialis, the germ cells appear to be of ectodermal origin, forming
at the posterior pole of the egg at the time of mesoderm ~egregation.~ Later in development
in A. domesticus, they become associated with the mesoderm of the second and third abdomi-
nal segments; in M. differentialis they migrate into the coelomic cavities of the first to eighth
abdominal segments, where they associate with the splanchnic wall and form a genital strand,
from which the gonad differentiates later in embryogenesis. In species where the germ cells
are first recognizable slightly later in embryonic development, they are associated with the
median walls of the dorsal cavities in the abdominal segments (e.g., L. migratoria, Blattella
germanica, P. americana). In each of these species, a genital ridge containing the germ cells
is formed on each side of the embryo. The typical genital rudiment in these insects, during or
just after involution of the germ band (anatrepsis), consists of a terminal filament membrane;
a mesodermal dorsal cell mass, ventral to the filament membrane; a central cell mass,
composed of primordial germ cells and mesodermal cells; a ventral cell strand of mesodermal
cells which are the primordia of the gonadal portion of the genital ducts; and a surrounding
epithelia1 membrane which envelops all of the above (Figure 6).
Among the endopterygotes, germ cell formation is well understood in the Diptera. The
germ cells are formed from pole cells, which are established very early in development at the
posterior pole of the embryo. In Drosophila, about 18 energids (cleavage nuclei) enter the
posterior pole plasma and are pinched off as pole cells. The pole cells continue to divide, to
Oogenesis and the Female Reproductive System

\ TERMINAL FILIMENTS

FIGURE 6. Diagrammatic representation of the early development (late embryonic stage) of the female reproduc-
tive system in an orthopteroid insect.(From Kaulenas, M. S., Insect Accessory Reproductive Structures. Function,
Structure, and Development, Springer-Verlag. Berlin, 1992. With permission.)

produce eventually between 37 and 71 cells? Only some of the pole cells migrate to the
presumptive gonads, which lie on either side of the gut and are mesodermally derived. The
final number of pole cells in the gonad has an upper limit of about 13 pole cells per gonad at
about stage 16 of ernbryogenesi~.~~ The pole cells which fail to reach the gonad probably
degenerate later on. The determination of the pole cells as presumptive germ cells depends
upon the interaction of the entering energids with the cytoplasm of the posterior pole cells
(polar plasm). Likely candidates for cytoplasmic elements important for germ cell determina-
tion are polar granules, which are concentrated at the posterior pole of late stage oocytes and
early embryos.22Functionally similar posterior pole plasms, which determine germ cell
differentiation, occur in the Coleoptera and the Hymenoptera. In the Coleoptera, germ cells
become distinguishable at the time of blastoderm formation, at the posterior end of the egg.
In most Hymenoptera, germ cells first become recognizable during gastrulation or later,
forming from the mesodermal tube. In many Lepidoptera, germ cells appear at the posterior
pole just after blastoderm formation.
Germ cells (oogonia) are the only cells that normally exhibit genetic programs that lead to
the construction of eggs.

IV. OOGENESIS
The first events in oogenesis include mitosis, the onset of meiosis, and ovariole differen-
tiation. In the panoistic ovary, all oogonia (except stem line oognoia) are transformed to
oocytes, whereas in the meroistic type oogonia generate both oocytes and nurse cells.

A. EARLY EVENTS IN OOGENESIS

Since more information is available concerning oogenesis in D. melanogaster than for any
other insect, this fruit fly will be used to illustrate the early events of ovarian development that
are shared by many evolutionary advanced insects with a polytrophic meroistic ovary.6Each
Drosophila ovariole contains a collection of egg chambers in which each oocyte is one
member of a clone of interconnected cells. In brief, four consecutive mitoses of the cystoblast
and its daughter cells in region 1 of each germarium give rise to the 16 germ-line cells
(cystocytes), which remain connected via intercellular bridges due to incomplete cytokinesis
(Figure 7). King et al." suggest that the interactions of the centrioles and fusomes during the
cystocyte divisions are responsible for the multiple-branched canal system that results. In
region 2, the individual 16-cell clusters become separated by invading somatic follicle cells.
One cell with four intercellular bridges (ring canals), the prospective oocyte, moves from a
central position in the cell cluster to a posterior location between regions 2 and 3 of the
germarium. This spatial reorganization of the 16 gem-line cells results in a polarization of the
cluster: the prospective oocyte becomes positioned posterior to the remaining 15 cells, which
differentiate into polyploid nurse cells. The forming follicle has established an anteroposterior
 Insect Reproduction

FIGURE 7. Germarium and young follicles in Drosophila oogenesis. The insert shows a diagram of the steps in
the production of a clone of 16 cystocytes. By a series of four mitoses, each followed by incomplete cytokinesis, a
branching chain of 16 interconnected cells is produced. Cell 1 represents the later oocyte which moves from a central
position in the cell cluster to a posterior location. FC, follicle cell; NN, nurse cell nucleus; O N oocyte nucleus.
Photograph courtesy of H. 0 . Gutzeit, Dresden.

axis. The nurse cells grow and simultaneously transfer cytoplasmic macromolecules to the
oocyte (see Section 1V.C). The follicle cells begin to form a monolayered epithelium around
the germ-line cells in region 3 of the germarium. This process starts at the posterior end of the
follicle. At the anterior end, a special group of follicle cells forms a stalk ("stalk cells"), thus
separating the follicle from the germarium and releasing it into the ~ i t e l l a r i u m . ~ ~
In the silkrnoth Bombyx mori, there are only three cystocytes divisions resulting into eight
germ-line cells (n = 23). In the braconid wasp Habrobracon juglandis, the final number of
cystocytes per cluster is 32; in other wasps, the number is not fixed but varies from 20-80.24
Fleas with polytrophic meroistic ovarioles (some species of the Hystrichopsylloidea) have
germ cell clusters consisting of 32 cells ( F ) which are generated by five mitotic cycles during
the pupal stage. One of the cells containing five intercellular bridges becomes the oocyte; the
others serve as nurse cells. However, nurse cells remain small and show the same ultrastruc-
. ~ ~ species of lacewings do not obey the N = 2" rule.6 For
tural characters as the o o ~ y t eCertain
example, in Chrysopa perla, egg chambers contain 12-14 cystocytes. In this case, first- and
second-generation cystocytes divide in synchrony, whereas at M, (see Figure 7), cells 3,4,7,
and 8 divide; the rest do not. In the earwigs (Dermaptera), each follicle in the vitellarium
Oogenesis and the Female Reproductive System

FIGURE 8. Hypothetical diagrammatic representation of germ cell cluster formation in subgroups of Hemiptera:
(1) development begins in one persisting germ cell; (2) the germ cell divides by mitosis, followed by incomplete
cytokinesis. One of the daughter cells is determined as a presumptive nurse cell (black nucleus), the other will be a
presumptive oocyte (white nucleus); (2 a-c) germ cell division in scale insects (Coccina); (3) in other groups, the
presumptive nurse cell and the presumptive oocyte divide, giving rise to a cluster of four germ cells arranged in a
rosette configuration; (3a-c) germ cell division in aphids (Aphidina); (4) further divisions of presumptive nurse cells
and oocytes; (4 a-b) germ cell division in bugs (Heterotera), where the oocyte subclone has its divisions limited.
Asterisk = region of microtubule matter. For further details see text. (Reprinted from King, R. C. and Biining J.,
Comprehensive Insect Physiology, Biochemistry, and Pharmacology, Vol. 1, Kerkut, G. A. and Gilbert, L. L, Eds.,
Copyright 1985, ch. 3. With permission from Pergarnon Press Ltd., Headington Hill Hall, Oxford 0 x 3 . OBW, UK.)

consists of one oocyte and one nurse cell, surrounded by a single layer of follicle cells.
Formerly, the polytrophic meroistic ovary of earwigs has been looked at as a parallel devel-
opment, but new findings indicate only one origin of the polytrophic meroistic ovary (see
Section II.B).8
In all hemipteran species with telotrophic ovarioles, the germ cells are also clearly sepa-
rated into nurse cells and oocytes. The oocytes are found at the base of the tropharium (see
Figure 2C), whereas the nurse cells occupy its upper parts. Two models of germ cell cluster
formation have been proposed. Both assume that independent stem cells in the apical region
of the tropharium fuse to form germ cell clusters in which basally located germ cells are
subsequently determined as oocytes. Based on data derived from several groups of Hemiptera,
a new model has been advanced which assumes that cluster formation begins with a single
cell. The first division is a differential mitosis, leading to a presumptive oocyte and a
presumptive nurse cell. In scale insects (Coccinea), the presumptive nurse cell divides and its
apical descendant will divide again. The final configuration is a rosette of four cells in which
the intercellular bridges stay close together. During rosette formation, the intercellular bridges
then vanish, and the trophic core forms (Figure 8). The last step in cluster formation is the
polyploidization of nurse cells. In other groups, the presumptive nurse cell and the presump-
tive oocyte divide, giving rise to a cluster of four germ cells arranged in a rosette configura-
tion. In aphids, germ cells continue to divide, and subclones of 2"oocytes and 2" nurse cells
14 Insect Reproduction

arise. Rosette formation, formation of the trophic core, and polyploidization of nurse cell
nuclei are the same as proposed for scale insects. Further divisions of presumptive nurse cells
and oocytes lead to the situation in bugs (Heteroptera) with a constant number of oocytes, but
an increasing number of nurse cells.
In snakeflies (Raphidioptera) and alderflies (Megaloptera, Sialidae), the same type of
telotrophic ovary occurs. Cluster formation starts with germ cell migration into the ovariole
anlage. The number of germ cells increases as more germ cells enter the anlage and as those
already there divide. Dividing stem cells undergo complete cytokinesis next to the terminal
filament (apical region), whereas germ cell clusters arise by incomplete cytokinesis more
basally. The clusters are disc-shaped and oriented at right angles to the long axis of the
ovariole. Each cluster presumably contains 2" cells, with five as the maximum number of
division^.^ Prospective oocytes do not differ from nurse cells in their ultrastructure, except for
one fact: nurse cells lose their cell membranes totally to form a syncytium, whereas prospec-
tive oocytes remain in their original status, which they acquired at the end of cluster divi~ions.~
In polyphage Coleoptera, germ cells migrate in the ovariole anlagen, and cluster formation
is started by mitosis of each germ cell. Each germ cell can undergo only a limited number of
mitoses, each followed by incomplete cytokinesis. Later mitoses are highly synchronized
within each cluster, but not between different clusters. In most species, the clusters are
oriented parallel to the long axis of the tropharium. A three-dimensional network of interstitial
cells keeps the nurse cell nuclei in place, when nurse cell-nurse cell membranes are r e d ~ c e d . ~
Oocytes develop at the base of the tropharium, primarily connected to nurse cells by an
intercellular bridge. However, recent investigations have shown that cluster formation is more
complex than has been assumed before, and that ramifications of clusters occur, even between
oocytes.

1. Oocyte Differentiation
Immediately after the cluster of 16 cystocytes is found in a Drosophila germarium, both
four-canal cells (cells 1 and 2) form synaptonemal complexes in their nuclei during the time
they pass through germarial region 2. The synaptonemal complexes form during zygonema,
are completed during pachynema, and are responsible for the synapsis of homologous chro-
mosomes during meiotic pr~phase.~ Since cells 1 and 2 start meiosis, they are called pro-
oocytes. Sometimes cells 3 and 4 also form synaptonemal complexes and enter meiosis,
whereas cells 5-16 fail to enter meiotic prophase. In the posterior region 2, one of the two four-
canal cells loses its synaptonemal complexes and enters the cycle of endomitosis characteristic
of nurse cells. The other cell continues to develop as an oocyte and retains its synaptonemal
complexes during the previtellogenic stage of oogenesis in the vitellarium (see Section IV.B).
The divergence of the two pro-oocytes takes place in the region where follicle cells first
surround the 16-cell clone. It is suggested that the first pro-oocyte to come into contact with
a follicle cell is the one that receives the critical stimulus that causes it to continue on the
oocyte developmental pathway. Such cellular interactions between the germ-line cells and the
somatic follicle cells have been studied for a long time.26.27More recent data show that the
correct cellular organization and determination of cells in the germarium seem to depend on
the activity of several genes. For example, in Drosphila, genes like egaliterian and Bicaudal-
D are apparently involved in cystocyte diversification, since in mutant follicles a nurse cell
differentiates instead of the oocyte. The genes dicephalic and spindle-C are required for the
correct spatial arrangement of the cystocytes. Moreover, the mutant dicephalic illustrates the
importance of early cellular interactions between somatic follicle cells and germ-line cells.28
However, it remains to be analyzed which kind of specific signals the pro-oocytes receive in
the germarium.
In some carabid beetles, all cystocytes of a clone form synaptonemal complexes, and
consequently all nurse cells enter meiotic prophase together with the presumptive oocyte. In
some telotrophic ovaries, the cell which differentiates as an oocyte also seems to depend upon
Oogenesis and the Female Reproductive System 15

its position relative to certain somatic tissues. These facts again are best explained by
involving signals passed to the cystocyte by adjacent somatic cells.
In all insects with panoistic ovaries, fully activated oocyte chromosomes are required. In
these cases, the oocyte chromosomes enter a lampbrush state in the diplotene stage of meiotic
prophase. Lampbrush chromosomes also occur in many insects possessing telotrophic meroistic
ovaries.

2. Endocrine Control of Early Oogenesis

In the blood-sucking bug Panstrongylus megistus, determination of mitosis, the onset of
meiosis, and ovariole differentiation take place during the last larval instar and begin 24-48
h after a blood meal.' Observation of the neurosecretory (ns) cells of the pars intercerebralis
during the last larval instar from the time of the blood meal until imaginal moult showed the
presence of four distinct ns ceIl types with different patterns of activity. In type A cells, the
ns material is released before the mitotic crisis, whereas in type A' cells it is discharged before
the onset of meiosis. Hence the ns cells appear to trigger both mitosis and meiosis.
Electrocoagulation of pars intercerebralis prevented ovarian development and caused its
degeneration. Destroying only type A ns cells resulted in an inhibition of the mitotic crisis, but
the onset of meiosis was unaffected. Removal of the prothoracic glands showed that ecdysteroids
do not trigger the onset of mitosis, but are necessary for that of meiosis. Hemolymph
ecdysteroid determinations demonstrated two peak values; the first coincided with the onset
of meiosis, the second preceded moulting. Juvenile hormone (JH) does not seem to intervene,
either in early gonial mitosis or in the initiation of meiosis. In L. migratoria, P. americana,
and Gryllus bimaculatus (Ensifera, Gryllidae), it was shown that the meiosis reinitiation in the
vitellogenic oocyte (see Section 1V.C) is preceded by an increase of free ecdysone of the
oocytes. In vitro, the reinitiation of meiosis can be determined by the addition of e c d y ~ o n e . ~ ~
Not only the germ cells but also the mesodermal elements surrounding them have to go
through certain steps in order to develop adult structures, and these events are controlled by
hormones. It was shown both in vivo and in vitro that growth and differentiation of nurse cells
and follicle cells require the presence of 20-hydroxyecdysone, whereas JH may suppress the
effects of 20-hydroxyecdysone. In conclusion, 20-hydroxyecdysone and JH are both neces-
sary for ovariole differentiation and function, but the two hormones must act separately and
successively.

3. Follicle Cell Differentiation

During oogenesis, the follicle cells follow a characteristic differentiation program. Three
processes have been studied particularly thoroughly:

1. The production of localized developmental signals which bind to their respective

receptors in the oocyte membrane. These signals play a role in the axial determination
of the embryo.30
2. The contribution of the follicle cells to the synthesis and uptake of yolk proteins (see
Section IV.C.3).
3. The formation of the egg shell and its genetic control (see Section 1V.D).

General follicle cell morphology has been well described for the cockroach P. americana.
The earliest follicle cells are of the squamous type, with the apical ends of the cells applied
closely to the oolemma. As the oocyte grows, the follicle cells rapidly increase in number and
gradually become cuboidal. The follicle cells send out processes which interdigitate with the
microvilli of the oocyte. The cuboidal shape is maintained until prior to vitellogenesis when
the follicular epithelium becomes columnar. Similar arrangements have been found also in
other species (e.g., L. migratoria, Galloisiana nipponensis [Grylloblattodea], and ants of the
genus Formica [Hymenoptera]).At the start of the vitellogenic phase, the cell shape changes
 Insect Reproduction

FIGURE 9. Diagrammatic representation of the follicular epithelium from a Drosophila ovary (vitellogenic
follicle). The basement membrane (bm) with laminin (lam) in circular orientation is partly removed to expose the
basal face of the follicle cells. Int, high concentrations of PSP integrin at the contact site of the cells; mf, parallel
. microfilament bundles below the cell membrane which extend in the same circular direction as laminin; ics,
intercellular space. Figure courtesy of H. 0.Gutzeit, Dresden.

again to assume a somewhat spherical (P. americana) or flattened character (L. migratoria)?
Ultrastructural studies showed that the cytoplasm of the follicle cells at the late previtellogenic
and vitellogenic stages contain large numbers of mitochondria, multivesiculated bodies, and
Golgi complexes, which are characteristics of a highly active tissue. The columnar cells are
well supplied with rough ER and large amounts of ribonucleoproteins. In some dipteran and
hymenopteran insects, in addition to septate desmosomes and adhesion plaques, intercellular
bridges interconnect adjacent follicle cells. The bridges appear to result from incomplete
cytokinesis and may serve to synchronize differentiation and function of the follicular epithe-
lium. Several authors have described the occurrence of gap junctions between adjacent follicle
cells (L. migratoria), as well as between follicle cells and oocyte (R. prolixus, Tribolium
destructor [Coleoptera, Tenebrionidae], D. melanogaster)? The junctions disappear during
the chorion formation phase and during atresia.
The typical transition of follicle cell morphology from cuboidal to columnar and flattened,
with large intercellular spaces, suggests that cytoskeletal changes (microtubuli and microfila-
ments) are responsible for the cell shape transformations. The maintenance of the columnar
shape is associated with a well-organized, cylindrical orientation of the microtubular
cytoskeleton (P. americana). A random distribution of the microtubules might facilitate the
transition to a more flattened morphology. Alterations in microtubular association seem to be
juvenile hormone dependent. In Drosophila, parallel microfilament bundles were shown to be
present at the basal side of the vitellogenic follicle cells facing the basement membrane
(Figure 9). The density of the microfilament bundles increases during the course of oogenesis.
Indirect evidence from a variety of experiments using proteolytic digestion of collagen and
inhibitor studies suggests that the microfilaments are required for the adhesion of the follicle
cells to the basement membrane. In the absence of parallel microfilaments, the cells lose their
epithelia1 character and round Components of the extracellular matrix may affect the
organization of the cytoskeleton. Consequently, the parallel microfilaments, together with the
extracellular matrix glycoprotein laminin (and possibly additional components of the base-
ment membrane), act in concert in shaping the follicle.
The total number of follicle cells associated with a single oocyte varies with the develop-
mental stage and with the size of the oocyte. A vitellogenic oocyte of Leucophaea has
approximately 27,000 investing follicle cells. Drosophila follicles when first formed have 80
cells, increasing to about 1200 in mature follicles?
In Drosophila, a rather interesting additional feature is found. Once the maximum follicle
cell number is reached, some of the follicle cells undergo a series of migrations (Figure 10).
Oogenesis and the Female Reproductive System

FIGURE 10. Diagrammatic representation of Drosophila midvitellogenic follicle, indicating follicle cell migratory
pathways (1-5). FC, follicle cell; NC, nurse cell; 0, oocyte; ON, oocyte nucleus. (From Kaulenas, M. S., Insect
Accessory Reproductive Structures. Function, Structure, and Development, Springer-Verlag, Berlin, 1992. With
permission.)

The various cell migrations are microtubule de~endent.~' In addition to the involvement of
microtubules, the large, steady electric currents which have been shown to traverse the
follicles are proposed to direct the follicle cell migration.32
In biological systems, electric current is carried by ions, not by electrons. Ion asymmetries
within the follicle, which may be associated with the electrical phenomena, have been
confirmed, e.g., in Drosophila and Hyalophora cecropia (Lepidoptera) meroistic ovaries. In
the Cecropia moth, a potential difference of about 6 mV is observed over the 30-pm-wide open
cytoplasmic bridge which interconnects the oocyte with a trophocyte. The oocyte is at positive
potential as related to the tro~harium.~~Most studies of currents associated with the vitellogenic
phase of an oocyte have reported an anterior inward current and a posterior outward directed
current, rotationally symmetric about the oocyte's long axis. In polytrophic ovaries, the
anterior to posterior currents are suggested to provide an electrophoretic force for distributing
negatively charged nurse cell products to the vitellogenic oocyte. In panoistic ovaries, currents
in this type of oocyte might play a role in the intracellular distribution of cell organelles or
products throughout the oocyte (see Section 1V.C).
Before or during the vitellogenic stage, the follicular epithelia] cells become polyploid. In
panoistic ovaries, an exact doubling of DNA during polyploidization is typical, whereas in
meroistic ovaries, polyploidization is generally not exact. For example, in Drosophila hydei,
DNA sequences for the ribosomal RNAs are severely under-replicated, and it has been
suggested that polyploidization increases the concentration of those genes which play impor-
tant roles in follicle cell function^.^ All the major functions of the follicle cells seem to be
juvenile hormone dependent.

4. Trophic Function of Nurse Cells

In both the polytrophic and the telotrophic ovaries, the nurse cell-oocyte syncytium is a
polarized structure, with the site of "nutrient" formation distinct and separate from the
18 Insect Reproduction

recipient cell, the oocyte. The most significant difference between the two ovary types is the
greater separation between donor and recipient poles in the telotrophic ovary, making it more
difficult to use diffusion as a transport mechanism for nutrients.
The trophic function of the nurse cell is enhanced by endoreplication of the DNA. In
Drosophila, the nurse cells begin their cycle of endomitotic DNA replication in region 3 of
the germarium. In the vitellarium, each nurse cell undergoes another seven replications. The
maximum level of polyploidy reached by Drosophila nurse cells is 2'O. In the giant moth
Antheraea polyphemus, each of the seven nurse cells reaches ploidy levels approaching 216.34
In most polytrophic meroistic ovaries, even in Drosophila, an apical-basal gradient in
polyploidization exists among nurse cells. Highest ploidy levels are found in basal nurse
cells.35Whereas in panoistic ovaries an exact doubling of DNA content during polyploidization
seems to be typical, in meroistic ovaries, polyploidization is generally not exact, but some
sequences (e.g., ribosomal RNA genes, histone genes, telomere sequences, satellite se-
quences) can be under-re~licated.~~ Very young nurse cell nuclei have distinct polytenic
(giant) chromosomes, but chromosomes cannot be distinguished in large nuclei, as different
sections are replicated to varying degrees. In the last endoreplication cycle, all sequences,
including the previously under-replicated, replicate fully. In telotrophic ovaries, trophocyte
nuclear DNA also undergoes multiple rounds of duplication - a total of seven in Dysdercus
intennedius (Heter~ptera).~'
In the case of polytrophic ovaries, all or most of the nurse cell cytoplasm is transferred to
the oocyte towards the end of oogenesis, whereas for telotrophic ovaries the process could be
more selective. Among the major products accumulated by oocytes are large quantities of
mitochondria. Also produced are massive stores of ribosomes, which are used by embryonic
cells during the early periods of development, when little or no rRNA synthesis takes place.
In most meroistic ovaries, the expanded ribosomal gene numbers in the polyploid nurse cells
provide sufficient templates for the massive rRNA synthesis. In a few cases, however,
additional extrachromosomal rDNA amplification is encountered, as in water beetles.38 A
large variety of nonribosomal transcripts is also synthesized in nurse cells and transferred to
the oocyte. Among the most interesting and important gene transcripts synthesized in nurse
cells and stored in oocytes are those which specify embryo p ~ l a r i t y . ~Other
~ . " ~mRNAs which
are transcribed in the nurse cells and later transferred to the oocyte include those for heat-
shock protein^.^'
Despite a substantial amount of work, the mechanisms of transport of macromolecules
from the trophic cells to the oocyte remain to be totally defined. In polytrophic ovaries, the
total nurse cell cytoplasm flows through the ring canals into the oocyte during the final phase
of vitellogenesis. Electrophysiological studies have shown that electrophoresis may regulate
the distribution of charged molecules between the nurse cells and oocyte. The electrophoretic
current from nurse cells to oocyte is driven by the voltage gradient produced by an egg
chamber (see Section VI.A.3). Since the equilibrium potential of the nurse cells is several mV
more negative than that of the oocyte, macromolecules carrying a net negative charge may be
carried by electrophoresis to the oocyte. However, some authors have failed to demonstrate
intercellular electrophoresis in movement of materials in polytrophic ovaries. The potential
difference between nurse cells and oocyte, therefore, may serve primarily as a regulatory gate
effect rather than providing the principal force for macromolecule transport? In a variety of
polytrophic ovaries, the nurse cell cytoplasmic streaming can be reversibly inhibited by
cytochalasins, so it is likely that microfilament contraction plays some role in the cytoplasmic
streaming phenomenon, possibly by squeezing the nurse cell contents into the ~ o c y t e . ~ ~
However, mechanisms for capturing specific regionalized compounds in the oocyte must also
exist.
In telotrophic ovarioles, the distance between the place of synthesis of macromolecules (the
trophic cells in the tropharium) and the place of deposition (the growing oocytes in the
vitellarium) can be enormous (see Figure 2). Therefore, most workers have assumed that
Oogenesis and the Female Reproductive System 19

molecules are actively transported to the oocyte. After the discovery of a system of parallel
microtubules in the nutritive cords of heteropterans, suggestions were advanced that these
organelles may play a role in the active transp~rt.~ However, various species of polyphage
Coleoptera have nutritive cords that lack microtubules, and in other insect species, the
microtubules in the nutritive cords are randomly oriented. Another favored mechanism for the
transport of molecules involves electrophoresis and a flow of material assisted by differences
in hydrostatic pressure between the trophic area and the oocyte, which may be created by ionic
current asymmetries around the o ~ a r i o l e sBesides
.~~ active transport, peristaltic movement of
the musculature of the epithelia1 sheath which surrounds the ovariole may cause some
cytoplasm to flow from the tropharium to the oocytes. Additional work is required to resolve
all the components acting during macromolecule transport in polytrophic as well as in
telotrophic ovaries.

B. PREVITELLOGENESIS
The period when young oocytes enlarge by incorporation of nurse cell material (see Section
IV.A.4) is called the previtellogenic growth phase, or just previtellogenesis. Previtellogenesis
is difficult to investigate because it takes place chiefly in the penultimate oocyte during
vitellogenesis of the terminal oocyte and is thus simultaneously subject to its own control and
the control exerted on the terminal oocyte. Generally, previtellogenesis begins in young
adults, late in pupal development, in nymph, or last instar larvae. However, in insects with
panoistic ovarioles, previtellogenic growth begins during earlier nymphal stages. In those
insect species which hibernate as adults, growth of the oocytes may be stopped at the
beginning of previtellogenesis. The arrest and the onset of previtellogenesis are part of the
adult diapause and may be under hormonal control1 (see Chapter by Hardie).

C. VITELLOGENESIS
Vitellogenesis is the most important metabolic event in the adult life of the female insect.
The vitellogenic growth phase begins when yolk spheres are first observed to accumulate in
the oocytes. Vitellogenesis often occurs in the adult insect but also may take place earlier. For
example, in Sialisflavilatera (Megaloptera), vitellogenesis begins early in pupal life. Such a
shift into preadult stages will become necessary when the adult lives only a few days, as in
species with polytrophic meroistic ovaries that do not feed as adults. In many insects, however,
vitellogenesis and egg production is dependent on food availability. An extreme example of
a cyclic yolk production with feeding as an initial trigger for vitellogenesis is found, e.g., in
Aedes, Phormia, or Rhodnius (anautogenous insects).
Vitellogenesis involves the production of female-specific proteins termed vitellogenins
(vg) and their entry into the oocyte. When vitellogenin is taken into the oocyte, it is processed
to vitellin (vn). The vitellogenins are mostly produced in the fat body but may be also
produced in the ovary. They are transported by the hemolymph, in which their titer is high
during vitellogenesis, and accumulate in the oocyte against a concentration gradient 20-100
times their concentration in the hemolymph. In most species, vitellins comprise 60-90% of the
total soluble egg yolk protein. As noted above, vitellogenesis occurs in the terminal oocyte
within an ovariole, yet in many species the process is highly synchronized among ovarioles
and between ovaries. The synchronization results in a production of egg batches. In some
females, vitellogenesis in the penultimate oocyte appears to be inhibited even after the
terminal oocyte has completed its yolk deposition and has become chorinated, provided that
the mature egg is not laid.

1. Vitellogenin and Vitellin Chemistry

Data on yolk protein chemistry have expanded since new techniques of DNA analysis are
providing cloned cDNA and genomic-DNA with which the relationships between vitellogenin
genes and the final secretion products can be studied (see Section IV.C.2).
20 Insect Reproduction

In all insect species investigated, vgs and vns are irnrnunochemically identical. They are
glycolipophosphoproteins of native molecular masses ranging between 190 and 650 kDa and
often are composed of several polypeptides of variable sizes. In some species, endogenous
proteolytic cleavage changes the pattern of vn peptides compared to vgs. In Leucophaea
maderae, B. germanica, and L. migratoria, there seem to be large precursor molecules that are
proteolytically cleaved either in the fat body, hemolymph, or oocyte it~elf.4~ Also, subtle
differences in vg and vn lipid and carbohydrate moieties may exist. However, the differences
between vg and vn are small and, therefore, the chemistry of both polypeptides will be
discussed together.
Cloning the vitellogenin genes enables the putative amino acid sequences of the primary
products to be determined; but only complex protein chemistry will unravel the processing and
modification of these molecules during their secretion, transport, uptake, and depo~ition.4~
Harnish and White46have characterized the vitellins of a number of insect species and report
the existence of three definable groups. The largest group (group I) comprises insects from
several orders, including Ephemeroptera, Orthoptera, Dictyoptera, Hemiptera, Demaptera,
Coleoptera, and Lepidoptera. The native proteins are between 380 and 470 kDa, and upon
denaturation two distinctly different size classes of subunits are released. The high molecular
mass group ranges from 100-180 kDa, the low mass group from 43-86 kDa. The simplest
patterns exhibit one polypeptide in each size class, but several in each class is very common.
Group I1 vitellins occur in the orders Hymenoptera (e.g., Apis mellifera) and from the more
ancient dipterans, the suborder Tipolomorpha (Aedes aegypti). In denaturing SDS gels, group
I1 vitellins release only high molecular weight polypeptides. The third type (group 111) is found
only in higher Diptera (e.g., Drosophila, Calliphora, Lucilia). These proteins appear to have
molecular masses of about 200 kDa and are composed entirely of small polypeptides of about
50 kDa. The evidence for this schema, however, is not entirely convincing.
Carbohydrate has been found covalently linked to purified vgs and vns in every instance
it has been sought. The average carbohydrate content is 1-1 1%. In many cases, mannose and
glucosamine were the only identified sugars involved. In group I proteins, the oligosaccha-
rides are attached only to the heavy subunit. Lipids are also integral to all vgs and vns
characterized (7-15%). Phospholipids, diacylglycerides, and cholesterol comprise the bulk
lipid components. Vns appear to contain less lipids than vgs. In locust eggs, conjugated
ecdysteroids have been found noncovalently bound to vitellin.
Esterified phosphate is another integral part of some vgs and vns. In L. maderae vitellogenin,
the covalently attached phosphorus is distributed in an uneven fashion among the five
subunits. Phosphorylation of vitellogenin occurs posttranslationally in the fat body endoplas-
mic reticulum.
The fat body of vitellogenic mosquitoes was found to synthesize and secrete another
protein, in addition to vitellogenins, that accumulated in developing oocytes. This 53-kDa
protein is glycosylated, and immunoblot analysis demonstrated the immunological identity of
the 53-kDa polypeptides from the fat body and the ovary.47In eggs of some lepidopteran
insects, vitellin comprises only half of the total yolk proteins, and the yolk contains significant
amounts of other kinds of proteins. Silkworm (B. mori) eggs contain a vitellin (M, 420 kDa)
belonging to group I which consists of two heavy subunits (178 kDa) and two light subunits
(42 kDa). The second major yolk protein group is composed of non-sex-linked serum proteins
with a molecular mass of 30 kDa. They are also produced in the fat body, released into the
hemolymph, and finally sequestered into developing oocytes. Bombyx 30-kDa proteins are a
mixture of three monomers (29.5 to 32 kDa) and contain various lipids and carbohydrates. The
third main protein of silkworm eggs is the so-called egg-specific protein, which is produced
by the ovary itself and accumulates especially in developing oocytes. The egg-specific protein
is a trimer (225 kDa) of two heavy subunits (72 kDa) and one light subunit (64 kDa).48
Oogenesis and the Female Reproductive System 21

2. Vitellogenin Genes
A large amount of information has been developed in the last decade on vitellogenin gene
sequences. Most of the information is on the genes of Drosophila. In Drosophila, three distinct
genes coding for yolk proteins are present, namely, YP1, YP2, and YP3. The genes were
shown to be single copy, with the YPl and YP 2 closely spaced and YP3 approximately 1000
kilobases (kb) distant on the X chromosome. YP1 and YP3 each have a single species of
transcript, of about 1.6 and 1.54 kb, respectively. YP 2 produces transcripts of two sizes, 1.59
and 1.67 kb. All three genes have been ~ e q u e n c e dWith
. ~ ~ the availability of base sequences
for the yolk protein genes, more recent work has been concentrating on their regulation40(see
also Section IV.C.3).
A. aegypti and L. migratoria are two other insects in which the yolk protein genes have been
cloned. In both, the primary transcripts are very large, over 6000 nucleotides in length. The
results on A. aegypti suggest that there may be a total of five different vitellogenin genes.50
Sequence information and analysis of any gene control region are not yet available. In L.
migratoria, two genes, VgA and VgB, coding for vitellogenins have been identified, with only
little homology between them (at the 5' ends)." Both are located in the X chromosome. The
homologous regions in their 5' flanking sequences may be important for their control by
juvenile hormone.

3. Vitellogenin Synthesis
The fat body is the major, and in many cases, the only site of vitellogenin synthesis. Among
some of the Holometabola (Diptera, Lepidoptera, Coleoptera), however, the ovarian follicle
cells are also involved in yolk protein production. In the Diptera, the same structural genes,
synthesizing identical proteins, are active in both the fat body and the follicle cells.
Three cell types are commonly found in the insect fat body - trophocytes, urocytes, and
mycetocytes. The trophocyte is the principal cell type and it functions in a metabolic and
storage capacity. The cells are characterized by the presence of lipid droplets, protein spheres,
and glycogen granules in a metabolically active cytoplasm.16 The majority of the trophocytes
are found at the periphery of the fat body lobe. This distribution allows the trophocytes to
absorb or release products efficiently into the hernolymph.
In the majority of insects, juvenile hormone appears to be the key element in the control
of yolk protein production (see Chapter by H ~ d i e )The . ~ ~primary mode of action for this
hormone is at the fat body by initiating vitellogenin synthesis, with a secondary function in
the regulation of yolk uptake by the ovary (Section IV.C.5). The induction of vg synthesis by
JH in the fat body provides a system of hormonal control of gene expression. In response to
JH stimulation, the nucleus of the fat body trophocytes enlarges, while the cytoplasm develops
extensive rough ER and Golgi complexes. At the macromolecular level, the cells undergo
rapid synthesis of DNA, RNA, and protein in response to JH. In L. migratoria, a primary
stimulation by JH or JH mimics results in a rapid synthesis of rRNA, while the accumulation
of vitellogenin mRNA can be detected only after a lag phase. A second dose of JH leads to
a more rapid accumulation and translation of vg mRNA, but lowers the production of rRNA.
The picture of JH action obtained in insect fat body parallels the finding for steroid hormone-
stimulated vitellogenin synthesis in vertebrate systems.52
Since JH is a terpenoid and therefore different in chemical structure from that of a steroid
hormone, it is of interest to know whether a specific JH receptor is present in the cytosol and
nuclei of the target tissues. Cytosolic receptors may function in the translocation of the
hormone to nuclear acceptor molecules, the latter being essential for the initiation of gene
tran~cription.~~ Cytosolic fat body preparations of L. maderae adults contained a population
of JH binding compounds with a high affinity (K, ca. 1 nM),which could not be found in
nymphal tissues. A JH binding compound with similar affinity was extracted from nuclei of
22 Insect Reproduction

vitellogenic fat body cells of L. maderae. Putative juvenile hormone receptors have also been
identified in locust (L. migratoria) fat bodies. In the absence of JH, the adipokinetic hormone
(AKH-I), which is involved in mobilizing diglycerides, may inhibit vitellogenin gene expres-
sion in the locust fat body.54Signals from the ovary are supposed to terminate vitellogenin
synthesis in the fat body since ovariectomized females continue to produce vitellogenin,
which accumulates in the hemolymph. These signals may operate via modulating the activity
of the corpora allata.5s
The majority of insects conform more or less to the regulating scheme described above, but
some display significant variations. For example, among Coleoptera, while JH is necessary to
set off the initial vitellogenic response, continued yolk production then becomes autonomous.
In many Lepidoptera, vitellogenesis appears to be a part of a programmed developmental
response to metamorphosis. Among the Hymenoptera, honeybee queens show no dependence
on JH or ecdysone for the production of vitellogenins. In some Hemiptera, ecdysone seems
to be responsible for triggering elevated levels of yolk protein production.
The induction of vitellogenin synthesis is normally limited to adult females. It is possible,
however, to induce vitellogenin synthesis in adult males and in nymphs with large doses of
juvenile hormone or JH analogues. This has been demonstrated for L. migratoria and several
species of Di~tyoptera.~~ In male fat bodies of some Diptera, vitellogenin synthesis could be
induced by 20-hydroxyecdysone, and not by JH.57Female- and male-produced vitellogenins
may be different in their polypeptide compositions.
Details of ovarian yolk protein production are best understood for Drosophila. Using a
radioactive labeled probe containing the coding regions of yolk protein genes and in situ
hybridization techniques, it has been shown that the follicular epithelium is the specific site
of vitellogenin synthesis. The maximum level of yolk protein synthesis by the follicle cells
occurs in early vitellogenic stages. The follicular epithelium contributes ca. 35% of the yolk
proteins 1 and 2 to the total oocyte content, but only about 10% of the yolk protein 3
polypeptide. Since all three yolk proteins are transcribed at similar rates, yolk protein 3 mRNA
seems to be destabilized in the ovarian follicle cells, accounting for the reduction in its steady
state level. In the housefly Musca domestica, at the start of vitellogenesis the fat body appears
to be the main site of vitellogenin synthesis; later, the dominant role is taken over by the
ovaries. Overall, the follicle cell contribution of yolk proteins to the oocyte exceeds that of the
fat body.
As in the Diptera, the yolk proteins of the Lepidoptera are synthesized in the fat body and
the ovarian follicle cells. In Lepidoptera, however, the follicle cell vitellogenins are the
product of genes different from those responsible for vg synthesis in the fat body. In a moth,
Plodia interpunctella, fat body and follicle cell yolk proteins show no immunological cross-
reactivity, either as native proteins or as individual subunits.
Diptera also handle the hormonal regulation of vg synthesis somewhat differently from
most other insects (see above). 20-Hydroxyecdysone appears to be the main hormonal trigger
in the activation of the vitellogenin genes, but JH is involved in facilitating 20-hydroxyecdysone
action in the fat body and in the regulation of yolk protein uptake by the ovary58(see Chapter
by Hardie). In mosquitoes, an oostatic hormone may act to inhibit vitellogenin p r o d u c t i ~ n . ~ ~

4. Vitellogenin Secretion
The mechanism for the export of the yolk proteins both from the fat body and the follicle
cells involves the usual route through the Golgi and exocytosis at the plasma membrane. The
carbohydrate moieties may confer a certain degree of stability to the protein subunits, ensuring
proper assembly or preventing aggregation prior to secretion.60In a cockroach, B. gennanica,
the vg precursor accumulates but is not secreted when the animal is treated with tunicamycin.
Similar observations were done on the export of fat body proteins in Galleria mellonella
(Lepidoptera). In dipteran follicle cells, the export mechanism can be disrupted by colchicine
and other microtubule inhibitors, suggesting an important role of these cytoskeletal compounds.
Oogenesis and the Female Reproductive System 23

Yolk proteins excreted from the follicle cells are not normally liberated into the hemolymph
at large, but possibly are presented directly to the oocyte surface.

5. Uptake of Vitellogenin by the Ovary

Because of the heterosynthetic nature of vitellogenins in most insects, the oocytes are
highly specialized for the specific accumulation of the fat body vitellogenins. The pathway for
internalization of vitellogenins and other yolk protein precursors is similar in all types of
ovarioles. The onset of vitellogenic uptake is characterized by the formation of gaps and
spaces between the follicle cells (Figure 11). Many researchers have utilized various dyes and
tracers to demonstrate that this interfollicular route of uptake is universal among insects.
Vitellogenins appear to concentrate in the perioocytic space and from there are later taken up
by micropinocytosis of the oocyte membrane. The enlargement of the interfollicular channels
occurs at the onset of vitellogenesis and has been termed patency. In R. prolixus, this condition
has been shown to be a development in response to juvenile hormone. In other insects, JH also
seems to be involved in yolk protein uptake, but in these cases the precise role of the hormone
is not entirely clear (see Chapter by Hardie). The follicle cells lose patency at the time of
cessation of vitellogenesis and the deposition of the chorion.
Current evidence clearly points to yolk protein uptake being a receptor-mediated process.
The uptake is highly selective for vgs, saturable, species specific, energy dependent, and
sensitive to conditions of pH, temperature, and divalent cation concentrations (Ca2+).Studies
on binding of vg to membranes isolated from follicles have been made on only a few insects.
Binding under equilibrium conditions demonstrated the existence of a saturable, single class
of binding sites of follicle or ovary membranes (except for the cockroach Nauphoeta cinerea,
where two separate binding sites for vg have been suggested). The dissociation constant (K,)
varied between 100 nM (for L. migratoria) to 13 nM (for Manduca sexta). In L. migratoria,
the vitellogenin receptor protein has been isolated and purified by immunaffinity chroma-
t~graphy.~' It is an acidic, negatively charged (p1 = 3.5) 180 kDa glycoprotein (nonreducing
conditions) with large amounts of N- and 0-linked oligosaccharides,among them neuraminic
acid, which has been found to be essential for receptor function. The vg receptor was localized
in oocyte membranes and in endocytotic vesicles. So far, vg receptors of two other insects
have been visualized using the ligand blotting technique, from N. cinerea (M, = 200 kDa) and
A. aegypti (M, = 205 kDa).62The fact that the receptors bind vg only when separated under
nonreducing conditions suggests that disulfide bonds within the receptor molecule are neces-
sary for its biological activity.
Receptor-bound vitellogenins accumulate in specialized regions of the surface membrane,
the coated pits, and are then internalized by the formation of coated vesicles (Figure 12).
Coated vesicles are suggested to be universal organelles for a specific macromolecular
transport in eukaryotic cells. The characteristic feature of the coated vesicles is an outer
proteinaceous polyhedral cage enclosing its membrane. The major protein of this cage (50%
of the total protein) is "clathrin." Three molecules of clathrin heavy chain (M, 180 kDa)
together with three molecules of clathrin light chain (33 and 36 kDa) form a subunit of the coat
called triskelion. The triskelions are self-assembledinto the polyhedral cage on the membrane
of the coated pits. Once the coated vesicle is formed and pinched off the plasma membrane,
rapid uncoating follows, releasing the vesicle and allowing the coat components to recycle to
the plasma membrane.62 The released vesicles fuse with endosomes. Within the endosomal
compartment, the adsorbed yolk precursors dissociate from the membrane to become homo-
geneously distributed in the lumen. As a final transformation, the transitional yolk body
changes into a mature yolk body with the crystallization of the vitellins. Mature yolk bodies
will store the yolk proteins until initiation of embryonic development, during which they are
utilized.
The developmental processes of vitellogenin synthesis and vg uptake by the maturing
ovaries occur independently of the presence of conspecific males. However, in Drosophila,
24 Insect Reproduction

FIGURE 11A

FIGURE 11B

FIGURES 11A and B. Follicular epithelium in Blatfella germanica oocytes. Sections at the equatorial zone in
oocytes from 3-day-old (A) and 5-day-old (B) females with large intercellular spaces (arrows) in (B). Photographs
courtesy of X. Belles, Barcelona.
Oogenesis and the Female Reproductive System

FIGURE 12. Schematic representation of the vitellogenin internalization pathway in mosquito oocyte: cl, clathrin;
cp. coated pit; CV,coated vesicle; end, vesicular endosome; FC, follicle cell; Itc, tubular endosome labeled positively
with anti-vg antibodies; mv, microvilli; myb, mature yolk body; rc, receptor; tyb, transitional yolk body; utc, tubula~
compartment labeled negatively with anti-vg antibodies; vg, vitellogenin. (Reproduced, with permission, from the
Annual Review of Entomology, Vol. 37, p. 217, 01992 by Annual Reviews Inc.)

it has recently been shown that mature males significantly accelerate the onset of vitellogen-
esis, and thus ovarian maturation overall, by about 4 days. Although the proximate stimulus
is not known, it is conceivable that social signals from the males (during courtship attempts)
elicit changes in the hormonal levels of females, thereby initiating the events leading to
vitellogenesis. Again, this effect of male behavior on female reproductive biology is similar
to that reported in vertebrate^.^^

D. CHORIONIZATION
When vitellogenesis is completed, the vitelline membrane and, later, the chorion (eggshell)
are formed (Figure 13).
26 Insect Reproduction

FIGURE 13. Follicle cells from a basal oocyte of Blattella germanica at late chorion formation. The perioocyte
the innerchorion layer(1CL) and the complex outer chorion layer (OCL)
space contains the vitelline membrane (VM),
showing columnar projections (P). IS, intercellular space; L, lipid droplet. Photograph courtesy of X. Belles,
Barcelona.

1. The Vitelline Membrane

Early observations suggested that the vitelline membrane was produced by the oocyte
itself. This may be the case in some species, for example, in grasshoppers and Lepid~ptera.~
However, more recent studies have demonstrated that the follicle cells secrete material which
forms the vitelline membrane in a large number of insects. Although differences in details of
this process have been encountered in all of the systems, vitelline membrane production is
preceded by an extensive hypertrophy of rough ER and formation of Golgi complexes,
followed by an accumulation of secretory granules (vitelline membrane body precursors) in
the apical zones of the follicle cells. In Simulium vittatum and other Diptera, the build-up of
the ER and Golgi vesicles occurs at midvitellogenesis.By the time that vitellogenesisis almost
complete, the Golgi complexes, containing both dense and fibrous materials, become even
more prominent, and the previtelline membrane secretory substance begins to be deposited
between the follicle cells and the oocyte. The secreted material then begins to coalesce and
gradually forms the vitelline membrane (see Figure 13). Most of the ultrastructural observa-
tions of vitelline membrane formation in exopterygotes support the above outline for the
process.
Electron microscopical studies on vitelline membrane formation in L. migratoria showed
that this structure is composed of two ultrastructurally distinguishable components, the
vitelline membrane bodies (VMB) and a fine granular material which eventually cements the
VMBs together to give the complete vitelline membrane. The observation that, in L. migratoria,
the first condensation of VMBs occurs in the vicinity of the oocyte membrane may indicate
that the oocyte is the source of VMB secretion, whereas the second vitelline membrane
component is clearly the product of the follicle cells. Other support for a dual origin of the
vitelline membrane has come from studies on Lepidoptera. In the butterfly Calpodes ethlius,
a distinct vitelline membrane is detectable at the end of the yolk uptake phase. It has an
electron-dense layer away from the oocyte and an electron-lucent layer apposing the oocyte.
Both the oocyte and the follicle cells contain coated vesicles, which appear to be in the process
of exocytosis. The electron-dense material seems to be exocytosed from the follicle cells,
whereas coated vesicles at the oocyte surface exocytose the electron-lucent granular material.
Oogenesis and the Female Reproductive System

micropylar
openings

inner opening

C vitelline membrane

---...-. .-..
endochorion

~ m i c r o n v l a r canal

FIGURE 14. Egg of Locusfa migraforia with a section through the chorion along the micropylar axis. (From
Gillott, C., Entomology, Plenum, New York, 1980, chap 19. With permission.)

Once fully formed, the vitelline membrane is completely electron dense.64 A definitive
decision on the source of the various contributors to the vitelline membrane cannot be made
until it can be demonstrated, with molecular biological techniques now becoming available,
that definitive vitelline membrane proteins are synthesized by the oocyte.

2. Chorion Formation
The chorion is usually secreted entirely by the follicle cells and can be seen to comprise
two main layers, an endochorion adjacent to the vitelline membrane and an exochorion
(Figures 13 and 14). In some insects, e.g., Acrididae, the shell takes on a third layer, the
extrachorion, as an oocyte moves through the common oviduct. Although the follicle cells are
mesodermal derivatives, the chorion is cuticle-like in nature and contains layers of proteins
and lipoproteins, some of which are tanned by polyphenolic substances released by the cells.
With chorion secretion, the follicle cells complete their duties and then die.
At the physiological level the chorion functions in protecting the oocyte from mechanical
stresses, such as from predators, as well as environmental stresses, such as dessication and
drowning, while at the same time permitting gas exchange and sperm ~enetration.~~ In some
species, a wax layer is formed immediately above the vitelline membrane by the coalescence
of oil droplets secreted by the follicle cells which renders the chorion waterproof. Viewed
from the perspective of the oocyte, the next structure is the basal or inner chorion layer
(crystalline chorion layer). The crystalline chorion layer, although flexible, puts a limit to the
volume that the oocyte can achieve. Speculations on other functions include a role in the
confinement of the wax layer, as well as allowing for gas exchange, through plastron
28 Insect Reproduction

respiration or directly. Distal to the basal chorion is the trabeculate layer which is considered
to be a part of the endochorion. The trabeculate layer is characterized by the presence of
cavities and pores. The small pores may be formed by the withdrawal of follicle cell processes
during andlor after the deposition of this layer. The cavities may interconnect and form
extensive channels. The channels may serve as air spaces and open to the exterior via
aeropyles (gas exchange). In other cases, the cavities are filled with a mucus-like substance
which serves as a reserve to surface-localized adhesive material used to attach eggs to the
substrate. The outermost layer of the endochorion, if present, is characterized by the presence
of lamellae, based on the helicoidal arrangement of stacks or fibrils. The lamellar layer may
be traversed by pores to the exterior. In many Lepidoptera, the lamella layer serves as the outer
portion of the shell. In most other insects, however, an exochorion is present. The exochorion
consists largely of mucoprotein and contains polysaccharides. In some insects, a ring of
follicle cells near the anterior end of the oocyte secretes no exochorion, so that a line of
weakness is created at this point, which facilitates hatching. Also, certain follicle cells appear
to have larger than normal microvilli which, when withdrawn after chorion formation, leave
channels to permit entry of sperm (micropyles; see Figure 14).
The molecular analysis of chorion formation has advanced rapidly. Recent reviews on the
biochemistry of chorion proteins, chorion gene structure, and chorion gene expression were
presented by Regier and K a f a t o ~and~ ~by Kaulenas9
In general, it is assumed that choriogenesis is independent of hormonal control and is
initiated at an appropriate late stage of vitellogenesis in response to local signals. However,
some information is available that shell formation might depend on brain neurohormones,
juvenile hormone, and 20-hydro~yecdysone.'.~~,~~ In L. migratoria, ecdysteroids are synthe-
sized by the epithelium of the follicle cells at around the time of chorion formation during a
short period of 8-12 h. However, ecdysteroid synthesis in follicle cells can also occur earlier,
particularly in ovoviviparous or viviparous species, such as in cockroaches and in Glossina.
Besides helping to regulate chorionization, at least part of these ecdysteroids enter the oocyte,
where they are mostly present as polar or apolar conjugates and seem to control the first events
in embryogenesis. In a cricket, G. bimaculatus, coincident changes in ecdysteroid production
for both the ovaries and the abdominal integument were observed.68The role of epidermal
ecdysteroids during oogenesis is not yet clear.

V. OVULATION AND OVIPOSITION

Ovulation includes the opening of both the oocyte follicle and interfollicular tissue, and
involves contractions of ovarioles, pedicels, and oviducts resulting in the expulsion of the egg
from the ovary into the oviducts. The follicle then collapses and the follicular cells undergo
autolysis. The degenerating follicular cells are generally referred to as a "corpus luteum", and
structures of this type have been described in several insect species.69Ovulation may occur
just before oviposition, or a larger period of time may elapse between the two processes.
Several species, including cockroaches and flies, are ovoviviparous and retain their eggs in the
genital pouch without supplying them with anything but water. Viviparous insects, on the
other hand, supply food to their progeny, either from modified follicle cells or from the
accessory glands.
Although in many insects ovulation is rapidly followed by oviposition, there are indications
that the two processes are separately'controlled. In R. prolixus, ovulation is a result of ovarian
motility induced by a neurosecretory myotropin released from the corpus cardiacum. The
hormone is released in mated females when mature eggs are present in the ovary, as signaled
by an increase in ecdysteroid titer in the hemolymph. In other species, ovulation seems to be
controlled in a similar way.
Oogenesis and the Female Reproductive System 29

In some insects, oocyte resorption, termed oosorption, may take place under various
unfavorable environmental conditions. Starvation or the lack of food, mating, or a suitable
place to oviposit are frequent causes, but factors such as temperature and change in photope-
riod, social life, or maternal care also induce oosorption. In general, oosorption occurs when
the external factors do not allow either the survival of eggs and larvae or egg deposition.*O
Oosorption may occur either in young previtellogenic oocytes or in vitellogenic oocytes, and
even in chorionated eggs. Oocytes may grow to a certain size and then stop, while the follicle
cells begin to change from cuboidal to irregular shapes. The transport of material from the
interfollicular spaces to the oocyte ceases, owing to the breakdown of the microvilli of the
oocytes and follicle cells. Hydrolytic enzymes produced in the follicle cells cause oocyte lysis,
breaking down first the protein and then the lipid yolk globules, penetrating the oocyte and
finally themselves degenerating. Often it can be observed that only certain follicles degener-
ate, while others continue to develop. The appearance of oosorption processes is apparently
caused by a decrease in the activity of the corpora allata, but the brain certainly intervenes too,
by regulating corpora allata functioning and/or acting humorally on the ovary.

VI. CONCLUDING REMARKS

The development of the female reproductive system is reasonably well understood at the
structural or morphological level across a wide range of insect species. Details of the molecu-
lar mechanisms involved in the development processes, and especially of the genetic control
of gene expression, have been explored in a much lower number of insects. Insect oogenesis
comprises many stages which are regulated by certain humoral factors, including neurohor-
mones, juvenile hormone(s), and ecdysteroids, whose importance and modes of intervention
vary depending on the species. Detailed knowledge on female reproductive biochemistry,
physiology, and endocrinology in a great many insects will be necessary, e.g., to provide a
basis for using insect hormones, hormone agonists, or hormone antagonists as "third genera-
tion pesticides" in insect pest control. Beyond that, ovarian development and oogenesis are
affected by environmental factors, including temperature, humidity, photoperiod, or the
finding of food. Therefore, it seems impossible to study the physiology and endocrinology of
insect reproduction without taking into account its ecological conditions.

REFERENCES
1. Raabe, M., Recent Developments in Insect Neurohormones. Plenum, New York, 1989, chap. 6.
2. Truckenbrodt, W., ijber die imaginale OvarvergroBerung im Zusammenhang mit der Physogastrie bei
Odontotermes badius Haviland (Insecta, Isoptera), Insectes Soc., 20, 21, 1973.
3. Gottanka, J. and Biining, J., Mayflies (Ephemeroptera), the most "primitive" winged insects, have telotrophic
meroistic ovaries, Wilhelm Roux Arch. Dev. Biol., 203, 18, 1993.
4. Gillott, C., Entomology, Plenum, New York, 1980, chap. 19.
5. Berry, S. J., Reproductive systems, in Fundamentals of Insecr Physiology. Blum, M. S., Ed., John Wiley &
Sons, New York, 1985, chap. 1 1 .
6. King, R. C. and Biining, J., The origin and functioning of insect oocytes and nurse cells, in Comprehensive
Insect Physiology, Biochemistry and Pharmacology, Vol. 1, Kerkut, G. A. and Gilbert, L. l., Ms., Pergamon,
Oxford, 1985, chap. 3.
7. Wigglesworth, V. B., The distribution of aeriferous tracheae for the ovaries of insects, Tissue Cell, 23, 57,
1991.
8. Biining, J., Germ cell cluster formation in insect ovaries, Int. J. Insect Morphol. Embryol., 22, 237, 1993.
9. Kaulenas, M. S., Insect Accessory Reproductive Structures. Function. Structure, and Development, Springer,
Berlin, 1992.
30 Insect Reproduction

10. Storto, P. D. and King, R. C., The role of polyfusomes in generating branched chains of cystocytes during
Drosophila oogenesis, Dev. Genet.. 10, 70, 1989.
11. King, R. C., Cassidy, J. D., and Rousset, A., The formation of clones of interconnected cells during
gametogenesisin insects, in Insect Ultrastructure, Vol. 1, King, R. C. and Akai, H., Eds., Plenum, New York,
1982, 3.
12. Dorn, A., Rademacher, J. M., and Sehn, E., Ecdysteroid-dependent development of the oviduct in last-
instar larvae of Oncopeltus fasciatus, J. Insect Physiol., 32,643, 1986.
13. Davey, K. G., The female reproductive tract, in Comprehensive Insect Physiology, Biochemistry and Phar-
macology, Vol. 1, Kerkut, G. A. and Gilbert, L. I., Eds.. Pergamon, Oxford, 1985, chap. 2.
14. Szopa, T. M. and Happ, G. M., Cytodifferentiation of the accessory glands of Tenebrio molitor. IX.
Differentiation of the spermathecal accessory gland in vitro, Cell Tissue Res., 222, 269, 1982.
15. Degrugillier, M. E., In vitro release of house fly, Musca domestica L. (Diptera: Muscidae), acrosomal
material after treatment with secretion of female accessory gland and micropyle cap substance, Int. J. lnsect
Morphol. Embryol., 14, 381, 1985.
16. Koeppe, J. K., Fuchs, M., Chen, T. T., Hunt, L.-M., Kovalick, G. E., and Briers, T., The role of juvenile
hormone in reproduction, in Comprehensive Insect Physiology. Biochemistry and Pharmacology, Vol. 8,
Kerkut, G. A. and Gilbert, L. I., Eds., Pergamon, Oxford, 1985, chap. 6.
17. Burns, E. L., Chiang, A.-S., Gadot, M., and Schal, C., Juvenile hormone regulation of the left colleterial
gland in intact and ovariectomized Blattella germanica L. (Dictyoptera: Blattellidae), J. Insect Physiol.. 37,
401, 1991.
18. Pau, R. N., Weaver, R. J., and Edwards-Jones, K., The regulation of cockroach oothecin synthesis by
juvenile hormone, Arch. Insect Biochem. Physiol. (Suppl.), 1, 59, 1986.
19. Ter Wee, T. J. and Stay, B., Termination of pregnancy and milk synthesis in the viviparous cockroach
Diploptera punctata: a role of juvenile hormone, Int. J. Invertebr. Reprod., 11, 59, 1987.
20. Raabe, M., Insect Reproduction: Regulation of successive steps, in Advances in Insect Physiology, Vol. 19,
Evans, P. D. and Wigglesworth, V. B., Eds., Academic Press, London, 1986.29.
21. Technau, G. M. and Campos-Ortega, J. A., Lineage analysis of transplanted individual cells in embryos
of Drosophila melanogaster. 111. Commitment and proliferative capabilitiesof pole cells and midgut progeni-
tors, Wilhelm Roux Arch. Dev. Biol.. 195. 489, 1986.
22. Hay, B., Jan, L. Y., and Jan, Y. N., Localization of vasa, a component of Drosophila polar granules, in
maternal-effect mutants that alter embryonic anteroposterior polarity, Development, 109, 425, 1990.
23. Gutzeit, H. O., von Seydlitz-Kunbach, E., and Neuschriier, R., How Drosophila follicles become spatially
organized and obtain their ovoid shape, hr. J. Insect Morphol. Embryol. 22, 335, 1993.
24. Hatakeyama, M., Sawa, M., and Oishi, K., Ovarian development and vitellogenesis in the sawfly, Arhalia
rosae ruficornis Jakovlev (Hymenoptera, Tenthridinidae), Invertebr. Reprod. Dev., 17, 237, 1990.
25. Buning, J. and Sohst, S., The flea ovary: ultrastructure and analysis of cell clusters, Tissue Cell, 20, 783,
1988.
26. Telfer, W. H., Development and physiology of the oocyte-nurse cell syncytium, Adv. Insect Physiol.. 11,223,
1975.
27. Gutzeit, H. O., Die Entwicklung der Eizelle bei Insekten. Ein Modellsystem fur die Entstehung von
Zellpolaritiit, Biol. Zeit. 20, 33, 1990.
28. Bohrmann, J., Frey, A., and Gutzeit, H. O., Observations on the polarity of mutant Drosophila follicles
lacking the oocyte, Wilhelm Roux Arch. Dev. Biol., 201, 268, 1992.
29. Lanot, R., Roussel, J.-P., and Thiebold, J. J., Ecdysteroids and meiotic reinitiation in oocytes of Periplaneta
americana (Dictyoptera) and Gryllus bimaculatus (Orthoptera), Invertebr. Reprod. Dev., 15, 69, 1989.
30. St. Johnston, D. and Nusslein-Volhard, C., The origin of pattern and polarity in the Drosophila embryo,
Cell, 68, 201, 1992.
3 1. Gutzeit, H. O., Transport of molecules and organelles in meroistic ovarioles of insects, Differentiation, 3 1,
155, 1986.
32. Overall, R. and Jaffe, L. F., Patterns of ionic current through Drosophila follicles and eggs, Dev. Biol., 108,
102, 1986.
33. De Loof, A., The meroistic insect ovary as a miniature electrophoresis chamber, Comp. Biochem. Physiol.
Sect. A, 74, 3, 1983.
34. Berry, Y. S., Maternal direction of oogenesis and early embryogenesis in insects, Annu. Rev. Entomol.. 27,
205, 1982.
35. Cardoen, J., Schoofs, L., Broekaert, D., van Mellaert, H., Verachtert, B., and de Loof, A., Polyploidization
and localisation of poly (A)' RNA in the different cell types of the vitellogenic meroistic ovary of the fleshfly,
Sarcophaga bullata. Histochenlistry, 85, 305, 1986.
36. Hammond, M. P. and Laird, C. D., Chromosome structure and DNA replication in nurse and follicle cells
of Drosophila melanogaster. Chromosoma, 91, 267, 1985.
37. Dittmann, F., Horner, R., and Engels, W., Endoploidization of tropharium nuclei during larval development
and the first gonocycle in Dysdercus intermedius (Heteroptera),Int. J. Invertebr. Reprod., 7, 279, 1984.
Oogenesis and the Female Reproductive System 31

38. Dittmann, F., Steinbriick, G., and Miinz, A., Amplification of tropharium rDNA in the telotrophic ovariole
of the bug, Dysdercus intermedius. Invertebr. Reprod. Dev., 17, 9, 1990.
39. Sander, K. L., Specification of the basic body pattern in insect embryogenesis, Adv. Insect Physiol.. 12, 125,
1976.
40. Frohnhofer, H. G. and Niisslein-Volhard, C., Organization of anterior pattern in the Drosophila embryo by
the maternal gene bicoid, Nature (W.), 324, 120, 1986.
4 1. Ambrosio, L. and Schedl, P., Gene expression during Drosophila melanogaster oogenesis: analysis by in situ
hybridization to tissue selections, Dev. Biol.. 105, 80, 1984.
42. Gutzeit, H. O., The role of microfilaments in cytoplasmic streaming in Drosophila follicles, J. Cell Sci., 80,
156, 1986.
43. Diehl-Jones, W. and Huebner, E., Pattern and composition of ionic currents around ovarioles of the
hemipteran, Rhodnius prolixus (Stahl), Biol. Bull., 176(S), 86, 1989.
44. Della-Cioppa, G. and Engelmann, F., The vitellogenin of Leucophaea maderae. Synthesis as a large
phosphorylated precursor, lnsect Biochem., 17,401, 1987.
45. Bownes, M., Expression of the genes coding for vitellogenin (yolk protein), Annu. Rev. Enfomol., 31, 507,
1986.
46. Harnish, D. G. and White, B. N., lnsect vitellins: identification,purification, and characterization from eight
orders, J. Exp. Zool., 220, 1, 1982.
47. Hays, A. R. and Raikhel, A. S., A novel protein produced by the vitellogenic fat body and accumulated in
mosquito oocytes, Wilhelm Roux Arch. Dev. Biol., 199, 114, 1990.
48. Zhu, J., Indrasith, L. S., and Yamashita, O., Characterization of vitellin, egg-specific protein and 30 kDa
protein from Bombyx eggs, and their fates during oogenesis and embryogenesis, Biochem. Biophys. Acta, 882,
427, 1986.
49. Yan, Y. L., Kunert, L. J., and Postlethwait, J. H., Sequencehomologies among the three yolk polypeptides
(Yp) genes in Drosophila melanogaster, Nucleic Acids. Res., 15, 67, 1987.
50. Gemmill, R. M., Hamblin, M., Glaser, R. L., Racioppi, J. V., Marx, J. L., White, B. N., Calvo, J. M.,
Wolfner, M. F., and Hagedorn, H. H., Isolation of mosquito vitellogenin genes and induction of expression
by 20-hydroxyecdysones, Insect Biochem., 16, 761, 1986.
51. Locke, J., White, B. N., and Wyatt, G. R., Cloning and 5' end nucleotide sequences of two juvenile
hormone-inducible vitellogenin genes of the African migratory locust, DNA. 6, 331, 1987.
52. Hagedorn, H. H. and Kunkel, J. G., Vitellogenin and vitellin in insects. Annu. Rev. Entomol.. 24,475, 1979.
53. Engelmann, F., Regulation of vitellogenesis in cockroaches, in Cockroaches as Models for Neurobiology:
Applications in Biomedical Research. Vol. 2, Huber, I., Masler, E. P,, and Rao, B. R., Eds., CRC, Boca Raton,
1990, 159.
54. Moshitzky, P. and Applebaum, S. W., The role of adipokinetic hormone in the control of vitellogenesis in
locusts, Insea Biochem., 20, 319, 1990.
55. Kempa-Tomm, S., Hoffmann, K. H., and Engelmann, F., Vitellogenins and vitellins of the Mediterranean
field cricket, Gryllus bimacularus: isolation, characterization and quantification,Physiol. Enfomol.. 15, 167,
1990.
56. Lamy, M., Vitellogenesis,vitellogenin and vitellin in the males of insects: a review, Int. J. Invertebr. Reprod.
Dev., 7, 31 1, 1984.
57. Huybrechts, R. and de Loof, A., Similarities in vitellogenins and control of vitellogenin synthesis within the
genera Sarcophaga, Calliphora, Phormia and Lucilia (Diptera), Comp. Biochem. Physiol., 72B, 339, 1982.
58. Kelly, T. J., Adams, T. S., Schwartz, M. B., Birnbaum, M. J., Rubenstein, E. C., and Imberski, R. B.,
Juvenile hormone and ovarian maturation in the Diptera: a review of recent results, lnsect Biochem., 17, 1089,
1987.
59. Borovsky, D., Thomas, B. R., Carlson, D. A., Whisenton, L. R., and Fuchs, M. S., Juvenile hormone and
20-hydroxyecdysone as primary and secondary stimuli of vitellogenesis in Aedes aegypti. Arch. Insect
Biochem. Physiol., 2, 75, 1985.
60. Kunke, J. G. and Nordin, J. H., Yolk proteins, in Comprehensive Insect Physiology, Biochemistry and
Pharmacology, Vol. 1, Kerkut, G. A. and Gilbert, L. I., Eds., Pergamon, Oxford, 1985, chap. 4.
61. Ferenz, H.-J., The locust oocyte vitellogenin receptor-function and characteristics, in Advances in Inverte-
brate Reproduction, Vol. 5, Hoshi, M. and Yamashita, O., Eds., Elsevier, Amsterdam, 1990, 103.
62. Raikhel, A. S. and Dhadialla, T. S., Accumulation of yolk proteins in insect oocytes, Annu. Rev. Entomol.,
37, 217, 1992.
63. Craddock, E. M. and Boake, C. R. B., Onset of vitellogenesis in female Drosophila silvestris is accelerated
in the presence of sexually mature males, J. Insect Physiol., 38, 643, 1992.
64. Griffith, C. M. and Lai-Fook, J., Structure and formation of the chorion in the butterfly, Calpodes. Tissue
Cell. 18, 589, 1986.
65. Regier, J. C. and Kafatos, F. C., Molecular aspects of chorion formation, in Comprehensive Insect
Physiology, Biochemistry and Pharmacology, Vol. 1, Kerkut, G. A. and Gilbert, L. I., Eds., Pergamon,
Oxford, 1985, chap. 5.
32 Insect Reproduction

66. Pascual, N., Cerdh, X., Benito, B., T o m h , J., Piulachs, M. D., and BeliCs, X., Ovarian ecdysteroid levels
and basal oocyte development during maturation in the cockroach Blattella germanica (L.), J. lnsect Physiol.,
38, 339, 1990.
67. BellCs, X., Cassier, P., Cerda, X., Pascual, N., AndrC, M., Rosso, Y., and Piulachs, M. D., Induction of
choriogenesis by 20-hydroxyecdysone in the German cockroach, Tissue Cell, 25, 195, 1993.
68. Weidner, K., Clan, M., Rieck, H., and Hoffmann, K. H., Developmental changes in ecdysteroid biosyn-
thesis in virro during adult life and embryogenesis in a cricket, Gryllus bimaculatus de Geer, Invertebr.
Reprod. Dev., 21, 129, 1992.
69. Griffith, C. M. and Lai-Fook, J., Corpus luteum formation and ovulation in the butterfly Calpodes, Tissue
Cell, 18, 783, 1986.
70. Wigglesworth, V. B., The Principles of Insect Physiology, Chapman and Hall, London, 1972, chap. 15.
 Chapter 2

INSECT MALE MATING SYSTEMS

Cedric Gillott

CONTENTS
I . Introduction .................................................................................................................
33

I1. Structure ......................................................................................................................

34
A. Internal Organs ..................................................................................................... 34
B . External Genitalia ................................................................................................. 36
C. Specialized Mating Systems ................................................................................. 37

I11. Development ................................................................................................................ 37

A. Embryonic Development ....................................................................................... 37
B . Postembryonic Development ................................................................................38
1. General ............................................................................................................. 38
2. Endocrine Regulation ....................................................................................... 38

IV. Functional Aspects ......................................................................................................39

A. Spermatogenesis and Sperm Storage ................................................................... 39
1. Spermatogenesis............................................................................................... 39
2. Sperm Release and Storage ............................................................................. 40
B . Accessory Gland Activity ..................................................................................... 40
1. Nature and Formation of Secretion ................................................................. 41
2. Endocrine Regulation and Effect of Mating ................................................... 41
C. Spermatophore Formation .................................................................................... 42
D. Seminal Fluid ........................................................................................................44
E. Fecundity-Enhancing and Receptivity-Inhibiting Chemicals .............................. 44
1. Source and Nature ............................................................................................ 45
2. Site and Mode of Action .................................................................................46
F. Other Functions .....................................................................................................47

Acknowledgments .................................................................................................................
49

References .............................................................................................................................
49

.
I INTRODUCTION
Among the problems that required solution in the evolution of the Insecta as a predominantly
terrestrial group was the bringing together of sperm and egg in the absence of an aquatic
environment. The solution came through the formation. in most insects. of a spermatophore
within which the sperm could be safely transferred to the female reproductive tract. avoiding
the risk of both desiccation and predation . In relatively few species. spermatophores are not
found and sperm transfer is achieved by means of an intromittent organ . Though sperm
production and transfer are the primary functions of the male reproductive system. a number

.
0-8493-6695-X/95/S0.00+$.50
O 1995 by CRC Press Inc.
34 Insect Reproduction

of secondary functions have evolved, including sperm storage, the generation of "signals,"
either physical or chemical, that induce significant changes in the female's fecundity andlor
receptivity, and the transfer of nutrients to the female.
This chapter will provide the structural and functional background for ensuing chapters that
deal with more specific aspects of male insect reproductive biology. In addition, it will focus
on selected aspects of the functions of the male reproductive system that are not covered
elsewhere but are of particular interest to the author. In keeping with the extreme morphologi-
cal diversity of the Insecta, the structural, physiological, and biochemical nature of the male
system is widely varied. It is not the purpose of this chapter to encyclopedically describe this
plethora of detail, but rather to note the generalities that have been established and to point
out areas where understanding is still weak.

11. STRUCTURE
A. INTERNAL ORGANS
The male internal organs comprise paired testes (fused to form a single median structure
in Lepidoptera), paired vasa deferentia and seminal vesicles, a median ejaculatory duct, and
in most species, accessory glands of varied origin and complexity (Figure 1).
Within each testis is a varied number of tubular follicles bound together by a connective
tissue sheath. Each follicle connects with a short vas efferens, the vasa efferentia from each
gonad opening either confluently or in a linear sequence into the vas deferens. Within each
follicle, groups of germ cells in various stages of spermatogenesis (see Section 1V.A) may be
seen in final instar larvae or adults. As well, glandular cells may occur which, on the basis of
their staining properties, have been proposed to produce the nourishment necessary for both
maturation of the sperm and their maintenance (storage) within the seminal vesicle. The
glandular cells commonly take the form of an ensheathing epithelium around the developing
germ cells, the whole structure being known as a "cyst." Though the cyst wall usually breaks
open in the final stages of spermatogenesis, the sperm within may remain as a bundle even
after insemination. In Acrididae (Orthoptera), one cyst cell differentiates as a "nurse cell"
during the spermatid stage.' The spermatids in each bundle then become oriented and embed
their heads into the cell, which produces a large quantity of mucoprotein, the sperm and
mucoprotein cap constituting a spermatodesm. Cantacuzbnel speculates that the cap does not
serve directly as a source of energy for the sperm while they are stored in the seminal vesicle;
rather, the enzymes which, she suggests, reside within it degrade the nutrients released by the
epithelium which lines the seminal vesicle. The literature also contains a number of reports
that implicate the testes as the site of production of chemicals that modify female fecundity
andlor receptivity, but these are more appropriately dealt with in Section 1V.E.
The vasa deferentia are typically assumed to be merely tubes for conducting the sperm from
the testes to the seminal vesicles; indeed, the latter are frequently dilations of the vasa
deferentia. However, light micro~copical~-~ and a few electron m i c r o s ~ o p i c a lstudies
~ - ~ ~ sug-
gest that these tubes have important glandular and phagocytic functions in some species.
Rojas-Rousse2and Gerber et aL5propose that the secretion is used to nourish the sperm within
the male reproductive tract, while the presence of numerous lysosomes in the vas deferens
cells of Drosophila melanogaster may indicate a role in digestion of aged or degenerate
sperm.ll This function is also proposed for the phagocytic cells in the upper vas deferens of
the lepidopterans Anagasta kiihniella8and Calpodes ethlius.12Other proposed functions of the
vas deferens secretion include involvement in spermatophore production in Trichoptera13and
the blister beetle, Lytta nuttalli,14 and production of the spermatodesm in Tettigoniidae
(Orth~ptera).~
In the great majority of insects, the ejaculatory duct is of ectodermal origin and, as such,
has a cuticular intima. However, in all Lepidoptera, Diadromus pulchellus (Hymenoptera),
Plecia nearctica (Diptera), and Nezara viridula (Herniptera) an anterior mesodermal component
Insect Male Mating Systems

FIGURE 1. Representative male reproductive systems (not to scale). A. Melanoplus sanguinipes (Orthoptera);
B . Tenebrio molitor (Coleoptera); C. Musca domestica (Diptera); D. Anagasta kiihniella (Lepidoptera). Abbrevia-
tions: AG, accessory glands; BAG, bean-shaped accessory gland; CS, cuticular simplex; D, duplex; ED, ejaculatory
duct; ES, ejaculatory sac; LHT, long hyaline tubule; LVD, lower vas deferens; SHT, short hyaline tubules; SV, seminal
vesicle; T, testis; TF, testis follicles; TAG, tubular accessory gland; UVD, upper vas deferens; VD, vas deferens;
WT. white tubules; 1-8, Eight regions of the noncuticular simplex. (A, original; B, redrawn from Dailey, P.J. et al.,
Journal ofMorphology, Vol. 166. Copyright O 1980John Wiley & Sons, Inc. Reprinted by permission of John Wiley
& Sons, Inc. C, redrawn from Luther S. West: The Housefly: Its Natural History. Medical Importance, and Control.
Copyright O 1950 Comstock Publishing Co., Inc. Used by permission of the publisher, Cornell University Press; D,
redrawn from diagram supplied by Dr. J.G. Riemann.)

occurs.15The term "simplex" is traditionally used for the ejaculatory duct of Lepidoptera, with
the corresponding sections being referred to as the anterior "noncuticular" and posterior
"cuticular" simplex regions; further, the noncuticular simpiex is subdivisible into as many as
seven or eight distinct segments (Figure ID),each with its own tinctorial and ultrastructural
characteristics (see, for example, References 12 and 16).
36 Insect Reproduction

Secretory cells of both ectodermal and mesodermal origin have been described from the
ejaculatory duct, and a variety of functions have been proposed for their products. In Lepi-
doptera and Coleoptera, components of the spermatophore are derived from the secretions.
The noncuticular simplex of some Lepidoptera also produces a sperm activator (see Section
IV.F).I7-l9In the midge Chironomusplumosus, which lacks accessory glands, seminal fluid is
produced by the ejaculatory duct cells.20An enzyme, esterase 6, produced in the anterior
ejaculatory duct of D. melanogastefll may have diverse roles in the species' reproductive
biologyz2(see Section 1V.F). Receptivity-inhibiting substances (see Section 1V.E) have also
been reported from the ejaculatory duct of Musca d o m e ~ t i c aand ~ ~Stomoxys ~ a l c i t r a n s . ~ ~
The forms and functions of the accessory glands (= collateral glands), which parallel the wide
diversity of external form and habits of the Insecta, have been considered in detail by Gillott.15 In
the great majority of insects, the glands are mesodermal and are named "mesadenia". This is clearly
the primitive arrangement, though, in a few groups, substitution of ectodermal for mesodermal
components in the reproductive system generally results in the formation of ectodermal glands
(ectadenia) (see Section III.A).Accessory glands are primitively absent in Thysanura, Ephemeroptera,
Plecoptera, Dermaptera, and most Odonata, and have been secondarily lost in many Diptera.
The accessory glands occur in most species as a single pair of tubular structures, though
in Coleoptera there are commonly two or three pairs, and in Thysanoptera and Acrididae
(Orthoptera) multiple pairs of tubules occur. In the latter arrangement, the cytology of a tubule
is generally uniform throughout its length, whereas in species with a single pair of tubules
there are often regional or intercellular differences. The secretion of the accessory glands,
indeed of individual tubules, is a complex mixture, and this is reflected in the variety of
functions that have been ascribed to it, including spermatophore, mating plug and seminal
fluid formation; fecundity enhancement and receptivity inhibition; sperm activation; and
supply of nutrients to the female. In view of the central importance of the accessory gland
secretion to the reproductive biology of the male, its nature and some of its major functions
are discussed as distinct topics in Section IV.

B. EXTERNAL GENITALIA
The male external genitalia include two components, the basic structures common to all
species and derived from the primary phallic lobes of the embryonic tenth (larval ninth)
abdominal segment25and secondary structures unique to groups or species formed on adjacent
segments. Because of the enormous diversity in form of the external genitalia among different
insect groups, only the basic plan will be considered here. For information on specific groups,
the reader is referred to the work of M a t ~ u d a . ~ ~
The phallic lobes arise as paired ectodermal outgrowths of the ventral surface of the
segment, though only in Ephemeroptera do they remain separate to form the paired penes seen
in the adult. In Thysanura the lobes meet in the midline to form a short tubular "penis," a
misnomer because the structure is not an intromittent organ. In Odonata, the genitalia on the
tenth segment are greatly reduced and, instead, secondary structures develop on the second
and third abdominal segments (see Section 1I.C). In most other insects, each phallic lobe is
divided to form a median mesomere and a lateral paramere. Between the fused mesomeres is
the gonopore from which the ejaculatory duct arises. Elongation of the mesomeres produces
an intromittent organ, the aedeagus, whose opening is the phallotreme and inner channel the
endophallus. In some insects, the endophallus is eversible through the phallotreme so that, in
effect, the gonopore becomes the external opening. Normally, the parameres differentiate into
elongate clasping organs. However, in some cases they fuse with the mesomeres to form the
phallobase. This can become an elongate structure, the phallotheca, that encloses the now
eversible aedeagus. This telescopic arrangement (i.e., phallotheca, aedeagus, and endophallus),
when everted by hemolymph pressure during copulation, may form a very elongate penis that
deposits sperm deep within the female system.
Insect Male Mating Systems 37

C. SPECIALIZED MATING SYSTEMS

According to M a t ~ u d ain, ~Ephemeroptera
~ the primary phallic lobes of the juvenile stage
are replaced by adult structures that develop within or in association with those of the nymph;
that is, phallic lobe development is biphasic in this order, in contrast to that in all other orders
in which continuous development towards the adult form occurs. Further, in this order each
imaginal phallic lobe remains separate and does not subdivide into mesomere and paramere.
Rather, each lobe develops as a penis, and from each gonopore an ejaculatory duct grows
anteriorly to fuse with the vas deferens on each side.
In Odonata, the phallic lobes do not develop to any significant degree so that males have
no aedeagus and only an inconspicuous ejaculatory duct. Thus, a unique method of sperm
transfer has evolved in this order.27Prior to mating, a male coils his abdomen forwards so that
the gonopore comes into contact with the intromittent organ on the third abdominal sternum,
and a spermatophore is transferred. During copulation, the male grasps the female's head
(Anisoptera) or prothorax (Zygoptera) by means of his terminal appendages (modified cerci).
The interlocking of the male and female parts may be extremely precise, thus playing an
important role in species isolation. The female then rotates her abdomen under that of the male
until her genital opening contacts the intromittentorgan when the spermatophoreis transferred.
Sperm transfer in Strepsiptera, many Cimicoidea, and at least one species of Anthocoridae
(Hemiptera) is achieved by hemocoelic Though the details vary, in this most
unusual form of insemination the penis penetrates the integument or vagina wall so that sperm
is deposited in the hemocoel, sometimes into a special structure, the spermalege. Eventually,
some of the sperm makes its way to the conceptacula seminales for storage, the rest being
phagocytosed either by hemocytes or by cells of the spermalege. HintonZ8suggested that
hemocoelic insemination may serve to provide females with nutrients to enable them to
survive for longer periods in the absence of suitable food.

111. DEVELOPMENT

A. EMBRYONIC DEVELOPMENT
The origin and development of the germ cells and gonads is varied, though two distinct
trends can be noted, namely, earlier segregation of the primordial germ cells and the restriction
of these cells to fewer abdominal segment^.^'.^^ In some Thysanura and Orthoptera, the germ
cells do not become distinguishable until they appear in the splanchnic walls of several
abdominal segments. In Locusta migratoria, for example, they arise in the second through
tenth pairs of abdominal somites although they persist only in pairs three to These
segmental groups of cells proliferate and fuse longitudinally to form the gonad on each side.
In contrast, in Dermaptera, Psocoptera, Homoptera, and many endopterygotes the germ cells
become obvious during blastoderm formation as roundish cells at the posterior end of the egg.
In due course, in exopterygotes the cells move anteriorly through the yolk to become enclosed
within the splanchnic mesoderm of the thirdfourth abdominal segments. The germ cells
subsequently separate into left and right halves, from which the testes develop. In endopterygotes,
the picture is similar except that division of the germ cells into two groups occurs prior to
migration through the y01k.~'.~*
In exopterygotes, the vasa deferentia develop from one or more posterior pairs of abdomi-
nal somites (e.g., those of segments nine and ten in p h a ~ m i d s while
~ ~ ) the ejaculatory duct is
an ectodermal invagination, usually arising behind the ninth sternum. In lower endopterygotes,
for example, Tenebrio molitor,6 embryonic development of the gonoducts is similar to that of
exopterygotes, and all the elements of the reproductive system can be identified at hatching.
In Diptera and Lepidoptera, the ductal components develop postembronically from a single
midventral or a bilateral pair, respectively, of genital imaginal discs that arise late in embryo-
g e n e ~ i s . ~In
~ -all
~ ' insects except, apparently, Acrididae (Orthoptera) the formation of the
38 Insect Reproduction

primary phallic lobes and subsequent development of the external genitalia occur during the
larval stage.26

B. POSTEMBRYONIC DEVELOPMENT
1. General
In exopterygotes, the reproductive system, like other organ systems, grows steadily during
the larval period; differentiation of the various components does not occur, however, until the
final juvenile stadium. In contrast, in endopterygotes very little growth occurs through the
larval period; rather, growth and differentiation occur simultaneously and are compressed into
the pupal stadium. "Differentiation," including in its broadest sense both spermatogenesis and
the organogenesis of the tubular components, is regulated hormonally as well as being affected
by various exogenous factors. The endocrine control of spermatogenesis is discussed by
Hardie in Chapter 5.

2. Endocrine Regulation
As with the differentiation of other tissues, it is changes in the hormone balance within the
insect that induce development of the reproductive tract. Though I am not aware of studies on
the phenomenon, it seems reasonable to propose, in light of what is known regarding hormonal
events during metamorphosis, that changes in hormone titers in earlier instars permit (and
possibly promote) the mitotic division of the undifferentiated reproductive tract cells and the
slight growth of the system seen especially in exopterygotes.
In the final juvenile instar, there occurs a marked decline in the level of circulating juvenile
hormone (JH), as well as one or more surges in the level of circulating e c d y ~ t e r o i d .This ~~.~~
change in the ratio of the two principal "players" permits the expression of adult characters,
though the specific roles of the two hormones are only slowly becoming clear. JH appears to
influence metamorphosis in two ways: first, by inhibiting the secretion of prothoracicotropic
hormone, ecdysteroid production and release is prevented; and second, though this is more
speculative, it may directly prevent the action of ecdysteroid at the organltissue l e ~ e l .
Our rather limited understanding of the involvement of hormones in preimaginal growth
and differentiation of the male reproductive system has recently been reviewed by H a ~ p . ~ '
Almost nothing is known about the site and mode of action of JH. In the locusts L. migratoria
and Schistocerca gregaria, implantation of two corpora allata into the abdomen of male fourth
(penultimate) instar nymphs leads to sterility.' However, this treatment or injection of syn-
thetic JH does not exert its effect on spermatogenesis, which proceeds normally, but prevents
differentiation of the vas deferens, which remains a thin, solid cord of cells.42These treatments
also prevent differentiation of the accessory whereas the reverse (i.e., allatectomy)
results in precocious metamorphosis of these structure^.^^ Perhaps more important,
Cantacuzi?ne'sl data show that the timing of the treatment is critical, the greatest effect being
achieved when insects are treated midway through the stadium. The accessory gland trans-
plantation experiments of G a l l o i ~ support ~ ~ . ~ ~this point. Implantation of these glands from
final instar nymphs of varied ages into adult hosts shows that only those glands from nymphs
2, 3, or more days into the instar are competent to differentiate. This critical period (days 2
and 3) coincides with a marked decrease in JH titer (and a small increase in the level of
ecdy~teroid)."~ As all these investigations utilized whole insects, it is unclear whether JH is
acting directly or indirectly on the tissues whose development it is influencing.
Mere absence of JH is insufficient for development to proceed; ecdysteroid (probably in
the form of ecdysterone) must be present both for growth and for differentiation of the
reproductive tract. The involvement of ecdysteroid has been investigated in several
endopterygotes, most notably Ephestia kiihniell~,~' Samia cynthia,1° Heliothis v i r e ~ c e n s ,
Bombyx m~ri,~O and T. m o l i t ~ r . ~In
' " ~Lepidoptera, both the sperm ducts and the genital discs
(which form the seminal vesicles [= upper vasa deferentia of some authors] and the remaining
Insect Male Mating Systems 39

components of the tract, respectively) grow and differentiateunder the influence of ecdysteroid.
Apparently, ecdysteroid exerts both direct and indirect effects. In S. cynthia1° and B. m~ri,~O
ecdysteroid is able to directly promote development of the tract in vitro, whereas in H.
virescens, ecdysterone alone has no e f f e ~ t . ~Development
~ , ~ ~ . ~ ~of the sperm ducts or genital
discs does occur when fat body or testis sheath is present in the culture medium in addition
to the hormone, or when the tissues are cultured with aqueous extract of fat body previously
exposed to ecdysteroid for 24 hours.
Only in T. molitor and H. virescens does the site and mode of action of the ecdysteroid
appear to have been s t ~ d i e d . ~In~ the
+ ~ bean-shaped
'-~~ glands (BAGs) of Tenebrio (Figure 1B)
there are two bouts of mitosis as growth occurs, the second of which coincides with the peak
of ecdysteroid in the pupal stage. In vitro studies showed that the second burst of mitosis
requires ecdysteroid for its oc~urrence?~ the hormone promoting the flow of cells from the G,
into the G, and S phases.52Though Grimnes and H a ~ showed p ~ ~ that the same concentration
of ecdysterone which produced maximum rates of mitosis also promotes formation of char-
acteristic secretions and adult-specific antigens in the BAGs, the latter activity in vivo is not
immediately induced by the ecdysterone but begins at the end of pupation when levels of the
hormone are In H. virescens genital discs, ecdysterone in combination with fat body or
testis sheath stimulates tritiated thymidine incorporation into DNA, suggestive of mitotic
divi~ion."~

IV. FUNCTIONAL ASPECTS

A. SPERMATOGENESIS AND SPERM STORAGE
1. Sperrnatogenesis
In insects, as in other animals, spermatogenesis comprises three phases: (1) the multipli-
cation phase (spermatocytogenesis); (2) the meiotic phase; and (3) the maturation phase
(spermiogene~is).~~ In many insects, this temporal sequence of events is "fixed" spatially; that
is, the phases occur in the distal, middle, and proximal regions, respectively, of each testis
follicle. In the multiplication phase, the spermatogonia (cells derived from germ cells)
undergo a species-specificnumber (usually, five to eight) of mitoses. For most insects (but not,
apparently, Diptera), these divisions (and hence the daughter spermatogonia) occur within a
"cyst," that is, a capsule of somatic cells formed initially around either a single spermatogo-
nium or a clone of daughter spermatogonia after one or two divisions. When mitosis is
complete, the spermatocytes, as the cells are now known, undergo two meiotic (maturation)
divisions. Generally, four spermatids result from each spermatocyte, each having a haploid
chromosome complement. However, especially among Diptera, variants of the meiotic pro-
cess occur, so that only one or two spermatids are formed from each spermatocyte. During
spermiogenesis, the spermatids differentiate into flagellated spermatozoa. At this time, the
enveloping cyst celis are rich in glycogen and may be supplying nutrients; in addition, they
phagocytose cytoplasmic remnants sloughed by the spermatids as they transform into sperma-
tozoa. In Acrididae (Orth~ptera)'.~~ and Cole~ptera,~'a mucopolysaccharide "cap," the
spermatodesm, comes to envelop the sperm heads near the end of spermiogenesis, though it
is by no means clear that this is a product of the cyst cells. According to Cantacuzbne,' the
cap is not a direct source of nutrients; rather, it contains enzymes that degrade nutritive
molecules secreted by the seminal vesicles into which the mature sperm are now transferred
and stored. In Tettigoniidae (Orthoptera), the spermatodesm is formed from secretion of the
intratesticular region of the vas deferens.'
In Lepidoptera, two types of sperm are produced, eupyrene (nucleate), which fertilize the
eggs, and apyrene (anucleate), whose largely speculative functions may include facilitating
eupyrene sperm movement from the testes to the vas deferens, assisting the eupyrene sperm
in their migration within the female reproductive tract, providing nourishment for the eupyrene
sperm, the female herself, or the zygote, and playing a role in sperm competition either by
40 Insect Reproduction

eliminating sperm deposited in previous matings or by preventing further matingsS8(and see

also Section 1V.F). The two types can be distinguished as early as the secondary spermatocyte
stage, following which their changing morphology follows very different paths.59 Eupyrene
sperm formation begins somewhat earlier than that of apyrene sperm, the former being
initiated in the last or penultimate larval instar, the latter in the last larval or pupal instar,
depending on the species. The eupyrene sperm, like the sperm of other types of insect, remain
in bundles until they have been transferred to the female tract; however, the apyrene sperm
bundles dissociate, releasing individual sperm, as they move out of the testis.
For most insects, formation of spermatogonia and spermatocytes is reported to occur in the
final nymphal or pupal instar, so that the testes of the adult contain only spermatids and
spermatozoa or only spermatozoa in those forms with a very short adult life. These observa-
tions led, more than 40 years ago, to the proposal that spermatogenesis was regulated by the
morphogenetic hormones, ecdysone and JH, the former promoting and the latter inhibiting the
process. However, in representatives from diverse orders, meiotic and even premeiotic phases
of spermatogenesis can be found in the adult t e ~ t i s .Dumser
~ ~ , ~ and
~ Da~ey@-~~%uggested that
the hormones affected only the rate of spermatogonial mitosis, not differentiation per se.
However, the subsequent discovery of ecdysteroids in adult males of several species and of
other effects of both e c d y s t e r o i d ~and
~ ~ ~JH" on germ cell maturation and viability has
reopened the question of the endocrine control of sperm production. See Chapter 5 for a full
discussion of this topic.

2. Sperm Release and Storage

In the majority of insects, the bundles of mature sperm are moved by peristalsis to the
seminal vesicle where they are stored until insemination. However, except in Lepidoptera,
information on this process is lacking. For several species of Lepidoptera, it has been observed
that release of sperm (both types) from the testis and its subsequent movement through the
upper vas deferens and seminal vesicle to the ductus ejaculatorius duplex (Figure 1D) shows
a daily rhythm (see Reference 64). This rhythm has since been shown to be circadian in
with the clock and its associated photoreceptor located in the testis-vas deferens
complex.66Correlated with the rhythm of sperm movement is cyclical secretory activity by the
epithelium of the upper vas deferens, though the purpose of this carbohydrate-rich secretion
remains unclear.@ However, in their passage toward the ductus ejaculatorius duplex, the
sperm undergo complex morphological changes, especially with respect to the fibrous sheath
that surrounds individual eupyrene sperm.67The significance of these changes remains un-
clear, but it may be speculated that they relate somehow to the ability of the sperm to fertilize.
The overall significance of this periodic sperm release and movement is related to the fact that
males normally mate but once a day, and at this time they ejaculate all the sperm stored in the
duplex. The timing of sperm release and movement (which is species specific) is such that the
sperm produced each day move into the duplex shortly after the end of the male's daily period
of receptivity to female sex pheromone. Thus, even if a male does mate, a substantial amount
of new sperm will be available for insemination the next day.65Though the usefulness of this
type of a system is readily apparent for species that have diurnal mating rhythms, it remains
to be determined how sperm release and storage is regulated in species whose males are more
opportunistic with respect to mating.

B. ACCESSORY GLAND ACTIVITY

As noted earlier, the accessory glands take on a vast array of forms, and their secretion a
spectrum of functions. Both these aspects have been extensively studied over the past two
decades, and the major conclusions are presented in reviews by H i n t ~ nLe0pold,6~
,~~ Chen,7O
Happ," and Gillott.15Also, the endocrine control of accessory gland activity has recently been
revie~ed.'~
Insect Male Mating Systems 41

1. Nature and Formation of Secretion

Numerous histochemical studies have shown the secretion to be a mixture or chemical
complex of protein, carbohydrate, and iipid, and electron microscopy has confirmed that the
glandular epithelium is designed for production of these materials. Also noteworthy is that
some of these early studies have demonstrated the regional variation that occurs both within
and between accessory gland tubules with respect to both amount and nature of secretory
components. Electrophoretic studies, in some cases combined with histochemistry, have
revealed that the secretion contains many distinct proteins (e.g., >40in the BAGSof T e n e b r i ~ ~ ~ ) ,
including glycoproteins and lipoproteins. Some of these proteins are enzymic in nature while
others, which may be quantitatively dominant, play a structural role (see below for details).
Free amino acids and small peptides have also been identified. Among the carbohydrates not
complexed with protein are glycogen, mucopolysaccharides (acidic, basic and neutral), glu-
cose, and inositol. Neutral lipids and phospholipids have been reported for some species.
Guanosine 3', 5'-cyclic monophosphate occurs in large amounts in the glands of Acheta
domesticus and several other Gryllinae (Orthoptera), where it may have a role in sperm
metabolism and a ~ t i v i t y .Uniquely,
~ ~ . ~ ~ the glands of male Hyalophora cecropia store massive
quantities of JH-I and JH-I1 which are transferred to the female during c ~ p u l a t i o n .Also,
~~-~~
the accessory glands of male Blaberoidea (Dictyoptera) accumulate uric acid (up to about 90%
of the dry weight in Blattella g e n n a n i ~ a ~which
~ ) , is then secreted as the outermost layer of
the spermatophore, possibly to dissuade the female from eating the spermatophore before
sperm evacuation has o c ~ u r r e d . However,
~ ~ . ~ the observation by Mullins and KeilSobthat
radiolabeled urate produced by males and ingested by females (as part of the spermatophore)
appears in the ootheca suggests that it may represent an important nitrogen resource for
reproductive success (and see Chapter 10).

2. Endocrine Regulation and Effect of Mating

Almost a11 investigations of the hormonal regulation of accessory gland activity have
focused on the role of JH. This is quite understandable in view of the demonstration,more than
50 years ago?' of the importance of the corpora allata in male reproductive activity and the
long-standing presumption that ecdysteroids would not occur in adults following the degen-
eration of the moult glands at metamorphosis. However, several demonstrations of ecdysteroids
in adult male^,^*-^^ combined with the realization that, in insects which are sexually mature at
eclosion, the accessory glands acquire their secretion during the pupal instar when the corpora
allata are inactive, raise the possibility that in some species ecdysteroids may be involved.
In males of most species studied, allatectomy prevents or greatly retards the accumulation of
secretion by the accessory glands, an effect that can be reversed by application of JH. Without
exception, the effect of JH is at the level of protein synthesis, and in the relatively few investigations
where it has been examined, JH has been shown to regulate the synthesis of specific proteins. For
example, in allatectomized Melanoplus sanguinipes the reduced ability of the long hyaline tubule
(Figure 1A) to accumulate secretion is due to the failure of the tubule to synthesize a glycoprotein,
LHPI, which comprises 51% of the total protein in the tubule of mature virgin males.85-86 Parallel
ultrastructural studies show that allatectomy causes changes in the protein-synthesizing machinery
of the gland cells and, in some cases, in the appearance of the s e c r e t i ~ n . ~ ~ - ~ ~
It must be emphasized that the majority of studies, in which whole insects have been used,
do not clarify whether the action of JH on the accessory glands is direct or indirect. A few
authors have used decapitation,%or have removed both the corpora allata and the cerebral
neurosecretory ~ystem,9'-~~ followed by JH application, or have used an in vitro system72.94-96
to confirm that JH acts directly on the glands. How and where JH acts in the accessory glands
is, at this time, a matter of conjecture, though Yamamoto et al?5 have obtained evidence
suggesting the existence of a membrane receptor protein for JH in D. melanogaster.
42 Insect Reproduction

Early reports that ecdysterone promoted protein-synthetic activity in accessory glands must
be treated cautiously because pharmacological, rather than physiological, doses of hormone
were e m p l ~ y e d .The
~ ~ .stimulation
~~ of RNA and protein synthesis by ecdysterone has been
observed under both in vivo and in vitro conditions in pharate and newly emerged adults of
Chilo partellus and Spodoptera l i t ~ r a . ~ ~These- ' O ~ are interesting observations, because in
other Lepidoptera ecdysteroid titers are known to be very low just prior to eclosion. Further,
as noted earlier (Section 1II.B.l), though ecdysteroid is required in T. molitor and B. mori to
render the accessory glands competent to produce secretion, when production begins at the
end of pupation, ecdysteroid levels are low; in other words, in these species protein synthesis
is not directly regulated by e c d y ~ t e r o i d s . ~ ~ . ~ ~
Only a handful of reports indicate that neurosecretory factors directly affect accessory
gland protein synthesis. For example, in Rhodnius prolixus, removal of the median neuro-
secretory cells reduces protein accumulation by the transparent accessory glands, an effect
which can be only partially overcome even by multiple doses of JH-I.93 In vitro studies
subsequently confirmed that a polypeptide from the brain stimulated protein synthesis in the
transparent accessory glands.Io2
An aspect of endocrine control that merits further examination i's the movement of proteins
between the accessory glands and the hemolymph. For example, in M. sanguinipes there are
immunologically similar proteins in the accessory glands, fat body, and hemolymph.lo3The
accessory glands can accumulate these proteins from the hemolymph in normal, but not in
allatectomized,males. In C. partellus, in vitro CO-cultureof accessory glands from adult males
with fat body and hemolymph from larvae injected several hours previously with [35S]-
methionine has demonstrated that the larval proteins are synthesized in the fat body, released
into the hemolymph, and are then accumulated by the accessory glands. Accumulation is
enhanced by ecdysterone or its agonist RH 5849 but inhibited by JH-I.Io4"Conversely, the
transparent accessory gland of Rhodnius synthesizes and releases into the hemolymph a 170-
kDa polypeptide,IWband Sevala and DaveylWbsuggest that the apparently redundant control
of protein accumulation in the transparent accessory gland by both JH%and neurose~retion'~~
may ultimately be explained by assigning a specific function, synthesis, or release, to each of
the hormonal factors.
That mating leads to enhanced synthesis of accessory gland secretion has been shown for
various However, to date there is no clear understanding of the control
pathway by which the mating effect is exerted. Baumann's study'0Ssuggested that in Droso-
phila funebris mechanical emptying of the gland was not the stimulus for renewed secretory
activity, and this author speculated that there might be "a neurohormonal influence." For D.
melanogaster, also, copulation-enhanced protein synthesis has been suggested to involve
neurohormonal factors based on the parallel effects of mating and JH a p p l i c a t i ~ n . In '~~
contrast, though JH is essential for the normal expression of protein synthesis in the accessory
glands of M. sanguinipes,lo7it does not mediate the copulation effect. This is readily seen in
mated but allatectomized M. sanguinipes which still exhibit a three-fold increase in accessory
gland protein synthetic activity compared with unmated allatectomized controls.lo7

C. SPERMATOPHORE FORMATION
Spermatophores are usually presumed to have evolved in association with the taking up of
terrestrial life by insect ancestors and have as their primary function the delivery of sperm to
the female reproductive tract. However, as Daveylo8pointed out, some marine Crustacea and
many aquatic Annelida produce a spermatophore, so that, for Insecta, this should not be
thought of as a new structure but rather as an existing struciure that has taken on a new
function. Though detailed descriptions of the mechanical aspects of spermatophore formation
are available for many species, aspects of the process such as the nature of the stimuli that
initiate it, its control and coordination, and its biochemical nature are relatively unexplored.
This is unfortunate, given the centrality of this method of sperm transfer in the evolutionary
Insect Male Mating Systems 43

success of insects and, from the pest management point of view, its importance in the life
history of species.
Generally, spermatophores have their most complex form in primitive groups; in advanced
endopterygotes they may be relatively simple or have been secondarily lost. Paralleling this
trend is a change in the site of formation of the spermatophore. GerberIogrecognized four
categories of spermatophore formation based on spermatophore complexity and site of
formation. In the most primitive, the first male-determined method, the complex spermato-
phore is formed in the ejaculatory duct or copulatory organ of the male. Such a method is
typical of orthopteroid insects. In the second male-determined method, which characterizes
some Hemiptera, Coleoptera, and a few Diptera, the spermatophore is again formed in the
copulatory sac, but the latter is everted into the bursa copulatrix of the female. After copula-
tion, the sac is withdrawn, leaving the usually less complex spermatophore in the bursa. In
Trichoptera, Lepidoptera, some Coleoptera, and a few Diptera, the spermatophore forms
directly in the female tract which thus determines its shape (first female-determined method);
the spermatophore is relatively simple but nonetheless, like those formed by the male-
determined methods, still encloses the sperm. In the second female-determined method, the
accessory gland secretions do not enclose the sperm; rather, they follow them into the female
genital tract where they may temporarily harden to form, for example, the mating plug of
mosquitoes and the honeybee and the sphragis of some Lepidoptera. It is speculated that these
"barriers" may prevent loss of semen or further transfer of sperm in a subsequent mating.
In most early analyses of spermatophore formation, only histological methods were em-
ployed, so that the origin and nature of spermatophore components remained unclear. A few
authors have used histochemistry or surgical removal of certain accessory gland structures to
determine the source and broad chemical nature of particular components (see Gillott15 for
references). However, it is only relatively recently that information on the biochemical nature
of individual spermatophore components and their mode of formation has become available.
For example, in T. molitor the use of monoclonal antibody techniques has permitted Happ's
group to trace three structural proteins (spermatophorins)from their site of production, distinct
cell types in the BAGS,to specific layers within the ~permatophore.~l@~~~ Amino acid analysis
of one of these spermatophorins showed that >25% of its residues were proline, which is in
keeping with the large amounts of this amino acid reported for other insect structural proteins
(in cuticle, egg shell, and ootheca) and for ~ollagen."~ Though having immunological
identicality, it is clear that the nature of the precursor accessory gland secretion may differ
from that of the spermatophorin. This was first noted by Grimnes and HappuOas a change in
solubility and electrophoretic mobility, leading these authors to speculate that proteolytic
cleavage of the precursor may have occurred. A proteolytic enzyme probably involved in the
formation of a spermatophorin has been identified in accessory gland secretions of M.
~ a n g u i n i p e sA
. ~precursor
~~ protein of m01 wt 85 kDa, which appears to be produced primarily
in short hyaline tubule 3 (Figure lA), is cleaved by a trypsin-like enzyme to form the
spermatophorin SP62 (m01 wt 62 kDa), a major component of the outer layer of the spermato-
phore. The major products of the white gland tubules of Melanoplus (Figure 1A) are also
spermatophorins, but it is not known whether they, too, are derived by proteolysis of higher
molecular weight precursors. The function of an aminopeptidasealso identified in Melanoplus
accessory gland secretion remains unknown.'13
Except in GryIlidae (Orthoptera), where a spermatophore is preformed by the male and
carried until copulation takes place, spermatophore formation begins only after copulation is
initiated, that is, the male has achieved the correct mating position. Sensory input, especially
tactile but for some species chemical and visual, triggers the process, which is under motor
neuronal control. For most insects studied, the central nervous system must be intact in order
for spermatophore formation to start. For example, in Locusta the head ganglia are necessary
at this stage to interpret input from the cerci, though decapitation 15 min or more after
initiation does not disrupt spermatophore formation.'14 Control of the subsequent stages of
44 Insect Reproduction

spermatophore formation resides in the terminal abdominal ganglion, removal of which causes
abrupt termination of the process. The presence of the female is necessary only at the start,
presumably related to the need for correct sensory input via the cerci. Separation of copulating
pairs beyond 15 min does not disrupt spermatophore formation though the structures are
malformed because, Gregoryii4suggested, the spermatophore desiccates and the spermathecal
duct serves as a mold for the spermatophore tube in this species.
In Teleogryllus commodus and perhaps other species of crickets where the spermatophore
is produced prior to mating, formation of the spermatophore is under circadian control; that
is, a new spermatophore is produced during each 24-h period.ii5According to Loher,li5who
employed a variety of surgical procedures, the pars intercerebralis region of the brain may
serve a dual function in regulating spermatophore formation. First, it may hormonally regulate
the synthesis of the raw materials in the accessory glands and, second, it may be the
coordinating center for release of the materials from the glands as the spermatophore is being
formed. However, Loher's workii5does not clarify whether the pars intercerebralis serves
only as a trigger, the rest of the process being regulated within the terminal abdominal
ganglion as in other insects, or whether it controls the entire event. The site of the circadian
control center is also undetermined but may be within the optic lobes, as their destruction leads
to random generation of spermatophores throughout the 24-h period.

D. SEMINAL FLUID
At the outset, it is important to clarify the term "insect seminal fluid" because, in contrast
to the mammalian situation in which seminal fluid is a secretion of the seminal vesicles, in
many insects the seminal vesicles are not glandular in nature. Potentially, the fluid that comes
to bathe the sperm may be derived from any or all glandular parts of the male system, and its
nature may change as secretions are added to or removed from it during insemination and
when it reaches the female system.
The following discussion will deal only with our somewhat limited knowledge of the
general composition and functions of insect seminal fluid. The nature and functions of specific
components will be examined in parts E and F of this Section. The obviously minute quantities
of semen together with the high density of sperm make the obtaining of seminal fluid samples
for analysis difficult. Further, depending on its source (e.g., ejaculate, spermatophore, or
spermatheca), its composition will vary. Perhaps not surprisingly, a variety of potential energy
sources for the sperm have been identified. For example, Apis mellifera seminal fluid contains
trehalose, glucose, and fructose though the latter becomes virtually undetectable 40 min after
ejaculation, perhaps because of its use by the sperm.ii6The three sugars occur throughout the
reproductive tract with the greatest concentration in the testes and penis bulb. Histochemical
studies have demonstrated the existence of glycogen in the seminal fluid of Periplaneta
americanaii7and S. g r e g ~ r i aThe
. ~ ~lipid
~ in the semen of A. mellifera is almost entirely of
spermatozoan origin rather than in the seminal fluid.Ii9The amino acids (free and bound) in
A. mellifera semen apparently resemble those of mammalian semen though their specific
origin (sperm or seminal fluid) and functions are undetermined.I2OWhile Blum et a1.Ii6report
that the semen of A. mellifera contains calcium, sodium, manganese, magnesium, copper, and
iron (of which only the last three are detectable in sperm), there appear to have been no
quantitative studies on the inorganic constituents of seminal fluid despite several reports on
the activation of sperm by buffers at various pH values.

E. FECUNDITY-ENHANCING AND RECEPTIVITY-INHIBITING CHEMICALS

In addition to its obvious function of sperm transfer, for many species mating has other
important effects, including the stimulation of egg production and the rendering of the female
unwilling to remate. In some species (for example, cockroachesi21)the stimulus given by
mating is physical in nature; that is, the insertion of the spermatophore stretches the wall of
the bursa copulatrix, which causes the female to become unreceptive. For others, chemicals
Insect Male Mating Systems 45

in the seminal fluid stimulate egg production andlor render females unreceptive. These
pheromones have been named fecundity-enhancing (FES) and receptivity-inhibiting sub-
stances (RIS), re~pectively.'~~
Both FES and RIS "inform" the female that she has been inseminated. For the FES, the
significance lies in the fact that in most insects unfertilized eggs are inviable; thus, it is critical
that oviposition not occur until a supply of sperm is available. For the RIS, the rendering of
the female unreceptive "guarantees" that only the first (fittest) male's sperm will be used to
fertilize the eggs. It may also provoke males to actively seek virgin females in the population
and, for females of some species, to switch their behavior from mate-seeking to food- and/or
oviposition-site seeking. Although the FES and RIS play distinct roles, it is appropriate to deal
with both concurrently in view of their similar sites of production and chemical nature. Indeed,
in some species it seems likely that the same substance may serve as both a FES and a RIS.
Gillott and FriedelIz2and GillottI5have provided detailed reviews of FES and RIS.

1. Source and Nature

In males of most species where FES or RIS have been identified, the accessory glands are
their source. However, in Musca, males of which lack such glands (Figure lC), the FESIRIS
is produced in the upper third of the ejaculatory duct.Iz3The testes apparently produce these
pheromones in H. c e c r ~ p i a ,~' ~r i~c k e t s , ' ~and R.~proli~us,'~'
~.'~ (though in the latter species,
an effect of the seminal fluid merely stretching the spermathecal wall cannot be ruled out). It
should also be noted that for none of these species is it clear whether glandular cells in the
gonads or the sperm are the source of the pheromone. In some Drosophila species, there is
evidence for the existence of both short- and long-term MS, the former being produced by the
accessory glands, the latter by the
With one exception, all FES and RIS characterized to date are peptides or proteins. For
example, the monocoitic substances reported from M. domestica, Cochliomyia hominivorux
and Phormia regina are peptides with molecular weights estimated at 750 to 3000 Da.I3O
Likewise, the RIS (PS-1) of D. funebris has a molecular weight of approximately 2700 Da and
comprises 27 amino acid r e s i d u e ~ .PS-1' ~ ~ exists in two forms, differing in a single amino acid
(valine or leucine) at position 2.l3l The sex peptides of D. melanogaster and D. sechellia,
which serve as both FES and RIS, each comprise 36 amino acids and differ from each other
at only three positions. Perhaps not surprisingly, the D. sechellia sex peptide stimulates
oviposition and inhibits receptivity in D. melanogaster, as well as in D. simulans and D.
mauritania, all four species belonging to the melanogaster species subgroup. However, the D.
melanogaster peptide has no effect on virgins of D. funebris and vice versa.132
On the other hand, the FES of M. ~anguinipes,'~~ L. r n i g r a t ~ r i a ,and
' ~ ~Aedes a e g ~ p t iare
l~~
proteins of molecular weight 30, 13, and 60 kDa, respectively. The FES of the crickets A.
d o r n e s t i ~ u sand
~~~T. . cornrnod~s'~~
~~~ is a prostaglandin synthetase enzyme complex which,
after transfer during mating, promotes prostaglandin production in the female reproductive
system.
Three D. melanogaster accessory gland proteins and the genes which code for them have
been studied by Wolfner's Of these, perhaps the most interesting is msP 355a, a
basic protein with many features typical of peptide pheromone and hormone precursors;
indeed, part of its amino acid sequence is similar to that of the egg-laying hormone (ELH) of
the mollusc Aplysia ca1iforni~a.l~~ In the accessory glands, msP 355a exists mainly in a form
of molecular weight 37 kDa. During mating, it is transferred to the female genital tract; some
of the 37-kDa fraction passes unchanged into the female's hemolymph, but the rest undergoes
rapid proteolysis, first to a 29-kDa, then to a 22-kDa fragment. However, the ELH-like
segment is retained by both these smaller m0lecu1es.l~~ The second protein in the trio is msP
355b, an acidic protein 90 amino acids in length and having a molecular weight of 11-14
kDa.l4I It, too, is transferred during mating, and some enters the female hemolymph. However,
that which remains in the genital tract is not proteolysed and, along with the msP 355a
46 Insect Reproduction

fragments, unstored sperm, and other secretions, is expelled from the genital tract some 2-3
h after copulation terminates. DiBenedetto et al.,I4Oon the basis of their analysis of the gene
that codes for it, have characterized msP 316, a small basic protein made up of 52 amino acids.
Prostaglandins have been identified from the male reproductive organs of B. r n ~ r iand ,~~~
greater levels of these compounds have been noted in mated females compared with virgins.143
However, it is not clear whether these increases are the result of direct transfer from the male
or are an effect of mating.

2. Site and Mode of Action

The most common effect of FES is to stimulate oviposition, though the manner in which
this is achieved is not well understood. Extracts of Locusta male accessory glands stimulate
contractions of the lateral oviduct, an effect that can be partially mimicked by octopamine and
forskolin, suggesting the involvement of octopaminergic re~ept0rs.l~~ And recently, specific
peptides with myotropic activity on the oviduct have been isolated from male accessory
glands, spermatophores and spermathecae of mated fern ale^.'^^.^^^ Obviously, oviposition is a
complex process, both mechanically speaking and in terms of when it occurs. Not surprisingly,
it is controlled both hormonally and neurally (for review, see Lange14'). Perhaps the role of
these male-derived substances is to prime the oviductal musculature so that, at the appropriate
time and place for egg laying, the nervous system can provide the fine control.
In contrast, in RhodniusI2' and T e l e ~ g r y l l u the
s ~ ~FES
~ acts on the spermatheca, which is
stimulated to produce a hormone of unknown nature (the "spermathecal factor") and prosta-
glandins, respectively. Similarly, in Hyalophora the FES is released into the bursa copulatrix
where it triggers production of a hormone.124For other species, it seems likely that the FES
passes through the wall of the female reproductive system to some other site of action. Thus,
in D. funebris, FES can be detected in all parts of the body 2 h after mating,148and in both M.
~anguinipesl~~ and M. d o m e s t i ~ asome
' ~ ~ male accessory gland proteins pass unchanged into
the hemolymph of the female (though in neither species is it clear whether the FES is one of
these).
The remaining links in the pathway culminating in egg laying are not well known. For a
number of species (references in References 15,147), the cerebral neurosecretory cells are
known to produce an ovulation- or oviposition-inducinghormone (a myotropin) whose release
may be facilitated by the FES. In Rhodnius, for example, ecdysteroids released from the ovary
as eggs mature can initiate electrical activity in, and release of myotropin from, specific
median neurosecretory cells. However, this occurs only in mated females, that is, in the
presence of the spermathecal factor which, it has been p r ~ p o s e d , ~may ~ ~ unmask
"~ aminergic
receptors within the brain allowing the ecdysteroids to act. In contrast, in B. mori the FES may
act on the terminal abdominal ganglion, whose spontaneous activity is increased after mating
or application of extracts of the male reproductive system.152a Further work showed that the
sensitivity of the ganglion to Ringer's solutions containing varied amounts of NaCl and KC1
is changed after mating, prompting the suggestion that the FES might act by altering the
permeability of the neural sheath surrounding the gang1i0n.l~~~
A second potential role for an FES is to enhance egg development. This has been proposed
by several authors, though only in a few species has a direct link been demonstrated.
B a ~ m a n n , using
l ~ ~ an in vitro system, showed that uptake of I4C-labeled amino acids into
proteins was increased in the ovaries of virgin D. funebris injected with FES. Decapitation
immediately after mating prevents this increase, leading Baumanr~l~~ to suggest that the FES
acts via the neuroendocrine system.
In mosquitoes, also, the FES promotes vitellogenesis, possibly by inducing the ovaries to
produce a hormone whose function is to stimulate release of egg development neurosecretory
hormone from the corpora ~ a r d i a c a .For ' ~ ~anautogenous mosquitoes, an additional means of
promoting egg development has been suggested,154namely, that the FES causes acceleration
Insect Male Mating Systems 47

of blood-meal digestion. Again, the evidence suggests that this effect is exerted via an
endocrine pathway. Even in nutritionally stressed female mosquitoes, accessory gland im-
plants trigger egg development, leading Klowden and Chambers155to suggest that the FES is
acting as a primer pheromone that switches the female's metabolic priorities from self-
sustenance to egg production.
In the bean weevil Acanthoscelides obtectus, the sole demonstrated role for the FES is
stimulation of egg development; that is, the pheromone does not promote egg laying.lS6
The demonstration that mated females carrying an hsP70-msP 355a fusion gene lay 20%
more eggs than controls has led Monsma et al.I4l to speculate that msP 355a may have some
role in egg production. The function of msP 355b is unknown. Like msP 355a, msP 316 has
features common to precursors of peptide hormones though its role remains unknown.I4O
Much less information is available on the site and mode of action of RIS, though, in all
instances, the pheromone exerts its influence via the neuroendocrine system. In H. cecropia,
the RIS, like the FES (with which it may be chemically identical), may stimulate the wall of
the bursa copulatrix to release a hormone.124Originally, it was proposed that this hormone
acted on the corpora cardiaca to stop release of the calling hormone; however, more recent
has ruled out any involvement of the corpora cardiaca in calling behavior, and the
remaining steps in the pathway remain unknown. The brain appears to be the site of action of
the RIS in Musca, as decapitated, decerebrated, or cervically ligated virgins will mate as many
as five times in an 8-h period.158Further, radiolabeled male material transferred during mating
accumulates in the head r e g i ~ n , ' ~ Oleading
J ~ ~ Leopold et a1.Is9to propose that the RIS binds
to receptor sites in the head (brain?) to induce refractoriness. Interestingly, in view of the
apparently identical nature of the RIS and the FES in Musca, refractoriness (which in this
insect is marked by withdrawal of the ovipositor) is induced by high concentrations, whereas
oviposition (requiring extension of the ovipositor) is triggered by low concentrations of the
pheromone.
In contrast, Gwadz's work160 suggests that in Aedes aegypti the RIS acts on the terminal
abdominal ganglion, with the brain, the suboesophageal ganglion, and the thoracic ganglionic
mass having no direct involvement in the control of sexual behavior.
An aspect worthy of study is the interaction between RIS and JH, the latter enhancing
receptivity as females become sexually mature. It is unlikely that RIS directly affects corpus
allatum activity as refractoriness is life-long, though a female goes through several cycles of
egg production, the latter correlated in many species with changes in corpus allatum activity.
As proposed earlier,15it may be that the RIS and JH compete for the same receptor sites within
the central nervous system, with the RIS being successful in normally monogamous species.
In this context, it is noteworthy that when less than normal amounts of RIS are transferred
during mating (e.g., by forcibly separating mating pairs or mating females to already multiply-
mated malesl6I), receptivity returns after a variable period of time.

F. OTHER FUNCTIONS
Several other functions have been ascribed to components of the accessory gland secretion
or to secretory products of other parts of the male tract. Most of these relate to maturation,
activation (motility), or energy metabolism of the sperm; in addition, for some species, the
spermatophore represents a source of nourishment for the female.
How sperm move from the spermatophore to the spermatheca is largely unknown though
it presumably involves either active movement on the part of the gametes or peristalsis of the
wall of the female tract so as to "squeeze" sperm out of the spermatophore andlor to draw
sperm up the spermathecal duct. In Rhodnius, there is good evidence for production by the
opaque accessory glands of a peristalsis-inducing s e ~ r e t i 0 n . Normally,
l~~ the spermathecae
begin to fill with sperm within 5 to 10 min of the end of mating. However, in females mated
to males whose opaque glands have been removed, the spermathecae are still empty even 5
h later. This is not due to a malformed spermatophore (which is a product of the transparent
48 Insect Reproduction

accessory glands) or to an action on the sperm (which even if killed can still be moved into
the spermathecae). In isolated preparations of the reproductive tract, the opaque gland secre-
tion induces peristalsis, ruling out central nervous control; however, D a ~ e yalso l ~ ~showed that
the secretion acts on the peripheral nervous system rather than directly on the musculature of
the reproductive tract. A similar function for the accessory gland secretion has been suggested
for several other species; in all cases, however, an effect on the sperm per se rather than the
tract musculature cannot be ruled out.
In Saturniidae (Lepidoptera), the noncuticular simplex (see Figure ID) produces a sperm
activator (it is not specified whether apyrene, eupyrene, or both types of sperm are affected).I8
The molecule, which is reported to be a peptide of molecular weight about 3100 Da,163
apparently diffuses through the spermatophore wall (already formed from secretions emanat-
ing further up the simplex) and may work by disrupting the sperm plasma membrane.Ibl A
detailed study of sperm activation of Bombyx mori has been undertaken by Osanai and
colleagues, who have demonstrated a remarkable proteolytic cascade within the spermato-
phore that affects both apyrene and eupyrene perm.'^^-^^' During spermatophore and seminal
fluid formation, various components are added from different regions of the ejaculatory duct,
the final (but key) participant being i n i t i a t ~ r i nan
, ~ endopeptidase
~~ released by the proximal
segment of the duct.166Four distinct roles have been identified for this enzyme. It digests the
coating around the apyrene sperm whose resultant activity then serves to stir the seminal
it also digests the intercellular glue that binds together the eupyrene sperm, leading
to their release and a~tivati0n.l~~ Its third role is to split proteins (produced elsewhere in the
simplex) on the C side of arginine r e ~ i d u e s and , ~ ~its
~ fourth role is to activate an arginine
carboxypeptidase produced in the ampulla.168The arginine then released by the action of this
e~opeptidasel~~ is hydrolyzed to ornithine and urea under the catalytic action of arginase from
the seminal vesicles.170From the ornithine is derived glutamate, then 2-oxoglutarate, which
serves both as a substrate for sperm respiration and as a promoter of pyruvate 0xidati0n.l~~
Trehalases have been detected in the accessory glands of male P. a m e r i ~ a n a and l ~ ~in the
BAGS and spermatophore of T.m01itor.I~~ Characterization of the enzyme174shows that it has
high specificity towards trehalose. However, the few published analyses do not suggest that
this sugar normally occurs in very significant amounts in insect seminal fluid. Thus, a role for
this enzyme, for example, in sperm metabolism, remains unclear.
The anterior ejaculatory duct of D. melanogaster produces an enzyme, esterase 6, that is
transferred to the female in the seminal fluid.21Esterase 6 appears to have two purposes: it may
be involved in lipid catabolism in the ejaculate, thus affecting sperm motility;175it was also
proposed22that the enzyme catalyzed the conversion of cis-vaccenyl acetate (produced in the
male's ejaculatory bulb and also transferred during mating) to cis-vaccenol, the latter then
being released by the female as an antiaphrodisiac. However, attempts to confirm this have
been u n s u c ~ e s s f u l . ~ ~ ~
In addition to the RIS described earlier, a second sex peptide has recently been identified
and characterized in the accessory glands of D. f ~ n e b r i s . lThis ~ ~ 63-amino acid-containing
molecule has several similar properties and sequence homologies with known protease
inhibitors, including the ability to inhibit acrosin, a trypsin-like endopeptidase associated with
the acrosome of mammalian sperm. Thus, Schmidt et speculate that this peptide tempo-
rarily inactivates the acrosomal proteases until the appropriate moment for egg fertilization.
Males of some species produce a very large spermatophore (e.g., in some Gryllidae and
Tettigoniidae [Orthoptera] it may be 40% of the male's body weight178),part or all of which
is eventually eaten by the female; in others, for example M. ~anguinipes,'~~ the male transfers
several spermatophores in sequence during a single mating. Though there have been claims
that in some species the spermatophore is digested within the female reproductive tract,
demonstrations of the existence of hydrolytic enzymes in this region are virtually nonexistent.
It has been shown, however, that material from the spermatophore does enter the female's
hemolymph and, in some species, the ovaries, leading to the proposal that the male may
Insect Male Mating Systems 49

thereby make a nutritional contribution to the female or, more specifically, to egg develop-
The subject of male nutrient investment is discussed by Boggs (Chapter 10).
ment.149-180-183

ACKNOWLEDGMENTS
Original work of the author cited in this review is supported by the Natural Sciences and
Engineering Research Council of Canada. Thanks are extended to Dr. J.G. Riemann for
provision of a diagram of the male flour moth reproductive system and to Mr. D. Dyck for
assistance in the preparation of the figure.

REFERENCES
I. Cantacuzkne, A.-M., Recherches morphologiques et physiologiques sur les glandes annexes dies des
O~thopttres.111. Modes d'association des spermatozoides d'olthoptbres, Z Zellforsch., 90, 113, 1968.
2. Rojas-Rousse, D., Description et fonctionnement de I'appareil genital interne de Diadromus pulcheNus
Wesmael (Hymenoptera:Ichneumonidae).I. Testicules, canaux deferents, glandes annexes, canaux collecteurs,
canal ejaculateur du mile, In!. J. Insect Morphol. Embryol., 1, 225, 1972.
3. Gerber, G.H., Church, N.S., and Rempel, J.G., The anatomy, histology, and physiology of the reproductive
systems of Lytta nuttalli Say (Coleoptera: Meloidae). I. The internal genitalia, Can. J. Zool., 49, 523, 1971.
4. Ramamurty, P.S., Histological studies of the internal organs of reproduction in Nezara viridula Fabr.
(Pentatomidae-Heteroptera, Insecta), Zool. Ant., 183, 119, 1968.
5. Gerber, G.H., Neill, G.B., and Westdal, P.H., The anatomy and histology of the internal reproductive organs
of the sunflower beetle, Zygogramma exclamationis (Coleoptera: Chrysomelidae), Can. J. Zool., 56, 2542,
1978.
6. Poels, A., Histophysiologie des voies gtnitales miles de Tenebrio molitor L. (Coleopttre: Tenebrionidae),
Ann. Soc. R. Zool. Belg., 102, 199, 1972.
7. Gerber, G.H., Reproductive behaviour and physiology of Tenebrio molitor (Coleoptera:Tenebrionidae).111.
Histogenetic changes in the internal genitalia, mesenteron, and cuticle during sexual maturation, Can. J. Zool.
54, 990, 1976.
8. Riemann, J.G. and Thorson, BJ., Ultrastructure of the vasa deferentia of the Mediterranean flour moth, J.
Morphol., 149, 483, 1976.
9. Cantacuzkne, A.-M., Origine et cancttres ultrastructuraux des cellules spermiophagesdu criquet mignteur
Locusta migratoria, J. Microsc. (Paris), 10, 179, 1971.
10. Szollosi, A. and Landureau, J.-C., Imaginal cell differentiation in the spermiduct of Samin cynthia (Lepi-
doptera). Responses in vitro to ecdysone and ecdysterone, Biol. Cell., 28, 23, 1977.
11. Bairati, A., Structure and ultrastructure of the male reproductive system in Drosophila melanogaster Meig.
11. The genital duct and accessory glands, Monit. Zool. Ital., 2, 105, 1968.
12. Lai-Fook, J., The vasa deferentia of the male reproductive system of Calpodes erhlius (Hesperiidae, Lepi-
doptera), Can. J. Zool., 60, 1172, 1982.
13. Khalifa, A., Spermatophore production in Trichoptera and some other insects, Trans. R. Entomol. Soc.
London, 100,449, 1949.
14. Gerber, G.H., Church, N.S., and Rempel, J.G., The structure, formation, histochemistry,fate and functions
of the spermatophore of Lyua nuttalli Say (Coleoptera: Meloidae), Can. J. Zool., 49, 1595, 1971.
15. Gillott, C., Arthropods-lnsecta, in Reproductive Biology of Invertebrates, Vol. 3 (Accessory Sex Glands),
Adiyodi, K.G. and Adiyodi, R.G., Eds., Oxford and IBH, New Delhi, 1988, chap. 12.
16. Riemann, J.G. and Thorson, BJ., Foliate and granule-secreting cells in the ejaculatory duct (simplex) of
the Mediterranean flour moth, J. Ultrastruct. Res., 66, 1, 1979.
17. Omura, S., Studies on the reproductive system of the male of Bombyx mori. 11. Post-testicular organs and
post-testicular behaviour of the spermatozoa, J. Fac. Agric. Hokkaido Imp. Univ., 40, 129, 1938.
18. Shepherd, J.G., Activation of satumiid moth sperm by a secretion of the male reproductive tract, J. Insect
Physiol., 20, 2107, 1974.
19. Herman, W.S. and Peng, P., Juvenile hormone stimulation of sperm activator production in male monarch
butterflies, J. Insect Physiol., 22, 579, 1976.
20. Wensler, RJ.D. and Rempel, J.G., The morphology of the male and female reproductive systems of the
midge, Chironomus plumosus L., Can. J. Zool., 40, 199, 1962.
2 1. Sheehan, K.B., Richmond, R.C., and Cochrane, BJ., Studies of esterase 6 in Drosophila melanogaster. 111.
The developmental pattem and tissue distribution, Insect Biochem., 9, 443, 1979.
22. Mane, S.D., Tomkins, L., and Richmond, R.C., Male esterase 6 catalyses the synthesis of a sex pheromone
in Drosophila melanogaster females, Science, 222, 419, 1983.
50 Insect Reproduction

23. Riemann, J.G., Moen, D.J., and Thorson, B.J., Female monogamy and its control in houseflies, J. Insect
Physiol., 13, 407, 1967.
24. Morrison, P.E., Venkatesh, K., and Thompson, B., The role of male accessory-gland substance on female
reproduction with some observations of spermatogenesis in the stable fly, J. Insect Physiol., 28, 607, 1982.
25. Snodgrass, R.E., A revised interpretation of the external reproductive organs of male insects, Smithson. Misc.
Collect., 132, 1, 1957.
26. Matsuda, R., Morphology and Evolution of the Insect Abdomen, Pergamon Press, New York, 1976, 534 pp.
27. Waage, J.K., Sperm competition and the evolution of odonate mating systems, in Sperm Competition and the
Evolution of Animal Mating Systems, Smith, R.L., Ed., Academic Press, New York, 1984, 251.
28. Hinton, H.E., Sperm transfer in insects and the evolution of haemocoelic insemination, Symp. R. Entomol.
Soc. London, 2, 95, 1964.
29. Carayon, J., Traumatic insemination and the paragenital system, Entomological Society of America, Lanham,
MD, Thomas Say Foundation Publ., 7, 81, 1966.
30. Carayon, J., Parthknog6nbse constante prouvb chez deux hkteropteres: le miride Campyloneura virgula et
I'anthocoride Calliodis maculipennis, Ann. Soc. Entomol. Fr., 25, 387, 1989.
3 1. Anderson, DJ., The development of hemimetabolous insects, in Developmental Systems: Insects, Vol. 1,
Counce, S.J. and Waddington, C.H., Eds., Academic Press, New York, 1972, chap. 3.
32. Anderson, D.T., The development of holometabolous insects, in Developmental Systems: Insects, Vol. 1,
Counce, S.J. and Waddington, C.H., Eds., Academic Press, New York, 1972, chap. 4.
33. Roonwal, M.L., Studies on the embryology of the African migratory locust, Locusta migratoria migratorioides
Reiche and Frm. (Orthoptera, Acrididae). 11. Organogeny, Philos. Trans. R. Soc. London. Ser. B, 227, 175,
1937.
34. Cavallin, M., hude descriptive du dkveloppement et de la diffkrenciation sexuelle de I'appareil genital chez les
embryons des phasmes Clitumnus extradentatus Br. et Carausius morosus Br., C. R. Acad. Sci., 268,2 189,1969.
35. Gehring, WJ. and Nothiger, R., The imaginal discs of Drosophila, in Developmental Systems: Insects, Vol.
2, Counce, S.J. and Waddington C.H., Eds., Academic Press, New York, 1973, 212.
36. Leclerq-Smekens, M., Organogenese et diffkrentiation des voies genitales miles internes des Itpidopt&ressur
la base d'obsewations effectukes chez Euproctis chrysorrhea (Lymantriidae),Ann. Soc. R. Zool. Belg., 104,
131, 1974.
37. Loeb, M.J., Development of isolated spermducts from Heliothis virescens (Lepidoptera)in vitro, Invertebr.
Reprod. Dev., 20, 67, 1991.
38. Steel, C.G.H. and Davey, K.G., Integration in the insect endocrine system, in Comprehensive Insect
Physiology. Biochemistry and Pharmacology, Vol. 8, Kerkut, G.A. and Gilbert, L.I., Eds., Pergamon Press,
Oxford, 1985, 1.
39. Wigglesworth, V.B., Historical perspectives, in Comprehensive lnsect Physiology. Biochemistry and Phar-
macology, Vol. 7, Kerkut, G.A. and Gilbert L.I., Eds., Pergamon Press, Oxford, 1985, 1.
40. Sehnal, F., Growth and life cycles, in Comprehensive Insect Physiology. Biochemistry and Pharmacology.
Vol. 2, Kerkut, G.A. and Gilbert L.I., Eds., Pergamon Press, Oxford, 1985, 1.
41. Happ, G.M., Maturation of the male reproductive system and its endocrine regulation, Annu. Rev. Entomol.,
37, 303, 1992.
42. Szollki, A., Imaginal differentiation of the spermiduct in acridids: effects of juvenile hormone, Acrida, 4,205,
1975.
43. Cantacuzkne, A.-M. and Girardie, A., Caractkres sexuels des adultes prothkttliques macs obtenus par
allatectomie de Locusta migratoria migratorioides (Orthopttre), C. R. Acad. Sci., 270, 2465, 1970.
44. Gallois, D., La diffkrenciation fonctionnelle des glandes annexes mlles de Locusta migratoria migratorioides
R. et F., Arch. Zool. Exp. G b , 122, 109, 1981.
45. Gallois, D., Control of cell differentiation in the male accessory reproductive glands of Locusta migratoria:
acquisition and reversal of competence to imaginal secretion, J. Insect PhysioL, 35, 189, 1989.
46. Baehr, J.-C., Porcheron, P., Papillon, M., and Dray, F., Haemolymph levels of juvenile hormone,
ecdysteroids and proteins during the last two larval instars of Locusta migratoria, J. Insect PhysioL, 25,415,
1979.
47. Nowock, J., Growth and metamorphosis in the testes of Ephestia kuhniella in vitro, J. Insect Physiol., 19,941,
1973.
48. Loeb, MJ., Growth and development of spermducts of the tobacco budworm moth Heliothis virescens, in
vivo and in vitro, Invertebr. Reprod. Dev., 19, 97, 1991.
49. Loeb, MJ. and Hakim, R.S., Development of genital imaginal discs of Heliothis virescens culture in vitro
with 20-hydroxyecdysone and fat body or testis sheaths, Invertebr. Reprod. Dev., 20, 181, 1991.
50. Shinbo, H. and Happ, G.M., Effects of ecdysteroidson the growth of the post-testicular reproductive organs
in the silkworm, Bombyx mori, J. Insect Physiol., 35, 855, 1989.
51. Szopa, T.M., Lenoir Rousseaux, J.-J., Yuncker, C., and Happ, G.M., Ecdysteroids accelerate mitoses in
accessory glands of beetle pupae, Dev. Biol., 107, 325, 1985.
Insect Male Mating Systems

52. Yaginuma, T., Kai, H., and Happ, G.M., 20-Hydroxyecdysone accelerates the flow of cells into the G, phase
and the S phase in a male accessory gland of the mealworm pupa (Tenebrio molitor), Dev. Biol., 126, 173,
1988.
53. Grimnes, K.A. and Happ, G.M., Ecdysteroids in vitro promote differentiation in the accessory glands of
male mealworm beetles, Experientia, 43, 906, 1987.
54. Yaginuma, H. and Happ, G.M., 20-Hydroxyecdysoneacts in the male pupa to commit accessory glands
toward trehalase production in the adult mealworm beetle (Tenebrio molitor), Gen. Comp. Endocrinol., 73,
173, 1989.
55. Roosen-Runge, E.C., The Process of Spermatogenesis in Animals, Cambridge University Press, Cambridge,
1977, 214 pp.
56. Szollosi, A., Ultrastructural study of the spermatodesm of Locusta migratoria migratorioides ( R . and F.):
acrosome and cap formation, Acrida, 3, 175, 1974.
57. Anderson, J.M., A cytological study of the testicular cyst cells in the Japanese beetle, Physiol. Zool., 23,308,
1950.
58. Silberglied, R.E., Shepherd, J.G., and Dickinson, J.L., Eunuchs: the role of apyrene sperm in Lepidoptera,
Am. Nat., 123, 255, 1984.
59. Lai-Fook, J., Testicular development and spermatogenesis in Calpodes ethlius Stoll (Hesperiidae, Lepi-
doptera), Can. J. Zool., 60, 1161, 1982.
60. Dumser, J.B. and Davey, K.G., Endocrinological and other factors influencing testis development in
Rhodniusprolixus, Can. J. Zool., 52, 101 1, 1974.
61. Dumser, J.B. and Davey, KG., The Rhodnius testis: hormones, differentiation of the germ cells, and duration
of the molting cycle, Can. J. Zool., 53, 1673, 1975.
62a. Dumser, J.B. and Davey, K.G., The Rhodnius testis: hormonal effects on germ cell division, Can. J. Zool.,
53, 1682, 1975.
62b. Dumser, J.B., The regulation of spermatogenesisin insects, Annu. Rev. Entomol., 25, 341, 1980.
63. Koeppe, J.K., Fuchs, M.S., Chen, T.T., Hunt, L.-M., Kovalick, G.E., and Briers, T., The role of juvenile
hormone in reproduction, in Comprehensive Insect Physiology, Biochemistry and Pharmacology, Vo1.8,
Kerkut, G.A. and Gilbert, L.I., Eds., Pergamon Press, New York, 1985, 165.
64. Riemann, J.G. and Giebultowin, J.M., Secretion in the upper vas deferens of the gypsy moth correlated
with the circadian rhythm of sperm release from the testes, J. Insect Physiol., 37, 53, 1991.
65. Thorson, BJ. and Riemann, J.G., Abdominally entrained periodicities of testis and vas deferens activity in
the Mediterranean flour moth, J. Insect Physiol., 23, 1189, 1977.
66. Giebultowia, J.M., Riemann, J.G., Raina, A.K., and Ridgway, R.L., Circadian system controlling release
of sperm in the insect testes, Science, 245, 1098, 1989.
67. Riemann, J.G. and Thorson, BJ., Sperm maturation in the male and female genital tracts of Anagasta
kiihniella (Lepidoptera: Pyralididae), Int. J. Insect Morphol. Embryol., 1, 11, 1971.
68. Hinton, H.E., Accessory functions of seminal fluid, J. Med. Entomol., 11, 19, 1974.
69. Leopold, R.A., The role of male accessory glands in insect reproduction, Annu. Rev. Entomol., 21, 199,1976.
70. Chen, P.S., The functional morphology and biochemistry of insect male accessory glands and their secretions,
Annu. Rev. Entomol., 29, 233, 1984.
71. Happ, G.M., Structure and development of male accessory glands in insects, in Insect Ultrastructure, Vol.
2, King, R.C. and Akai, H., Eds., Plenum Press, New York, 365, 1984.
72. Gillott, C. and Gaines, S.B., Endocrine regulation of male accessory gland development and activity, Can.
Entomol., 124, 87 1, 1992.
73. Happ, G.M., Yuncker, C., and Dailey, PJ., Cytodifferentiationin the accessory glands of Tenebrio molitor.
VII. Patterns of leucine incorporation by the bean-shaped glands of males, J. Exp. ZooL, 220, 81, 1982.
74. Fallon, A.M. and Wyatt, G.R., Cyclic guanosine 3'. 5'-monophosphate. High levels in the male accessory
gland of Acheta domesticus and related crickets, Biochim. Biophys. Acta, 41 1, 173, 1975.
75. Fallon, A.M. and Wyatt, G.R., Guanylate cyclase in the accessory gland of the cricket, Acheta domesticus,
J. Insect Physiol., 23, 1037, 1977.
76. Shirk, P.D., Dahm, K.H., and Roller, H., The accessory sex glands as the repository for juvenile hormone
in male cecropia moths, Z Naturforsch. Teil C. 31, 199, 1976.
77. Shirk, P.D., Bhaskaran, G., and Roller, H., The transfer of juvenile hormone from male to female during
mating in the cecropia silk moth Hyalophora cecropia, Experientia, 36, 682, 1980.
78. Roth, L.M. and Dateo, G.P., Jr., Uric acid in the reproductive system of males of the cockroach Blattella
germanica, Science, 146, 782, 1964.
79. Roth, L.M. and Dateo, G.P., Jr., Uric acid storage and excretion by accessory sex glands of male
cockroaches, J. Insect Physiol., 11, 1023, 1965.
80a. Roth, L.M., Uricose glands in the accessory sex gland complex of male Blattaria, Ann. Entomol. Soc. Am.,
60,1203, 1967.
80b. Mullins, D.E. and Keil, CB., Paternal investment of urates in cockroaches, Nature, 283, 567, 1980.
52 Insect Reproduction

8 1. Wigglesworth, V.B., The function of the corpus allatum in the growth and reproduction of Rhodniusprolixus,
Q. J. Microsc. Sci., 79, 91, 1936.
82. Delbecque, J.-P., Weidner, K., and Hoffmann, K.H., Alternative sites for ecdysteroid production in insects,
Invert. Reprod. Dev., 18, 29, 1990.
83. Gee, J.D., Whitehead, D.L., and Koolman, J., Steroids stimulate secretion by insect Malpighian tubules,
Nature (London), 269, 238, 1977.
84. Hoffmann, K.H. and Behrens, W., Free ecdysteroids in adult male crickets, Gryllus bimaculatus, Physiol.
Entomol., 7, 269, 1982.
85. Cheeseman, M.T. and Gillott, C., Identification and partial characterization of the major secretory protein
of the long hyaline gland in the male grasshopper, Melanoplus sanguinipes, Insect Biochem., 18, 135, 1988.
86. Cheeseman, M.T. and Gillott, C., Corpus allatum and corpus cardiacum regulation of long hyaline gland
protein synthesis in the male grasshopper, Melanoplus sanguinipes, Gen. Comp. Endocrinol., 72,416, 1988.
87. Odhiambo, T.R., Site of action of the corpus allatum hormone at the cellular level in Schistocerca gregaria,
Acta Trop., 23, 264, 1966.
88. De Loof, A. and Lagasse, A., The ultrastructure of the male accessory reproductive glands of the Colorado
beetle, Z Zellforsch., 130, 545, 1972.
89. Couche, G.A. and Gillott, C., Development of secretory activity in the long hyaline gland of the male
migratory grasshopper, Melanoplus sanguinipes (Fabr.) (Orthoptera: Acrididae), lnt. J. Insect Morphol.
Embryol., 16, 355, 1987.
90. Ogiso, M. and Takahashi, S.Y., Trehalases from the male accessory glands of the American cockroach:
developmental changes and the hormonal regulation of the enzymes, Gen. Comp. Endocrinol., 55,387, 1984.
9 1. Hartmann, R., Der Einfluss endokriner Faktoren auf die manlichen Akzessorischen Drusen und die Ovarien
bei der Keulenheuschrecke Gomphocerus rufus L. (Orthoptera, Acrididae), 2. Vgl. Physiol., 74, 190, 1971.
92. Hartmann, R., Wolf, W., and Loher, W., The influence of the endocrine system on reproductive behavior
and development in grasshoppers, Gen. Comp. Endocrinol. Suppl., 3, 518, 1972.
93. Barker, J.F. and Davey, K.G., Neuroendocrine regulation of protein accumulation by the transparent
accessory reproductive gland of male Rhodnius prolixus, In!. J. Invertebr. Reprod., 3,291, 1981.
94. Glauser and Chen, unpublished data, in Schmidt, T., Stumm-Zollinger, E., and Chen, PS., Protein
metabolism of Drosophila melanogaster male accessory glands. 111. Stimulation of protein synthesis follow-
ing copulation, lnsect Biochem., 15, 391, 1985.
95. Yamamoto, K., Chadarevian, A., and Pellegrini, M., Juvenile hormone action mediated in male accessory
glands of Drosophila by calcium and kinase C., Science, 239,916, 1988.
96. Gold, S.M.W. and Davey, K.G., The effect of juvenile hormone on protein synthesis in the transparent
accessory gland of male Rhodnius prolixus, Insect Biochem., 19, 139, 1989.
97. Bar-Zev, A. and Kaulenas, M.S., Nucleic acid metabolism in adult tissues of Gromphadorhina in response
to ecdysterone: quantitative effects, J. lnsect Physiol., 21, 623, 1975.
98. Herman, W.S. and Barker, J.F., Ecdysterone antagonism, mimicry, and synergism of juvenile hormone
action on the monarch butterfly reproductive tract, J. Insect Physiol., 22, 643, 1976.
99. Sridevi, R., Bajaj, P., and Dutta-Gupta, A., Ecdysteroid stimulated protein synthesis in the male accessory
reproductive glands of Spodoptera litura, lnvertebr. Reprod. Dev., 14, 177, 1988.
100. Sridevi, R., Bajaj, P., and Dutta-Gupta, A., Hormonal regulation of macromolecular synthesis in testes and
accessory reproductive glands of Spodoptera litura during post-embryonic and adult development, Indian J.
Exp. Biol., 27, 699, 1989.
101. Ismail, P. and Dutta-Gupta, A., Effect of 20-hydroxyecdysone and inhibitors on the protein synthesis in
male accessory reproductive glands of Chilo partellus, Biochem. Arch., 6, 321, 1990.
102. Barker, J.F. and Davey, K.G., A polypeptide from the brain and corpus cardiacum of male Rhodnius
prolixus which stimulates in vitro protein synthesis in the transparent accessory reproductive gland, Insect
Biochem., 13, 7, 1983.
103. Friedel, T. and Gillott, C., Extraglandular synthesis of accessory reproductive gland components in male
Melanoplus sanguinipes, J. Insect Physiol., 22, 1309, 1976.
104a. Ismail, P. and Dutta-Gupta, A., In vitro uptake of larval haemolymph proteins by male accessory reproduc-
tive glands of the stem borer, Chilo partellus, lnvertebr. Reprod. Dev., 20, 193, 1991.
104b. Sevala, V.L. and Davey, K.G., The transparent accessory reproductive gland secretes a polypeptide into the
hemolymph of male Rhodnius prolixus, Insect Biochem., 2 1 , 215, 1991.
105. Baumann, H., The isolation, partial characterization,and biosynthesis of the paragonial substances, PS-I and
PS-2, of Drosophila funebris, J. Insect Physiol., 20, 2181, 1974.
106. Schmidt, T., Stumm-Zollinger, E., and Chen, P.S., Protein metabolism of Drosophila melanogaster male
accessory glands. Ill. Stimulation of protein synthesis following copulation, lnsect Biochem., 15, 391, 1985.
107. Cheeseman, M.T. and Gillott, C., Control of copulation-enhanced protein synthesisin the long hyaline gland
of the male grasshopper, Melanoplus sanguinipes, Arch. lnsect Biochem. Physiol., 11, 13, 1989.
108. Davey, K.G., Reproduction in the lnsecrs, Oliver and Boyd, Edinburgh, 1965, 96 pp.
Insect Male Mating Systems 53

109. Gerber, G.H., Evolution of the methods of spermatophore formation in pterygotan insects, Can. Enromol.,
102, 358, 1970.
110. Grimnes, K.A. and Happ, G.M., A monoclonal antibody against a structural protein in the spermatophore
of Tenebrio molitor (Coleoptera), Insect Biochem., 16, 635, 1986.
11 1. Grimnes, K.A., Bricker, CS., and Happ, G.M., Ordered flow of secretion from accessory glands to specific
layers of the spermatophore of mealworm beetles: demonstration with a monoclonal antibody, J. Exp. Zool.,
240,275, 1986.
112. Shinbo, H., Yaginuma, T., and Happ, G.M., Purification and characterization of a proline-rich secretory
protein that is a precursor to a structural protein of an insect spermatophore, J. Biol. Chem., 262,4794, 1987.
113. Cbeeseman, M.T., Gillott, C., and Ahmed, I., Structural spermatophore proteins and a trypsin-like enzyme
from the accessory reproductive glands of the male grasshopper, Melanoplus sanguinipes, J. Exp. Zool., 255,
193, 1990.
114. Gregory, G.E., On the initiation of spermatophore formation in the African migratory locust, Locusta
migratoria migratorioides Reiche and Fairmaire, J. Exp. Biol., 42, 423, 1965.
115. Loher, W., Circadian control of spermatophore formation in the cricket Teleogryllus commodus Walker, J.
Insect Physiol., 20, 1155, 1974.
116. Blum, M.S., Glowska, Z., and Taber, S., 111, Chemistry of the drone honey bee reproductive system. 11.
Carbohydrates in the reproductive organs and semen, Ann. Entomol. Soc. Am., 55, 134, 1962.
117. Vijayalekshmi, V. and Adiyodi, K.G., Accessory sex glands of male Periplaneta americana (L.). 111.
Histochemistry of the mushroom-shaped and conglobate glands, Indian J. Exp. Biol., 11, 521, 1973.
118. Odhiambo, T.R., The architecture of the accessory reproductive glands of the male desert locust. I. Types
of glands and their secretions, Tissue Cell, 1, 155, 1969.
119. Blum, MS., Bumgarner, J.E., and Taber, S., 111, Composition and possible significance of fatty acids in
the lipid classes in honey bee semen, J. Insect Physiol., 13, 1301, 1967.
120. Novak, A.F., Blum, M.S., Taber, S., 111, and Luizzo, J.A., Separation and determination of seminal plasma
and sperm amino acids of the honey bee, Apis mellifera, Ann. Entomol. Soc. Am., 53, 841, 1960.
121. Roth, L.M., The stimuli regulating reproduction in cockroaches, Colloq. Int. C.N.R.S., 189, 267, 1970.
122. Gillott, C. and Friedel, T., Fecundity-enhancing and receptivity-inhibiting substances produced by male
insects: a review, in Advances in Invertebrate Reproduction, Vol. 1, Adiyodi, K.G. and Adiyodi, R.G., Eds.,
Peraiam-Kenoth, Karivellur, India, 1977, 199.
123. Leopold, R.A., Cytological and cytochemical studies on the ejaculatory duct and accessory secretion in
Musca domestica, J. Insect Physiol., 16, 1859, 1970.
124. Riddiiord, L.M. and Ashenhurst, J.B., The switchover from virgin to mated behavior in female Cecropia
moths: the role of the bursa copulatrix, Biol. Bull. Woods Hole Mass., 144, 162, 1973.
125. Loher, W. and Edson, K., The effect of mating on egg production and release in the cricket, Teleogryllus
cornmodus, Entomol. Exp. Appl., 16, 483, 1973.
126. Destephano, D.B. and Brady, U.E., Prostaglandin and prostaglandin synthetase in the cricket, Acheta
domesticus, J. Insect Physiol., 23, 905, 1977.
127. Davey, K.G., Copulation and egg-production in Rhodniusprolixus: the role of the spermathecae, J. Exp. Biol.,
42, 373, 1965.
128. Manning, A., The control of sexual receptivity in female Drosophila, Anim. Behav., 15, 239, 1967.
129. Merle, J., Fonctionnement ovarien et rkceptivitk sexuelle de Drosophila melanogaster ap&s implantation de
fragments de I'appareil genital mile, J. Insect Physiol., 14, 1159, 1968.
130. Nelson, D.R., Adams, T.S., and Pomonis, J.G., Initial studies on the extraction of the active substance
inducing monocoitic behavior in house flies, black blow flies, and screw-worm flies, J. Econ. Entomol., 62,
634, 1969.
131. Baumann, H., Wilson, KJ., Chen, P.S., and Humhel, R.E., The amino acid sequence of a peptide (PS-l)
from Drosophila funebris: a paragonial peptide from males which reduces the receptivity of the female, Eur.
J. Biochem., 52, 521, 1975.
132. Chen, PS., Biochemistry and molecular regulation of the male accessory gland secretions in Drosophila
(Diptera), Ann. Soc. Entomol. Fr. (N.S.), 27, 231, 1991.
133. Friedel, T. and Gillott, C., Male accessory gland substance of Melanoplus sanguinipes: an oviposition
stimulant under the control of the corpus allatum, J. Insect Physiol., 22, 489, 1976.
134. Lange, A.B. and Loughton, B.G., An oviposition-stimulating factor in the male accessory reproductive gland
of the locust, Locusfa rnigratoria, Gen. Comp. Endocrinol., 57, 208, 1985.
135. Fuchs, M.S., Craig, G.B., Jr., and Despommier, D.D., The protein nature of the substance inducing female
monogamy in Aedes aegypti, J. Insect Physiol., 15, 701, 1969.
136. Destephano, D.B., Brady, U.E., and Lovins, R.E., Synthesis of prostaglandin by reproductive tissue of the
house cricket, Acheta domesticus, Prostaglandins, 6, 71, 1974.
137. Destephano, D.B., Brady, U.E., and Woodall, L.B., Partial characterization of prostaglandin synthetase in
the reproductive tract of the male house cricket, Acheta domesticus, Prostaglandins, 11, 261, 1976.
54 Insect Reproduction

138. Loher, W., Ganjian, I., Kubo, I., Stanley-Samuelson, D., and Tobe, S.S., Prostaglandins: their role in egg-
laying of the cricket Teleogryllus commodus, Proc. Natl. Acad. Sci. U.S.A., 78, 7835, 1981.
139. Monsma, S.A. and Wolfner, M.F., Structure and expression of a Drosophila male accessory gland gene
whose product resembles a peptide pheromone precursor, Genes Dev., 2, 1063, 1988.
140. DiBenedetto, AJ., Harada, H.A., and Wolfner, M.F., Structure, cell-specific expression, and mating-
induced regulation of a Drosophila melanogaster male accessory gland gene, Dev. Biol., 139, 134, 1990.
141. Monsma, S.A., Harada, H.A., and Wolfner, M.F., Synthesis of two Drosophila male accessory gland
proteins and their fate after transfer to the female during mating, Dev. Biol., 142, 465, 1990.
142. Yamaja Setty, B.N. and Ramaiah, T.R., Isolation and identification of prostaglandins from the reproductive
organs of male silkmoth, Bombyx mori L., Insect Biochem., 9, 613, 1979.
143. Yamaja Setty, B.N. and Ramaiah, T.R., Effect of prostaglandins and inhibitors of prostaglandin biosynthe-
sis on oviposition in the silkmoth Bombyx mori, Indian J. Exp. Biol., 18, 539, 1980.
144. Lafon-Cazal, M., Gallois, D., Lehouelleur, J., and Bockaert, J., Stimulatory effects of male accessory-
gland extracts on the myogenicity and the adenylate cyclase activity of the oviduct of Locusta migratoria, J.
Insect Physiol., 33, 909, 1987.
145. Paemen, L., Schoofs, L., and De Loof, A., Presence of myotropic peptides in the male accessory glands of
Locusta migratoria, J. Insect Physiol., 36, 861, 1990.
146. Paemen, L., Schoofs, L., Proost, P., Decock, B., and De Loof, A., Isolation, identification and synthesis of
Lom-AG-myotropin 11, a novel peptide in the male accessory reproductive glands of Locusta migratoria,
Insect Biochem., 21,243, 1991.
147. Lange, A.B., The neural and hormonal control of locust oviducts and accessory structures, Adv. Comp.
Endocrinol., 1, 109, 1992.
148. Baumann, H., Biological effects of paragonial substances, PS-I and PS-2, in females of Drosophila funebris,
J. Insect Physiol., 20, 2347, 1974.
149. Friedel, T. and Gillott, C., Contribution of male-produced proteins to vitellogenesis in Melanoplus sanguinipes,
J. Insect Physiol., 23, 145, 1977.
150. Terranova, A.C., Leopold, R.A., Degrugillier, M.E., and Johnson, J.R., Electrophoresis of the male
accessory secretion and its fate in the mated female, J. Insect Physiol., 18, 1573, 1972.
151a. Ruegg, R.P., Orchard, I., and Davey, K.G., 20-Hydroxyecdysone as a modulator of electrical activity in
neurosecretory cells of Rhodnius prolixus, J. Insect Physiol., 28, 243, 1982.
151b. Orchard, I., Ruegg, R.P., and Davey, KG., The role of central aminergic neurons in the action of 20-
hydroxyecdysone on neurosecretory cells of Rhodnius prolixus, J. Insect Physiol., 29, 387, 1983.
152a. Yamaoka, K. and Hirao, T., Stimulation of virginal oviposition by male factor and its effect on spontaneous
nervous activity in Bombyx mori, J. Insect Physiol., 23, 57, 1977.
152b. Yamaoka, K., The central nervous function in ovipositional behaviour of Bombyx mori with special reference
to the spontaneous nervous activity, in Advances in Invertebrate Reproduction, Vol. 1, Adiyodi, K.G. and
Adiyodi, R.G., Eds., Peralam-Kenoth, Karivellur, India, 1977, 414.
153. Borovsky, D., The role of the male accessory gland fluid in stimulating vitellogenesisin Aedes taeniorhynchus,
Arch. Insect Biochem. Physiol., 2, 405, 1985.
154. Downe, A.E.R., Internal regulation of rate of digestion of blood meals in the mosquito, Aedes aegypti, J.
Insect Physiol., 21, 1835, 1975.
155. Klowden, M.J. and Chambers, G., Male accessory gland substances activate egg development in nutrition-
ally stressed Aedes aegypti mosquitoes, J. Insect Physiol., 37, 721, 1991.
156. Huignard, J., Quesneau-Thierry, A., and Barbier, M., Isolement, action biologique et evolution des
substances paragoniales contenues dans le spermatophore d'Acanthoscelides obtectus (ColCopttre), J. Insect
Physiol., 23, 351, 1977.
157. Sasaki, M., Riddiford, L.M., Truman, J.W., and Moore, J.K., Re-evaluationof the role of corpora cardiaca
in calling and oviposition behaviour of giant silk moths, J. Insect Physiol.. 29, 695, 1983.
158. Leopold, R.A., Terranova, A.C., and Swilley, E.M., Mating refusal in Musca domestica: effects of repeated
mating and decerebration upon frequency and duration of copulation, J. Exp. Zool., 176, 353, 1971.
159. Leopold, R.A., Terranova, A.C., Thorson, BJ., and Degrugillier, M.E., The biosynthesis of the male
housefly accessory secretion and its fate in the mated female, J. Insect Physiol., 17, 987, 1971.
160. Gwadz, R.W., Neuro-hormonal regulation of sexual receptivity in female Aedes aegypti, J. Insect Physiol.,
18, 259, 1972.
161. Smith, P.H., Gillott, C., Barton Browne, L., and van Gerwen, A.C.M., The mating-induced refractoriness
of Lucilia cuprina females: manipulating the male contribution, Physiol. Entomol., 15, 469, 1990.
162. Davey, K.G., The migration of spermatozoa in the female of Rhodnius prolixus Stil, J. Exp. Biol., 35,694,
1958.
163. Shepherd, J.G., A polypeptide sperm activator from male saturniid moths, J. Insect Physiol., 21, 9, 1975.
164. Shepherd, J.G., Sperm activation in saturniid moths: some aspects of the mechanism of activation, J. Insect
Physiol., 20, 232 1, 1974.
Insect Male Mating Systems 55

165. Osanai, M., Aigaki, T., and Kasuga, H., Arginine degradation cascade as an energy-yielding system for
sperm maturation in the spermatophore of the silkworm, Bombyx mori, in New Horizons in Sperm Cell
Research, Mohri, H., Ed., Japan Scientific Society Press, Tokyo, 1987, 185.
166. Aigaki, T., Kasuga, H., and Osanai, M., A specific endopeptidase,BAEE esterase, in the glandula prostatica
of the male reproductive system of the silkworm, Bombyx mori, Insect Biochem., 17, 323, 1987.
167. Osanai, M., Kasuga, H., and Aigaki, T., Induction of motility of apyrene spermatozoa and dissociation of
eupyrene sperm bundles of the silkworm, Bombyx mori by initiatorin and trypsin, Invertebr. Reprod. Dev., 15,
97, 1989.
168. Aigaki, T., Osanai, M., and Kasuga, H., Arginine carboxypeptidase activity in the male reproductive glands
of the silkworm, Bombyx mori, Insect Biochem., 18, 295, 1988.
169. Kasuga, H., Aigaki, T., and Osanai, M., System for supply of free arginine in the spermatophore of Bombyx
mori. Arginine-liberating activities of contents of male reproductive glands, Insect Biochem., 17, 3 17, 1987.
170. Osanai, M., Aigaki, T., Kasuga, H., and Yonezawa, Y., Role of arginase transferred from the vesicula
seminalis during mating and changes in amino acid pools of the spermatophore after ejaculation in the
silkworm, Bombyx mori, Insect Biochem., 16, 879, 1986.
17 1. Osanai, M., Aigaki, T., and Kasuga, H., Energy metabolism in the spermatophore of the silkmoth, Bombyx
mori, associated with accumulation of alanine derived from arginine, Insect Biochem., 17, 71, 1987.
172. Takahashi, S.Y., Higashi, S., Minoshima, S., Ogiso, M., and Hanaoka, K., Trehalases from the American
cockroach, Periplaneta americana: multiple occurrence of the enzymes and partial purification of enzymes
from male accessory glands, In!. J. Invertebr. Reprod., 2, 373, 1980.
173. Yaginuma, T. and Happ, G.M., Trehalase from the bean-shaped accessory glands and the spermatophore
of the male mealworm beetle, Tenebrio molitor, J. Comp. Physiol. B, 157, 765, 1988.
174. Ogiso, M., Shinohara, Y., Hanaoka, K., Kageyama, T., and Takahashi, S.Y., Further purification and
characterization of trehalases from the American cockroach, Periplaneta americana, J. Comp. Physiol. B, 155,
553, 1985.
175. Gilbert, D.G., Ejaculate esterase 6 and initial sperm use by female Drosophila melanogaster, J. Insect
Physiol., 27, 641, 1981.
176. Vander Meer, R.K., Obin, M.S., Zawistowski, S., Sheehan, K.B., and Richmond, R.C., A reevaluation
of the role of cis-vaccenyl acetate, cis-vaccenol and esterase 6 in the regulation of mated female sexual
attractiveness in Drosophila melanogaster, J. Insect Physiol., 32, 681, 1986.
177. Schmidt, T., Stumm-Zollinger, E., Chen, PS., Biihlen, P., and Stone S.R., A male accessory gland peptide
with protease inhibitory activity in Drosophila funebris, J. Biol. Chem., 264, 9745, 1989.
178. Gwynne, D.T., Male nutritional investment and the evolution of sexual differences in Tettigoniidae and other
Orthoptera, in Orthopteran Mating Systems, Gwynne, D.T. and Morris, G.K., Eds., Westview Press, Boulder,
CO, 1983, 337.
179. Pickford, R. and Gillott, C., Insemination in the migratory grasshopper, Melanoplus sanguinipes (Fab.),
Can. J. Zool., 49, 1583, 1971.
180. Boggs, C.L. and Gilbert, L.E., Male contribution to egg production in butterflies: Evidence for transfer of
nutrients at mating, Science, 206, 83, 1979.
181. Huignard, J., Transfer and fate of male secretions deposited in the spermatophoreof females ofAcanthoscelides
obtectus Say (Coleoptera Bmchidae), J. lnsect Physiol., 29, 55, 1983.
182. Markow, T.A. and Ankney, P.F., Drosophila males contribute to oogenesis in a multiple mating species,
Science, 224, 302, 1984.
183. Boucher, L. and Huignard, J., Transfer of male secretions from the spermatophore to the female insect in
Caryedon serratus (01.): analysis of the possible trophic role of these secretions, J. lnsect Physiol., 33, 949,
1987.
 Chapter 3

SEX DETERMINATION IN INSECTS

Roger L. Blackman

CONTENTS
I. Introduction .................................................................................................................
57

11. General Aspects of Sex Determination in Insects ....................................................58

A. XX/XY Systems ..................................................................................................
58
B. XX/XO Systems ...................................................................................................
59
C. Multiple Sex Chromosome Systems .................................................................... 61
D. "Multiple Factor" Systems ..................................................................................
63
E. Haplodiploid Sex Determination ..........................................................................63
F. The Molecular Basis of Sex Determination ......................................................... 64
G. Dosage Compensation .........................................................................................
67

111. Sex Determination in Different Groups of Insects ..................................................... 67

A. Apterygota .............................................................................................................
67
B. Primitive Exopterygota ........................................................................................ 67
C. The Orthopteroid Orders ..................................................................................... 68
D. The Hemipteroid Orders ....................................................................................... 71
E. Neuropteroidea and Coleoptera ...........................................................................75
F. Hymenoptera .......................................................................................................
77
G. The Panorpoid Orders ........................................................................................... 78

IV. Evolution of Sex Chromosomes and Sex Determination in Insects ..........................85

References ............................................................................................................................
86

I. INTRODUCTION
Sex determination is the process by which the gender of a bisexual organism becomes
fixed, so that the individual progeny develops either as a son or a daughter. As is the case with
other fundamental biological processes, evolution has in the course of time produced a
seemingly infinite variety of ways of achieving this one essentially simple objective, and
classical genetic and cytogenetic observations have, over the years, combined to display a
bewildering diversity of sex-determiningmechanisms. Much of this work has been on insects,
from the first recognition of sex chromosomes in the heteropteran Pyrrhocoris apterus,'
through the classic experiments of bridge^^-^ on Drosophila and Golds~hmidt~.~ on Porthetria
dispar, to the recent molecular work elucidating the hierarchy of regulatory genes responsible
for the sex of fruit f l i e ~ . ~ . ~
In the general literature on sex determination, two works stand out?,1° each with a radically
different approach to the subject. Both cover the full range of sex-determining systems, but
the cytogeneticist Whiteg gives pride of place to evolutionary changes in the sex chromo-
somes, whereas the evolutionary geneticist Bulllo pays more attention to the underlying

0-8493-6695-X/95/50.MkS.50
8 1995 by CRC Press. Inc.
58 Insect Reproduction

mechanisms. The problem with reviewing sex determination in insects at the present point in
time is that, for all orders except Diptera, the greater part of the evidence is cytological, and
even the most basic information about the genetic systems involved is not usually available.
It is nevertheless worthwhile to follow the lead of Nothiger and Steinmann-Zwicky,ll and look
for general principles of sex determination that could be applicable to all insects, and possibly
to all biparental organisms.
In this chapter I shall start by reviewing the main types of sex determination found in
insects, then outline what is known about sex chromosome systems in each insect order, and
end with some discussion of the evolutionary implications of what we now know about sex
determination.

11. GENERAL ASPECTS OF SEX DETERMINATION IN INSECTS

The sex of an insect is almost always determined genetically. Hermaphroditism, in the
sense of the same genotype producing functional male and female organs in the same
individual, seldom occurs in insects; it seems to have evolved only once, in one genus of scale
insects (Icerya). An oft-quoted second example of insect hermaphroditism,in the termiticolous
phorid group Termitoxeriinae, has now been refuted.I2 (True hermaphrodites should not be
confused with gynandromorphs and intersexes - genetically abnormal individuals - which
are of common occurrence in insects.) Environmental factors may sometimes influence the
genetic determination of sex (see BergerardI3for review), but the type of environmental sex
determination that occurs widely in reptiles, for example, where the sex of an individual is
decided by the environment of the egg after it has been fertilized, seems to be rare in insects.

A. XXIXY SYSTEMS
Most bisexual organisms produce a 1:1 sex ratio, and this can be achieved simply by having
one sex (the heterogametic sex) produce two genetically different types of gamete, and the
other sex (the homogametic sex) produce gametes of only one of these types. The two types
of gamete carry different sex factors (here given the notation S, and S,), which segregate from
one another in Mendelian fashion in the meiosis of the heterogametic sex:

s,s, X s,s, (parents) + 1:l s,s, and sls2(progeny)

The heterogametic sex is the male in most insects, but female in Lepidoptera, Trichoptera, and
some Diptera.
Although these sex factors are conventionally regarded as alternative alleles at the same
locus, it is usually the case that only one sex factor plays an active part in the determination
of sex. The other "sex factor" may merely be the corresponding site on a homologous
chromosome: e.g., a nonfunctioning (null) allele, or the location at which the sex factor is
inserted, in the case of a transposable element.
In some cases, sex factors may be inherited as single genes, recombining freely with other
genes on the same chromosome pair, although they are more often than not tightly linked to
other genes involved in sex differentiation. Very often, however, chromosomes carrying sex
factors are cytologically distinct (heteromorphic), so that the inheritance of sex can be
observed cytologically:

XX X XY (parents) + 1:1 XX and XY (progeny)

X and Y chromosomes usually pair at meiosis before segregating to opposite poles, but there
is normally little or no recombinational exchange between them. This is called an XXIXY, or
XY (male) sex determination system. When the female is the heterogametic sex, the sex
chromosomes are sometimes termed Z and W, and the system called ZWIZZ:
Sex Determination in Insects 59

ZW (female parent) X ZZ (male parent) 4 1:l ZZ and ZW (progeny)

but this terminology has now been largely abandoned in the literature on insect cytogenetics
as an unnecessary complication. The fact that the female is the heterogametic sex can be
indicated by putting the heterozygous genotype first; i.e., XY/XX or XY (female) sex
determination.
At this stage it is important to address two common misconceptions about sex chromo-
somes, which can easily hinder understanding of sex-determining mechanisms and their
evolution. The first point concerns the common notation of heteromorphic sex chromosomes
throughout both plant and animal kingdoms as X and Y, which might be thought to imply
some degree of homology, not just at the sex-determining loci but of the chromosome as a
whole, across major groups of organisms. On the contrary, there is no doubt that heteromor-
phic sex chromosomes have evolved many times independently in different taxa.I4 The Y
chromosome in particular can sometimes be an extremely labile structure, apparently under-
going cycles of degeneration and regeneration within taxa. These will be discussed further
later, and in order to emphasize this lability it is sufficient here to note that what seem to be
major changes in sex chromosome constitution, such as the formation of a Y chromosome de
novo (a "neo-Y"), can be found even within a single species.
The second common misconception is that the Y chromosome always has a dominant,
male-determining function. This is certainly the case in mammals and in some insects, but the
more general condition in insects with XXJXY systems is for the Y chromosome to have an
essentially passive role, influencing sex merely by segregating opposite the X at meiosis. The
sex of the zygote is then determined by the balance between the actions of regulatory genes
on the X chromosome and on the autosomes. This is roughly equivalent to the "genic balance"
model developed by Bridges3from his work on Drosophila. The zygote must be homozygous
for a sex factor in order to be female (XX), so this has also been termed a "recessive-X
~ystem".'~
Genetic studies are obviously needed to establish for certain whether the system operating
in any one species is based on a dominant Y or genic balance. For species with heteromorphic
sex chromosomes, deductions are possible by observing the sex of individuals with abnormal
sex chromosome constitutions. The two most informative abnormalities are XXY and XO. If
XXY individuals of a species which normally has an XXIXY system are male, then there is
obviously a dominant Y factor operating, but if XXY is female, then the Y chromosome is
likely to be sexually inert, and the X has an active, although recessive, role. Frequently,
aberrant individuals completely lacking a Y chromosome (i.e., with X 0 constitution) are at
least viable enough to observe their sex; such individuals will be female in dominant-Y
systems, but male in genic balance systems, as the latter require XX for female determination.
Evidence like this is available for only relatively few insects, mostly Diptera, in which occur
both dominant-Y (e.g., Phormia regina, Lucilia cuprina15) and recessive-X systems (e.g.,
Drosophila melanoga~ter,~ Glossina palpali~'~). However, there are good reasons for believ-
ing that genic balance is the most general condition in insects, stemming from the widespread
occurrence of XXIXO sex determination in many insect order^.^

B. XXJXO SYSTEMS
In organisms with heteromorphic sex chromosomes, X-Y recombination is usually sup-
pressed, and the Y chromosome tends to be more degenerate than the X, often having few or
no functional alleles. This degeneration of the Y is generally perceived as a progressive
evolutionary phenomenon.I0Various explanations for this have been offered.l4.I7For example,
because the Y is permanently heterozygous and nonrecombinant, selection must act at the
level of the entire chromosome, so it evolves as "an asexual component of an otherwise sexual
genome".18 Deleterious mutations (often nonfunctional alleles) will tend to accumulate in the
absence of recombination by the process known as "Muller's ratchet,"lg which may be
 Insect Reproduction

(a) "Muller's ratchet" (h) "Hitchhiking"

deleterious
mutation

beneficial
mutation

FIGURE 1. Degeneration of a Y chromosome by operation of (a) Muller's "ratchet," and (b) "hitchhiking." In a
population of Y chromosomes free from recombination (left), some Ys will have one or more mutations to
nonfunctional alleles (black segments), with different Ys mutated at different loci. If the selective disadvantage per
locus is small and population size is small enough relative to the mutation rate, the class of Y chromosomes with no
mutations may be lost by chance and, because there is no recombination, it cannot be restored. Next the class canying
just one mutation becomes vulnerable to chance loss, and so on, so that as time goes on the mean numbers of mutations
per Y chromosome gradually increases. If a favorable Y-linked mutation should occur, however (a),it could spread
rapidly through the population by selection, carrying with it any nonfunctional alleles that happen to be present on
the Y chromosome in which it originated ("hitchhiking"), leading to fixation of nonfunctional alleles and accelerating
the degenerative process.

accelerated by a "genetic hitchhiking" effect18 (Figure 1). However, there is still no entirely
satisfactory explanation.
The end point of such an evolutionary process may be the complete loss of the Y
chromosome, so that males are XO. An XX/XO sex determination system works in exactly
the same way as an XX/XY system, with the X moving to one pole in male meiosis, so that
sperm are either with or without an X:

XX (female parent) X X 0 (male parent) + 1:1 XX and X 0 progeny

Obviously, since there is no Y, such a system must be based on genic balance. XXIXO sex
determination occurs in almost all orders of insects, both primitive and advanced. It is almost
certainly the ancestral form of sex determination of orthopteroid insects and probably of other
major orders such as Hemiptera and Coleoptera. It is even possible that the ancestors of all
insects had X 0 males, and that all insect Y chromosomes have arisen de novo.
Y chromosomes in insects often seem to arise as a result of centric fusion between an X
chromosome and an autosome (e.g., Figure 2). A recently formed, autosomally derived Y
chromosome (a neo-Y) is often easily recognized because it is likely to be still homologous
with the autosomal part of the neo-X, and therefore synapses with it at meiosis. This homology
may be gradually lost in the course of evolution as X-Y recombination becomes suppressed,
and secondary structural and genetic changes occur independently on both the neo-X and the
neo-Y.
Clearly, a neo-Y chromosome cannot carry a dominant factor for male determination, and
sex determination in such cases must be based on genic balance, as in the ancestral XXIXO
system. Neo-XY systems occur most frequently in groups in which XXIXO systems are
common; these include the orthopteroid orders, Odonata, the hemipteroid orders, and Co-
leoptera. Each of these groups will be discussed later. In all of them, there are also species with
multiple sex chromosomes.
Sex Determination in Insects

< plus qy

Autosome

FIGURE 2. White'sg model of the origin of neo-XY sex determination from an X 0 condition in the heterogametic
sex. After breakage near the centromeres of an X and an autosome, centric fusion occurs, creating a neo-X
chromosome. When this fusion has reached fixation in the population (i.e., when the original unfused Xs no longer
occur), the original homologue of the autosome involved in the fusion will be confined to the male line and will act
as a "neo-Y", segregating opposite the neo-X at meiosis. (White9believed that centromeres were never situated at
the extreme ends (telomeres) of chromosomes, and therefore his models assume arm breaks and exchanges, followed
by loss of a minute chromosome. Alternative models of centric fusion involving breakage within centromeres are
discussed by John and H e ~ i t t . ' ~ ~ )

C. MULTIPLE SEX CHROMOSOME SYSTEMS

Chromosomal differences between male and female insects normally involve only one
chromosome pair (X and Y, or just the X in XX/XO systems), but there are numerous cases
in most insect orders where the difference between the sexes involves a larger number of
chromosomes. The most common "multiple" sex chromosome systems have just two Xs (the
notation used for the male condition X,X,Y, or X,X,O, and the notation for the sex determi-
nation system is X,X,X,X2/X,X2Y,or X,X,X2X2/X,X20),although species are known with
almost any number of Xs from 1 to 6 (and in one extreme case, 12), and any number of Ys
from 1 to 6.
The origins of X,X,X2X2/X,X2Ysex determination are usually fairly simply explained,
with good evidence provided by the way in which the X,, X,, and Y associate at metaphase
I of spermatogenesis. Figure 2 showed the origin of a neo-XY condition by centric fusion,
resulting in a metacentric X (i.e., with the centromere near the center of the chromosome) and
an acrocentric Y (with the centromere at one end). If a further centric fusion should occur, this
time between the Y chromosome and another acrocentric autosome, then an X,X,Y condition
will arise, with a metacentric Y (Figure 3).
Alternatively, an X,X2Y condition may be derived directly from an XO, by a reciprocal
translocation between the X and one member of an autosome pair, the other member of the
autosome pair becoming a neo-Y (Figure 4).
The other common derivation of a multiple sex chromosome system is that found in
organisms with holocentric chromosomes (i.e., chromosomes with diffuse centromeric activ-
ity), such as the Hemiptera and Dermaptera, where X,, X,, etc., have almost certainly arisen
simply by dissociation (fission) of the single original X chromosome into two or more parts,
which still segregate to the same pole at meiosis (Figure 5). Such X dissociations have
occurred in both XXIXY and XX/XO systems, a single X dissociation giving X,X,X,Xfl,X,Y
and X,X,X,X~,X,O, respectively. Multiple Ys have also arisen by dissociation in some
species. It is characteristic of multiple sex chromosome systems formed by dissociation that the
X,, X,, etc. are much smaller than the original X, and that there is no accompanying change in
the number of autosomes.
Multiple sex chromosomes are cytogenetically interesting, because they show very clearly
the fixation of different types of chromosomal rearrangement, but they have little or no
 Insect Reproduction

-
plus c@
Autosome (lost)

FIGURE 3. Origin of an X,X,Y condition in the heterogametic sex (leading at fixation to an X,X,X,XdX,X,Y sex
determination mechanism), by centric fusion between an acrocentric Y and an acrocentric autosome, to give a
metacentric "neo-Y" ("neo" because part of it is recently derived from an autosome), and a neo-X,. The arrangement
of sex chromosomes on the spindle of the first meiotic division, with homologous sections associated, is shown
diagrammatically. (Adapted from White, M. Animal Cytology and Evolution, 3rd ed., Cambridge University Press,
Cambridge, U.K., 1973.)

.....
......
......
......
.: :.....
......
:::.
......
......
......
......
....... ......
......
......
......
.......
.....
...... ......
......
......
......
......
......
.......
...... ......
......
....
......
......
..... X2

neo-y

Autosome
1
FIGURE 4. Origin of an X,X,Y condition in the male (leading at fixation to X,X,X,X~X,X,Y sex determination)
from an XXIXO system, by reciprocal translocation between a single metacentric X in the male and a metacentric
autosome. The arrangement on the spindle of the first meiotic division, with homologous sections associated
terminally, and X, and X, moving to the opposite pole from the Y chromosome, is shown diagrammatically. (Adapted
from White, M. Animal Cytology and Evolution, 3rd ed., Cambridge University Press, Cambridge, U.K., 1973.)

FIGURE 5. Multiple X chromosomes (in [b] and [c]) are derived from an XY system (a) by simple dissociation.
Segregation of Xs and Y at spermatogenesis is represented diagrammatically, with autosomes not shown. Usually in
such systems the Xs and Y do not associate, but show "distance" (or "touch-and-go") pairing.
Sex Determination in Insects 63

significance in relation to the molecular genetic basis of sex determination. Of more interest
in this respect are the so-called "multiple factor" systems of certain Diptera.

D. "MULTIPLE FACTOR" SYSTEMS

Species with multiple sex chromosomes are nevertheless still likely to have only a single
sex-determining locus - for example, on just one of the Xs in a multiple X system. But more
complex sex-determination systems are known where one species may have sex-determining
factors at several different loci, and sometimes even different individuals within the same
population have their sex factor on different chromosomes. Such complex systems cannot be
detected cytogenetically, and they have only been recognized in genetically well-studied
organisms with an abundance of genetic markers. In insects, this means certain species of
Diptera.
The type of multiple factor system most commonly observed in Diptera and found in
members of both the Nematocera and the Cyclorrhapha, can be symbolized in a general way
as follows

Females Males
SlSl S2S2 Slsl s2s2
SlSl S2s2

S, and S, behave as dominant male-determining factors, segregating, respectively, at the two

different loci 1 and 2, and always restricted to the male line. This 2-locus system, with S, and
S,, can be generalized to any number of loci:

Females Males
....
S I S l S2S2 S3S3 Slsl S2S2 S3S3....
sls, S2s2s,s ,....
s,s, s2s2S,s ,....

So for any species with a multiple factor system, there is always just one female genotype,
homozygous for all sex factor loci, and n distinct male genotypes, each heterozygous at just
one of the n loci. Sometimes a sex factor locus may be on a cytologically distinct sex
chromosome, so that S,, for example, is manifestly a Y chromosome; but sex factors may
equally occur on chromosomes that are in all other respects autosomal, in which case there is
no obvious cytological difference between the sexes. Dipteran geneticists have termed the
autosomal male-determining factors "M factors". Green20suggested that M factors at different
loci in any one species were perhaps all actually the same gene, transposed to several different
sites in the genome. The evidence now strongly favors this interpretation in several cases
(discussed later under Diptera). This explains the exclusive nature of their occurrence in
individual males, but makes the term "multiple sex factors" something of a misnomer.
This kind of sex determination may occur in other insects, especially where heteromorphic
sex chromosomes have not been detected or do not occur regularly, but the necessary genetic
evidence is lacking for other orders apart from Diptera.

E. HAPLODIPLOID SEX DETERMINATION

No general survey of sex determination in insects would be complete without mention of
haplodiploidy, of which Hymenoptera are, of course, the leading exponents. Haplodiploidy
may also be general to Thysanoptera, and is found in some species of Homoptera and
Coleoptera (as well as occurring widely in mites and ticks). The genetic mechanisms involved
have been the subject of much speculation (reviewed by C r o ~ i e r ~Indeed,
~ ~ ~ ~at) first
. sight, it
is difficult to see how any genetic mechanism at all can be operating, as haploid males simply
64 Insect Reproduction

have half the female dose of all genes. However, several hypotheses based on multiple sex
factors have been suggested, and one has some experimental foundation.
It has long been known that, in certain Hymenoptera, inbreeding results in diploid male^,^^-^^
indicating that not only haploids but diploid homozygotes are male. This can be explained if
there are multiple sex factors (S,S2S,...) - possibly alternative alleles at a single locus -
segregating in opposition:

Females Males
SlS2, SIS,, SzS,, ... S,, S,, S,, ... (if eggs unfertilized)
or S$,, S,S2, S$,, ... (in inbred populations)

The sex factors appear to complement one another, and this has therefore been termed a
complementary sex-determining mechanism.
While this explanation fits members of several groups of Hymenoptera very well (e.g.,
Habrobracon, Apis, Neodiprion, and Solenopsis), it cannot apply generally, because some
other Hymenoptera inbreed considerably, yet fail to produce diploid males. To accommodate
this problem, the hypothesis can be modified to involve multiple loci.21The theory is that
diploids would then have to be homozygous at all loci in order to be male, and this would only
be likely after long-term intensive inbreeding.

F. THE MOLECULAR BASIS OF SEX DETERMINATION

Thus there is a variety of ways in which sex can be determined in insects. Sex factors can
apparently determine either maleness or femaleness, can be dominant or recessive in their
action, can be single or multiple, and can occur on sex chromosomes or autosomes. How can
all this be explained in molecular terms?
Our knowledge of the molecular biology of sex determination is almost entirely restricted
to D. melanogaster, but at least in this one species some of the details of the mechanism are
now worked out. The key gene is Sexlethal (Sxl), which is located on the X chromosome. This
gene is essential for determination of females, but completely functionless in males, so that
it can be eliminated by mutation without affecting the male phenotype. There is a maternal
gene, daughterless (da), that has to be active for Sxl to function, because a mutation at the da
locus causes mothers to produce only sons; however, da is not normally involved in sex
determination. Activation of the female-determining function of Sxl in Drosophila is in fact
dependent in some way on the ratio between the number of X chromosomes and the number
of sets of autosomes (henceforth X:A). Thus, if X:A is 1.0 (as in diploid eggs with two X
chromosomes), Sxl produces an active product that causes the embryo to develop as female,
but if X:A is 0.5 (e.g., diploid eggs with XY or XO) then Sxl is silent and the embryo develops
as male.
The genes repressing Sxl in male eggs have not been identified, but in Drosophila they must
be located on the autosomes, because a genotype with one X chromosome and one set of
autosomes (X + A) is female, whereas the addition of another autosome set (X + AA) results
in a male. The molecular nature of the X:A signal is a source of continuing ~peculation.~~
ChandraZ7proposed that the normal, diploid forms of both sexes produce the same limited
number of repressor (R) molecules; two X chromosomes can bind all the R molecules so that
Sxl can be transcribed, but one X chromosome leaves sufficient R molecules to repress Sxl.
One currently favored form of this hypothesisz8has the X:A signal produced by two or more
X-linked genes (e.g., sis-a and sis-b in Figure 6) whose products act as "numerator elements,"
and are titrated by certain autosomal products ("denominator elements"), so that a sufficient
concentration of sis products to promote transcription of an active product by Sxl will only be
achieved in females (Figure 6).29.30
Sxl regulates the differentiation of the female tissues, in Drosophila acting entirely through
its control of a locus on chromosome 3, transformer (tra), which is also only functional in
Sex Determination in Insects

Female Determination Male Determination

Mother

product needed 0 .
to activate S?d

autosomal repressor
genes (postulated) m/ lm]
products bind
to repressor molecules

... chromosome
t
non-functional
product

lZq-1
- ........ . . ...
.-...
autosome 3

FIGURE 6. Simplified model of a possible mechanism for genetic control of sex determination in Drosophila
melanogaster. Female determinationand differentiation (left) depends on the product of the key gene Sex lethal (Sxl).
The product of the maternal gene daughterless (da) needs to be present for Sxl to be active, but this is normally
supplied to both male and female eggs. It is thought that unidentified autosomal genes produce a similar concentration
of repressor molecules ("denominator elements": R) in both sexes. Female eggs, with two X chromosomes, produce
twice as many "numerator elements" (i.e., products of X-linked loci such as sis-a and sis-b) as male eggs, so that there
is an excess of unbound molecules to promote the female-determining activity of Sxl. The active product of Sxl
influences the transcription of the product of the autosome 3 gene transformer (tra), which in turn acts on the
doublesex locus (dsx: see text and Figure 7).

females. The tra product collaborates with the product of another gene (tra-2) to control the
expression of another locus on chromosome 3, doublesex (dsx) (Figure 6). The dsx locus is
active in both sexes and provides the double switch mechanism necessary to ensure that
development proceeds only as either one sex or the other; it consists of two cistrons, d s p and
dsd, only one of which functions in each sex. In female eggs (i.e., when both tra and tra-2
are active), dsd is active and its products repress the male sex differentiation genes, whereas
in male eggs the products of dsx" repress female sex differentiation genes.
It is now known that regulation at all the three main stages occurs at the level of RNA
splicing$ that is to say, the primary gene products are the same in both sexes, but they are
"edited" by the splicing out of different sections (introns) of RNA to produce the male- and
female-specific messenger RNAs (Figure 7). The male-specific messenger RNAs of both Sxl
and tra include a stop codon which truncates the open reading frame so that the transcript is
nonfunctional. This explains why mutational loss of these genes has no effect in males.
66 Insect Reproduction

Primary gene products

Female-specific M
splicing 9 Default splicing
A
1121415161718
v
STOP

U
Female-specific
splicing 9 Default splicing

. . .:.
or STOP
tra-2

Female-specific &X
splicing 9 Default splicing

FIGURE 7. Production of sex-specificmessenger RNAs (mRNAs) from the primary gene products of the Sexlethal
(Sxl), rransformer (tra), and doublesex (dsx) genes of Drosophila melanogaster by differential splicing. Primary
transcripts of these genes are shown in the center; the boxes represent coding regions (exons), the horizontal lines
joining them represent introns (which do not form part of an active product), and the female-specificand male-specific
patterns of splicing are depicted, respectively, above and below the structures of the primary transcripts. The mRNAs
generated by this process are depicted to left (female) and right (male) of the primary transcripts. In males the mRNAs
result from the default pattern of splicing. which in the cases of Sxl and tra includes a stop codon rendering the mRNA
nonfunctional. The female-specific product of Sxl regulates its own activity by positive feedback, and regulates tra
activity by promoting the female-specific product of that gene, which in turn plays its part in directing the female-
specific pattern of splicing of the dsx gene. (Adapted from Baker, B. Annu. Rev. Gener.. 17, 345, 1983.)

Thus, sex determination in Drosophila depends on a hierarchical system of regulatory

genes. Can such a mechanism be generally applicable, given the apparently diverse systems
of sex determination found in other organisms? Nothiger and Steinmann-Zwicky " speculated
on various mutations in the regulatory system that could, together with changes in the sex
chromosomes, explain most of the variations observed in insects. They regarded the action of
the double switch gene, dsx, as likely to be basic to the sex determination of all insects, and
therefore not capable of functional mutation. They postulated that the primitive system in
insects was probably represented by species in which the male is heterozygous at a single sex-
determining locus. Using the notation introduced earlier in this chapter, in dominant-Y
systems the sex factor S, acts as the male determiner by repression of the key gene Sxl, whereas
the sex factor S, does not code for functional product, thus allowing Sxl to be active. If
heteromorphic sex chromosomes are involved, then the male sex factor S, (or strictly S,, since
its effect is dominant) would be located on the Y chromosome, but it could be transposed to
different locations in the genome, or copied to different locations to form a "multiple factor"
(M) system. Recessive-X systems, which occur commonly and widely in insects, are expli-
cable in terms of a genic balance as already discussed for Drosophila; the repressor gene is
on an autosome, and Sxl is only activated if two X chromosomes are present, i.e., in individuals
homozygous for S,.
It is possible to interpret other, less common, types of sex determination in terms of
mutations of the key gene Sxl, of its repressor, or of the maternal gene da. Some of these
special cases will be referred to later in this chapter.
Sex Determination in Insects 67

G. DOSAGE COMPENSATION
Organisms in which the Y has little or no homology with the X or does not exist at all (XO),
have a gene dosage problem. An XX female has two copies of every X-linked gene, while an
XY or X 0 male has only one copy. Mammals compensate for this by inactivation of one of
the two X chromosomes in female somatic tissues. However, in Drosophila, where polytene
chromosomes make it easy to study the level of transcriptional activity, dosage compensation
is achieved in a different way. Both X chromosomes are active in female tissues, but the
transcription rate is only half that of the single X chromosome in males, which produces just
as much RNA as the two Xs in females put t~gether.~"' The hyperactivity of X-linked genes
in male Drosophila appears to be due to a set of genes (msl) which are inhibited when the key
gene Sxl is active and are therefore only functional in males8
The only other information on dosage compensation in insects is in Orthoptera. Rao and
Ali32showed that both X chromosomes in hepatic cecal cells of female Acheta domesticus
were euchromatic (i.e., transcriptionally active), and provided some evidence - using an
indirect measure of transcriptional activity of unproven reliability - that the single X
chromosome in the male may be hyperactive, as in Drosophila. On the other hand, females
of the mole cricket Gryllotalpa fossor (=africana?) seem to have only one arm of one X
chromosome transcriptionally active in hepatic cecal cell~,3~,"which resembles the system in
mammals. However, there is evidence that activity or inactivity of the X chromosomes in
Orthoptera may differ among tissues.35
In Lepidoptera, the limited evidence available from the differential activity of sex-linked
loci suggests that members of this order may manage without a dosage compensation mecha-
nism. Indeed, Johnson and Turner36suggested that in mimetic butterflies the dosage differential
may be used to advantage, in order to limit expression of a polymorphism to the female sex.

111. SEX DETERMINATION IN DIFFERENT

GROUPS OF INSECTS
A. APTERYGOTA
Of the four most primitive extant orders of insects, only the Collembola have been studied
sufficiently to warrant generalization, and in these the male is the heterogametic sex and is
normally X0,37J8 but in Neanuridae, species with X 0 and others with XY are known.39
Presumably the XY species are neo-XY, but there is no cytological evidence to confirm this.
In Neanura monticola, with X 0 males, the X chromosome shows considerable polymorphism
with large amounts of heterochromatin (probably repetitive, noncoding DNA) in high altitude
population~.~~ In the Thysanura, Themobia domestica possibly has X,X20males.4O In Protura,
on the other hand, no instances of an XXKO system have been reported; very few of the
species examined had morphologically differentiated sex ~hromosomes.4'~~~ No representa-
tives of the Diplura seem to have been examined cytologically.

B. PRIMITIVE EXOPTERYGOTA
XX/XO sex determination predominates in the Odonata, possibly in the Ephemer~ptera~~
(although these are poorly studied), and certainly in the main Orthopteroid orders (Dictyoptera-
Phasmida-Orthoptera). Where an XXIXY system occurs in these groups, it is usually clear that
it is a neo-XY system, formed by fusion of an X with an autosome (Figure 2), so that the neo-
Y is homologous with a large part of the neo-X. In the anisopteran families of Odonata, for
example (reviewed by Kia~ta,"~")most species are XXIXO, but there are apparent neo-XY
systems in 15 species scattered through the families Gomphidae, Aeschnidae, Cordaliidae, and
Libellulidae, representing about 4% of the dragonflies then studied. Species with neo-XY
generally have, as might be expected, one less autosome pair than related species with an X 0
68 Insect Reproduction

system; e.g., Aeshna crenata has 2n = 28 and X 0 males, whereas A. grandis has 2n = 26 and
neo-XY males.46 Kia~ta,4~ followed by Tyagi,4' explained an evolutionary decrease in the
number of autosomes in the family Gomphidae as a succession of fusions and translocations
between the neo-Y, the neo-X, and autosomes, the outcome of each step being a secondarily
derived X 0 system with one fewer autosome pairs. However, there is no clear cytogenetic
evidence that the sex chromosomes are involved in these changes of karyotype.

C. THE ORTHOPTEROID ORDERS

In the Plecoptera, which are generally thought to be an orthopteroid order that retains
primitive features, several species in different genera have X 0 males, always with a very large
metacentric X chromosome that moves in a highly characteristic way in the first meiotic
division.48 XY males (presumably neo-XY) are only recorded for one species (Perla
(=Paragnetina) immarginata), but some species of Perla have a multiple X chromosome
system apparently derived from XX/XO, males being X,X20, and in Perlodes there are three
species known with X,X,X30 males. The two or three Xs in these species are much smaller
than the single X of X 0 and XY males in related species, and associate together in the first
meiotic division. Their mode of origin is a mystery, as there is no simple way in which a large
metacentric chromosome can give rise to several smaller elements.
The cytogenetics of the other orthopteroid orders has been comprehensively reviewed by
H e ~ i t (Orthoptera)
t~~ and WhiteS0 (Grylloblattodea, Dictyoptera, Isoptera, Phasmida,
Dermaptera, and Embioptera), so information about the sex chromosome systems of these
groups will only be summarized here and up-dated. Dictyoptera (Blattodea + Mantodea),
Phasmida, Orthoptera, and Embyoptera all seem to be primitively XXIXO, whereas in
Isoptera and Dermaptera XX/XY predominates and is possibly the primitive condition.
In Blattodea (cockroaches), males seem to be invariably X 0 where both sexes have been
k a r y ~ t y p e d Whiteso
.~~ suggested that this stability of the sex determination system could be
due to the fact that the X is almost always metacentric, and therefore not so readily available
for centric fusion with an autosome to generate a neo-XY system. Yet it is difficult to see why
this argument does not equally apply to Phasmida, which also have a metacentric X yet
frequently develop a neo-XY system.
Isoptera (termites) are generally thought to have arisen from primitive Blattodea but,
whereas the most primitive cockroach examined cytologically has XX/X0,52 the most primi-
tive extant termites seem to be mainly XX/XY.53Nevertheless, some species do have XX/
XO?4 and fusions and translocations between sex chromosomes and autosomes are so com-
mon in termite^^^"^ that XX/XO could still be the primitive c o n d i t i ~ n . ~ ~
Several species of Kalotermitidae in southern U.S. and the Caribbean form remarkable
chains or rings of up to 19 chromosomes in male meiotic n~clei,~'-~O often involving more than
half the total chromosome complement. The chains are thought to be due to a series of
reciprocal translocations involving both the sex chromosomes and the autosomes of one
chromosome set, these changes being restricted entirely to the male line, so that all the
chromosomes involved function together as a multiple Y chromosome complex (Figure 8).
Females are structurally homozygous and form normal bivalents at meiosis. The genetic
consequences of such an arrangement are quite profound; for example, they restrict many
alleles to males and increase the genetic similarity of offspring to the same-sex parent and to
same-sex siblings. The idea that this unusual system has played a significant part in the
development of eusociality in termites61is, however, somewhat undermined by the fact that
the most extreme rearrangements are found in only a few of the more primitive termites.
Mantodea also seem to have an XX/XO mechanism with a metacentric X chromosome as
the primitive condition, but members of the largest subfamily Mantinae consistently have an
X,X,X2X2:X,X2Ymechanism that has aroused considerable interest among cytogeneticists.
White62proposed that this was derived from XXIXO by translocation between the X and a
metacentric autosome (Figure 4), and all subsequent evidence has been consistent with this
Sex Determination in Insects 69

FIGURE 8. Diagram illustrating how a series of reciprocal translocations, involving one member of each of six
autosome pairs and the Y chromosome (a), could lead to a ring of linked chromosomes in male meiosis of the termite
Incisirermes schwarrzi. The reciprocal interchange set is stippled, and only occurs in males. Chromosome pairs not
involved in translocations (right) form normal bivalents. Sizes of chromosomes are arbitrary; the X and Y chromo-
somes have not actually been distinguished from the autosomes or from each other in this species. (Adapted from
Syren, R. and Luykx, P,,Nature (London), 266, 167, 1987.)

hypothesi~.~~ All Mantinae have a remarkably consistent chromosome complement, with 2n

(male) = 27 (only one exception is known - see below). Presumably the X,X,Y system had
a single origin in the evolution of this subfamily, and some of the species with X,X2Y males
currently placed in other subfamilies are perhaps wrongly classified. However, the African
mantid genus Compsothespis has an X,X2Ysystem with much smaller sex chromosomes, and
2n (male) = 23; at least in this case, an independent origin seems likely (see WhiteS0for further
details). It is not at all apparent why this system has proved so successful for mantids. The
mechanism itself does not seem very efficient; complications often seem to arise in correctly
forming an X,X2Y trivalent in the first division of spermatogenesis, and consequently in
correctly segregating the X, and X, into one daughter spermatocyte and the Y into the other.
~ ~ for Mantis religiosa that the first meiotic division of those
Callan and J a ~ o b sshowed
spermatocytes that fail to form the X,X,Y trivalent is inhibited, thus preventing the formation
of aneuploid sperm. Liebenberg et al.@ reported a single male of Polyspilota aeruginosa
(Mantinae) with 2n = 28 instead of the usual 2n = 27, and an X,X2Y,Y, mechanism. The origin
of the extra Y (neo-Y) in this one aberrant case is unclear.
Whitesolisted 57 species of Phasmida (stick insects) with identified sex chromosomes, of
which 49 are reported to have X 0 males - undoubtedly the primitive condition - and 7
species (in 6 separate genera) have a neo-XY system arising through fusion of an X with an
autosome (see Figure 2). The one other species studied, Didymuria violescens, occurs in
Southeast Australia, where it has at least 10 chromosomal races, occupying contiguous
distribution areas, and including both X 0 and neo-XY forms.6s Several independent origins
of a neo-XY system can be traced from X 0 ancestors.50
In the well-known parthenogenetic stick insect Carausius morosus, males and masculin-
ized females (intersexes or sex mosaics) appear occasionally in laboratory cultures, and their
numbers can be enhanced by various treatments, e.g., subjecting the eggs to high (30°C)
temperature,'j6centrifuging the eggs,'j7X-irradiating egg or o ~ c y t e sor, ~injecting
~ the mother
with pterine derivative^.^^ Females and masculinized females have three metacentric chromo-
somes that are regarded as sex chromosomes because of their behavior in meiosis. Males lack
one of these sex chromosomes or a segment of one of them.'O The method of sex determination
is difficult to work out because the female karyotype is highly aberrant due to its long history
of parthenogenesis. It has been suggested that C. morosus originated as a triploid or tetra-
p10id.~~ However, whatever their origins, both sex chromosome and autosome complements
70 Insect Reproduction

are now aneuploid, and cannot be regarded as comprising any particular number of chromo-
some sets. Male determination presumably occurs because of a change in the genic balance
between factors on the sex chromosomes and on the autosomes (so that, assuming that the
molecular model established for Drosophila applies, the key female-determining gene Sxl is
repressed). It is not clear why intersexes, which retain the female karyotype, arise under
certain conditions; one possibility is that high temperatures, etc. prevent splicing of female-
specific messenger RNAs. General inactivation of the sex chromosomes by
heterochromatinization has been suggested70 to function in sex determination, but such
heterochromatinization has only been observed in germ-line (spermatogonial) interphase
nuclei, and it is not known whether it occurs in embryonic somatic cells.
Only eight species of Embioptera have been studied cytologically (four in each of the
families Oligotomidae and Embiidae), and all have an odd number of chromosomes in male
somatic cells, indicating that sex determination is probably XX/XO, with the X chromosomes
large and m e t a c e n t r i ~Nothing
.~~ is known about sex determination in Zoraptera.
H e ~ i tcomprehensively
t~~ reviewed the extensive cytogenetic studies that have been car-
ried out on the Orthoptera proper (Saltatoria). Since Hewitt's review, there have been signifi-
cant contributions on the sex chromosome systems of neotropical Acridoidea (about 200
species72),the acridoid subfamilies Catant~pinae,'~and Pam~haginea,~~ Indian Orthoptera (30
species7s), and certain Tettigon~idea?~-~~ XX/XO sex determination is found in the great
majority of species in all subdivisions of the order, both primitive and advanced, and is
undoubtedly the primitive condition for the Orthoptera as a whole. The only exception is the
relic group Grylloblattodea, with XY males in the only two species studied,'O but in the face
of all the other evidence, this must be regarded as a derived state. About 8% of species have
X Z X Y or XlXlX2X,/XlX2Ysystems, which occur in every major subdivision of the group
and are usually clearly evolved secondarily from an XX/XO condition by centric fusion
(Figures 1,2). Two cases are known, one in Eumastacoidea ("Morabinae species P45b")79and
the other in Tettigonoidea (Callicrania ~ e o a n e i of
) ~ the
~ neo-X being formed by "tandem
fusion" of an autosome to the centromeric end of the original X. In both these cases, the neo-
Y forms a terminal connection with the neo-X at meiosis, and this neo-XY bivalent divides
equationally at first meiotic division, so that the X and Y do not segregate until the second
division ("postreductional meiosis"). The mantid type of origin of an XlX2Ysystem, directly
from XX/XO by translocation between an X and an autosome, is not known to occur in
Saltatoria, perhaps because autosomes in this order are predominantly acrocentric? making
centric fusions a more likely occurrence.
The neo-X produced by centric fusion between an acrocentric X and an acrocentric
autosome is likely to be large and metacentric, and the neo-Y (the original autosome) is
acrocentric (see Figure 1); the majority of cases of neo-XY systems in Saltatoria have sex
chromosomes of this form (see Table 8 in H e ~ i t t ~Likewise,
~). neo-X,X,Y males produced as
a result of a Y-autosome fusion have a metacentric X, and Y and an acrocentric X, (Figure
2); again, the majority of X,X,Y systems in Saltatoria conform to this pattern. This may reflect
the recent origin of many of these systems because, once a neo-Y is formed, it is subject to
very different evolutionary pressures from the original autosome. Several species have been
studied that have both X 0 and neo-XY p o p ~ l a t i o n s ; ~presumably
~ , ~ ~ , ~ ~ the
- ~ ~neo-XY system
is only very recently established in such populations, and in some cases the early stages of
differentiation of the neo-Y from its homologue, now part of the neo-X, can be observed. The
neo-Y may acquire heterochromatic segments, and pairing between the neo-X and neo-Y may
become restricted to terminal regions, so that crossing-over is limited, paving the way for
further differentiation of the genetic role of the neo-Y from that of its former homologue.
In time, as discussed earlier, the neo-Y is likely to degenerate; an example of this may be
the Gryllacridoid genus Dolichopoda, where the "neoW-XYsystem is possibly as old as the
genus itself, and all species studied have a large metacentric X and a small dot-like Y.83
However, no instance has yet been identified in Orthoptera of the complete loss of a neo-Y,
Sex Determination in Insects 71

to revert to an X 0 system, which suggests that the neo-Y may acquire and retain some
functional male-linked loci.49
The earwigs (Dermaptera) seem to stand somewhat apart from the other orthopteroid
orders, and this is reflected in their chromosomes, which have diffuse centromeric activity like
those of Hemiptera, and in their sex determination system, as the primitive condition for the
group seems to be XXIXY rather than XX/XO. Only two species with X 0 males are recorded,
belonging to different families.84Multiple sex chromosomes are very common, occurring in
about half the species that have been karyotyped, with similar frequency of incidence in all
families. Multiple Xs have probably arisen by simple dissociation of the existing X chromo-
somes, as in other insects with holocentric chromosomes (Figure 5). They form a close cluster
on the spindle at first meiotic division, and all move together to one pole, while the X moves
to the opposite pole. The ubiquitous earwig Foficula auricularia is unusual in having two
alternative Y chromosomes, one of which ("Y,") is mitotically unstable so that it tends to
accumulate in number, and individual males may have up to four copies (XY,Y2Y,Y2).
Mosaic males have been recorded with different numbers of Y chromosomes in the cells of
each testisg4

D. THE HEMIPTEROID ORDERS

The Psocoptera are generally regarded as close to the basal hemipteroid stock, and all the
32 species so far examined cytologicallysSseem to have XX/XO sex determination. Nothing
is known about the sex-determining mechanisms of biting and sucking lice (Mallophaga and
Siphunculata), as no sex chromosomes have been identified in any of them. When a female
human louse (Pediculus humanus) is mated with a single male, the sex ratio of the progeny
is strongly biased toward one or other sex, and unisexual broods are common.86Contrary to
White? no information is available about the progeny of females mated more than once, and
it seems likely on the available evidence that maternal factors are involved in the determina-
tion of sex in lice, as in certain Diptera (e.g., Chrysomya).
Most species of Heteroptera have XX/XY sex determinati~n,~' but there are some groups
- e.g., 124 species in the related families Coreidae and Alydidae - that are almost exclu-
sively XX/XO.ggX 0 males also predominate in the supposedly more primitive Heteroptera
(Gerromorpha; but see Calabrese and Tallericog9),and Ueshimas7 concluded that the XY
system in Heteroptera, despite its widespread occurrence, is derived from a primitive X 0
condition. Nokkala and N ~ k k a l aon , ~the other hand, argued that XY was ancestral. Clearly,
XY systems are ancient and well-established in terrestrial Heteroptera; the X and Y chromo-
somes generally show little or no evidence of the homology expected of a neo-XY system and
undergo a characteristic pattern of meiotic behavior in which they usually segregate at the
second division (for details, see Whiteg(pp. 620-62 1) or Ueshimag7).The scattered occurrence
of X 0 species within genera must surely be due to secondary loss of the Y chromosome, and
such loss may have occurred early in the evolution of many families of terrestrial Heteroptera.
However, this does not rule out the possibility that the common ancestor of all Heteroptera was
XO, as in Psocoptera, and that XX/XO sex determination may be the primitive condition in
some families of Gerromorpha. The problem can only be resolved when the cytology of
members of the most primitive groups, Enicocephalomorpha and Dipsocoromorpha, now
thought to have a sister-group relationship with all other Heter~ptera,~' as well as of the relic
family Peloridiidae (suborder Coleorrhyncha), have been examined. The only information for
these groups so far is for one species of Dipsocoromorpha, males of which were tentatively
recorded as X0.92Multiple sex chromosomes are common in Heteroptera, and may be derived
from either XY or X 0 systems. Apparently they are in most cases due to dissociation of the
X chromosome into two or more smaller parts, which group together on the spindle of the
second meiotic division and move en bloc to one pole (see Figure 5). In some species, the
number of X chromosomes varies; the best-known example is the bedbug Cimex lectularius,
where the number of separate X elements varies from 2 to 15.
72 Insect Reproduction

Messthaler and T r a ~showed

t ~ ~ that the Y chromosome was heterochromatic and therefore
transcriptionally inactive in all stages of spermatogenesis of the milkweed bug, Oncopeltus
~ can be little doubt that the Y chromo-
fasciatus. Despite the reservations of T h o m a ~ ?there
some in Heteroptera is genetically inert, and that sex determination is based, as in most insects,
on a "recessive-X" (i.e., genic balance) system. Otherwise it would be impossible to explain
how the secondary loss of the Y chromosome could occur in so many groups without
concomitant loss of genetic viability.
Sex determination in Homoptera-Auchenorrhyncha is predominantly XX/X0.95,96A few
species with XY males occur within genera and subfamilies that are otherwise exclusively
XO, and in such cases it is often clear that the Y is a neo-Y; i.e., the homologue of an autosome
that has recently fused with the X. Such a neo-Y pairs with the autosomal part of the neo-X
in meiosis and segregates from it at the first divi~ion.~' In several species of Oncopsis, both
X 0 and neo-XY males occur in the same or different populations; the XY state results from
fusion of the X chromosome with a different autosome in each species.98
The Homoptera-Sternorrhyncha include some of the most specialized hemipteroid fami-
lies, and the basic system of sex determination is often obscured, especially in groups with
well-developed parthenogenesis. The Psylloidea are the least reproductively specialized, and
here again sex determination is predominantly XXKO. Only three species with XY males
have been found in a total of 39 species examined, all apparently recently derived from X 0
by X chromosome-autosomefu~ion.~~-lO~ The Aleyrodoidea have received very little attention
from cytogeneticists. On the basis of early cytological work on three species,lo2J03and the
observation that males are only produced in laboratory populations by unmated females, the
general presumption is that all male aleyrodids are haploid? It would be preferable to have this
confirmed for more species before assuming that haplodiploidy is of general occurrence in this
group. The factors invoking male determination are unclear, but the cytological mechanism
in those species studied seems to be the same as in Hymenoptera, with meiosis replaced by
a single mitotic division, each primary spermatocyte giving rise to only two spermatids.
Populations of Trialeurodes vaporariorum seem to have an approximately 1:l sex ratio in
field populations in both Europe and North America,Io4which is unusual for a haplodiploid
system. However, thelytoky is a complicating factor in interpreting sex ratios in this species.
The populations originally introduced from North America to England consisted almost
exclusively of thelytokous females,Io3and it is not known what proportion (if any) of females
reproduce thelytokously in present-day populations. In Bemisia tabaci, which has not been
studied cytologically, thelytoky is unknown and the number of males produced seems to be
temperature dependent.los
It is impossible to do justice here to the remarkable sex determination systems of scale
insects (Coccoidea), and for details the reader is referred to the authoritative reviews by
NU^.^^^.'^^ All the different systems are believed to have evolved from an ancestral XXIXO
system which is still found in some members of the more primitive families (Ortheziidae,
Margarodidae, Phenaeolaechiidae). Some of the margarodids (Icerya and four closely related
genera) have evolved male haploidy, and in some species of Icerya there is the further
development of hermaphroditism, with morphologically female individuals maturing haploid
sperm and diploid ova in an ovotestis. In hermaphrodite Ice~ya,fertilization is usually between
eggs and sperm of the same individual; nevertheless, some eggs apparently remain unfertilized
and give rise to functional haploid males. In all the more advanced families of Coccoidea, the
paternal set of chromosomes is rendered inactive in most tissues by heterochromatinization
during the development of male embryos (the "lecanoid" and "Comstockiella" systems; see
NurIo6). It seems that sex in these families is determined maternally rather than by the
genotype of the zygote, because the sex ratio is greatly affected by the age of the female at
mating and by environmental conditions such as temperature.'07 It is not clear, however,
whether the inactivation occurs after, and as a consequence of, the embryo already having
Sex Determination in Insects 73

been determined as male, as in Sciaridae (see Diptera, below), or whether the inactivation
process itself provides the mechanism for male determination.Io7
Bulllo pointed out that the evolution of these advanced coccoid systems from XXIXO is
something of a mystery, because the heterochromatinized paternal chromosomes are elimi-
nated in spermatogenesis, so that all sperm carry only the maternal genome. There is thus no
genetic polymorphism among sperm to serve as a basis for sex determination, which effec-
tively means that the advanced coccoid systems can never have coexisted with a system such
as XXIXO, and must therefore have evolved through a form of sex determination without male
heterogamety. There are a few coccid species (e.g., Lachnodius eucalypti) without identifiable
sex chromosomes, and which do not undergo heterochromatinization of one chromosome set
in the male (2N-2N of NurIM).Nur thought that these were probably derivatives from forms
with heterochromatinization,but Bull's argument makes it more likely that they are represen-
tative of this intermediate stage, evolved from XXIXO prior to the origin of
heterochromatinization, which is in line with the original views of Brown.Io8 Haig has
developed a model for the evolution of the advanced coccoid systems based on sex ratio
theory .log
Aphids (Aphididae) all have XX/XO sex determination. An XXKY system would be an
impossibility for these cyclically parthenogenetic insects, because most species exist through
the summer as all-female, thelytokous populations, and during this period the Y chromosome
would have "nowhere to go". Aphids produce males parthenogenetically. To develop as X 0
males, oocytes have to lose half the sex chromatin of the parent female. This is achieved in
a single egg maturation division, as in the thelytokous production of females, but the X
chromosomes pair during prophase1'Oand then undergo a sort of "mini-meiosis" on their own,
first separating the products of pairing and then dividing equationally with the autosomes, all
on the spindle of the single maturation d i v i ~ i o n . ~ l This
l . ' ~ ~peculiar cytological mechanism for
male determination is of special interest because it is normally triggered by environmental
conditions and mediated by a low level of juvenile hormone in the haemolymph; males can
be induced by treatment with precocene, which destroys the corpus allatum, and inhibited by
the juvenile hormone analogue kin~prene."~ The environmental factors are normally photo-
period (actually the length of the dark phase) and temperature in Aphidinae, but may be
nutritional in other groups, and in some species, males appear spontaneously or after a
genetically programmed number of thelytokous generations.Il4
The spermatogenesis of aphids is also relevant to their sex determination, because the
fertilized eggs must all develop as thelytokous females, so all the sperm from X 0 males must
carry an X chromosome. This is achieved by a peculiar first meiotic division in which the X
is stretched on the spindle before passing into one of the daughter spermatocyte nuclei, after
which the daughter nucleus without an X degenerates.Il5
Multiple X chromosome systems occur in some aphids, apparently as a result of dissocia-
tion of the original X, and the separate elements all behave in the same way in m e i o ~ i s . l ' ~ * ' ~ ~
The greenideine species Schoutedenia ralumensis (=lutes) has what was presumably origi-
nally an X,X,X2X2/X,X,0 system, but it has become modified in a remarkable way by
consistent association or temporary fusion of one member of an autosome pair with X, and
the other member of the same pair with X,.lk8Male determination necessarily retains both
these autosomal homologues (AA), so males receive the two elements X, + A and X, +A. One
of the X chromosomes (it is not clear whether it is X, or X,) then has to lose its connection
with the autosome at anaphase I of spermatogenesis, so that males can transmit one X + A and
one X to the next generation (Figure 9). How this peculiar system evolved as a stable
mechanism for sex determination is something of a mystery.
Multiple X chromosome systems with X,X2 males also occur in the primitive aphidoid
families Phylloxeridae and Adelgidae (see BlackmanH9for review), and show some unusual
features in the few species studied. In particular, there seem to be species in each group which
have evolved a potential for male-linked inheritance "by proxy," which overcomes the total
 Insect Reproduction

1 Spermatocyte ll nucleus lacking

0 xl and xz degenerates

FIGURE 9. Sex chromosome-autosome associations in the aphid Schoutedenia ralumensis (=S.lurea). For simpiic-
ity only, the X chromosomes and the pair of autosomes (AA) associated with them are shown. Female somatic cells
(a) have four long chromosomes of unequal length, representing X,, (X, + A), X, and (X, + A). Male somatic cells
and spermatogonia (h) have the longest two chromosomes, which are (X, + A) and (X, + A). In spermatogenesis, at
prophase of the first meiotic division, the autosomes attached to X, and X,, being homologous, pair in parallel (c,d).
When the cell divides (anaphase), either X, or X, loses its connection with the autosome (e; shown here as X,, but
it is uncertain which). The lost autosome passes into one daughter cell which lacks both X chromosomes and
degenerates. The other daughter cell has both X chromosomes, with one autosome still attached to one of them (0;
it divides equationally to give spermatids with the same chromosome constitution. Presumably, for the system to be
stabilized, oogenesis must somehow result in oocytes with the complementary arrangement; i.e., if spenn have (X,+
A) and X,, oocytes will have X, and (X, + A). It is not known how this is achieved. (Based on Hales, D. Chromosoma
98, 295, 1989.)

absence of males during the parthenogenetic (thelytokous)part of the life cycle, by having two
cytologically distinct types of all-female line; one leading eventually to male production and
the other to sexual females. In Phylloxera caryaecaulis, studied by the pioneer cytogeneticist
T. H. Morgan,120one member of the smaller "pair" of X chromosomes seems to be limited to
the male-producing line, and behaves differently from the other in its pairing relationships
during sex determination and spermatogenesis (Figure 10). In the adelgid Gilletteella (=Adelges)
cooleyi, Steffan12' found one member of the longer pair of X chromosomes dissociated into
two parts in about 50% of thelytokous females, and in the somatic cells of males, but not in
sexual females. Further work is needed on these groups to confirm and extend these findings.
The last hemipteroid order to be considered is the Thysanoptera (thrips), both suborders of
which (Terebrantia, Tubulifera), on the basis of the few species that have been studied
cytologically, have haploid males.122.123
The cytological mechanism involved is not very clear,
but seems to differ from that of Aleyrodoidea and Hymenoptera, and must be independently
derived. Instead of meiosis being replaced by a single mitotic division, as in other insects with
haploid males, two meiotic divisions are retained; the first is apparently equational, giving rise
to two similar-sized spermatocytes, but the second produces one large functional spermatid
Sex Determination in Insects

Female-Producing Male-Producing
Line Line

Polar plate of

+ +
Stem mother's egg

#- Somatic metaphase of

R.#
parthenogenetic generation Ot3
+
Polar plate of Polar plate of
6 egg
J

Anaphase
9 egg + +
Somatic metaphase
of 6 e-
Somatic metaphase O O Q

of sexual 9

Anaphase I
sexual
egg

and oocyte 9 line 6 line

nucleus

FIGURE 10. Chromosome cycle of Phylloxera caryaecaulis, redrawn from M~rgan.~~O Autosomes are shown
black, X chromosomes white, except for one member of the smaller pair (X,) in the male line, which is stippled to
show its differential behavior and possible role in sex determination. In the line leading to production of sexual
females (left), small and large X chromosomes seem to be consistently associated, in the somatic cells of both
parthenogenetic and sexual females and throughout oogenesis. In the line leading to male production (right), the small
and large X chromosomes are likewise associated throughout the parthenogenetic phase, but during maturation of
eggs destined to become male, the Xs exchange partners, so that the two large X chromosomes form one pair, and
the small Xs another. Consequently, at maturation division of male eggs, the small and large X chromosomes
segregate from each other independently. Males have X,X,O; half of them apparently have X, and X, associated
together as in females, and half have them separate. Sperm with separate X, and X, are believed to give rise to the
parthenogenetic line that will produce the males of the next bisexual generation.

and one much smaller one that rapidly degenerates.lZ2As in the Aleyrodoidea, the factors
invoking male determination are unclear; sex ratios show considerable variation within and
between species,lZ4but the interpretation of these in genetic terms is complicated by the
occurrence of thelytokous parthenogenesis in many of the best-studied species.Iz5 In
Elaphrothrips tuberculatus, females have unisexual broods, the males being produced vivipa-
rously and the females oviparously; more males seem to be produced when the offspring are
larger and fitter in the favorable nutritional conditions of spring.lZ6

E. NEUROPTEROIDEA AND COLEOPTERA

The Neuropteroidea (Megaloptera, Raphidioptera, and Plannipennia) are generally re-
garded as an early branch in the phylogeny of the endopterygote insects, but no species have
 Insect Reproduction

FIGURE 11. Diagrammatic drawings of first meiotic metaphase of male of (a) the neuropteran Macroneurus
appendiculatus, showing "distance pairing" of X and Y chromosomes, and (b) the megalopteran Neohermes
filicornis, showing X and Y forming a bivalent like the "parachute bivalent" (Xy,) of Coleoptera. Structure of the
parachute bivalent is shown in (c). (Based on Hughes-Schrader. S. Chromosoma. 81, 307, 1980.)

been found with XXIXO. Almost all species studied seem to have XXIXY sex determination,
with a few showing multiple X systems. The X and Y chromosomes of Raphidioptera and
Plannipennia (=Newoptera sensu stricto) behave in a very consistent fashion during spermato-
genesis (Figure l la). They are both small chromosomes that apparently lack any homology,
because they never pair to form a bivalent in the first meiotic division, and regularly take up
positions in opposite halves of the spindle before segregating into the daughter spermato-
c y t e s . 9 ~In~ the
~ ~ two species of Megaloptera that have been studied, however, the X and Y
chromosomes form a bivalent that positions itself with the autosomes on the equator of the
spindle and segregates synchronously with them at the first meiotic division.128The Y
chromosome is much smaller than the X, and in one species the bivalent looks very like the
"parachute" bivalent (Xy,) found in Coleoptera (Figure l lb,c, and see below).
Thus, the sex chromosome systems of the Neuropteroidea seem to provide useful phylo-
genetic evidence pointing to a sister-group relationship between Raphidioidea and Plannipennia,
and also supporting the often-held view (e.g., Henning129)that the Megaloptera are the sister
group to the Coleoptera.
The Coleoptera show a great diversity of sex chromosome systems, although the underly-
ing genetics of sex determination may well be far less variable, and is likely to be based on
a recessive-X mechanism, except where male haploidy has evolved. Coleopteran cytogeneti-
cists have accumulated information about the sex chromosome systems of over 2500 species.
Fortunately, the comprehensive reviews by Smith and VirkkiI3O and Virkkil3I mean that only
a brief overview and some updating are necessary here.
The peculiar symbols used in the literature on beetle sex chromosomes are somewhat
daunting to the nonspecialist, but can be simply explained. They symbolize the appearance
and behavior of the sex chromosomes in the first meiotic division of the male beetle. Sex
chromosome symbols are written together if there is any sort of pairing between them to form
a bivalent (e.g., XY), but separated by a plus sign (e.g, X+Y) in the much rarer cases where
they do not pair. The Y chromosome is usually very small in Coleoptera, and this is indicated
by writing Xy instead of XY. In most Polyphaga with Xy, the minute Y is attached by both
its arms to the larger X, so that it resembles a parachutist suspended below the "canopy"
formed by the X (Figure 1lc). The formation and structure of the parachute has recently been
studied by silver staining;132it is believed to have a role in assisting the regular segregation
of the X and Y at first meiotic division. When the Xy bivalent takes this form, then a subscript
Sex Determination in Insects 77

"p" (for parachute) is added: Xy,. XX/XO systems in Coleoptera are represented by a single
X, rather than as XO. Systems with multiple small Y (=y) chromosomes involved in a single
parachute are written Xyy,, Xyyy,, etc.
Two main types of neo-XY system occur in beetles; those with a large Y, probably derived
from an X 0 system by X-autosome fusion (e.g., Figure 2), and those where an autosome has
apparently undergone a reciprocal translocation with either the X, or the y, of an Xy, system
to give an "X,neoX-neoY,", or some other complex system in which the original parachute
has elements (neo-X, neo-Y) associated with it in the first division of m e i o ~ i s . ' ~ ' . ' ~ ~
The more primitive beetles (Adphaga) differ from the Polyphaga in that Xy, systems are
virtually absent except in a few Dytiscidae (records of Carabidae with Xy, are apparently
q~estionablel~~). XXIXO is most frequent in Adephaga, occurring in about 53% of species,
with 29% having XX/XY (or XX/XY).'~~ XY systems predominate, however, in the carabid
genus Bembidion (176 out of the 205 species examined135).Tiger beetles (Cicindelidae)
mostly seem to have multiple X systems, with 2, 3, or 4 X chrom~somes.l~~
More than half of over 2000 species of Polyphaga examined cytologically have Xy, sex
chromosome systems, which are well represented in every major family, and are generally
thought to be the ancestral condition for the whole suborder. Whether Xy, is the primitive
condition for all beetles is not quite so clear, because of its rarity in Adephaga, although the
recent finding of a sex parachute in a megalopteran makes it more likely. Only single species
have been examined in each of the two primitive beetle suborders Archostemmata and
Myxophaga, and they may both be ~ntypical.'~~,'~'
Since Virkki's 1984 review,13' there have been significant studies on the sex chromosome
systems of C h r y s ~ m e l i d a e , ' ~
Histeridae,laIndian
~J~~ Staphylinidae,I4' Indian C u r c u l i ~ n i d a e , ~ ~ ~
32 other Indian beetle species,143Tenebri~nidae,'~~ ~ ~ 50 Russian beetle
B r ~ c h i d a e , 'and
species.146
Apparently the related order Strepsiptera is still cytologically unknown.

F. HYMENOPTERA
All the Hymenoptera except the few species that are thelytokous have haploid males,
produced from unfertilized eggs. The origin of haplodiploidy in this group presumably
therefore dates back to its inception in the early Mesozoic or late Palaeozoic. There seems little
doubt that this form of sex determination has been the key factor enabling the development
of eusociality in the higher groups of the order.I4'
Possible genetic mechanisms underlying haplodiploid sex determination have already been
outlined. At least two different models are necessary to fit the observed facts, one involving
multiple alleles at a single locus, and the other involving multiple l o ~ i . ~ The
~ . single-locus
~~~.'~~
mechanism (see p. 64) results in up to 50% of the fertilized eggs in inbred populations
developing as diploid males, which generally have low viability and fertility.21J50Diploid
males have been reported from several species of Tenthredinoidea, Ichneumonoidea, Apoidea,
and Formic~idea.'~~ Not all the reported instances can be attributed to inbreeding, but the
single-locus model seems to be established for one or more species in each of the above-cited
subfamilies, suggesting that it is ancestral to the Hymenoptera as a whole; e.g., the sawfly
(Neodiprion nigroscutum, the braconid Habrobracon hebetor, the ichneumonid Diadromus
p~lchellus,~5' the honeybee Apis mellifera, and the fire ant Solenopsis i n v i ~ t a . ' ~ ~
If single-locus sex determination occurs generally in the ichneumonid and braconid para-
sitoids reared and released as biological control agents, they may suffer from reduced viability
if inbred populations are used, because of diploid male produ~tion.'~~ In several species of
Chalcidoidea, however, in which sibmating is common in nature, inbreeding has not led to the
male-biased sex ratios that would be expected if diploid males were being produced, and a
multiple-locus model seems to be necessary. A single-locus scheme also does not seem to
explain sex determination in six species of meliponine bees, which did not produce diploid
males when ~ i b m a t e d , although
'~~ diploid males were later obtained in another meliponine
78 Insect Reproduction

species.ls4 Neither single-locus nor multiple-locus models seem applicable to the bethyloid
Goniozus nephantidis, which typically has within-brood mating and hence marked inbreed-
ing.155Generalizations would be unwise in the present state of knowledge, but it seems that
single-locus sex determination is likely to occur in Hymenoptera that generally practice
outbreeding, or in the higher social groups where the production of diploid males can be
controlled; for example, diploid male honeybee larvae are eaten by workers about 72 h after
eclosi~n.~~~
There have been estimates of the number of sex-determining alleles for several species,
either by crossing different lines (9 alleles, in H. hebetor), or by a statistical calculation based
on the incidence of diploid males in natural populations (99-19 in A. mellifera, depending on
population size;Is720 in Melipona compressipes f a ~ c i c u l a t a ;15
~ ~in~ S. invictalS2).
The pteromalid (chalcidoid) wasp Nasonia vitripennis has on occasions produced fully
fertile diploid males in laboratory cultures, but will not do so in response to intensive
inbreeding.158It is difficult to explain sex determination in this species, even as a multiple-
locus me~hanism.'~ Nasonia has been studied particularly with regard to the ability of the
female wasp to manipulate sex ratios by controlling sperm access to eggs, and in the course
of those studies several apparently extrachromosomal factors were discovered that influence
sex. One of these is of particular interest because it is transmitted paternally, but then
inactivates the paternal chromosome set by heterochromatinization in the fertilized egg, so
that genomically haploid, all-male broods are 0btai11ed.I~~ It thus mimics the normal mode of
sex determination of some scale insects and of sciarid flies. The transmitting agent has now
been identified as an accessory (B) chromosome, termed the paternal sex ratio or PSR
c h r o m o ~ o m e .In
~ ~effect,
~ . ~ ~the
~ PSR chromosome "jumps" from one haploid set to another
at the expense of the chromosomes with which it is associated and is thus an extreme example
of "selfish" DNA.
Most gall-forming Cynipoidea have two generations per year, one thelytokous and the
other bisexual. For several common species it has been shown that females of the unisexual,
thelytokous generations differ in the eggs that they lay, producing either only haploid (male)
eggs or only diploid (female) eggs.162The females of the bisexual generation are also of two
types, one giving rise only to the male producers of the next unisexual generation, the other
only to the female producers. Thus each female of the bisexual generation has grandchildren
of only one sex. Possible underlying genetic mechanisms were discussed by C r o ~ i e r . ~ ~

G. THE PANORPOID ORDERS

It is generally agreed that the remaining insect orders -including Lepidoptera and Diptera
- form a monophyletic group, with the Mecoptera close to its main stem. It is therefore of
interest that, whereas only a minority of species in the higher panorpoid orders have an
XX/XO system, X 0 males are found in all species of Mecoptera so far examined except one
(which has a clearly derived multiple sex chromosome system, with X,X,Y males)? It seems
likely, therefore, that the variety of sex chromosome systems found in the remaining orders
were all derived from an XXIXO system.
Very little is known about sex determination in the highly specialized order Siphonaptera
(fleas), which is placed somewhat uncertainly in the panorpoid complex. Of the four species
examined, two were probably XXIXY, and two apparently had multiple sex c h r o m o s ~ m e s ; ~ J ~ ~
males of one of these latter were X,X,Y, and of the other possibly XlYlX,Y2.
At some stage in one of the two main branches of panorpoid evolution - that leading to
the Trichoptera and Lepidoptera - the XX/XO system was replaced by a system involving
female heterogamety (XYIXX or ZWEZ). S ~ o m a l a i n e ndiscussed
'~~ the similarity between
trichopteran and lepidopteran sex chromosome systems. Both Trichoptera and Lepidoptera
have numerous small holocentric chromosomes, with sex chromosomes that are hardly
distinguishable in either mitotic or meiotic cell divisions, so much of the early work on sex
determination in Lymantria dispaP and Bombyx m ~ r i was l ~ ~done without any cytological
Sex Determination in Insects 79

information. R o b i n ~ o n ' s list

' ~ ~of chromosome numbers of over 1000 species of Lepidoptera
has no information on sex chromosomes. However, Smith's167discovery that all or part of the
unpaired Y (or W) chromosome in females is heterochromatic in interphase nuclei - and
hence that nuclei of female Lepidoptera with XY sex chromosome constitution contain a
dense "sex chromatin body" that is not found in male cells - provided a simple method for
determining the sex chromosome system. Traut and M o ~ b a c h e rand ' ~ ~E n n i ~between
l ~ ~ them
found a sex chromatin body in cell nuclei of females of 151 out of a total of 185 species
studied. Where females of a species had no sex chromatin body, female somatic or oogonial
cells of that species generally had an odd number of chromosomes, one less than in the male,
indicating an XO/XX (or ZO/ZZ) sex chromosome system. X 0 females are more common in
some families (e.g., Lasiocampidae, Yponomeutidae, Noctuidae) than others. Genera in
several families contain both XY and X 0 species.169A very few species have one or two
heterochromatic bodies in male as well as female nuclei, presumably due to presence of
constitutive heterochromatin on other chromosomes apart from the Y.
Multiple sex chromosomes (females with XY,Y,) have been found in two species of
Pyraloidea and two of Tortri~oidea.'~~ The two Y chromosomes, probably arising by simple
dissociation, both associate with the X in the first meiotic division of oocytes to form a
trivalent. In female somatic cell nuclei of three of the four species, it was possible to observe
two sex chromatin bodies, corresponding to Y, and Y,.
Some species of Solenobia (Psychidae) have thelytokous races, and a special mechanism
is necessary to ensure that progeny are all heterogametic like their mother. In S. triquetrella,
which has both diploid and tetrapioid thelytokous races, the nucleus that develops as an
embryo is derived by fusion of two of the four nuclei resulting from meiosis; as this fusion
is always between nonsister nuclei, the sex chromosome heterozygosity is preserved.171
Even though very few species have been studied in any detail, it is clear that the genetic
mechanisms involved in sex determination in Lepidoptera must be almost as variable as they
are in Diptera (see below). In the silkworm B. mori, it has long been known that the Y
chromosome of the female carries a dominant female-determining gene.16sDiploid, triploid
and tetraploid individuals are female as long as there is at least one Y chromosome, even when
the X:Y ratio is 3:l. Intersexes do not occur. Thus any male-determining factors on the X or
on autosomes must be very weak in their effect. On the other hand, there is abundant evidence
from Goldschmidt's work that sex determination in L. dispar depends on a delicate and
evolutionarily labile balance between the strengths of a male-determining factor or factors on
the X chromosomes and one or more female-determining factors, probably on the Y (reviewed
by White;9 but see also Clarke and Ford17,).
In several lepidopteran families, including some La~iocampidael~~ and S a t ~ r n i d a e ,in~ ~ ~ . ~ ~ ~
the same superfamily as Bombyx, the occurrence of XO/XX sex chromosome systems seems
to rule out any mechanism based on a dominant Y-borne sex factor and, where closely-related
X 0 and XY species occur, it seems likely that the Y has little or no role in sex determination.
In Ephestia, at least half the Y chromosome can be lost without any effect on sex determina-
tion174(although the female-determining factor could still be located on the remaining half).
It seems possible, therefore, that the vital and dominant role of the silkworm Y chromosome
is somewhat unusual.
The other main branch of the panorpoid complex - that leading to the higher Diptera -
also seems to have undergone major changes in the mechanism of sex determination early in
its evolution. The most primitive group of Diptera-Nematocera, the Tipuloidea, includes
species with XY males, such as Pales (=Nephrotoma) ferruginea, in which the X and Y
behave in an almost identical fashion to those of Neuroptera-Plannipennia; i.e., they do not
form a bivalent in the first division of meiosis, and take up positions in opposite halves of the
spindle ("distance pairing"), before segregating to the p01es.l~~ Even in Tipuloidea, however,
there is evidence of a change in the sex determination mechanism which forms the basis for
the variety of systems in the Diptera as a whole. In P.fermginea, XXY individuals are male,176
80 Insect Reproduction

indicating that sex determination is based on a dominant-Y mechanism, which is unlike all the
other insect groups covered so far.
The X and Y in most tipulids are very small, and in Tipula caesia and T. pruinosa they have
"disappeared; it seems probable that the sex chromosomes in these two species, or at least
the Y chromosome, have fused with members of a pair of autosomes, so that one of the three
pairs of autosomes now bears the sex-determining locus? A similar change seems to have
occurred in the two species of the tipulid subfamily Limnobidae (=Limoniinae) studied by
Wolf:177Dicranomiya (=Limonia)tnnotata and Thaumastoptera calceata. A species of Liriope
in the family Ptychopteridae, which is placed phylogenetically somewhere between Tipulidae
and Psychodidae, provides support for this idea;178it has heteromorphic sex chromosomes (X
and Y), but they have "acquired" homologous regions so that they pair to form a bivalent in
meiosis, suggesting that they are a neo-X and neo-Y formed by translocation or fusion with
a pair of autosomes. In this case, the size of the original Y, or the size differential between the
original X and Y, was presumably large enough to ensure that the neo-X and neo-Y are
recognizably heteromorphic.
In the Psychodoidea, which appear to be a branch of the dipteran phylogeny arising
between the Tipuloidea and Culicoidea, only a few species of sand flies (Psychodidae) have
been studied c y t ~ l o g i c a l l y , but
~ ~ ~these
- ~ ~provide
~ a similar picture to the Tipuloidea. Most
species have 2n = 6 or 2n = 8 without recognizable sex chromosomes, but one (Phlebotomus
pemiciosus) had 2n = 10 including a small heteromorphic pair of sex chromosomes~79-
presumably the more primitive condition.
In Culicoidea, where many more species have been studied cytogenetically, only the
Chaoboridae and the culicid subfamily Anophelinae have heteromorphic sex chromosome^.^^^.'^^
The Chaoboridae and one anopheline species (Chagasia bathana) have 2n = 8, with acrocen-
tric X and Y, whereas other Anophelinae and all other Culicidae studied have 2n = 6, perhaps
as a result of sex chromosome-autosome fusion. X and Y chromosomes in most mosquito
species can, however, be distinguished by their different patterns of staining with Giemsa or
quinacrine ("G, C, or Q bands"; e.g., Newton et al.Ix4),and slight intraspecific or interspecific
differences size can often be attributed to different-sized blocks of constitutive heterochroma-
tin (repetitive, noncoding DNA) (e.g., Mezzanotte et aI.lg5).Anopheles X and Y chromosomes
have extensive heterochromatic regions.lX3
The dominant, male-determining locus (M) of Aedes aegypti is on one member of the
shortest chromosome pair near the centromere,Ix6and has been similarly located in several
species of Culex,Ix3but in one strain of C. tritaeniorhynchus in Japan, "M" is on one of the
longest chromosome pair.Ig7Thus, the sex determinant can alter its position in the genome, a
phenomenon which comes into its own in the next superfamily, Chironomoidea.
Members of the other family usually included in Culicoidea, the Dixidae, lack heteromor-
phic sex chrorno~omes,~~~ as in the Chironomoidea.
Sex determination in Chironomoidea-or at least, in Simuliidae and Chironomidae, since
the Ceratopogonidae are little studied - is characterized by two features: (1) there are almost
always three chromosome pairs, none of which are heteromorphic; and (2) there is usually a
dominant male-determining factor that can apparently occur almost anywhere in the genome
and often differs in location between closely related species (Figure 12). In the Eusimulium
vemum complex alone, for example, five out of the six chromosome arms are involved in sex
determination in different species and sibling species.'8g In the E. aureum species group,
which is unusual in having 2n = 4, either of the two chromosome pairs may serve as the sex
chromosomes, and sex factors may occur in any of the four chromosome arms.Ig0
Sometimes the location of the sex factor varies within species, in which case sex determi-
nation operates as a multiple factor system. For example, the Australian Chironomus oppositus
species complex includes one form, whitei, which is apparently polymorphic for four different
sex factor locations, with up to three locations occurring in any one population.lgl Many other
examples are now available which support the idea that the sex-determining locus in
Sex Determination in Insects

FIGURE 12. Diagrammatic illustration of the mobility of the male-determining factor in blackflies (Simuliidae).
The diagram also shows the standard notation used by blackfly cytogeneticists for the six arms of the three
chromosome pairs (I, 11, and 111) of the normal blackfly chromosome complement. Any of these six arms can function
as the sex chromosomes, due to transposition of the male-determining factor (M) between chromosomes.The location
of M could also be switched from one arm to the other of the same chromosome by a pericentric inversion; for
example, an inversion of the section bracketed by a dotted line on chromosome I (although paracentric inversions -
those not involving the centromere - are much more common in blackflies).

Chironomoidea - and in many of the higher Diptera discussed below - acts as, or is
associated with, a transposable element, and can thus be excised and moved to multiple
locations in the gen~me.~O
The location of the male sex factor can sometimes be detected cytologically in polytene
chromosomes by minor differences in the banding pattern, or by sex-linked inversions.
Inversions arise when sections of the chromosome of varying length are excised and then
reinserted in the chromosome the "wrong way around". Individuals heterozygous for an
inversion can be detected by examining the sequence of bands in the polytene chromosomes,
which is inverted in a section of one chromosome in comparison with its homologue. If an
inversion is close to or encompasses the sex factor locus, as seems to happen very frequently
in blackflies, then it will be partially or completely sex-linked.192 The frequency of sex-linked
inversions also fits the idea of a transposable element being involved in sex determination,
because the breakpoints for inversions can also be the sites of excision or insertion of
transposable elements.193
R o t h f e l ~ considered
l~~ that the ancestral condition for Simuliidae and related groups was
a complete absence of differentiation of the chromosome carrying the male-determining factor
(the notation used for undifferentiated, homomorphic sex chromosomes by simuliid cytoge-
neticists is )dY,). While this may well be true, it is also possible for the )dYo condition to be
secondary; if, for example, a sex locus associated with an inversion is transposed out of the
inversion to a new genomic site, so that the inversion is no longer ~ e x - l i n k e d . ~ ~ ~ . ~ ~ ~
In Chironomus tentans, males are normally heterozygous for a dominant male sex factor,
but one population was found that seemed to have female heterogamety; this was interpreted,
on the basis of crosses between populations, as due to a dominant female sex f a ~ t o r . ' ~ . ' ~ ~
Nijthiger and Steinmann-Zwicky1Ipostulated that this situation might arise by a null mutation
of the key gene Sxl, accompanied by loss of the dominant male sex factor (M). It has also been
suggested, however, that a model involving a weakened male determiner could provide a
better explanation of the published result^.^^'.^^^
The remaining groups of Nematocera all have an achiasmate male meiosis, a feature that
links them cytogenetically with the higher Diptera. The sex chromosomes of Thaumaleidae
and Bibionoidea are usually small, do not form a bivalent, and show "distance pairing" in the
first meiotic division, as in tip~1ids.I~~ The Mycetophilidae also have XY males, but the related
82 Insect Reproduction

families Sciaridae and Cecidomyidae have developed remarkably aberrant chromosome sys-
tems, with more chromosomes in the germ line than in the soma (reviewed in detail by
White9). Neither of these families have Y chromosomes, so that the genic balance between X
chromosomes and autosomal factors must be the basis for sex determination.
In Sciara, male somatic cells are XO, but the germ line is XX and, after passing through
a highly peculiar spermatogenesis, the sperm are homogametic and all carry two X chromo-
somes. Oocytes are normal, with a single X, so all zygotes have three X chromosomes, with
the potential to develop as either sex. Either one or two of the three X chromosomes are
eliminated from presumptive somatic cells at the seventh or eighth cleavage division, to
determine the soma of the embryo as either female (XX) or male (XO), respectively. (Germ-
line cells later lose one X chromosome, irrespective of the sex of the embryo, so that they are
XX in both sexes.) The sex of the offspring - i.e., whether one or two X chromosomes are
eliminated from somatic cells - depends entirely on the genetic constitution of the mother.
Certain species of Sciara are monogenous, i.e., they invariably have unisexual progenies, so
that there are two kinds of mother, male producing and female producing. The latter are
thought to be heterozygous for a dominant factor (F), presumably acting through the cyto-
plasm of the egg to cause the soma of the embryo to develop as female. The ratio of male- to
female-producing mothers in such species, and hence the resulting sex ratio, is approximately
1:l. Thus, sex is inherited genetically, but the inheritance is displaced back to the maternal
generation. The genetic mechanism could be a null mutation of the daughterless gene or its
equivalent, as discussed for Chrysomya below. Other species of Sciara have females that
normally produce progeny of both sexes, however, and sex determination in other genera of
Sciaridae has still hardly been studied, so it would be unwise to generalize. Haig'99 reviewed
the chromosome system of Sciara coprophila and developed a model for its evolution based
on sex ratio theory.
Cecidomyidae have even more aberrant chromosome systems, with numerous extra ("E")
chromosomes in the germ line that are eliminated from somatic cells in early cleavage
division^.^ In most species studied, there are six chromosomes in male somatic cells and eight
in female somatic cells, so that the sex chromosome system is XlXlX2X2/XlX20. This is the
case in the hessian fly, Mayetiola destr~ctor,2~.20' despite early reports of eight chromosomes
in the somatic cells of both sexes. As in Sciara, sex is determined by a maternal factor rather
than by male heterogamety. Males although X,X,O are homogametic, producing only XlX2
sperm, so that zygotes are all XlX1X2X2.In male embryos, two X chromosomes (one X, and
one X,) are eliminated from presumptive somatic cells at a separate, later cleavage division
than that at which the E chromosomes are eliminated; e.g., in Wachtliella persicariae, E
chromosomes are eliminated at the fourth cleavage division and X chromosomes at the
seventh.202
As in Sciaridae, many cecidomyids have unisexual pro genie^,^^^^^ but others have the
same mothers producing both male and female progeny, and the system by which sex is
controlled is unclear. Heteropeza pygmaea is best studied in this respect, but Heteropezinae
differ from other cecidomyids in that the male somatic cells appear to be haploid, with five
chromosomes, whereas female somatic cells have ten chromosomes. However, this only
applies when the progeny are produced pedogenetically; in H. pygmaea, females reproducing
as adults lay only female-determined eggs, but these have five chromosomes as in
pedogenetically produced male Sex determination therefore cannot be based on
haplodiploidy, and does not seem to have a genetic basis at all. Went and Camenzind205
cultured larval ovaries of H. pygmaea in vitro, using as culture medium the hemolymph of
larvae that had been previously kept in different nutritional environments, and were able to
show that the sex of the progeny was dependent on the nutritional conditions experienced by
the mother during development.
The more primitive groups of Brachycera have received very little attention from cytoge-
neticists. In the Tabanoidea, Rhagionidae and Stratiomyidae have XY males where these have
Sex Determination in Insects 83

been studied,206whereas in Asiloidea, the asilid Dasyllis (=Laphria) grossa is reported to have
an XXIXO system.207In the more advanced groups of Brachycera (=Cyclorrhapha), there has
been detailed work on sex determination mechanisms of representatives of five families:
Phoridae (Megaselia scalaris), Muscidae (Musca domestica), Calliphoridae (Chrysomya
ru$facies Lucilia cuprina), Tephritidae (Ceratitis capitata), and, of course, Drosophilidae (D.
melanogaster, D. miranda). These five families span four superfamilies of Cyclorrhapha, so
may be fairly representative of the range of mechanisms in the higher Diptera as a whole.
In the phorid fly M. scalaris, X and Y chromosomes are not morphologically differentiated,
and the male-determining factor (M) is capable of being located on any of the three chromo-
some pairs,208much as in chironomids. In laboratory strains the chromosome (Y) carrying the
M factor could be distinguished from its homologue (X) using a combination of cytogenetic
and molecular technique^.^^ The segment of the Y chromosome carrying M was found to be
conserved in comparison with the homologous region of X, when two unrelated strains were
compared. Nevertheless, when the two strains were crossed, four cases were found where the
M factor had moved to a different chromosome. The frequency of this change was about
0.06%, which is comparable with known rates of movement of transposable elements in other
organisms.210The conservation of the M-containing chromosomal region observed in pure
strains perhaps indicates that a specific location is favored under certain circumstances, and
this could be the first step in the differentiation of new heteromorphic sex chromosomes.
The housefly M. domestica provides some particularly interesting examples of evolution
of sex-determining systems in progress. It was fully reviewed by Bull,lo but since then there
have been further interesting developments. Earlier European work established that sex
determination in houseflies was XXIXY, with heteromorphic sex chromosomes and a presum-
ably dominant male sex factor on the Y. Apart from the Y-borne sex factor, X and Y
chromosomes seem to have few or no functional coding regions and are heterochromatic. In
strains of non-European origin, however, various sex factors have been found on the auto-
somes, especially a male determiner (M) near the centromere on autosome 3, and a female
determiner (F) on autosome 4 which is epistatic to (i.e., ovemdes) any number of male-
determining factor~.~" In continental Europe, samples from Denmark to Sicily taken in 1975-
1981 showed a latitudinal cline: north European populations were all XXIXY, whereas in
south and central Italy all populations were XXIXX with sex determined autosomally, the X
being totally neutral with regard to sex determination. In southern France, Yugoslavia, and
northern Italy, intermediate, mixed populations occurred with all combinations of X and Y in
either A very similar north-south cline was found in Japan.213The changes in sex
determination mechanisms in both southern Europe and Japan are believed to be recent.
Various models (e.g., Jayakar2I4)have been advanced to explain this phenomenon; possibly
climatic influences are involved, or perhaps the driving force is selective insecticide pressure,
as there is now good evidence that pyrethroidJDDT resistance (the "knockdown factor," Kdr)
is genetically linked with the male-determining locus on autosome 3.215However, recent
changes have also occurred in the sex-determination system of housefly populations in
southeast England, involving an apparent increase in frequency of a male factor on the X
c h r o m ~ s o m e , ~and
~ ~there
. ~ ~ 'is no evidence that loci associated with insecticide resistance (or,
in fact, any other functional genes) occur on the housefly X chromosome. Or has the
resistance-conferring gene also been transposed to the X from autosome 3, along with the male
sex factor? This seems quite possible, since a laboratory housefly strain in Australia was
shown to have DDT resistance linked to a male sex factor, but in this case on autosome 2.218
Few other Muscidae have been studied cytogenetically. Recent work on Hydrotaea
meriodionalis indicates a similar story to M. domestica, with a dominant autosomal male
determiner in some populations and others with XY males.219In the closely related
Anthomyiidae, the cabbage root fly, Delia radica (=Hylemyia brassicae), is now known to
have a male-determining factor on an autosome (chromosome 6), whereas D. antiqua has a
small heteromorphic X and Y.220
84 Insect Reproduction

Sex determination in houseflies, with male determiners at various locations in the genome,
and the presence of a dominant female determiner in some populations, seems a very different
mechanism to the Drosophila system, based on an X:A ratio, but there are ways of deriving
one from the other fairly simply. Nothiger and Steinmann-Zwickyl' suggested, for example,
that the dominant female determiner (F) could be a mutation of the key gene Sxl to an
irrepressible condition, so that it cannot be turned off by M. A similar conclusion was reached
by Inoue and H i r o y i ~ h i ; ~their
~ ' model for housefly sex determination incorporates the
discovery of a mutation tra, closely linked with F on autosome 4; when this is present in the
mother, it causes progeny to develop as males even in the absence of any M factors.
In fact, a genic balancelrecessive-X system does operate in another muscid, the tsetse fly
G. palpalis, which also resembles Drosophila in that the Y chromosome carries some loci that
are necessary for sperm viability, but is not involved in sex determination.16
Blowflies (Calliphoridae) generally have small, heteromorphic, and mainly heterochro-
matic, sex chromosomes. The Y chromosome in L. cuprina carries a dominant male sex factor,
located near its c e n t r ~ m e r e .In ~ ~Calliphora
~ . ~ ~ ~ erythrocephala, however, the small hetero-
chromatic pair are no longer sex chromosomes, and the male-determining locus is on one of
the other chromosomes, where it is recognizable as a small heterozygosity of the chromomere
pattern of the polytene chromosome.224And in the monogenic blowfly C. rufifacies, sex is
controlled by a dominant female determining factor (F') in the mother, Flfmothers producing
only daughters andflmothers producing only sons, which are therefore also of ff genotype.
U l l e r i ~ h , 2in~some
~ elegant experimental work, transplanted pole cells (primordial germ cells)
between female embryos of Flf and ff genotypes. The resultant mothers were germ-line
mosaics for Flfand ff, and both the donor and recipient genotypes were expressed, resulting
in a mixture of male and female progeny. Thus, the F'gene product is synthesized by the germ-
line cells themselves, rather than by maternal somatic cells. Ullerich also did pole cell
transplantations between male and female embryos. These resulted in germ-line mosaics that
were completely fertile and heterosexual; the donor cells underwent sex reversal and devel-
oped as male or female according to their mother's genotype and irrespective of their own
genotype. Thus, a genotypically male germ cell can develop as a functional oocyte in a female
host, a genotypically female germ cell can develop as a functional sperm in a male host, and
sex is determined solely by regulatory factors provided by maternal somatic cells.
Nothiger and Steinmann-Zwickyll postulated that F'in Chrysomya is similar or identical
to the daughterless gene (da)of Drosophila, which is necessary in the mother in order for the
key gene Sxl to be active (Figure 6). If f is the null (mutant) allele da-, then in homozygous
condition it will render Sxl of embryonic germ cells inactive, so that all progeny will be sons.
DNA sequence homology has now been demonstrated between the da gene of Drosophila and
a polytene band on the Chrysomya chromosome that carries the F ' l o ~ u s strongly
, ~ ~ ~ support-
ing this hypothesis.
Tephritidae mostly have heteromorphic sex chromosomes (XXIXY), and in several cases
a dominant-Y system has been demonstrated, e.g., in the medfly C. ~ a p i t a t aX,X,Y . ~ ~ ~ males
occur in some s p e c i e ~ , 9 Indian
. ~ ~ ~ species in four genera of Trypetinae apparently have
homomorphic sex chromosomes,229and female heterogamety (XYKX) has been demon-
strated in some Australian species.230In C. capitata, the sex chromosomes are almost entirely
heterochromatic, and the Y chromosome can suffer large deletions without any obvious ill
effect; the male-determining factor is located on its long arm close to the c e n t r ~ m e r eSome
.~~~
repetitive DNA sequences that are specific to or concentrated in the Y chromosome of C.
capitata were recently is0lated.~3~
The sex determination mechanism of D. melanogaster was discussed earlier in this chapter,
and there are numerous recent r e v i e ~ s . ~ .AS . ~ ~ ~Drosophila
~ ~regards - ~ ~ ~ other than D.
melanogaster, the most interesting developments have been with D. miranda, a species in the
obscura group which has an XlX2Y system, the X, and the Y being recently derived (i.e., a
neo-X and neo-Y) by translocation to the original Y chromosome of one member of the third
Sex Determination in Insects 85

autosome pair found in the closest relatives (D. pseudobscura, D. persimilis), leaving its
homologue as a neo-X. Chromosome 3 of D. pseudobscura/persimilis is also homologous to
the right arm of chromosome 2 of D. melanogaster. Thus, a very comprehensively mapped
chromosome segment has quite recently become a neo-Y, providing considerable scope for
study of the degenerative changes that follow from its permanent heterozygosity, and the
consequent accumulation of nonfunctional alleles. Comparisons of the neo-Y and its recently
homologous neo-X have particularly shown that the neo-Y has acquired inserted DNA
sequences that are not present in the neo-X, and appear to represent a novel transposable
element that may be involved in the degenerative p r o ~ e s s . ~ ~ ' - ~ ~ ~

IV. EVOLUTION OF SEX CHROMOSOMES AND SEX

DETERMINATION IN INSECTS
It has taken quite a lot of words to provide an outline review of the many and various
methods by which sex determination is achieved in the different orders of insects. It is clearly
important to distinguish between the sex chromosome systems, which display to the cytoge-
neticist a remarkable diversity in their form, behavior, and extent of evolutionary change
within and between groups, and the underlying molecular mechanisms, which may perhaps
show less variation. However, for most insect groups, the molecular genetics of the sex-
determining process is still merely a matter of speculation or extrapolation from the paradigm
of Drosophila.
Much of the discussion on sex chromosome evolution has centered on the Y chromosome.
The ideas about the progressive evolutionary degeneration of the Y chromosome discussed
earlier in this chapter were developed primarily with regard to vertebrate systems, particularly
mammals where the Y chromosome bears a dominant male-determining l o ~ ~ sThe .
various models that have been proposed241all assume a primitive condition where the sex
chromosomes are undifferentiated, homologous, and in fact essentially autosomal except at
the sex-determining locus, but in time become progressively differentiated as the Y acquires
noncoding DNA and the X acquires a system of dosage compensation.14 Bullioreviewed the
evidence for progressive sex chromosome differentiation in other groups including insects,
and concluded that it could be applied more generally. Nothiger and Steinmann-Zwicky"
postulated that the various genetic mechanisms for sex determination in higher Diptera arose
by mutations from a primitive state with undifferentiated sex chromosomes and a dominant
male determiner, as in some mosquitoes.
However, when insects are looked at as a group, certain qualifications to the model of
progressive sex chromosome differentiation and Y degeneration are necessary. First, a model
that is applicable to the genic balance systems that seem to predominate in insects has not yet
been developed.I4 Second, the XXIXO system predominates in the lower insect orders, and
must be the ancestral condition for several major groups, if not for the class Insecta as a whole.
Thus, for many insects with XXIXY, if not the majority, the XY condition has arisen by a
major chromosomal rearrangement, rather than by a progressive, gradual change from a
primitively undifferentiated state. The subsequent evolution of the neo-Y and the homologous
region of the neo-X may be comparable in many ways to the process of mammalian sex
chromosome differentiation, but the presence from the start of a fully differentiated X
chromosome, coupled with the lack of sex determiners on the Y, must have consequences that
need to be fully addressed. It would be instructive to compare the molecular changes in the
recently acquired neo-XY system of Drosophila miranda with the changes following a recent
acquisition of "autosomal" sex determination due to the transposition of a male determiner
(M) to a new location, e.g., in certain mosquito species or, very recently, in certain populations
of M. domestica.
Third, although there are examples of loss of homology between a neo-Y and a neo-X, and
evidence of accumulation of nonfunctional alleles and repetitive DNA on the Y chromosome,
86 Insect Reproduction

there is a lack of information about the circumstances which determine whether a neo-XY
becomes an evolutionarily stable XY system, or proceeds inevitably towards complete degen-
eration and eventual loss of the Y chromosome. In the cytologically well-studied Orthoptera,
which show much evolutionary change in the sex chromosomes including numerous examples
of de novo acquisition of XY systems, there are no clear cases where the Y chromosome has
been secondarily lost. It seems that a stable condition may sometimes be reached, where it is
advantageous to have sex-linked genes retained on the Y chromosome. In Coleoptera, Xy,
systems with a "degenerate Y" are very ancient and show great evolutionary stability. In
Heteroptera, secondary loss of Y chromosomes seems to have occurred many times in the
course of evolution at the family level, but not between closely related species, indicating that
it does not happen fast or frequently.
This leads to the fifth and final point of qualification, which was discussed by Feraday et
al.'95 specifically with respect to the evolution of the sex chromosomes of Simuliidae. There
has been a tendency to regard sex chromosome differentiation as an inevitable sequence of
events, under the influence of mutation and random drift, rather than as an adaptive process.
In Simuliidae, any of the three chromosome pairs can be heterozygous for the male-determin-
ing sex factor. Usually the only cytological differentiation between the "X" and the "Y" is in
the form of inversions, which may be sex-linked but do not form part of any progressive
evolutionary sequence of sex chromosome differentiation.lg5Whitegpointed out that if certain
autosomal alleles are polymorphic and exert different selective pressures in the two sexes, then
it is advantageous to have them linked to the sex chromosomes. In Orthoptera this may be
accomplished by centric fusions between sex chromosomes and autosomes to give neo-XY
systems.242In those Diptera which have single locus, dominant male sex factors, the linkage
may be more easily obtained by transposing the sex locus to another position in the genome.
Selective advantage is thus important in establishing a new sex chromosome system, and
presumably continues to influence the nature and extent of any subsequent differentiation of
the X and Y chromosomes.

I. Henking, H., Untersuchungen iiber die ersten Entwichlungsvorgange in die Eiern der Insekten. 11. ~ b e r
spermatogCnltse und deren Beziehung zur Entwicklung bei Pyrrhocoris apterus. 2. Wiss. Zool. Abt. A., 51,
685, 1891.
2. Bridges, C. B., The origin of variations in sexual and sex-limited characters, Am. Not., 56, 51, 1922.
3. Bridges, C. B., Sex in relation to genes and chromosomes. Am. Nat., 59, 127, 1925.
4. Bridges, C. B., Cytological and genetic basis of sex, in Sex and Internal Secretions, 2nd ed., Allen, C., Ed.,
Williams & Wilkins, Baltimore, 1939, 15.
5. Goldschmidt, R. B., The Mechanism and Physiology of Sex Determination. Methuen, London, (translation
by W. J. Dakin), 1923.
6. Goldschmidt, R. B., Die Sexuellen Zwischenstufen, Monogr. Gesamtgeb. Pjlanzen Tiere, Vol. 23, Springer,
Berlin, 1931.
7. Baker, B. S. and Belote, J. M., Sex determination and dosage compensation in Drosophila melanogaster,
Annu. Rev. Genet., 17, 345, 1983.
8. Baker, B. S., Sex in flies: the splice of life, Nature (London), 340, 521, 1989.
9. White, M. J. D., Animal Cytology and Evolution, 3rd ed., Cambridge University Press, Cambridge, U.K.,
1973.
10. Bull, J. J., Evolution of Sex-Determining Mechanisms. Benjamin/Cummings Publishing, Men10 Park, CA,
1983.
11. Nothiger, R and Steinmann-Zwicky, M,, A single principle for sex determination in insects, Cold Spring
Harbor Symp. Quant. Biol.. 50, 615, 1985.
12. Disney, R. H. L. and Cumming, M. S., Abolition of Alamirinae and ultimate rejection of Wasmann's theory
of hermaphroditism in Temitoxeniinae (Diptera, Phoridae), Bonn. 2001. Beitr., 43, 145, 1992.
Sex Determination in Insects 87

13. Bergerard, J., Environmental and physiological control of sex determination and differentiation, Annu. Rev.
Entomol., 17, 57, 1972.
14. Charlesworth, B., The evolution of sex chromosomes, Science, 251, 1030, 1991.
15. Ullerich, F.-H., Geschlechtschromosomen und Geschlechtsbestimmungbei einigen Calliphoren (Calliphoridae,
Diptera), Chromosoma, 14, 45, 1963.
16. Southern, D. I., Chromosome diversity in tsetse flies, in Insect Cytogenetics, Blackman, R. L., Hewitt, G. M,,
and Ashbumer, M,, Eds., Blackwell, Oxford, 1980,225.
17. Steinemann, M. and Steinemenn, S., Degenerating Y chromosome of Drosophila miranda: a trap for
retrotransposons, Proc. Natl. Acad. Sci. U.S.A., 89, 7591, 1992.
18. Rice, W. R., Genetic hitchhiking and the evolution of reduced genetic activity of the Y chromosome,
Genetics, 116, 161, 1987.
19. Charlesworth, B., Model for evolution of Y chromosomes and dosage compensation, Proc. Natl. Acad. Sci.
U.S.A., 75, 5618, 1978.
19a. John, B. and Hewitt, G. M., Patterns and pathways of chromosome evolution within the Orthoptera,
Chromosoma, 25, 40, 1968.
20. Green, M. M., Transposable elements in Drosophila and other Diptera, Annu. Rev. Genet.. 14, 109, 1980.
21. Crozier, R. H., Hymenoptera, Animal Cytogenetics 3, Insecta 7, Gebriider Bomtraeger, BerlinIStuttgart,
1975.
22. Crozier, R. H., Evolutionary genetics of Hymenoptera, Annu. Rev. Entomol., 22, 263, 1977.
23. Whiting, P. W., Multiple alleles in complimentarysex determination of Habrobracon. Genetics. 28,365, 1943.
24. Mackensen, O., Viability and sex determination in the honey bee Apis mellifica, Genetics, 36, 500, 1951.
25. Smith, S. G. and Wallace, D. R., Allelic sex determination in a lower hymenopteran, Neodiprion nigroscutum
Midd., Can. J. Genet. Cytol.. 13,617, 1971.
26. Steinmann-Zwicky, M., Amrein, H., and Nothiger, R., Genetic control of sex determination in Drosophila.
Adv. Genet., 27, 189, 1990.
27. Chandra, H. S., Sex determination: a hypothesis based on noncoding DNA, Proc. Natl. Acad. Sci. U.S.A..
82, 1165, 1985.
28. Parkhurst, S. M., Bopp, D., and Ish-Horowicz, D., X:A ratio, the primary sex-determining signal in
Drosophila, is transduced by helix-loop-helix proteins, Cell, 63, 1179, 1990.
29. Torres, M. and Sanchez, L., The sisterless-b function of the Drosophila gene scute is restricted to the stage
when the X:A ratio determines the activity of Sex-lethal. Development, 113, 715, 1991.
30. Keyes, L. N., Cline, T. W., and Schedl, P., The primary sex determination signal of Drosophila acts at the
level of transcription, Cell, 68, 933, 1992.
3 1. Stewart, B. and Merriam, J., Dosage compensation, in The Genetics and Biology of Drosophila. Ashburner,
M. and Wright, T. R. F., Eds., 2d, Academic Press, LondonhJew York, 1980, 107.
32. Rao, S. R. V. and Ali, S., Insect sex chromosomes 6: a presumptive hyperactivation of the male X
chromosome in Acheta domesticus, Chromosoma, 86, 325, 1982.
33. Rao, S. R. V. and Arora, P., Insect sex chromosomes. 111: differential susceptibility of homologous X
chromosomes of Gryllotalpafossor to 'H-Urd-induced aberrations, Chromosoma, 74, 241, 1979.
34. Sarkar, S. and Rao, S. R. V., Insect sex chromosomes XI. 'H-TdR induces random aberrations in the X
chromosomes of GryIIotnlpafossor (Orthoptera), Mutat. Res., 282, 113, 1992.
35. Nagl, W., The sex chromatin of Tachycines asynamorus (Orthoptera) and its implications. Cytologia, 38,107,
1973.
36. Johnson, M. S. and Turner, J. R. G., Absence of dosage compensation for a sex-linked enzyme in butterflies
(Heliconius), Heredity, 43, 71, 1979.
37. Nufiez, O., Cytology of Collembola, Nature (London), 194, 946, 1962.
38. Kiauta, B., Review of the germ cell chromosome cytology of Collembola, with a list of chromosome numbers
and data on two species new to cytology, Genen Phaenen, 13, 89, 1970.
Cassagnau, P., Les chromosomespolytknes de Neanura monticola Cassagnau (Collembola). I . Polymorphisme
dcologique du chromosome X, Chromosoma, 46, 343, 1974.
Charlton, H. H., The spermatogenesis of Lepisma domestica, J. Morphol., 35, 381, 1921.
Fratello, B., Citotassonomia dei Proturi (Insecta, Apterygota), Atti Congr. Naz. Ital. Entomol. Siena. 9,267,
1972.
42. Fratello, B. and Sabatini, M. A., Chromosome studies in Protura Eosentomoidea, in Third Int. Sem.
Apterygota, Dallai, R., Ed.,University of Siena, Italy, 1989, 167.
43. Mol, A. W. M., Notes on the chromosomes of some western European Ephemeroptera, Chromosome
Information Service, 24, 10, 1978.
44. Kiauta, B., Sex chromosomes and sex-determining mechanisms in Odonata, with a review of the cytological
conditions in the family Gomphidae, and references to the karyotype evolution in the order, Genetica, 40,127,

45. Kiauta, B., Synopsis of the main cytotaxonomic data in the order Odonata, Odonatologica (Utrecht), 1,73,
1972.
88 Insect Reproduction

46. Oksala, T., Zytologische Studien an Odonata. I. Chromosomenverhaltnisse bei der Gattung Aeschna mit
besonderer Beriicksichtigung der postreduktionellen Teilung der Bivalente, Ann. Acad. Sci. Fenn. A N. 4.1.1943.
47. Tyagi, B. K., Cytotaxonomy of the genus Onychogomphus Selys (Odonata: Anisoptera, Gomphidae), with
special reference to the evolution of the sex-determiningmechanism and the reduced chromosome number in
the family Gomphidae, in Proc. 1st Indian Symp. Odonatology. Mathavan, S., Ed., Madurai Kamaraj
University, Madurai, India, 1985, 217.
48. Matthey, R. and Aubert, J., LRs chrOmosomes des Pltcoptkres, Bull. Biol. Fr. Belg., 81, 202, 1947.
49. Hewitt, G. M., Orthoptera, Animal Cytogenetics 3, Insecta 1, Gebriider Borntraeger, BerlinIStuttgart. 1979.
50. White, M. J. D., Blanodea, Mantodea, Isoptera, Grylloblattodea, Phasmatodea, Dermaptera, Embioptera,
Animal Cytogenetics 3, Insecta 2, Gebriider Borntmeger, BerlinIStuttgart, 1976.
5 1. Cohen, S. and Roth, L. M., Chromosome numbers of the Blattaria, Ann. Entomol. Soc. Am., 63, 1520.1970.
52. Luykx, P., X0:XX sex chromosomes and Robertsonian variation in the autosomes of the wood-roach
Cryptocercus punctulatus (Dictyoptera: Blattaria: Cryptocercidae), Ann. Entomol. Soc. Am., 76, 518, 1983.
53. Luykx, P., A cytogenetic survey of 25 species of lower termintes from Australia, Genome. 33, 80, 1990.
54. Tyagi, B. K., Review of the cytotaxonomy of Isoptera (Insecta) with a description of the male germ cell
chromosomes of Microcerotermes beesoni Snyder from the Dehradun Vallet, India, Indian Rev. Life Sci.. 7,
263, 1987.
55. Vincke, P. P. and Tilquin, J. B., A sex-linked ring quadrivalent in Termitidae (Isoptera), Chromosoma. 67,
151, 1978.
56. Fontana, F., Cytological analysis of the chromosome complement of Kalotermesflavicollis Fabr. (Isoptera,
Kalotermitidae). The Sex determining mechanism, Cytologia, 47, 147, 1982.
57. Syren, R. M. and Luykx, P., Permanent segmental interchange complex in the termite Incisitermes schwarzi,
Nature (London), 266, 167, 1987.
58. Syren, R. M. and Lyukx, P., Geographic variation of sex-linked translocation heterozygosity in the termite
Kalotermes approximatus Snyd. (Insecta: Isoptera), Chromosoma, 82, 65, 1981.
59. Syren, R. M. and Luykx, P., Experimental hybridization between chromosomal races in Kalotermes
approximatus, a termite with extensive sex-linked translocation heterozygosity. Chromosoma, 83,563, 1981.
60. Luykx, P. and Syren, R. M., Multiple sex-linked reciprocal translocations in a termite from Jamaica,
Experientia. 37, 819, 1981.
61. Luykx, P. and Syren, R. M., The cytogenetics of Incisitermes schwani and other Florida termites, Socio-
biology, 4, 191, 1979.
62. White, M. J. D., The evolution of sex chromosomes I. The X0 and X,X,Y mechanisms in praying mantids,
J. Genet., 42, 143, 1941.
63. Callan, H. G. and Jacobs, P. A., The meiotic process in Mantis religiosa males, J. Genet., 55,200, 1957.
64. Liebenberg, H., Fossey, A., and Jacobs, D. H., An unexpected sex chromosome mechanism in a South
African mantid Polyspilota aeriginosa Goez, Caryologia, 44, 195, 1991.
65. Craddock, E., Chromosomal evolution and speciation in Didymuria, in Genetic Mechanisms of Speciation
in Insects, White, M . J . D., Ed., Australia and New Zealand Book Co., Sydney, 1974, 24.
66. BergCrard, J., Intersexualit15experimentale chez Carausius morosus Br. (Phasmidae), Bull. Biol. Fr. Belg.,
95, 273, 1961.
67. Pijnacker, L. P., Effect of centrifugation of the eggs on the sex of Carausius morosus Br., Nature (London),
210, 1184, 1966.
68. Pijnacker, L. P., Effects of X-rays on different meiotic stages of oocytes in the parthenogenetic stick insect
Carausius morosus Br., Mutat. Res., 13, 251, 1971.
69. I'Helias, C. and Boulanger-Sandrin, J., Influence des pttrines kduites et de I'hormone juvenile sur
I'intersexualitt du Phasme Carausius morosus, Ann. Endocrinol., 37, 189, 1976.
70. Pijnacker, L. P. and Ferwerda, M. A., Sex chromosomes and origin of males and sex mosaics of the
parthenogenetic stick insect Carausius morosus, Chromosoma, 79, 105, 1980.
71. Pijnacker, L. P. and Harbott, J., Structuralheterozygosity and aneuploidy in the parthenogenetic stick insect
Carausius morosus Br. (Phasmatodea: Phasmatidae), Chromosoma, 76, 165, 1980.
72. Mesa, A., Ferreira, A., and Carbonell, C. S., Cariologie de 10s acridoides neotropicales, estado actual de
su conocimiento y nuevas contribucones, Ann. Soc. Entomol. Fr., 18, 507, 1982.
73. Yadav, J. S. and Yadav, A. S., X-autosome fusion in catantopine grasshoppers (Acridoidea: Orthoptera),
Cytobios, 61, 21, 1990.
74. Bugrov, A. G., [Neo-XY sex chromosome determination in grasshoppers Asiotmethis heptaopamicus (Zub.)
and Atrichotmethis semenovi (Zub.) (Orthoptera: Pamphagidae)], Tsitologiya, 28, 117, 1986.
75. Yadav, J. S. and Yadav, A. S., Chromosome number and sex-determining mechanisms in 30 species of
Indian Orthoptera, Folia. Biol. (Krakow), 34, 277, 1986.
76. Fernandez-Piqueras, J., Rojo Garcia, E., and Sentis Castano, C., A tandem fusion origin of a neo XY sex
determining mechanism in the long-horned Callicrania seoanei (Bol.), Heredity, 47, 397, 1981.
77. Fernandez-Piqueras, J., Rodriguez Campos, A., Sentis Castano, C., and Rojo Garcia, E., Sex chromo-
some evolution in the polytypic species Pycnogaster cucullata (Chap.), Heredity, 50, 217, 1983.
Sex Determination in Insects 89

78. Sentis Castano, C. and Fernandez-Piqueras, J., Nature and distribution of heterochromatinized regions in
the chromosomal races of Pycnogaster cucullata (Insecta, Orthoptera), Genetica, 72, 127, 1987.
79. White, M. J. D., Blackith, R. E., Blackith, R. M., and Cheney, J., Cytogenetics of the viatica group of
morabine grasshoppers: I. The "coastal" species, Austr. J. Zool., 15, 263, 1967.
80. Hewitt, G. M. and John, B., Inter-population sex chromosome polymorphism in the grasshopper Podisma
pedestris. 11. Population parameters, Chromosoma, 37, 23, 1972.
81. Bella, J. L., Westerman, M., Lopez Fernandez, C., Delatorre, J., Ruhio, J. M., and Gosalvez, J., Sex
chromosome and autosome divergence in podisma (Orthoptera) in Westem Europe, Genet. Sel., 23,5, 1991.
82. Sands, V. E., The neo-XY system of Catantops humilis (Acrididae: Catantopinae) in Malaysia, Biol. J. Linn.
Soc., 39, 269, 1990.
83. Saltet, P., Les Dolichopodes de Corse (Orthoptera - Raphidophoridae) 1' Ctude cytologique preliminaire,
Bull. Soc. Hist. Nat. Toulouse, 104, 165, 1967.
84. Webb, G. C. and White, M. J. D., A new interpretation of the sex-determiningmechanism of the European
earwig, Forficula auricuiaria. Experientia. 26, 1387, 1970.
85. Meinander, M., Halkka, O., and Sderland, V., Chromosomal evolution in the Psocoptera, Not. Entomol..
54, 81, 1974.
86. Buxton, P. A,, The Louse. Edward Amold, London, 1939.
87. Ueshima, N., Hemiptera. 11: Heteroptera, Animal Cytogenetics 3, Insecta 6, Gebriider Bomtraeger,
Berlin/Stuttgart. 1979.
88. Muramoto, N., A chromosomal study of thirty Japanese heteropterans (Heteroptera),Genetica, 49,37, 1978.
89. Calabrese, D. M. and Tallerico, P., Cytogenetic study in males of Nearctic genera of Gerridae (Hemiptera:
Heteroptera), Proc. Entomol. Soc. Wash., 86, 354, 1984.
90. Nokkala, S. and Nokkala, C., The occurrence of the X 0 sex chromosome system in Dicryonota tricornis
(Schr.) Tingidae Hemiptera and its significance for concepts of sex chromosome system evolution in
Heteroptera, Hereditas, 100, 299, 1984.
91. Schuh, R. T., The influence of cladistics on heteropteran classification,Annu. Rev. Entomol.. 31, 67, 1986.
92. Cobben, R. H., Evolutionary Trends in Heteroptera. I. Eggs. Architecture of the Shell, Gross Embroylogy and
Eclosion, Center for Agricultural Publication and Documentation, Wageningen, 1968, 374.
93. Messthaler, H. and Traut, W., Phases of sex chromosomeinactivation in Oncopeftusfasciarus and Pyrrhocoris
apterus (Insecta, Heteroptera), Caryologia, 28, 501, 1975.
94. Thomas, D. B., Jr., Chromosome evolution in the Heteroptera (Hemiptera): Agmatoploidy versus aneu-
ploidy, Ann. Entomol. Soc. Am., 80, 720, 1987.
95. Emeljanov, A. F. and Kirillova, V. I., Trends and modes of karyotype evolution in the Cicadina (Homoptera).
I. Karyotypic peculiarities and evolutionary changes in the karyotypes of cicadas of suprafamily Cicadelloidea,
Entomol. Rev. (USSR), 69,62, 1990.
96. Emeljanov, A. F. and Kirillova, V. I., [Trends and types of karyotype evolution in Cicadina (Homoptera).
11. Peculiarities and evolutionary changes of the karyotypes in the superfamilies Cercopoidea, Cicadoidea,
Fulgoroidea and in the Cicadina as a whole, Entomol. Rev. (USSR), 71, 59, 1992.
97. den Hollander, J., The chromosomes of Niloparvata lugens Stal. amd some other Auchenorrhyncha,
Cytologia, 47, 227, 1982.
98. John, B. and Claridge, M. F., Chromosome variation in British populations of Oncopsis (Hemiptera:
Cicadellidae), Chromosoma. 46, 77, 1974.
99. Maryahska-Nadachowska, A., Kuznetsova, V. G., and Warchalowska-Sliwa, E., Karyotypes of Psyllina
(Homoptera). 1. New data and check list, Folia Biol. (Krakdw), 40, 15, 1992.
100. Maryanska-Nadachowska, A. and Hodkinson, I. D., Karyotypes of Psylloidea. 11. Chromosome numbers
of nine Mediterranean species from Mallorca (Spain), Folia Biol. (Krakdw), 41, 1, 1993.
101. Marya'nska-Nadachowska, A., Glowacka, E., and Warchalowska-Sliwa, E., Karyotypes of Psylloidea
(Homoptera). 111. Chromosome numbers of eight species belonging to the families Aphalaridae, Psyllidae,
Homotomidae and Triozidae, Folia Biol. (Krakdw). 41, 7, 1993.
102. Schrader, F., Sex determination in the white-fly (Trialeurodes vaporariorum), J. Morphol., 34, 267, 1920.
103. Thomsen, M., Studien iiber die Parthenogenese bei einigen Cocciden und Aleurodiden, 2. Zellforsch.
Mikrosk. Anar.. 5, 1, 1927.
104. van Lenteren, J. C. and Noldus, P. J. J., Whitefly-plant relationships: behavioural and ecological aspects,
in Whiteflies: Their Bionomics, Pest Status and Management, Gerling, D., Ed.,Intercept, Andover, U.K.,
1990, 47.
105. Sharaf, N. and Batta, Y., Effect of some factors on the relationship between the whitefly Bemisia tabaci
Genn. (Homopt., Aleyrodidae) and the parasitoid Eretmocerus mundus Mercet (Hymenopt., Aphelinidae), Z.
Angew. Entomol., 99, 267, 1985.
106. Nur, U., Unusual chromosome systems in scale insects, in Insect Cytogenetics, Blackman, R. L., Hewitt,
G. M., and Ashbumer, M., Eds., Blackwell, Oxford, 1980, 97.
107. Nur, U., Chromosomes, sex-ratios and sex determination, in Armored Scale Insects, Their Biology, Natural
Enemies and Control, Rosen, D., Ed.,Elsevier, Amsterdam, 1990, 179.
90 Insect Reproduction

108. Brown, S. W., Adaptive status and genetic regulation in major evolutionary changes of coccid chromosome
systems, Nucleus, 20, 145, 1977.
109. Haig, D., The evolution of unusual chromosomal systems in coccoids: extraordinary sex ratios revisited, J.
Evol. Biol., 6, 69. 1993.
110. Blackman, R. L. and Hales, D. F., Behaviour of the X chromosomes during growth and maturation of
parthenogenetic eggs of Amphorophora tuberculata (Homoptera, Aphididae),in relation to sex determination,
Chromosoma, 94, 59, 1986.
11 1. Orlando, E., Sex determination in Megoura viciae Buckton (Homoptera, Aphididae), Monit. Zool. Ital.
(N.S.), 8, 61, 1974.
112. Blackman, R. L., Chromosomes and parthenogenesis in aphids, in Insect Cytogenetics, Blackman, R. L.,
Hewitt, G. M., and Ashburner, M., Eds., Blackwell, Oxford, 1980, 133.
113. Hales, D. F. and Mittler, T. E., Chromosomal sex determination in aphids controlled by juvenile hormone,
Genome, 29, 107, 1987.
114. Kawada, K., Polymorphism and morph determination, in Aphids, Their Biology, Natural Enemies and
Control, 2A, Minks, A. K. and Harrewijn, P,, Eds., Elsevier, Amsterdam, 1987, 255.
115. Blackman, R. L., Spermatogenesis in the aphid Amphorophora tuberculata (Homoptera, Aphididae),
Chromosoma. 92, 357, 1985.
116. Blackman, R. L., Stability of a multiple X chromosome system and associated B chromosomes in birch
aphids (Euceraphis spp., Homoptera: Aphididae), Chromosoma, 96, 318, 1988.
117. Blackman, R. L., The chromosomes of Lachnidae, Acta Phytopathol. Entomol. Hung., 25, 273, 1990.
118. Hales, D. F., The chromosomes of Schoutedenia lutea (Homoptera, Aphidoidea, Greenideinae), with an
account of meiosis in the male, Chromosoma, 98, 295, 1989.
119. Blackman, R. L., Aphid cytology and genetics, in Evolution and Biosystematics of Aphids, Szelegiewicz, H.,
Ed., Ossolineum, Warsaw, 1985, 17 1.
120. Morgan, T. H., The predetermination of sex in phylloxerans and aphids, J. Exp. Zool.. 19, 285, 1915.
121. Steffan, A. W., Zum Generation- und Chromosomenzyklus der Adelgidae (Homoptera: Aphidina), Verh.
Dtsch. Zool. Ges.. 1967, 762.
122. Bournier, A., Contribution B 1'Ctude de la parthtnogenbse des Thysanoptbres et de sa cytologie, Arch. Zool.
Exp. Gen., 93, 219, 1956.
123. Risler, H. and Kempter, E., Die Haploidie der Mhnchen und die Endopolyploidie in einigen Geweben von
Haplothrips (Thysanoptera), Chromosoma, 12, 351, 1961.
124. Mound, L. A., Patterns of sexuality in Thysanoptera, in 1991 Conference on Thrips (Thysanoptera): Insect
and Disease Considerations in Sugar Maple Management, U.S.D.A. Forest Service, North Eastern Forest
Experimental Station General Tech. Rep. NE-161, University Park, PA, 1991.2.
125. Ananthakrishnan, T. N., Bioecology of Thrips, Indira Publishing House, Oak Park, Mf, 1984.
126. Crespi, B. J., Sex-ratio selection in a bivoltine thrips. 1. Conditional sex-ratio manipulation and fitness
variation, Evolution, 42, 1199, 1988.
127. Hughes-Schrader, S., Distance segregation and compound sex chromosomes in mantispids (Neuroptera:
Mantispidae), Chromosoma. 27, 109, 1969.
128. Hughes-Schrader, S., Segregational mechanisms of sex chromosomes in Megaloptera (Neuropteroidea),
Chromosoma, 8 1, 307, 1980.
129. Hennig, W., Insect Phylogeny (English translation by Pont, A. C.), Wiley, Chichester, U.K., 1981.
130. Smith, S. G. and Virkki, N., Coleoptera, Animal Cytogentics 3, Insecta 5, Gebriider Borntraeger,
BerlinIStuttgart, 1978.
131. Virkki, N., Chromosomes in evolution of Coleoptera, in Chromosomes in Evolution of Eukaryotic Groups,
Vol. 2, Sharma, A. K. and Sharma, A., Eds., CRC Press, Boca Raton, E, 1984, 41.
132. Virkki, N., Mazzella, C., and Denton, A., Silver staining of the coleopteran Xy, sex bivalent, Cytobios, 67,
45, 1991.
133. Postiglioni, A., da Silva, A., de Leon, R., and de Vaio, E. S., Three species of Helipodus (Coleoptera,
Curculionidae) with different karyotypes and sex chromosome systems, Genetica, 75, 213, 1987.
134. Serrano, J. and Yadav, J. S., Chromosome numbers and sex-determining mechanisms in adephagan
Coleoptra, Coleopt. Bull.. 38, 335, 1984.
135. Maddison, D. R., Chromosomal diversity and evolution in the ground beetle genus Bembidion and related
taxa (Coleoptera: Carabidae: Trechitae), Genetica, 66, 93, 1985.
136. Yadav, J. S., Kondal, K., and Yadav, A. S., Cytology of Cicindela (Myriochile) undulata and C. (M.)
fastidiosa with a summary of chromosomal data on the Cicindelidae, Cicindela, 17, 1, 1985.
137. Mesa, A. and Fontanetti, C. S., The chromosomes of a primitive species of beetle, Ytu zeus (Coleoptera,
Myxophaga, Tonidincolidae), Proc. Acad. Nat. Sci. Philadelphia, 137, 102, 1985.
138. Petitpierre, E. and Segarra, C., Chromosomal variability and evolution of Chrysomelidae (Coleoptera),
particularly that of Chrysomelinae and palaearctic Alticinae, Entomography, 3, 403, 1985.
139. Petitpierre, E., Recent advancesin the evolutionary cytogeneticsof the leaf beetles (Coleoptera,Chrysomelidae),
Entomography, 6,433, 1989.
Sex Determination in Insects 91

140. Yadav, J. S. and Dange, M. P., Chromosomal investigations on eight species of histerids (Coleoptera:
Histeridae), Elytron (Barcelona), 3, 103, 1989.
141. Yadav, J. S. and Dange, M. P., Chromosome number and sex determining mechanisms in twenty species
of Indian rove beetles (Staphylinidae:Polyphaga), Cell Chromosome Res.. 10, 23, 1987.
142. Gill, T. K., Gulati, M., and Pajni, H. R., Chromosome numbers in Indian weevils (Coleoptera: Curculionidae),
Coleopt. Bull., 44, 437, 1990.
143. Yadav, J. S., Burra, M. R., and Dange, M. P., Chromosome number and sex determiningmechanism in 32
species of Indian Coleoptera (Insecta), Nat. Acad. Sci. Lett. (India), 12, 93, 1989.
144. Juan, C. and Petitpierre, E., Chromosome numbers and sex-determining systems in Tenebrionidae (Co-
leoptera), in Advances in Coleopterology, Zunino, M., Belles, X., and Blas, M.. Eds., European Association
of Coleopterology, Barcelona, 1992.
145. Handa, S. M. and Kochhar, N., Cytology of bmchids: I. Chromosome number and sex mechanism of seven
cytologically unknown species of bmchids, Res. Bull. Panjab Univ. Sci., 37, 145, 1987.
146. Yadav, J. S., Lyapunova, E. A., and Vorontsov, N. N., Chromosome numbers and sex-chromosome
mechanisms in fifty species of Coleoptera from USSR, Folia Biol. (Krakow), 34, 269, 1986.
147. Trivers, R. L. and Hare, H., Haplodiploidy and the evolution of social insects, Science, 191, 249, 1976.
148. Luck, R. F., Stouthamer, R., and Nunney, L., Sex determination and sex ratio patterns in parasitic
Hymenoptera, in Evolution and Diversity of Sex Ratio in Haplodiploid Insects and Mites, Wrensch, D. L. and
Ebbert, M. A., Eds., Chapman and Hall, Engelwood Cliffs, NJ, 1993.
149. Cook, J. M., Experimental tests of sex determination in Goniozus nephanridis (Hymenoptera: Bethylidae),
Heredity, 71, 130, 1993.
150. Stouthamer, R., Luck, R. F., and Werren, J. H., Genetics of sex determination and the improvement of
biological control using parasitoids, Environ. Entomol., 21, 427, 1992.
151. Periquet, T. G., Hedderwick, M. P., El Agoze, M., and Poirie, M., Sex determination in the hymenopteran
Diadromus pulchellus (Ichneumonidae): validation of the one-locus multi-allele model, Heredity, 70, 420,
1993.
152. Ross, K. G. and Fletcher, D. J. C., Genetic origin of male diploidy in the fire ant Solenopsis invicta
(Hymenoptera: Formicidae), and its evolutionary significance, Evolution, 39, 888, 1985.
153. Kerr, W. E. and Nielsen, R. A., Sex determination in bees (Apinae), J. Apic. Res., 6, 3, 1967.
154. Kerr, W. E., Sex determination in bees. XXI. Number of XO-heteroalleles in a natural population of
Melipona compressipes fasciculata (Apidae), Insectes Soc.. 34, 274, 1987.
155. Cook, J. M., Sex determination in the Hymenoptera: a review of models and evidence, Heredity, 71, 421,
1993.
156. Woyke, J., Sex determination, in Bee Genetics and Breeding, 1st edition, Rinderer, T. E. (Ed.),Academic
Press, Orlando, FL, 1986, 91.
157. Adams, J., Rothman, E. D., Kerr, W. E., and Paulino-Sim6es, Z. L., Estimation of the number of sex
alleles and queen matings from diploid male frequencies in a population of Apis mellifera. Genetics, 86,583,
1977.
158. Skinner, S. W. and Werren, J. H., The genetics of sex determination in Nasonia vitripennis. Genetics, 94,
598, 1980.
159. Werren, J. H., Nur, U, and Eickbush, D., An extrachromosomal factor causing loss of paternal chromo-
somes, Nature (London), 327.75, 1987.
160. Werren, J. H., The paternal sex-ratio chromosome of Nasonia, Am. Nat., 137, 392, 1991.
161. Reed, K. M., Cytogenetic analysis of the paternal sex ratio chromosomeof Nasonia vitripennis, Genome. 36,
157, 1993.
162. Folliot, R., Contribution h I'etude de la biologie des cynipides gallicoles (Hymenoptbres, Cynipoidea), Ann.
Sci. Nat. Zool. Biol. Anim. Zool. Ser., 12(6), 407, 1964.
163. Bayreuther, K., Die Cytogenetik zweier norddeutscher Populationen von Nosopsyllus fasciatus Bosc.
(Aphaniptera), Chromosoma. 27, 20, 1969.
164. Suomalainen, E., Achiasmatische Oogenese bei Trichopteren, Chromosoma, 18, 201, 1966.
165. Tazima, Y., Genetics of the Silkworm, Logos Press, Bristol, 1964.
166. Robinson, R., Lepidoptera Genetics, Pergamon Press, Oxford, 1971.
167. Smith, S. G., Heteropycnosis as a means of diagnosing sex, J. Hered., 36, 194, 1945.
168. Traut, W. and Mosbacher, G. C., Geschlechtschromatin bei Lepidopteren, Chromosoma, 25, 343, 1968.
169. Ennis, T. J., Sex chromatin and chromosome numbers in Lepidoptera, Can. J. Genet. Cytol., 18, 119, 1976.
170. Suomalainen, E., On the sex chromosome trivalent in some lepidopteran females, Chromosoma, 28, 298,
1969.
171. Seiler, J. and Schaffer, K., Untersuchungen iiber die Entstehung der Parthenogenesebei Solenobia triquetrella
F.R. (Lepidoptera, Psychidae). 11. Analyse der diploidparthenogenetischen S. triquetrella, Verhalten,
Aufzuchtresultate und Zytologie, Chromosoma, 11, 29, 1960.
172. Clarke, C. and Ford, E. B., Intersexuality in Lymantria dispar (L.). A reassessment, Proc. R. Soc. London.
Ser. B. 206, 381, 1980.
92 Insect Reproduction

173. Gupta, M. L. and Narang, R. C., Karyotype and meiotic mechanism in muga silkmoths, Anrheraea compta
Roth. and A. assamensis (Helf.) (Lepidopten: Saturnidae), Genetica. 57, 21, 1981.
174. Traut, W., Weith, A., and Traut, G., Structural mutants of the W chromosome in Ephestiu (Insects,
Lepidoptera), Genetica, 70, 69, 1986.
175. Fuge, H., The 3-dimensional architecture of chromosome fibres in the cranefly. 11. Amphitelic sex univalents
in meiotic anaphase I, Chromosoma, 91, 322, 1985.
176. Ullerich, F. H., Bauer, H., and Dietz, R., Geschlechtsbestimmung bei Tipuiiden (Nematocera; Diptera),
Chromosoma, 15, 591, 1964.
177. Wolf, E., Die Chromosomen in der Spermatogenese einiger Nematocera, Chromosoma, 2, 192, 1941.
178. White, M. J. D., Cytological evidence on the phylogeny and classification of the Diptera, Evolution, 3,252,
1949.
179. White, G. E. and Killick-Kendrick, R., Polytene chromosomes of the sandfly Lutzomyia longipalpis and the
cytogenetics of Psychodidae in relation to other Diptera, J . Entomol. Ser. A, 50, 187, 1975.
180. Troiano, G., Heterozygous heterochromatin in giemsa C-banded chromosomes of Clogmia albipuncrata
(Telmatoscopus albipunctatus) (Diptera: Psychodidae), Caryologia, 41, 201, 1988.
181. Kreutzer, R. D., Modi, G. B., Tesh, R. B., and Young, D. G., Brain cell karyotypes of six species of New
and Old World sand flies (Dipten: Psychodidae), J. Med. Entomol., 24,609, 1987.
182. Kitzmiller, J. B., Genetics, cytogenetics and evolution of mosquitoes, Adv. Genet., 18, 315, 1976.
183. White, G. B., Academic and applied aspects of mosquito cytogenetics, in Insect Cytogenerics, Blackman,
R. L., Hewitt, G. M., and Ashburner, M., Eds., Blackwell, Oxford, 1980, 245.
184. Newton, M. E., Southern, D. I., and Wood, R. J., X and Y chromosomes of Aedes aegypti (L.) distinguished
by Giemsa C-banding, Chromosoma. 49, 41. 1974.
185. Meuanote, R., Ferrucci, L., and Contini, C., Identificationof sex chromosomes and characterization of the
heterochromatin in Culisera longiaerolata (Macquart 1838). Genetica, 50, 135, 1979.
186. Newton, M. E., Wood, R. J., and Southern, D. I., Cytological mapping of the M and D loci in the mosquito,
Aedes aegypti (L.), Genetica, 48, 137, 1978.
187. Baker, R. H. and Sakai, R. K., Male-determining factor on chromosome 3 in the mosquito, Culex
tritaeniorhynchus, J. Hered., 67,289, 1976.
188. Frizzi, G., Contini, C., and Mameli, M., Ulteriori recherche citogenetiche sui Dixidae della Sardegna, Atti
Assoc. Genet. Iral., 11, 286, 1966.
189. Brockhouse, C., Sibling species and sex chromosomes in Eusimulium vernum (Diptera: Simulidae), Can. J.
Zool., 63, 2145, 1985.
190. Leonhardt, K. G. and Feraday, R. M., Sex chromosome evolution and population differentiation in the
Eusimulium aureum group of black flies, Genome, 32, 543, 1989.
191. Martin, J. and Lee, B. T. O., A phylogenetic study of sex determiner location in a group of Australasian
Chironomus species (Diptera, Chironomidae), Chromosoma, 90, 190, 1984.
192. Rothfels, K. H., Chromosomal variability and speciation in black flies, in Insect Cytogenetics, Blackman,
R. L., Hewitt, G. M,, and Ashburner, M., Eds., Blackwell, Oxford, 1980, 207.
193. Smith, P. a. and Corces, V. G., Drosophila transposable elements: mechanisms of mutagenesis and
interactions with the host genome, Adv. Genet., 29, 229, 1991.
194. Mason, G. F., Sex chromosome polymorphism in the Simulium tuberosum complex (Lundstrom) (Diptera:
Simulidae), Can. J. Zool.. 62, 647. 1984.
195. Feraday, R. M., Leonhardt, K. G., and Brockhouse, C. L., The role of sex chromosomes in black fly
evolution, Genome, 32, 538, 1989.
196. Thompson, P. E. and Bowen, J. S., Interactions of differential primary sex factors in Chironomus tentans,
Genetics, 70, 49 1, 1972.
197. Feraday, R. M., Weak male-determining genes and female heterogarnety in Chironomus tentans. Can. J.
Genet. Cytol., 26, 748, 1984.
198. Martin, J. and Lee, B. T. 0..Are there female heterogametic strains of Chironomus tenrans Fabricius?, Can.
J. Genet. Cytol., 26, 743, 1984.
199. Haig, D., The evolution of unusual chromosome systems in sciarid flies: intragenomic conflict and the sex
ratio, J. Evol. Biol., 6, 249, 1993.
200. Stuart, J. J. and Hatchett, J. H., Cytogenetics of the hessian fly. I. Mitotic karyotype analysis and polytene
chromosome correlations, J. Hered., 79, 184, 1988.
201. Stuart, J. J. and Hatchett, J. H., Cytogenetics of the hessian fly. 11. Inheritance and behaviour of somatic
and germ-line-limited chromosomes, J. Hered.. 79, 190, 1988.
202. Geyer-Duszyh, I., Experimental research and chromosome elimination in Cecidomyidae (Diptera).
Chromosoma, 11,499, 1959.
203. Kozlova, L. V., [On monogeny of gall midges, Aphidoletes aphidimyza Rond. (Diptera: Cecidomyidae)],
Nauchn. Tr. Leningr. Ord. Skh. Inst., No. 374, 11, 1979.
204. Went, D. F. and Camenzind, R., Sex determination in the dipteran insect Heteropeza pygmaea, Genetica,
52/53, 373, 1980.
Sex Determination in Insects 93

205. Went, D. F. and Camenzind, R., Haemolymph-dependent sex determination in a paedogenetic gall midge,
Natunvissenschafen, 64, 276, 1977.
206. Boyes, J. W., The chromosomes of Rhagionidae, Stratiomyidae and Xylomyidae (Diptera), Can. J. Genet.
Cytol., 15, 255, 1973.
207. Metz, C. W., Chromosome studies in the Diptera. IV. Incomplete synapsis in Dasyllis grossa. Biol. Bull.
(Wood's Hole, Mass.), 43, 253, 1922.
208. Mainz, F., The genetics of Megaselia scalaris Loew (Phoridae): a new type of sex determination in Diptera,
Am. Nut., 98, 415, 1964.
209. Willhoeft, U. and Traut, W., Molecular differentiation of the homomorphic sex chromosomes in Megaselia
scalaris (Diptera) detected by random DNA probes, Chromosoma. 99, 237, 1990.
210. Traut, W. and Willhaft, U., A jumping sex determining factor in the fly Megaselia scalaris, Chromosoma,
99,407, 1990.
21 1. Diibendorfer, A., Hiltiker-Kleiner, D., and Nothiger, R., Sex determination mechanisms in dipteran
insects: the case of Musca domestica. Semin. Dev. Biol.. 3, 349, 1992.
212. Franco, M. G., Rubini, P. G., and Vecchi, M., Sex-determinantsand their distributionin various populations
of Musca domestica L. of western Europe, Genet. Res., 40, 279, 1982.
213. Tomita, T. and Wada, Y., Multifactorial sex determination in natural populations of the housefly (Musca
domestica) in Japan, Jpn. J. Genet., 64, 373, 1989.
214. Jayakar, S. D., Some two-locus models for the evolution of sex-determining mechanisms, Theor. Popul.
Biol.. 32, 188, 1987.
215. Shono, T. and Scott, J. G., Autosomal sex-associated pyrethroid resistance in a strain of housefly (Diptera:
Muscidae) with a male-determining factor on chromosome three, J. Econ. Entomol., 83, 686, 1990.
216. Denholm, I., Franco, M. G., Rubini, P. G., and Vecchi, M., Identification of a male determinant on the X
chromosome of a housefly (Musca domestica L.) population in south-eastEngland, Genet. Res., 42.31 l, 1983.
217. Denholm, I., Franco, M. G., Rubini, P. G., and Vecchi, M., Geographical variation in house fly (Musca
domestica L.) sex determinants within the British Isles, Genet. Res., 47, 19, 1986.
218. Kerr, R. W., Inheritance of DDT resistance in a laboratory colony of the house fly, Musca domestica. Aust.
J. Biol. Sci., 23, 377, 1970.
219. Loeschke, V., Nielsen, B. 0..and Andersen, D., Chromosomal variation, segregation and sex determination
in Hydrotaea meridionalis (Diptera: Muscidae), Hereditas, 118, 229, 1993.
220. Samoylov, Yu. B., [Genetic control for the cabbage root fly 11. Localization of male determining factor in the
cabbage root fly Delia brassicae Bouche], Genetika, 21, 1810, 1985.
221. Inoue, H. and Hiroyishi, I., A maternal effect sex transformation mutant of the housefly Musca domestica,
Genetics, 12,469, 1986.
222. Maddern, R. H. and Bedo, D. G., Properties of the sex chromosomes of Lucilia cuprina deduced from
radiation studies, Genetica. 63, 203, 1984.
223. Bedo, D. G. and Foster, G. G., Cytogenetic mapping of the male-determining region of Lucilia cuprina
(Diptera: Calliphoridae), Chromosome, 92, 344, 1985.
224. Ribbert, D., Die Polyt;inchromosomender Borstenbildungszellen von Calliphoraeryrhrocephula, Chromosomu,
21, 296, 1967.
225. Ullerich, F.-H., Analysis of sex determination in the monogenic blowfly Chrysomya rujifacies by pole cell
transplantation, Mol. Gen. Genet., 193,479, 1984.
226. Clausen, S. and Ullerich, F.-H., Sequence homology between a polytene band in the genetic sex chromo-
somes of Chrysomya rufifacies and the daughterless gene of Drosophila melanogaster, Natunvissenschaften,
77, 137, 1990.
227. Lifschitz, E. and Cladera, J. L., Ceratitis capirata: cytogenetics and sex determination,in Fruit Flies. Their
Biology, Natural Enemiesand Control (World Crop Pests 38). Robinson, A. S. and Harper, G., Eds., Elsevier,
Amsterdam, 1989, 3.
228. Solferini, V. N. and Morgante, J. S., X,X,X,X,:X,X,Y mechanism of sex determination in Anastrepha
bistrigata and A. serpentina (Diptera: Tephritidae), Rev. Bras. Genet., 13, 201, 1990.
229. Grewal, J. S. and. Kapoor, V. C., Karyotypes of some fruitfly species (Tephritidae) of India, in Fruit Flies
of Economic Importance (CEC/IOBCInt. Symp.. Rome, 7-IOApril 1987), Cavallaro, R., Ed.,A. A. Balkema,
Rotterdam, 1989,237.
230. Bush, G. L., Female heterogamety in the family Tephritidae (Acalyptrata, Diptera),Am. Nar., 100, 119, 1966.
231. Anleitner, J. E. and Haymer, D. S., Y-Enriched and Y specific DNA sequences from the genome of the
Mediterranean fruit fly, Ceratitis capitata, Chromosomu, 101, 271, 1992.
232. Hodgkin, J., Drosophila sex determination: a cascade of regulated splicing, Cell, 56, 905, 1989.
233. Slee, R. and Bownes, M., Sex determination in Drosophila melanogaster, Q. Rev. Biol., 65, 175, 1990.
234. Torres, M. and Sanchez, L., The segmentation gene runt is needed to activate Sex-lethal, a gene that controls
sex determination and dosage compensation in Drosophila, Genet. Res.. 59, 189, 1992.
235. Steinmann-Zwicky, M., How do germ cells choose their sex? Drosophila as a paradigm, BioEssays. 14,513,
1992.
94 Insect Reproduction

236. Cline, T. W., The Drosophila sex determination signal: how do flies count to two?, Trends Genet., 9, 385,
1993.
237. Steinemann,M. and Steinemann,S., Evolutionary changes in the organization of the major LCP gene cluster
during sex chromosomal differentiation in the sibling species Drosophila persimilis, D. pseudobscura and D.
miranda, Chromosoma, 99, 424, 1990.
238. Ganguly, R., Swanson, K. D., Ray, K., and Krishnan, R., A BamHI repeat element is predominantly
associated with the degenerating NEO-Y chromosome of Drosophila miranda but absent in Drosophila
melanogaster genome, Proc. Natl. Acad. Sci. U.S.A., 89, 1340, 1992.
239. Steinemann, M., Steinemann, S., and Lottspeich, F., How Y chromosomes become genetically inert, Proc.
Natl. Acad. Sci. U.S.A., 90, 5737, 1993.
240. Ohno, S., Evolution of sex chromosomes in mammals, Annu. Rev. Genet., 3, 495, 1969.
241. Jablonka, E. and Lamb, M. J., The evolution of heteromorphic sex chromosomes, Biol. Rev. Camb. Philos.
Soc.. 65, 249, 1990.
242. Charlesworth,D. and Charlesworth, B., Sex differences in fitness and selection for certain fusions between
sex chromosomes and autosomes, Genet. Res., 35,205, 1980.
 Chapter 4

HORMONES AND REPRODUCTION

Jim Hardie

CONTENTS
I. Introduction .................................................................................................................
95

Hormones and Females ............................................................................................... 96

A. Oogenesis ..............................................................................................................96
1. Cockroaches ..................................................................................................... 96
2. Dipterans ......................................................................................................... 97
a. Mosquitoes .................................................................................................. 97
b. Flies .............................................................................................................
98
3. Hemipterans .....................................................................................................99
4. Homopterans ....................................................................................................99
B. Regulation of Sex Pheromone Release .............................................................. 101

111. Hormones and Males ................................................................................................ 101

A. Spermatogenesis.................................................................................................. 102
B. Sperm Release and Maturation ........................................................................... 103
C. Gonadal Development ........................................................................................ 103

IV. Summary ....................................................................................................................

103

Acknowledgments ...............................................................................................................
103

References ...........................................................................................................................
104

I. INTRODUCTION
This chapter concerns the endocrine control of reproduction in female and male insects.
The major gonadotropic hormones are juvenile hormones (JHs) and ecdysteroids, the same
hormones that control metamorphosis and moulting. This parsimony in hormone complement
occurs because the windows of sensitivity for development and reproduction are, to an extent,
isolated from each other and the temporal distribution of hormone receptors differs between
different tissues. Thus, during larval and pupal stages, JHs and ecdysteroids are responsible
for development, while in the adult (and in certain instances preadult) stages they take on
gonadotropic functions. Juvenile hormones are synthesized and released from the corpora
allata which are present in all insect stages. The corpora allata are possibly the only site of
synthesis, although recently male accessory glands have been reported to produce JH.' In
contrast, during larval and pupal stadia, ecdysteroids are synthesized and released, for the
main part, from the prothoracic glands, which, with certain exceptions (e.g., in Apterygota
which alternately moult and reproduce as adults and in solitary locusts), atrophy at the final
moult. In adult females, ecdysteroid synthesis occurs in the ovaries, or more specifically in the
follicle cells of the ovaries, and in males they are synthesized by the testes, while other

0-8493-6695-X/95/$0.MkSSM
O 1995 by CRC Press. Inc.
96 Insect Reproduction

possible sites for production have been ~uggested.~

With today's more sensitive techniques for
hormone isolation and identification together with molecular biological procedures, the con-
trol of JH and ecdysteroid titers and their actions at the gene level are becoming more
under~tood.~ Recent reviews of the hormonal control of insect reproduction include references
4 through 7.

11. HORMONES AND FEMALES

The endocrine control of reproduction in female insects varies with species; this is, perhaps,
not surprising as the reproductive strategies also differ (e.g., oviparous, ovoviparous, vivipa-
rous, sexual, parthenogenetic). The present chapter will concentrate on presenting some of the
different endocrine strategies with reference to recent investigations of some well-researched
insects. Chapter 1 describes the events of oogenesis with some details on endocrine effects on
female accessory glands.

A. OOGENESIS
1. Cockroaches
In all species of cockroaches investigated, including oviparous, ovoviparous, and vivipa-
rous species, JH has been shown to play a major gonadotropic role. Mated females of the
oviparous American cockroach, Periplaneta americana, undergo the cyclical production of
oothecae (egg cases containing up to 16 eggs) every 4 to 5 days. Removal of the corpora allata
from immature females prevents any previtellogenic ovarian development, while in mature
females it prevents further ootheca formation by inhibiting vitellogenin synthesis and ~ p t a k e . ~
Reimplantation of corpora allata or treatment with JH restores the reproductive c y ~ l e . ~The
.'~
precise corpora allata control of the cycle of ootheca production has been the subject of many
studies. In vitro culture of corpora allata under conditions that allow the synthesis of JH (JH
111 in this species) has shown a corresponding cycle of JH synthesis and release." More
recently, it has been shown that in vivo titers of JH I11 show concomitant cyclical rises, with
peak titers occurring during vitellogenesis and low titers at the point where oothecae are
deposited.I2Surgical techniques, more refined than complete allatectomy, have been imple-
mented and show that after unilateral allatectomy (removing one of the pair of corpora allata),
ootheca production continues, but at a slightly lower rate, while severing all the nervous
connections to the corpora allata had a similar effect.13 The corpora allata effects on ootheca
production could, therefore, be accommodated by the loss of one corpus allatum and, more
important, the cyclical activity appeared to be driven by hemolymph-borne factors. However,
the same study showed that the cyclical production of oothecae could be reinstated in
allatectomized females by treatment with a potent JH analogue which also speeded up the
ootheca cycle in intact insects. The study concluded that the ootheca production cycle was not
driven by the cycle of JH synthesis and release, but that JH sewed only to control the speed
of an endogenous reproductive cycle: lower JH levels slowed the cycle while higher levels
speeded it up; in the absence of JH the cycle stopped.13 Control of JH synthesis in vivo may
well be effected by peptide factors such as allatostatins (which inhibit JH synthesis by the
corpora allata), recently identified in the brain of the viviparous cockroach, Diploptera
punctata, and shown to be effective in P. americana.14
Ecdysteroids are produced by the ovaries of adult co~kroaches,~~ but their role has not been
fully elucidated. It has been proposed that the CO-occurrenceof ecdysteroid and choriogenesis
might indicate a role at this stage in egg development. They may have an inhibitory efect on
JH secretion, and their continued presence in the ootheca indicates that they supply the
requirements of the embryo^.'^.'^
Hormones and Reproduction

v+
Adult Eodysis Blood Meal

ma~ng--Err,l
brain
l
corpus allaturn
I -
behaviour
fat body O W
1
TMOF
i+
competent fat body resting W H
CCSF

t oocyts maturation
WTULOQENlN -----#-vit~ltOgOni~ OVW
l

FIGURE l. Factors regulating ovarian development in the mosquito, Aedes aegypti. CCSF = corpus cardiacum-
stimulating factor; MNC = medial neurosecretory cells: EDNH = egg development neurosecretory hormone;
TMOF = trypsin-modulating oostatic factor. (Modified from Hagedom, H. Comprehensive Insect Physiology, Bio-
chemistry and Pharmacology, Kerkut, G. and Gilbert, L., Eds. Pergamon, Oxford, 1985 page 165.)

2. Dipterans
In a number of Diptera, ecdysteroids replace JH as the major gonadotropic hormone
stimulating vitellogenin synthesistuptake.

a. Mosquitoes
The hormonal control of egg maturation in Aedes aegypti, an anautogenous mosquito
(requiring a blood meal to develop the first and subsequent egg batches) has been studied
inten~ively.~ A summary of the endocrine interactions is shown in Figure 1. Adult emergence
triggers the release of JH from the corpora allata.I8JH then acts on the ovary, which contains
the primary follicles, to stimulate growth to around twice the original size and form a "resting
stage" ovary which is complete within 3 days of emergence. This previtellogenic development
of the ovary also involves the production of endocytic organelles by the oocytes, which then
become competent for protein uptake.19The "resting stage" ovary now exerts an inhibition of
JH secretion,20but does not develop further until a blood meal has been taken. This initial rise
in JH titer also stimulates mating behaviorZ1and induces a competence in the fat body to
respond' to ecdysteroids. The fat body remains unresponsive if allatectomy is performed at
, ~ ~in vitro experiments confirm that JH promotes the competence required
adult e c l o s i ~ nand
for an ecdysteroid response.23
After a blood meal, egg development neurosecretory hormone (EDNH)" is released from
the corpora cardiaca, having been produced in the protocerebrum by the medial neurosecre-
tory cells, and acts on the "resting stage" ovary to induce the production of e ~ d y s o n eThis
.~~
EDNH release, also, involves a corpus cardiacum-stimulating factor (CCSF) which is pro-
duced by the resting o ~ a r y .Ecdysone
~ ~ . ~ ~is secreted into the hemolymph and converted to 20-
hydroxyecdysone, which then acts on the competent fat body to stimulate vitellogenin synthe-
sis and release. Additionally, the 20-hydroxyecdysoneinduces the separation of new follicles,
the secondary follicles, from the g e r m ~ i aControl
. ~ ~ of the endocytotic uptake of vitellogenin
by the oocytes has yet to be elucidated, but it is possible that both 20-hydroxyecdysone and
an unidentified brain hormone effect vitellogenin uptake.29After a short delay, blood meals
also induce a 24- to 36-h pulse of elevated JH titer which coincides with decreased levels of
98 Insect Reproduction

JH-esterase activity.I8 This meal-induced JH pulse stimulates the new follicles to develop to
the resting stage and a renewed competence of the fat body to respond to ecdysteroids. The
next blood meal reinitiates the egg maturation cycle. Further blood meals are usually taken
only after egg batches are laid. If taken before oviposition, the mature postvitellogenic oocytes
produce an oostatic hormone, providing an inhibitory feedback which prevents further ovarian
development and vitellogene~is.~~ This oostatic hormone has recently been characterized as a
decapeptide which inhibits synthesis of a trypsin-like enzyme in the midgut of female A.
aegypti; it thus prevents the blood-meal digestion and, indirectly, vitell~genesis.~~
Aedes atropalpus is an autogenous mosquito species which does not require a blood meal
to produce the first egg batch. Nevertheless, hormonal control of the ovarian cycle appears
similar to the anautogenous A. aegypti except that emergence is the stimulus for EDNH release
and development of the first egg batch.5 Indeed, peptides have now been isolated that induce
ecdysteroid release from ovaries (i.e., GDNH-like) of both species.32

b. Flies
Both JH and ecdysteroids are again implicated in the hormonal control of egg maturation
in the higher Diptera (Cyclorrhapha), and finer details are still being revealed. The major JH
produced by the corpus allatum of adult Cyclorrhaphan flies (Drosophila melanogaster,
Calliphora vomitoria) has recently been identified as the bisepoxide of JH III.33.34
In an anautogenous colony (requiring a protein meal before egg development) of the black
blowfly, Phormia regina, it appears that JH is required for the uptake of vitellogenin by the
~ o c y t e Treatment
.~~ of sugar-fed adult females with JH I11 or a JH analogue resulted in a
proportion of follicles with oocytes containing an opaque material. Immunological procedures
showed that this opaque material was not vitellogenin, and it was concluded that protein
uptake by the oocytes had been stimulated, but, in the absence of vitellogenin, the opaque
material comprised other hemolymph proteins. Sugar-fed flies also retained low levels of
ecdysteroids, but after a protein meal, ecdysteroids increased, the ovaries being the major
source, and vitellogenin uptake by the oocytes followed.36Previous experiments had shown
that treatment with precocene I1 (a pro-allatocidin compound which prevents JH synthesis by
the corpus allatum) inhibited oocyte development beyond the previtellogenic stage, but did not
prevent vitellogenin synthesis or release into the hemolymph, albeit levels were lower than
controls.37Recently it has been shown that a peptide factor from the midgut is released in liver-
fed females, which stimulates neurosecretory cells in the brain to initiate the neuroendocrine
cascade leading to o o g e n e ~ i sA. ~summary
~ of the proposed endocrine interaction for this fly
species is presented in Figure 2 and includes a possible factor released from the brain which
induces synthesis and release of ecdysteroids from the ovaries (modified from References 36
and 38).
It has been shown in other flies that both the fat body and the ovaries produce ~ i t e l l o g e n i n . ~ ~ . ~ ~
In Drosophila, genes for three vitellogenins (termed yolk proteins 1,2, and 3) are expressed
in the fat body and follicuiar cells of the ovary but the regulation of expression differs between
the tissues. Experiments with ligated abdomens showed that ecdysteroids and JHs could
induce vitellogenin formation but, in addition, JH promoted vitellogenin uptake by the
oocytes. Further studies on the relative amounts of the three vitellogenins indicated that JH
stimulated normal synthesis and uptake in the ovary but abnormal synthesis by the fat body,
while ecdysteroids had no effect on the ovary but induced normal synthesis by the fat body.5
This simplistic model may have to be modified in view of more recent work which indicates
the presence of other factors mediating nutritional effects on vitellogenin production and
~ p t a k e . ~By
~ .comparison,
~' in the housefly Musca domestica, ecdysteroids and a JH analogue
have been shown to induce vitellogenin mRNA in both the fat body and the ovaries. However,
the JH analogue was the least potent of the two and proved more effective on the ovary than
on the fat body.40
Hormones and Reproduction

Protein Meal

corpus wdiacum corpus allaturn

l

FIGURE 2. Factors regulating ovarian development in the black blowfly,Phormia regina. (Modifiedfrom Yin et a1.S3&)

3. Hernipterans
The blood-sucking bug Rhodnius prolixus was used by Wigglesworth to identify JH as a
hormone controlling both metamorphosis and reproducti~n.~~ He demonstrated that the re-
moval of the corpus allatum prevented vitellogenesis in the ovaries of adult females. This
insect has continued to be utilized for the examination of reproductive endocrinology, particu-
larly by Davey and colleagues, and our knowledge of humoral events controlling ovarian
development and oviposition has become increasingly detailed. JH probably stimulates
vitellogenin synthesis in the fat body, but the study of vitellogenin uptake by the oocytes has
taken research ~ r e c e d e n c eIn
. ~mated
~ females, the JH has a three-fold action in the stimulation
of vitellogenin uptake by the terminal oocytes. Vitellogenins are large protein molecules, and
their access to the oocyte cell membrane is facilitated by JH-induced changes in shape of the
surrounding follicle cells and the resulting enlargement in the intercellular spaces (increased
"patency"; see Chapter However, the presence of JH is also necessary during follicle cell
development in order that cells become competent to respond to later increases in JH t i t e ~ - . ~ ~
The uptake of vitellogenin by the oocyte is a calcium-dependent receptor-mediated response,
and the receptor binding of vitellogenin is also enhanced by JH.46Recent studies also provide
evidence of a neural inhibition of corpus allatum activity coming from the brain.47
In addition to the gonadotropic effects of JH, Rhodnius females possess abdominal neuro-
secretory organs which produce an antigonadotropic hormone (oostatic h ~ r m o n e ) .Ovaries ~~.~~
containing mature oocytes in the pedicels stimulate the release of the antigonadotropin, which
counters the effect of JH on the vitellogenin uptake by the oocytes. The changes in patency
induced by JH are inhibited in the presence of antigonadotr~pin.~~
Ten neurosecretory cells have been identified in the pars intercerebralis of the brain, which
produce a peptidergic myotropic factor that initiates ovulation (movement of the fully devel-
oped oocyte from the ovary) and subsequently o v i p o s i t i ~ n . Unmated
~ ~ . ~ ~ females retain eggs
in the ovarian pedicel for long periods, resulting in a much lower oviposition rate than mated
females, and it is the presence of a spermathecal factor in mated insects that facilitates the
release of the oviposition hormone.53However, it appears to be the production of ecdysteroids
by the ovary, 5 days after a blood meal, that induces the release of the myotropic ovulation
hormone from axon terminals in the corpora ~ a r d i a c a .The ~ ~ ecdysteroids
-~~ do not act directly
on the neurosecretory cells but indirectly via aminergic neuronss7while more recent investiga-
tions indicate that there is also a circadian influence over these endocrine events.58

4. Homopterans
Aphids form one of the few groups of insects that commonly alternate between partheno-
genetic and sexual reproduction (cyclical parthenogene~is).~~
While there are species (andlor
100 Insect Reproduction

clones) that will reproduce only by parthenogenesis, there are none that reproduce solely by
sexual means. In a number of species, both the mode of reproduction and ovarian development
in the parthenogenetic females is is controlled by the same hormone, JH. Photoperiodic cues
are most frequently found to determine the reproductive type: long days result in the formation
of parthenogenetic females (virginoparae), while short days promote the development of the
sexual forms (oviparae and male^).^
Work in the late 1970s indicated that the medial neurosecretory cells of the protocerebrum
(called the group I cells in aphid^)^' were involved in the determination of female embryos as
virginoparae or o ~ i p a r a eAblation
.~~ of these cells by radiocautery, in long-day reared vetch
aphids, Megoura viciae, induced the production of "short-day" oviparae rather than the
expected virginoparae. The observations indicated that a factor produced by these protocerebral
cells promoted the appearance of virginoparae, and it was proposed that the factor acted
directly upon the developing embryos in the ovarioles. The factor was termed "virginoparin"
by In contemporary studies, topical application of natural juvenile hormones or JH
analogues was found to modify embryonic development such that female embryos that were
photoperiodically determined as sexual females (oviparae) were redirected to develop as
parthenogenetic females (~irginoparae).~"-" In addition to the switch induced in ovary type,
there was a concommitant induction of winged or partially winged forms, many of which were
sterile, but judicious application of JH produced seemingly normal virginoparous females that
were capable of normal reprod~ction."~~' Thus, "virginoparin" and JH had the same effect on
the development of female embryos and promoted their development as virginoparae; they
were "long-day" hormones.
It was later shown that "virginoparin" and JH were part of the endocrine pathway for the
induction of ~irginoparae.~~ It was shown that corpus allatum volume (this is a single fused
gland in the aphid) differed, from day 10 onwards, between short-day- and long-day-reared
females that were producing oviparae or virginoparae, respectively. When the group I cells
were ablated in long-day insects, and the progeny switched from virginoparae to oviparae,
there was a concomittant change in the corpus allatum volume such that it became equal in
size to the short-day-reared insect corpora allata. Aphids that underwent sham cautery re-
mained similar to untreated controls; the corpora allata were the same size, and they continued
to produce virginoparae. It appears that the group I cells regulate JH synthesis via the corpus
allatum. Assessment of JH titers in the aphids showed only JH I11 present and, although the
levels were low, there were higher JH titers in the long-day- than the short-day reared aphids.69
There are other subtleties to aphid reproduction in that the parthenogenetic females un-
dergo precocious ovarian development which results in the telescoping of generations.'O The
two ovaries of adult virginoparae (mother) comprise a number of ovarioles containing em-
bryos (daughters) in various stages of development. The most mature of these embryos already
contain ovaries with developing embryos (granddaughters). Parthenogenetic aphids are also
viviparous and give birth to live young which, at birth, already have ovaries with ovarioles that
contain one or two developing embryos. In the black bean aphid Aphis fabae, oocyte devel-
opment and embryogenesis (of the granddaughter generation) within the embryonic (daughter
generation) ovaries begins during the mother's fourth in~tar.~' Decapitation, which included
the removal of the corpora allata, drastically reduced the rate of oocyte differentiation, but this
could be restored by JH application. In addition, the growth of the terminal (daughter)
embryos is reduced almost completely by decapitation, but again is restored by JH treatment.
However, this stimulation of embryogenesis is not reflected in a shortening of the pre-
reproductive period or in the initial rate of reproduction, possibly because other endocrine
factors are involved in part~rition.~~
It appears that JH has both a role in determining the parthenogenetic aphid morph and in
the regulation of ovarian development in that morph. Additionally, as in other insects, it
regulates metarnorph~sis.'~ However, in most insects the metamorphic role of JH terminates
Hormones and Reproduction 101

at the final moult, when JH often takes on a reproductive role. Perhaps uniquely in aphids,
adult form is determined by low JH titers in the third (penultimate) instar larva and JH can then
take on a reproductive role, precociously, in the last larval instar. Control of ovarian devel-
opment in the sexual females has not been researched, but it will differ. Oocyte differentiation
in the germarium occurs later, in the second larval instar (rather than prenatally as in
parthenogenetic forms), with vitellogenesis following throughout the later instars, but
embryogenesis requires mating, fertilization,and a period of diapause development. Presump-
tive subunits of vitellogenin are also produced by parthenogenetic females, the ovaries of
which produce embryos, not vitellogenic eggs.74

B. REGULATION OF SEX PHEROMONE RELEASE

The efficiency of mate location by male insects is increased dramatically by the production
of species-specific, volatile cues that can be detected at a distance and used directionally as
well as on contact.75The production/release of these sex pheromones is intermittent, and
recent studies indicate a precise endocrinological control. Barth76used behavioral techniques
to demonstrate that allatectomy prevented sex pheromone release in some cockroaches, but in
other species had no effect. This study formed the basis for the suggestion of neuroendocrine
regulation of "calling" (pheromone release) behavior in species with a long adult life span and
which mated repeatedly, but with periods of unreceptivity, while there would be no such
control in species with a short, nonfeeding adult stage where mating occurs at emergence.
Evidence is accumulating that JH may be crucial for both pheromone synthesis and release in
the German cockroach, Blattella gerrnanica, which mates repeatedly,77.78and the brown-
banded cockroach, Supella longipalpa, which mates only once but lives for several months.79
JH also appears to regulate sex pheromone production in the female true armyworm, Pseudaletia
unipunctata, which mate after a period of migration, as well as in male responsiveness to
fern ale^.^^.^^ At least in the case of the female moth, the action of JH is indirect via a brain
factor, possibly via pheromone-biosynthesis-activating peptide (PBAN) (see below).
In a number of lepidopterans, a PBAN has been located in the brain-suboesophageal
ganglion complex, first in the corn earworm, Helicoverpa (Heliothis) z e ~ . The ~ * peptide has
now been shown to comprise a 33-amino acid sequence in H. zea,g3while Kitamura et a1.84.85
have identified two other PBANs, of similar size, in the silkworm, Bombyx mori. An 18-amino
acid peptide from the common armyworm, Pseudaletia (= Leucania) separata, showed
similar pheromone synthesis effects.86The latter peptide shares a common Phe-Ser-Pro-Arg-
Leu-NH2 terminal sequence with the identified PBAN molecules, but also induces cuticular
melanization and acts as a weak diapause hormone.86The PBAN family of peptides requires
further elucidation, and cross-reactivity between different bioassay systems raises questions
about the primary actions in vivo.86.87 It is also of interest that the PBAN from H. zea stimulates
pheromone biosynthesis in the Hessian fly, Mayetiola destructor (a cecidomyiid dipteran),
and PBAN-like activity has been reported in the cockroach, B. germanica, and the locust,
Locusta m i g r ~ t o r i a . In
~ ~the
. ~ housefly,
~ Musca domestica, however, ecdysteroids are impor-
tant not only for vitellogenesis (see above) but also for pheromone p r o d u c t i ~ n .In~ ~this
species, sex pheromone production can be artificially induced in males by ecdysteroid treat-
ment, while JH has no effect on pheromone production by either sex.

111. HORMONES AND MALES

The physiological control of spermatogenesis in males has been less well studied than has
oogenesis in females. This is partly because mature sperm are often present at eclosion of adult
males, and spermatogenesis has occurred under the hormonal conditions required for meta-
morphosis of the somatic cells?l Indeed, experiments on R. prolixus indicated precisely this
and that spermatogenesis continued autonomously, at a basal rate, in the adult under conditions
102 Insect Reproduction

of low JH and ecdysteroid~.~~ Spermatogenesis involves the mitotic cell divisions necessary
for the development of fully formed spermatocytes, followed by meiotic division and differ-
entiation of the sperm (spermiogenesis; see Chapter 2). The hormonal regulation of events is
not fully understood, and evidence is often contradictory. A number of contradictions un-
doubtedly occur as some studies focus upon the resumption of testis development after a
period of interrupted development (which can include spermatocyte autolysis) during diapause,
while others tackle normal development under nondiapausing conditions where spermatoge-
nesis is often arrested during meiotic prophase in early larval insects, only being resumed in
late larval (or later) stages.93There is no reason to assume that endocrine events leading to
renewed development are identical. In lepidopterans and some other species, two types of
sperm are produced (see also Chapter 2) -eupyrene (nucleate) sperm which fertilize the egg
and apyrene (nonnucleate) sperm whose function is not known.94Earlier work tended to look
for effects of JH and the results are again contradi~tory,4-~.~~ but more recent investigations
have tended to concentrate on the effects of ecdysteroids.
The testes of male insects, like the ovaries of females, have been shown to produce
ecdysteroids. Although their contribution to overall titers is minimal, actions at a more local
level could be i m p ~ r t a n t ? A
~ -brain
~ ~ ecdysiotropic factor is also reported to induce ecdysteroid
secretion by testes.99

A. SPERMATOGENESIS
Spermatocysts from diapausing male silkmoth pupae can be stimulated to develop by
ecdysteroids in vitro, but this effect is indirect and primes the cysts to respond to a large
molecular weight chemical (macromolecular factor), found in pupal hemolymph, by increas-
ing the sheath permeability.lOO,lO1 Similar observations have been made on the codling moth,
Cydia pomonella, but the response was irregular and incomplete,102and ultrastructural inves-
tigations of ecdysteroid effects on the testicular sheath did not indicate an increased perme-
ability, but an increased metabolic activity and glycogen accumulation.103In the latter insect,
JH appears to counter the effects of ecdysteroid on spermatogenesis, and topically applied JH
analogue induced spermatogenesis arrest in nondiapause-destined larvae, while allatectomy
renewed spermatogenesis in diapause-destined larvae."'" In the cabbage armyworm, Mamestra
brassicae, ecdysteroid levels are 4.4 times higher in testes of nondiapausing than diapausing
pupae. CO-culture of testis and spermatocysts, both from diapausing pupae, resulted in
spermiogenesis and correlated with raised ecdysteroids in culture medium, but proteins are
also released from the cultured testes and may play a part in spermiogenesis.lo5
In nondiapausing tobacco budworm larvae, Heliothis virescens, the initiation of meiosis in
young eupyrene spermatocysts occurred in vitro, apparently without ecdysteroids, but requir-
ing fetal calf serum (possibly providing a macromolecular-likefactor) and testis sheath.Io6The
testis sheath factor was not an ecdysteroid nor was it species specific, as reciprocal stimulation
occurred with gypsy moth (Lymantria dispar) testis sheath.lo6However, in common army-
worm, P. (=Leucania) separata, spermatogenesis may be autonomous, since no factors have
been found to affect s p e r m i ~ g e n e s i s . ' ~ ~ ~ ~ ~ ~
In nondiapausedestined tobacco hornworm, Manduca sexta, eupyrene spermatocytes remain
in meiotic prophase from early larval stages to the last larva or pupa. Coincident with the
postwandering ecdysteroid peak, they recommence development to meiotic meta~hase.9~ This
effect can be mimicked in isolated abdomens by 20-hydroxyecdysone injections but not in vitro,
indicating the ecdysteroid effect may be indirect. In the testes of diapausing pupae, ecdysteroid
titers remain low, while in nondiapausing pupae they rise to a peak around day 10,returning to low
levels before adult ecl0sion.9~During diapause, mitosis and spermatocytedevelopment continues,
but apyrene spermatogenesis is arrested with the lysis of premeiotic apyrene spermatocytes, and
no apyrene spermatids are found. On the other hand, eupyrene meiosis precedes, but is followed
by lysis after differentiation of the eupyrene spermatids. It appears that the lack of ecdysteroids in
diapausing pupae allows partial spermatogenesisbut leads to resorption of the products at different
Hormones and Reproduction 103

stages of apyrene and eupyrene spermatogenesis. In the European corn borer, Ostrinia nubilalis,
apyrene spenniogenesis is directly stimulated by ecdysone and 20-hydroxyecdysone in vitro.

B. SPERM RELEASE AND MATURATION

In Lepidoptera, the release of clonal sperm bundles from the testes into the upper vas
deferens occurs in the pharate adult, the release being a cyclical, gated event under circadian
c ~ n t r o l .l ~l In J Mediterranean flour moth Ephestia kiihniella, release of eupyrene, but not
~ ~the
apyrene, sperm bundles from the testes was inhibited by injection of 20-hydro~yecdysone.~~~
More recent studies on the gypsy moth L. dispar indicate that the release of mature eupyrene
sperm is delayed between day 5 of pupation (when the reproductive tract and sperm are fully
differentiated) and day 9 by high titers of circulating ecdysteroids.l13Infusion or injection of
20-hydroxyecdysone further delayed the onset of sperm release in a dose-dependent fashion
but did not interfere with the rhythm or gating of release. In addition, exogenous ecdysteroids
did not inhibit the release of sperm once the rhythm had been initiated.Il3Thus it appears that
the the decline in endogenous ecdysteroid levels on day 7 after pupation are the cue for sperm
release to begin, but they do not control the cyclical release. The disparity between the
ecdysteroid inhibition of sperm release, after cyclical release had started, in E. kiihniella but
not L. dispar, may be due to differences in doses used.Il3
In the male silkmoth B. mori, apyrene sperm are present as individual cells in the seminal
vesicles while eupyrene sperm remain in bundles. Prior to spermatophore formation in the
ejaculatory duct, the apyrene sperm become mobile while the eupyrene sperm undergo
maturation, separation to single cells, and activation in the spermatophore. These events are
evoked by an endopeptidase, initiatorin, secreted by cells in the ejaculatory duct."4 The
precise regulation of this enzymic activation remains to be elucidated.

C. GONADAL DEVELOPMENT
Testis fusion occurs in the last larval instar of lepidopterans, but not in isolated abdomens
with low ecdysteroid titers. Testis fusion has been found to require ecdy~teroids?~J~~ In
addition, the development of larval spermducts into the seminal vesicles and upper vas
deferentia in the pupa requires e~dysteroids."""~However, this may be an indirect effect via
factors produced by the fat body and testis sheath, as isolated spermducts fail to develop in
the presence of e c d y ~ t e r o i d sThe
. ~ ~role
~ of ecdysteroids in regulating the development of male
accessory glands has recently been reviewed by Happ7 (see also Chapter 2).

IV. SUMMARY
The examples of hormonal regulation of reproduction in insects demonstrate the wide
variation in control mechanisms. This variation occurs even between species with similar life
strategies, e.g., blood-feeding mosquitoes and Rhodnius. The same two major hormones
appear to be involved in all insect species so far investigated, but their roles differ (e.g.,
ecdysteroids stimulates vitellogenin synthesis in Diptera but the release of a myotropin in
Rhodnius, while JH stimulates vitellogenin synthesis and uptake in many other insects). The
idea of a unified mechanism of endocrinological control in females or males may not be
feasible. However, some of the apparent differences in hormonal control of oogenesis between
Dipteran species may be due to variation in endogenous hormone titers and the timing of
experiments.l2ICurrent studies of insect reproduction are tending to concentrate on the role
of neurosecretory peptides and the molecular biology of hormone synthesis and action.

ACKNOWLEDGMENTS
I thank the Biotechnology and Biological Sciences Research Council for financial support
and Stuart Reynolds and Rob Storer for comments on an earlier draft.
 Insect Reproduction

REFERENCES
1. Borovsky, D., Carlson, D.A., Hancock, R.G., Rembold, H., and van Handel, E., De novo synthesis of
juvenile hormone 111 and I by the accessory glands of the male mosquito, Insect Biochem. Molec. Biol., 24,
437, 1994.
2. Delbecque, J.-P., Weidner, K., and Hoffmann, K.H., Alternative sites for ecdysteroid production in insects,
Invertebr. Reprod. Dev., 18, 29, 1990.
3. Wyatt, G.R., Gene regulation in insect development, Invertebr. Reprod. Dev., 20, 1, 1991.
4. Koeppe, J.K., Fucbs, M., Chen, T.T., Hunt, L.-M., Kovalick, G.E., and Briers, T., The role of juvenile
hormone in reproduction, in Comprehensive Insect Physiology, Biochemistry and Pharmacology, Kerkut,
G.A. and Gilbert, L.I., Eds., Pergamon, Oxford, Vol. 8, 1985, 165.
5. Hagedorn, H.H., The role of ecdysteroids in reproduction, in Comprehensive Insect Physiology, Biochem-
istry and Pharmacology, Kerkut, G.A. and Gilbert, L.I., Eds., Pergamon, Oxford, Vol. 8, 1985, 205.
6. Raabe, M., Insect reproduction, Adv. Insect Physiol., 19, 29, 1986.
7. Happ, G.M., Maturation of the male reproductive system and its endocrine regulation, Annu. Rev. Entomol..
37, 303, 1992.
8. Bell, W.J., Dual role ofjuvenile hormone in the control of yolk formation in Periplaneta americana. J. Insect
Physiol., 15. 1279, 1969.
9. Chen, D.H., Robbins, W.E., and Monroe, R.E., The gonadotropicaction of Cecmpia extracts in allatectomised
American cockroaches, Experientia, 18, 577, 1962.
10. Girardie, A., ~ t u d ebiometrique de ia croissance ovarienne ap&s ablation et implantation de corpora allata
chez Periplaneta americana, J. Insect Physiol.. 8, 199, 1962.
I l . Weaver, RJ., Pratt, G.E., and Finney, J.R., Cyclic activity of the corpus allatum related to gonadotrophic
cycles in adult female Periplaneta americana, Experientia, 31, 597, 1975.
12. Edwards, J.P., Chambers, J., Short, J.E., Price, N.R., Weaver, RJ., Abraham, L., and Walter, C.M.,
Endogenous juvenile hormone 111 levels and in vitro rates of hormone biosynthesis by corpora allata during
the reproductive cycle of adult female Periplaneta americana, in Chromatography and Isolation of Insect
Hormones and Pheromones. McCaffery A.R. and Wilson, I.D., Eds. Plenum Press, New York, 1990, 3.
13. Weaver, R.J. and Edwards, J.P., The role of the corpora allata and associated nerves in the regulation of
ovarian cycles in the oviparous cockroach Periplaneta americana. J. Insect Physiol.. 36, 51, 1990.
14. Weaver, RJ., Profile of the responsiveness of corpora allata from virgin female Periplaneta americana to
an allatostatin fom Diploprera punctata, J. Insect Physiol., 37, 11 l, 1991.
15. Weaver, R.J., Strambi, A., and Strambi, C., The significance of free ecdysteroids in the haemolymph of
adult cockroaches, J. Insect Physiol., 30, 705, 1984.
16. Chiang, A.-S., Burns, E.L., and Schal, C., Ovarian regulation of cyclic changes in size and activity of corpus
allatum cells in Blattella germanica, J. Insect Physiol.. 37, 907, 1991.
17. Pascual, N., Cerdi, X., Benito, B., Tomb, J., Piulachs, M.D., and BellCs, X., Ovarian ecdysteroid levels
and basal oocyte development during maturation in the cockroach Blattella germanica (L.), J. Insect Physiol.,
38, 339, 1992.
18. Shapiro, A.B., Wheelock, G.B., Hagedorn, H.H., Baker, F.C., Tsai, L.W., and Schooley, D.A., Juvenile
hormone and juvenile hormone esterase in adult females of the mosquito Aedes aegypti, J. Insect Physiol., 32,
867, 1986.
19. Raikhei, A.S. and Lea, A.O., Hormone-mediated formation of the endocytotic complex in mosquito oocytes,
Gen. Comp. Endocrinol.. 57, 422, 1985.
20. Rossignol, P.A., Feinsod, F.M., and Speilman, A., Inhibitory regulation of corpus allatum activity in
mosquitoes, J. Insect Physiol., 27, 651, 1981.
21. Gwadz, R.W., Lounibos, L.P., and Graig, G.B., Precocious sexual receptivity induced by a juvenile
hormone analog in females of the yellow fever mosquito, Aedes aegypti, Gen. Comp. Endocrinol.. 16, 47,
1971.
22. Flanagan, T.R. and Hagedorn, H.H., Vitellogenin synthesis in the mosquito: the role of juvenile hormone
in the responsiveness to ecdysone, Physiol. Entomol., 2, 173, 1977.
23. Ma, M., Zhang, J.-Z., Gong, H., and Gwadz, R., Permissive action of juvenile hormone on vitellogenin
production by the mosquito, Aedes aegypti, J. Insect Physiol., 34, 593, 1988.
24. Lea, A.O., The medial neurosecretory cells and egg maturation in mosquitoes, J. Insect Physiol., 13, 419,
1967.
25. Hagedorn, H.H., Shapiro, J.P., and Hanaoka, K., Ovarian ecdysone secretion is controlled by a brain
hormone in an adult mosquito, Nature. 282,92, 1979.
26. Borovsky, D., Release of the egg development neurosecretory hormone in Aedes aegypti and Aedes
taeniorhynchus induced by an ovarian factor, J. Insect Physiol.. 28, 31 1, 1982.
27. Lea, A.O. and Van Handel, E., A neurosecretory hormone-releasing factor from ovaries of mosquitoes fed
blood, J. Insect Physiol.. 28, 503, 1982.
Hormones and Reproduction 105

28. Beckermeyer, E.F. and Lea, A.O., Induction of follicle separation in the mosquito by physiological amounts
of ecdysterone, Science, 209, 819, 1980.
29. Koller, C.N. and Raikhel, A.S., Initiation of vitellogenin uptake and protein synthesis in the mosquito (Aedes
aegypti) ovary in response to a blood meal, J. Insect Physiol.. 37, 703, 1991.
30. Else, J.G. and Judson, C.L., Enforced egg-retention and its effects on vitellogenesis in the mosquito Aedes
aegypti, J. Med. Entomol., 9, 527, 1972.
31. Borovsky, D., Carlson, D.A., Griffin, P.R., Shabanowitz, J., and Hunt, D.F., Mass spectrometry and
characterisation of Aedes aegypti trypsin modulating oostatic factor (TMOF) and its analogues, lnsect
Biochem. Molec. Biol., 23, 703, 1993.
32. Matsumoto, S., Brown, M.R., Suzuki, A., and Lea, A.O., Isolation and characterization of ovarian
ecdysteroidogenic hormones from the mosquito, Aedes aegypti, lnsect Biochem., 19, 651, 1989.
33. Altaratz, M., Segal, D., Richard, D.S., Gilbert, L.I., and Applebaum, S.W., Juvenile hormone production
by wild type and apterous4mutant Drosophila melanogaster corpora allata in vitro, in Insect Neurochemistry
and Neurophysiology 1989, Borkovek, A.B. and Masler, E.P., Eds., Humana Press, Clifton, NJ, 1990, 333.
34. Duve, H., Thorpe, A., Yagi, KJ., Yu, C.G., and Tobe, SS., Factors affecting the biosynthesis and release
of juvenile hormone bisepoxide in the adult blowfly Calliphora vomitoria, J. Insect Physiol., 38,575, 1992.
35. Stoffolano, J.G., Li, M.-F., Zou, B.-X., and Yin, C-M., Vitellogenin uptake, not synthesis, is dependent on
juvenile hormone in adults of Phormia regina (Meigen), J. lnsect Physiol., 38, 839, 1992.
36. Yin, C.-M., Zou, B.-X., Yi, S.-X., and Stoffolano, J.G., Ecdysteroid activity during oogenesis in the black
blowfly, Phormia regina (Meigen), J. Insect Physiol., 36, 375, 1990.
37. Yin, C.-M., Zou, B.-X., and Stoffolano, J.G., Precocene I1 treatment inhibits terminal oocyte development
but not vitellogenin synthesis and release in Phormia regina (Meigen), J. Insect Physiol.. 35, 465, 1989.
38. Yin, C.-M., Zou, B.-X., Yi, M.-F. and Stoffolano, J.G., Discovery of a midgut peptide hormone which
activates the endocrine cascade leading to oogenesis in Phormia regina (Meigen), J. lnsect Physiol., 40,283,
1994.
39. Bownes, M. and Reid, G., The role of the ovary and nutritional signals in the regulation of fat body yolk
protein gene expression in Drosophila rnelanogaster, J. Insect Physiol., 36,471, 1990.
40. Agui, N., Shimadu, T., Izumi, S., and Tomino, S., Hormonal control of vitellogenin mRNA levels in the
male and female housefly, Musca domestica, J. Insect Physiol., 37, 383, 1991.
41. Bownes, M., The role of juvenile hormone, ecdysone and the ovary in the control of Drosophila vitellogen-
esis, J. Insect Physiol., 35, 409, 1989.
42. Wigglesworth, V.B., The function of the corpus allatum in growth and reproduction of Rhodnius prolixus,
Q. J. Microsc. Sci., 79, 91, 1936.
43. Davey, K.G., Hormonal control of vitellogenin uptake in Rhodnius prolixus Stil. Am. Zool., 21,701, 1981.
44. Pratt, G.E. and Davey, KG., The corpus allatum and oogenesis in Rhodniusprolixus (Stil.). Ill. The effects
of allatectomy, J. Exp. Biol., 56, 201, 1972.
45. Abu-Hakima, R. and Davey, K.G., Two actions of juvenile hormone on the follicle cells of Rhodnius
prolixus StSI., Can. J. Zool., 53, 1187, 1975.
46. Wang, Z. and Davey, K.G., Characterization of yolk protein and its receptor on the oocyte membrane in
Rhodnius prolixus, lnsect Biochem. Molec. Biol., 22, 757, 1992.
47. Davey, KG. and Chiang, R.G., The effect of severing the doral vessel on egg production in Rhodnius
prolixus, Arch. lnsect Biochem. Physiol., 11, 139, 1989.
48. Heubner, E. and Davey, K.G., An antigonadotropin from the ovaries of the insect Rhodnius prolixus StSI.,
Can. J. Zool., 51, 113, 1973.
49. Davey, KG. and Kuster, J.E., The source of an antigonadotropin in the female of Rhodniusprolixus Still.,
Can. J. Zool., 59, 761, 1981.
50. Davey, K.G. and Heubner, E., The response of follicle cells of Rhodnius prolixus to juvenile hormone and
antigonadotropin in vitro, Can. J. Zool., 52, 1407, 1974.
51. Kriger, F.L. and Davey, K.G., Identified neurosecretory cells in the brain of female Rhodnius prolixus
contain a myotropic peptide, Can. J. Zool., 62, 1720, 1984.
52. Davey, K.G. and Kriger, F.L., Variations during the gonotrophic cycle in the titer of the myotropic ovulation
hormone and the response of the ovarian muscles in Rhodnius prolixus. Gen. Comp. Endocrinol.. 58, 452,
1985.
53. Davey, K.G., Copulation and egg production in Rhodniusprolixus: the role of the spermathecae,J. Exp. Biol.,
42, 373, 1965.
54. Ruegg, R.P., Kriger, F.L., Davey, K.G., and Steel, C.G.H., Ovarian ecdysone elicits release of a myotropic
ovulation hormone in Rhodnius (Insects: Hemiptera), Int. J. Inverrebr. Reprod., 3, 357, 1981.
55. Ruegg, R.P., Orchard, I., and Davey, K.G., 20-Hydroxy-ecdysoneas a modulator of electrical activity in
neurosecreory cells of Rhodnius prolixus, J. Insect Physiol., 28, 243, 1982.
56. Davey, KG. and Prasher, A.K., Increased synthesis of proteins is closely coupled to release in the myotropic
neurosecretory cells of Rhodnius prolixus. Insect Biochem., 20,215, 1990.
106 Iiuect Reproduction

57. Orchard, I., Ruegg, R.P., and Davey, K.G., The role of central aminergic neurons in the action of 20-
hydroxy-ecdysone on neurosecretory cells of Rhodnius prolixus, J. Insect PhysioL, 29, 387, 1983.
58. Ampleford, E.J. and Davey, K.G., Egg laying in the insect Rhodniusprolixus is timed in a circadian fashion,
J. Insect Physiol., 35, 183, 1989.
59. Dixon, A.F.G., Aphid Ecology. Blackie, Glasgow, 1985.
60. Lees, A.D., The control of polymorphism in aphids, Adv. Insect Physiol., 3, 207, 1966.
61. Johnson, B., A histological study of neurosecretion in aphids, J . Insect Physiol., 9, 727, 1963.
62. Steel, C.G.H. and Lees, A.D., The role of neurosecretion in the photoperiodic control of polymorphism in
the aphid Megoura viciae. J. Exp. Biol., 67, 117, 1977.
63. Steel, C.G.H., Neurosecretory control of polymorphism in aphids, in Phase and Caste Determination in
Insects, Endocrine aspects, Liischer, M., Ed., Pergamon, Oxford, 1976, 117.
64. Mittler, T.E., Nassar, S.G., and Staal, G.B., Wing development and parthenogenesis induced in progenies
of kinoprene-treated gynoparae of Aphis fabae and Myzus persicae, J. Insect Physiol.. 22, 1717, 1976.
65. Hardie, J., Juvenile hormone and photoperiodically controlled polymorphism in Aphis fabae: prenatal effects
on presumptive oviparae, J. Insect Physiol., 27, 257, 1981.
66. Hardie, J. and Lees, A.D., The induction of normal and teratoid viviparae by juvenile hormone and kinoprene
in two aphid species, Physiol. Entomol., 10, 65, 1985.
67. Corbitt, T.S. and Hardie, J., Juvenile hormone effects on polymorphism in the pea aphid, Acyrthosiphon
pisum. Entomol. Exp. Appl., 38, 131, 1985.
68. Hardie, J., The corpus allatum, neurosecretion and photoperiodically controlled polymorphism in an aphid.
J. Insect Physiol.. 33, 201, 1987.
69. Hardie, J., Baker, F.C., Jamieson, G.C., Lees, A.D., and Schooley, D.A., The identification of an aphid
juvenile hormone, and its titre in relation to photoperiod, Physiol. Entomol., 10, 297, 1985.
70. Hardie, J. and Lees, A.D., Endocrine control of polymorphism and polyphenism, in Comprehensive Insect
Physiology. Biochemistry and Pharmacology. Kerkut, G.A. and Gilben, L.I., Eds., Pergamon, Oxford, Vol.
8, 1985, 441.
71. Hardie, J., Juvenile hormone stimulation of oocyte development and embryogenesis in the parthenogenetic
ovaries of an aphid, Aphis fabae, Int. J. Invertebr. Reprod. Dev., 11, 189, 1987.
72. Hardie, J., Mallory, A.C.L., and Quashie-Williams, C.A., Juvenile hormone and host-plant colonization by
the black bean aphid, Aphis fabae, Physiol. Entomol., 15, 331, 1990.
73. Lees, A.D., The development of juvenile hormone sensitivity in alatae of the aphid Megoura viciae, J. Insect
Physiol.. 26, 143, 1980.
74. Ishikawa, H. and Matsuka, M., Hemolymph proteins of chestnut aphid, Lachnus tropicalis, Comp. Biochem.
Physiol., 748, 521, 1983.
75. Tamaki, Y., Sex pheromones, in Comprehensive Insect Physiology, Biochemistry and Pharmacology, Kerkut,
G.A. and Gilbert, L.I., Eds., Pergamon, Oxford, Vol. 9, 1985, 145.
76. Barth, R.H., Insect mating behavior: endocrine control of a chemical communication system, Science, 149,
882, 1965.
77. Schal, C., Burns, E.L., and Blomquist, GJ., Endocrine regulation of female contact sex pheromone
production in the Gennan cockroach, Blattella germanica, Physiol. Entomol., 15, 81, 1990.
78. Schal, C., Bums, E.L., Gadot, M., Chase, J., and Blomquist GJ., Biochemistry and regulation of
pheromone production in Blattella germanica (L.) (Dictyoptera, Blattellidae), Insect Biochem.. 21.73, 1991.
79. Smith, A.F. and Schal, C, Corpus allatum control of sex pheromone production and calling in the female
brown-banded cockroach, Supella longipalpa (F.) (Dictyoptera: Blattellidae), J. Insect Physiol., 36, 251,
1990.
80. Cusson, J. and McNeil, J.N., Involvement of juvenile hormone in the regulation of pheromone release
activities in a moth, Science, 243, 210, 1989.
81. Cusson, J., Yagi, K., Tobe, S.S., and McNeil, J.N., Identification of release products of corpora allata of
male and female armyworm moths, Pseudaletia unipuncm. J. Insect Physiol., 39, 775, 1993.
82. Raina, K.A. and Klun, J.A., Brain factor control of sex pheromone production in the female corn earworm
moth, Science, 225, 531, 1984.
83. Raina, K.A., Jaffe, H., Kempe, T.G., Blacher, R.W., Fales, H.M., Riley, C.T., Klun, J.A., Ridgway, R.L.,
and Hayes, D.K., Identification of a neuropeptide hormone that regulates sex pheromone production in
female moths, Science. 244, 796, 1989.
84. Kitamura, A., Nagasawa, H., Kataoka, H., Ando, T. and Suzuki, A., Amino acid sequence of pheromone-
biosynthesis-activating neuropeptide-I1 (PBAN-11) of the silkworm, Bombyx mori, Agric. Biol. Chem.. 54,
2495, 1990.
85. Kitamura, A., Nagasawa, H., Kataoka, H., Inoue, T., Matsumoto, S., Ando, T., and Suzuki, A., Amino
acid sequence of pheromone-biosynthesis-activatingneuropeptide (PBAN) of the silkworm, Bombyx mori,
Biochem. Biophys. Res. Commun., 163, 520, 1989.
Hormones and Reproduction 107

86. Matsumoto, R., Yamashita, O., Fonagy, A., Kurihara, M., Uchiumi, K., Niagamine, T., and Mitsui, T.,
Functional diversity of a pheromonatropic neuropeptide: induction of cuticular melanization and embryonic
diapause in lepidopteran insects by Pseudaletia pheromonotropin, J. Insecf Physiol., 38, 847, 1992.
87. Lafont, R., Reverse endocrinology, or "hormones" seeking functions, Insect Biochem., 21,697, 1991.
88. Foster, S.P., Bergh, J.C., Rose, S., and Harris, M.O., Aspects of pheromone biosynthesis in the Hessian
fly, Mayetiola destructor (Say), J. Insect Physiol., 37, 899, 1991.
89. Raina, K.A. and Menn, JJ., Pheromone biosynthesis activating neuropeptide: from discovery to current
status, Arch. Insect Biochem. Physiol., 22, 141, 1993.
90. Blomquist, GJ., Adams, T.S., Halarnkar, P.P., Gu, P., Mackay, M.E., and Brown, L.A., Ecdysteroid
induction of sex pheromone biosynthesis in the housefly, Musca domestica -are other factors involved?, J.
Insect Physiol., 38, 309, 1992.
91. Dumser, BJ., The regulation of spermatogenesis in insects, Annu. Rev. Entomol., 25, 341, 1980.
92. Dumser, B.J. and Davey, K.G., The Rhodnius testis: hormonal effects on germ cell division, Can. J. Zool.,
53, 1682, 1975.
93. Friedlander, M. and Reynolds, S.E., Meiotic metaphases are induced by 20-hydroxyecdysone during
spermatogenesis of the tobacco hornworm, Manduca sexta, J. Insect Physiol., 34, 1013, 1988.
94. Leviatan, R. and Freidlander, M., The eupyrenelapyrene dichotomous spermatogenesisof Lepidoptera. I.
The relationship with postembryonic development and the role of the decline in juvenile hormone titre towards
pupation, Dev. Biol., 68, 515, 1979.
95. Koolman, J., Scheller, K., and Bodenstein, D., Ecdysteroids in the adult male blowfly CaNiphora vicina.
Experientia.. 35, 134, 1979.
96. Loeb, MJ., Woods, C.W., Brandt, E.P., and Borkovec, A., Larval testes of the tobacco budworm: a new
source of insect ecdysteroids, Science, 218, 896, 1982.
97. Loeb, MJ., Brandt, E.P., and Birnbaum, MJ., Ecdysteroid production by testes of the tobacco budworm,
Heliothis virescens, from last larval instar to adult, J. Insect Physiol.. 30, 375, 1984.
98. Friedlander, M. and Reynolds, S.E., Intratesticular ecdysteroid titres and the arrest of sperm production
during pupal diapause in the tobacco hornworm, Manduca sexta, J. Insect Physiol., 38, 693, 1992.
99. Loeb, MJ., Brandt, E.P., Woods, C.W., and Borkovec, A., An ecdysiotropicfactor from brains of Heliofhis
virescens induces testes to produce a immuno-detectable ecdysteroids in vitro, J. Exp. Zool., 243,275, 1987.
100. Kambysellis, M. and Williams, C.M., In vitro development of insect tissues. 1. A macromolecular factor
prerequisite for silkworm spermatogenesis, Biol. Bull., 141, 527, 1971.
101. Kambysellis, M. and Williams, C.M., In vitro development of insect tissues. 1. The role of ecdysone in the
spermatogenesis of silkworms, Biol. Bull., 141, 541, 1971.
102. Friedlander, M. and Benz, G., Control of spermatogenesis resumption in post-diapausing larvae of the
codling moth, J. Insect Physiol., 28, 349, 1982.
103. Friedlander, M., 20-Hydroxyecdysoneinduces glycogen accumulation within the testicular sheath during in
vitro spermatogenesisrenewal in diapausing codling moths (Cydia pomonella), J. Insect Physiol. 35.29, 1989.
104. FriedPnder, M., Juvenile hormone and regulation of dichotomous spermatogenesis during larval diapause
of the codling moth, J. Insect Physiol., 28, 1009, 1982.
105. Shimizu, T., Moribayashi, A., and Agui, N., In vitro analyses of spermiogenesis and testicular ecdysteroids
in the cabbage armyworm,Mamesfra brassicae L. (Lepidoptera: Noctuidae), Appl. Entomol. Zool., 20.56.1985.
106. Giebultowicz, J.M., Loeb, MJ., and Borkovec, A.B., In vitro spermatogenesis in lepidopteran larvae: role
of the testis sheath, J. Invertebr. Reprod. Dev., 11, 21 1, 1987.
107. Shimizu, T., Yagi, S., and Kuramochi, K., Regulation of spermiogenesis in the common armyworm,
Leucania separafn (Lepidoptera: Noctuidae), Appl. Enromol. Zool., 23, 156, 1988.
108. Shimizu, T., Yagi, S., and Agui, N., The relationship of testicular and hemolymph ecdysteroid titer to
spermiogenesis in the common myworm, Leucania separata. Entomol. Exp. Appl., 50, 195, 1989.
109. Gelman, D.B., Woods, C.W., and Borkovec, A.B., Effects of ecdysone and 20-hydroxyecdysone on apyrene
spermiogenesis in the European corn borer, Ostrinia nubilalis. J. Insect Physiol., 34, 733, 1988.
110. Riemann, J.G., Thorson, BJ., and Ruud, R.L., Daily cycle of release of sperm from the testes of the
Mediterranian flour moth, J. Insect Physiol.. 20, 195, 1974.
111. Giebultowia, J.M., Riemann, J.G., Raina, A.K., and Ridgway, R.L., Circadian system controlling the
release of sperm in insect testes, Science. 245, 1098, 1989.
112. Thorson, BJ. and Riemann, J.G., Effect of 20-hydroxyecdysone on sperm release from the testes of the
Mediterranian flour moth, Anagasta kuehniella (Zeller), J. Insect Physiol., 28, 1013, 1982.
113. Giebultowicz, J.M., Feldlaufer, M., and Gelman, D.B., Role of ecdysteroids in the regulation of sperm
release from the testis of the gypsy moth, Lymantria dispar, J. Insect Physiol., 36, 567, 1990.
114. Osanai, M., Kasuga, H., and Aikagi, T., Induction of sperm motility of apyrene sperm and dissociation of
eupyrene sperm bundles of the silkworm, Bombyx mori by initiatorin and trypsin, Invertebr. Reprod. Dev., 15,
97, 1989.
108 Insect Reproduction

115. Nowock, J., Induction of imaginal differentiation by ecdysone in the testes of Ephestia kuehniella, J. Insect
Physiol.. 18, 1699, 1972.
116. Szollosi, A. and Landureau, J.-C., Imaginal cell differentiation in the spermduct of Samia cynthia (Lepi-
doptera). Responses in vitro to ecdysone and ecdysterone, Biol. Cell., 28, 23, 1977.
117. Shinbo, H. and Happ, G.M., Effects of ecdysteroids on the growth of the post-testicular reproductive organs
in the silkworm, Bombyx mori, J . Insecr Physiol., 35, 855, 1989.
118. Shimizu, T., Development of spermduct and seminal vesicle during phmte adult of cabbage armyworm,
Mamestra brassicae, Invertebr. Reprod. Dev., 15, 221, 1989.
119. Loeb, MJ., Growth and development of spermducts of the tobacco budworm, Heliothis virescens, in vivo and
in vitro, Invertebr. Reprod. Dev., 19, 97, 1991.
120. Loeb, MJ., Development of isolated spermducts from Heliothis virescens (Lepidoptera) in virro, Invertebr.
Reprod. Dev., 20.67, 1991.
121. Kelly, TJ., Adams, T.S., Schwartz, M.B., Birnbaum, MJ., Rubenstein, E.C., and Imberski, R.B.,
Juvenile hormone and ovarian maturation in the diptera: a review of recent results, Insect Biochem., 17, 1089,
1987.
 Chapter 5

FATAL ATTRACTION: THE DISRUPTION OF MATING

AND FERTILIZATION FOR INSECT CONTROL
Richard Wall

CONTENTS
I . Introduction ...............................................................................................................
109

I1. Sterile Insect Technique ............................................................................................

110
A. The Principles of Control by SIT ....................................................................... 110
B . Practical Control by SIT .....................................................................................
112
1. Screwworm Eradication in North America ...................................................113
2. Screwworm Eradication in North Africa ......................................................114

I11. Genetic Control ......................................................................................................... 115

A . Introduction ......................................................................................................... 115
B . Genetic Sexing Systems .....................................................................................115
1. General ........................................................................................................... 115
2 . Genetic Control of the Australian Sheep Blowfly ........................................ 116

IV . Autosterilization ........................................................................................................ 118

A. The Principles of Control by Autosterilization .................................................. 118
B . Attractants and Sterilants ....................................................................................120
C . Autosterilizing Systems ...................................................................................... 121
1. General ......................................................................................................... 121
2. Control of Tsetse Fly by Autosterilization .................................................... 121
3. Development of an Autosterilizing System for Housefly Control ...............122

V. Disruption of Behavior .............................................................................................. 123

V1. Conclusions ...............................................................................................................123

Acknowledgments ............................................................................................................... 125

References ...........................................................................................................................
125

.
I INTRODUCTION
The range of powerful insecticides now available provides the means of controlling the vast
majority of insect pests. However. complete reliance on these chemicals brings with it a
variety of associated problems. These include the development of resistance by the target pest
species. a variety of unintentional effects on nontarget organisms. the presence of pesticide
residues in food. and high costs both of development and deployment.' As a result. there is
growing concern that in the future the judicious use of chemical insecticides must. at the very
least. be supplemented by the development of a range of noninsecticidal techniques for pest

.
0-8493-6695-X/951$0.OOtSS50
8 1995 by CRC Press Inc.
110 Insect Repr.oduction

In the search for effective and acceptable alternatives to chemical control, considerable
attention has been directed toward insect reproduction. Mate location and identification,
copulation, insemination and fertilization, external physical and chemical stimuli, and the
internal physiological mechanisms that regulate each step present an enormous range of
features which are available for disruption or impairment. Disruption of component parts of
the process of reproduction allows the insect's own reproductive behavioral repertoire to be
turned against itself, to bring about genetic death. These autocidal control techniques have
several advantages over simple killing systems, including being highly species specific so that
they have minimal impact on the nontarget fauna.
The techniques so far developed for control by manipulation of insect reproduction are
those which produce a reduction in fertility. This has been effected principally by the release
of males which have been rendered sexually sterile, altered genetically, or otherwise treated
to disrupt reproduction in the natural population. In the case of sterile males, the released
individuals compete for matings in the wild population, and the eggs of wild females fertilized
by their sperm fail to hatch. The release of males with heritable genetic deficiencies4is a step
in sophistication beyond the use of individuals that have been sterilized by the induction of
dominant lethal mutations with radiation because it has the advantage that the resultant effects
may persist in the population for several generation^.^ Finally, the development of systems that
attract and impair the reproduction of individuals in the field overcomes the need to mass rear
and release sterilized or genetically modified insects. Such devices are known as autosterilizing
systems and are based on the integration of a chemosterilant or a compound which can produce
a similar end result through transovarial effects on eggs or larvae with an attractant into a
single control device which can be placed in the field.
This chapter presents an overview of these techniques and the principles behind them. It
focuses particularly on Diptera of medical and veterinary importance and seeks, wherever
possible, to give detailed examples of practical control operations in the field.

11. STERILE INSECT TECHNIQUE

The release of sterilized male insects to suppress a wild population was first proposed in
the late 1 9 3 0 It~ is
~ now referred to as the sterile insect technique or SIT. Irradiated sterile
males are released into a wild population in numbers sufficient to allow them to obtain a large
proportion of the matings with fertile females. Eggs fertilized by the sperm of the irradiated
males fail to hatch. As a result, with continued release of sterile males, the population of wild
insects is eventually driven to extinction.

A. THE PRINCIPLES OF CONTROL BY SIT

The effectiveness of the release of sterile males depends critically on the ratio of wild
females to sterile males released. At the simplest level, in a population of 100 fertile females
and an equal number of fertile males, if 900 sterile males are released to give a total male
population of 1000, on average only 100 X (100/1000) = 10 fertile matings will occur. If each
fertile female produces only a single female offspring, a further release of the same number
of sterile males would make it unlikely that any fertile matings would occur in the second
generation.'
The effectiveness of the suppression is influenced by the competitiveness of the released
males, immigration of already mated females into the release area, and the recovery capacity
or seasonal changes in the target populati~n.~These factors can be incorporated into a simple
equation8 which can be used to explore the effects of sterile male release:
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control

GENERATION NUMBER
FIGURE 1. The number of wild insects of either sex remaining in a population subjected to the release of a constant
number of sterile males, equivalent to the initial number of fertile females (solid line). Also, the change in ratio of
released males (R) to wild (W) insects of either sex (dashed line). There is no immigration (M = 0).The wild
population would have remained constant if there had been no intervention, and density-dependent factors do not
operate to give it any tendency to recover (D = 1). The released sterile males are assumed to be fully sterile and fully
competitive with wild males (C = 1).

W is the number of wild individuals of either sex in the population each generation n,
assuming an equal sex ratio. Sterilization is considered to be complete, and R is the number
of completely sterile males released. C is the probability of a released male mating with a wild
female, relative to that of a wild male doing so, where a value of 1 represents full competi-
tiveness. M is the number of fertile, mated, immigrant females. These are assumed to be
refractory to further mating. D simulates the tendency of the target population to increase or
decrease due to seasonal or density dependent factors. When D = 1 and R = 0,each adult
female produces an average of one female offspring and with no immigration (M = 0) the
population size remains constant. Sterile females released are not included in the equation
since they are not expected to affect the dynamics of the population.
Using the equation to examine the effects of altering the various parameters shows that
with a constant rate of release of sterile males (R), the population declines towards
eradication because as the number of wild individuals falls, the R:W, ratio increases
(Figure 1). However, the rate at which eradication of the wild population is achieved
would be retarded if the competitiveness of released males was lower than that of wild
males (Figure 2).
The rate of eradication is also strongly affected by the immigration of fertile females since,
even if the released males sterilize all the indigenous females, this can never push the
reproducing population below the number of inseminated female immigrants per generation.
The rate of reduction due to sterile male release may also be affected in the short term by
underlying increases or decreases in the target population, but more important, may be
permanently prevented if density-dependent regulatory factors are strong enough to compen-
sate for the maximum sterility which can be achieved. Finally, in practical terms, the initial
size of the wild population is of importance, since low density populations require the release
of smaller numbers of sterile males to achieve a given rate of reduction.
The major advantages of the technique are that it is species specific, so that in most cases
it presents no environmental hazards. In addition, it may be an effective means of "mopping
up" the residual population left after insecticidal methods have achieved the maximum
possible effect, since even at low population densities, the behavioral mechanisms that bring
the sexes together for mating mean that released male flies are likely to be able to locate the
remaining fertile females.
 Insect Reproduction

o1 I I I

1 .75 .5 .25 .l .05

COMPETIIVENESS OF RELEASED MALES
FIGURE 2. The number of generations required to reduce a wild population to less than 1, by the release of a
constant number of sterile males, equivalent to the initial number of feltile females. Released males differ in their
competitiveness with wild males (C). There is no immigration and D in equation 1 is unity.

B. PRACTICAL CONTROL BY SIT

As the foregoing discussion shows, there are numerous practical imperatives which must
be achieved for the effective use of SIT. Most critically, to ensure success, the release area
must be isolated as far as possible to protect against immigration. The fact that this is seldom
achievable makes effective control difficult. For example, suppression of populations of the
blood-sucking pest the stable fly Stomoxys calcitrans, with sterilized males was attempted in
central F l ~ r i d aSterilization
.~ was brought about by exposure of pupae to 2 krad from a
source. Adults were allowed to emerge in the laboratory and released in areas of high cattle
density. Each day sterile flies were released, initially at equal and, eventually, at double the
estimated initial wild population. Despite daily losses of about 35% of the released males
through mortality and emigration and the fact that the released males were only about half as
competitive as wild males, after one generation, over 85% sterility was found in the native
population. With continued release, sterility rapidly approached loo%, while the field popu-
lation fell in size by 97%. Although sterility remained high in the release area for a number
of weeks after the last release of sterile males, subsequent reinfestation was rapid?
The usefulness of the sterile insect technique for the control or eradication of tsetse flies
(Glossina spp.), the vectors of African trypanosomiasis, has been demonstrated by several
experiments and field project^.'^-'^ However, the extensive rearing facilities and logistic
support required to breed and release large numbers of insects make application of SIT
expensive. This is particularly so when more than one species or strain of a single species need
to be controlled. For example, control of Glossina palpalis palpalis was achieved in central
Nigeria, through the release of more than 1.5 million sterile males in an area of 1500 km2.I3
Males were released at a ratio of at least 10 sterile to each fertile wild male, for at least 3
generations. Control was assisted by the fact that G. p. palpalis populations are concentrated
in areas of riverine habitat, thus allowing the release of sterile males to be focused in these
areas. The release area was maintained free of G. p. palpalis for 3 years by deploying
insecticide trap and target barriers around the cleared area. However, the problem of animal
trypanosomiasis was not eliminated because of the presence of the sympatric species Glossina
tachinoides. Additionally, when dealing with pests that are harmful to man or animals such
as tsetse flies, the release of hundreds of thousands of potential vectors, although sterilized,
could increase substantially the transmission of disease. Therefore, the wild population must
be reduced significantly with insecticides or traps prior to the release.
The competitiveness of the males reared for release is often difficult to maintain under mass
rearing regimens in the laboratory. For example, attempts to control the mosquito Anopheles
quadrimaculatus in Florida involved the release of 433,600 irradiated males over a period of
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 113

14 months.14No reduction in mosquito numbers and no reduction in fertility of wild females

resulted. It was concluded that the failure to suppress the wild population was the result of the
fact that the laboratory strain used was too inbred to be competitive with wild males. Similar
lack of success was recorded in an attempt to control the mosquito Aedes aegypti, lack of
competitiveness on this occasion being attributed to too high a radiation dose.15
One further disadvantage of sterile male release and all other sterilizing systems over
simply killing the target pest with insecticides is the delay in suppressing the population. If
a pest has already reached economic damage levels, considerable further losses would occur
before the sterilized flies has any affect on the wild population.
In addition to the above examples, control by release of sterile males has been attempted
for the suppression of populations of numerous other species of pest Diptera, such as the
Mediterranean fruit fly Ceratitis capitata,16 horn flies Haematobia irritans,I7 the blowfly
Lucilia sericata,I8 and the heel fly Hypoderma lineatum.lg However, in many cases, while
trials have given promising results, the technique has been found too expensive to be used on
a larger scale in the field, or little more than temporary, local population suppression has been
achieved. The major exception to this pattern, however, has been the use of the sterile insect
technique for the eradication of the New World screwworm fly Cochliomyia hominivorax
from North America and North Africa.

1. Screwworm Eradication in North America

The screwworm fly C. hominivorax is an obligate primary parasite of warm-blooded
animals, including humans.20Adult females lay batches of up to 350450 eggs on the edges
of open wounds or in body orifices. On hatching, within 24 hours the maggots start to feed,
burrowing into the living tissue.21The resulting extensive wound may rapidly lead to the death
of the struck
The natural range of the fly extends from the southern states of the U.S. through Central
America and the Caribbean Islands to northern Chile, Argentina, and Uruguay. Each summer
the fly used to spread north and west into more temperate zones from its overwintering areas
near the U.S./Mexican border. The fly was of greatest significance as a pest of livestock,
necessitating the continued costs of vigilance, treatment and control. In the epidemic year of
1935 in Texas, there were approximately 230,000 cases in livestock and 55 in humans.23
In 1954 the first full-scale operation to eradicate C. hominivorax was mounted on the island
of Curagao, 50 miles off the coast of Venezuela. Eradication was achieved in 6 months by
swamping the wild population with about 800 sterile males per square mile per week with
screwworm files sterilized with X-rays.24Attention was then turned to removal of the screw-
worm fly from mainland U.S. The fly was eradicated first of all from the Florida peninsula
in 1957 through 1959, assisted by the unusually cold winters of 1957 and 1958. Pupae 5 days
old were exposed to 60Coradiation and both sexes were released as adults at a peak rate of
about 500 km2/week.25
In subsequent stages, the fly was progressively pushed south from areas of the U.S. Flies
were produced at a large rearing factory in Texas capable of producing 200 million flies per
week. These were sterilized with 137Csat a minimum dose of 5500 rads and released from the
air as adults. Eventually a 3 100-km long barrier zone was established against reinvasion, from
Texas to California along the Mexico b ~ r d e r . ~ ~ , ~ ~
Between 1972 and 1976, however, the number of infestations of C. hominivorax recorded
in the U.S. increased dramati~ally.~~ This may have been the result of the intense selection
pressure generated by the massive scale of the release of irradiated screwworm flies, leading
to rapid evolution of a strain of wild flies that no longer mated readily with the released males.
In addition, factory rearing may have favored the selection of a domesticated strain unable to
compete with wild males.27The outbreak in Texas in 1976 was estimated to have cost
consumers and producers between U.S. $280-370 million. Nevertheless, effective control of
C. hominivorax in the U.S. was reimposed in 1979, and no screwworm infestations have been
 Insect Reproduction

FIGURE 3. The number of confirmed cases of New World screwworm fly infestation recorded each month in
Libya in 1990 and 1991 (solid line); the number of sterile screwworm flies released each month in 1991 (dashed line).
(Modified from FAO New World Screwworm Newsletter 1991, 1.)

detected since 1984.28,29The U.S. Department of Agriculture program has subsequently been
directed against the fly in Mexico, Puerto Rico, Vieques, and the Virgin I s l a n d ~ , 2and
~ . ~in
~
1991 Mexico was declared officially free of screwworm fly.
The effects of weather on the outcome of the SIT control campaigns has been the subject
of recent debate. Weather has been recognized to affect screwworm fly populations; higher
temperatures and humidity increase the survival of this subtropical species and affect under-
lying rates of population increase.24It is notable that the successful screwworm SIT control
campaigns in the U.S. coincided with the particularly cold winters of 1957, 1958, and 1962.
It has been suggested that a contributory factor to the breakdown of screwworm control in the
U.S. between 1972 and 1976 may have been the favorable warm winters in those years. These
may have allowed flies to invade the control area and overwinter, or may have enabled
undetected residual populations to expand rapidly.30Furthermore, it was suggested that given
the presence of undetected residual populations, new outbreaks may well occur when climatic
conditions again favor the pest. The existence of residual populations has been disputed,
however, on the grounds that they could not remain undetected, given the fecundity and
mobility of C. hominivorax and the clinical severity of screwworm infestations. It was further
suggested that there were no grounds for supposing that climatic factors are critical to the
success of screwworm eradication, especially in subtropical areas such as southern
Nevertheless, by influencing the initial size of the target population and the underlying rate
of population increase, climatic factors will inevitably affect the ease with which a population
can be suppressed by SIT.

2. Screwworm Eradication in North Africa

In 1988 C. hominivorax screwworm flies were discovered in an area 10 km south of Tripoli
in Libya.32They probably arrived with a shipment of contaminated livestock from South
America. This was the first known established population of this species outside the Americas.
The fly quickly spread to infest about 25,000 km2. In 1989 there were about 150 cases of
myiasis by C. hominivorax, but by 1990, a total of 12,068 confirmed cases of screwworm fly
myiasis were recorded and, at its peak, almost 3000 cases were seen in the single month of
September 1990 (Figure 3). It was estimated that if unchecked the infestation could cost the
Libyian livestock industry about U.S. $30 million per year and the North African region
approximately U.S. $280 million per year.33 Considerable alarm was expressed about the
effects of allowing the fly to remain, since it is well adapted for rapid dispersaP4 and could
have spread throughout North Africa and possibly into more favorable subSaharan habitats.35
Fatal Attraction: the Disruption of Mating and Fertilizationfor Insect Control 115

As a result, an international program to eradicate the fly was established, coordinated by the
Food and Agriculture Organization of the United Nations.
C. hominivorax were reared at the Tuxtla factory in Mexico, where they were sterilized by
exposure to gamma radiation as late stage pupae. These were then transported in refrigerated
vehicles to Mexico City, and then air-freighted to Libya. On arrival, the boxes were stored in
environmentally controlled trailers and a sugar-based food was supplied to sustain the emerg-
ing flies. When approximately 80% of the adults had emerged, boxes of 1600 pupae were
dispersed at about three to ten boxes per minute, from light aircraft flying at 240 kmlh and at
an altitude of 500 m. The aircraft flew along predetermined flight paths about 4 km apart. The
boxes opened in midair or on impact with the ground, releasing the newly emerged sterile
adults.33
The screwworm fly population in Libya was able to overwinter, though it showed a
substantial fluctuation in response to weather (Figure 3). After a trial program in December
1990, full-scale releases began in February 1991, at the time when the screwworm fly
population was at its seasonal minimum. Flies were released at initial rates of 3.5 million per
week, escalating to peak rates in July 1991 of 40 million per week, at densities ranging from
500-1200 per km2. By May 1991 an area of 41,000 km2 was being treated each week. The
release of sterile males was backed up by extensive control operations on the ground,
including surveillance teams and quarantine stations to prevent movement of infested animals
outside the infested area. In 1990 a total of 12,068 confirmed cases of C. hominivorax
infestation has been recorded, but in 1991 only 6 cases were seen, the last occurring in April
(Figure 3). By November, after 6 months with no further cases, the release of sterile flies was
terminated and eradication a n n ~ u n c e dThe
. ~ ~ final cost of the program was estimated to be
approximately U.S. $64 million.
The outstanding success of this sterile male release operation can be attributed in part to
the ready availability of large numbers of screwworm flies and access to the already developed
U.S. and Mexican expertise. In addition, the success of the operation was due to the absence
of immigration, the confinement of the outbreak population in North Africa by the ocean to
the north and desert to the south, and to the fact that in Libya, C. hominivorax was probably
close to the edge of its climatic range, a marked trough in abundance occurring at unfavorable
times of year.

111. GENETIC CONTROL

A. INTRODUCTION
Advances in SIT stimulated interest in the development of alternative mechanisms by
which insects could be rendered effectively sterile. This interest has resulted in the identifi-
cation of a wide range of genetic mechanisms that can be used to manipulate the genetic
composition of natural insect population~.~ These can broadly be divided into two categories.
The first include mechanisms that have an effect for one generation only after release, and
include dominant lethal mutations such as those induced by radiation or chemosterilants,
cytoplasmic incompatibility, and hybrid sterility. The second includes inherited partial steril-
ity, recessive lethal genes, and chromosome translocations. These mechanisms lead to inher-
itable distortions which, therefore, continue to exert an effect on the population for some
generations after release. The advantages of sterility effects in the target population, prolonged
over several generations, have concentrated attention on the second category of genetic
mechanism, and translocations have, to date, appeared to be the most useful of these.

B. GENETIC SEXING SYSTEMS

1. General
Chromosome breakage is a common result of the irradiation of cells. If the broken sections
of two nonhomologous chromosomes rejoin to the wrong partners, the result is called reciprocal
116 Insect Reproduction

translocation. Meiosis in individuals heterozygous for such translocations may then result in
the production of gametes with an unbalanced, lethal, genetic complement. When these
fertilize normal gametes, nonviable embryos are produced. The heterozygote is referred to as
semisterile. For a single translocation, half of the progeny of a mating between a heterozygous
and normal individuals are usually affected. A homozygote, which inherits the same translo-
cation from both parents, may have impaired fertility or viability. Nevertheless, identification
and selection of such homozygotes may allow them to be reared in sufficient numbers in the
laboratory for their release in to the field. Every mating between an individual homozygous
for the translocation and a normal individual will then result in the production of heterozy-
gotes, which at meiosis will produce a proportion of nonviable gametes. When the semisterile
heterozygote mates, only a proportion of its offspring will mature. The rearing and release of
insects carrying semisterilizing autosomal translocations as a means of pest control was first
proposed by S e r e b r ~ v s k y . ~ ~
The three main types of translocations that can occur are between an autosome (non-sex
chromosome) and the Y chromosome, an autosome and X chromosome, or between two
autosomes, In the first case, semisterility is inherited through the male. In the second, it can
be inherited through both the male and female, but only female homozygotes are possible. In
the third case, the translocation can be inherited through both sexes and both can be homozygotes.
The artificial linkage of deleterious selectable genes to sex, using Y-autosome transloca-
tions is known as a genetic sexing system. The development of genetic sexing systems may
be of considerable value in the separation of the sexes in laboratory cultures, allowing males
to be isolated prior to release in conventional SIT programs.38Hence, they may be particularly
important in mosquitoes where males do not bite and mass release of biting females would be
unacceptable. Genetic sexing systems have been proposed for a number of species of Diptera,
including the housefly Musca dorne~tica,~~ the tsetse flies G. austeni and G. rnor~itans,"~.~~the
mosquitoes Culex tritaeniorhynch~s~~,~~ s ~ ~the Mediterranean fruit fly C.
and A. a l b i m a n ~ and
c a p i t ~ t a , although
4 ~ ~ ~ functioning genetic sexing systems have been established for relatively
few.48The use of Y-autosome translocations for control has been evaluated most comprehen-
sively in Australia for the sheep blowfly Lucilia ~ u p r i n a . 4 ~ ~ ~ ~

2. Genetic Control of the Australian Sheep Blowfly

The blowfly L. cuprina is the most important myiasis pest of sheep in A u ~ t r a l i aAnnual
.~~
costs of production losses, prevention, and treatment were estimated to be approximately $149
million in 1985 and, in the absence of effective preventative measures, the sheep industry
would be nonviable over much of the present grazing land in Australia. The primary means
of combating flystrike in Australia has been with insecticides. However, concerns over
growing insecticide resistance resulted in the initiation of research into SIT and genetic
control.
It was found that exposure of L. cuprina to radiation can induce a range of mutations
including translocations where one or more sections of the Y chromosome are swapped with
sections of one or more autosomes. Males possessing the translocation can be identified and
selected in the laboratory. Possession of the Y-linked translocation confers partial sterility, and
a proportion of the eggs produced in a cross between a modified male and a wild female will
fail to h a t ~ h . 4Among
~ . ~ ~ the individuals that do hatch, the deleterious translocation is carried
by the male progeny but not the females.
However, in a further refinement of the technique, it was found to be possible to translocate
wild type alleles of autosomal mutations affecting eye color onto the Y chromosome. The
recessive mutations are carried on the nontranslocation chromosome set by the genetically
modified males and in heterozygotic females, but are not expressed because of the presence
of the dominant wild type alleles. In female homozygotes, the mutations are expressed as
white or yellow eye color. These females lack the light-filtering pigments that give the normal
blowfly eye its reddish-brown color, so the homozygous mutant flies are effectively blind in
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 117

FEU) FEMALE RELEASED MALE

I
Normal

.-*
Partially sterile and carries
eye mutations on non-
partial sterility translocated chromosomes.

RELEASED MALE

W+

Partially sterlle and carries

Partially sterile eye mutations in
heterozygous form
1
,,Partial Sterility

Combined effect of sterility

and blindness causes up to MALE OFFSPRING FEMALE OFFSPRING
98% genetic death.
All partially sterile and many Most females blind. Rest
carry heterozygous eye carry eye mutations in
mutations. hetarozygous form.

b & l e heterozygote; normal White eye homozygote Yellow eye homozygote

vision but likely to produce (Mind) (blind).
Mind progeny.

FIGURE 4. Transfer of genes (white eye W*, yellow eye Ye+, and the white and yellow eye mutations W and Ye)
and translocated chromosomes from genetically modified male Lucilia cuprina released in the field.

daylight. When heterozygous females mate with the genetically altered males, a large percentage
of their female progeny are homozygous for one or more of the eye pigment mutations. The
blind females can readily be reared in the laboratory, but die rapidly in the field. The system
is termed a female-killing (FK) system or the genetically impaired female technique (GIFT)?
Hence, release of the modified strains causes genetic death, partly from semisterility caused
by the chromosome rearrangement and partly from death of the female descendants of the
released males due to homozygosity for the mutations (Figure 4). An additional development
(not shown in Figure 4) is that the released males also carry inversions to try to ensure
maintenance of the linkage of the genes to the translocation and also to contribute an additional
degree of potential sterility from females which inherit the inversions in heterozygous form.
Computer simulations indicated that a theoretical maximum death rate of 98% per generation
could be achieved by release of males possessing the chromosome abnormalities and the eye
color rnutation~.~~ Furthermore, genetic death from semisterility and homozygosity should
118 Insect Reproduction

persist in the wild population for several generations after cessation of releases, giving this
control system a considerable advantage over conventional SIT in which suppression ceases
when release stop^.^^.^^ At lower release rates GIFT would be expected to result in more rapid
suppression of the target population than SIT.
In field trials on Flinders Island, which is about 40 km2and 27 km off the Australian coast,
34,000 modified male L. cuprina were released per km2each week between August 1985 and
May 1986. The induced rate of genetic death peaked at 87%, 6 months after the trial began
and the blowfly population declined from 345 females per hectare in October 1985 to less than
1 female per hectare in May 1986, when releases were terminated. The population remained
at below 4 females per hectare for the following 10 month^.^^.^^
However, population suppression with the release of genetically modified males also
suffers from many of the problems associated with conventional SIT, particularly the immi-
gration of females from outside the release area and the noncompetitiveness of released
males.s7While the Flinders Island trial showed that it is possible to reduce an isolated Lucillia
population by this method, further trials in the much larger Furneaux Island group encountered
significant problems. Difficulties were experienced in rearing the 15 million modified flies
required per week to swamp the wild population, and problems also arose from the breakdown
of the genetic strains under large-scale rearing, due to the spontaneous recombination of
genes.58The instability of genetic-sexing systems has also been recorded in a wide variety of
other insect species.59As a result of these considerations in combination with the expense of
maintaining large-scale rearing facilities, to date removal of L. cuprina from large areas of the
Australian mainland has not been attempted.

IV. AUTOSTERILIZATION
Many of the problems associated with the need to mass rear insects for the release of
sterilized or genetically modified males could be overcome by the use of devices which attract
and disrupt the fertility of wild flies in the field. Such devices are known as autosterilizing
systems. These devices may either sterilize flies directly, using a chemosterilant, or produce
an equivalent end result via transovarial effects on eggs, larvae, or pupae. The principles that
apply to the use of autosterilization systems are quite different from conventional killing
systems or SIT. However, the potential for effective and efficient suppression of pest popu-
lation~by autosterilization is in some respects much greater, although it does not have the
advantage of the steri1e:fertile male ratio improving as the wild population declines, as it does
with SIT.

A. THE PRINCIPLES OF CONTROL BY AUTOSTERILIZATION

A device that simply attracts and kills greater numbers of males is unlikely to have any
impact on the target pest population, since in most insect populations females mate infre-
quently, males are capable of inseminating several females, and the operational sex ratio,
therefore, is heavily biased towards males. As a result, even a very small proportion of
surviving males are usually capable of finding and inseminating all the available females. This
is a critical problem for traps baited with chemicals, such as pheromones, or sound, that attract
only males. Attracting, sterilizing, and then releasing males only would be more effective than
attracting and killing these males, since the sterile males would mate with fertile wild females.
The effects would then be similar to conventional SIT. Autosterilizing males also has the
benefits of reduced costs and complexity when compared with mass-rearing and release.
However, it would be difficult to achieve high sterile-to-fertile male ratios and, in most
circumstances, attracting and sterilizing males only would not bring about suppression of the
population as effectively as simply attracting and killing both sexes with insecticide.
Attracting and sterilizing females only would have no advantage over killing them at the
same rate with insecticide. The attraction and sterilization of both sexes, however, would be
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 119

TIME (WEEKS)

FIGURE 5. The change in abundance (log number) in a simulated housefly population that is untreated (solid line)
or treated with insecticides that kill 90% of each generation (thin dashed line) or sterilants that sterilize 90% of each
generation (thick dashed line). (Modified from Borkovec, A. Advances in Pest Control Research, Vol. 7, Interscience,
New York.)

the most effective ta~tic.~O*~' For example, if a housefly population was treated with insecticides
that kill 90% per generation, too few individuals would be killed to overcome the high
potential for increase (Figure 5). However, if the population was exposed to material that
sterilized 90% of both sexes of each generation, because sterile females are effectively
genetically dead and there are nine sterile to each fertile male attempting to obtain a mating
with each remaining fertile female, on average only 1% of the matings are between fertile
females and fertile males and produce viable offspring (Figure 5). Hence, sterilization would
be expected to eradicate the population in four generation^.^^
The value of sterilization of both sexes over killing depends on the proportion of the
population that can be attracted and sterilized each generati~n.~' At low rates of kill or
sterilization there is little to choose between the two techniques, but as a greater and greater
number of individuals are affected, the relative advantage of sterilization of both sexes over
killing increases steeply (Figure 6). This is because the proportion of fertile matings is the
product of the proportions of males and females not entering traps.63As a result, the availabil-
ity of highly attractive baits and traps is critical to the development of practical autosterilizing
systems.
Autosterilizing systems have a number of further theoretical advantages over insecticidal
treatment or mass-rearing and release. Differential survivorship of fertile and sterile males and

PERCENTAGE KIUEO OR STERILISED

FIGURE 6. Relative superiority of sterilization over killing for control, plotted against the percentage sterilized or
killed. Superiority is calculated as the ratio of the proportion of the population reproducing after treatment with
insecticides to the propoltion of the population reproducing after treatment with sterilant. (Modified from Knipling, E.
U.S.D.A. Agriculture Handbook, No. 512, U.S. Department of Agriculture, Washington, D.C., 1979,659.)
120 Insect Reproduction

residual fertility of males visiting the baits make much less difference to the outcome of
autosterilizing systems than they do to a conventional sterile release program, because much
of the effect comes from the sterilizing of the trapped females. In addition, with sterilization
of wild insects in the field, no additional insects are being added to the system.60Using
autosterilizing devices would also reduce the strength of selection for behavioral resistance to
attractants compared with killing traps, since attracted sterilized flies would mate with many
of the unattracted individuals.
The action of food-bated traps containing insecticide would be expected to be density
independent with respect to pest mortality. In contrast, the relative increase in efficiency of
sterilization of both sexes over killing would increase at lower pest densities, because of the
continued ability of individuals to find mates. The operation of density-dependent population
regulation also would add to the advantage of autosterilization over insecticide use." In
populations which experience high levels of density-dependence, a smaller absolute number
of individuals will need to be attracted and sterilized because of the lower density at which
the population equilibrates, although the percentage of the population that would need to be
attracted and sterilized per day would remain the same.61However, as with other control
methods based on the induction of sterility, immigration into the control area would be
especially detrimental to the use of auto~terilization.~~

B. ATTRACTANTS AND STERILANTS

As discussed earlier, the relative advantages of using an autosterilizing system depend on
the nature and potency of the attractant, since this determines how many and whether only one
or both sexes can be attracted. Attraction can be achieved either with olfactory
behavior-controlling chemicals (semiochemicals) or visual or auditory cues. Among the
semiochemicals, sex pheromones, particularly those produced by Lepidoptera, have received
most a t t e n t i ~ n . ~However,
~.~' most of the pheromones identified are produced by females for
the stimulation of males. Additionally, among the Diptera most pheromones, at best, have been
found to have only relatively low volatility. The hydrocarbons concerned often serve as
contact sex-recognition chemicals, arrestants, and mating stimuli for the male and induce male
copulatory activity when used in physiological amount^.^^-^' Pheromones produced by males
for female attraction have been identified in few flies, with the exceptions of fruit flies of the
genera D a c u ~and ~~Drosophila. Since attraction of males alone is of only limited use in an
autosterilizing system, nonpheromonal semiochemicals, the kairomones, particularly those used
by insects to locate host animals and plants for feeding or oviposition and which can attract both
sexes or predominantly females, may be of considerably greater value as attractant^.^^.^^
Vision has been shown to be of considerable importance in the behavior of many species
of i n s e ~ t .A ~~wide
. ~ ~range of features, such as shape, size, color, and movement have been
found to contribute to the attraction of a number of species of flies to baited traps. However,
of these, the responses to color have been shown to be of particular i m p ~ r t a n c e ?In ~ -general,
~~
attraction to blue, black, and red hues have been identified as most attractive in hematopha-
gous Diptera, such as tabanids, blackfly, and m o ~ q u i t o e sIn . ~ contrast,
~ diurnal herbivorous
insects have been found generally to be attracted to yellow hues with reflectivities between
500-580 nm.75 The development of attractive baits for insects has been the subject of
extensive work, which has been recently comprehensively reviewed by Muirhead-Thomp~on.~~
The theoretical advantages of autosterilization have prompted the search for chemical
sterilants that are effective and environmentally acceptable. Ideal sterilants should sterilize
both sexes of a target species or, alternatively, separate male and female sterilants should be
developed and used simultaneously. Sterilization should be lifelong and not impair mating
competitiveness. The materials used should affect only the target pest, either by being
biochemically species specific or by using a selective physical method of application.
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 121

A vast number of chemicals have been inve~tigated.~~ However, many of the most potent
materials, such as the alkylating agentsg2or the thioaziridine sterilant b i ~ a z i r ,are
~ ~potent
, ~ ~ not
only against insects but have such high mammalian toxicity as to be unsafe for deployment
in the field. A potent but entirely insect-specific sterilant, which acted against both sexes,
would represent a major advance in the development of autosterilizing systems.
Considerable interest has been given to the use of pathogenic viruses as pesticide^.^^
However, as yet, few appear to have shown practical promise as sterilants.
The search for alternative compounds has resulted in intensive studies of hormonal disrup-
tion of fertility or reproductive behavior using insect growth regulators ( I G R s ) . ~Amongst
~,~~
these the juvenile hormones and chitin synthesis inhibitors are of particular interest. The
juvenile hormones, their synthetic mimics, and the chitin synthesis inhibitors have long been
known to have a range of effects that might be exploited for control purposes, including
disruption of normal embryogenesis,molt inhibition and disruption interference with diapause,
stimulation of precocious egg development, and, most commonly, the derangement of meta-
morphosis. Although they have the advantages of being highly species specific and having low
persistence in the environment, as yet only a small number have found commercial application
for insect population suppression. As used at present, the mode of action of the IGRs currently
available means that they generally act as "third generation pesticideswg8in terms of popula-
tion suppression, rather than in the manner of sterilants, but as a result of their considerable
potential for further development, they are currently the focus of intensive r e s e a r ~ h . ~ ~ - ~ ~

C. AUTOSTERILIZING SYSTEMS
1. General
Some early successes in the development of practical autosterilizing systems incorporating
chemosterilants were made for use for the control of stable hou~eflies,~~and mo~quitoes.~~
A combination of chemosterilant and sweetened bait was used to control houseflies M. domestica
in poultry house^.^^^^^ However, their development and use in the field has been limited by the
high mammalian toxicity of the chemosterilants currently available and particularly by the
relatively low percentage of the target population that can be attracted by the baits available.
For control of an insect population with baits, the number that need to be treated per day
is dependent on the potential for increase of that p0pulation.9~With an insect such as a blowfly
or screwworm fly, where a single female may produce batches of 200 to 400 eggs several
times during its lifetime, approximately 20 to 40% of the females in the population need to
be killed per day to bring about its elimination. If both males and females could be sterilized,
approximately 10-20% of each sex would need to be affected per day. This level of attraction,
however, is seldom achievable with the baits currently available.
In contrast, the relatively low rate of reproduction of tsetse flies makes them good
candidates for control by trapping and autosterilization. Imposed increases in daily mortality
of only 2 4 % are required to bring about their eventual eradi~ation,~~ and baits available at
present make this level of attraction a practical reality.Im

2. Control of Tsetse Flies by Autosterilization

Initial experiments showed that it was possible to sterilize tsetse flies in the field with the
chemosterilant metepa,'O1and that both males and females could be attracted using host odors.
This was shown to be capable of suppressing an island population of two species of tsetse.lo2
However, metepa was considered to be too toxic for use in large-scale trials in the field. More
recently it has been shown that the unusual mode of reproduction of tsetse flies favors the use
of a juvenile hormone mimic, pyriproxyfen, as a practical chem~sterilant.~~~J"
The tsetse fly has an advanced form of adenotrophic viviparity. After mating, a single egg
is ovulated and retained in the oviduct where it hatches and undergoes three larval instars
 Insect Reproduction

FIGURE 7. A tsetse F2-trap7' with sterilizer attached and detail of the sterilizer, which consists of a wire frame
covered with mutton cloth dipped in pyriproxyfen solution. The frame is inserted into a plastic casing and attached
to the apex of the trap. Flies enter at the base of the trap and move up through the trap towards the light, where they
are focused by a mesh cone into the sterilizer tower. They leave via the sterilizer, after contacting the pyriproxyfen-
treated cloth. (Adapted from Hargrove, J. and Langley, P. Bull Entomol. Res., 80, 397, 1990.)

feeding on a nutritive secretion produced by the uterine glands of the female. Larviposition
occurs when the fully mature larva is ready to pupate.Io5Following suitable formulation in an
oil, the compound is absorbed through the adult female cuticle on tarsal contact. In the female,
pyriproxyfen is transferred to the larva "in utero." The third instar larva appears to be normal,
but after pupariation, metamorphosis is disrupted and the pupa dies.
Only very small amounts of pyriproxyfen are required, and a single treatment will ensure
that the female is effectively sterilized for life. Laboratory studies have shown that males
making contact with a treated surface can transfer sterilizing doses to females when they mate.lo4
A field trial conducted in Zimbabwe showed that pyriproxyfen can be used in traps for
tsetse control.106Tsetse entering traps, baited with host-odor olfactory attractants, were forced
to brush against material dosed with pyriproxyfen to effect their escape (Figure 7). Emergence
rates from puparia of G.m. morsitans and G. pallidipes fell to 30% and 2.7%, respectively. The
decline in birth rate was estimated as sufficient to cause a population reduction to 106 of its
original level if the autosterilization program was maintained for 1 year.

3. Development of an Autosterilizing System for Housefly Control

In addition to attracting sufficiently large numbers to the bait, a further practical obstacle
to the disruption of reproduction for autosterilization with IGRs is inducing flies to pick up
an effective dose of the compound. For example, significant egg hatch inhibition in the
housefly, M. domestica, following contact with surfaces treated with a 20% suspension
concentrate of the chitin synthesis inhibitor triflumuron, requires tarsal contact for at least one
hour.Io7Such concentrations and contact times are not realistic for a working autosterilizing
device. Nevertheless, recent studies have shown that it is possible to induce female houseflies
to pick up an effective dose by presenting triflumuron on sugar-baited targets.lo8Direct effects
on females may be a result of ingestion of triflumuron combined with tarsal contact allowing
absorption of the chemical across the cuticle. In addition, male houseflies exposed to sugar and
triflumuron treated targets are also capable of affecting the reproductive performance of
unexposed females. The effects of triflumuron on males may be through direct impairment of
spermatogenesis or insemination ability, or exposed males may simply pick up sufficient
triflumuron on their bodies to allow them to pass on an effective dose to normal females during
mating. Experimental studies have shown that after exposure of both sexes to triflumuron on
sugar-baited targets, egg hatch may be reduced to less than 4% and, for those larvae that do
eclose, less than 3% pupariate, giving a cumulative mortality of 98.8%.Io9If maintained, this
level of mortality would be sufficient to suppress a field population of houseflies. This
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 123

autosterilizing system is currently being tested for housefly control in poultry houses with
encouraging results.

V. DISRUPTION OF BEHAVIOR
Identification and isolation of the complex interaction of olfactory, visual, physiological,
and tactile cues used by insects to locate their mates and complete successful mating and
insemination may allow them to be used to provide inappropriate stimuli to manipulate and
disrupt reproductive behavior. For example, in many species of Diptera, males form persistent
swarms near a prominent marker. The males may pursue, court, or capture and copulate with
females passing briefly through the swarm.110It has been demonstrated that blood-seeking
females of the mosquito Cx. tritaeniohynchus can be attracted to artificial swarm markers and
killed."'
Following initial copulation, females of most species of insect become unreceptive to
further mating attempts by males for substantial periods, if not for life. This is believed to be
caused, at least in the short term, by the transfer of a receptivity-inhibiting substance, passed
in the form of an accessory gland secretion during c o p u l a t i ~ n . ~Identification
~~J~~ of these
chemicals and discovery of a means of contaminating virgin females in the field could be used
to switch off their receptivity, effectively rendering them sterile.
In tsetse flies (Glossina spp.), the contact sex pheromone of the female induces the initial
stage of male copulatory behavior. This includes arrestment of locomotion, orientation over
the surface of the female, extension of the hypopygium, and engagement of the genitalia.l14.115
It has been shown that the presence of synthetic tsetse sex pheromone on female decoys sewn
onto screens in the field will induce copulatory behavior in males that make contact with
them.11G120 However, it has not been possible to make any practical use of this behavior, using
either insecticide or chemosterilant added to the decoys, since insufficient males can be
attracted. In contrast, pheromones have been used effectively, as attractants and mating
disruptants for male Lepidoptera, in a number of field trial^.'^^-'^^
There is clearly considerable scope for further work in the development of techniques that
disrupt the mating behavior of insects by providing inappropriate stimuli or misdirecting
behavior. However, it should be noted that techniques that simply block or disrupt some stage
of the reproductive process act as direct alternatives to insecticides and have no inherent
superiority as in some respects do SIT, genetic control, and autosterilization.

VI. CONCLUSIONS
The efficacy of the release of sterile or genetically altered strains for insect population
suppression is considerable. The techniques are most appropriate for use against pest popula-
tions that exist at naturally low densities, or are subject to substantial seasonal population
density fluctuations. They may also be used to prevent the establishment of immigrant
populations in new areas or as a follow-up technique against populations that have already
been reduced by other methods. However, the costs associated with the scale on which these
techniques must be used render them impractical and uneconomic in most instances and
against most pest species. As a result, in the 50 years since the techniques were first proposed,
they have found relatively limited practical application. Only where it is feasible to eradicate
an isolated population of a particularly virulent pest, or to eradicate it from an area which will
subsequently be protected by a barrier zone, will these techniques be cost effective. Screw-
worm eradication in North America and Mexico and more recently in North Africa are the
classic examples.
In contrast, the use of autosterilizing devices baited with pheromone or kairomone attrac-
tants and a sterilizing agent seem to have substantially greater potential for widespread
124 Insect Reproduction

application in field control programs. Autosterilization has a number of advantages over the
other autocidal control techniques. The sterilization of both sexes at a specific rate would be
expected to achieve more efficient suppression than the release of sterile or genetically altered
males at the same rate or killing both sexes at the same rate with in~ecticides.~~.~'
Autosterilizing
systems provide a means of achieving many of the benefits of the release of sterile or
genetically altered males without the need to mass rear and release. However, considerable
costs may be incurred in the deployment and maintenance of sterilizing devices, particularly
in areas of terrain with difficult access. Nevertheless, they are, theoretically, highly cost
effective systems of autocidal pest control.
Autosterilizing systems may be particularly valuable where the aims of the control program
are less comprehensive than total eradication. In practical terms, autosterilization would be
most effective as a control technique when used against insects with low birth rates and high
adult survivorship and against isolated populations. The latter may be most easily found in
relatively closed systems such as poultry houses, barns, or grain storage silos. They also may
be important components of integrated pest management systems. For example, an
autosterilizing system could be used in combination with the release of sterile or genetically
impaired males. It could also be used in conjunction with the release of insect biocontrol
agents such as predators or parasitoids, so long as the autosterilizing device either did not
attract these or the chemosterilant used was inactive against them. The use of conventional
insecticides or traps is unhelpful with these techniques because they kill sterilized and fertile
insects equally.
Advances in the development of autosterilizing systems for a range of pest insects are
highly dependent on the identification of potent attractants and on the development of
powerful but entirely insect-specific sterilants, both of which should affect males and females.
The isolation of a range of chemical kairomones and visual attractants shows considerable
promise.80 While relatively few attractants yet appear to be able to lure sufficiently large
numbers to overcome the high intrinsic rates of increase of populations of many species of pest
Diptera, strategic timing of the deployment of an autosterilizing device so that it has maximum
impact on the target populations may contribute to its efficient ~ s e .Strategic
~ ~ ~deployment
- ~ ~ ~
requires a detailed understanding of the seasonal population dynamics of the target species.
In the development of autosterilizing systems, less progress has been made in the search
for effective sterilants. A great deal of interest has been given to the use of IGRs. However,
the most common effects of juvenile hormones and juvenile hormone mimics are the disrup-
tion of various aspects of metamorphosis. They do not generally affect the egg production of
higher Diptera. The development of successful autosterilizing systems for tsetse control has
been possible, using a juvenile hormone mimic, but it is the unusual viviparous habit of the
tsetse which permits disruption of metamorphosis through treatment of the adult female. A
similar mode of application is unlikely to be effective, therefore, for other flies. Greater
success for Diptera may be achieved using the chitin synthesis inhibitors, such as
triflurn~ron.~~~-~@'The considerable benefits of specificity and environmental safety that would
be gained through using an IGR or other biochemical sterilant, should direct intensive research
towards this area. While it might initially appear almost impossible to affect male fertility
using IGRs, the aim of affecting males could be achieved by the development of delivery
systems which attract and allow males to pick up sufficient material to enable them to transfer
effective doses to females during subsequent mating. The males thereby would be rendered
effectively sterile. The use of sex pheromones, as behavioral arrestants for males, may have
an important role in this context.
The manipulation of the reproductive physiology and behavior of insects to bring about
their destruction has the potential for highly effective suppression of pest populations. How-
ever, the development of autocidal techniques for pest control is complex. Detailed informa-
tion relating to the ecology, genetics, behavior, and physiology needs to be obtained for every
pest species to be controlled. Basic information is also needed concerning the seasonal
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 125

fluctuations, mobility, rates of increase, and, in particular, the extent to which the attempts to
suppress a target population are likely to be negated by density-dependent compensation. The
growing recognition of the need to develop more specific, effective, and environmentally
sensitive pest control techniques will increase the urgency of these areas of research.

ACKNOWLEDGMENTS
I am grateful to Dr. M. J. R. Hall and an anonymous referee for their helpful comments on
this chapter. The financial support of a Royal Society University Research Fellowship is
gratefully acknowledged.

REFERENCES
I. Curtis, C. F. and Lines, J. D., Insecticides in the management of insect vectors of tropical disease, Insect
Sci. Applic., 8, 709, 1987.
2. Worthington, E. B., Insecticides, Nature, 234, 55, 1971.
3. Strong, L. and Wall, R., The chemical control of livestock parasites: problems and alternatives, Parasitol.
Today, 6,291, 1990.
4. Dame, D. A., Control of insects of veterinary importance by genetic techniques, Prev. Vet. Med., 2,515, 1984.
5. Foster, G. G., Weller, G. L., James, W. J., Paschalidis, K. M., and McKenzie, L. J., Advances in sheep
blowfly genetic control in Australia, in IAEMFAO Int. Symp. Management Insect Pests: Nuclear and Related
Molecular and Genetic Techniques, International Atomic Energy Agency/Food and Agricultural Organiza-
tion, Vienna, 1992.
6. Bushland, R. C. and Hopkins, D. E., Experiments with screw-worm flies sterilized by X-rays, J. Econ.
Entomol., 44, 725, 1951.
7. Knipling, E. F., The Basic Principles of Insect Population Suppression and Management, U.S.D.A. Agricul-
ture Handbook No. 512, U.S. Department of Agriculture, Washington, D.C., 1979, 659.
8. Curtis, C. F., Genetic control: growth industry or lead balloon?, Biol. J. Linn. Soc.. 2, 359, 1985.
9. La Brecque, G. C., Meifert, D. W., and Weidhaas, D. E., Potential of the sterile-male technique for the
control or eradication of stable flies, Stomoxys calcitrans Linnaeus, in Sterility Principlefor Insect Control,
IAEAFAO, Vienna, 1974,449.
10. Dame, D. A. and Schmidt, C. H., The sterile-male release technique against tsetse flies, Glossina spp., Bull.
Entomol. Soc. Am., 16, 24, 1970.
11. Williamson, D. L., Dame, D. A., Gates, D. B., Cobb, P. E., Bakuli, B., and Warner, P. V., Integration of
insect sterility and insecticides for control of Glossina morsitans morsitans Westwood (Diptera: Glossinidae)
in Tanzania. V. The impact of sequential releases of sterilized flies, Bull. Entomol. Res., 73, 391, 1983.
12. Takken, W., Olandunmade, M. A., Dengwat, L., Feldmann, H. H., Onah, J. A., Tenabe, S. O., and
Hamann, H. J., The eradication of Glossina palpalis palpalis (Robineau-Desvoidy)(Diptera: Glossinidae)
using traps, insecticide-impregnatedtargets and the sterile insect technique in central Nigeria, Bull. Entomol.
Res., 76, 275, 1986.
13. Olandunmade, M. A., Feldmann, U., Takken, W., Tenabe, S. O.,Hamann, H. J., Onah, J., Dengwat, L.,
Van Der Vloedt, A. M. V., and Gingrich, R. E., Eradication of Glossina palpalis palpalis (Robineau-
Desvoidy) (Dipten: Glossinidae) from agropastoral land in central Nigeria by means of the sterile insect
technique, in Sterile Insect Techniquefor Tsetse Control and Eradication, IAEMAO, Vienna, 1990, 5.
14. Weidhaas, D. E., Schmidt, E. L., and Seabrook, E. L., Field studies on the release of sterile males for the
control of Anopheles quadrimaculatus, Mosq. News. 22, 283, 1962.
15. Morlan, H. B., McCray, E. M., and Kilpatrick, J. W., Field tests with sexually sterile males for control of
Aedes aegypti, Mosq. News, 22,295, 1962.
16. Mellado, L., La tecnica de rnachos esteriles en el control de la mosca Mediterrano. Programas realizados en
Espana, in Sterility Principle for lnsect Control or Eradication, IAEMAO, Vienna, 1971, 49.
17. Kunz, S. E., Graham, M. R, Hogan, B. F., and Eschle, J. L., Effect of releases of sterile horn flies into
a native population of horn flies, Environ. Entomol., 3, 159, 1974.
18. MacLeod, J. and Donnelly, J., Failure to reduce an isolated blowfly population by the sterile males release
method, Entomol. Exp. Appl., 4, 101, 1961.
19. Kunz, S. E., Drummond, R. O., and Weintraub, J., A pilot test to study the use of the sterile insect technique
for eradication of cattle grubs, Prev. Vet. Med., 2, 523, 1984.
126 Insect Reproduction

20. James, M. T., The Flies That Cause Myiasis in Man, U.S. Dep. Agric. Misc. Publ., No. 631, 1947, 175.
21. Thomas, D. B. and Mangan, R. L., Oviposition and wound visiting behaviour of the screwworm fly,
Cochliomyia hominivorar (Diptera: Calliphoridae), Ann. Entomol. Soc. Am., 82, 526, 1989.
22. Humphrey, J. D., Spradbery, J. P., and Tozer, R. S., Chrysomya beuiana: pathology of old world screw-
worm fly infestations in cattle, Exp. Parasitol., 49, 381, 1980.
23. Dove, W. E., Myiasis of man, J . Econ. Entornol.. 30, 29, 1935.
24. Bushland, R. C., Screwworm research and eradication, Bull. Entomol. Soc. Am., 21, 23, 1975.
25. Baumhover, A. H., Husman, C. N., Skipper, C. C., and New, W. D., J. Econ. Entomol.. 52, 1202, 1959.
26. Knipling, E. F., The eradication of the screwworm fly, Sci. Am., 203, 54, 1960.
27. Bush, G. L., Neck, R. W., and Kitto, G. B., Screwworm eradication: inadvertent selection for noncompeti-
tive ecotypes during mass rearing, Science. 193, 491, 1976.
28. Graham, 0.H., Symposium on eradication of the screwwormfrom the United Statesand Mexico, Misc. Publ.
Entomol. Soc. Am.. 62. 1985, 1.
29. Krafsur, E. S., Whitten, C. J., and Novy, J. E., Screwworm eradication in North and Central America,
Parasitol. Today, 3, 131, 1987.
30. Readshaw, J. L., Screwworm eradication a grand delusion?, Nature, 320,407, 1986.
3 1. Krafsur, E. S., Townson, H., Davidson, G., and Curtis, C. F., Screwworm eradication is what it seems,
Nature, 323, 495, 1986.
32. Gabaj, M. M., Wyatt, N. P., Pont, A. C., Beesley, W. N., Awam, M. A. Q., Gusbi, A. M., and Benhaj,
K. M., The screwworm fly in Libya: a threat to the livestock industry of the Old-World, Vet. Rec., 125,347,
1989.
33. Lindquist, D. A., Abusowa, M., and Hall, M. J. R., The New World screwworm fly in Libya: a review of
irs introduction and eradication, Med. Vet. Entomol.. 6, 2, 1992.
34. Hightower, B. G., Adams, A. L., and Alley, D. A., Dispersal of released irradiated laboratory-reared
screwworm flies, J. Econ. Entomol., 58, 373, 1965.
35. Hall, M. J. R. and Beesley, W. N., The New World screwworm fly in North Africa, Pestic. Outlook, 1, 34,
1990.
36. FAO, New World Screwworm Newsletter, FAO/Screwworm Emergency Centre for North Africa (SECNA),
Rome, 1991, 1.
37. Serebrovsky, A. S., On the possibility of a new method for the control of insect pests, Zool. Zh., 19,618, 1940
[Russian].
38. Busch-Peterson, E., Genetic sex separation in sterile insect technique pest management programmes, with
special reference to the medfly Ceratitis capitara (Weid.), in Fruit Flies of Economic Importance, Cavalloro,
R., Ed.,CECAOBC Int. Symp. 7-10 April, 1987, Rome, Italy, 1989, 225.
39. Wagoner, D. E., Nickel, A. C., and Johnson, 0. A., Chromosomal translocation heterozygotes in the house
fly, J. Hered., 60, 301, 1969.
40. Curtis, C. F., Southern, D. I., Pell., P. E., and Craig-Cameron, T. A., Chromosome translocations in
Glossina austeni, Genet. Res. Camb., 20, 101, 1972.
41. Curtis, C. F., Langley, P. A., Mews, A. R., Offori, E. D., Southern, D. I., and Pell, P. E., Sex ratio distortion
and semi-sterility in the progeny of irradiated Glossina morsitans. Genet. Res. Camb., 21, 153, 1973.
42. Baker, R. H., Reisen, W. K., Sakai, R. K., Hayes, C. G., Aslamkhan, M., Saifuddin, U. T., Mahmood,
F., Perveen, A., and Javed, S., Field assessment of mating competitiveness of male Culex tritaeniorhynchus
carrying a chromosomal aberration, Ann. Entomol. Soc. Am., 72, 75 1, 1979.
43. Baker, R. H., Reisen, W. K., Sakai, R. K., Rathor, H. R., Raana, K., Azra, K., and Niaz, S., Mating
behaviour and competitivenessof males carrying a complex chromosomal aberration,Ann. Entomol. Soc. Am.,
73, 58 1, 1980.
44. Suguna, S. G., Seawright, J. A., Joslyn, D. J., and Rabbani, M. G., Homozygous pericentric inversions and
other studies of inversions on chromosome 3 in the mosquito Anopheles albimanus, Can. J. Genet. Cytol.. 23,
57, 1981.
45. Rossler, Y., Recombination in males and females of the Mediterranean fruit fly (Diptera: Tephritidae) with
and without chromosomal aberrations, Ann. Entomol. Soc. Am., 75,619, 1979.
46. Robinson, A. S. and Van Heemert, C., Ceratitis capitata -a suitable case for genetic sexing, Genetica. 58,
229. 1982.
47. Saul, S. H., Genetic sexing in the Mediterraneanfruit fly Ceratitis capitara (Weidemann)(Diptera: Tephritidae):
conditional lethal translocations that preferentially eliminate females, Ann. Enromol. Soc. Am., 77,280, 1984.
48. Kaiser, P. E., Seawright, J. A., Dame, D. A., and Joslyn, D. J., Development of a genetic sexing system
for Anopheles albimanus, Econ. Entomol., 71, 766, 1978.
49. Whitten, M. J., Insect control by genetic manipulation of natural populations, Science, 171, 682, 1971.
50. Whitten, M. J., Foster, G. G., Vogt, W. G., Kitching, R. L., Woodburn, T. L., and Konovalov, C., Current
status of genetic control of the Australian sheep blowfly, Lucilia cuprina Weidemann (Diptera:Calliphoridae),
Proc. XV lnt. Congr. Enromol., Washington, D.C., 1976, 129.
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 127

5 1. Foster, G. G., Vogt, W. G., and Woodburn, T. L., Genetic analysis of field trials of sex-linked translocation
strains for genetic control of the Australian sheep blowfly Lucilia cuprina Weidemann, Aust. J. Biol. Sci., 38,
275, 1985.
52. Foster, G. G.,The sheep blowfly genetic control programme in Australia, Insect and Pest Control Newsletter,
FAOIIAEA, 43, 23, 1989.
53. Foster, G. G., Kitching, R. L., Vogt, W. G., and Whitten, M. J., Sheep blowfly and its control in the pastoral
ecosystem of Australia, Proc. Ecol. Soc. Aust., 9, 213, 1975.
54. Whitten, M. J., The use of genetically-selectedstrains for pest replacement or suppression, in Genetics in
Relation to Insect Management, Hoy, M. A. and McKelvey, J. J., Eds., The Rockefeller Foundation, New
York, 1979, 31.
55. Foster, G. G., Vogt, W. G., Woodburn, T. L., and Smith, P. H., Computer simulation of genetic control.
Comparison of sterile male and field-female killing systems, Theor. Appl. Genet., 76, 870, 1988.
56. Davidson, S., Sheep blowfly control by genetic sabotage, Rural Res., 145, 19, 1990.
57. Vogt, W. G., Woodburn, T. L., and Foster, G. G., Ecological analysis of field trials conducted to assess
the potential of sex-linked translocation strains for genetic control of the Australian sheep blowfly, Lucilia
cuprina (Weidemann), Aust. J. Biol. Sci., 38, 259, 1985.
58. Foster, G. G., Chromosomal inversions and genetic control revisited: the use of inversions in sexing systems
for higher Diptera, Theor. Appl. Genet., 81, 619, 1991.
59. Busch-Peterson, E. and Baumgartner, H., Simulation of instability in genetic-sexingstrains: effects of male
recombination in association with other biological parameters, Bull. Entomol. Res.. 81, 11, 1991.
60. Langley, P. A. and Weidhaas, D., Trapping as means of controllingtsetse, Glossina spp. (Diptera:Glossinidae):
the relative merits of killing and sterilization, Bull. Enromol. Res., 76, 89, 1986.
61. Wall, R. and Howard, J., Autosterilization for control of the housefly Musca domestica. J. Theor. Biol., 117,
431, 1994.
62. Borkovec, A. B., Insect chemosterilants, Vol. 7, Advances in Pest Control Research, Interscience, New York,
1966, 143 pp.
63. Knipling, E. F., The Basic Principles of Insect Population Suppression and Management, U.S.D.A. Agricul-
ture Handbook, No. 512, U.S. Department of Agriculture, Washington, D.C., 1979, 659.
64. Barclay, H. J. and Mackauer, M., The sterile insect release method for pest control: a density dependent
model, Environ. Entomol.. 9, 810, 1980.
65. Barclay, H. J., Pheromone trapping models for pest control: effects of mating patterns and immigration,Res.
Popul. Ecol., 26, 303, 1984.
66. Klassen, W., Ridgway, R L., and Inscoe, M., Chemical attractants in integrated pest management programmes,
in Insect Suppression with Controlled Release Pheromone Systems. Vol. 1, Kydonieus, A. F. and Beroza, M,,
Eds., CRC Press, Boca Raton, FL, 1982, 13.
67. Sanders, C. J., Monitoring spruce budworm population density with six traps, Can. Ent., 120, 1175, 1988.
68. Bartell, R J., Shorey, H. H., and Barton-Browne, L., Pheromonal stimulation of the sexual activity of
males of the sheep blowfly Lucilia cuprina (Calliphoridae) by the female, Anim. Behav.. 17, 576, 1969.
69. Carlson, D. A., Mayer, M. S., Silhacek, D. L., James, J. D., Beroza, M., and Bierl, B. A., Sex-pheromone
attractant of the house fly: isolation, identification and synthesis, Science, 174, 76, 1971.
70. Carlson, D. A., Langley, P. A., and Huyton, P., Sex pheromones of the tsetse fly: isolation identification
and synthesis of contact aphrodisiac, Science, 201, 750, 1978.
71. Adams, T. S. and Holt, G. G., Effects of pheromone components when applied to different models on male
sexual behaviour in the housefly Musca domestica, J. Insect Physiol., 33, 9, 1987.
72. Fletcher, B. S., Storage and release of a sex-pheromoneby the Queensland fruit fly, Dacus tryoni (Diptera:
Trypetidae), Nature, 219,631, 1968.
73. Wall, R. and Langley, P. A., From behaviour to control: the development of trap and target techniques for
tsetse fly population management, Agric. Zool. Rev., 4, 137, 1991.
74. Coppedge, J. R., Broce, A. B., Tannahill, F. H., Goodenough, J. L., Snow, J. W., and Crystal, M. M.,
Development of a bait system for suppression of adult screwworms, J. Econ. Entomol., 71, 483, 1978.
75. Prokopy, R. J. and Owens, E. D., Visual detection of plants by herbivorous insects. Annu. Rev. Entomol..
28, 337, 1983.
76. Allan, S. A., Day, J. F., and Edman, J. D., Visual ecology of biting flies. Annu. Rev. Entomol., 32,297,1987.
77. Green, C. H. and Flint, S., An analysis of colour effects in the performance of the F2 tnp against Glossinn
pallidipes Austen and G. morsitans morsitans Westwood (Diptera: Glossinidae), Bull. Entomol. Res., 76,409,1986.
78. Holloway, M. P. T. and Phelps, R. J., The responses of Stomoxys spp. (Diptera: Nuscidae) to traps and
artificial host odours in the field, Bull. Entomol. Res.. 81, 51, 1991.
79. Wall, R., Green, C., French, N., and Morgan, K., Development of an attractive target for the sheep blowfly
Lucilia sericata, Med. Vet. Entomol., 6, 67, 1992.
80. Muirhead-Thompson, R. C., Trap Responses of Flying Insects. The Influence of Trap Design on Capture
Efficiency. Academic Press, London, 1991,287.
128 Insect Reproduction

81. Fye, R. L. and LaBreque, G. C., Bibliography of Arthropod Chemosterilants, ARS S-93, Agricultural
Research Service, U.S. Department of Agriculture, 1976, 54 pp.
82. La Brecque, G. C., Studies with three alkylating agents as house fly sterilants, J . Econ. Entomol.. 54, 684,
1961.
83. Langley, P. A. and Carlson, D. A., Laboratory evaluation of bisazir as a practical chemosterilant for tsetse
control, Bull. Entomol. Res., 76, 583, 1986.
84. Hall, M. J. R. and Langley, P. A., Development of a system for sterilizing tsetse flies, Glossina spp., in the
field, Med. Vet. Entomol., 1, 201, 1987.
85. Moore, N. F., King, L. A., and Possee, R. D., Viruses of insects, lnsect Sci. Appl., 8, 275, 1987.
86. Schniderman, H. A., Insect hormones and insect control, in Insect Juvenile Hormones Chemistry and Action,
Academic Press, London, 1972, 3.
87. Ruscoe, C. N. E., lnsect control by hormones, in Biology in Pest and Disease Control, Price Jones, D. and
Solomon, M. E., Eds., Blackwell, Oxford, 1974, 147.
88. Williams, C. M., Third-generation insecticides, Sci. Am.. 217, 13, 1967.
89. Friedel, T., Hales, D. F., and Birch, D., Cyromazine-induced effects on the larval cuticle of the sheep
blowfly, Lucilia cuprina: ultrastructural evidence for a possible mode of action, Pestic. Biochem. Physiol., 31,
99, 1988.
90. Langley, P. A., Felton, T., and Oouchi, H., Juvenile hormone mimics as effective sterilants for the tsetse
fly Glossina morsitans morsitans, Med. Vet. Entomol., 2, 29, 1988.
91. Advances in Regulation of Insect Reproduction, Bennettova, B., Gelbic, I., and Soldan, T., Eds., Institute of
Entomology, Czechoslovakian Academy of Science, Academia Praha, Prague. 1992, 329.
92. Insect Juvenile Hormone Research. Fundamental and Applied Approaches. Chemistry Biochemistry and
Mode of Action., Mauchamp, B., Couillaud, F., and Baehr, J. C., Eds., lnstitut National de la Recherche
Agronomique, Paris, 1992,296.
93. L a Brecque, G. C., Meifert, D. W., and Rye, J., Jr., Experimental control of stable flies Stomoxys calcitrans
(Diptera: Muscidae) by release of chemosterilized adults, Can. Entomol.. 104, 885, 1971.
94. La Brecque, G. C., Fye, R. L., and Morgan, J., Jr., Induction of sterility in adult house flies and stable flies
by chemosterilization of pupae, J. Econ. Enromol., 65, 751, 1972.
95. Ikeshoji, T. and Yap, Y. Y., Monitoring and chemosterilization of a mosquito population, Culex quefasciatus
(Diptera: Culicidae) by sound traps, Appl. Entomol. Zool., 22,474, 1987.
96. La Brecque, G. C. and Meifert, D. W., Control of house flies (Diptera: Muscidae) in poultry houses with
chemosterilants, J. Med. Enromol., 3, 232, 1966.
97. Meifert, D. W. and La Brecque, G. C., Integrated control for the suppression of a population of houseflies,
Musca domestica L., J. Med. Entomol., 8, 43, 1971.
98. Weidhaas, D. E. and Haile, D. G., A theoretical model to determine the degree of trapping required for insect
population control, Bull. Entomol. Soc. Am., 24, 18, 1978.
99. Hargrove, J. W., Tsetse: the limits to population growth, Med. Vet. Entomol., 2, 203, 1988.
100. Vale, G. A., Bursell, E., and Hargrove, J. W., Catching-out the tsetse fly, Parasitol. Today, 1. 106, 1985.
101. House, A. P. R., Chemosterilization of Glossina morsitans morsitans Westwood and G . pallidipes Austen
(Diptera: Glossinidae) in the field, Bull. Entomol. Res., 72, 529, 1982.
102. Vale, G. A., Hargrove, J. W., Cockbill, G. F., and Phelps, R. J., Field trials of baits to control populations
of Glossina morsitans rnorsitans Westwood and G. pallidipes Austen (Diptera: Glossinidae), Bull. Entomol.
Res., 76, 179, 1986.
103. Langley, P. A., Felton, T., and Oouchi, H., Juvenile hormone mimics as effective sterilants for the tsetse
fly Glossina rnorsitans morsitans, Med. Vet. Entomol., 2, 29, 1988.
104. Langley, P. A., Felton, T., Stafford, K., and Oouchi, H., Formulation of pyriproxyfen, a juvenile hormone
mimic, for tsetse control, Med. Vet. Entomol., 4, 127, 1990.
105. Denlinger, D. A. and Ma, W. C., Dynamics of the pregnancy cycle in the tsetse Glossina morsitans. J. Insect
Physiol.. 20, 1015, 1974.
106. Hargrove, J. W. and Langley, P. A., Sterilizing tsetse (Diptera: Glossinidae) in the field: a successful trial,
Bull. Entomol. Res., 80, 397, 1990.
107. Howard, J. and Wall, R., The effects of triflumuron, a chitin synthesis inhibitor, on the house fly, Musca
domestica (Diptera: Muscidae), Bull. Entomol. Res., 85, 71, 1995.
108. Howard, J. and Wall, R., The use of triflumuron on sugar-baited targets for autosterilization of the house
fly, Musca domestica, Ent. Exp. Appl., 74, 1995.
109. Howard, J. and Wall, R., Autosterilization of the house fly, Musca domestica, using the chitin synthesis
inhibitor Triflumuron on sugar-baited targets, Med. Vet. Entomol., 10, 1995.
110. Downes, J. A., The swarming and mating flight of Diptera, Annu. Rev. Entomol.. 14, 271, 1969.
11 1. Bidlingmeyer, W. L. and Hem, D. G., The range of visual attraction and the effect of competitive visual
attractants upon mosquito (Diptera: Culicidae) flight, Bull. Entomol. Res., 70, 321, 1980.
Fatal Attraction: the Disruption of Mating and Fertilization for Insect Control 129

112. Gillott, C., Arthropods - Insecta. Reproductive Biology of Invertebrates, Vol. 3, Accessory Sex Glands,
Adiyodi, G. G. and Adiyodi, R. D., Oxford, New Dehli, 1988, 319.
113. Smith, P. H., Gillott, C., Barton Browne, L., and van Genven, A. C. M., The mating induced refractoriness
of Lucilia cuprina females: manipulating the male contribution, Physiol. Entomol., 15, 469, 1990.
114. Huyton, P. M., Langley, P. A., Carlson, D. A., and Coates, T. W., The role of sex pheromones in initiation
of copulatory behaviour by male tsetse flies, Glossina morsitans morsitans. Physiol. Entornul.. 5,243, 1980.
115. Huyton, P. M., Langley, P. A., Carlson, D. A., and Schwan, M., Specificity of contact sex pheromones
in tsetse flies, Glossina spp., Physiol. Entomol., 5, 253, 1980.
116. Langley, P. A., Coates, T. W., Carlson, D. A., Vale, G. A., and Marshall, J., Prospects for autosterilization
of tsetse flies, Glossina spp. (Diptera: Glossinidae), using sex pheromone and bisazir in the field, Bull.
Entomol. Res., 72, 319, 1982.
117. Hall, M. J. R., The orientation of tsetse flies to pheromone-baited decoy 'females' in the field, Bull. Entomol.
Res., 77, 487, 1987.
118. Hall, M. J. R., Characterisation of the sexual responses of male tsetse flies, Glossina morsitans morsitans to
pheromone-baited decoy 'females' in the field, Physiol. Entomol., 13, 49, 1988.
119. Hall, M. J. R. and Langley, P. A., The responses of individual males in an isolated population of Glossina
morsitans morsitans Westwood (Diptera: Glossinidae) to pheromone-baited decoy 'females', Bull. Entomol.
Res., 79, 319, 1989.
120. Wall, R., Sexual responses of males of Glossina morsitans morsitans and G. pallidipes to traps and targets
in the field, Bull. Entomol. Res.. 79, 335, 1989.
121. Critchley, E. R., Campion, D. G., McVeigh,L. J., Hunter-Jones, P., Hall, D. R., Cork, A., Nesbit, B. F.,
Marrs, G. J., Jutsum, A. R., Hosny, M. M., and El-Syed, A. N., Control of pink bollworm, Pectinophora
gossypiella Saunders (Lepidoptera: Gelechiidae) in Egypt by mating disruption using an aerially applied
microencapsulated pheromone formulation, Bull. Entomol. Res., 73,289, 1983.
122. Webb, R. E., Tatman, K. M., Leonhardt, B. A., Plimmer, J. R., Boyd, V. K., Bystrak, P. G., Schwalbe,
C. P., and Douglass, L. W., Effects of aerial application of racemic disparlure released on male trap catch
and female mating success of Gypsy Moth (Lepidoptera: Lymantriidae), J. Econ. Entomol., 81, 268, 1990.
123. Webb, R. E., Leonhardt, B. A., Plimmer, J. R., Tatman, K. M., Boyd, V. K., Cohen, D. L., Schwalbe,
C. P., and Douglass, L. W., Effects of racemic disparlure released from grids of plastic ropes on mating
success of Gypsy Moth (Lepidoptera:Lymantriidae) as influenced by dose and by population density, J. Econ.
Entomol., 83, 910, 1990.
124. Wall, R., French, N., and Morgan, K. L., Sheep blowfly population control: development of a simulation
model and analysis of management strategies, J. Appl. Ecol.. 30, 743, 1993.
125. Wall, R., French, N., and Morgan, K. L., Predicting the abundance of the blowfly Lucilia sericata Meigen
(Diptera: Calliphoridae), Bull. Entomol. Res., 83,431, 1993.
 Chapter 6

PARTHENOGENESIS IN INSECTS WITH PARTICULAR

REFERENCE TO THE ECOLOGICAL ASPECTS
OF CYCLICAL PARTHENOGENESIS IN APHIDS
A. F. G. Dixon

CONTENTS
I. Introduction ............................................................................................................... 131

11. Evolution of Parthenogenesis....................................................................................

132

111. Parthenogenesis in Aphids ........................................................................................

133

IV. Ecology of Cyclical Parthenogenesis ....................................................................... 134

A. Telescoping of Generations ................................................................................ 134
B. Polyphenism ........................................................................................................ 135
1. Reproduction .................................................................................................. 136
2. Migration ........................................................................................................
136
3. DefenseICleaning ........................................................................................... 137
4. Estivation/Hibernation ................................................................................... 137
C. Complex Life Cycles ..........................................................................................137
D. Facultative Tracking of the Environment .......................................................... 138

V. Conclusions ...............................................................................................................
138

Acknowledgments ...............................................................................................................
139

References ........................................................................................................................... 139

I. INTRODUCTION
The term "parthenogenesis" was coined by Richard O ~ e n in ' ~1849, who defined it as the
successive production of procreating individuals from a single ovum. He also viewed the
increase in numbers by parthenogenesis as similar to growth, a view which was supported by
H u ~ l e yin~1858,
~ and much later by Jan~en,5~ who stressed that in apomictic parthenogens
natural selection acts at the level of a clone, which he designated the "evolutionary indi-
vidual." That is, a clone produced by parthenogenesis is equivalent to the body of a sexually
reproducing organism although split up into a number of physically separate units. Etymologi-
cally, the word parthenogenesis means "reproduction by a virgin."
Gametic reproduction either involves genetic mixing, as in normal sexual reproduction and
automictic parthenogenesis, or no genetic mixing, as in apomictic parthenogenesis. The
distinction between apomictic and automictic parthenogenesis is blurred because there are
some forms of automictic parthenogenesis (functional apomictics) in which genetic mixing is
s u p p r e s ~ e d Parthenogenesis
.~~~~~ is widespread in the Insecta, occurring in Coleoptera,
Collembola, Diptera, Embioptera,Ephemeroptera, Hemiptera, Isoptera, Lepidoptera, Orthoptera,

0-8493-6695-X/95/S0.M)+SS5O
Q 1995 by CRC Press. Inc.
132 Insect Reproduction

Psocoptera, Strepsiptera, Trichoptera, and Thysan~ptera.~~ Indeed, parthenogenesis was first

experimentally confirmed by Bonnet,14working on aphids in the 18th century, and virgin birth
was observed in aphids by Leeuwenhoek in the 17th century. However, in birds and mammals
parthenogenesis is completely absent, and it is extremely rare in molluscs. A major challenge
is to develop a theory to account for the phylogenetic distribution of parthenogenesis.The high
incidence among insects may be because they are relatively immobile compared with larger
organisms and therefore less able to seek refuge from adverse climatological and biological
conditions. Under these conditions, parthenogenesis may have been at a selective advantage
because it conserves general purpose genotypes.62However, the comparable dispersal ability
but markedly different incidence of parthenogenesis in aphids and marine molluscs tends to
refute this suggestion.

11. EVOLUTION OF PARTHENOGENESIS

Reasons generally cited for the abandonment of sex are that periodic shortages of males in
small local populations would favor females that are parthenogenetic; parthenogens are better
able to conserve particular genotypes and so more easily exploit extreme environments, or
parthenogenesis offers advantages for the colonization and rapid exploitation of ephemeral
habitats.22.25.27,62.85.90 This applies to parthenogenetic ability that is under genetic control, e.g.,
References 43 and 50. A very different approach is to see parthenogenesis in terms of the
advantages for microorganisms or cytoplasmic elements, transmitted solely in the female line,
of manipulating the sex ratio in favor of females.86
Hymenoptera have diploid females and haploid males, i.e., sex is dependent on whether an
egg is fertilized, which is under the control of the mother. Some mothers, however, produce
only females without fertilization (thelytoky). Interestingly, the completely parthenogenetic
Encarsia and Trichogramma can be rendered permanently bisexual by treatment with antibi-
otics. There is no evidence that this is genetically controlled, but there is evidence of
extrachromosomal inheritance. The males that develop after treatment with antibiotics pro-
duce sperm and mate with females, but are unable to inseminate them. It is suggested that the
parthenogenetic females lay unfertilized eggs and that microorganisms cause the restoration
of diploidy.logAlthough the female-biased sex ratios of Hymenoptera are explicable in terms
of an evolutionary stable strategy of the insect's genome, under certain conditions this strategy
and adaptations of the insect's symbionts coincide.86The presence of ovarially transmitted
microorganisms or cytoplasmic elements capable of specifically killing males appears to be
widespread in insects. There are examples known from Coleoptera, Diptera, Hemiptera,
Hymenoptera, and L e p i d ~ p t e r aTherefore,
.~~ their role in determining sex ratios and, more
important for this review, their possible role in the evolution of parthenogenesis, should not
be neglected.
Deleterious or nonfunctional alleles accumulate in continuously parthenogenetic strain^^^.^^
as they have no way of reducing the overall load. That is, mutation load cannot be less than
that already existing in the organism's at present least-loaded lines. This is known as Muller's
ratchet p r i n ~ i p l e .The
~ , ~ expectation
~ is that exclusively parthenogenetic reproduction should
lead, despite compensating mutation,94to rapid extinction on an evolutionary time scale, with
the clonal species that do evolve not giving rise to higher taxonomic c a t e g o r i e ~ .However,
~~.~~
as pointed out by Maynard Smith6' and W i l l i a m ~ ,there ' ~ ~ are notable challenging exceptions.
The whole of the rotifer order Bdelloidea, with its several families and many genera and
species, and the freshwater gastrotrich order Chaetonotoidea are exclusively parthenoge-
~ case of the diploid parthenogenetic brine shrimp, Artemia, the diversity and the
n e t i ~In. ~the
5 million years' existence of the parthenogenetic lineage is thought to have been sustained by
automictic recombinati~n.~~
Apomictic parthenogenesis is thought to have evolved either from automictic parthenogen-
esis in which the meiotic division is retained43.90Jw or abruptly in its full form.10sBecause of
Parthenogenesis in Insects 133

the imperfect cytogenetic mechanism involved in automictic parthenogenesis, it initially

confers a lower fitness than either pure sexuality or apomixis, most strikingly because of poor
egg hatch, but also the retention of mating behavior in spite of nearly complete absence of
ma1es.21~58~64~90~92
These disadvantages might explain why automicts are relatively rare and
apomicts, which do not incur these costs, f l o ~ r i s h . ~ . ~ ~
That apomictic parthenogenesis may have evolved abruptly in its full form is supported by
both aphids and Daphnia having genes that suppress meiosis. A genetic change results in the
failure to produce sexual forms and may result in the coexistence of cyclic parthenogenetic
and continuously parthenogenetic clones. In response to environmental stimuli, some clones
produce sexuals (holocyclic), others some males and parthenogenetic females (androcyclic),
and others only parthenogenetic females (anholocyclic). In the case of the aphid Myzus
persicae, these life cycle traits are inherited monofactorily, with androcycly recessive to
holocycly, which allows the exchange of genes between androcyclic and holocyclic clones
and the generation of other anholocyclic clone^.^.^^ Further studies on the inheritance of these
life history traits are needed for a better understanding of the evolution of apomictic parthe-
nogenesis and in particular cyclical parthenogenesis.
Alternatively, as exposing parthenogenetic strains of some species of Hymenoptera to high
temperatures also induces the production of males, it has been suggested that this might form
the basis for the seasonal control of sexual reproduction in the cyclical parthenogenesis of
organisms like aphids. There is some evidence that the maternally transmitted symbionts in
aphids have a nutritive role, but they have not yet been shown to have a causative role in
gametogenesis or sex determination, although they are known not to enter male and soldier
embryos of some but not all

111. PARTHENOGENESIS IN APHIDS

Most aphids show cyclical parthenogenesis in which there is several generations of parthe-
nogenetic reproduction between each bout of sexual reproduction. Owen7$thought sexual
reproduction occurred on the exhaustion of the spermatic force associated with fertilization.
However, a hundred years earlier, BonnetI4had correctly attributed the appearance of sexuals
to the effect of long chilly autumnal nights. In parthenogenetic reproduction each ovum is
produced mitotically (apomixis), and as a consequence the offspring are identical. This view
was challenged by C ~ g n e t t i who
, ~ ~ claimed that there is an exchange of alleles between
homologous chromosomes within the prophase nucleus of the oocyte, a process known as
endomeiosis, which generates variation within clones. This led to a controversy, which has
been reviewed by Bla~krnan~-~' and Pagliai-Bonvicni and C ~ g n e t t iSuffice
. ~ ~ it to say, exhaus-
tive cytological and morphological studies and even one using DNA finger~rinting?~ have
failed to reveal any significant intraclonal variation.
Four strains of the raspberry aphid, Amphorophora idaei Borner, differentially infest
cultivars of raspberry. The aphid strains result from segregation at two gene loci and the plant
resistance to aphids from different combinations of 14 gene^.^^.^^ Similarly, in Acyrthosiphon
pisum (Harris), Metopolophium dirhodum (Walker), M. persicae (Sulzer), Schizaphis graminum
(Rhondani), and Sitobion avenue (F.), particular genotypes are better adapted to living on
certain plant species or cultivars within the aphid's host range.65-74-80~81~91~100~101
Thus, contrary
to Blakeley'sI3 claim, there is support for the idea that genetic variation among host plants is
important for the retention of sex in aphids. That is, sex and the resultant genetic recombina-
tion are at a selective advantage because all the other organisms that aphids interact with are
evolving.93However, the increasing use of sterile males and lures baited with sex pheromones
for capturing males for controlling pests, combined with the widespread planting of geneti-
cally uniform crops, would appear to tip selection in favor of parthenogenesis in insect pests.
Although sex in aphids has advantages in heterogeneous environments, nevertheless con-
tinuously parthenogenetic species of aphids have rapidly adapted to a wide range of conditions.
134 Insect Reproduction

This has been achieved by mutation and is particularly well illustrated by the history of
Therioaphis trifolii forma maculata (Monell) in North America. In 10 years, a continuously
parthenogenetic clone introduced into New Mexico from the Mediterranean area colonized
most of the southern areas of North America and evolved biotypes that were able to thrive on
previously resistant cultivars of alfalfa and also became resistant to certain insecticides, and
in the northern areas it even started to reproduce sexually and overwinter as eggs. Therefore,
the lack of genetic recombination did not prevent the spotted alfalfa aphid from adapting to
new ~ o n d i t i o n s . ' ~The
, ~ ~success
. ~ ~ of this parthenogen, as in other pest aphids, comes from
infesting crops that tend to be spatially invariant habitats, in which a single clone may gain
dominance3I and there is a mutation rate sufficient to allow it to adapt to changes in its largely
man-made and relative homogeneous habitat.
Within the family Aphididae there is only one exception to the notion that exclusively
parthenogenetic forms do not give rise to higher taxonomic categories - that is, the Tramini.
This tribe consists of 4 genera and 32 species,39and it would be of considerable interest to
know more about this group's taxonomic status and evolutionary history. In addition, only 1
out of 30 species of aphids is continuously parthenogenetic." The potentially greater rate of
increase of a continuously parthenogenetic mutant76should rapidly result in the competitive
elimination of cyclically parthenogenetic strains. Therefore, there must be a markedly com-
pensating advantage of sex.61This, as indicated above, is thought to be the production of rare
recombinant genotypes that can survive in the face of new adversity. In retaining cyclical
parthenogenesis, aphids have the advantages of both parthenogenesis and sex. The prevalence
of cyclically parthenogenetic over continuously parthenogenetic species of aphids is a conse-
quence of the balance between the extinction of asexual populations and the origination by
fixation of asexual mutants.23

IV. ECOLOGY OF CYCLICAL PARTHENOGENESIS

Once parthenogenetic, certain lines of development become possible, in particular, the
telescoping of generations, polyphenism, and host alternation. Aphids have developed these
life history traits, which to a large extent accounts for their success.

A. TELESCOPING OF GENERATIONS
The strong relationship between the rates of increase and the size of organisms indicates
that aphids have a faster rate of increase than one would expect. They achieve rates of increase
that are more characteristic of organisms the size of mites, i.e., one to two orders of magnitude
smaller. 32-42 Most organisms first grow and then switch to reproduction, and an important
feature that constrains their rate of increase is the time it takes them to develop from birth to
maturity.
Aphids, as a consequence of being parthenogenetic, have developed a different strategy.
They invest in both somatic and gonadal growth during their larval development with soma
growing logistically and the gonads exponentially.19 At birth aphids already have embryos
developing in their gonads, and their most advanced embryos have also started to develop
gonads. This telescoping of generations is characteristic of aphids. That is, throughout larval
development aphids simultaneously invest in growth of soma and gonads such that on
becoming adult most aphids are ready to give birth, i.e., they do not indulge in a bang-bang
reproductive strategy.84What favors the simultaneous commitment to growth and reproduc-
tion during larval development in aphids?
If there is no constraint on the relative rate of development of the gonads, it is best to first
grow and then reproduce, i.e., a bang-bang strategy.84Kindlmann and D i ~ o nargue ~ ~ for a
constraint on the rate of development of the gonads. Fecundity is the conversion of gonadal
tissue into offspring containing their own gonads. Gonads are crucial for the future, while
soma only assimilates and then dies. The best strategy for maximizing growth rate is to use
Parthenogenesis in Insects 135

all the energy gained from the soma to keep the gonadal developmental rate maximal. After
moulting to the adult stage, aphids do not grow anymore and all the energy assimilated is
utilized for reproduction. Thus there must be an optimal somatic (S,) to gonadal (g,) ratio for
an adult. If the ratio were bigger there would be a surplus of energy, as the gonads would be
unable to utilize all of the energy because of the constraint on the rate of development of the
gonads (R). If the ratio were smaller, then there would be insufficient energy to ensure the
maximum rate of development of the gonads. Therefore, either energy would be wasted or
gonadal development rate would be submaximal during the adult stage if s,/g, is not optimal.
Embryos assimilate nutrients over the whole surface of their body.I6 Therefore, their
prenatal developmental time (i.e., the period from ovulation to birth) is unlikely to be affected
by the size of their gonads. Assuming that the maximum rate of development of the gonads,
R, applies to the embryos, then whether an embryo can achieve birth size before its mother
reaches maturity depends on when during its mother's development it is ovulated. If the
prenatal developmental time is longer than the mother's larval developmental time, then it is
advantageous for ovulation to occur and development to commence before the birth of the
mother. If not, then the embryo would have to complete its development after its mother
becomes adult, which would lead to further increase in the size of the mother's gonads during
adulthood and break the optimal balance between sA/gA.If ovulation occurs before a mother
is born, then she can give birth just after reaching maturity. In the case of aphids, ovulation
occurs in the grandmother such that each individual actually consists of individuals of three
generations telescoped together. After birth of offspring the mother's gonads are smaller, the
optimal balance between sA/gAis broken, and there is now surplus energy to support further
growth of the gonads, allowing maturation of other embryos and ovulation.
That aphids generally produce their offspring singly at intervals in time (iteroparous) rather
than all at once (semeloparous) can be similarly explained. The adult soma provides enough
energy to cover the respiration costs of a body of size s,+g, and the Rg, needed for the growth
of the gonads. However, if there is a constraint on the rate of development of the gonads, R,
then on giving birth, gonad size decreases to g where g <g, and only requires sufficient energy
to support Rg < RgA.Thus a certain amount of energy is unused until the gonads reach the size
g, again. Therefore, it is advantageous to keep the size of the gonads as close to g, as possible
in order to use most of the energy assimilated by the soma. This is achieved by the anterior
germaria ovulating first and ovulation occurring over a prolonged period so that there are
differences in the developmental status of embryos within an ovariole and, for example,
between the basal embryos in each of the o ~ a r i o l e sIn ~ way, aphids are able to produce
. ~ this
offspring continuously (iteroparously) and maintain the size of their gonads close to g,.
What evidence is there for a constraint on development? Under congenial conditions aphids
generally take approximately a week to develop from birth to maturity. In contrast, many other
insects take approximately 3 weeks. However, if one takes into consideration that an aphid
starts developing inside its grandmother, then the actual development time is 2.5 times longer
than it takes an aphid to develop from birth to maturity, i.e., approximately 3 weeks. Therefore,
there appears to be a minimum "time" required for development. Telescoping of generations
enabled aphids to overcome this constraint on development and achieve rates of increase more
typical of much smaller organisms. Similarly, the oocytes in the gonads of the immature stages
of the parthenogenetic cecidomyid fly Heteropezapygmaea may develop parthenogenetically,
which results in the larvae reproducing, a phenomenon known as paedogenesis.Io3Although
different from the telescoping of generations, paedogenesis in omitting the adult stage also
circumvents the constraint on development.

B. POLYPHENISM
An aphid life cycle usually consists of a sequence of parthenogenetic generations and ends
in sexual reproduction. Normally, individual aphids are short-lived, and experience only a
small part of the total range of environmental and nutritional conditions experienced by the
136 Insect Reproduction

LlPOlDAL RESERVES L QENERATlON

FIGURE 1. Diagrammatic representation of the relationships: (A) between gonadal size and lipoidal reserves in
individuals of generations 1 to 3; and (B) the size of the offspring born to individuals in generations 1 to 3.

species during a season. This has favored the evolution of generation-specific reproductive
strategies rather than a single but flexible strategy for all morphs of a specie^.^'.^^.^^ Individuals
of each generation are differentially adapted for survival, reproduction, and migration. How-
ever, this division of labor in which individuals specialize in one or other of these functions
imposes constraints in terms of resource allocation, physiology, and structure for carrying out
other functions. This concept of the changing importance of roles over time emphasizes the
functional aspect of polyphenism and largely avoids the use of the confusing array of names
for the various morphs.

1. Reproduction
Early in the season, while plants are actively growing, they are a rich source of food for
aphids, but generally become less suitable as they cease growing. Associated with this change
in host quality is a programmed change within clones of aphids in which individuals of the
first generation have proportionally larger gonads and smaller fat reserves than those of later
generation^.^^,^^^ First-generation individuals also tend to produce rapidly many small off-
spring and the later generations fewer but larger offspring (Figure 1). Large offspring are
better able to survive on poor quality hosts than are small ~ffspring.~'

2. Migration
As plants eventually die or are killed, aphids need to disperse to ensure their long-term
survival. In species that show alary dimorphism, minor movements on a plant and between
adjacent plants are mainly undertaken by unwinged individuals, with the winged individuals
migrating over greater distances.
Tactile stimulation associated with crowding60,83deterioration in host a combina-
tion of host quality and ~ r o w d i n g , 4day
~ . ~length,Io8
~ or a combination of these factod6 can
induce the development of winged forms:By responding to a number of stimuli rather than
one, aphids can possibly achieve a closer and more reliable tracking of environmental
conditions. As it takes a generation or more for a change of form, aphids tend to respond to
cues that enable them to anticipate the onset of adverse conditions; however, on reaching
maturity, they only migrate if conditions are ad~erse.~3 This enables them to produce the more
fecund and faster developing nonmigratory forms29J07while the host plant is favorable and,
as it becomes unfavourable, to switch to producing migratory forms.
Parthenogenesis in Insects 137

3. DefenseICleaning
The existence of a "soldier" morph in species of Colophina, Pemphigus, and Pseudoregma
is a particularly good example of the division of labor within a clone, with some individuals
specialized for defense and in some cases even cleaning. They can make up 13% of a colony,
and are usually very short-lived, may not feed or reproduce, and clean and defend the colony
against insect Similarly, approximately 50% of the aphids in the large galls
produced by Astegopteryx styracicola are "bitters" and possibly afford the gall protection
against predators like squirrel^.^-^,^^ Because a colony is likely to consist mainly of the
descendants of a founder aphid (i.e., it is a clone), investing in soldiers, which seize and kill
insect enemies or clean honeydew, shed skins or dead aphids from the colony could be
advantageous because it increases the fitness of the clone.

4. EstivationIHibernation
In summer, many plants, especially shrubs and trees, cease growing and, until their leaves
become senescent at the onset of autumn, are poor hosts for aphids. Several species of aphids
estivate during the summer. The second generation of sycamore aphids, Drepanosiphum
platanoidis, mature with small gonads and large fat body and may not reproduce for up to 8
week^.^^.^^ The aphid Periphyllus testudinaceus, also living on sycamore, estivates as a first
instar nymph. Poor nutrition induces this aphid to produce small flattened nymphs whose
bodies are covered and fringed with minute plates. These nymphs attach themselves very
closely to the surface of leaves and can only be removed with great difficulty by predators.
Many species can overwinter viviparously. In some species there is a special overwintering
form, called a hiemali~."~For example, about a fifth of the autumnal apterae of the lettuce root
aphid (Pemphigus bursarius) can survive in the soil without a host plant for 48 weeks at 3°C.3R
They have poorly developed gonads and a well-developed fat body. The environmental cue
that induces the development of the hiemalis in this root-feeding aphid is low temperature. The
hiemalis remains in hibernation until soil temperatures increase and its host plants resume
growing the following spring.53

C. COMPLEX LIFE CYCLES

Seasonally, 10% of aphids switch between unrelated host plants. This phenomenon, known
as host alteration or heteroecy, has long attracted interest. Bonnet14 suspected it existed, and
it was first described by Walker?$ The typical life cycle involves migration between a woody
host, on which sex occurs and eggs are laid (the primary host) and an herbaceous plant, where
reproduction is solely parthenogenetic (the secondary host).
Why aphids host-alternate was first addressed by M o r d v i l k ~ .Clearly,
~ ~ - ~ ~the twice-yearly
migratory cycle with associated mortality is a cost, especially in a taxon such as aphids with
poor host location abilities. Even though a clone can be represented by many host locators,
nevertheless only an average of 1 in 100, or even 1000, are thought to survive each host
transfer.89Why then do aphids host alternate? Throughout the literature of the last century,
there are two themes: (1) as aphids can produce many generations in the course of a season
they are able to exploit the complementary growth patterns of herbaceous and woody plants;
and (2) although it is advantageous for aphids to shift all life cycle stages to the phylogeneti-
cally younger and nutritionally more favorable herbaceous plants, the morphs produced in
spring have undergone such extensive morphological and physiological changes to living on
woody plants that it is impossible for them to live on other plants.'' An extensive review of
host alternation in aphids63and an experimental analysis of host alternation in C a ~ a r i e l l a ~ ~
give little support to the second hypothesis. It is concluded that such obligate resource
switching occurs when changing hosts is predictably more profitable than remaining on the
same plant species. The coexistence of plants displaying asynchronous phenologies supplied
138 Insect Reproduction

the potential, and the ability of aphids to produce a number of highly prolific generations in
quick succession, which amplify small differences in performance on the different plants,
supplied the means of exploiting the potential.

D. FACULTATIVE TRACKING OF THE ENVIRONMENT

Generation-specific strategies have enabled aphid clones to track very closely the predict-
able seasonal trends in habitat quality. Although at any one time the habitat is generally good
or poor, there is nevertheless a great deal of variability. It is probably no coincidence,
therefore, that many aphids have a facultative ability to vary their investment in gonads and
energy reserves.
In the unwinged individuals of each generation, in many aphids there is a directly propor-
tional trade-off in the investment in gonads and lipoidal reserves, which is independent of size.
Individuals with small gonads are better able to resist periods of starvation and seek out high
quality feeding sites, whereas those with large gonads are better able to exploit the high quality
habitats. Although the range in investments in gonads and lipoidal reserves in each clone
appears to be fixed, the proportion with small gonads and large lipoidal reserves and vice
versa, is variable, but under maternal control. In harsh habitats mothers produce proportionally
more individuals with small gonads and large lipoidal reserves and vice versa, but the range
remains the
Similarly, winged individuals show an intraclonal trade-off between reproductive invest-
ment and size of the fat body, which is also independent of size. Winged aphids with small
gonads have a greater urge to migrate, may fly longer and more frequently, and are better able
to survive starvation than aphids with large gonads. The short-distance migrants consist
mainly of individuals with large gonads, and the long-distance migrants should consist mainly
of individuals with small gonads. If they settle on a poor host they tend to retain functional
wing muscles longer than when they settle on a good host. The degree of differential migratory
ability shown by an aphid clone will probably reflect both the within and between year
heterogeneity of the environment. In favorable environments, individuals with a high repro-
ductive investment ("risk takers"), will be fitter than those with a low reproductive investment
("risk averse"), and vice versa.36*59.97.98
Alary dimorphism and intraclonal tactical diversity in reproductive investment is particu-
larly characteristic of one of the ten subfamilies of the Aphididae, the Aphidinae, and may
partly account for their success, with 60% of aphid species belonging to this subfamily.

V. CONCLUSIONS
The high incidence of parthenogenesis in the Insecta compared to many other groups of
animals is puzzling. Although ovarially transmitted microorganisms are undoubtedly impli-
cated in the evolution of parthenogenesis in some Hymenoptera, this is not the case for other
insect orders where parthenogenesis appears to be under genetic control. Parthenogenesis is
thought to be advantageous in extreme habitats because it conserves general-purpose geno-
types and in ephemeral habitats because it enhances their colonization and rapid exploitation.
In adopting cyclical parthenogenesis, aphids benefit from both sex and parthenogenesis.
The genetic variation generated by recombination during sexual reproduction enables them to
keep up with the generation of habitat heterogeneity, and parthenogenesis permits the prolific
propagation of the more successful genotypes. Parthenogenesis also enabled aphids to over-
come the developmental constraint on the rate of increase by beginning their development
inside their grandmother. Short individual life span also made it possible for aphids to develop
generation-specific strategies, which along with a facultative ability to vary their investment
in gonads and energy reserves enabled them to track very closely seasonal trends in habitat
quality. Their high rate of increase amplifies small differences in performance on different
Parthenogenesis in Insects 139

species of plants and so makes host plant specialization and host alternation advantageous.
Although the "altruistic" behavior of soldier aphids in defending a colony has rightly attracted
a lot of attention, it should not be allowed to obscure the fact that this is only a part of a
functional polyphenism involving defence, migration, reproduction, and survival, which is
likely to have been selected for because it increased fitness at the clonal level. Elton40
compared phenotypic plasticity to a conjuror's magic kettle, dispensing the beverage the
environment demands, which is particularly apt for aphids.
Given that cyclical parthenogenesis is a very effective mechanism for coping with both
long- and short-term environmental uncertainty, it is relevant to ask: why is it so rare? The
view that cyclical parthenogenesis is advantageous if the individual's life spans a small part
of the seasonal range of conditions is supported by the occurrence of cyclical parthenogenesis
in cladocera and aphids. Taxa with longer lived individuals lack the ability to develop the
temporal correspondence of strategy with so characteristic of aphids. However, a
short life span is common to many other groups of insects that do not show cyclical parthe-
nogenesis.

ACKNOWLEDGMENT
The author is indebted to Dr. Richard Sequeira for reading and commenting on the
manuscript.

REFERENCES
l. Aoki, S., Colophina clematis (Homoptera, Pemphigidae), an aphid species with 'soldiers', Konryi. 45, 276,
1977.
2. Aoki, S., Further observations on Astegopreryx sryracicola (Homoptem:Pemphigidae) an aphid species with
soldiers biting man. Konryi. 47. 99. 1979.
3. Aoki, S., Akimoto, S., and Yamane, D., Observations on Pseudoregma alexanderi (Homopten, Pemphigidae),
an aphid species producing pseudoscorpion-like soldiers on bamboos, Konryli. 49, 355, 1981.
4. Aoki, S., Yamane, S., and Kiuchi, M., On the biters of Astegopteryx styracicola (Homoptera, Aphidoidea),
Kontyri, 45, 563, 1977.
5. Bell, G., The Masterpiece of Nature: the Evolution and Genetics of Sexualiry, Croom Helm, London, 1982.
6. Bell, G., Recombination and immortality of the germ line, J. Evol. Biol., 1, 67, 1988.
7. Benton, T. G. and Foster, W. A., Altruistic housekeeping in a social aphid, Proc. R. Soc. Lond. B., 247, 199,
1992.
8. Blackman, R. L., Biological approaches to the control of aphids, Phil. Trans. R. Soc. Lond. B., 274, 473,
1976.
9. Blackman, R. L., Early development of the parthenogenetic egg in three species of aphids (Homopten:
Aphididae), Int. J. Insect Morphol. Embryol.. 7, 33, 1978.
10. Blackman, R. L., Stability and variation in aphid clonal lineages, Biol. J. Linn. Soc., 11, 259, 1979.
11. Blackman, R. L., Chromosomes and parthenogenesis in aphids, in Insect Cytogenetics, Blackman, R. L.,
Hewitt, G. M., and Ashbumer, M,, Eds., Symp. Roy. Entomol. Soc. Lond.. 10, 133, 1980.
12. Blackman, R. L., Species, sex and parthenogenesis in aphids, in The Evolving Biosphere. Forey, P. L., Ed.,
Cambridge University Press, Cambridge, 1981, 75.
13. Blakely, N., Biotic unpredictability and sexual reproduction: do aphid genotype-host genotype interactions
favour aphid sexuality?, Oecologia (Berlin), 52, 396, 1982.
14. Bonnet, C., Trait6 d'insectologie ou Observations sur les Pucerons. Chez Durand, Paris, 1745.
15. Briggs, J. B., The distribution, abundance, and genetic relationships of four strains of the rubus aphid
(Amphorophora rubi (Kalt)) in relation to raspberry breeding, J. Hort. Sci., 40, 109, 1965.
16. Brough, C. N. and Dixon, A. F. G., Follicular sheath (ovarian sheath) structure in virginoparae of the vetch
aphid, Megoura viciae Buckton (Homopten: Aphididae), Int. J. Insect Morphol. Embryol., 18, 217, 1989.
17. Brough, C. N. and Dixon, A. F. G., Reproductive investment and the inter-ovariole differences in embryo
development and size in virginopame of the vetch aphid, Megoura viciae. Entomol. Exp. Appl., 52,215,1989.
140 Insect Reproduction

18. Brough, C. N. and Dixon, A. F. G., Intraclonal trade-off between reproductive investment and size of fat
body in the vetch aphid, Megoura viciae Buckton, Funct. Ecol., 3, 747, 1989.
19. Brough, C. N., Dixon, A. F. G., and Kindlmann, P., Pattern of growth and fat content of somatic and
gonadal tissues of virginoparae of the vetch aphid, Megoura viciae, Entomol. Exp. Appl., 56, 269, 1990.
20. Browne, R. A., Population genetics and ecology of Artemia: insights into parthenogenetic reproduction,
TREE, 7, 232, 1992.
21. Buda, V. G., Mozuraitis, R. L., and Myarchaitis, V. P., Parthenogenesis and signal behavior of the
butterflies Lithocolletis emberizaepennella Bouche (Lepidoptera Gracillariidae), Dokl. Acad. Nauk SSSR.
319, 512, 1991.
22. Bulmer, M. G., Cyclical parthenogenesis and the cost of sex, J. Theoret. Biol., 94, 197, 1982.
23. Charlesworth, B., Reply to Bulmer on the cost of sex with cyclical parthenogenesis, J. Theoret. Biol., 94,
223, 1982.
24. Carvalho, G. R., Maclean, N., Wratten, S. D., Carter, R. E., and Thurston, J. P. O., Differentiation of
aphid clones using DNA fingerprints from individual aphids, Proc. Roy. Soc. h n d . B, 243, 109-1 14, 1991.
25. Clark, W. C., The ecological implicationsof parthenogenesis, in Perspectives in Aphid Biology, Lowe, A. D.,
Ed., Enromol. Soc. New Zeal. Bull., No. 2, 103, 1973.
26. Cognetti, G., Citogenetica della partenogenesi negli Afidi, Arch. Zool. Ital., 46, 89, 1961.
27. Cuellar, O., Animal parthenogenesis, Science. 197, 837, 1977.
28. Dickson, R. C., Development of the spotted alfalfa aphid population in Nolth America, Int. Kong. Entornol.
(Wien 1960), 2, 26, 1962.
29. Dixon, A. F. G., Fecundity of brachypterous and macropterous alatae in Drepanosiphum dixoni (Callaphididae,
Aphididae), Entomol. Exp. Appl.. 15, 335, 1972.
30. Dixon, A. F. G., Seasonal changes in fat content, form, state of gonads and length of adult life in the sycamore
aphid Drepanosiphum platanoids (Schr.), Trans. Roy. Entomol. Soc. Lond., 127, 87, 1975.
31. Dixon, A. F. G., Aphid Ecology. Blackie, Glasgow, 1985.
32. Dixon, A. F. G., Individuals, populations and patterns, in Individuals, Populations and Patterns in Ecology.
Leather, S. R., Watt, A. D., Mills, N. J., and Walters, K. F A., Eds., Intercept, Andover, 1994, 449.
33. Dixon, A. F. G., Burns, M. D., and Wangboonkong, S., Migration in aphids: response to current adversity,
Nature, 220, 1337. 1968.
34. Dixon, A. F. G. and Dharma, T. R., Number of ovarioles and fecundity in the black bean aphid, Aphis fabae.
Entomol. Exp. Appl., 28, 1, 1980.
35. Dixon, A. F. G. and Glen, D. M., Morph determination in the bird cheny-oat aphid, Rhopalosiphum padi
L., Ann. Appl. Biol.. 68, 11, 1971.
36. Dixon, A. F. G. and Howard, M. T., Dispersal in aphids, a problem in resource allocation, in Insect Flight:
Dispersal and Migration, Danthanarayana, W . ,Ed.,Springer-Verlag, Berlin, 1986, 145.
37. Dixon, A. F. G. and Wellings, P. W., Seasonality and reproduction in aphids, Int. J. Invertebr. Reprod., 5,
83, 1982.
38. Dunn, J. A., The survival in soil of apterae of the lettuce root aphid, Pemphigus barsarius (L.), Ann. Appl.
Biol., 47, 766, 1959.
39. Eastop, V. F., Worldwide importance of aphids as virus vectors, in Aphids as Virus Vectors, Harris, K . F. and
Maramorosch, K., Eds., Academic Press, New York, 1977, 4.
40. Elton, C., Animal Ecology and Evolution, Oxford University Press, Oxford, 1930.
41. Fakatsu, T. and Ishikawa, H., Soldiers and male of an eusocial aphid Colophina a m lack endosymbiont:
implication for physiological evolutionay interaction between host and symbiont,J. Insect Physiol., 38,1033,1992.
42. Fenchel, T., Intrinsic rate of natural increase: the relationship with body size, Oecologia (Berlin), 14, 317,
1974.
43. Fuyama, Y., Genetics of parthenogenesisin Drosophila melanogaster. XI. Characterizationof a gynogenetically
reproducing strain, Genetics, 114, 495, 1986.
44. Geiler, H., Eichhornchen, Sciurus vulgarisfuscoater Altum. 1876, als Vertilger von Blattlausen (Aphidae),
Saugentierkundl. Mitt., 4, 13, 1956.
45. Hille Ris Lambers, D., Polymorphism in Aphididae, Annu. Rev. Entomol., l l , 47, 1966.
46. Howard, M. T. and Dixon, A. F. G., The effect of plant phenology on the induction of alatae and the
development of populations of Metopolophium dirhodum (Walker), the rose-grain aphid, on winter wheat,
Ann. Appl. Biol., 120, 203, 1992.
47. Hurst, G. D. D., Majerus, M. E. N., and Walker, L. E., Cytoplasmic male killing elements in Adalia
bipunctata (Linnaeus) (Coleoptera: Coccinellidae), Heredity, 69, 84, 1992.
48. Huxley, T. H., On the agamic reproduction and morphology of Aphis - Part 1, Trans. Linn. Soc., 22, 193,
1858.
49. Ishikawa, H., Biochemical and molecular aspects of the aphid endocytobiosis, in Insect Endocytobisis:
Morphology, Physiology, Genetics, Evolution, Schwemmerl, W. and Gassner, G.. Eds., CRC Press, Boca
Raton, 1989, 123.
Parthenogenesis in Insects 141

50. Innes, D. J. and Herbert, P. D. N., The origin and genetic basis of obligate parthenogenesis in Daphnia
pulex, Evolution. 42, 1024, 1988.
51. Iwasa, Y., Pessimistic plant: optimal growth schedule in stochastic environments, Theor. Popul. Biol.. 40,
246, 1991.
52. Janzen, D. H., What are dandelions and aphids? Am. Nat., l l l, 586, 1977.
53. Judge, F. D., Overwintering in Pemphigus bursarius (L.), Nature, 216, 1041, 1967.
54. Kindlmann, P. and Dixon, A. F. G., Developmental constraints in the evolution of reproductive strategies:
telescoping of generations in parthenogenetic aphids, Funct. Ecol.. 3, 531, 1989.
55. Kindlmann, P. and Dixon, A. F. G., Optimum body size: effects of food quality and temperature, when
reproductive growth rate is restricted, with examples from aphids, J. Evol. Biol., 5, 677, 1992.
56. Knight, R. L. and Alston, F. H., Pest resistance in fruit bleeding, in Biology in Pest and Disease Control,
Price Jones, D. and Solmon, M. E., Eds., Blackwell, Oxford, 1974, 73.
57. Kundu, R. and Dixon, A. F. G., Evolution of complex life cycles in aphids, J. Anim. Ecol., 64, 246, 1995.
58. Lamb, R. Y. and Willey, R. B., Are parthenogenetic and related bisexual insects equal in fertility?Evolution,
33, 774, 1979.
59. Leather, S. R., Wellings, P. W., and Dixon, A. F. G., Habitat quality and the reproductive strategies of the
migratory morphs of the bird cheny-oat aphid, Rhopalosiphum padi (L.) colonizing secondary host plants,
Oecologia (Berlin), 59, 302, 1983.
60. Lees, A. D., The production of the apterous and alate forms in the aphid Megoura viciae Buckton, with special
reference to the role of crowding, J. Insect Physiol., 13, 289, 1967.
61. Lloyd, D. G., Benefits and handicaps of sexual reproduction, Evol. Biol., 13, 69, 1980.
62. Lynch, M., Destabilizing hybridization, general-purpose genotypes and geographic parthenogenesis, Q. Rev.
Biol.. 59, 259, 1984.
63. Mackenzi, A. and Dixon, A. F. G., An ecological perspective of host alternation in aphids (Homoptera:
Aphidinea: Aphididae), Entomol. Gener., 16, 265, 1991.
64. Marescalchi, O., Pijnacker, L. P., and Scali, V., Cytology of parthenogenesisin Bacillus whitei and Bacillus
lynceorum (Insecta Phasmatodea), Invertebr. Reprod. Develop., 20, 75, 1991.
65. Markkula, M., Studies on the pea aphid Acyrthosiphonpisum Harris (Hom. Aphididae) with special reference
to the difference in the biology of the green and red forms, Ann. Agric. Fenn., 2, 1, 1963.
66. Matsuka, M. and Mittler, T. E., Enhancement of alata production by an aphid, Myzus persicae, in response
to increases in day length, Bull. Fac. Agric. Tamagawa Univ. Tokyo, 18, 1, 1978.
67. Maynard Smith, J.,The evolution of recombination,in The Evolution of Sex, Michod, R. E. and Levin, B. R.,
Eds., Sinauer, Sunderland, MA, 1988, 106.
68. Meglitsch, P. A. and Schram, F. R., Invertebrate Zoology, 3rd ed., Oxford University Press, Oxford, 1991.
69. Mordvilko, A. K., Zur Biologie und Morphologie der Pflanzenlause (Fam. Aphididae Pass.), Horae Soc.
Entomol. Ross., 31, 1, 1900.
70. Mordvilko, A. K., Zur Biologie und Morphologie der Pflanzenlause (Fam. Aphididae Pass.), Horae Soc.
Entomol. Ross., 33, 162, 1901.
7 1. Mordvilko, A. K., The evolution of cycles and the origin of heteroecy (migration) in plant-lice, Ann. Mag.
Nut. Hisr.. 2, 570, 1928.
72. Mordvilko, A., On the evolution of aphids, Arch. Naturgesch. N.S., 3, 1, 1934.
73. Mukai, T., The genetic structure of natural populationsof Drosophila melanogaster. 1. Spontaneous mutation
rates of polygenes controlling variation, Genetics, 50, 1, 1964.
74. Miiller, F. P., Biotypen und Unterarten der "Erbsenlaus" Acyrthosiphon pisum Hams, Z. Pflanzenkr.
Pjlanzenschutz. 69, 129. 1962.
75. Muller, H. J., The relation of recombination to mutational advance, Mutation Res.. 1, 2, 1964.
76. Newton, C. and Dixon, A. F. G., The cost of switching from asexual to sexual reproduction in an aphid,
Entomol. Exp. Appl.. 47, 283, 1988.
77. Nielson, M. W. and Don, H., A new virulent biotype of the spotted alfalfa aphid in Arizona, J. Econ.
Entomol., 67, 64, 1974.
78. Owen, R., On Parthenogenesis or the Successive Production of Procreating Individuals from a Single Ovum.
John van Voorst, London, 1849.
79. Pagliai-Bonvicni, A. M. and Cognetti, G., Partenogenesi e variabilita genetica negli afidi, Mem. Soc.
Entomol. Ital., 60, 63, 1982.
80. Puterka, G. J. and Peters, D. C., Inheritance of greenbug virulence to Gb2 and Gb3 resistance genes in
wheat, Genome, 32, 109, 1989.
81. Puterka, G. J. and Peters, D. C., Sexual reproduction and the inheritance of virulence in the greenbug,
Schizaphis graminum (Rondani), in Aphid-Plant Genotype Interactions. Campbell, R. K . and Eikenbary,
R. D., Eds., Elsevier, New York, 1990, 289.
82. Ruvinsky, A. O., Perelygin, A. A., Lobkov, Yu. I., and Belyaeu, D. K., Factors organizing and maintaining
polymorphism in a cyclic parthenogenetic species: Daphnia pulex, Heredity, 57, 15, 1986.
142 Insect Reproduction

83. Shaw, M. J. P., Effects of population density on alienicolae of Aphis fabae Scop. 1. The effect of crowding
on the production of alatae in the laboratory, Ann. Appl. Biol.. 65, 191, 1970.
84. Sibly, R. M. and Calow, P., Physiological Ecology of Animals. An Evolutionary Approach, Blackwells
Scientific Publications, Oxford, 1986.
85. Stalker, H. D., On the evolution of parthenogenesis in Lonchoptera (Diptera), Evolution, 10, 154, 1956.
86. Stouthamer, R., Luck, R. F., and Hamilton, W. D., Antibiotics cause parthenogenetic Trichogramma
(Hymenoptera/Trichogrammatidae)to revert to sex, Proc. Natl. Acad. Sci. U.S.A.. 87, 2424, 1990.
87. Suomalainen, E., Saura, A., and Lokki, J., Evolution of parthenogenetic insects, Evol. Biol., 9,209, 1976.
88. Suomalainen, E., Saura, A., and Lokki, J., Cytology and Evolution in Parthenogenesis, CRC Press, Boca
Raton, FL, 1987.
89. Taylor, L. R., Migration and the spatial dynamics of an aphid, Myzuspersicae, J. Anim. Ecol., 46.41 l, 1977.
90. Templeton, A. R., The prophecies of parthenogenesis, in Evoluh'on and Genetics of Life Histories, Dingle,
H. and Hegmann, P., Eds., Springer, Hamburg, 1982, 75.
91. Ueda, N. and Takada, H., Differential relative abundance of green-yellow and red forms of Myzus persicae
(Sulzer) (Homoptera: Aphididae) according to host plant and season, Appl. Entomol. Zool., 12, 124, 1977.
92. Uyenoyama, M. K., On the evolution of parthenogenesis: a genetic representation of the "cost of meiosis,"
Evolution, 38, 87, 1984.
93. Van Valen, L., A new evolutionary law, Evol. Theory. I, 1, 1973.
94. Wagner, G. P. and Gabriel, W., Quantitative variation in finite parthenogenetic populations: what stops
Muller's ratchet in the absence of recombination, Evolution, 44, 715, 1990.
95. Walker, F., Remarks on the migration of Aphides, Ann. Mag. Natur. Hist., 1, 372, 1848.
96. Walters, K. F. A., Brough, C., and Dixon, A. F. G., Habitat quality and reproductive investment in aphids,
Ecol. Entomol., 13, 337, 1988.
97. Walters, K. F. A. and Dixon, A. F. G., Migratoly urge and reproductive investment in aphids: variation
within clones, Oecologia (Berlin). 58, 70, 1983.
98. Ward, S. A. and Dixon, A. F. G., Spreading the risk, and the evolution of mixed strategies: seasonal variation
in aphid reproductive biology, Adv. Invertebr. Reprod., 3, 367, 1984.
99. Watt, A. D. and Dixon, A. F. G., The role of cereal growth stages and crowding in the induction of alatae
in Sitobion avenae and its consequences for population growth, Ecol. Entomol., 6, 441, 1981.
100. Weber, G., On the ecological genetics of Sitobion avenae (F.) (Hemiptera, Aphididae). Z. Angew. Entomol.,
100, 100, 1985.
101. Weber, G., On the ecological genetics of Metopolophium dirhodum (Walker) (Hemiptera, Aphididae), 2
Angew Entomol.. 100,451, 1985.
102. Wellings, P. W., Leather, S. R., and Dixon, A. F. G., Seasonal variation in reproductive potential: a
programmed feature of aphid life cycles, J. Anim. Ecol., 49, 975, 1980.
103. Went, D. F., Paedogenesis in the dipteran insect Heteropeza pygmaea: an interpretation, Int. J. Invertebr.
Reprod., 1, 21, 1979.
104. Went, D. F., Parthenogenetic strategies in insect reproduction, Adv. Invertebr. Reprod., 3, 303, 1984.
105. White, M. J. D., Animal Cytology and Evolution, 3rd ed., Cambridge University Press, Cambridge, 1973.
106. Williams, G. C., Natural Selection: Domains, Levels, and Challenges, Oxford University Press, New York,
1992.
107. Wratten, S. D., Reproductive strategy of winged and wingless morphs of the aphids Sitobion avenae and
Metoplophiurn dirhodum, Ann. Appl. Biol., 86, 319, 1977.
108. Yagamuchi, H., Biological studies on the todo-fir aphid Cinara todicola Inouye with special reference to its
population dynamics and morph determination, Bull. Gov. For. Exp. Sm. Tokyo, 283, 1, 1976.
109. Zchori-Fein, E., Roush, R. T., and Hunter, M. S., Male production induced by antibiotic treatment in
Encarsia formosa (Hymenoptera: Aphelinidae), an asexual species, Experentia, 48, 102, 1992.
 Chapter 7

FACTORS AFFECTING FECUNDITY. FERTILITY.

OVIPOSITION. AND LARVIPOSITION IN INSECTS
Simon R Leather .
CONTENTS
I . Introduction ............................................................................................................... 144

I1. Fecundity and Fertility ..............................................................................................144

A . Ovariole Number ................................................................................................ 144
B . The Role of Nutrition ......................................................................................... 145
1. Larval Nutrition and Adult Size .................................................................... 145
a. Adult Size ................................................................................................. 146
i. Size and Fecundity .............................................................................. 146
ii. Size and Fertility ................................................................................. 147
2 . Adult Nutrition ............................................................................................... 147
a. Adult Longevity and Fecundity ................................................................ 148
b . Adult Longevity and Fertility ................................................................... 148
3. Host Plant Quality .......................................................................................... 149
a. Nitrogen .................................................................................................... 149
b. Seasonal Patterns in Foliar Nitrogen Levels ............................................ 150
C . Age at Mating ..................................................................................................... 151
D . Copulation Frequency ......................................................................................... 152
1. Sex Ratio ........................................................................................................ 153
E. Abiotic Factors .................................................................................................... 154
1. Temperature ................................................................................................... 154
2. Photoperiod .................................................................................................... 155
3. Humidity ......................................................................................................... 155
F. Flight and Dispersal ............................................................................................ 156

I11. Oviposition and Larviposition .................................................................................. 158

A . Deterrents and Attractants .................................................................................. 159
B . Offspring Fitness ................................................................................................. 159
1. Maternal Choice ............................................................................................. 160
2 . Host Plant Phenology ..................................................................................... 162
3. Clutch Size ..................................................................................................... 163
a. Adult Life Span ........................................................................................ 163
b . Host Quality .............................................................................................. 164
4 . Offspring Size ................................................................................................ 164

IV . Conclusions ............................................................................................................... 166

Acknowledgment ................................................................................................................. 166

References ........................................................................................................................... 166

.
0-8493-6695-X/95/50.00+5.50
Q 1995 by CRC Press Inc .
144 Insect Reproduction

I. INTRODUCTION
The production of its offspring is arguably the most important event in the life span of an
insect. It is affected by a number of factors, many of which have already been discussed in
the preceding chapters. This chapter examines critically the role that reproductive load and
host plant quality have on the reproductive patterns of herbivorous insects. Most of the
examples used will refer to the Lepidoptera and the Hemiptera, as these two groups have been
extensively studied and the strategies displayed are representative of those shown by insect
groups that deposit eggs and live young, respectively. This does not imply that other insect
groups will be ignored; examples from other groups will be discussed to demonstrate how
similar the strategies employed by insects are, regardless of phylogenetic classification.

11. FECUNDITY AND FERTILITY

Insect fecundity and fertility, the number of offspring produced and the viability of those
offspring, are affected by a number of factors and these are discussed at length below. The
number and viability of offspring produced by an individual insect is of great importance to
the population dynamics of the species as a whole as well as to the individual concerned. The
evolution of reproductive strategies (Chapter 10)and specific oviposition behaviors are results
of the factors that affect fecundity and fertility, although to a certain extent, oviposition
behavior can influence fecundity and is intrinsically linked with reproductive strategies. There
is some confusion in the literature between potential fecundity and achieved (realized)
fecundity. Potential fecundity can be defined in a number of ways, but is most often some
measure of the reproductive capacity of an insect that is measured before the reproductive
event occurs, e.g., number of eggs in the reproductive tract. Achieved fecundity is the actual
number of offspring produced by an insect during its life span. The difference between
potential and achieved fecundity can be substantial. Achieved fecundity depends upon poten-
tial fecundity and various ecological factors, particulary nutrition.I0 Nutrition affects size,
which is one of the major factors affecting fecundity (see II.B.1.a.i). The number of ovarioles
within an insect has a large bearing on the potential fecundity of insects and it is useful to
discuss the likely implications of variability in this characteristic before dealing with any other
factor.

A. OVARIOLE NUMBER
Insects are characterized by the possession of two ovaries within which are a number of
ovarioles. Ovariole number in insects is not always a constant for either family or species. In
some groups, e.g., Lepidoptera, it is nearly always a constant, with eight being the norm?7
however, in some rare individuals of the butterfly Heliconius charitonius, six, seven, and nine
ovarioles have been reported.60Ovariole number also varies slightly in the butterfly Colias
philodice eriphyle: individuals with eight, nine, and twelve ovarioles have been observed.1ss
No explanation for this phenomenon has been sought and it is so rare as to have no bearing
on reproductive strategies. There is considerable variation between species and within species
in other groups, e.g., Aphididae and D i ~ t e r a . ~Within
~ . ' ~ ~those insect species with a variable
number of ovarioles, ovariole number is positively related to
For some insect groups such as the Diptera, the number of ovarioles is positively related
to the weight of the The number of ovarioles in the mosquito Aedes punctor is
strongly correlated with wing length (an indicator of size), but wing length is less well
correlated with achieved fecundity.168In other insects, e.g., Melanoplus grasshoppers, ovariole
number is variable but is strongly related to size. In addition, the number of ovarioles is equal
to the number of eggs laid in a pod, so ovariole number in this case is a good indicator of
achieved fecundity.12a
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 145

Other insects, such as aphids, show no satisfactorily demonstrated significant relationship

between weight and ovariole number,132,21L.221 but the more ovarioles an individual possesses,
the more fecund that individual is likely to be.132Thus, within the same ovariole number class,
large individuals are more fecund than smaller ones, but a female with six ovarioles and the
same weight as one with ten ovarioles would be less fecund than the latter. Ovariole number
in aphids is both a function of generational morph,L39,221 and of host quality. Ovariole number
in the majority of the aphid species studied is high at the beginning of the life cycle, namely
those generations emerging from eggs or, as in the case of Rhopalosiphum padi, in the second
generation from the egg. Ovariole number decreases as the season progresses, but shows an
increase in the autumnal generations. The morphs associated with the highest host quality
available, expanding buds and leaves in the spring and senescing leaf tissue in the autumn, are
those with the greatest numbers of o v a r i o l e ~By. ~ having
~ ~ ~ ~a large flexibility in reproductive
potential, these aphids are able to fully exploit this high host quality and rapidly increase their
numbers before the leaves mature and host quality decreases or the leaves drop. Aphids
exploiting plants with a low host quality tend to have fewer ovarioles and to be less fe-
cundL32.139 (see below).

B. THE ROLE OF NUTRITION

Nutrition, i.e., the quality and amount of food experienced by an insect during larval and
sometimes adult life, has a major effect on both fecundity and fertility. In some insects, e.g.,
within the Lepidoptera, it has been considered that adult nutrition is of little importance to
fecundity and that the major influence is from nutrition afforded to the larvae. Other insects,
e.g., are considered to be affected by both adult and juvenile nutrition. The following
sections consider how both adult and larval nutrition affect fecundity and fertility and also
consider how substances not generally thought to be nutritious can also affect these life history
parameters.

1. Larval Nutrition and Adult Size

The importance of larval nutrition cannot be underestimated. Whether the insect feeds as
an adult or not, the size that it attains as an adult is dependent on the nutrition experienced by
an immature larva. The size (weight) an adult insect achieves is generally a good indicator of
its fecundity. It must be noted, however, that although adult size is almost invariably correlated
with the individual's potential fecundity, it is not always correlated with achieved fecundityL2'
(see 1.a). It is those factors which affect both potential and achieved fecundity that will be
discussed below.
If the stored product moth Cadra cautella is crowded as a larva, then fecundity can be
reduced by as much as 67%.8'Even more dramatic responses to larval nutrition can be seen
in other Lepidoptera. For example, when larvae of Heliothis zea were fed on corn, 92% of the
eggs that were produced by the subsequent adult females were fertile. However, if reared on
cotton squares, adult females were totally infe~-tile.'43
The sawfly Diprion pini is strongly affected by the host plant as a larva; even clones of the
same species of host plant have a strong effect on larval performance. For example, when
reared as a larvae on four different clones of Pinus sylvestris, the fecundity of the subsequent
adults was highly affected. Those host plants that allowed good larval growth and survival
were associated with large, highly fecund fern ale^.^ Larval feeding is also very important in
determining ovarian development and subsequent fecundity of the sawfly Pristiphora
erichsonii?'
In general, if insects are reared under poor conditions of larval nutrition, e.g., crowding or
poor-quality food plants, then the adults that arise are smaller and tend to be less fecund than
those adults arising from larvae developing under better conditions. For example, larvae of the
chrysomelid beetle Chrysomela knabi, when reared on a less suitable willow host, e.g., Salix
146 Insect Reproduction

nigra under low moisture conditions, were smaller and less fecund as adults than when reared
on S. h u m ~ l i sLarval
. ~ ~ competition in the predatory stoneflies Megarcys signata and Kogatus
modestus results in the production of small adults that are less fecund than the large adults that
arise when larvae are reared under low density conditions.17'

a. Adult Size
The size that an insect achieves as an adult is dependent on a number of factors, many due
to larval nutrition (see above). The size of the adult insect has many important life history
implications, ranging from effects of adult life span, mating success, fecundity, etc. In the
following sections, the effect that adult size has on fecundity and fertility are discussed. Other
effects of size on insect reproduction are discussed elsewhere, e.g., Chapter 9.
i. Size and Fecundity
Until recently it was believed that for most arthropods increased weight and/or size resulted
in increased f e c ~ n d i t y . ' ~ Certainly,
.~'~ this has been reported for many insects, especially
aphids. However, this relationship is not as straightforward as might at first appear.l2I Many
studies have used potential fecundity (i.e. the number of eggs or offspring within an insect at
the adult moult) as the index of fecundity for their estimate of lifetime fecundity. In those
insects that have produced all their eggs or embryos by the time adulthood is reached, such
as in the pine looper moth Bupalus piniarius,lR and the sycamore aphid Drepanosiphum
platanoidis, I4O this is indeed an adequate prediction. However, in species where ovulation does
not cease after the adult moult and the number of steps between the dependent variable
(offspring number) and independent variable (weight) is increased, then this relationship
becomes less robust. For example, using embryo or large embryo counts as indicators of
fecundity in aphids is a common p r a c t i ~ e . ~In~an. ~aphid
~ . ~such
~ as R. padi, in which ovulation
continues after the adult moult and the initial fecundity estimate can be increased by 135%,'14
then this method would produce a highly erroneous result.
The sizetfecundity relationship is affected by a number of factors, many of which can be
linked either directly or indirectly with host quality. The effect of host quality on this
relationship can act through the larval stage as well as on the reproductive adult stage.
Moreover, even if large insects of the same species are more fecund than smaller ones of the
same species, confounding factors can occur. For example, large adults of the sawfly Neodipnon
sertifer produce more eggs than small adults, but fewer of their eggs are fertile.89In some
aphids (e.g., Aphis fabae), the relationship between size and fecundity is indirect and acts
mainly through reproductive rate and mortality.197That is, small aphids have a faster rate of
reproduction than large aphids, but do not live as long and are thus less fecund. In Lepidoptera,
even more confusion can arise. For example, adult weight in the pine beauty moth Panolis
j7ammea is a good indicator of the potential fecundity.Il5However, this has little bearing on
the number of eggs that are actually laid, with the achieved fecundity sometimes being as little
as 1% of the potential fecundity.135In fact, fewer than 15% of laboratory reared P. flammea
lay even 70% of their maximum potential number of eggs.12' In field conditions this is likely
to be even lower,33and the maximum fecundity estimate for field P. flammea is 28% of its
maximum reproductive p0tentia1.l~~In other Lepidoptera, for example the moth Epiphyas
posmittana, there is a good correlation between adult weight and achieved fecundity when the
moths are fed, but not if they are starved.83
As an adjunct to the above it is interesting to consider the effect of male size on female
fecundity. For example in Drosophila melanogaster, females that mate with small males have
a greater fecundity than those that have mated with large males. This apparently contradicts
the theory that large males are fitter than small males and enjoy greater mating success
(Chapter 9), but it appears that small males copulate longer and transfer more sperm than large
males.'77
Factors Affecting Fecundity, Fertility, Oviposition, and Lurviposition in Insects 147

ii. Size and Fertility

The size of a female insect can have marked effects on the fertility of the eggs laid. For
example, large individuals of the codling moth Cydia pomonella lay more fertile eggs than
~ I nbutterfly Pararge aegeria, small eggs tend to be infertile, but since small
small ~ n e s . ~ the
eggs also tend to be laid at the end of the reproductive period, it may be that female age
influences egg fertility rather than egg weight per se.224This area requires further study.

2. Adult Nutrition
Adult insects can be divided into two main categories - those that feed after becoming
adult and those that do not feed, or if they do, feed only minimally. The former are usually
long lived and obtain the nutrients required for reproduction as adults. The latter are usually
short lived and have laid down their reserves for reproduction during larval life, as is the case
with many Lepidoptera. There are, of course, exceptions. Aphids, for example, feed during
adult life but are generally short lived, and larval (nymphal) experience plays a large part in
how they function as adults.55Some adult Lepidoptera feed on pollen and even blood61.115 and
not solely on carbohydrates.
Adult feeding has for some time been thought only important in those species of insects that
are relatively long lived as an adult or which require a specific food source unobtainable as
a larvae to commence reproduction, e.g., the blood meal of a female mosquito. Insect groups
such as the Lepidoptera, where larval development is long and reproductive reserves are laid
down during the larval period, have been considered to be essentially nonfeeding as adults
despite the fact that many species are known to be nectar feeding.23
However, significant effects in Lepidoptera have been demonstrated in several studies. For
example, carbohydrate intake has been shown to increase longevity and egg production in the
butterfly Coliasphilodice eurytheme, the moth Euxoa messoria, and the checkerspot butterfly
Euphydiyas editha.30.160.'91 In addition, the pine beauty moth P. jlammea, when given access
to water, is twice as fecund as when deprived of a water source, and if given a carbohydrate
source is three times as fecund.Il5E. messoria is more than three times more fecund when fed
on honey solution than when deprived of water.30 The potato tuber moth Phthorimaea
operculella lays almost four times as many eggs as unfed moths when fed on sucrose
~olution,~' and the moth Heliothis virescens shows a 50% increase in fecundity when fed on
a sucrose s01ution.l~~ If H. virescens is unable to obtain nectar in the field, its reproductive
capacity decreases as the moth ages.224a Like P. flammea, if fed it is able to maintain its body
~ ~field, the spruce budworm Choristoneura
weight and devote its reserves to r e p r o d u c t i ~ nIn. ~the
fumiferana enhances adult fecundity by feeding on honeydew.1s0
Adult feeding in Lepidoptera is thus important in determining realized fecundity. Unfed
adult E. postvittana are 30% less fecund than moths that have access to normal levels of adult
nutrition, but interestingly the fertility of the eggs is unaffected, suggesting that extra nitrogen
is not required as an adult.83Similarly, the moth Zeiraphera canadensis lived longer when fed
on a sugar solution and was more fecund than those fed on water alone. However, the eggs
of fed moths had a fertility of only 84% compared with one of 95% in the unfed moths. As
overall fecundity was increased, the fed moths laid 88 fertile eggs per female compared with
61 fertile eggs per female for unfed moths.29Since the oviposition period was increased but
not the oviposition rate, it is possible that the moths ran out of sperm, and, not being in a
position to mate for a second time, started to lay infertile eggs. In the case of P. jlammea, the
oviposition period was also increased by adult feeding, but no increase in the daily oviposition
rate was seen.lI5
The armyworm Spodoptera exempta is unable to realize its potential fecundity without
water to achieve hydration and maturation of the o o c y t e ~In . ~addition,
~ S. exempta is able to
utilize supplementary lipid and protein reserves following carbohydrate uptake and this results
148 Insect Reproduction

in increased f e c ~ n d i t yIt. ~might,

~ therefore, be thought that a nitrogen supplement to the diet
of adult Lepidoptera would result in increased fecundity. However, adding amino acids to the
diet of the butterfly Euploea core actually resulted in a decrease in fecundity, whereas sucrose
resulted in an increase.92Insects that are predators as adults must supplement their nitrogen
reserves. The predatory bug Anthocoris confusus, although able to lay eggs even when fed at
suboptimal rates, are most fecund when fed on aphids at a rate of 6 a p h i d ~ l d a yOn
. ~ ~the other
hand, another aphid predator, the green lacewing Chrysoperla carnea was shown to be almost
twice as fecund in the absence of prey as when prey was present, although there was no
difference in preoviposition ~ e r i 0 d . l ~ ~

a. Adult Longevity and Fecundity

Longevity is a further complicating factor in the sizelfecundity relationship for a number
of insect species. For example, in the planthopper Prokelisia marginata, a significant relation-
ship between eggs produced and adult female weight has been dem~nstrated."~ However, this
relationship was obtained by dividing total fecundity by total longevity and using the figure
obtained as eggs per female per day. This is not strictly correct, as the pre- and postreproductive
periods were ignored and both are of great importance in determining insect f e ~ u n d i t y . ~ ~ ~ . ~ ~ ~ . ' ~ ~
Longevity and the factors affecting longevity appear to be the most important factors
influencing achieved fecundity in the Lepidoptera. In P. flammea there is no relationship
between weight and achieved fecundity, but a strong positive correlation between longevity
and achieved fecundity exists.Il5 In addition, there is no relationship between longevity and
weight, but a significant relationship between availability of adult food source and longev-
ity.lI5 In Euxoa messoria there is a significant relationship between pupal weight and both
achieved fecundity and longevity. However, if the moths are fed, the relationship between
weight and longevity disappears, although longevity and achieved fecundity remain corre-
latede30Increased fecundity has also been found to be correlated with increased longevity in
H. virescens and Sesamia n o n ~ g r i o d e s , ~and . ' ~ ~with the availability of an adult food source
in the pierid butterfly C. philodice eurytheme.'gl Larval host plants can also affect longevity
and this in turn affects the achieved fecundity of S. nonagriodes and P. flammea in both the
laboratory and the field.3,116.135 It is interesting to note that virgin moths lived longer than those
that had been forced to mate in conditions when suitable oviposition sites were not avail-
able.135Those mated moths that delayed mating because of being presented with a poor host
quality sacrificed life span in favor of producing fewer, but fitter eggs.Iz7

b. Adult Longevity and Fertility

The phenomenon of a decline in offspring fitness as the mother ages is not confined to
insects; human beings show the same general relationship. In aphids, (which, like humans,
give birth to live young) although there is no evidence that the incidence of malformed and
dead offspring increases as the mother reaches the end of her lifespan, it has been suggested
that small aphids may resorb their embryos to a greater extent than larger aphids.59Certainly
at high temperatures the frequency of aphid still births is increased.76
In Lepidoptera a decline in the fertility of the eggs produced as the mother ages is a
common occurrence. Whether this is a result of sperm depletion or the inability of the mother
to provision all her eggs is an open question. Fertility of the eggs of the moth C. cautella
declines with increasing female age, from 80%at day 1 to 20%by day 8.x1The fertility of eggs
of the butterfly P. aegeria also decrease with the age of the mother.224In the moth E.
postvittana egg fertility also changes with the age of the mother, but unlike the previous two
examples, the relationship is not linear. Egg fertility is greatest halfway through the oviposi-
tion period, and is lowest at the beginning and at the end of the lifespan of the moth.41In the
moth Choristoneura murinana, the color of the eggs is indicative of when in the reproductive
period of the mother they have been laid; they are green at the beginning of the reproductive
period and yellow at the end, and at the same time the fertility decreases from 96 to 65%.62
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 149

3. Host Plant Quality

Host quality can be defined as those plant attributes, chemical or physical, that contribute
either negatively or positively to the fitness of the insect population or individual that feeds
upon the plants tissues.'25
Host quality can affect the fecundity of an herbivorous insect in two ways; it can affect the
fecundity (achieved and potential) of the adult arising from the larvae feeding on it, or actually
affect the achieved fecundity of the adult insect meeting that individual plant for the first time.
The latter case is almost certainly just as common as the former.
The fecundity of the spear-marked black moth, Rheumaptera hastata, which feeds on Pinus
nigra, was affected in two ways by the quality of its host plant. On hosts that had previously
been attacked by conspecifics, the sex ratio was skewed in favor of males, and those females
that were produced were less fecund than those produced on undamaged host plants.222In this
case damaged plants were of poorer quality than undamaged plants. The fecundity of the pine
processionary moth Thaumetopoea pityocampa was also affected by host plant variation in
this way.1°
The grain aphid, Sitobion avenue, is more fecund on the ears of cereal plants than on other
parts of the plant, and that is where it is most often found.2J6The bird cherry aphid, R. padi,
is more fecund on young grass and cereal plants than on mature plants, and on mature plants
it is most fecund on the stem, and this too, is where it is most often found.53.129 The cabbage
aphid, Brevicoryne brassicae, is also strongly affected by host quality, being apparently most
fecund on leaves high in n i t r ~ g e n . ~ ~ ~ . ~ ~ ~
In times of nutrient stress, the aphid Metopolophium dirhodum reduces embryo growth and
resorbs the youngest embryos, so as to safeguard immediate reproduction for when conditions
improve.82When the quality of the host plant improves, embryo formation begins anew. Host
quality greatly affects the fecundity of adult aphids, both in terms of what happens to adults
arriving on a novel host plant, and in terms of what they were reared upon. An aphid reared
on a poor-quality host plant but feeding on a good-quality host plant will be more fecund than
it would have been if it had remained on its original host plant. This has been demonstrated
with R. padi, transferred from oats to Timothy grass, Phleum pratense.111-122 A further
complication occurs depending on the morph of the mother (whether winged or not), the
offspring of alate mothers being more fecund than the offspring of apterous mothers. The
effect of maternal experience is also seen in Lepidoptera. Larvae of the small ermine moth,
Yponomeuta evonymellus, survive and develop better when transferred as neonate larvae from
poor host plants to good host plants, but do not develop as well as those larvae that have been
transferred from good-quality plants to similarly good-quality plants. The reverse is also true,
larvae transferred from good-quality plants to poor-quality plants still develop faster than
those larvae reared on poor-quality plants for their entire developmental period.13'

a. Nitrogen
Nitrogen in its various forms is without doubt the most important single foodstuff to an
insect, being as it is, the basis for the building blocks of life, protein. Hence, as well as having
many important effects on fecundity and fertility per se, it also affects reproductive strategies
directly and indirectly, by influencing host preferences, life span, etc.Iz5The debatable point,
particularly when considering herbivorous insects, is the way in which the nitrogen content
of a particular food plant is measured, and how what is being measured affects the insect in
question. Plant nitrogen content is a major determinant of host quality, although not solely so.
Host plant quality affects herbivorous insects in a number of ways, many of which have been
reviewed elsewhere.lZ5
One aspect of food quality, the level of nutrient nitrogen, is known to be important in
several species. For example, the host quality of larval food plant affects the fecundity of the
adult gypsy moth, Lymantria dispar. Those adults arising from hosts with low nitrogen levels
are less fecund than those reared on high nitrogen level hosts.% Hosts plants that are high in
150 Insect Reproduction

nitrogen result in more fecund individuals of the armyworm, S. f r u g i p e r d ~ . ' ~ ~

Spruce bud-
worm larvae that are reared on old, nitrogen-poor foliage, develop into small adults of reduced
fecundity when compared to those arising from larvae reared on young, nitrogen-rich foliage.21
When reared on the grass Holcus lanatus with artificially enhanced nitrogen levels, the
leafhoppers Dicranotropis hamata, Elymana sulphurella, Encelis incisus and Zyginidia
saitellaris were all more fecund as adults than those reared on plants with low levels of
nitrogen.178The collembolan Folsomia candida was more fecund when fed on a host rich in
nitrogen.205Adults of the cinnabar moth Tyria jacobaeae were larger and produced more eggs
in areas where their host plant Senecio jacobaea had high nitrogen levels. In addition, larval
survival was better on plants high in nitrogen than on those plants which contained low levels
of foliar nitrogen.207
Nitrogen rich plant parts are important to the plant, and they are often protected by high
levels of plant secondary compounds, e.g., tannins.'O In some instances, plant secondary
compounds are also important in determining insect fecundity. The fecundity of the scale
insect, Fiorinia externa, is increased by terpenoid alcohols as is that of another scale insect,
Nuculaspis tsugae, both of which feed on species of T ~ u g a . However,
'~~ the fecundity of F.
externa is negatively correlated with acyclic terpenoids, whereas that of N. tsugae is positively
correlated with the same compound^.'^"

b. Seasonal Patterns in Foliar Nitrogen Levels

Seasonal changes in plant nitrogen levels have had a marked effect on the evolution of
insect reproductive strategies. This is particularly true in the case of sap-feeding insects, which
feed on an extremely dilute solution and have nitrogen acquisition as one of their major
objectives. With nitrogen at a premium it is in sap-feeding insects that the importance of host
quality to migratory and reproductive strategies may be most easily demonstrated. As herba-
ceous host plants age, their quality as a food source as measured by aphid performance in
terms of growth and fecundity, shows a decline. This is seen in several species; for example,
the asparagus aphid Brachycorynella asparagi is less fecund on old leaves than it is on young
ones,227and the flower thrips Frankliniella occidentalis also shows a variable fecundity
depending on host growth stage.203
The soluble nitrogen content of plant tissues varies throughout the year.52,53,98.220 In the
temperate climates of spring, when the leaves are growing rapidly, foliar nitrogen content is
high. This falls during the summer months as the leaves mature, and rises again (at least in
the phloem) in the autumn, as the leaves begin to senesce and nitrogen reserves are mobilized
within the plant in preparation for leaf fall.
As would be expected, this pattern of events has affected the life history strategies of a
number of tree-dwelling aphids. For example, the bird cherry-oat aphid R. padi, a host
alternating aphid, feeds and reproduces on its primary host Prunus padus while soluble
nitrogen levels are high in the spring.53As soluble nitrogen levels fall, winged forms are
produced in response to this deterioration in host quality and these winged forms migrate to
their secondary graminaceous hosts where host quality is higher.130In autumn, as host quality
on these grass hosts declines, they return to their primary host whose quality has once again
i m p r o ~ e d . The ~ ~ Uroleucon gravicorne, although not strictly a host alternating aphid,
~ ~ Japhid
usually moves from its overwintering host Solidago spp., to plants of several annual Erigeron
species. Like R. padi, it responds to changes in soluble nitrogen levels of its host plant and
when nitrogen levels are low, reproductive performance is also low. Soluble nitrogen content
declines throughout the season on Erigeron spp. and growth and developmental rates, as well
as the fecundity of U, gravicorne, decrease at the same time.Is8
The sycamore aphid D. platanoidis also has to face the same deterioration in host quality
as that faced by the host alternating R. padi. However, instead of migrating to a host plant of
a higher host quality status, D. plantanoidis reduces its metabolic rate and enters a summer
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 151

aestivation until the host quality of its food plant once again becomes suitable for growth and
. ~ ~ ~ ~ ~tree-dwelling aphid, the birch aphid Euceraphis punctipennis, has
r e p r o d u ~ t i o nAnother
resolved the same problem by exploiting the fact that birch trees (Betula spp.) have a certain
number of shoot tips that produce new leaves throughout the spring and summer months.
These are high in soluble nitrogen content, and the adult birch aphids track these growing
points throughout the season, moving within the tree and even between trees when necessary,
in order to deposit their offspring on suitable food Aphid species on annual hosts
are also greatly affected by the changes in soluble nitrogen concentration which occur as their
hosts age. The cabbage aphid B. brassicae shows both a steady decline in growth rate as their
Brussels sprout host plants age and a corresponding decline in reproductive c a p a ~ i t y . ~ ~ ~ . ~ ~ ~
Not all sap feeders show this same dependence on young flushing tissue or nitrogen-rich
senescent tissue. However, in all cases a nitrogen-rich feeding site is required. Even within the
aphid species already discussed, some differences in reproductive strategy are shown. The
aphid R. padi is frequently found on older cereal plants in the summer months, but is habitually
found on the basal parts of the stem where its growth and reproductive rates are highest.I2"his
is also the area of the mature cereal plant where nitrogen levels are high.94
In the case of other sap suckers such as the delphacid P. marginata, which feeds on the
same plant species (Spartina spp.) the whole year, a response is still seen to soluble nitrogen
levels. The insects move to vegetation growing along the sides of streams, which is of a higher
quality in the spring and early summer, and return to overwinter on the nutritionally less
suitable marshland Spartina where greater protection is afforded.43Thus, like the birch aphid,
these insects track their food plant resources. The psyllids, Psylla peregrina and Psylla
subferruginea, also show this ability to track nitrogen-rich resources. In spring, they are
closely associated with the expanding buds of their host Crataegus monogyna. After bud burst
is completed, the only sources of high nitrogen flush are in the growing shoots and inflores-
cences, and it is on these points that the psyllids aggregate.'" Insects that feed throughout the
season on one portion of their host plants, such as the membracid Publilia reticulata on
Veronia noveboracensis, must either increase their feeding rate to maintain the same level of
activity, go into summer aestivation as in D. platanoidis, or restrict their feeding to that portion
of the leaf that affords the best nutrition. In the case of P. reticulata, feeding is confined to
the major cross-veins where sap flow is brisk; this enables the insect to maintain a high
reproductive rate.25Other membracids with relatively long generation times have hatching
times such that nymphal development takes place at those times of the year when soluble
nitrogen levels are high. The oak tree hopper Platycotis vitata has two generations a year, one
in spring and the other in autumn, while adult females diapause in the winter and summer
depending on their generation, and the eggs are laid in spring and autumn respectively to take
advantage of the flushes in soluble nitrogen in their host trees.'03
Among chewing insects, a large number of species have adopted flush feeding strategies
to achieve high growth rates which lead to increased size and greater fecundity. A problem
faced by flush feeding insects is that young plant tissue is often protected by high levels of
secondary metabolites, which must be detoxified by the young larvae. The winter moth
Operophtera brumata, feeding on oak, is a bud feeder.'O Others begin feeding on flush foliage
and then move onto older, more abundant tissues once they have successfully survived the
early, highly vulnerable establishment phase. For example, newly hatched larvae of the pine
beauty moth P. flammea feed on young, newly expanding needles, but then move onto the
previous year's growth to complete their devel~pment,~'~ only returning to the current year's
foliage in the late summer if no older needles are

C. AGE AT MATING
The age at which an insect mates can have profound influences on a number of life history
parameters of the female, for example, the life span of females of the pine beauty moth
 Insect Reproduction

-0 2 4 6 8 10 12 14 16
Age at mating (days)

FIGURE 1. The effect of age at mating on the fecundity of the moths Busseola fusca, m, Panolisflammea, +, and
Pectinophora gossypiella, *. (Based on References 135, 141, and 204).

"
0 2 4 6 B 10 12
Age at mating (days)

FIGURE 2. The effect of age at mating on the fertility of Busseolafusca, M, Chilo partellus, *, Panolisflammea,
+, and Pectinophora gossypiella, 0.(Based on References 15, 135, 141, and 204.)

P. jlammea is reduced by a delay in mating.13sMany lepidopteran species display protandry,

i.e., the males emerge first. It has been proposed that this ensures that females are mated
sooner rather than later and thus minimizes any detrimental effects likely to follow a delay in
mating.74+223 Certainly, in many species of Lepidoptera, a delay in mating results in both a
decrease in achieved fecundity (Figure 1) and also in fertility (Figure 2). Interestingly, P.
jlammea exhibits protogyny, i.e., the females emerge before the males, but this is probably a
response to the unsettled weather conditions likely to be experienced by this insect in early
spring as virgin females that experience a delay in mating realize more of their potential
fecundity than those females that mate and then experience a delay in oviposition due to cold
weather.135
A delay in mating has a fairly straightforward effect on the fertility rate of Lepidoptera
(Figure 2), i.e., the greater the delay, the fewer fertile eggs that are laid. However, fecundity,
despite showing this same general pattern (Figure I), can, as the period of delayed mating is
extended, show a slight increase; mating imposes a cost in females in term of life span and
also causes an increase in prereproductive period as the eggs are fertilized.12' However, if the
number of fertile eggs is calculated, it can be seen that the delay in mating does indeed reduce
effective fecundity. It is thus in the interests of a female insect to mate as quickly as possible.

D. COPULATION FREQUENCY
Copulation frequency can have marked effects on achieved fecundity in many insects. Most
work has been done on Lepidoptera and accordingly most of the examples presented are from
Factors Ajfecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 153

that group. Multiple mating is common in a number of lepidopteran species and this is seen
as an investment, in terms of nutrient transfer from the males to the females receiving the
spermat~phores~~ (Chapter 10). However, in other species there is sperm competition and
although multiple mating may take place, the female oviduct may be blocked by the first
corner's spermatoph~re.'~~ In the field, as one might expect, fertility is generally high,
suggesting that the strategies used by the insects are successful. For example, a 3-year study
(1956-1958) of the moth Rhyacionia buoliana, showed that egg fertility was 99.64,94.5 and
10096, respecti~ely.~~
In the black swallowtail butterfly Papilio polyxenes asterius, multiple matings increased
both the fecundity and fertility of the femalesL40and this was also the case in the moth E.
p o s ~ i t t a n a In
. ~ other
~ Lepidoptera, multiple matings are required to keep female fecundity at
a constant rate; for example, if females of Euphydryas chalcedona are only mated once, then
egg output steadily declines over the life span of the adult.'a5In other species, the relationship
is not simple; for example, in Adoxophyes orana, one or two extra copulations increases the
fertility of the eggs laid but more than this number of copulations results in a reduction of
fertilit~.~"In yet other species, multiple mating has no effect on fecundity, e.g., Danaus
plexippus.lg5 In fact, in one study which involved 103 different species of butterfly from
around the world, no correlation between multiple mating and high egg counts was found.64
Multiple mating in the crickets Gryllodes sigillatus and Gryllodes veletis, offset the costs of
reproduction incurred by the females, and also increased fecundity as a result of nutrient
tran~fer.~'
Copulation frequency on the part of the male may also affect female achieved fecundity.
The more times male H. virescens have mated, the less sperm they transfer and the more
infertile eggs the female will produce.g0This is also seen in the codling moth C. pomonella.
The more times a male has mated, the more infertile eggs that the female produces, but the
more times a female has mated, the more eggs she will prod~ce."~ The females, therefore, have
to take the risk that by multiple mating they increase their fecundity, but that at the same time
their chance of encountering a male that has mated more than once also increases, as does their
chance of laying infertile eggs. In some species, the females are able to assess the mating
condition of the male and mate more often when they have copulated with males who have
repeatedly mated, e.g., H. charitonius females are able to assess that the spermatophores are
smaller than normal and therefore contain less nitrogen.22

1. Sex Ratio
The sex ratio perceived by an insect can have marked effects on its fecundity and fertility.
The skewing of sex ratio is basically the idea behind pheromone disruption schemes as it is
felt that if the proportion of males is reduced then at least the fertility, and hopefully the
fecundity, of the females remaining in the population will be reduced.11,38.107.225h
The results are not always straightforward. For example, the pine beauty moth P.flammea,
which occurs in a 1:l ratio in nature, is actually most fecund when the females are outnum-
bered by males 3: 1 (Figure 3). On the other hand, the greatest proportion of fertile eggs are
produced when the sex ratios are equal (Figure 4). If the appropriate calculation is done, then
it can be seen that more fertile eggs are produced overall when the females are present in the
same numbers as the males. This apparently strange strategy may be an adaptation to the fact
that the females of P. jlammea, unlike many other Lepidoptera, emerge before the males.126
In the Sphingid moth Manduca sexta, the situation is reversed, here females were most fecund
at a sex ratio of 1:1 (Figure 3) but their fertility decreased linearly over the range. Thus, the
greatest number of fertile eggs was produced at a sex ratio of five males to one female (Figure
3). In another Lepidopteran species, Spodoptera litura, (unfortunately not tested over sex
ratios where males were in excess) fecundity declined in much the same way as the previous
two but fertility increased with increasing female bias (Figure 3). In another
lepidopteran species, Plodia interpunctella, the number of unmated females was recorded in
 Insect Reproduction

Sex ratio (%females)

FIGURE 3. The effect of sex ratio on the fecundity of Heliothis virescens, 0,Manduca sexta, m and Panolis
flammea, m. (Based on References 84, 123, and 192.).

01 I
0 20 40 60 80 100
Sex ratio (%females)

FIGURE 4. The effect of sex ratio on the fertility of Epiphyas postvinana, m, Manduca sexta, *, Panolisflammea,
+, and Spodoprera litura, 0.(Based on References 41, 123, 166, and 192).

respect to sex ratio. Over the range of six males to each female to an even sex ratio, all females
were successfully mated. However, once the ratio was biased in favor of females, the
proportion of unmated females rose to 20%. This was not a linear response.*6
Sex ratio thus has marked effects on female fecundity and fertility of eggs deposited by
adult Lepidoptera. This is likely to apply to other insect groups.

E. ABIOTIC FACTORS
1. Temperature
Temperature is perhaps the most important abiotic factor influencing living organisms. It
is particularly important in affecting insect life history parameters such as developmental
period, life span, pre-reproductive delay, fecundity, and fertility, as well as flight and behavior.
Basically speaking, all insects show the same response to temperature: there is a lower
threshold below which reproduction does not occur and an upper threshold beyond which
reproduction ceases. Fecundity increases between these two thresholds up to an optimal point,
usually linearly, with the turnover point of the curve being related to the normal climatic
distribution of the insect (Figure 5). For example, aphids from temperate regions have an
optimal temperature for reproduction at about 20°C whereas aphids from subtropical regions
have an optimal temperature for reproduction and development at 25OC. This is also seen
within other groups, e.g., the moth S. nonagriodes has an optimum temperature for reproduc-
tion at 21°C while P. flammea has an optimum temperature for reproduction at 18OC (Figure
5). The response shown between the lower threshold and the optimal point can be extremely
linear; for example, in P. flammea, the correlation between temperature and egg hatch over
this range is 99.99%.136
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 155

f rnn

Temperature oC

FIGURE 5. The effect of temperature on the fecundity of insects. (Acyrthosiphon pisum, 0. Diuraphis noxia, *,
+
Ostrinia nubilalis, , Panolisflammea, B, Rhopalosiphum padi, 0,Sesamia nonagriodes, *. (Based on References
1, 3, 80a, 104, 11 1, and 116.)

Fertility can also be affected by high temperatures; for example, the number of malformed
and still born nymphs born to aphids increases as the upper temperature threshold is ap-
pr~ached.~~

2. Photoperiod
Photoperiod is one of the most reliable indicators available for insects to distinguish
forthcoming changes in season. It is thus not surprising to find that photoperiod has marked
effects on insects. In aphids, photoperiod affects the morph pr~duced~~.'~'' so that the morphs
leading on to the diapause stage are initiated; it acts as signal for diapause preparation in the
moth Cotesia r ~ b e c u l a land
~ ~ also for several butterfly and beetle species133.
Photoperiod also has marked effects on the reproductive behavior of insects, in particular,
aphids. In the case of aphids, this is more a result of the fact that as day length decreases,
aphids produce sexual and sexual-producing morphs which are generally less fecund than the
spring and summer forms.55Temperature modifies the effect of photoperiod and the response
to temperature and photoperiod is not the same for all aphid species. For example, the aphid
Myzus persicae is more fecund under long day lengths at 24°C than at 18°C. Brachycaudus
helichrysi is more fecund under short day conditions at 18°C than at 24°C but the situation
is reversed under long day conditions.227aHowever, the reproductive activity of the aphid
Hyperomyzus lactucae at 22OC is unaffected by changes in ph~toperiod.'~~
The reproductive behavior of Lepidoptera is also affected by photoperiod. For example, the
moth Plutella maculipennis will only copulate in the dark but oviposition behavior is unaf-
fected by light conditions.lsl However, fecundity in this species is affected by the photoperiod
experienced by the adult. For example under short day conditions, i.e., 12 h light, the mean
fecundity is only 37 eggs per female, whereas under long day conditions (16 h light), fecundity
is twice that.87The photoperiod experienced by moths during their larval development can also
affect their fecundity. For example both C. pomonella and Grapholithafunebrana are more
fecund when exposed to long day lengths during their larval periods than if they are exposed
to short day length^.^'
Photoperiod is thus an important factor in determining the fecundity of an insect species,
but as photoperiod is seasonally and geographically determined, the effects are inextricably
linked with particular stages of the life cycle that are seasonally adapted to a particular
photoperiod.

3. Humidity
Humidity has been shown to have marked effects on insect fecundity and fertility, although
the results are not always straightforward. For example, apterous (nonwinged) adults of the
 Insect Reproduction

2001 ' 1
40 50 60 70 80 90
Relative Humidity

FIGURE 6. The effect of relative humidity on the fecundity of Heliothis zea at 21.I0C, 8 , 26.7"C. 0,and 32.6OC,
*. (Based on Ellington, J.J. and El-Sokkari, A., S. W. Entomol., 1 1 , 177, 1986.)
aphid R. padi were shown in choice chamber experiments to prefer more humid conditions
(>70% RH).Il1However, in experiments in which fecundity was measured, the aphids were
shown to be more fecund at 35% RH.117a The experimental host plants on which the aphids
were reared all received adequate water, and it is possible that this somehow altered host plant
quality so as to benefit the aphids feeding on plants at low relative humidities.
Adults of the moth H. zea are more fecund at higher relative humidities (Figure 6) than at
lower relative humidities and the percentage hatch of its eggs is also greater at higher relative
humidities than at lower.65However, this may not be a result of reduced fertility at the lower
relative humidities but rather an expression of the ability of the eggs to survive desiccation.

F. FLIGHT AND DISPERSAL

As discussed earlier, migration, whether it be by active flight, ballooning or by walking
must impose a cost on reproduction, if only because of the energy used for this purpose rather
than for offspring production. Foraging and oviposition behavior, which are components of
flight and dispersal, are also affected by the egg load of the mother. In general, females with
a high egg load search more intensively, accept hosts of a lower quality, spend less time
searching the host that they land on, and, in gregarious species, lay larger egg clutches.153
It is interesting to note that in many insects the decision to invest in a migratory form is
usually caused by a deterioration in host quality. For example, the light brown apple moth E.
postvittana produces smaller adults when the food available to the larvae is suboptimal. These
smaller insects have a significantly lower wing loading than large individuals and are more
adapted to dispersal and presumably the ability to locate additional suitable or better quality
hosts for their o f f ~ p r i n g .This
~ ~ . ~is~seen in a number of other lepidopteran species. The
velvetbean caterpillar Anticarsia gemmatalis, if deprived of food for any period during its
larval development, produces adult moths with a low wing loading and consequently greater
powers of dispersal. The fall armyworm S.frugiperda, the corn earworm Helicoverpa zea, and
the armyworm Spodoptera latifascia, also show this response to larval ~tarvation.~
One of the more obvious responses to changes in host quality evinced by those insect
species which exhibit alary polymorphism, is the switch from apterous nonwinged forms to
winged forms. As well as being influenced by changes in photoperiod and temperature, wing
formation in these insects is also influenced by changes in food quality and by rearing density.
Crowding during development in nongregarious species is usually an indicator that host
quality is likely to decline in the near future and the production of a winged dispersive form
in these circumstances is adaptive. This change to the production of winged forms is common
in many species of Homoptera. For example, many planthoppers, including P. marginata,
produce winged forms in response to nymphal crowding, as do many species of aphid^.^^,^^ It
would be reasonable to expect that insects capable of dispersal respond to a deterioration in
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 157

host quality by producing either a specifically adapted morph, or utilizing a set of behavioral
patterns to escape adverse conditions. This then would maximize reproductive potential while
operating under the constraints of crowding or poor food. To illustrate this hypothesis,
examples of a few, but well-documented examples from the literature will be used.
In those aphid species where the possession of wings is the norm, migratory flight is taken
when host quality deteriorates locally. For example, the sycamore aphid D. platanoidis
responds to crowded conditions during development by leaving its host plant with little or no
reproduction taking place, preferring to reproduce on an unexploited resour~e.~' In those
aphids that show alary polymorphism, the winged morphs also colonize habitats of a higher
host quality than that on which they were reared. On arrival at the new host plant they produce
apterous offspring which are able to exploit the new habitat effecti~ely.~~' These apterae are
also able to detect and anticipate deterioration in their habitat through cues such as crowding
and lowered foliage nitrogen levels, they, in turn produce alatae and are able to fly to other
hosts.57
Host location is, however, a risky process, and although the quality of their present host
plant may be deteriorating, it will not always become totally unsuitable for reproduction and
feeding. Thus, it is not to the advantage of the species as a whole for all of the alatae to disperse
from the current host plant. A flexible response should be an advantage. This is certainly the
case within some aphids. Winged R. padi and S. avenue are produced with a variable number
of o v a r i ~ l e s ,that
l ~ ~is, the reproductive investment of the offspring of an apterous mother on
a host plant of deteriorating quality is not a constant. The reproductive investment of aphids
is linked with their migratory strategies. Alate aphids with a small number of ovarioles take
off more readily, do so at a steeper angle, and delay wing muscle autolysis for longer than
those aphids with a greater number of ovarioles. In addition, those aphids with a lower number
of ovarioles possess more chemosensory organs and are thus better able to locate host plants.
Thus, those individuals with a low reproductive investment are better suited to long distance
dispersal than those with a greater reproductive investment.210
In addition to these differences, aphids with smaller reproductive investments are able to
withstand starvation for longer periods of time than those with a large reproductive invest-
ment.210Moreover, aphids with a large number of ovarioles (i.e., greater reproductive invest-
ment) are more likely to die before reaching the adult stage.213The number of individuals
within each ovariole class produced by the mother is adjusted so as to maximize the fitness
of their offspring. Thus, on poor quality hosts where survival of apterous forms is likely to be
low, a higher proportion of those that are born have a low number of ovarioles when compared
to those born on high quality hosts.211Alate aphids landing on high-quality hosts produce
proportionately more offspring with a large number of ovarioles than do the wingless apterae.
In addition, when aphids autolyze their wing muscles, their reproductive output becomes
similar to that of apterous morphs.55
When apterous aphids come under the influence of nutritional stress, they are less able to
avoid the effects than winged forms. They are able to disperse to a certain extent by walking,
but this is likely to be costly in terms of time, and the aphid is likely to undergo a period of
starvation. The black bean aphid A. fabae can survive starvation as an adult for up to 3 days.
This was shown to reduce life time fecundity and longevity. However, to compensate for this
potential loss of fitness, the adult aphids increased their rate of reproduction above normal,
when favourable host conditions became a~ai1able.l~~
After being produced in response to host adversity, active winged insects migrate in search
of new habitats. Not all the plants selected by migrants are of the same quality, and the
postalighting response of a migrating insect is influenced by features of the host it has landed
on. Characteristically, a complex suite of responses is found in herbivorous For
example, it has been shown that prealighting behavior often depends on different cues than
oviposition behavior. The butterflies Eurema brigitta and Eurema herla, which are monopha-
gous on Cassia mimosoides, land on plants with leaves of similar shapes, size, and color. They
158 Insect Reproduction

then use chemical and textural cues to confirm their choice of host, leaving for a further period
of search if the appropriate oviposition stimulants are not received.'44Similar or more complex
patterns occur in other plant-feeding insects. In aphids, plants are probed and sampled; the
decision to remain is a result of the balance between food plant stimuli, resources remaining
for migration, and those available for reproducti~n.~~ This is itself strongly influenced by the
availability of wing muscles that can be autolyzed after settling, as is common in many
species. In the aphid R. padi, alates that land on poor quality hosts are relatively less likely
to autolyze their wing muscles, thus giving themselves the option to produce a few offspring
and still make a further migratory flight.'38It is noticeable that the emigrants of this species,
which are produced in early summer when habitat quality is generally higher, are less variable
in terms of ovariole number and do not show this adaptation.I3$
The seasonal adaptations of aphids to their expected environment are best demonstrated by
looking at two studies involving the bird cherry aphid R. padi. The dispersal and reproductive
strategies of the three winged morphs can be compared. These are (1) the emigrants (arising
from the primary host and migrating to the secondary host in spring), (2) the alate exules
(which migrate between secondary hosts during the summer), and (3) the gynoparae (arising
from the secondary host and migrating to the primary host in a ~ t u m n ) . ~Several
~ . ' ~ ~dispersal
and reproductive characteristics have been examined. The emigrants, which migrate into a
high-quality habitat that is nitrogen-rich with rapidly growing graminaceous hosts, have a high
reproductive investment with most of their embryos in an advanced stage of development and
wing muscles that tend to autolyse sooner, rather than later. The alate exules on the other hand,
which are produced in times of adversity, i.e., crowded nymphal development or low host
plant quality) and are likely to be surrounded by equally poor-quality hosts, have a low
reproductive investment and wing muscles that are not programmed to autolyse quickly. They
are thus able to delay wing muscle autolysis if the host on which they land is of a poorer quality
than that from which they emigrated. The environment to which they migrate is unpredictable
and they are adapted to this unpredictability. The gynoparae, on the other hand, which are
migrating to a less common host, (of good, but temporally limited quality due to the immi-
nence of leaf fall), have a low fecundity but produce their offspring very quickly once they
have settled. Their fat reserves are high which is appropriate, as they may take some time to
locate a suitable host. Each morph appears to be well adapted to fit the expected environment.
Although aphids are so versatile, they are not the only insects that can respond to host
quality in such a flexible manner. The pine beauty moth P.flammea is also able to assess host
quality by monitoring the monoterpene composition of the plant and it will avoid
poor quality hosts if possible. If the choice of host plant is restricted, then the female moths
will eventually lay eggs on the low-quality hosts, but only after a prolonged prereproductive
delay.135Among butterflies, if the reproductive cost of dispersal is low, then many species will
move between host patches so as to maximize offspring fitness. This has been shown in the
cabbage butterfly Pieris rapae, which, after initiating oviposition on a suitable host plant, will
eventually leave for another host patch when its supply of chorionated eggs is exhausted.34
Habitat quality, that is, the relative proportion of suitable host plants, also affects dispersal.
The checkerspot butterfly E. editha, for example, will leave patches with few or none of its
preferred host, Pedicularis semibarbata, but will remain in those patches where P. semibarbata
is ab~ndant.'~~Thus, herbivorous insects respond to detrimental changes in the quality of their
host plants in a number of different ways, all of which are likely to maximize their fitness and
that of their offspring. After dispersal has occurred, the insect is faced with another problem:
the exploitation of the host to which it has migrated.

111. OVIPOSITION AND LARVIPOSITION

Upon arrival at its new host, a reproducing female insect must deposit its young, be they
immobile eggs or mobile larvae. A number of decisions have to be taken at this stage and these
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 159

are affected by the plant, not just in terms of its suitability as a host but also its attractiveness
to the female insect. Although it is in the interest of the insect to produce as many offspring
as possible in as short a time as possible, its responses are mediated by the quality of the host
plant. The relationship between oviposition preference in insect herbivores and their offspring
performance is recorded to range from good to poor, although most evidence points to a good
c o r r e l a t i ~ n . ~The
~ ~reasons
. ~ ~ . ~for
~ ~why this relationship is not as good as human observers
would expect is probably linked to how preference and performance are related genetically,
and has been fully reviewed e l s e ~ h e r eThe . ~ ~concept of perceived host quality has been
discussed in some detai1,lz5but it is graphically illustrated by the oviposition behavior of the
two chrysomelid beetles, Phratora vitellinae and Galerucella lineola, which both feed on
Salix spp. G. lineola lays its eggs on species of Salix which are low in salicylates and on which
the larvae are able to grow and develop better than if they were feeding on salicylate-rich
plants. P. vitellinae, on the other hand, lays its eggs on willow species high in salicylates on
which its larvae are at a disadvantage in terms of growth and development. However, the
larvae of P. vitellinae have a better defense against predators as they are able to sequester
salicylates from their host plant and produce a secretion which is high in these distasteful
compounds, when attacked. Larvae of G. lineola and P. vitellinae feeding on hosts with low
concentrations of salicylates are, however, virtually defen~eless.~~ It is also felt that although
the correlation between oviposition preference and offspring performance is not perfect, the
majority of insects requiring specialist hosts can depend on plant chemistry in order to select
the most suitable host for the development and survival of their o f f ~ p r i n g . ~ ' . ' ~ ~

A. DETERRENTS AND ATTRACTANTS

It is probable that one of the major ways in which an insect assesses the quality of a
particular plant is by the volatile compounds released by the tissues of that plant. These
compounds can act as either deterrents or attractants, or also as oviposition stimulants. It
would be expected that those compounds acting as deterrents would be correlated with poor
herbivore performance and that those acting as attractants would be positively correlated with
herbivore performance. However, evidence is mounting to indicate that deterrents are not
always correlated with negative postingestion effects as would logically be expected.16 In-
stead, it has been suggested that avoidance responses to these chemicals have evolved under
selection pressures that are not related to herbivore performance in respect to the deterrent.17
Insects with a wide host range appear to be markedly affected by some of these compounds,
e.g., iridoid glycosides, which often have strong deterrent effects.la2
The situation with those compounds that act as oviposition stimulants andlor attractants is
somewhat more clear-cut. It is possible that certain volatile chemicals, although not directly
acting as nutrition sources, are correlated with the presence or absence of true nutrients, such
as protein and amino acids. For example, the spruce budworm C. fumiferana is stimulated to
oviposit on host plants by the presence of the monoterpenes alpha and beta pinene,laYas is the
pine beauty moth P. jlammea which lays more eggs on pine plants with high beta to alpha
pinene ratio^."^.^^^ The pyralid moth Dioryctria amatella lays more eggs on pine trees
containing high levels of monoterpene~.~~ The cinnabar moth, T.jacobaeae on the other hand,
lays its eggs on plants high in organic nitrogen and sugars, once it has orientated on a host plant
using the secondary compounds as primary stirnulant~.~~' The butterfly P. rapae lays its eggs
on tall plants high in nitrogen, although the primary stimulants are various glucosinolates,~44
and the monarch butterfly is primarily stimulated to oviposit by the presence of cardenolide
g1u~osides.l~~

B. OFFSPRING FITNESS
Offspring fitness can be measured in a number of ways, but generally speaking, high larval
growth rates, along with rapid larval development and high larval survival (leading to a large
fecund adult) are indicative of high fitness and a good quality host plant.
160 Insect Reproduction

1. Maternal Choice
There is much debate as to whether egg distribution on a single host plant indicates sites
where offspring fitness is enhanced. A recent review suggests that ovipositing insects are
relatively poor at discerning suitable host plants for their offspring,35and this is supported to
some extent by other authors.200However, the real area of debate is whether plant chemistry
is the major determinant of host plant selection and offspring fitness. The suggestion that
deterrents may have little to do with overall offspring fitnessL6is indeed a tenable hypothesis,
but chemical attractants do appear to be more positively correlated with offspring f i t n e s ~ . ~ ~ ~ , ~ ~ ~
It has been suggested that rather than host plant chemistry (equalling quality) being the
criterion for host plant selection, other abiotic factors may be the determining measure.35
Moore, Myers, and Eng,L56 working with the western tent caterpillar Malacosoma califomicum
pluviale, suggested that microclimate was more important in determining host selection than
host quality. They questioned the idea that egg distribution reflected host quality variation
within the host tree, as they found that eggs were laid mostly on the sunny side of the tree.
From this they suggested that it was not host chemistry that determined oviposition site but
temperature. However, leaves on the sunny side of the tree had higher nitrogen levels and the
male pupae produced from those leaves were heavier than those reared on leaves from the
shady side of the tree. It is thus possible that the female moths were somehow responding to
the nitrogen content of the leaves rather than to the microclimate.
Some insects show a relatively straightforward relation between host quality and host plant
selection; for example, adults of the leaf-folding sawfly Phyllocolpa spp., lay their eggs on
those Salix lasiolepis bushes on which their offspring will grow and develop fastest.77In other
insects, the relationship between the stimuli attracting the insect to oviposit and the suitability
of the host for offspring growth and survival is not so apparent. An excellent example of this
type of relationship between oviposition preference and host suitability is shown by the
monarch butterfly D. plexippus. Adult females are stimulated to oviposit by cardenolides in
their milkweed host plants. However, the larvae are adversely affected by very high levels of
cardenolides in individual Asclepias spp. Females lay their eggs on plants with intermediate
cardenolide content, which is also most suitable for the survival rate of their larvae.229Many
insects appear to be capable of making quite subtle choices of host in relation to plant
chemistry. The chrysomelid beetle Paroposis atomaria, for example, has larvae that perform
best in terms of growth and survival on Eucalyptus hosts that are high in nitrogen irrespective
of the content of essential oils and the adult beetles choose oviposition sites a~cording1y.l~~
The pine beauty moth P. jZammea is greatly affected by the quality of the pine trees it
encounters. Like many species of Lepidoptera, it is able to detect differences among hosts (the
cues used involve the monoterpene profiles of the plants, more eggs being laid on plants with
a high beta to alpha pinene ratio than on plants with a low and to select those
individual plants on which their offspring have a greater fitness in terms of growth and
I ~ ~ of both P.JZammea and Crocidosema plebejana
fecundity when they become a d ~ 1 t s . Adults
have been shown to have marked preferences for host plants that will maximize their capacity
for increase (Figure 7) and this clearly implies that females select their host plants in a way
that maximizes larval growth and survival. There is no evidence available yet concerning
oviposition preferences relative to phenology in the former case as the eggs are laid when the
plants are still dormant. Adults of the cinnabar moth, Tyria jacobaeae, which oviposit on
growing plants of tansy ragwort S. jacobaea, lay more eggs on individual plants with high
nitrogen contents,207and the noctuid moth Helicoverpa armigera uses cues such as flowering
status of the various host plants to determine its oviposition choices. Flower structures are
more suitable for larval growth and survival than vegetative structures due to their high
nitrogen content and low levels of alkaloids and other secondary corn pound^.^^
However, sometimes the ovipositing insect does indeed appear to be unable to distinguish
the host plant of higher quality. For example, larvae of the pine beauty moth P.flammea grow,
develop, and have higher survival rates on pine trees that are water stressed.2L7The adult
Factors Affecting Fecundity, Fertility, Oviposition, and Lantiposition in Insects 161

Host suitability

Figure 7. Relationship between preference index of adults of the moth PanolisJammea, m, and Crocidosema
plebejana, 0,and suitability of the tree for growth of their larvae. (Based on References 86a and 117.)

moths, however, although able to distinguish between pine trees of differing suitability
because of genetic differences or because they have previously suffered insect a t t a ~ k , I ~are ~J~'
unable to distinguish between plants of the same genetic material which are in different states
of water stress.217This may, however, be because the host plant stimuli on which P. flammea
relies upon to determine the quality of its host (monoterpenes) is unaffected by water stress.
On the other hand, some insects are more adept when faced with this situation. There is, for
example, a strong relationship between oviposition preference and larval performance in the
shoot-galling sawfly, Euura l a ~ i o l e p i sIn . ~this
~ case, plants that are under water stress are not
suitable hosts for the larvae of E. lasiolepis, and the adult females are able to distinguish
between stressed and unstressed ~ 1 a n t s .This l ~ ~ may be because E. lasiolepis lays its eggs
inside the host plant and can use its ovipositor to detect stimuli relating to host plant quality,
whereas P. jlammea lays its eggs externally on the needles and has a less intimate association
with the host plant.
Plants may appear suitable for offspring in one aspect, such as high nitrogen levels, but they
may have secondary disadvantages. For example, host quality can be reduced by the feeding
activity of other insects. The weevil Cyrtobagous salvinae and the moth Samea multiplicalis,
both feed on the aquatic plant Salvina molesta, and both prefer to lay eggs on buds high in
nitrogen. However, C. salvinae avoids plants that have been damaged, particularly if they have
been damaged by S. multiplicalis. The weevils also select their host plants by assessing the
number of buds available and avoiding plants with few buds so as to maximize the potential
food supply of their o f f ~ p r i n g .Other
~ ~ . ~insects
~ ~ show similar sensitivities. The bruchid beetle
Callosobruchus maculatus, which feeds on cowpea, lays its eggs in the field before harvest.
Younger, smaller pods are preferred as oviposition sites, and these are indeed more suitable
for the 1 a r ~ a e .Inl ~addition,
~ the female is able to determine whether another beetle has already
laid its eggs on a pod and to assess the number of eggs already present. The adult females avoid
laying eggs on pods with a high egg load and thus ensure that larval competition is reduced.
Similarly, the moth Cactoblasriscactorum, lays its eggs on those Opuntia plants that its larvae
grow well on. Unlike some insects, it lays more eggs on previously attacked plants than
unattacked plants. In this case, the signs of a successful previous attack may be an indicator
of a high quality host plant.l6I
The perception of host plant quality can be changed by the previous experience of an insect.
In a choice test, the leaf-mining fly Liriomyza trijolii preferred tomato plants which had a high
foliar nitrogen content on which their larvae survived and developed better than foliage which
was low in nitrogen. However, flies with no experience of high quality hosts (those high in
nitrogen), showed no preference at first. After exposure to a high quality plant, however, a
strong preference for high quality plants was evinced. It was postulated that in nature, L.
trijolii uses plants that maintain or increase in acceptability, and disperses relatively quickly
162 Insect Reproduction

from areas containing plants of a low acceptability (and quality). As conditions worsen and
host deprivation increases, then nutritionally subthreshold plants once again become accept-
able as host plants.lS2
It would appear from the evidence presented above that, on the whole, mother does indeed
know best and oviposition errors may be attributable to factors other than host quality - to
mothers making the best of a bad job or a result of factors unappreciated by the observer.
Whatever the reason or reasons, further study in this area is required.

2. Host Plant Phenology

Plant phenology also affects insect oviposition strategies. The tephritid flies Tephritis
bardanae and Cerajocera tussilaginis time their oviposition so that the establishment and
rapid growth of their early instar larvae coincides with the two main periods of physiological
activity of their host - the development of the flower heads of the burdock Arctium minus.
T. bardanae lays its eggs in the flower heads at flowering, and C. tussilaginis at achene
maturation. At both these times, nutrient availability is at a maximum and the young larvae
position themselves in those structures where the greatest nutrient flux occurs.193Another
tephritid, Eurosta solidaginis, samples the ramets of its host, Solidago altissimma, and lays its
eggs on those hosts at the phenological stage most suitable for the growth of its l a r ~ a e . ~
Adult leaf miners must be extremely sensitive to leaf quality since their offspring are
confined to one leaf for the whole of their development, and a poor selection will mean very
high mortality. In addition to the usual details of host selection, they may have to deal with
the fact that plants often show a defensive response to attack by leaf miners by shedding leaves
earlier than n ~ r r n a l For.~~ these
~ ~reasons,
~ ~ ~ as well as nutritional ones, the age of the leaf
which they lay their eggs on or in, is very important in oviposition choice. Lithocolletis
quercus, a miner of oak leaves, selects its leaves in a way that appears to minimize the chance
of early death for its larvae.' Other insects are constrained in their choice of host plant due to
lack of suitable leaves. Those that feed on deciduous trees in the autumn are in a particularly
difficult position. For example, the aphid Periphyllus califomiensis only colonizes those
maple trees whose leaves are orange-yellow. Trees with red leaves are avoided. Orange-
yellow leaves remain on the tree longer, and this ensures that the aphids are able to complete
their development before leaf fall.'* On the other hand, the bird cherry aphid R. padi, which
colonizes the bird cherry tree P. padus in autumn, shows no preference for different ages of
1 e a ~ e s .However,
I~~ it times the production of the morphs that colonize trees in the autumn in
response to the temperature it experiences in July. This cue is, in fact, an excellent predictor
of leaf In addition, although these autumn morphs do not feed as adults,113they are able
to distinguish the relative suitability of their tree hosts for their offspring's offspring. If the
host is unsuitable they will leave, if able to do so, or if unable (due to wing muscle autolysis),
they will produce offspring that lay fewer eggs on that particular tree.Il8
The tropical satyrid butterfly Mycalesis perseus lays its eggs on plants that are high in
nitrogen, but only chooses the youngest leaves on that plant as oviposition sites. It inhabits
savannah grassland, which is prone to premature drying, and this behavior ensures that eggs
are laid only on those grasses that are likely to live longest. This gives its offspring the
maximum possible time in which to deve10p.l~~ A similar scenario is seen in the oviposition
strategy of the butterfly E. chalcedona. In laboratory trials, the females prefer to lay their eggs
on the shrub Scrophularia californica. This is nutritionally superior in terms of larval growth
rates to an alternative host plant, the shrub Diplacus auranticus. However, D. auranticus is
more drought-tolerant than S. califomica, and in the wild E. chalcedona will lay a sizeable
proportion of its eggs on the former shrub. Although D. auranticus is nutritionally inferior, its
drought-tolerance means that it is more persistent in dry years and means that in those years
larval survival will be assured. Thus host use reflects a tradeoff between nutritional quality and
resource availability ( p e r s i s t e n ~ e ) . ~ ~ ~
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 163

Once the insect has made the decision to produce its offspring on a particular host plant,
then a number of reproductive options are still open to it, and these are greatly dependent on
the quality of that host plant.

3. Clutch Size
Clutch size, i.e., the number of eggs or offspring produced in a reproductive event, has
attracted a great deal of attention from evolutionary ecologists. The general conclusion among
vertebrate zoologists is that large clutches are characteristic of short-lived, fast developing
animals exhibiting little parental care and high offspring mortality. Small clutches, on the
other hand, are thought to be characteristic of long-lived, slow maturing animals with a high
investment in parental care and high offspring survival.202Clutch size in birds, for example,
is often closely related to prey/food availability; in years following good food conditions,
clutch size tends to be higher.173If clutch size is high and the food conditions experienced by
the adults are poor, then there will be an increased mortality among the hatchlings since the
more eggs that are laid by the bird, the smaller each hatchling will be and its chances of
surviving are decreased.174The evolution of insect clutch size has undoubtedly been shaped
by similar factors, and this assumption is clearly apparent in the literature.
Clutch size in insects represents a trade off between offspring fitness and adult dispersal.
A detailed review of the costs and benefits of varying clutch size in insects is presented by
Godfray.s0 It is worth considering, however, some of the basic assumptions behind current
models.
Clutch size is related to a number of factors: the frequency of oviposition or larviposition,
the size of the mother, the number of eggsloffspring produced, and the quality and amount of
the host. Several authors have addressed the question of clutch size using models,147.169.170and
their conclusions are in broad agreement with those reached by the vertebrate zoologists. A
major difference is that some authors believe that one of the most important factors determin-
ing clutch size in insects is not offspring success, but the need to achieve maximum egg or
larval deposition.170 This does not contradict the theory that host quality is a major factor in
determining clutch size as it has been shown that host quality affects all the factors listed
above.Iz7

a. Adult Life Span

The rate of offspring production is not a constant over the life of the insect, and the patterns
of clutch size variation are to a certain extent determined by this fact. In nature, female insects
do not generally achieve their maximum possible life span,I2l and the offspring produced at
the beginning of the reproductive period are therefore of greater value than those produced
later, (and thus, in some species, those represented by development of eggs in the upper
reproductive tract). If this is true, then most female insects would be expected to have a large
initial burst of reproduction, and then to have a reproductive rate that would fall away
gradually with increasing age. In fact, female insects usually show a pattern of reproduction
which starts at a low level as the female first begins to reproduce, builds to a peak (the time
taken depending on the probable life span of the insect), and then drops off to a low level again.
Despite this pattern, however, the result is that approximately half the reproductive capacity
of a female insect is achieved in the first third of its reproductive life. This pattern is seen in
long-lived Lepidoptera such as D. p l e ~ i p p u sin , ~relatively
~~ short-lived Lepidoptera such as
P. ~7arnrnea,I~~ and also in short-lived aphids and bugs such as A. fabae and Oncopeltus
f a s c i a t u ~ . ~ ~ Those
. ' ~ ~ . ~insects
~ ' with a high probability of early death lay most of their eggs
either in one clutch or on the first night of oviposition. An example of this strategy is seen in
Chilo partellus, which lays most of its eggs on its first night of oviposition, with clutch size
becoming smaller each succeeding night.I5Some insects with relatively immobile females are
constrained to lay their eggs in one large reproductive burst. For example, the gypsy moth
164 Insect Reproduction

L. d i ~ p a rand
l ~ ~the bird cherry ermine moth Y. e v o n y r n e l l ~ sboth
~ ~ ~lay their eggs clutches in
one large reproductive event. Such insects adjust their reproductive effort in other ways and
must also show specific adaptations to avoid predation on the large egg masses.lgOFemale
insects that lay their eggs in such large clusters tend to have a high realized fecundity.33This
may compensate for the risks entailed in laying all their eggs on the same host rather than
spreading the risk by moving from one host to another, and the increased probability of death
for the mother before oviposition is completed.
Cluster-laying (i.e., the deposition of offspring in aggregations) in butterflies is associated
with aposematic coloration of eggs and larvae. A large number of cluster-laying butterflies
have been shown to be relatively unpalatable as both larvae and adults (e.g., Pieris brassicae,
Euphydryas, Battus spp, and Eumeas ~ p p . )However,
.~~ this is not the whole story as some
aposematic butterflies lay eggs singly and this has been suggested to be a response to
competition for limited larval food resources.14

b. Host Quality
Does host quality affect clutch size? If offspring produced earlier in the reproductive life
of an insect are of greater value, then on a good quality host it would be expected that peak
offspring production would be reached sooner rather than later, as it is to the advantage of the
female to deposit its offspring on a good quality host before she dies. On a poor quality host,
the peak should be achieved later, as the insect has to trade off the risks of waiting for an
improvement in host quality vs. an investment in dispersal to seek out a better quality host.
Some insects are not as flexible in their strategies as others. For example, in the flies Dacus
jarvisi and Dacus tryoni, clutch size is related to food availability. Dacus jarvisi normally
feeds on large fruit and lays large egg clutches, whereas D. tryoni normally feeds on small fruit
and consequently lays only a few eggs on each. When presented with the fruit of the opposite
size class, they are unable to change their ~ t r a t e g i e sThe
. ~ ~ pine beauty moth P. flammea, on
the other hand, when laying eggs on a good host plant, has a short prereproductive delay, lays
a greater proportion of its eggs in the first three days of reproduction, and lives longer than
when ovipositing on a poor quality host.135The pipevine butterfly Battus philenor is also able
to adjust its clutch size in response to differences in host quality, with more eggs per clutch
being deposited when host quality is g ~ o d . " ~ . " ~
Thus, as predicted, herbivorous insects tend to lay large clutches early in their reproductive
life when host quality is high, which will maximize both female reproductive success and
offspring fitness. On poor quality hosts, clutches are small and are produced at a slower rate
than on good quality hosts. In general, clutch size is adjusted so as to maximize reproductive
effort. This can be done by adjusting the size of the offspring produced.lZ7

4. Offspring Size
One of the ways in which an insect can manipulate its reproductive investment is by having
different sizes of offspring. In some insects, for example, large females produce large eggs,
e.g., Parapediasia t e t e r e l l ~ large
, ~ ~ ~females being produced as a result of good larval
nutrition. Egg size variability within a species is generally a response to host or habitat quality.
Theoretically, the more investment there is in reproductive effort per egg or larva (e.g., by
having parental care or large offspring), the greater the likelihood of survival of those
offspring.28Models show that the more eggs produced per clutch, the smaller those eggs will
be.6 Theory states that the larger the offspring are at birth, the greater are the advantages to
be gained. Large egg size in Lepidoptera and other insects confers an advantage on the
offspring, in that they are able to combat poorer conditions.224Thus, on a poor quality host,
or in situations where the eggs are laid at a distance from the host, insects hatching from large
eggs or being deposited as large live young will have greater reserves and be able to establish
more successfully than smaller insects.lOlIn two grasshopper species, Chorthippus brunneus
and Chorthippus parallelus, insects arising from big eggs do indeed grow at a faster rate than
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 165

those arising from small eggsiwOn a good quality host, large size is not as critical as rapid
growth and development is afforded by the host plant. Thus, theoretically, insects depositing
offspring on a poor host should deposit fewer, but larger offspring than those depositing
offspring on a good quality host. The pine beauty moth P. jlammea lays fewer, larger eggs on
Alaskan lodgepole pine (a poor host), than it does on south coastal lodgepole pine (a good
host).I2' The feeding habitats of the insect can also affect the size of eggs laid. For example,
in the Japanese Skipper butterflies (Lepidoptera:Hesperiidae) where some species are polypha-
gous and others less so, the more polyphagous species (which are likely to encounter a wider
variety of hosts of differing quality), lay larger eggs and are less fecund than the species with
the more restricted diets.162Although there are many exceptions, there is a general tendency
for larger insect species to produce relatively large and for generalist feeders to produce
larger eggs than specialist^.^'^ The phenomenon of producing many offspring but of a small
size on a good host is also seen within as well as between species. Aphids, in general, produce
fewer but larger offspring when host quality is
Egg size in insects generally shows a decrease in size as the mother ages, e.g., in Lepi-
doptera such as H. zea and Spodoptera ornithogalli2 and in the pentatomid bug, Graphosoma
l i n e a t ~ mIn ~ ~ bug Dysdercus fasciatus, both clutch size and egg size decrease as the
. ~the
mother ages.lo2However, in the moth P. teterella, although clutch size declines over the life
span of the mother, egg size remains constant.i48In this species, fertility is inversely related
to egg size in that larger eggs are less viable, however, despite this, large females are fitter than
small females due to the fact that although they produce larger eggs overall, they produce more
viable eggs than small females.i48
The pine beauty moth P. jlammea provides a very good illustration of the effects of both
maternal age and host quality on egg size. The size of the eggs laid decreases as the mother
ages.I2' When confined to hosts of differing quality, the female moths respond by delaying the
onset of oviposition on some hosts, and when they do begin to oviposit, their eggs are larger
on the poor quality hosts than on the better quality hosts. In addition, the greater the clutch
size, the smaller the eggs produced.lZ7This phenomenon is also found in other insect groups.
Egg size of the beech leaf-mining weevil, Rhynchaenus fagi is smaller when adult nutrition
is poor, but is much greater when food quality improves? Modeling work on two grasshopper
species, Myrmeleotettix maculatus and C. brunneus, also shows that egg size should decline
with maternal age as reproductive investment becomes increasingly risky.12
Egg size is frequently affected by the size of the mother as well as by her age. For example,
in the butterfly Euploea core corrinne, egg weight decreases with maternal age and is also
correlated with maternal weight.92The maternal weight is in turn an indicator of host quality
of the l a r ~ a e .However,
~ ~ , ~ ~ another study on the same species showed no correlation between
larval diet and the weight of eggs produced by the resultant females.92Egg weight in the
butterfly P. aegeria is also positively correlated with female size and decreases as the mother
agesJmFurther, in the butterflies D. plexippus, Lusiommata petropolitana, Lusiomrnata
maera, Lasiommata megera, Lopinga achine, and P. rapae crucivora, egg weights decrease
as the mother ages.1m106J95 This is seen in a large number of insects in other groups. For
example, in the beetle C. maculatus, egg size decreases with maternal age and large eggs are
more viable than small ones.214
If host quality is ignored by the observer, then apparently anomalous results can be found.
For example, the size of eggs of the butterfly P. rapae were found in one study to be positively
correlated with maternal age and inversely correlated with adult size, which appears to run
contrary to the general rule.* Large individuals were more fecund but laid smaller eggs.
However, the small butterflies that then arose were generally present as adults in poor quality
habitats and thus produced large eggs that produced larvae able to withstand the poor
conditions.* Another apparent contradiction is shown by L. dispar. Small eggs were produced
in forests where there were large numbers of defoliated trees and hosts were of poor quality.Iw
This was explained by the fact that small eggs gave rise to less mobile larvae which tended
166 Insect Reproduction

to remain on the hosts on which they were deposited. Since suitably foliated host trees are in
short supply under these conditions, it is to the advantage of the larvae to remain where they
were laid, rather than use energy dispersing to another tree which is just as likely to be of poor
host quality as the one they had left. Large eggs are produced when conditions are good and
hosts are in abundance, as is the case when a small insect population is in the process of
building up. Under these circumstances, it may be to the advantage of the larvae to disperse
to new hosts. Large larvae are more mobile and thus disperse successfully.10g
In the Lepidoptera, the relationship between offspring size and fitness is not immediately
clear. In the butterfly P. aegeria, no fitness function was found in relation to egg size, that is,
large larvae did not survive better than small larvae. However, egg size did decrease with
maternal age,224possibly allowing more eggs to be laid. The checkerspot butterfly E. editha,
when fed as adults, were found in one study to keep their egg weights constant over their entire
life span, in contrast to unfed females which showed the expected decline in egg weight.'57
However, in other studies Lepidoptera were also shown to have a decline in egg weight with
maternal age even when fed.127Clearly, egg weight can alter in response to selection pressure
over evolutionary time, but the variability with relation to the host plant within a species or
individual is also surprisingly large and apparently adaptive. In this context, it is relevant to
note that egg size in Lepidoptera may be related to the number of larval instars and the size
of the adult. Dyaf13showed that in many lepidopteran species there is an approximate doubling
of volume between each instar. Thus, by having an egg double the size of a related species,
the number of larval instars can be reduced by one and an adult of the same size still produced.

IV. CONCLUSIONS
Despite the large amounts of literature pertaining to the subject, it is apparent that many
of the factors affecting fecundity, fertility, and offspring deposition in insect are still relatively
little understood. Although the mechanics of oviposition and many of the biological con-
straints, e.g., sex ratio, larval nutrition, etc., affecting fecundity and fertility are well known,
the effect that the various interactions between host quality and abiotic factors have on them
requires further research.

ACKNOWLEDGMENT
It is a pleasure to thank Nigel Straw for his perceptive and helpful comments on earlier
drafts of this chapter.

REFERENCES
1 . Aalbersberg, Y.K., Du Toit, F., van der Westinizen, M.C., and Hewitt, P.H., Development rate, fecundity
and lifespan of apterae of the Russian wheat aphid, Diuraphis noxia (Mordvilko) (Hemiptera: Aphididae),
under controlled conditions. Bull. Entomol. Res., 77, 629, 1987.
2. Adler, P.H., Willey, M.B., and Bowen, M.R., Temporal ovipositionpatterns of Heliarhis tea and Spodoptera
ornithogalli. Entomol. Exp. Appl., 58, 159, 1991.
3. AI Salti, M.N., Influence de la temperature pendant la vie imaginale sur les potentialites reproductrices de
I'espece Sesamia nonagriodes Lefebvre (Lep., Noctuidae). Acta Oecol. Oecol. Appl., 5, 103, 1984.
4. Anderson, S.S., McCrea, K.D., Abrahamson, W.G., and Hartzel, L.M., Host genotype choice by the ball
gallmaker Eurosta solidagensis (Diptera: Tephritidae). Ecology, 70, 1048, 1989.
5 . Angelo, MJ. and Slansky, F., Body building insects: trade-offs in resource allocation with particular
reference to migration. Fla. Enromol.. 67, 22, 1984.
6. Atkinson, D.A. and Begon, M., Reproductive variation and adult size in two CO-occumnggrasshopper
species. Ecol. Enromol., 12, 119, 1987.
F~tctorsAffecting Fecundity, Fertility, Oviposition, and Luwiposition in Insects 167

7. Auerbach, M. and Simberloff, D., Oviposition site preference and larval mortality in a leaf-mining moth.
Ecol. Entomol., 14, 131, 1989.
8. Auger, M.A., Geri, C., Jay-Allemand, C., and Bastien, C., Comestibiliti: de differents clones de pin
sylvestre pour Diprion pini L. (Hym., Diprionidae). I. Incidence de la consommation des aiguilles de
differknts clones de pin sylvestre sur le developpement de Diprion pini L. J . Appl. Entomol., 1 10,489, 1990.
9. Bale, J.S., Bud burst and success of the beech weevil, Rhynchaenus fagi: feeding and oviposition. Ecol.
Entomol., 9, 139, 1984.
10. Battisti, A., Host-plant relationships and population dynamics of the pine processionary caterpillar
Thaumeropoea pityocampa (Dennis and Schiffermuller). 1.Appl. Entomol., 105, 393, 1988.
11. Beevor, P.S., Mumford, J.D., Shah, S., Day, R.K., and Hall, D.R., Observations on pheromone baited mass
trapping for control of cocoa pod borer, Conopomorpha cramerella, in Sabah, East Malaysia. Crop Prof., 12,
134, 1993.
12. Begon, M. and Parker, G.A., Should egg size and clutch size decrease with age? Oikos, 47,293, 1988.
12a. Bellinger, R.G. and Pienkowski, R.L., lnterspecific variation in ovariole number in Melanopline grasshop-
pers (Orthoptera: Acrididae). Ann. Entomol. Soc. Am., 78, 127, 1985.
13. Bennetova, B. and Fraenkel, G., What determines the number of ovarioles in a fly ovary? J . Insect. Physiol.,
27,403, 1981.
14. Benson, W.W., Resource partitioning in passion vine butterflies. Evolution, 32, 493, 1978.
15. Berger, A., Egg weight, batch size and fecundity of the spotted stalkborer, Chilopartellus in relation to weight
of females and time of oviposition. Entomol. Exp. Appl., 50, 199, 1989.
16. Bernays, E.A. and Chapman, R.F., Evolution of plant deterrence to insects. In Perspectives in Chemore-
ception and Behavior, (Eds., R.F. Chapman, E.A. Bernays, and J.G. Stoffolano), Springer-Verlag. New York,
1987, 159.
17. Bernays, E.A. and Cornelius, M., Relationship between deterrence and toxicity of plant secondary com-
pounds for the alfalfa weevil Hypera brunneipennis. Entomol. Exp. Appl., 64, 289, 1993.
18. Bevan, D. and Brown, R.M., Pine Looper Moth, Forestry Commission Forest Record 119, HMSO, London,
1978.
19. Bintcliffe, EJ.B. and Wratten, S.D., Antibiotic resistance in potato cultivars to the aphid Myzus persicae.
Ann. Appl. Biol., 100, 382, 1982.
20. Blackman, R.L., Specificity in aphidlplant genetic interactions with particular reference to the role of the
alate colonizer. In Aphid-Plant Genotype Interactions, (Eds., R . K . Campbell and R.D. Eikenbary), Elsevier,
Amsterdam, 1990, 25 1.
21. Blais, J.R., Effects of the destruction of the current year's foliage of Balsam fir on the fecundity and habits
of flight of the spruce budworm. Can. Entomol., 85,446, 1953.
22. Boggs, C.L., Selection pressures affecting male nutrient investment at mating in heliconiine butterflies.
Evolution, 35, 931, 1981.
23. Boggs, C.L., Ecology of nectar and pollen feeding in Lepidoptera. In Nutritional Ecology of Insects, Mites.
Spiders, and Related Invertebrates (Eds., F. Slansky and J.G. Rodriguez), John Wiley & Sons, New York,
1987, 369.
24. Boggs, C.L. and Watt, W.B., Population structure of Pierid butterflies. IV. Genetic and physiological
investment in offspring by male Colias. Oecologia, 50, 320, 1981.
25. Bristow C. M., Differential benefits from ant attendance to two species of Homoptera on New York iron
weed. J. Aninz. Ecol., 53, 715, 1984.
26. Brower, J.H., PIodia interpunctella: effect of sex ratio on reproductivity. Ann. Entomol. Soc. Am., 68, 847,
1975.
27. Burpee, D.M. and Sakaluk, S.K., Repeated matings offset costs of reproduction in female crickets. Evol.
Ecol., 7, 240, 1993.
28. Calow, P., Economics of ontogeny - adaptational aspects. In Evolutionary Ecology. (ed. B. Shorrocks),
Blackwell Scientific Publications, Oxford, 1984, 8 1.
29. Carroll, A.L. and Quiring, D.T., Sucrose ingestion by Zeiraphera canadensis Mut. and Free. (Lepidoptera,
Tortricidae) increases fecundity and lifetime fecundity, but not oviposition rate. Can. Entomol., 124, 335,
1992.
30. Cheng, H.H., Oviposition and longevity of the dark-sided cutworm, Euxoa niessoria (Lepidoptera: Noctuidae),
in the laboratory. Can. Entomol., 104, 919, 1972.
3 1. Chew, F.S., Coevolution of pierid butterflies and their cruciferous food plants. 11. The distribution of eggs on
potential foodplants. Evolution, 31, 568, 1977.
32. Chew, F.S and Robbins, R.K., Egg laying in butterflies. In The Biology of Butteg7ies, (Eds., R.1. Vane-
Wright and P.R. Ackery), Symposium of the Royal Entomological Society of London, Number 1I, Academic
Press, London, 1984, 65.
33. Courtney, S.P., The evolution of egg clustering by butterflies and other insects. Am. Nat., 123,276, 1984.
34. Courtney, S.P., Why insects move between host patches - some comments on risk-spreading. Oikos, 47,
112, 1986.
168 Insect Reproduction

35. Courtney, S.P. and Kibota, T.K., Mother doesn't know best: selection of hosts by ovipositing insects. In
Insect-Plant Interactions, Volume I1 (Ed., E.A. Bernays), CRC Press, Boca Raton, 1990, 161.
36. Craig, T.P., Itami, J.K., and Price, P.W., A strong relationship between oviposition preference and larval
performance in a shoot-galling sawfly. Ecology, 70, 1691, 1989.
37. Craig, T.P., Price, P.W., and Itami, J.K., Resource regulation by a stem-galling sawfly on the arroyo
willow. Ecology, 67, 419, 1986.
38. Critchley, B.R., Chamberlain, DJ., Campion, D.G., Attique, M.R., Ali, M., and Ghaffar, A., Integrated
use of pink bollworm pheromone formulations and selected conventional insecticides for the control of the
cotton pest complex in Pakistan. Bull. Entomol. Res., 81, 371, 1991.
39. Danthanarayana, W., Factors determining variation in fecundity of the light brown apple moth, Epiphyas
postvittana (Walker) (Tortricidae). Aust. J. Zool., 23,439, 1975.
40. Danthanarayana, W., Environmentally cued size variation in light-brown apple moth, Epiphyaspostvi~ana
(Walk.) (Tortricidae), and its adaptive value in dispersal. Oecologia, 26, 121, 1976.
41. Danthanarayana, W. and Gu, H., Multiple mating and its effect on the reproductive success of female
Epiphyas postvirtana (Lepidoptera: Tortricidae). Ecol. Entomol., 16, 169, 1991.
42. Denno, R.F. and Grissell, E.E., The adaptiveness of wing-dimorphism in the salt marsh inhabiting planthopper
Prokelisia marginata (Homoptera: Delphacidae). Ecology, 60, 221, 1979.
43. Denno, R.F. and McCloud, E.S., Predicting fecundity from body size in the planthopper, Prokelisia
marginata (Homoptera: Delphacidae). Environ. Entomol., 14, 846, 1985.
44. Denno, R.F., Larsson, S., and Olmstead, K.L., Role of enemy-free space and plant quality in host-plant
selection by willow beetles. Ecology, 7, 124, 1990.
45. Derr, J.A., Alden, B., and Dingle, H., Insect life histories in relation to migration, body size and host plant
assay: a comparative study of Dysdercus. J. Anim. Ecol., 50, 181 1981.
46. Deseo, K.V., Study of factors influencing the fecundity and fertility of codling moth, (Laspeyresia pomonella
L. Lepid.; Tortr.). Acta Phytopathol. Acad. Sci. Hung., 6, 247, 1971.
47. Deseo, K.V. and Saringer, G., Photoperiodic effect on fecundity of Laspeyresia pomonella, Grapholitha
funebrana and G. molesra: the sensitivity period. Entomol. Exp. Appl., 18, 187, 1975.
48. Dewar, A.M., Assessment of methods for testing varietal resistance to aphids in cereals. Ann. Appl. Biol., 87,
183, 1977.
49. Dixon, A.F.G., Reproductiveactivity of the sycamore aphid, Drepanosiphum platanoides (Schr.) (Hemiptera,
Aphididae). J. Anim. Ecol., 32, 33, 1963.
50. Dixon, A.F.G., The effect of population density and nutritive status of the host on the summer reproductive
activity of the sycamore aphid, Drepanosiphum platanoides (Schr.). J. Anim. Ecol., 35, 105, 1966.
51. Dixon, A.F.G., Population dynamics of the sycamore aphid Drepanosiphurnplatanoides (Schr.) (Hemiptera:
Aphididae): migratory and trivial flight activity. J. Anim. Ecol., 38, 585, 1969.
52. Dixon, A.F.G., Quality and availability of food for a sycamore aphid population. In Animal Popularions in
Relation to their Food Resources, (Ed.,A. Watson), Blackwell Scientific Publications, Oxford, 1970, 277.
53. Dixon, A.F.G., The life cycle and host preferences of the bird cherry-oat aphid, Rhopalosiphum padi (L.) and
their bearing on the theories of host alternation in aphids. Ann. Appl. Biol., 68, 135, 1971.
54. Dixon, A.F.G.. Reproductive strategies of the alate morphs of the bird cherry-oat aphid Rhopalosiphum padi
(L.). J . Anim. Ecol., 45. 1976.
55. Dixon, A.F.G., Aphid Ecology, Blackie, London, 1985, Chapter 6.
56. Dixon, A.F.G. and Dharma, T.R., Number of ovarioles and fecundity in the black bean aphid, Aphis fabae.
Entomol. Exp. Appl., 28, 1, 1980.
57. Dixon, A.F.G. and Glen, D.M., Morph determination in the bird cherry-oat aphid, Rhopalosiphum padi (L.).
Ann. Appl. Biol., 68, 11, 1971.
58. Dixon, A.F.G. and Wellings, P.W., Seasonality and reproduction in aphids. Int. J. Inv. Rep., 5, 83, 1982.
59. Dixon, A.F.G. and Wratten, S.D., Laboratory studies on aggregation size and fecundity in the black bean
aphid, Aphis fabae (Scop.). Bull. Entomol. Res., 61, 97, 1971.
60. Dunlap-Pianka, H.L., Ovarian dynamics in Heliconius butterflies: correlations among daily oviposition
rates, egg weights, and qualitative aspects of oogenesis. J. Insect Physiol., 25, 741, 1979.
61. Dunlap-Pianka, H.L., Boggs, C.L., and Gilbert, L.E., Ovarian dynamics in Heliconiine butterflies: pro-
grammed senescence versus eternal youth. Science, 197, 487, 1977.
62. Du Merle, P. and Brunet, S., From green to yellow or yellowish white: egg-colour changes in relation to
oviposition rank in the fir budworm Choristoneura murinana (Hb.) (Lep.. Tortricidae). J . Appl. Entomol.. 11 I,
342. 1991.
63. Dyar, H.G., The number of moults of lepidopterous larvae. Psyche, 5,420, 1890.
64. Ehrlich, A.H. and Ehrlich, P.R., Reproductive strategies in the butterflies: mating frequency, plugging and
egg number. J. Kans. Entomol. Soc., 51, 666, 1978.
65. Ellington, JJ. and El-Sokkari, A., A measure of the fecundity, ovipositional behavior and mortality of the
bollworm, Heliothis tea (Boddie) in the laboratory (Lepidoptera, Noctuidae). S.W. Entomol., 11, 177, 1986.
Factors Affecting Fecundity, Fertility, Oviposition, and Lurviposition in Insects 169

66. Elliott, W.M., A method of predicting short term population trends of the green peach aphid, Myzus persicae
(Homoptera: Aphididae), on potatoes. Can. Entomol., 105, 11, 1973.
67. El-Sherif, S., Gomaa, A.A., and Hemeida, I.A., Effect of adult diet and mating on egg laying capacity and
longevity of potato tuber moth, Phthorimaea operculella Zeller. Z. Angew. Entomol., 87, 170, 1979.
68. Evans, H.F., The population dynamics of Anthocoris confusus in a laboratory cage system. J. Anim. Ecol.,
45, 773, 1976.
69. Fatzinger, C.W. and Merkel, E.P., Oviposition and feeding preferences of the southern pine coneworm
(Lepidoptera, Pyralidae) for different host-plant materials and observationson monotexpenes as an oviposition
stimulant. J. Chem. Ecol., l l, 689, 1985.
70. Feeny, P.P., Seasonal changes in oak leaf tannins and nutrient as a cause of spring feeding by winter moth
caterpillars. Ecology, 5 1, 565, 1970.
7 1. Firempong, S. and Zalucki, M.P., Host plant selection by Helicoverpa armigera (Hubner) Lepidoptera:
Noctuidae); the role of certain plant attributes. Aust. J . Zool., 37, 675, 1990.
72. Fitt, G.P., Variation in ovariole number and egg size of species of Dacus (Diptera: Tephritidae and their
relation to host specialization. Ecol. Entomol.. 15, 255, 1990.
73. Fitt, G.P., Comparative fecundity, clutch size, ovariole number and egg size of Dacus tryoni and D. jarvisi,
and their relationship to body size. Entomol. Exp. Appl., 55, 11, 1990.
74. Ford, E.B., Butterflies. Collins, London, 1945.
75. Forno, I.W. and Bourne, AS., Oviposition by the weevil Cyrtobagous salviniae Calder and Sands when its
host plant, Salvinia molesta is damaged. J. Appl. Entomol., 106, 85, 1988.
76. Frazer, B.D., Life tables and intrinsic rates of increase of apterous black bean aphids and pea aphids, on broad
bean (Homoptera: Aphididae). Can. Entomol., 104, 1717, 1972.
77. Fritz, R.S. and Nobel, J., Host plant variation in mortality of the leaf-folding sawfly on the arroyo willow.
Ecol. Entomol.. 15, 25, 1990.
78. Furuta, K., Host preferences and population dynamics in an autumnal population of the maple aphid,
Periphyllus californiensis Shinji (Homoptera: Aphididae). Z Angew. Entomol., 102, 93, 1986.
79. Gilbert, L.E., The biology of butterfly communities. In The Biology of BurterJies, (Eds. R.I. Vane-Wright
and P.R. Ackery), Symposium of the Royal Entomological Society Number 11, Academic Press, 1984, 41.
80. Godfray, H.CJ., The evolution of clutch size in invertebrates. Oxford Surveys in Evolutionary Biology, 4,
117, 1987.
80a. Gohari, H. and Hawlitzky, N., Acivitt reproductrice de lay pyrale du mais, Ostrinia nubilnlis, Hbn. (Lep.,
Pyralidae) B basses temperatures constantes. Agronomie, 6, 91 1, 1986.
8 1. Gordon, D.M. and Stewart, R.K., Demographic characteristicsof the stored products moth Cadra cautella.
J. Aninl. Ecol., 57, 627, 1988.
82. Gruber, K. and Dixon, A.F.G., The effect of nutrient stress on development and reproduction in an aphid.
Entomol. Exp. Appl., 47, 23, 1988.
83. Gu, H. and Danthanarayana, W., The role of availability of food and water to the adult Epiphyas
postvittana, the light brown apple moth, in its reproductive performance. Entomol. Exp. Appl., 54, 101, 1990.
84. Guerra, A.A., Wolfenbarger, A., and Garcia, R.D., Factors affecting reproduction of the tobacco budworm
in the laboratory. J. Econ. Entomol., 65, 1341, 1974.
85. Gunn, A. and Gatehouse, A.G., Effects of the availability of food and water on reproduction in the African
armyworm, Spodoptera exempta. Phys. Entomol., 10, 53, 1985.
86. Gunn, A. and Gatehouse, A.G., The effect of adult feeding on lipid and protein reserves in African
armyworm, Spodoptera exempta, moths before and during reproduction. Phys. Entomol.. l l, 423, 1986.
86a. Hamilton, J.G. and Zalucki, M.P., Interactions between a specialist herbivore, Crocidosem plebejana, and
its host plants Malva parviflora and cotton, Gossypium hirsutum - oviposition preference. Entomol. Exp.
Appl., 66, 207, 1993.
87. Harcourt, D.G. and Cass, L.M., Photoperiodism and fecundity in Plutella maculipennis (Curt.). Nature, 2 10,
217, 1966.
88. Harris, P., Natural mortality of the pine shoot moth Rhyacionia buoliana (Schiff.) in England. Can. J. Zool.,
38, 755, 1960.
89. Heliovaara, K. and Vaisanen, R., Changes in population dynamics of pine insects induced by air pollution.
In Population Dynamics of Forest Insects, (Eds., A.D. Watt, S.R. Leather, M.D. Hunter, and N.A.C. Kidd),
Intercept Press, Andover, 1990, 209.
90. Hennebeny, TJ. and Clayton, T.E., Tobacco budworm moths (Lepidoptera: Noctuidae): effect of time of
emergence, male age, and frequency of mating on sperm transfer and egg viability. J. Econ. Entomol., 78,379,
1985.
91. Heron, R.J., The reproductive capacity of the larch sawfly and some factors of concern in its measurement.
Can. Entomol.. 98, 56 1, 1966.
92. Hill, CJ., The effect of adult diet on the biology of butterflies. 2. The common crow butterfly, Euploea core
corinna. Oecologia, 8 1, 258, 1989.
170 Insect Reproduction

93. Hill, CJ. and Pierce, N.E., The effect of adult diet on the biology of butterflies. 1. The common imperial
blue, Jaln~enusevagoras. Oecologia, 81, 249, 1989.
94. Honek, A., Host plant energy allocation to and within ears, and abundance of cereal aphids. J. Appl. Entomol.,
110, 68, 1990.
95. Horton, D.R., Performance of a willow-feeding beetle, Chrysomela knabi Brown, as affected by host species
and dietary moisture. Can. Entomol., 121, 777, 1989.
96. Hough, J.A. and Pimentel, D., Influence of host foliage on development, survival and fecundity of the gypsy
moth. Environ. Entomol., 7, 97, 1978.
97. Imms, A.D., A General Textbook of Entomology, Ninth ed., Chapman & Hall, London, 1957.
98. James, W.O., An Introduction to Plant Physiology, Oxford University Press, Oxford, 1973, Chapter 7.
99. Jones, R.E., Hart, J.R., and Bull, G.D., Temperature, size and egg production in the cabbage butterfly, Pieris
rapae L. Aust. J. Zool., 30, 223, 1982.
100. Karlsson, B., Variation in egg weight, oviposition rate and reproductive reserves with female age in a natural
population of the speckled wood butterfly, Pararge aegeria. Ecol. Entomol., 12,473, 1987.
101. Karlsson, B. and Wiklund, C., Egg weight variation in relation to egg mortality and starvation endurance
of newly hatched larvae in some satyrid buttertlies. Ecol. Entomol., 10, 205, 1985.
102. Kasule, F.K., Egg size increases with maternal age in the cotton stainer bugs Dysdercus fasciatus and D.
cardinalis (Hemiptera: Pyrrhocoridae). Ecol. Enromol., 16, 345, 1991.
103. Keese, M.C. and Wood, T.K., Host-plant mediated geographic variation in the life history of Plafycotis
vittata (Homopten: Membracidae). Ecol. Entomol., 16, 63, 1991.
104. Kenten, J., The effect of photoperiod and temperature on reproduction in Acyrthosiphon pisum (Hams) and
on the forms produced. Bull. Entomol. Res., 46, 599, 1955.
105. Kidd, N.A.C. and Tozer, D.J., On the significance of post-reproductive life in aphids. Ecol. Entomol., 10,
357, 1985.
106. Kimura, K. and Tsubaki, Y., Egg weight variation associated with female age in Pieris rapae crucivoro
Boisduval (Lepidoptera; Pieridae). Appl. Entomol. Zool., 20, 500, 1985.
107. Koyama, J., Teruya, T., and Tanaka, K., Eradication of the Oriental fruit fly (Diptera: Tephritidae) from
the Okinawa Islands by male annihilation technique. J. Econ. Entomol.. 77,468, 1984.
108. Labine, P.A., The population biology of the butterfly, Euphydryas editha. IV. Sperm precedence - a
preliminary report. Evolution, 20, 580, 1966.
109. Lance, D.R., Host-seeking behavior of the gypsy moth: the influence of polyphagy and highly apparent host
plants In Herbivorous insects -Host Seeking Behaviour and Mechanisms (Ed., S. Ahmad), Academic Press,
New York, 1983, 201.
110. Larsson, F.K., Female longevity and body size as predictors of fecundity and egg length in Graphosoma
lineatum L. Dtsch. Entomol. Z., 36, 329, 1989.
111. Leather, S.R., Aspects of the Ecology of the Bird Cherry-Oat Aphid. Rhopalosiphum padi (L.). Unpublished
Ph.D. Thesis, University of East Anglia, Nonvich, U.K., 1980.
112. Leather, S.R. Reproduction and survival: a field study of gynoparaeof the bird cherry-oat aphid, Rhopalosiphum
padi (Homopten; Aphididae) on its primary host Prunuspadus. Ann. Entomol. Fenn., 47, 131, 1981.
113. Leather, S.R., Do gynoparae and males need to feed? An attempt to allocate resources in the bird cherry oat
aphid, Rhopalosiphum padi. Entomol. Exp. Appl., 31, 386, 1982.
114. Leather, S.R., Evidence of ovulation after adult moult in the bird cherry-oat aphid, Rhopalosiphum padi.
Entomol. Exp. Appl.. 33, 348, 1983.
115. Leather, S.R., The effect of adult feeding on the fecundity, weight loss and survival of the pine beauty moth,
Panolisflammea (D & S). Oecologia, 65, 70, 1984.
116. Leather, S.R., Factors affecting pupal survival and eclosion in the pine beauty moth, Panolis flammea
(D & S). Oecologia, 63, 75, 1984.
117. Leather, S.R., Oviposition preferences in relation to larval growth rates and survival in the pine beauty moth,
Panolis flammea. Ecol. Entomol., 10, 2 13, 1985.
117a. Leather, S.R., Atmospheric humidity and aphid reproduction. Z. Angew. Entomol., 100, 510, 1985.
118. Leather, S.R., Host monitoring by aphid migrants: do gynoparae maximise offspring fitness? Oecologia, 68,
367, 1986.
119. Leather, S.R., Insects on bird cherry. I. The bird cheny ermine moth, Yponomeuta evonymellus (L.)
(Lepidoptera: Yponomeutidae). Entomol. Gaz., 37, 209, 1986.
120. Leather, S.R., Pine monoterpenes stimulate oviposition in the pine beauty moth, Panolisflammea. Entomol.
Exp. Appl., 43, 295, 1987.
121. Leather, S.R., Size, reproductive potential and fecundity in insects. Things aren't as simple as they seem.
Oikos, 51, 386, 1988.
122. Leather, S.R., Do alate aphids produce fitter offspring? The influence of maternal rearing history and morph
on life-history parameters of Rhopalosiphum padi (L.). Funcr. Ecol., 3, 237, 1989.
123. Leather, S.R., Sex-ratio and reproductive success in the pine beauty moth, Panolis flammea (Den. and
Schiff.) (Lep., Noctuidae). J. Appl. Entomol., 109, 200, 1990.
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 171

124. Leather, S.R., The role of host quality, natural enemies, competition and weather in the regulation of autumn
and winter populations of the bird cherry aphid. In Population Dynamics of Forest Insects, (Eds., A.D.Watt,
S.R. Leather, M.D. Hunter, and N.A.C. Kidd), Intercept Press, Andover, 1990, 35.
125. Leather, S.R., Life history traits of insect herbivores in relation to host quality. In Insect-Plant Interactions,
Volume V, (Ed.,E.A. Bemays), CRC Press, Boca Raton, 1994, 176.
126. Leather, S.R. and Barbour, D.A., The effect of temperature on the emergence of pine beauty moth, Panolis
flammea Schiff. (Lep., Noctuidae). Z. Angew. Entomol.. 96, 445448, 1983.
127. Leather, S.R. and Burnand, A.C., Factors affecting life-history parameters of the pine beauty moth, Panolis
flammea (D & S): the hidden costs of reproduction. Funcr. Ecol., 1, 331, 1987.
128. Leather, S.R. and Dixon, A.F.G., The effect of cereal growth stage and feeding site on the reproductive
activity of the bird-cheny aphid, Rhopalosiphum padi. Ann. Appl. Biol., 97, 135, 1981.
129. Leather, S.R. and Dixon, A.F.G., Growth, survival and reproduction of the bird-cheny aphid, Rhopalosiphum
padi, on its primary host. Ann. Appl. Biol., 99, 115, 1981.
130. Leather, S.R. and Dixon, A.F.G., Secondary host preferences and reproductive activity of the bird cherry-
oat aphid, Rhopalosiphum padi. Ann. Appl. Biol., 101, 219, 1982.
131. Leather, S.R. and Mackenzie, G.A., Factors affecting the population development of the bird cheny ermine
moth, Yponomeuta evonymella (L.). The Entomologist, 113, 81, 1994.
132. Leather, S.R. and Wellings, P.W., Ovariole number and fecundity in aphids. Entomol. Exp. Appl., 30, 128,
1981.
133. Leather, S.R., Walters, K.F.A., and Bale, J.S., The Ecology of Insect Overwinrering, Cambridge University
Press, Cambridge, U.K., 1993.
134. Leather, S.R., Ward, S.A., and Dixon, A.F.G., The effect of nutrient stress on life history parameters of the
black bean aphid, Aphis fabae Scop. Oecologia, 57, 156, 1983.
135. Leather, S.R., Watt, A.D.,and Barbour, D.A., The effect of host plant and delayed mating on the fecundity
and lifespan of the pine beauty moth, Panolisflammea (Dennis and Schiffermuller)(Lepidoptera:Noctuidae):
their influence on population dynamics and relevance to pest management. Bull. Entomol. Res., 75,641, 1985.
136. Leather, S.R., Watt, A.D., and Entwistle, P.F., The effect of temperature on oviposition and egg hatch in
the pine beauty moth, Panolisflammea (D & S): simulation and prediction of field and laboratory populations.
For. Ecol. Manage.. in press.
137. Leather, S.R., Watt, A.D., and Forrest, G.I., Insect-induced changes in young lodgepole pine (Pinus
contorta): the effect of previous defoliation on oviposition, growth and survival of the pine beauty moth,
Panolisflammea. Ecol. Entomol.. 12,275, 1987.
138. Leather, S.R., Wellings, P.W., and Dixon, A.F.G., Habitat quality and the reproductive strategies of the
migratory morphs of the bird cherry-oat aphid, Rhopalosiphum padi (L.), colonizing secondary host plants.
Oecologia, 59, 302, 1983.
139. Leather, S.R., Wellings, P.W., and Walters, K.F.A., Variation in ovariole number within the Aphidoidea.
J. Nar. Hist., 22, 381, 1988.
140. Lederhouse, R.C., The effect of female mating frequency on egg fertility in the black swallowtail, Papilio
polyxenes asterius (Papilionidae). J. Lepid. Soc., 35, 266, 1981.
14 1. Lingren, P.D., Warner, W.B., and Henneberry, TJ.,Influence of delayed mating on egg production, egg
viability, mating and longevity of female pink bollworm, (Lepidoptera:Gelechiidae). Environ. Entomob, 17,
86, 1988.
142. Liu, S.S. and Hughes, R.D., The influence of temperature and photoperiod on the development,survival and
reproduction of the sowthistle aphid, Hyperomyzus lactucae. Entomol. Exp. Appl., 43, 31, 1987.
143. Lukefahr, M.J. and Martin, D.F., The effect of various larval and adult diets on the fecundity and longevity
of the bollworm, tobacco budworm and the cotton leafworm. J. Econ. Enromol., 57, 233, 1964.
144. Mackay, D.A. and Jones, R.E., Leaf shape and the host-finding behaviour of two ovipositing monophagous
butterfly species. Ecol. Enromol., 14, 423, 1989.
145. McClure, MS., Reproduction and adaptation of exotic hemlock scales (Homoptera: Diapsidae) on their new
and native hosts. Environ. Entomol., 12, 1811, 1983.
146. McClure, M.S. and Hare, J.D., Foliar telpenoids in Tsuga species and the fecundity of scale insects.
Oecologia, 63, 185, 1984.
147. Mangel, M., Oviposition site selection and clutch size in insects. J. Marh. Biol., 25, 1, 1987.
148. Marshal, L.D., lntraspecific variation in reproductive effort by female Parspediasia teterella (Lepidoptera:
Pyralidae) and its relation to body size. Can. J. Zool., 68, 44, 1990.
149. Messina, FJ., Influence of cowpea pod maturity on the oviposition choices and larval survival of a bruchid
beetle Callosobruchus macularus. Entomol. Exp. Appl.. 35, 241, 1984.
150. Miller, W.E., Reproductive enhancement by adult feeding effects of honeydew in imbibed water on spruce
budworm. J. Lepid. Soc., 43, 167, 1989.
151. Moller, J., Investigations on a laboratory culture of the diamond-back moth, Plutella maculipennis (Curt.)
(Lep., Tineidae). 11. Influence of larval density on life history panmeters of larvae, pupae and adults. J. Appl.
Entomol., 105,425, 1988.
172 Insect Reproduction

152. Minkenberg, O.P.J.M. and Fredrix, M.J.J., Preference and performance of an herbivorous fly, Liriomyzn
trijolii (Diptera: Agromyzidae), on tomato plants differing in leaf nitrogen. Ann. Entomol. Soc. Am., 82,350,
1989.
153. Minkenberg, O.PJ.M., Tatar, M., and Rosenheim, J.A., Egg load as a major source of variability in insect
foraging and oviposition behaviour. Oikos, 65, 134, 1992.
154. Montgomery, M.E. and Wallner, W.E., The gypsy moth - a westward migrant. In Dynamics of Forest
Insect Populations: Patterns, Causes, Implications, (Ed., A.A. Berryman), Plenum Press, New York, 1988,
353.
155. Moore, GJ., Host plant discrimination in tropical satyrine butterflies. Oecologia, 70, 592, 1986.
156. Moore, L.V., Myers, J.H., and Eng, R., Western tent caterpillars prefer the sunny side of the tree, but why?
Oikos, 51, 321, 1988.
157. Moore, R.A. and Singer, M.C., Effects of maternal age and adult diet on egg weight in the butterfly
Euphydryas editha. Ecol. Entomol.. 12, 401, 1987.
158. Moran, N.A., Seasonal shifts in host usage in Uroleucon gravicome (Homoptera: Aphididae) and implica-
tions for the evolution of host alternation in aphids. Ecol. Entomol., 8, 371, 1983.
159. Morrow, P.A. and Fox, L.R., Effects of variation in Eucalyptus essential oil yield on insect growth and
grazing damage. Oecologia, 45, 209, 1980.
160. Murphy, D.D., Launer, A.E., and Ehrlich, P.R., The role of adult feeding in egg production and population
dynamics of the checkerspot butterfly Euphydryas editha. Oecologia, 56, 257, 1983.
161. Myers, J.H., Monro, J., and Murray, N., Egg clumping, host plant selection and population regulation in
Cactoblastis cactorum (Lepidoptera). Oecologia, 5 1, 1981.
162. Nakasuji, F., Egg size of skippers (Lepidoptera: Hesperiidae) in relation to their host specificity and to leaf
toughness of host plants. Ecol. Res., 2, 175, 1987.
163. Nealis, V.G., Diapause and the seasonal ecology of the introduced parasite Cotesia (Apanteles) rubecula
(Hymenoptera: Braconidae). Can. Entomol., 117, 333, 1985.
164. Newton, C. and Dixon, A.F.G., Methods of hatching the eggs and rearing the fundatrices of the English grain
aphid, Sitobion avenue. Entomol. Exp. Appl., 45, 277, 1987.
165. Noguchi, H., Mating frequency, fecundity, and egg hatchability of the smaller tea tortrix moth, Adoxyphyes
sp. (Lepidoptera: Tortricidae). Jpn. J. Appl. Entomol. Zool., 25, 259, 1981.
166. Otake, A. and Sakuratami, Y., Egg production of the adult tobacco cutworm, Spodoptera litura (Lepi-
doptera: Noctuidae) under different sex ratios and population processes of the larvae. Appl. Entomol. Zool.,
7, 190, 1972.
167. Oyeyele, S.O. and Zalucki, M.P., Cardiac glycosides and oviposition by Danaus plexippus on Asclepias
fruiticosa in south-east Queensland (Australia), with particular notes on the effect of plant nitrogen content.
Ecol. Entomol., 15, 177, 1990.
168. Packer, M J . and Corbet, P.S., Size variation and reproductive success of female Aedes punctor (Diptera:
Culicidae). Ecol. Entomol.. 14, 297, 1989.
169. Parker, G.A. and Begon, M., Optimal egg size and clutch size: effects of environment and maternal
phenotype. Am. Nat., 128, 573, 1986.
170. Parker, G.A. and Courtney, S.P., Models of clutch size in insect oviposition. Theor. Popul. Biol., 26, 27,
1984.
171. Peckarovsky, B.L. and Cowan, C.A., Consequences of larval intraspecific competition to stonefly growth
and fecundity. Oecologia, 88, 277, 1991.
172. Pencoe, N.L. and Martin, P.B., Fall armyworm (Lepidoptera: Noctuidae) larval development and adult
fecundity on five grass hosts. Environ. Entomol., l l, 720, 1982.
173. Petty, SJ., Fur and feather - tawny owls in 1988. Entopath News, 92, 7, 1989.
174. Pianka, E.R., Evolutionary Ecology, 2nd ed., Harper & Row, New York, 1978, Chapter 5.
175. Pilson, D. and Rausher, M.D., Clutch size adjustment by a swallowtail butterfly. Nature, 333, 361, 1988.
176. Pilson, D. and Rausher, M.D., In response to Tabor. Oikos, 55, 136, 1989.
177. Pitnick, S., Male size influences mate fecundity and remating interval in Drosophila melanogaster. Anim.
Behav., 41, 735, 1991.
178. Prestidge, R.A., Instar duration, adult consumption, oviposition and nitrogen utilization efficiencies of
leafhoppers feeding on different quality food (Auchenorrhyncha:Homoptera). Ecol. Entomol., 7, 91, 1982.
179. Preszler, R.W. and Price, P.W., Host quality and sawfly populations: a new approach to life table analysis.
Ecology. 69, 2012, 1988.
180. Pritchard, I.M. and James, R., Leaf mines: their effect on leaf longevity. Oecologia, 64, 132, 1984.
18 1. Pritchard, I.M. and James, R., Leaf fall as a source of leafminer mortality. Oecologia, 64, 140, 1984.
182. Puttick, G.M. and Bowers, M.D., Effect of qualitative and quantitative variation in allelochemicals on a
generalist insect: iridoid glycosides and the southern armyworm. J. Chem. Ecol., 14, 335, 1988.
183. Raworth, D.A., McFarlane, S., Gilbert, N., and Frazer, B.D., Population dynamics of the cabbage aphid,
Brevicoryne brassicae (Homoptera: Aphididae) at Vancouver, British Columbia. 111. Development, fecundity
and morph determination vs aphid density and plant quality. Can. Entomol., 116, 879, 1984.
Factors Affecting Fecundity, Fertility, Oviposition, and Larviposition in Insects 173

184. Renwick, J.A.A., Chemical ecology of oviposition in phytophagous insects. Experientia, 45, 223, 1989.
185. Rutowski, R.L. and Gilchrist, G.W., Courtship, copulation and oviposition in the chalcedon checkerspot,
Euphydryas chalcedona (Lepidoptera: Nymphalidae). J. Nut. Hist., 21, 1109, 1987.
186. Sibly, R. and Monk, K.A., A theory of grasshopper life cycles. Oikos, 48, 186, 1987.
187. Slansky, F., Food consumption and reproduction as affected by tethered flight in female milkweed bugs
(Oncopeltus fasciatus). Entomol. Exp. AppL, 28, 1980.
188. Springer, P. and Boggs, C.L., Resource allocation to oocytes: heritable variation with altitude in Colias
philodice eriphyle (Lepidoptera). Am. Nat., 127, 252, 1986.
189. Stadler, E., Host plant stimuli affecting oviposition behavior of the eastern spruce budworm. Entomol. Exp.
Appl., 17, 176, 1974.
190. Stamp, N.E., Egg deposition patterns in butterflies: why do some species cluster their eggs rather than deposit
them singly? Am. Nut., 115, 367, 1980.
191. Stem, V.M. and Smith, R.F., Factors affecting egg production and oviposition in populations of Colias
philodice eurytheme Boisduval (Lepidoptera: Pieridae). Hilgardia, 29, 41 1, 1960.
192. Stewart, J.G. and Philogene, B.J.R., Reproductive potential of laboratory-reared Manduca sexta (Lepi-
doptera: Sphingidae) as affected by sex ratio. Can. Enromul., 115, 295, 1983.
193. Straw, N.A., The timing of oviposition and larval growth by two tephritid fly species in relation to host-plant
development. Ecol. Entomol., 14, 443, 1989.
194. Sutton, R.D., The effect of host plant flowering on the distribution and growth of hawthorn psyllids
(Homoptera: Psylloidea). J. Anim. Ecol., 53, 37, 1984.
195. Svard, L. and Wiklund, C., Fecundity, egg weight and longevity in relation to multiple matings in females
of the monarch butterfly. Behav. Ecol. Sociobiol., 23, 39, 1988.
196. Tauber, C.A. and Tauber, M.J., Phenotypic plasticity in Chrysoperla - genetic variation in the sensory
mechanism and in correlated reproductive traits. Evolution, 46, 1754, 1992.
197. Taylor, L.R., Longevity, fecundity and size; control of reproductive potential in a polymorphic migrant, Aphis
fabae Scop. J. Anim. Ecol., 44, 135, 1975.
198. Taylor, M.F.J. and Forno, I.W., Oviposition preferences of the Salvinia moth Samea multiplicalis Guenee
(Lep., Pyralidae) in relation to host plant quality and damage. J. Appl. Entomol., 104, 73, 1987.
199. Thomas, C.D., Butterfly larvae reduce host plant survival in vicinity of alternative host plants. Oecologia, 70,
113, 1986.
200. Thompson, J.N., Evolutionary ecology of the relationship between oviposition preference and performance
of offspring in phytophagous insects. Entomol. Exp. Appl.. 47, 3, 1988.
201. Thompson, J.N., Coevolution and the evolutionary genetics of interactions among plants and insects and
pathogens. In Pests. Pathogens and Plant Communities, (Eds., J.J. Burdon and S.R. Leather), Blackwell
Scientific Publications, Oxford, 1990, 249.
202. Tinkle, D.W., Wilbur, H.M., and Tilley, S.G., Evolutionary strategies in lizard reproduction. Evolution, 24,
55, 1969.
203. Trichilo, P.J. and Leigh, T.F., Influence of resource quality on the reproductive fitness of flower thrips
(Thysanoptera: Thripidae). Ann. Entomol. Soc. Am., 81, 64, 1988.
204. Unnithan, G.C. and Paye, S.O., Factors involved in mating, longevity, fecundity and egg fertility in the
maize stem-borer, Busseola fusca, (Fuller) (Lep., Noctuidae). J. Appl. Entomol., 109, 295, 1990.
205. van Amelsvoort, P.A.M. and Usher, M.B., A method for assessing the palatability of senesced leaf litter
using Folsomia candida (Collembola: Isotomidae). Pedobiologia, 33, 193, 1989.
206. van der Kraan, C. and van der Straten, M., Effects of mating rate and delayed mating on the fecundity of
Adoxophyes orana. Entomol. Exp. Appl., 48, 15, 1988.
207. van der Meijden, E., van Zoolen, A.M., and Soldaat, L.L., Oviposition by the cinnabar moth, Tyria
jacobaeae, in relation to nitrogen, sugars and alkaloids of ragwort, Senecio jacobaea. Oikos, 54, 337, 1989.
208. van Emden, H.F. and Bashford, M.A., A comparison of the reproduction of Brevicoryne brassicae and
Myzus persicae in relation to soluble nitrogen concentration and leaf age (leaf position) in the Brussels sprout
plant. Entomol. Exp. Appl., 12, 351, 1969.
209. van Emden, H.F. and Bashford, M.A., The performance of Brevicoryne brassicae and Myzus persicae in
relation to plant age and leaf amino acids. Entomol. Exp. Appl., 14, 349, 1971.
21 0. Walters, K.F.A. and Dixon, A.F.G., Migratory urge and reproductive investment in aphids: variation within
clones. Oecologia, 58, 70, 1983.
21 1. Walters, K.F.A., Brough, C., and Dixon, A.F.G., Habitat quality and reproductive investment in aphids.
Ecol. Entornol., 13, 337, 1988.
212. Ward, S.A., Leather, S.R., and Dixon, A.F.G., Temperature prediction and the timing of sex in aphids.
Oecologia, 62, 230, 1983.
21 3. Ward, S.A., Wellings, P.W., and Dixon, A.F.G., The effect of reproductive investment on pre-reproductive
mortality in aphids. J. Anim. Ecol., 52, 305, 1983.
2 14. Wassermann, S S . and Asami, T., The effect of maternal age upon fitness of progeny in the southern cowpea
weevil, CaNosobruchus maculatus. Oikos, 45, 191, 1985.
174 Insect Reproduction

215. Wassermann, S.S. and Mitter, C.. The relationship of body size to breadth of diet in some Lepidoptera. Ecol.
Entomol., 3, 115, 1978.
216. Watt, A.D., The effect of cereal growth stages on the reproductive activity of Sitobion avenue and
Metopolophium dirhodum. Ann. Appl. Biol., 91, 147, 1979.
217. Watt, A.D., The performance of the pine beauty moth on water-stressed lodgepole pine plants: a laboratory
experiment. Oecologin, 70, 578, 1986.
2 18. Watt, A.D., The effect of shoot growth stage of Pinus contorta and Pinus sylvestris on the growth and survival
of Panolisflammeo larvae. Oecologia, 72, 429, 1987.
2 19. Watt, A.D. and Leather, S.R., The pine beauty in Scottish lodgepole pine plantations. In Dynamics of Forest
Insect Populations, (Ed. A. A. Benyman), Plenum Publishing Corporation, New York, 1988, 243.
220. Webb, J.W. and Moran, V.C., The influence of the host plant on the population dynamics of Aciuin
russellae (Homoptera: Psyllidae). Ecol. Entomol., 3, 313, 1978.
221. Weliings, P.W., Leather, S.R., and Dixon, A.F.G., Seasonal variation in reproductive potential: a pro-
grammed feature of aphid life cycles. J. Anim. Ecol., 49, 975, 1980.
222. Werner, R.A., Influence of host foliage on development, survival, fecundity and oviposition of the spear-
marked black moth, Rheumaptera hasrara (Lepidoptera: Geometridae). Can. Entomol., 111, 317, 1979.
223. Wiklund, C. and Fagerstriim, T., Why do males emerge before females? A hypothesis to explain the
incidence of protandry in butterflies. Oecologia, 3 1, 153, 1977.
224. Wiklund, C. and Persson, A., Fecundity, and the relation of egg weight variation to offspring fitness in the
speckled wood butterfly Pararge aegeria, or why don't female butterflies lay more eggs? Oikos, 40.53.1983.
224a. Willers, J.L., Schneider, J.C., and Ramaswamy, S.B., Fecundity, longevity and caloric patterns in female
Heliothis virescens: changes with age due to flight and supplemental carbohydrate.J. Insect. Physiol., 33,803-
808, 1987.
225. Williams, K.S., The coevolution of Euphydryas chalcedona butterflies and their larval host plants. 111.
Oviposition behavior and host plant quality. Oecologia, 56, 336, 1983.
225a. Willson, H.R. and Trammel, K., Sex pheromone trapping for control of codling moth, Oriental fruit moth,
lesser appleworm and three tortricid leafrollers in a New York apple orchard. J. Econ. Entomol., 73,291,1980.
226. Wratten, S.D., Aggregation in the birch aphid, Euceraphispunctipennis (Zett.) in relation to food quality. J.
Anim. Ecol., 43, 191, 1974.
227.. Wright, L.C. and Cone, W.W., Population statistics for the asparagus aphid, Brachycorynella asparagi
(Homoptera: Aphididae) on different ages of asparagus foliage. Environ. Entomol.. 17, 699, 1988.
227a. Wyatt, I.J. and Brown, S., The influence of light intensity, daylength and temperature on increase rates of
four glasshouse aphids. J. Appl. Ecol., 14, 391, 1977.
228. Zalucki, M.P., The effects of age and weather on egg laying in Dannusplexippus L. (Lepidoptera: Danaidae).
Res. Popul. Ecol., 23, 3 18, 1981.
229. Zalucki, M.P., Brower, L.P., and Malcolm, S.B., Ovipositionby Dannusplexippus in relation to cardenolide
content of three Asclepias species in the southeastern USA. Ecol. Entomol., 15, 231, 1990.
 Chapter 8

PROTANDRY AND MATE ACQUISITION

Christer Wiklund

CONTENTS
I. Introduction ............................................................................................................... 175

11. Protandry Theory ....................................................................................................... 176

A. General ..............................................................................................................176
B. Male Benefit of Protandry .................................................................................. 178
C. Female Benefit of Protandry .............................................................................. 179
D. Conflicts Between Males and Females .............................................................. 179

111. Tests of Protandry Theory ........................................................................................ 180

A. Comparison Between Theoretical Models ......................................................... 180
B. Tests of Assumptions of Protandry Theory ....................................................... 182
C. Male vs. Female Cooperation/Conflicts Over Protandry ................................... 185

IV. Protandry and Sexual Size Dimorphism ...................................................................

188

V. Protandry in Relation to Monandry/Polyandry ........................................................ 190

VI. Protandry in Relation to Sex Ratio ...........................................................................

191

VII. Time and Life History .............................................................................................. 192

Acknowledgment ................................................................................................................. 194

References ......................................... ......... . ............ . ......... . . . . ..................... 195

I. INTRODUCTION
Mate acquisition can be a problem for females as well as males. However, as a rule sexually
active males greatly outnumber females, and hence, the male-biased operational sex ratio
makes mate finding a more severe problem for the average male compared to the average
female.
A variety of mate location systems have been observed in insects and include behaviors
where males spend there entire life in search of females to territoriality and lek polygyny.'
Territoriality and lekking are mate location systems that usually entail the opportunity for
interference competition between males for females, and hence, can lead to selection for large
male body size as in many mammals where males as a rule are larger than Mate
location systems where males search incessantly for females can be regarded as a situation
similar to scramble competition, where large size does not seem to confer any obvious
advantage.I2-l3Although the size of males may have important consequences for mate acqui-
sition, males also compete for females in the time dimension. In short, the fact that the

0-8493-6695-X/95/$O.M)+$.50
43 1995 by CRC Press. Inc.
176 Insect Reproduction

operational sex ratio is almost invariably male biased means that receptive females are in
shortage and hence males should time their appearance so that they maximize the number of
receptive females they encounter during their lifetime.
This was realized by Darwin14 who wrote,

It is certain that amongst almost all animals there is a struggle between the males for the
possession of the female. This fact is so notorious that it would be superfluous to give
instances. Hence, the females have the opportunity of selecting one out of several
males, ... Thus the males of our migratory birds generally arrive at their places of
breeding before the females, so that many males are ready to contend for each female.
... The majority of the male salmon in our rivers, on coming up from the sea, are ready
to breed before the females. So it appears to be with frogs and toads. Throughout the
great class of insects the males almost always are the first to emerge from the pupal state,
so that they generally abound for a time before any females can be seen. The cause of
this difference between the males and females in their periods of arrival is sufficiently
obvious. Those males which first migrated into any country, or which in the spring were
first ready to breed, or were the most eager, would leave the largest number of offspring;
and these would tend to inherit similar instincts and constitutions. It must be borne in
mind that it would have been impossible to change very materially the time of sexual
maturity in the females, without at the same time interfering with the period of the
production of the young - a period which must be determined by the seasons of the
year.

In addition, Darwint4quotes Wallace saying, "that in proportion as the individual moth

(Bombyx mori) is finer, so is the time required for its metamorphosis longer; and for this reason
the female, which is the larger and heavier insect, from having to carry her numerous eggs,
will be preceded by the male, which is smaller and has less to mature". Darwin goes on to say

Now as most insects are short-lived, and as they are exposed to many dangers, it would
manifestly be advantageous to the female to be impregnated as soon as possible. This
end would be gained by the males being first matured in large numbers ready for the
advent of the females; and this again would naturally follow, as Mr. A.R. Wallace has
remarked, through natural selection; for the smaller males would be first matured, and
thus would procreate a large number of offspring which would inherit the reduced size
of their male parents, whilst the larger males from being matured later would leave
fewer offspring.

This quotation may be taken as a case in point supporting Alan Grafen'sI5 assertion that
"Darwin (1 871) discovered almost everything important now known about sexual selection
and did so without measurement."
Darwin's general statement that males usually emerge before females "throughout the great
class of insects" might make further verification superfluous, but for the sake of record I still
feel that it can be useful to point out that this pattern, known as protandry, seems to be virtually
ubiquitous in most insect taxa, e.g., mayflies,16 katydids,I7-l8solitary bees and wasp^,^^-^^
b ~ t t e r f l i e s , and
~ ~ -m
~ o~ s q u i t o e ~ . ~ ~

11. PROTANDRY THEORY

A. GENERAL
Although Darwin14advanced a perfectly sound explanation for protandry, his appreciation
of the phenomenon seems not to have been generally realized, and from the late 19th up to
the middle of the 20th century a variety of hypotheses were advanced to account for the
Protandry and Mate Acquisition

relatively early emergence of males, including (1) prevention of inbreeding,22(2) enhance-

ment of the selective process by which unfit males are removed during the prereproductive
period,Ig (3) reduction of prereproductive death of females through facilitation of their rapid
fertilization after e c l o s i ~ n , and
~ ~ -(4)
~ ~improvement of male reproductive success because
individuals emerging early would have access to a larger number of receptive females than
males emerging late, a "rediscovery" of Darwin's idea.23
Wiklund and FagerstromZ6reviewed these ideas and pointed out that prevention of inbreed-
ing would be equally well served if females emerged before males, and so does not seem to
be capable of explaining the ubiquity of the relatively earlier emergence of males. The idea
that males emerge early so as to put themselves on the real "cutting edge" of selection
facilitating the weeding out of the relatively unfit is a notorious "for the good of the species"
explanation and as such utterly un-Darwinistic. Moreover, the idea that males emerge before
females so as to minimize prereproductive death of females is also group-selectionist,because
it explicitly states that early emerging males are risking their own prereproductive death so as
to minimize that of the females, the female sex being recognized as the more important for the
propagation of the species.
Although Darwin's appreciation of protandry has obvious merit, there still are aspects of
the early emergencelappearance of males that warrant further analysis; for example, the
statement that "those males which annually first migrated into any country, or which in the
spring were first ready to breed, or were the most eager, would leave the largest number of
offspring". On face value, this statement implies that early males do better than late males, and
hence, natural selection should lead to a situation where all males appear before the first
female. Hence, Darwin's theory, in a strict sense, predicts that the whole male population
should emerge before the onset of the female emergence period. This leaves the field open to
alternative predictions of how the male population should emerge in relation to that of the
female population. The prediction of how male and female emergence curves should be timed
relative to one another is greatly facilitated by mathematical modeling, and from the late 1970s
up to the present, a number of models have been advanced to explain the adaptive significance
of p r ~ t a n d r y . ~ ~These
. ~ O - ~models
~ all ask the question, given that we know the shape of the
emergence curve for the female population, how should the male population emerge in time.
Basically, these models fall into two categories that can be called "fixed distribution models"
and "flexible distribution models", respectively. The fixed distribution models assume that the
distribution of emergence times has a fixed shape,26.30-31.34-36 while the flexible distribution
models assume that there are absolutely no constraints on the shape of the male emergence
~ ~ r v e The . ~ fixed
~ - ~distribution
~ . ~ ~ models assume that, although the mean time of male
emergence is under selective control, the shape of the male and female emergence curves are
not. Instead, the shape of the emergence curves is the result of genetic factors interacting with
environmental factors like t e m p e r a t ~ r e . ~ ~By-~O contrast,
+ ~ ~ - ~the
~ flexible distribution models
assume that, in addition to the location in time of the emergence curve, the shape is also
determined by election.^^-^^.^^ The fixed distribution models conclude that the fitness distri-
bution around the male emergence peak is somewhat asymmetrical, which means that the
mean emergence time that is favored is not the emergence time that results in the highest
fitness (although it is reasonably close). The flexible distribution models result in a predicted
emergence curve for the male population where all males are expected to have equal mating
success, and hence, represents an Evolutionarily Stable Strategy (ESS).
Apart from the different optimization criteria, the two kinds of models are based on the
same set of assumptions: viz., that (1) male and female emergence curves are under genetic
control; (2) all females become fertilized (but see Zonneveld and met^^^); (3) Females mate
only once; (4) females always mate on the day of eclosion; (4) males are capable of multiple
mating; and (5) males are endowed with a given mortality rate which is independent of age.
Moreover, both models also assume that all males active on the same day have equal
opportunity of finding and mating with a virgin female, and hence, both kinds of models
 Insect Reproduction

I- 1 b

5 10 15 20 '/h (days)

FIGURE 1. Protandry calculated according to an optimality model assuming that the mean time of emergence for
the male population is under selective control, whereas the shape of the emergence curves of male and females are
not. Protandry (X) as a function of death rate of males (Ilk)when the duration of male and female emergence periods
are equal (here given by od and a?, the standard deviations of the population eclosion curves; hence 95% of the
population emerged within a time span equaling 40). (From Wiklund, C. and Fagerstrijm. T. 1977. Oecologia
(Berlin), 31:153-158.With permission.)

assume use of the optimization criterion that males are selected for maximizing the number
of matings during their lifetime, an end which is proportional to the number of females they
encounter and inversely proportional to the number of competing males.
Before proceeding to the quantitative predictions of the different models, it is necessary to
point out that the domain of protandry in insects is limited to systems where species have
discrete non-overlapping generations. Indeed, protandry cannot occur in environments where
reproductively active males and females are present throughout the year.

B. MALE BENEFIT OF PROTANDRY

The fixed distribution model presented by Wiklund and Fager~trijm?~ which assumed the
variance around the mean time of eclosion to be given, yielded the result that males emerging
at the peak of the male emergence curve, should maximize their expected number of matings
if the male population emerged some time before the female population (Figure 1). Moreover,
protandry should increase with the life expectancy of males, and with the duration of the
emergence period. In species where the life expectancy is only about 3 days, and virtually all
females and males emerge during a 5-day period, males should, on average, maximize the
number of expected matings by having their emergence curve peaking 1 day before that of
females. On the other hand, in species where the life expectancy of males is some 20 days,
and the duration of the emergence period of the male and the female population approaches
a month, the difference in time of mean emergence between the sexes should be approximately
10 days (Figure 1).
The flexible distribution models by Iwasa et al.32and Parker and C ~ u r t n e yassuming
,~~ that
the distribution of male emergence times can respond without any constraints to sexual
selection (so leading to an "ideal free" distribution), arrive at very similar predictions, hence,
giving considerable robustness to the general hypothesis that protandry can be viewed as a
mate acquisition strategy by males. The main difference between the quantitative predictions
of the two kinds of models is that the fixed distribution models predict that the very early and
very late males have lower fitnesses, whereas the flexible distribution models predict a slightly
smaller amount of protandry, and that the male eclosion curve should exhibit a rather abrupt
truncation point some time after the peak of male emergence because the loss of male mating
Protandry and Mate Acquisition 179

0 10 20 30 40 50 60 70 80 90 100
DAY NUMBER (t)

FIGURE 2. Protandry calculated according to a game theory model assuming that both the location and the shape.
of the male emergence curve are under selective control. Curve a is the female emergence curve, curve b is the male
emergence curve, and curve c is the male presence curve, calculated for the case where the total number of males and
females are 100, and the difference in preemergence and postemergence mortality is small. (From Iwasa, Y.,
Odendaa1,F.J.. Murphy, D.D., Ehrlich, P.R., and Launer, A.E. 1983. Theor. Popul. Biol. 23:363-379. With permission.)

opportunity is more severe for males emerging several days after the male emergence peak
than for males emerging as many days before (Figure 2).

C. FEMALE BENEFIT OF PROTANDRY

The models discussed up to now have all been male chauvinistic, asking the question of
how males should time their emergence to maximize their mating opportunities when the
emergence curve of the female population is given. However, even if the approximate time
of year when females should emerge may reasonably be given by the timing of the appearance
of suitable host plants etc., cf. Darwin'si4 assertion that "it would have been impossible to
change very materially the time of sexual maturity in the females, without at the same time
interfering with the period of the production of the young - a period which must be
determined by the seasons of the year", it seems reasonable to assume that also females can
be selected to time their period of emergence relative to that of males. So females may be
selected to emerge so as to minimize death before reproduction. This is best achieved by
females emerging at the time when there is a maximum of living males in the population. Since
even very short-lived insects do have some life expectancy, the peak of the male presence
curve must always occur some time after that of the male eclosion curve (cf. Figure 2), and
hence, it is easy to understand that females benefit from emerging later than males. In other
words, protandry may be viewed also as a female mating strategy by which the time they spend
unmated is minimized.30

D. CONFLICTS BETWEEN MALES AND FEMALES

Although both males and females benefit from protandry, it is conceivable that there could
be considerable conflict between the sexes as to how profound the difference in emergence
between the sexes should be. However, when comparing the optimal degree of protandry
between the sexes, Wiklund and Fagerstrom30 found that there seemed to be virtually no
conflict between the sexes for cases where the eclosion periods were similar for the two sexes
(Figure 3). Using a slightly different approach, were ESS models for optimal time of emer-
gence where calculated for males and females separately, Zonneveld and M e t ~ , also
3 ~ arrived
at the same general conclusion, i.e., that although male and female ESS's were not identical,
there is only a small conflict of interest.
180 Insect Reproductioti

FIGURE 3. The degree of conflict between the sexes in the decision when to emerge (X) as a function of the
standard deviation of male ( 0 8 ) and female ( a ? ) eclosion curves for different values of the life expectancy of males
(1A). Positive X-valuessignify that the degree of protandry that is optimal for the males is less than that being optimal
for the females; negative X-valuessignify the reverse. (From Fagerstrom, T. and Wiklund, C. 1982. Oecologia (Berlin)
52:164-166. With permission.)

111. TESTS OF PROTANDRY THEORY

A. COMPARISON BETWEEN THEORETICAL MODELS

There are several ways in which protandry theory can be tested, but unfortunately the exact
quantitative predictions are difficult to test in the field because the development time of insects
is so dependent on temperature. Natural selection can certainly act on how insects respond to
temperature, and so there will be individual variation in growth rate even under constant
temperature; however, since insects are so strongly ectothermic, variation in growth rate
between individuals will be superseded by the action of temperature. For instance, the actual
observation of protandry in nature bears witness to the fact that the development time of males
is shorter than that of females (Figure 4). However, the exact difference in time between the
peak of male and female eclosion is likely to be strongly influenced by temperature. If a drastic
cold spells occurs just after the peak of male emergence, the difference in mean time of
emergence is likely to be quite accentuated. However, if the weather is cold up to the point
of the male emergence peak, but warm thereafter, the difference in time of emergence between
the sexes is likely to be considerably smaller.
The flexible distribution models of Iwasa et al.32 and Parker and C ~ u r t n e y and , ~ ~ the
optimization model of Wiklund and F a g e r ~ t r o mand ~ ~ Fagerstrom and Wikl~nd,~O were
compared by B ~ l m e r , who
~ ' found difficulty in accepting that natural selection could result
in a emergence curve that is abruptly truncated after its maximum, as predicted by the flexible
distribution models. Bulmer says, "It is difficult for selection to alter the shape of the
distribution of a polygenic metric character because recombination tends to restore approxi-
mate normality each generation (Bulmer3'). It is particularly difficult to envisage a mechanism
which could produce a distribution with an abrupt truncation point as required in the ideal free
distribution". Moreover, even if the prediction of a specific truncation of the male emergence
curve is somewhat relaxed, Bulmer31argues that "it seems unlikely that either the shape or the
Protandry and Mate Acquisition 181

-~ a ~ " ' l " l

81
~ l "
A
' l i l
male presence
l l
\ a male emergence

---
t': female emergence
7
1 AA
- m(t)

-.-
A *

- I
l
'
l

\ A
- \

i
l

-
.
8
A I
l
'l A
1
7
I $
I
- l \l A

- A '8
I \
\
$A

l '\ A

- l' '88
8

0 2 4 6 8 10 1 2 1 4 1 6 1 8 2 0 2 2 2 4 2 6 2 8 3 0 3 2 3 4 3 6
DAY NUMBER ( t )

FIGURE 4. Number of males present, emerging females, and emerging males each day for Euphydryas editha data
(points) and theoretical values (lines) when the preemergence mortality is 0.04 (From Iwasa, Y., Odendaal, F.J.,
Murphy, D.D., Ehrlich, P.R.,and Launer, A.E. 1983. Theor. Popul. Biol. 23:363-379. With permission.)

variance of the distribution of male emergence times will be able to respond to selection in
the way predicted by (flexible distribution model) theory" [(flexible distribution model) added
for clarification by the present author].
Moreover, examination of emergence times for males and females from the field have
consistently failed to give evidence of any truncation point in the eclosion of the male
population (Figures 4 and 5; Iwasa et al.;32Parker and C ~ u r t n e yBaughman
;~~ et al.36).Hence,
the exact prediction of the flexible distribution models about the shape of the male distribution
curve have not been corroborated by observation in the field. Moreover, the empirical data on
emergence of male and female populations show that they are strongly overlapping, hence,
lending little support to the strict interpretation of Darwin's statement predicting that the
whole male population should emerge before the onset of female emergence. Additionally,
B a ~ g h m a nwas
~ ~ able to test the mating success of early, intermediateiy, and late emerging
males of Euphydryas editha by brushing male genitalia with fluorescent dust of three different
colors, and later collecting females to see if the dust color from the early emerging group of
males was more commonly found on the outer part of female genitalia. However, the result
was that male mating success appeared to be independent of time of emergence, showing again
that Darwin's strict prediction that early emerging males have higher mating success does not
hold true. However, Baughman's results do not allow any conclusion as to whether the flexible
distribution models of Iwasa et al.32 and Parker and C ~ u r t n e y or, ~ ~the fixed distribution
models of Wiklund and F a g e r ~ t r o mand
~ ~ Fagerstrom and Wiklund?O hold true, as the first
predicts (or rather is based on the assumption) that all males have equal mating success, the
latter assumes that males emerging at the peak of the male emergence curve should have
maximal mating success.
 Insect Reproduction

1 3 5 8 10 12 15 17 18 19 22 24 26 29
DAY

FIGURE 5. Graphical representation of male and female emergence patterns for Euphydryas editha in 1981. (From
Baughman, J.F., Murphy, D.D., and Ehrlich, P.R. 1988. Theor. Popul. Biol. 33:102-113. With permission.)

B. TESTS OF ASSUMPTIONS OF PROTANDRY THEORY

Apart from testing quantitative predictions, another way to test the general hypothesis that
protandry is a mate acquisition strategy is to test the relevance of its assumptions. As pointed
out previously, males cannot emerge before females in environments when reproductively
active males and females are found throughout the year. Hence, insofar as the earlier emer-
gence of males relative to females is associated with a cost of some kind, males should not
be expected to have shorter development time compared to females in populations without
discrete generations (i.e., where there can be no protandry). A case in point is served by the
speckled wood butterfly Pararge aegeria, which has a Palearctic distribution. In southern
Sweden and England, this species can spend the winter either as half grown larvae or in the
pupal stage, both of which have a bivoltine life cycle ( G ~ d d a r dWiklund,
;~~ unpublished).
However, although P. aegeria consequently exhibits four flight periods annually, the different
generations are largely nonoverlapping. In Spain, adults of the speckled wood cannot be found
from November to February, but are multivoltine through the rest of the year with different
generations overlapping quite strongly (Garcia-Barros, unpublished). On the island of Ma-
deira, P. aegeria flies throughout the year with little, if any, seasonality. In accordance with
assumptions of protandry theory, populations from southern Sweden and England exhibit
strong protandry, whereas populations from Spain and Madeira do not exhibit significant
protandry at all (Figure 6; Nylin et al.40).This is quite remarkable as there is no variation in
the degree of sexual size dimorphism between the four populations - males weight being
some 80% of female weight in all four populations (Table 1; Nylin et al.40).
Another general prediction of protandry theory, again assuming that shorter development
time and higher growth rate carries a cost, is that male development time should be shorter
than that of females only when individuals become sexually active immediately following
development. Hence, in butterflies that overwinter as adults and do not mate until after
hibernation, males should not be expected to emerge before females in the summer. However,
protandry should be expected in the spring with males emerging from hibernation earlier in
the season relative to females. This kind of life cycle is exhibited by the brimstone butterfly,
which emerges from the pupa in midsummer but where individuals do not mate until after
hibernation. As predicted, the development time from egg to adult is not shorter for males
Protandry and Mate Acquisition 183

Development time (days) in P. aeqeria

Sweden England Spain Madeira

53

48

43

38

FIGURE 6. Total development time (mean +/- S.E.) for the two sexes in four potentially multivoltine populations
of Pararge aegeria as measured in two replicates. Males are shown to the left in each pair (cohort) and the differences
between the sexes show the degree of protandry. Populations from south Sweden and England represent more strongly
seasonal environments that Spain or Madeira, and here protandry was significant. (From Nylin, S., Wiklund, C.,
Wickman, P.-O., and Garcia-Barros, E. Ecology, 74: 1414-1427. With permission.)

TABLE 1
Sexual Differences in Weight (means + S. E.) at Different
Developmental Stages in Cohorts of Five Populations
of Pararge aegeria

Pupal weight Female/Male

Cohort Sex N (nig) pupal weight ratio

Central Sweden Males 15 140.7 + 2.9

Females 20 173.0+1.9 1.23
South Sweden I Males 24 137.9 + 3.2
Females 30 165.7 + 3.2 1.20
South Sweden II Males 13 153.5 + 3.7
Females 10 172.0 + 3.8 1.12
England I Males 34 148.9+ 1.5
Females 24 185.5 + 2.8 1.25
England II Males 10 137.3 + 1.3
Females 14 163.6 + 2.0 1.19
Spain I Males 20 134.4 + 2.4
Females 25 169.1 + 3.2 1.25
Spain II Males 12 142.6 + 3.3
Females 7 170.1 +5.2 1.19
Madeira Males 24 132.7 + 2.0
Females 20 153.1+2.1 1.15

Note: Sexual dimorphism in weight was highly significant (p <0.01 - p <0.001; ANOVA) in all
experiments. Roman numerals indicate replicate number.

From Nylin, S., Wiklund, C., Wickman, P.-O., and Garcia-Barros, E. Ecology. 74: 1414-1427.
With permission.
 Insect Reproduction

TABLE 2
Pupal Weights (mean + S. E) of
Diapausing and Nondiapausing
Leptidea sinapis a t 23OC

Pupal weight
(mg) N

Diapause Females 58.6 + 0.9 26

Males 56.1 + 1.2 21
Difference 2.5
Non-diapause Females 55.5 + 0.9 22
Males 51.7 + 1.2 21
Difference 3.8

From Wicklund, C. and Solbreck, C. 1982. Evolution

36:56-62. With permission.

-m
m
4
0
3
.-
L Males
z 2 Females
-0

E
1

0
40 41 42 43 44
Development time (days)

FIGURE 7. Development time of Gonepteryx rhamni from egg to adult when reared at 20°C and a 20 h daylength.
The mean development time was 41.7 +l- 0.3 days for the 12 males and 41.6 +l- 0.4 days for the 8 females. Note
that this butterfly emerges during summer and does not mate until after hibernation, and so protandry is not relevant
at the time when the adults emerge (Wiklund and Lindfors, in preparation.)

compared to females, but in the spring males can be seen on the wing much earlier than
females (Table 2; Figures 7 and 8; Wiklund and Lindfors, unpublished).
Yet another way to test whether protandry is a mate acquisition strategy or not is to compare
alternative hypothesis for the shorter development time of males relative to females. This can
be done in bivoltine species in which protandry is effectuated differently in the two genera-
tions. The hypothesis that protandry is a mate acquisition strategy is based on the idea that the
difference in time of emergence between the sexes is selected for, per se. However, in most
insects males are smaller than females, and so the shorter development time of males might
be explained by the simple fact that it takes less time to acquire a smaller soma. These two
hypotheses yield different predictions for the degree of protandry in the two generations for
a butterfly that overwinters in the pupal stage like the wood white Leptidea sinapis. In Sweden,
the first generation of this butterfly emerges in the middle of May, and the first eggs are laid
in late Maylearly June. These early eggs give rise to larvae that develop directly and produce
pupae from which a second generation of adults is formed that can be seen on the wing in late
Julylearly August (late eggs laid by first generation females yield larvae that develop into
overwintering pupae). The eggs laid by the second generation females give rise to larvae that
 -
Protandry and Mate Acquisition

l00 - H

80 -
a-
s
2

g
4-
46
43-
$:
20 -
0 1 . 1 . 1 . 1 . 1

0 10 M 30 10 M
April May

FIGURE 8. Number of male and female Gonepreryx rhamni observed during 1 hour counts by car along roads in
central Sweden during April (days 1-30) and 1-10 of May (days 31-40) in 1992. The first males were observed on
April 9, whereas the first females were observed on May 3, i.e., protandry measured as the difference in time of
appearance amounted to more than 3 weeks. (Wiklund and Lindfors, in preparation.)

produce overwintering pupae from which the adult butterflies emerge the following spring.
The wood white can only survive the winter in the pupal stage, and interestingly, development
is synchronized during the winter. This is evidenced both by the observation that the pupae
spend the winter in an undeveloped stage, being semitransparent when held against a strong
light, and because there is no difference in time of emergence between the pupae formed in
JuneNuly by the offspring from the first generation of adults and those formed in August by
the offspring from the second generation of adults. This means that the early emergence of
males in the directly developing summer generation is brought about both by male larvae and
pupae, having shorter development time compared to females. However, in the overwintering
generations, the earlier emergence of males is brought about only because of the difference
in postdiapause development time between male and female pupae. Hence, if males emerge
before females only because they are smaller, the difference in time of appearance between
the sexes should be considerably smaller in the overwintering spring generation than in the
directly developing summer generation. However, if the difference between the sexes in time
of emergence is selected for, per se, protandry should be expected to be equal in the two
generations, given equal costs, regardless of the fact that it is brought about differently.
Indeed, when reared in the laboratory under a constant temperature of 23OC, the males emerge
some 2 to 2.5 days earlier than females both in the diapausing and the nondiapausing
generations (Figure 9). Hence, the hypothesis that protandry in the wood white is an incidental
side effect of the sexual size dimorphism was not supported, whereas the hypothesis that the
difference betwen the sexes in time of emergence is selected for, per se, was supported. The
observation that protandry is similar in directly developing and diapausing generations (and
that it is brought about by a relative increase in the sex difference in postdiapause development
time) has also been made in the black swallowtail butterfly Papiliopolyxenes (Lederhouse et
al.41)and for a number of Swedish pierid and satyrid butterflies (Wiklund and F ~ r s b e r g ~ ~ ) .

C. MALE VS. FEMALE COOPERATION/CONFLICTS OVER PROTANDRY

As previously mentioned, theoretical models predict that males and females both have
similar interests relating to how many days earlier males should emerge prior to females. The
case of the bivoltine wood white butterfly can also be used to analyze this aspect of protandry
because the results suggest that the two sexes take turns in realizing protandry. When
overwintering pupae of L. sinapis are kept at 4OC diapause is broken after approximately 2.5
months, after which the duration of the time pupae are kept at low temperature does not seem
 Insect Reproduction

Difference in pupal development time

Difference in brml development time

FIGURE 9. Sex differences in egg-larval and pupal development times for Leptidea sinapis under diapause and
nondiapause development, respectively. The degree of protandry is similar in the two generations although it is
brought about only by the sex difference in postdiapause development time in the diapausing generation, whereas it
is the accumulated sex difference in development time from egg, larval, and pupal development in the non-diapausing
generation. (From Wiklund, C., and Solbreck, C. 1982. Evolution 36:56-62. With permission.)

to affect the postdiapause development time of male pupae (Figure 10). However, postdiapause
development time is strongly dependent on the time spent at low temperature, and the
statement that female pupae take some 2 to 2.5 days longer to produce adults after hibernation
holds true only when pupae are kept at 4OC between 2.5 to 4 months. When female pupae are
kept at low temperature for longer than that, postdiapause development time becomes shorter
with increasing time spent in 4OC (Figure 10). Hence, when females are kept at low tempera-
ture for 8 months, the difference in postdiapause development time between the sexes is only
about 0.5 days, i.e., equal to the sex difference in pupal development time in the directly
developing generation (cf. Figure 9). The observation that females, but not males, vary their
postdiapause development time indicates that it is the female sex that effectuates protandry in
the diapausing generation by not developing a maximum speed for intermediate times spent
at low temperature. Given that the same time difference in the appearance of the sexes is
optimal in both generations of the wood white butterfly, it appears that the degree of protandry
brought about by females corresponds to the optimal one of 2 to 2.5 days when the pupae
hibernate for an intermediately long period of time. Wiklund and S01breck~~ have argued that
up to 4 months spent at 4OC corresponds to the average winter duration in Sweden, whereas
longer times at low temperatures suggest that winter is longer than usual. This implies that
spring will be later than usual, and given that it is beneficial for females to emerge so early
in the season that their offspring will have time to develop directly, females may have to make
a trade off between emerging early and effectuating optimal protandry. According to this
scenario, females opt for optimal protandry for average duration of the winter period, whereas
they effecuate suboptimal protandry after unusually long winters to avoid the penalty of
emerging too late in the season.
According to the idea that it is important for females not to emerge too late in the season,
the second generation of females will be even more pressed for time because their offspring
must complete development and pupate before the autumn temperatures get too cold for
development. Hence, it is unlikely that the female sex will effectuate protandry in the directly
Protandry and Mate Acquisition

Chilling time (months)

FIGURE 10. Postdiapause development times for male and female pupae of Leptidea sinapis in relation to the
length of chilling ( + 4°C) period. (From Wiklund, C. and Solbreck, C. 1982. Evolution 36:56-62. With permission.)

developing generation. Hence, the males will have to shorten their development time if they
are to emerge before the females and, as pointed out by Singer,44given that males and females
grow at a similar rate, protandry can only be achieved by the male sex becoming smaller.
Although few issues in evolutionary biology have proved less easily tractable than that of
optimal size (cf. Sibly and C a l o ~ ~insofar
~ ) , as optimal male size is concerned, it ought to be
achieved by the individuals from the first generation of wood whites that enter the diapause
developmental pathway and have some 2 months longer to complete development before the
winter compared to the offspring from the second generation. Indeed, when comparing
development times of males and females from the two generations of wood whites, it turns out
that the difference in egg-larval development times between the sexes was only some 0.5 days
for the diapausing individuals, whereas it was some 1.5 for the directly developing larvae (cf.
Figure 9). When comparing the difference in weight between male and female pupae, the
difference was considerably larger in the directly developing generation (albeit not signifi-
cantly so), which suggests the tendency that it is the male sex that shortens its development
time relative to the female sex, and in doing so becomes smaller (Table 2). Hence, it may be
that males in the directly developing generation have to make a trade off between optimal size
and achieving optimal protandry. Since the degree of protandry achieved in the directly
developing generation was very similar to that in the diapausing generation, this can be
interpreted as the males favoring the achievement of optimal protandry to that of optimal size.
This section has been based on studies of one butterfly, but the pattern that both sexes
achieve protandry is also exhibited by the green-veined white butterfly Pieris napi, in which
females also seem to achieve to effectuate protandry in the overwintering generations, with
males effectuating protandry in the directly developing generation (according to the same
argumentation as for the wood white; cf. Forsberg and W i k l ~ n d ~However,
~). there is one
interesting difference between protandry in the two generations of the two butterflies, insofar
as the time difference in emergence of the sexes is smaller in the directly developing
generation of P. napi, whereas the difference in size between the sexes is just as profound as
 Insect Reproduction

DIRECT DEVELOPMENT

1.2

DIAPAUSE DEVELOPMENT
V)

Protondry (days)

FIGURE 11. The relationship between malelfemale size dimorphism and protandry. For directly developing
generations: y = 0 . 0 2 ~+ 0.03; r = -0.327; n = 8; p c0.3: For diapause developing generations: y = -0.02~+ 0.99;
r = -0.51 1 ; n = 14;p <0.05, one-tailed test. Values for pierids are indicated by ( 0 )and values for satyrids are indicated
by (0): The numbers refer to the following species (2)Pieris brassicae; ( 3 )Pieris rapae, (4)Pieris napi, (5) Pontia
daplidice, ( 6 ) Anthocharis cardamines, (7) Colias hyale, (9) Colias palaeno, (1 1 ) Leptidea sinapis, (19) Pararge
aegeria, (20) Lasiommata megera, (21)Lasiommata maera, (22) Lasiommata petropolitana, (23)Lopinga achine.
(From Wiklund, C. and Forsberg, J . 1991. Oikos 60:373-381. With permission.)

in the diapausing generation. This may suggest that males of the green-veined white, when
having to make a trade off between achieving optimal size and protandry, favor achieving
optimal size to achieving optimal protandry. This, in turn, may be understandable in terms of
the different mating systems and the influence of relative male size on reproductive success
in different matings systems, as will be discussed below.

IV. PROTANDRY AND SEXUAL SIZE DIMORPHISM

In two families of butterflies, Wiklund and F0rsbe1-g~~ have shown that relative male size
increases with the degree of female polygamy. The causal mechanism underlying this pattern
is suggested to be sperm competition, where large males are reproductively favored because
the size of the spermatophore that males transfer to females at mating is proportional to male
size, and the time following mating, during which females are unreceptive to male courtship
is, in turn, proportional to the size of the spermatophore the female received at mating
(Rut~wski;~' O b e r h a ~ s e rSviird
; ~ ~ and W i k l ~ n dKaitala
; ~ ~ and Wiklund, submitted). As pointed
out by "If males are to emerge before females as a result of selection for protandry,
they must have shorter development times than females, and hence will be smaller, if they
grow at the same rate as larvae". Hence, a negative relationship between malelfemale size
dimorphism and protandry should be expected, especially among directly developing genera-
tions where development is uninterrupted and males can only emerge before females by
shortening their development time, in the process becoming smaller than females. However,
a comparative study of six pierid and two satyrid butterflies failed to give any evidence of a
negative relationship between malelfemale size dimorphism and protandry (Figure 11;Wiklund
and F o r ~ b e r g ~ ~ ) .
Protandry and Mare Acquisition 189

FIGURE 12. Larval development times of males and females of Pieris napi under direct and diapause develop-
ment, respectively, at three different temperatures 17OC and a 22 h daylength, at 20°C and a 22 h daylength, and
pooled results for larvae reared at 23°C and 18:OO. 18:15, 18:30, 18:45 and 19:00 h daylengths. Although directly
developing males complete their development several days before their brothers, the mean weight of directly
developing pupae is always higher than that of diapausing pupae (although usually not significantly heavier). (From
Wiklund, C., Nylin, S., and Forsberg, J. 1991. Oikos 60:241-250. With permission.)

The absence of such a negative relationship can only be understood if Singer's44assump-

tion of male and female larvae growing at the same rate is violated. This explanation is
corroborated by a study on the growth rate and development times of male and.female larvae
of the green-veined white butterfly P. napi, under direct and diapause development. In this
species, where males are actually larger than females (i.e., they have larger mass and longer
wings compared to females at eclosion), the development time of males is indeed longer than
that of females in individuals that develop according to a univoltine life cycle and diapause
in the pupal stage before emerging as sexually active adults the following spring. However,
when individuals develop directly, male development is considerably shorter than that of
females, although the malelfemale size dimorphism remains unaltered. In fact, sibling males
that develop directly become larger than brothers that enter diapause development while still
completing their development a couple of days earlier (Figure 12). Wiklund et al.50showed
this by rearing larvae of P. napi at the critical photoperiod, at which a proportion of the larvae
entered direct development and part entered diapause development, which means that the
results shown in Figure 12 relate to differences between individuals that were experiencing
virtually identical temperature (and photoperiodic) conditions being reared in the same
climatic chamber. These results show that larvae of P. napi exhibit considerable sex-related
plasticity in growth rate, a phenomenon that has been demonstrated also in P. aegeria (Nylin
et al.51).This capacity for varying growth rate is surprising, in view of the belief that insect
larvae as a rule are heavily preyed upon and hence might be expected to develop at some
maximum rate. However, the demonstration that butterfly larvae do have this ability can only
be understood in terms of high growth rates carrying some kind of cost, as has been indeed
shown in birds and mammals (Clutton-Brock et aL8). Wiklund et al.50envisage a cost in terms
of high growth rates being associated with higher vulnerability to disease and higher
susceptability to death from starvation, but these hypotheses are as yet untested.
A negative relationship between malelfemale size dimorphism should also be expected
among diapausing generations, but here the causal relationship could easily be the reverse of
that relevant for directly developing generations. For example, among species that overwinter
in the pupal stage, protandry results only because of the sex difference in postdiapause
 Insect Reproduction

DIRECT DEVELOPMENT

Female polygamy

FIGURE 13. The relationship between protandry and female polygamy: For directly developing generations: y =
- 0 . 6 9 ~+ 2.3; r = -0.248; n = 8; p ~ 0 . 3For
. diapause generations: y = - 3 . 1 4 ~+ 7.14; r = -0.521; n = 14; p <0.05.
Values for pierids are indicated by ( 0 ) and values for satyrids are indicated by (0). the numbers refer to species as
in Figure 1 1 . (From Wiklund, C. and Forsberg, J. 1991. Oikos 60:373-381. With permission.)

development time, and it stands to reason that the morphogenesis of an adult butterfly from
the undifferentiated state of an overwintering pupa takes less time for a small soma compared
to a larger one. Indeed, a comparative study of nine pierid and five satyrid butterflies did
demonstrate a negative relationship betwen malelfemale size dimorphism and protandry in
diapausing generations (Figure 11). In view of the phenotypic plasticity in growth rate of
larvae in the directly developing generations, the demonstrated negative relationship between
malelfemale size dimorphism in the diapausing generations suggest that post-diapause devel-
opment rate of pupae is less phenotypically plastic.

V. PROTANDRY IN RELATION TO MONANDRYPOLYANDRY

All of the first generation of protandry models use the important assumption that females
mate only once. When females are monandrous and always mate soon after eclosion, this
strongly compresses the time when males can find females to mate with, and so favors early
male emergence because the penalty of emerging late (when no receptive females can be
found) is more severe than emerging early (when males face the risk of prereproductive death
before the appearance of the first females). In a general sense, selection for protandry should
be relaxed under mating systems where females mate more than once. Hence, a negative
relationship might be expected between protandry and the degree of female polygamy.
A comparative analysis of protandry in relation to female polygamy failed to give any
significant relationship for directly developing generations of six pierid and two satyrid
butterflies, whereas a weak negative relationship was found under diapause development in
nine pierid and five satyrid butterfly species (Figure 13; Wiklund and F o r ~ b e r g ~Although
~).
the benefit conferred upon males by emerging early decreases with increasing degree of
Protandry and Mate Acquisition 191

females polygamy, selection for protandry may still be important insofar as mating with virgin
females is more beneficial than mating with nonvirgin females. The observation that the
number of eggs laid per day decreases with age in many insects suggests this (cf. Labine;52
Karlsson and W i k l ~ n d ;Svard
~ ~ . ~and
~ Wikl~nd;".~~ Z ~ n n e v e l d ~and
~ ) ,may hence explain the
relative lack of a strong negative relationship between protandry and polyandry. Indeed, a
recent theoretical model shows that the influence of female polygamy on protandry seems to
be relatively small, as a result of the fact that the reproductive value of females decreases with
female age (Zonne~eld~~). Moreover, Zonneveld's model shows that the influence of female
polygamy also decreases with decreasing life expectancy of females, which is but another way
in which the importance of mating with young females is accentuated on the part of males.
It is obvious that protandry, sexual size dimorphism, and female polygamy are interrelated
in several ways, conferring that the direction of causality may be difficult to discern. However,
in some cases the direction of causality seems clear. For instance, it is conceivable that the
mating system maintained by females affects the optimal time of male eclosion in relation to
females, whereas it seems less likely that the degree of protandry should affect female mating
frequency.
However, the relationship between development time (and protandry) and size may be
more variable between species. For obvious reasons, development time should have conse-
quences for the size acquired, and if growth rate were constant, a given development time
would result in a given predictable size. Hence, in species where there is a conflict between
both optimal development time and size, "trade offs" have to be made. In this situation, it is
conceivable that the mating system maintained may have consequences for the order of
priority. For example, it is possible that males belonging to strictly monandrous species place
priority on emerging early (i.e., favoring protandry at suboptimal size), whereas males
belonging to polyandrous species may put priority on achieving large size and emerging at a
suboptimal time. Data on the direction of "trade offs" made in the directly developing
generations of the monandrous L. sinapis and the polyandrous P. napi are consistent with this
view (Wiklund and S ~ l b r e c kForsberg
;~~ and W i k l ~ n dWiklund
;~~ and F ~ r s b e r g ~ ~ ) .

VI. PROTANDRY IN RELATION TO SEX RATIO

In temperate areas, directly developing generations may be severely pressed for time, and
as a consequence many alleged bivoltine species are likely to be only partially so. The
establishment of what proportion of individuals that do develop directly requires some sort of
rearing, because the relatively common observation that the second flight generation holds
more individuals than the first does not allow the conclusion that all of the offspring from the
first generation developed directly. Indeed, in Sweden, the second flight generation of P. napi
seems to be larger than the first, but outdoor rearing of larvae showed that all cohorts of eggs
laid by females in the first generation produced some individuals that entered the univoltine
developmental pathway and produced diapausing pupae (Wiklund et al.50).The fact that many
second generations that develop directly are hard pressed for time is of relevance here because
it may interfere with the ability of second generation males to achieve protandry, and, as a
result, may influence male propensity to develop directly.
The flexible distribution models for protandry of Iwasa et al.32show that not only should
males start to emerge before females, but also cease to emerge before females (B~lmer;~'
Iwasa et al.;32Parker and C ~ u r t n e y ~More
~ ) . specifically, even if the female eclosion curve is
smooth with one peak, the male eclosion curve should be abruptly truncated before female
emergence has ceased (cf. Figure 2). In temperate insects having partial second generations,
we predict that males should enter diapause development at an earlier date than should
females, as a result of late emerging males being penalized in terms of fewer mating oppor-
tunities, and so postpone emergence until next season. This prediction follows logically from
 Insect Reproduction

DAY LENGTH (HOURS)

FIGURE 14. The difference between males and females in the propensity to develop directly in response to
photoperiod, (left) as predicted by theory, and (right) as shown by Pieris napi reared at 23OC and the daylengths 18:00,
18:15, 18:30, 18:45 and 19:00 h. The number of individuals reared are shown at the bottom of the figure. (From
Wiklund, C., Wickman, P.-0.. and Nylin, S. 1992. Evolution 46519-528. With permission.)

theories on optimal timing of eclosion of males relative to females, when applied to partially
bivoltine insect populations.
The decision to diapause or to develop directly is usually mediated by response to environ-
mental stimuli of which daylength is the most important, but also temperature can play a role
(Danilev~kii;~~ Tauber et al.;58DanksS9).Hence, we predict that the mechanism by which
males enter diapause development at an earlier date than females will be that of the male
propensity to enter diapause development being shifted towards longer day lengths compared
to that of females (Figure 14). More specifically, we predict that the difference in diapause
propensity between the sexes should increase when approaching the point where all individu-
als enter diapause development, because directly developing males will be most severely
penalized when few or no females develop directly. This means that a higher percentage of
females should develop directly at any given photoperiod in the interval of day lengths that
produce mixed broods, i.e., broods in which some of the individuals develop directly and
others enter diapause development. As a result, partial second generations should be female
biased, on the assumption that the primary sex ratio is unity. As a corollary, first generations
of populations exhibiting partial bivoltinism should be male biased because some males of the
first generation are from the previous year.
These predictions have now been supported in a number of ways. Firstly, Wiklund et
showed that a higher proportion of females of P. napi developed directly when reared in
climatic chambers under constant photoperiod and temperature conditions that produced
mixed broods. The higher propensity for females to develop directly was also demonstrated
by outdoor rearings of cohorts of larvae of P. napi and Pieris rapae that were initiated
throughout the season (Figure 15). Moreover, the prediction that partial second generations
should be female biased was supported by laboratory rearings at constant temperature of P.
napi (Pieridae), P. aegeria (Satyridae), and Polygonia c-album (Nymphalidae) under critical
day length conditions, producing female-biased sex ratio under direct, and male-biased sex
ratio under diapause development (Table 3).

VII. TIME AND LIFE HISTORY

This whole chapter has really been about time. In Stephen Vogel's61 book Life's Devices,
Woody Allen is quoted as having said that "Time is nature's way of keeping everything from
happening all at once", but as may have been demonstrated by this chapter, time may play
other roles in the lives of insects. In short-lived animals that die before they show any evidence
of senescence (as evidenced by the good fit of models that assume daily survival, probability
is independent of age), it is important that all of the important events of life happen, if not "all
at once", still within a very short time period.
Protandry and Mate Acquisition

Date of larval hatch

&Y June July *ug

FIGURE 15. Percent directly developing males and females of Pieris rapae and Pieris napi reared outdoors
throughout the season. Cohorts of approximately40 larvae were reared, and the hatching date of each cohort is shown
above the bars. (From Wiklund, C., Wickman, P.-0.. and Nylin, S. 1992. Evolution 46519-528. With permission.)

TABLE 3
Number of Males and Females that Enter Direct or Diapause Development under
Constant Temperature and Photoperiod Conditions that Produce Mixed Broods in
Three Species of Butterfly from Three Different Subfamilies, namely Pieris napi
(Pieridae; Pierinae), Polygoniu c-album (Nymphalidae, Nymphalinae)
and Pararge aegeria (Nymphalidae; Satyrinae)

Development
Direct Diapause
Species Males Females Males Females P

P. napi 189 295 152 86 <0.001

P. c-album 141 155 91 43 <0.001
P. aegeria 64 95 56 34 <0.001

From Wiklund, C., Wickman, P.-0.. and Nylin, S. 1992. A sex difference in the propensity to enter directtdiapause
development: a result of selection for protandry. Evolution 46519-528. With permission.
194 Insect Reproduction

For animals with a short life expectancy, time becomes an essential currency that has a
fundamental influence on the life of both males and females. For instance, it is likely to
influence the opportunity for female mate choice, an issue that has been debated ever since
Darwin raised it in 1871. In butterflies, the degree of female hesitance to accept courting males
varies from species like Anthocharis cardamines, P. napi, and Coenonympha tullia, in which
some 90% of courtships involving virgin males result in mating to species like Coenonympha
pamphilus, where the corresponding figure is around 60% (Wiklund and F o r ~ b e r gForsberg
;~~
and W i k l ~ n dW; ~i~~ k m a n ~ Lack
. ~ ~ )of
. strong female mate choice can be rationalized in terms
of females being selected to minimize the time spent in unmated condition, conferring that
females pay a time cost for every male they reject. In accordance with the concept of life
expectancy constraining the female option to be selective in their choice of mates, W i c k ~ n a n ~ ~
recently showed that females of the relatively short-lived satyrid C. tullia fly up at, and solicit
courtship from, passing males, whereas females of the relatively more long-lived congeneric
C. pamphilus do not and often resist male mating attempts and wait to mate until they have
located a territorial male (which typically can be found close to specific landmarks). Hence,
the option for female mate choice is inversely proportional to female longevity (cf. J a n e t ~ s ; ~ ~
Thornhill and Alcock;' Hubbel and J ~ h n s o nReaP8).;~~
In systems where females are short-lived and their option for mate choice is limited, male
mating success is strongly dependent on their ability to locate receptive females. In species
where females mate only once, the timing of male appearance becomes an important factor
influencing the number of females that males have the opportunity to mate with. In accordance
with the flexible distribution models for protandry, it is easy to understand that the penalty on
males that emerge too late is more severe than that on males that emerge too early. This is
particularly highlighted in butterflies where males locate females before they emerge, and sit
(sometimes in clusters) and wait for the adult female to emerge from the pupa as in some
Heliconius species (Gilbert69)and the lycaenid Jalmenus evagoras (Elgar and Pierce70).
As a result of the benefit of protandry, males will often be severely pressed in a number
of different ways. For instance, since sexual selection seems to favor large relative male size
in polyandrous butterflies, these males are pressed to become larger than females in a shorter
time period, which can only be achieved by males increasing their growth rate which
presumably carries a cost. Moreover, in species where the adults emerge during the summer
but do not reproduce until after hibernation, as in Gonepteryx rhamni, females can use their
energy stores accumulated during summer and autumn for surviving the winter. Although
females are able to mate immediately upon becoming active in spring, they seem not to have
devoted energy to reproduction prior to mating, as evidenced by the fact that they ususally do
not start to lay eggs until about a week after mating (Wiklund and Lindfors, in preperation).
Conversely, males not only have to survive the winter, but their being able to mate directly
in spring requires that they have their ejaculatory ducts filled with sperm and accessory
substances at that time, which means that their energy reserves during the winter have been
devoted both to survival and reproduction.
In conclusion, many life history variables are related to male emergence, like variablity in
larval growth rate in relation to the acquisition of large size, producing sperm early during
hibernation so as to be ready to mate early in spring, and male reluctance to develop directly
in partially bivoltine species. This provides evidence that protandry has far-reaching effects
on insect life history and represents a truly fundamental aspect of mate acquisition and
reproductive success in insects.

ACKNOWLEDGMENT
I am grateful to Peter Abrams for comments on a previous version of this paper.
Protandry and Mate Acquisition

REFERENCES
1. Thornhill, R. and Alcock, J. 1983. The Evolution of Insect Mating Systems. Harvard University Press,
Cambridge, MA.
2. Clutton-Brock, T.H. and Hawey, P.H. 1977. Primate ecology and social organisation. J. Zool. 183:l-39.
3. Clutton-Brock, T.H., Harvey, P.H., and Rudder, B. 1977. Sexual dimorphism, socionornic sex ratio and
body weight in primates. Nature (London) 269:797-800.
4. Clutton-Brock, T.H., Guinness, F.E., and Albon, S.D. 1985. Parental investment and sex differences in
juvenile mortality in birds and mammals. Nature (Lond.) 313:131-133.
5. Shine, R. 1978. Sexual size dimorphism and male combat in snakes. Oecologia (Berlin) 33:269-277.
6. Shine, R. 1979. Sexual selection and sexual size dimorphism in the amphibia. Copeia 1979:297-306.
7. Berry, J.F. and Shine, R. 1980. Sexual size dimorphism and sexual selection in turtles. Oecologia (Berlin)
41:185-191.
8. Clutton-Brock, T.H. 1985. Selection in relation to sex. In: Bendall, D.S. (Ed.), Evolution from Molecules to
Men. Cambridge University Press, Cambridge, pp. 457-482.
9. Clutton-Brock, T.H. (Ed.) 1988. Reproductive Success: Studies of Individual Variation in Contrasting
Breeding Systems. University of Chicago Press, Chicago.
10. Partridge, L., Ewing, A, and Chandler, A. 1987a. Male size and mating success in Drosophila melanogaster:
the roles of male and female behaviour. Anim. Behav. 35:555-562.
11. Partridge, L., Hoffman, A., and Jones, J.S. 1987b. Male size and mating success in Drosophila melanogaster
and D. pseudoobscura under field conditions. Anim. Behuv. 35:468476.
12. Persson, L. 1985. Asymmetrical competition: are larger animals competitively superior? Am. Nat. 126:261-
266.
13. Bengtsson, J. 1988. Smaller zooplankton species are not superior in exploitative competition: a comment on
Persson. Am. Nat. 129:928-931.
14. Darwin, C. 1871. The Descent of Man and Selection in Relation to Sex. J . Mumy, London.
15. Grafen, A. 1987. Measuring Sexual Selection: Why bother? In: Bradbury, J.W. and Andersson, M.B. (Eds.)
Sexual selection: Testing the alternatives. John Wiley and Sons, Chichester, pp. 221-233.
16. Richards, O.W. 1927. Sexual selection and allied problems in the insects. Biol. Rev. 2:298-364.
17. Wedell, N. 1993. Protandry and mate assessment in the wartbiter Decticus verrucivorus. Behav. t c o l .
Sociobiol. 3 1:301-308.
18. Simmons, L.W., Llorens, T., Schinzig, M., Hosken, D., and Craig, M. 1994. Sperm competition selects for
male mate choice and protandry in the bushcricket, Requena verticalis. Anim. Behav. 47: 117-122.
19. Demoll, R. 1908. Die Bedeutung der Proterandrie bei Insekten. Zool. Jahrb. Abt. Syst. Oekol. Geogr. T i c e
26:621-628.
20. Stephen, W.P., Bohart, G.E., and Torchino, P.F. 1969. The Biology and External Morphology of Bees.
Oregon State University Press, Corvallis, OR.
21. Evans, H.E. and West-Eberhard, M.J. 1970. The Wasps. University of Michigan Press, Ann Arbor.
22. Petersen, W. 1892. ijber die Ungleichzeitigkeit in der Erscheinung der Geschlechter bei Schmetterlingen.
Zool. Jahrb. Abt. Syst. Oekol. Geogr. Tiere 6:671-679.
23. Petersen, B. 1947. Die geografische Variation einiger Fennoskandischer Lepidopteren. Zool. Bid. Uppsala
25:329-53 1.
24. Shapiro, A.M. 1970. The role of sexual behavior in density-related dispersal of pierid bunerflies. Am. Nat.
104:367-372.
25. Scott, J.A. 1986. The butteflies of North America. Stanford University Press, Stanford, CA.
26. Wiklund, C. and Fagerstrom, T. 1977. Why do males emerge before females? A hypothesis to explain the
incidence of protandry in bunerflies. Oecofogia (Berlin) 3 1: 153-158.
27. Nielsen, H.T. and Nielsen, E.T. 1953. Field observations on the habits of Aedes taeniorhynchus. Ecology
34:141-156.
28. Ford, E.B. 1945. Butteflies. Collins, London.
29. Lundgren, L. and Bergstrom, G. 1975. Wing scents and scent-released phases in the courtship behaviour
of Lycaeides argyrognomon. J. Chem. Ecol. 1:399-412.
30. Fagerstrom, T. and Wiklund, C. 1982. Why do males emerge before females? Protandry as a mating
strategy in male and female butterflies. Oecologia (Berlin) 52:164-166.
31. Bulmer, M.G. 1983. Models for the evolution of protandry in insects. Theor. Popul. Biol. 23:314-322.
32. Iwasa, Y., Odendaal, F.J., Murphy, D.D., Ehrlich, P.R., and Launer, A.E. 1983. Emergence patterns in
male butterflies: A hypothesis and a test. Theor. Popul. Biol. 23:363-379.
33. Parker, G.A. and Courtney, S.P. 1983. Seasonal incidence: adaptive variation in the timing of life history
stages. J. Theor. Biol. 105:147-155.
196 Insect Reproduction

34. Zonneveld, C. and Metz, J.A.J. 1991. Models on butterfly protandry: virgin females are at a risk to die.
Theor. Popul. Biol. 40:308-321.
35. Zonneveld, C. 1992. Polyandry and protandry in butterflies. Bull. Math. Biol. 54:957-976.
36. Baughman, J.F., Murphy, D.D., and Ehrlich, P.R. 1988. Emergence patterns in male checkerspot butter-
flies: testing theory in the field. Theor. Popul. Biol. 33:102-113.
37. Bulmer, M.G. 1980. The Mathematical Theory of Quantitative Genetics. Oxford University Press, London.
38. Baughman, J.F. 1991. Do protandrous males have increased mating success?The case of Euphydryas editha.
Am. Nut. 138:536-542.
39. Goddard, M.J. 1967. Broods of the speckled wood (Pararge aegeria aegerides Stgr.). The Entomologist
100:241-254.
40. Nylin, S., Wiklund, C., Wickman, P.-O., and Garcia-Barros, E. 1993. Absence of trade-offs in life-history
evo1ution:the case of sexual size dimorphism and protandry in Pararge aegeria. Ecology. 74: 1414-1427.
41. Lederhouse, R.C., Finke, M.D., and Scriber, J.M. 1982. The contributions of larval growth and pupal
duration to protandry in the black swallowtail butterfly, Papilio polyxenes. Oecologia (Berlin) 53:296-300.
42. Wiklund, C. and Forsberg, J. 1991. Sexual size dimorphism in relation to female polygamy and protandry
in butterflies: a comparative study of Swedish Pieridae and Satyridae. Oikos 60:373-38 1.
43. Wiklund, C. and Solbreck, C. 1982. Adaptive versus incidental explanations for the occurrence of protandry
in a butterfly, Leptidea sinapis. Evolution 36:56-62.
44. Singer, M.C. 1982. Sexual selection for small size in male butterflies. Am. Nat. 119:440443.
45. Sibly, R.M. and Calow, P. 1986. Physiological Ecology of Animals: An Evolutionary Approach. Blackwell,
Oxford.
46. Forsberg, J. and Wiklund, C. 1988. Protandry in the green-veined white butterfly, Pieris napi. Funct. Ecol.
2:8 1-88.
47. Rutowski, R.L. 1980. Courtship solicitation by females of the checkered white butterfly Pieris protodice.
Behav. Ecol. Sociobiol. 7:113-117.
48. Oberhauser, K. 1989. Effects of spermatophores on male and female monarch butterfly reproductive success.
Behav. Ecol. Sociobiol. 25:237-246.
49. Svard, L. and Wiklund, C. 1989. A comparative study on ejaculate mass in relation to female monan-
drylpolyandry. Behav. Ecol. Sociobiol. 24: 395402.
50. Wiklund, C., Nylin, S., and Forsberg, J. 1991. Sex-related variation in growth rate as a result of selection
for large size and protandry in a bivoltine butterfly, Pieris napi. Oikos 60:241-250.
51. Nylin, S., Wickman, P.-O., and Wiklund, C. 1989. Seasonal plasticity in growth and development in the
speckled wood butterfly, Pararge aegeria. Biol. J. Linn. Soc. 38:155-171.
52. Labine, P.A. 1966. The population biology of the butterfly Euphydryas editha. VIII. Oviposition and its
relation to patterns of oviposition in other butterflies. Evolution 22:799-805.
53. Karlsson, B. and Wiklund, C. 1984. Egg size variation in satyrid butterflies: adaptive versus historical.
"Bauplan", and mechanistic explanations. Oikos 43:391-400.
54. Karlsson, B. and Wiklund, C. 1985. Egg weight variation in relation to egg mortality and starvation
endurance of newly hatched larvae in some satyrid butterflies. Ecol. Entomol. 10:205-211.
55. Svard, L. and Wiklund, C. 1988. Fecundity, egg weight and longevity in relation to multiple matings in
females of the monarch butterfly. Behav. Ecol. Sociobiol. 23:3943.
56. Svard, L. and Wiklund, C. 1991. The effect of ejaculate mass on female reproductive output in the European
swallowtail butterfly, Papilio machaon. J. Insect. Behav.
57. Danilevskii, A.S. 1965. Photoperiodism and Seasonal Development of Insects. Oliver and Boyd, Edinburgh.
58. Tauber, MJ., Tauber, C.A., and Masaki, S. 1986. Seasonal Adaptations of Insects. Oxford University
Press, Oxford.
59. Danks, H.V. 1987. Insect Dormancy: An Ecological Perspective. Biological Survey of Canada, Ottawa.
60. Wiklund, C., Wickman, P.-O., and Nylin, S. 1992. A sex difference in the propensity to enter direct/
diapause development: a result of selection for protandry. Evolution 46:519-528.
61. Vogel, S. 1989. Life's devices: The Physical World of Animals and Plants. Princeton University Press,
Princeton, NJ.
62. Wiklund, C. and Forsberg, J. 1985. Courtship and male discrimination between virgin and mated females
in the orange tip butterfly, Anthocharis cardamines. Anim. Behav. 34:328-332.
63. Forsberg, J. and Wiklund, C. 1989. Mating in the afternoon: time-saving in courtship and remating by
females of a polyandrous butterfly, Pieris napi. Behav. Ecol. Sociobiol. 25:349-356.
64. Wickman, P.-0.1986. Courtship solicitation by females of the small heath butterfly, Coenonympha pamphilus,
and their behaviour in relation to male territories before and after copulation. Anim. Behav. 34:153-157.
65. Wickman, P.-0. 1992. Mating systems of Coenonympha butterflies in relation to longevity. Anim. Behav.
44:141-148.
66. Janetos, A.C. 1980. Strategies of female mate choice: a theoretical analysis. Behav. Ecol. Sociobiol. 7: 107-
112.
Protandry and Mate Acquisition 197

67. Hubbel, S.P. and Johnson, L.K. 1987. Environmental variance in lifetime mating success, mate choice, and
sexual selection. Am. Nut. 130:91-112.
68. Real, L. 1990. Search theory and mate choice. I. Models of single sex discrimination.Am. Nut. 136:376-404.
69. Gilbert, L.E. 1975. Ecological consequences of a coevolved mutualism between butterflies and plants. In:
Gilbert, L.E. and Raven, P.R. (Eds.) Coevolurion of Animals and Plants. Texas University Press, Austin, pp.
2 10-240.
70. Elgar, M.A. and Pierce, N.E. 1988. Mating success and fecundity in an ant-tended lycaenid butterfly. In:
Clutton-Brock, T.H.(Ed.) Reproductive Success. University of Chicago Press, Chicago, pp. 59-75.
 Chapter 9

SWARM-BASED MATING SYSTEMS

Athol McLachlan and Rachel Neems

And thick and fast they came at last and more and more and more
Lewis Carroll. Alice's Adventures in Wonderland

CONTENTS
I. Introduction ...............................................................................................................
199

11. How Are Swarms Formed? ...................................................................................... 201

111. Why Do Insects Swarm? .......................................................................................... 201

IV. Body Size And Mating Success In Swarming Insects ............................................. 203
A. Body Size ............................................................................................................ 203
B. Alternative Mating Options ................................................................................ 206
C. The Importance of Swarm Size ..........................................................................207
D. The Role of the Larval Habitat in Determining Body Size in the Adult .......... 2 11

V. Summary ....................................................................................................................
21 1

Acknowledgments ............................................................................................................... 2 12

References ...........................................................................................................................2 12

I. INTRODUCTION
Swarms of flying insects, sometimes numbering many millions of individuals (Figure l),
are a common sight. They have been recorded in the following taxa at least: beetles, termites,
dragonflies, bees, ants, caddis flies, stone flies, mayflies, and among the Diptera in the
Trichoceridae, Empididae, Culicidae, Ceratopogonidae, Simulidae, Chaoboridae, Bibionidae,
and Chironomidae. In many of these taxa, such assemblages are composed entirely of male
insects, and the obvious question is what function, if any, they serve. Good evidence exists that
swarms composed of males are mating assemblages. Typically, these are aggregations of
males displaying to attract females which enter the swarm to acquire a mate (reviewed by
Thornhill and Alcockl). Such displays are necessary properties of a lek,2 a type of mating
system more familiar among birds and mammals. Leks are peculiar mating systems and,
despite being widespread, they do not constitute a common way for animals to acquire mates
(reviewed by Krebs and Davies3).They are peculiar because males seem to have nothing to
offer the female but their sperm. Males do not defend a resource sought by females as is the
common alternative method of acquiring mates (see Parker"). In insect leks, there is the added
peculiarity that there is little evidence of females exercising any choice of mate within the lek.
Instead, the emphasis shifts to competition between males. In a spectrum of l e k ~ from ,~

0-8493-6695-W95/S0.oOcS.50
0 1995 by CRC Press, Inc. 199
 Insect Reproduction

FIGURE 1. A swarm of lake flies (Chaoboridae) on Lake George (Uganda). (Photograph by L. MacGowan).

extreme emphasis on female choice as seen in some bird species to extremes of malelmale
competition, swarming insects considered here occupy a position near the latter end of the
spectrum.
In a recent review, Bradbury and Davies? express the opinion that in insects ". .. the heavier
emphasis on male-male conflict when compared to avian leks cries out for detailed study".
Over the last 10 years, there have been several studies on insect swarms as arenas for sexual
selection. Leks of the lovebug (Bibionidae), mayflies (Ephemeroptera), and the nonbiting
midges (Chironomidae) have received attention. The mayflies and chironomids are of particu-
lar interest because adults do not feed, or at least are not obligate feeders (Burtt et al.'). Adults
are, therefore, entirely concerned with reproduction, which should make them easier subjects
for studies of sexual selection. Here, we consider leks in these taxa and particularly chirono-
mid leks. Because females do not seem to exercise choice in the matter of partner, we leave
aside the question of epigamic selection and consider only intrasexual selection between
competing males. This decision diverts the emphasis away from the evolution of male
adornment, which is the focus of work on lekking vertebrates. What has become known as the
"paradox of the leY8stems from this emphasis. The paradox exists because it is not clear why
females should show such a strong preference for males offering little but their sperm. This
is a question which does not apply to the insects considered here. The absence of elaborate
ornamentation in males compared with females among swarming insects is evidence for the
shift away from female choice.
We give prominence to body size as a factor determining mating success in swarms of
insects. The emphasis on size has impeccable antecedents (see Darwin? Wallace,Io and
Simpson"). There is no sign of size-assortative mating in the taxa we consider (Bibionidae,
Swarm-Based Mating Systems 201

Ephemeroptera, and Chironomidae), so sexual selection on male body size is not complicated
by considerations of female size and fecundity. This is another reason for leaving females
aside in this Chapter. We consider swarming in a variety of insects, but focus progressively on
the Diptera and finally on the nonbiting midge Chironomus plumosus, as a model swarming
insect. We intend that much of what is said about C. plumosus be extrapolated, not only to
other chironomid species, but also to male mating swarms of insects in general. Time will tell
whether these generalizations are incautious.

11. HOW ARE SWARMS FORMED?

There is sufficient evidence about sensory physiology in the Diptera at least for a reason-
able understanding of how swarms are formed and how they attract females. Swarming
evidently depends upon a visual search by males for landmarks followed by a visual station
keeping over the landmark. A landmark is simply any visual contrast in the environment. A
bush, tree, track, or pond serve equally well. The elaborate antennae of the male, seen in many
families of Diptera, are sound detectors that respond to the wing beat sounds of the female and
come into play only after the female has entered a swarm. There is neither evidence that the
male antenna responds to the higher frequency sounds made by flying males to aid the initial
formation of a swarm, nor is there any evidence that the less elaborate antenna of the female
is capable of detecting sound at all. She seems to find the swarm entirely on visual cues and
is captured there by a male using a sound cue. The different mating behaviors of the sexes are
clearly reflected in sexual dimorphism of their sensory equipment (Figure 2). The observa-
tions reported here have been reviewed by Downes,12but they have a much earlier origin. In
1901, Sir Hiram Maxim, better known for his invention of the machinegun, conducted an
elegant series of experiments published in the Times (London) newspaper. He clearly demon-
strated that the female flight sound was attractive to the male mosquito (reported by Roth et.
aI.l3). If the male antenna is necessary to detect the female, it follows that the females lacking
such equipment may not be able to detect an individual male at all. This is one reason to
suppose that female choice is not a prominent aspect of mating in swarming Diptera.

111. WHY DO INSECTS SWARM?

In this section, we consider questions about how these swarm-based leks have evolved.
Such questions hinge on understanding what the individual male stands to gain in terms of
mating success, by entering a lek as compared to remaining on his own. There are two main
families of hypotheses to explain why it is advantageous for males to lek. These hypotheses
center around the "hotspot" and the "hotshot" models (reviewed by Krebs and Davies3).
"Hotspots" are places where males aggregate because females are likely to be common there.
Such places include female emergence sites, as seen in the lovebug, or where female territories
overlap as in many bird species. The "hotshot" model concerns the case, again observed
among birds, where males aggregate around an especially successful competitor and improve
their chances of mating by association with him. Chironomid swarms, and perhaps swarming
Ephemeroptera as well (and possibly many others), present a difficulty because, at first sight,
neither of these models seems to be applicable. We propose the following testable hypothesis.
Females are unable to detect a single male, but are able to see a dancing swarm of males. This
is basically the old idea of "stimulus pooling," which Bradbury2has criticized on the grounds
that joining an aggregation is unlikely to increase the individual males chances of mating.
However, we propose that there is a critical minimum swarm size below which the males
chances of attracting a mate are zero. There may also be a critical maximum swarm size above
which the costs of intrasexual competition become excessive. If we are correct, the idea of
stimulus pooling may be applicable to insect swarms. The test requires the determination of
 Insect Reproduction

FIGURE 2. Some examples of sexual dimorphism sensory equipment. Part A is a chironomid, Chironomus bellus
(a) front view of the male head; (b) antenna of male; (c) and (d) the same for the female (Modified from Freeman,
P. Bull. Br. Mus. Nar. Hist.. 4, 1 , 1955. With permission.) Part B is a mosquito (Aedes) showing head of male and
female. Part C depicts male and female of a bibionid fly, Bibio albipennis (Figures B and C modified from An
Inrroducrion to the Study of Insects, copyright 1954 and renewed 1982 by Donald J. Borror and Dwight M. Delong,
reproduced by permission of Saunders College Publishing.)

the range of swarm sizes in a species and the relationship between swarm size and mating
success. Some of these matters are taken up in the following section.
There is a second facet to the evolution of swarm-based mating in insects. This concerns
the landmark over which swarms form. Landmarks seem central to the formation of such
swarms. Indeed, Downes12 describes experiments on mosquitos where swarm size was ma-
nipulated by altering the number of landmarks available. It is widely observed that a swarm
will disappear if the landmark is hidden by the observer. So, an understanding of the evolution
of such swarms requires information of the origin and use of landmarks. It is intuitively
reasonable to suppose that the landmark has evolved from the pond as a lekking site. This idea
is feasible both because females emerge from the pond and because the pond represents a
resource to the females as an oviposition site. Furthermore, males of most species emerge
before females (protandry), so swarming over the home pond to await the arrival of mates
makes sense (for review of protandry, see Wiklund and FagerstomI4).The landmark, and not
Swarm-Based Mating Systems 203

the female, thus, becomes the focus of male lekking. The landmark can be seen as symbolic
of an ancestral resource. This is the view taken by Thornhill and Alcock.' Downed2 has coined
the phrase "landmark swarming" for the type of insect lek we are interested in. We suggest
that the term be retained but broadened to include the idea of the lek. What Bradbury5 calls
arbitrary lekking sites may be landmark swarming as we define it here. He suggests that
female choice of such sites may have evolved in precisely the same way as for the preference
of other male traits, that is, through a closed feedback loop between female choice and male
preference. This argument seems not entirely satisfactory because females have become the
searchers in lekking species. This point cannot easily be ignored because it raises the difficulty
that, in cases of anisogamy, it ought to be the male that has surplus resources to put to
searching behaviour (reviewed by Parker"). So, there are a number of unresolved problems in
understanding the evolution of landmark swanning and of lekking behavior in general.

IV. BODY SIZE AND MATING SUCCESS IN SWARMING INSECTS

The males of most species of animal spend their adult lives fighting between themselves
for mates. Since large size carries advantages in such contests, size lies at the center of studies
on intrasexual selection in males. In this section, we take up the question of body size in
swarming insects and show how new evidence puts a different light on topics such as the
evolution of landmark swarming, the role of landmarks and of social interactions in swarm
formation. Much of the evidence presented comes from studies of swarming chironomids.
Most species produce a cohesive, dancing swarm very amenable to investigation.

A. BODY SIZE
The mating success and fitness of males of all swarming insects thus far studied is found
to be strongly influenced by size. In some cases, as in the lovebug Plecia nearctica15and the
mayfly Epeorus longimanus,16 it is the larger males that acquire a disproportionate number of
matings. In others, for example among many species of chironomid,17the smaller than average
male is most successful (Figure 3A and B). Size selective pressures on the male can lead to
the sexual size dimorphism so widespread among animals (for review see Adams and
GreenwoodI8). Specific differences in what constitutes the successful male are presumably
due to differences in the method adopted by males to capture females. Details of mate capture
and malelmale competition appear to be unknown in the lovebugs and mayflies. For the
chironomids, we have some idea of what happens to account for the success of small males.
It is well known that small flying animals, like small aircraft, have a better turning moment
and greater acceleration than larger ones. An increased aerobatic ability in smaller male
insects may therefore help in capturing mates on the wing.lg A direct test of this idea would
be the measurement of the turning moment and acceleration of males in a swarm. Okubo et
achieved direct measurement using cinefilm of swarms of the cecidomyiid midge Anarete
pritchardii and found that some individuals had far greater acceleration than others. Unfor-
tunately, they did not relate this finding to body size. An indirect way of relating together size
and mating success is to weigh the wing muscles and regress this measure against wing length
as a correlate of size. The expected allometric relationship for weight against length (see
Calder2') is

where
M = mass
X = length
a = constant
 Insect Reproduction

N.54 Swarm
10.

M a t i n g pairs

Wing l e n g t h (mm)

FIGURE 3A.

L-
(%l
&&&-
&
Frequency

r Random

Mating

Body length (mm)

FIGURE 3B.

FIGURES 3A and 3B. Size distribution of males in the unmated male swarm compared with males caught paired
with females in (A) the midge Chironomusplumosus (Modified from Neems, R.M., McLachlan, A.J., and Chambers,
R. Anim. Behav. 40,648, 1990. With permission.) and (B) the mayfly Epeorus longimanus (Modified from Flecker,
A.S., Allan, J.D., and McClintock, N.L.Holarctic Ecol. l l , 280, 1988. With permission.)

This is the reasoning adopted by Conveyz2and by Marden23for dragonflies. The relation-

ship between wing length and flight muscle mass in C. plumosus is unexpected in that it is a
straight line (y = 0 . 2 3 ~- 0.52); that is
Swarm-Based Mating Systems

FIGURE 4. The relationship between wing length and dry weight of flight muscles in male Chironomusplumosus.
Each point represents a single male, with n being the total number of flies measured. (Modified from Neems, R.M.,
Ph.D. thesis, University of Newcastle upon Tyne, U.K., 1990.)

and not M = ax3 (Figure 4). This means that large male C. plumosus do not have the expected
increase in muscle mass predicted from the cubic formula. The reciprocal conclusion is that
small males are invested with a disproportionate amount of flight muscle. Hence, they may
gain in aerobatic performance, not only through being smaller per se, but also in having a
proportionately larger flight machinery.
In other words, in cases of size specific mating success it is not the most common size that
does best. This is a puzzle, because selection ought to result in males that are most successful
also being the most common. Among mammals, the explanation may be that few males ever
reach large size, so the advantages to possessors of these genes are probably seldom realized
(see Parker24).Among damsel flies, small males have a better short-term mating success but
a short life span so their lifetime reproductive success (LRS) is reduced in favour of larger
corn petit or^.^^ In the case of short-lived insects, longevity is not such an important component
of fitness and in chironomids, the LRS of small males is better than that of larger ones. We
use LRS as an approximation to fitness (W) so, for chironomids small males are fitter than
large. This conclusion takes into account four components of fitness: mating success, stamina,
fertility, and longevity (Neems et. Figure 5). The explanation as to why the most fit size
of male is not also the most common may lie in a different part of the life cycle. The major
 Insect Reproduction

I
Size

FIGURE S. Lifetime reproductive success (W) as a function of body size in Chironomusplumosus (Modified from
Neems, R.M., Ph.D. thesis, University of Newcastle upon Tyne, U.K., 1990.)

part of a chironomids life is spent as a larva and among larvae competing for food and space,
large size may be favored. The resulting selection for large size in larvae might account for
the discrepancy between the fittest size and the commonest size among adults. We have yet
to investigate this possibility.

B. ALTERNATIVE MATING OPTIONS

What has been said about body size and fitness is only part of the story. Among chirono-
mids, males are able to adopt a variety of alternative mating strategies depending upon body
size and species. These alternative mating behaviors give evidence of social interactions in the
swarm and de-emphasise Downes' view of the landmark as the sole organizing force in cases
of landmark swarming (Downes12).The most extreme examples are seen in such species as
Bryophenocladius vernalis and Procladius crassinervis. As in most chironomid species,
mating involves females entering a swarm singly to emerge, almost immediately, paired with
a male. Pairs fall out of the swarm and can be captured when they alight on the ground. In these
species, the smaller males in the population are poor at sustaining flight and spend much of
their time on the ground near a swarm made up of larger males. Some of the females attracted
to the swarm evidently settle on the ground because these small males are not found in samples
of the swarm but appear in samples of malelfemale pairs caught on the ground (Figure 6 ) .
These are small species. In bigger ones, smaller males do not lose their ability to fly, indeed
they may be more aerobatic than larger individuals. An example of alternative mating
strategies is seen in Chironomus plumosus, where males assort themselves into different
swarms depending upon body size. Small males can form satellite swarms around a swarm
composed of larger males keeping station over a landmark.27Males in these satellite swarms
appear tq intercept females on their way to the main swarm. As in the case of the small males
that do not swarm, the existence of satellite swarms opens the possibility of phenotype-
determined alternative mating strategies with small individuals avoiding competition without
jeopardizing their chances of mating. We must now ask why any of the males undertake the
energetically costly flight at all. Why do they all not alight on the ground and await the arrival
of a female as do small males of B. vernalis and P. crassinervis? The answer is not difficult
Swam-Based Mating Systems

Wing length class

FIGURE 6. The distribition of wing size among males of the midge Bryophenocladius vernalis (a) in mating pairs
and (b) in the male swarm. (Modified from McLachlan, A.J., Ecol. Entomol. 11, 237, 1986. With permission.)

to guess. If too many males join satellite swarms or alight on the ground, there is no signal
to attract females. An Evolutionarily Stable Strategy (ESS) occurs where the payoff to each
option is equal. In other words, we are seeing a frequency-dependent alternative option
strategy.28
We have said enough to show that body size is at the center of mating performance in
lekking male insects. Since it seems so important, it is desirable to quantify the intensity of
sexual selection operating on body size. This has been attempted for d r a g ~ n f l i e sand
~ ~ more
. ~ ~have attempted to measure selection intensity in C. plumosus
recently for D r ~ s o p h i l aWe
using the same methods. Calculations depend upon the assumption that variance in mating
success for males of different sizes is directly related to the intensity of selection. In principle,
how much mated males differ in size from those in the male population as a whole is
calculated. An example of the procedure for C. plumosus is shown in Table 1. The outcome
(Table 2) shows that sexual selection does appear to significantly affect body size in this species.

C. THE IMPORTANCE OF SWARM SIZE

We have already shown that an individual's body size is important in determining his
success in the swarm and, as with any other group of animals, what everybody else is doing
has consequences for the individual as well. We now turn to the question of size of group, or
the swarm in this case, on an individual's reproductive success. All the male insects of a
particular species in an area do not normally aggregate into a single large swarm, but instead
form many discrete aggregations of different size. In an attempt to explain why a range of
swarms exist, we model the costs and benefits to the individual of joining swarms of various
sizes. On the one hand, the larger the swarm the better the chances of avoiding detection by
a predator. This is because, as in Hamilton's "selfish herd, an individual can more effectively
hide in a crowd.3' On the other hand, because of low levels of intrasexual competition, the
smaller the swarm the better a males' chances of acquiring a mate. There ought, therefore, to
be a balance between the payoff to small vs. large swarm size. We proceed with caution
because swarm size may not be stable in the wild. One reason for this, suggested by S i b l ~ , ~ ~
is that group size may become larger than optimal due to excess individuals attempting to join
a successful group.
208 Insect Reproduction

TABLE 1
An Example of the Procedure used for Estimating
the Intensity of Sexual Selection of
Male Body Size in Chironomus plumosus

Mean male wing

length (mm) Paired status Class frequency

4.2711 1 0.03
4.3220 0 0.97

Note: z is mean size; P, variance around z; W, mean fitness; s, selection

differential; i, selection intensity in standard deviation units. Hence,
z = 4.3205. P = 0.0148, W = 0.03 and z, = 4.2711; s, is z,, -z and
i are s,/P.

From Neems, R.M., Ph.D. thesis, University of Newcastle upon Tyne,

U.K., 1990.

TABLE 2
A Summary of the Parameters Governing the Intensity
of Sexual Selection, z is the Mean Size, P is the Variance
Around z; N, Total Sample Size; z,, Mean Size After
Selection; and i, Selection Intensity

Sample z P N z, i

1 4.3205 0.0148 95 4.2711 -0.4081

2 4.301 1 0.0175 68 4.1577 -1.0840
3 4.3044 0.0144 122 4.3394 -0.2917
4 4.0072 0.0187 126 3.9090 -0.7181

To determine whether an optimal swarm size can be demonstrated in the case of chirono-
mids, we gathered data on the rate of predation by Emphis.33 These flies are active predators
of the C. plumosus swarms we studied in Northumberland. We determined predation rates by
counting the number of Emphis males leaving the swarm with a chironomid over a 40 minute
period. These records were used to calculate a figure for the risk of predation per individual
chironomid male (Ppred), obtained by dividing the number of attacks by the number of males
in a swarm. At the same time, mating pairs leaving the swarm were counted to obtain a figure
for the probability of mating per male (Pmale). Again, the number of pairs was divided by the
number of males in the swarm to generate a figure for the probability of mating.
Combining these data, we arrive at a predicted, combined fitness payoff (W) to individual
males to joining swarms of different sizes (Figure 7 and Box 1). The smallest swarms give the
highest pay-off in fitness. A swarm size of about 100-500 males is optimal while those of
around 4000 individuals give the minimum pay-off. The smallest swarms are not the common-
est swarm size observed in the wild which is about ten times larger (1000-2000 males).
This outcome seems to justify Sibly's concern, but thus far we have left considerations of
body size aside. As there is a strong covariance between body size and swarm size (Figure 8),
this variable must be incorporated into the model. As it stands, the optimal swarm size is
calculated from the combined payoff to the average male. We know that males differ widely
in competitive ability, and the size of the swarm that is optimal for a small male may not be
optimal for a large male. In this way, a range of swarm sizes should be expected rather than
just one optimal swarm size (Figure 9). We may, in fact, be dealing with an Ideal Free
Swarm-Based Mating Systems

2000 4000 6000

Swarm size ( N I

FIGURE 7. The predicted combined payoff (W) to individual males in joining swarms of different sizes.

Swarm size

FIGURE 8. The relationship between log swarm size and body size in male Chironomus plumosus. Each point
represents the average body size in a different swarm. (Modified from Neems, R.M., Ph.D. thesis, University of
Newcastle upon Tyne, U.K., 1990.)
 Insect Reproduction

FIGURE 9. The payoff (W) to large males (LM) and small males (SM) in joining swarms of various sizes . The
optimal size of swarm for each size of male is indicated by an arrow.

Distribution of unequal competitor^,^^ where the best competitors (small males) occupy the
best patches (small swarms).
For variance in male body size to be evolutionarily stable, the payoff to males of all sizes
must be equal. We started this chapter by showing that the payoff is not equal to all males but
that small size is favored. This is an unresolved dilemma. Recall that we believe the discrep-
ancy in fitness between the commonest and the smallest males may be corrected in the larval
stage so that over the life cycle as a whole, the two strategies can persist despite an unequal
payoff in the adult part of the life cycle. This is not yet tested, but something more can be said
here about the immature stages of the life cycle.

Box 1

The number of females attracted to a swarm increases with swarm size (r = 0.93, P <0.001).
However, since the attraction rate does not increase in proportion to swarm size, the individual
males' probability of mating (P,,,) decreases with swarm size (but increases again slightly
above a size of 4000). The curvilinear nature of this relationship is described by the regression
equation

Predatory attack rate also increased with swarm size, but again the probability of predation
to the individual(P,,) declines significantly as swarm size increases due to a dilution effect.
This relationship is described by the linear equation

P,, = 0.0263 - 0.3 X 10-5N (4)

Equations (3) and (4) can be combined to provide a measure of "mating success" per
evening (W) in a common currency that takes into account the fact that the individual might
be captured by a predator while in the swarm

W = P,", X p,, (5)

where

P,,, = l - Pp=d
Swarm-Based Mating Systems 211

As Figure 7 in the main text shows, W declines as swarm size increases up to a value of
about 4000 males but increases thereafter. In spite of the greater safety of larger swarms,
mating success is greatest in the smallest swarms. These relationships are summarized in the
figure below.

swarm size

BOX 1 FIGURE. The calculation of optimal swarm size in C. plumosus.

D. THE ROLE OF THE LARVAL HABITAT IN

DETERMINING BODY SIZE IN THE ADULT
Our conjecture is that under conditions of intense crowding, competition for resources is
asymmetric, with larger larvae doing better than smaller ones. If size has a heritable compo-
nent, which it almost certainly does, and if large larvae always give rise to large adults and
small larvae to small adults, then size determined in the larval habitat may have an important
bearing on size in the adult. It is often observed in insects that smaller adults emerge from
larvae reared under crowded condition^.^^ The immature stages of many insects experience
very variable levels of crowding because the habitat is a temporary one and crowding is
correlated with progress of the habitat towards extinction. Examples are the conditions
experienced by larvae of Drosophila in a rotting apple, Calliphora in carrion, and larvae of
the genus Chironomus in rainpools. In some Chironomus species, as in some toads (see
Duellman and T r ~ e and
b ~McLachlan3'
~ for reviews), the story is more complex because giant
adults as well as dwarfs are produced when crowding is extreme. We do not yet have data on
the relationship between size of individual larvae and adults in the chironomids, nor do we
have data on differential survival by size among larvae under different levels of competition.
However, it is an obvious thing to investigate and seems accessible to study.

V. SUMMARY
The majority of insect swarms are aggregations of flying male animals, maintaining station
over a landmark of no value to the female. We consider such swarms to be leks, but they differ
from the classical lek seen in vertebrates in at least two important respects. First, there is little
evidence of female choice and, as a consequence, the focus of interest moves from the
evolution of male ornamentation to questions of competition between males. Second, there are
peculiarities about the evolution of swarming in insects which do not easily fit any of the
existing models depending upon the distribution of females. Swarming may have evolved
from a system originally dependent on female distribution. The occurrence of the lek depends
upon male choice of a suitable swarming site or landmark where males aggregate to create a
signal to attract dispersed females. The landmark may represent the home pool or stream as
a site where emerging females were likely to be encountered. We propose that the term
"landmark swarming" be retained for such insect leks.
212 Insect Reproduction

For our test species C. plumosus, we have the data to develop models for the prediction of
optimal swarm size. This turns out to depend upon a compromise between costs of predation
and benefits of mate acquisition. We also consider the mechanisms of mate acquisition within
the swarm. Both swarm size and mating success hinge on body size in the male sex. The larva
is the stage of the life cycle where the environmental component to size in the adult is entirely
determined. We identify selection on size in the larval stage, the evolution of the insect lek,
and the mechanisms of malelmale competition as areas for further work.

ACKNOWLEDGMENTS
We owe many thanks to John Lazarus for help in developing the mathematical models
considered here and for writing the computer programs used in the modeling section. We
thank Nicholas Davies for his helpful comments on earlier versions of this chapter.

REFERENCES
1. Thornhill, R. and Alcock, J., The Evolution of Insect Mating Systems, Harvard University Press, London,
1983.
2. Bradhury, J.W., The evolution of leks, in Natural Selection and Social Behaviour, Alexander, R.D. and
Tinkle, D.W., Eds., Blackwell Scientific Publications, Oxford, 1981, 12.
3. Krebs, J.R. and Davies, N.B., Behavioural Ecology, An evolutionary approach, Third ed., Blackwell
Scientific Publications, Oxford, 1991.
4. Parker, G.A., Evolution of competitive mate searching, Annu. Rev. Entomol., 23, 173, 1978.
5. Bradbury, J.W. Contrasts between insects and vertebrates in the evolution of male display, female choice,
and lek mating, in Experimental Behavioural Evolution, Ecology and Sociobiology, Holldobler, G. and
Lindauer, M., Eds., Fortschr. Zool. Gustav Fischer Verlag, Stuttgart, 31, 273, 1985.
6. Bradbury, J.W. and Davies, N.B., Relative roles of intn- and intersexual selection, in Sexual Selection:
Testing the Alternatives, Bradbury, J.W. and Andersson, M.B., Eds., Wiley, New York, 1987, 143.
7. Burtt, E.T., Perry, R.J., and Mclachlan, AJ., Feeding and sexual dimorphism in adult midges, Holarctic
Ecol., 9, 27, 1986.
8. Kirkpatrick, M. and Ryan, MJ., The evolution of mating preferences and the paradox of the lek. Nature,
350, 33, 1991.
9. Darwin, C., The Descent of Man and Selection in Relation to Sex, J . Murray, London, 1871.
10. Wallace, A.R., Darwinism: An Exposition of the Theory of Natural Selection with Some of its Applications,
Macmillan, London, 1889.
11. Simpson, G.G., The Major Features of Evolution. Columbia University Press. New York, 1953.
12. Downes, J.A., The swarming and mating flight of Dipten, Annu. Rev. Entomol., 14, 271, 1969.
13. Roth, M., Roth, L.M., and Eisner, T.E., The allure of the female mosquito, Nut. Hist., 75, 27, 1966.
14. Wiklund, C., and Fagerstom, T., Why do males emerge before females? A hypothesis to explain the
incidence of protandry in butterflies, Oecologia, 31, 153, 1977.
15. Thornhill, R., Sexual selection within mating swarms of the lovebug Plecia nearctica (Dipten: Bibionidae),
Anim. Behav.. 28,405, 1980.
16. Flecker, A.S., Allan, J.D., and McClintock, N.L., Male body size and mating success in swarms of the
mayfly Epeorus longimanus,Holarctic Ecol., 11, 280, 1988.
17. Mclachlan, AJ. and Allen, D.F., Male mating success in Diptera: advantages of small size, Oikos, 48, 11,
1987.
18. Adams, J. and Greenwood, PJ., Why are males bigger than females in pre-copula pairs of Gammarus
pulex?, Behav. Ecol. Sociobiol., 13, 239, 1983.
19. Mclachlan, AJ., Survival of the smallest: advantages and costs of small size in flying animals, Ecol.
Entomol., l l, 237, 1986.
20. Okubo, J., Chiang, J.C., and Ebbesmeyer, C.C., Acceleration field of individual midges, Anarete pritchardi
(Diptera: Cecidomyiidae), within a swarm, Can. Entomol., 109, 149, 1977.
2 1. Calder, W.A., Size, Function and Life History, Harvard University Press, London, 1984.
Swam-Based Mating Systems 213

22. Convey, P., lnfluences on the choice between territorial and satellite behaviour in male Libellula quadrimaculata
Linn. (Odonata: Libellulidae), Behaviour, 109, 125, 1989.
23. Marden, J.H., Bodybuilding dragon flies; costs and benefits of maximizing flight muscle, Plzysiol. Zool., 62,
505, 1991.
24. Parker, G.A., Arms race in evolution: an ESS to the opponent- independent costs game, J. Theor. Biol., 101,
619, 1983.
25. Banks, M J . and Thompson, DJ., Lifetime mating success in the damselfly Coenagrion puella, Anim.
Behav., 22, 1175, 1985.
26. Neems, R.M., Lazarus, J., and Mclachlan, AJ., Lifetime reproductive success in a swarming midge:
stabilising selection for male body size. Nature. 1995 (submitted).
27. Neems, R.M., The Role of Body Size in the Mating Systems of Midges (Chironomidae), Ph.D. thesis,
University of Newcastle upon Tyne, U.K., 1990.
28. Maynard Smith, J., Evolution and the Theory of Games, Cambridge University Press, Cambridge, 1982.
29. Arnold, SJ. and Wade, M.J., On the measure of natural and sexual selection: applications, Evolution, 38,
709, 1984.
30. Wilkinson, G.S., Equilibrium analysis of sexual selection in Drosophila melanogaster, Evolution, 41, 11,
1987.
31. Hamilton, W.D., Geometry for the selfish herd, J. Theor. Biol., 31, 295, 1971.
32. Sibly, R.M., Optimal group size is unstable, Anim. Behav., 31, 947, 1983.
33. Neems, R.M., Lazarus, J., and McLachlan, AJ., Swarming behavior in male chironomid midges: a cost
- benefit analysis, Behav. Ecol.. 3, 285, 1992.
34. Fretwell, S.D., Populations in a Seasonal Environment. Princeton University Press, Princeton, NJ.,1972.
35. Nicholson, A.J., The self-adjustment of populations to change, in Population Studies: Animal Ecology and
Demography, Warren, K.B., Ed., Cold Spring Harbor Symposium in Quantitative Biology, 22, 1958, 153.
36. Duellman, W.E. and Trueb, L., Biology of Amphibians. McGraw-Hill, New York, 1986.
37. McLachlan, A.J., Animal populations at extreme densities: size dimorphism by frequency dependent selec-
tion in ephemeral habitats,Funct. Ecol., 3, 633, 1989.
 Chapter 10

MALE NUPTIAL GIFTS: PHENOTYPIC CONSEQUENCES

AND EVOLUTIONARY IMPLICATIONS
.
Carol L Boggs

CONTENTS
I . Description Of Male Nuptial Gifts ........................................................................... 216
A. Definition ............................................................................................................
216
B. Prey Items and Regurgitants............................................................................... 216
C. Accessory Gland Products .................................................................................. 217
1. Nonspermatophore Secretions ....................................................................... 217
2. Spermatophores .............................................................................................. 217
D. Body Parts and Hemolymph ............................................................................... 219
E. Sperm ..................................................................................................................
219

I1. Overview of Adaptive Functions of Male Nuptial Gifts ........................................

219

I11. Physiological Costs and Benefits of Male Nuptial Gifts:

Effects on Resource and Time Budgets ...................................................................220
A. Female Perspective .............................................................................................220
B. Male Perspective .................................................................................................
222

IV . Effects of Nuptial Gifts on Demographic Fitness Components .............................. 223

A. Age-Specific Fecundity Patterns ..................................................................223
B. Survival Patterns ................................................................................................. 224

V . Effects of Nuptial Gifts on Population Structure ..................................................... 226

A . Age Structure ...................................................................................................... 226
B. Dispersal .............................................................................................................. 226
C. Effective Population Size ...................................................................................227

V1. Patterns of the Evolution of Nuptial Gifts ...............................................................228

A. Origins: Phylogenetic History ............................................................................ 228
B. Current Variation and Future Prospects: Heritability ........................................ 229
C . Interaction of Nuptial Gifts with Mate Competition and Mate Choice ............ 229
1. Sex Roles and Sexual Selection: Theory ...................................................... 230
2. Experimental Evidence: Nuptial Gifts. OSR. and Mate Competition .......... 231
3. Mate Choice Based on Nuptial Gifts ............................................................ 232
D. Interaction of Nuptial Gifts with Mating Systems
and Security of Paternity ....................................................................................233
E Interaction of Nuptial Gifts with Age-Specific Fecundity: 235
Fecundity Enhancement ...................................................................................... 235
F. Nonindependence of Adaptive Function: Interactions
Between Fecundity Enhancement and Mating Systems
Effects of Nuptial Gifts ......................................................................................235

0-8493-6695-WY5/5O.W+Ss5O
. .
O 1995 by CRC Press Inc
216 Insect Reproduction

VII. Future Prospects ........................................................................................................ 236

Acknowledgments ............................................................................................................... 237

References ........................................................................................................................... 237

I. DESCRIPTION OF MALE NUPTIAL GIFTS

A. DEFINITION
Male insects of many species give a nuptial gift to females. Nuptial gifts, as used here, are
potentially nutritious substances given to the female by the male in conjunction with mating
and used by the female in her resource budget. Such gifts can be as extreme as allowing the
female to eat part of the male,I4 as behaviorally complex as presenting a female with a prey
item on which she feeds during c~pulation?~ or as mundane as passing nutritive accessory
gland fluid into the female's reproductive tract along with spenx8 Thus, nuptial gifts are an
investment by the male in a female andtor her prezygotic offspring with potential selective
consequences. Activities such as guarding offspring9or feeding offspringi0are a postzygotic
investment and are not included in my definition of nuptial gifts. Tallamyli reviews postzygotic
investment in insects.
Male nuptial gifts are widespread across insect orders, including those with a range of
feeding habits and in a diversity of environments. Groups and the type of donation are
tabulated in several recent review^.^.'^-^^ Such tables therefore will not be duplicated here.
Nuptial gifts have ramifications throughout the biology of both males and females. Indi-
vidual-level physiology and behavior, as well as population demography and genetics, may
be affected in ecological and evolutionary time. Much of the work to date has focused on only
a few aspects of nuptial gifts, in part because nuptial gift giving is an excellent vehicle to test
sexual selection theory. However, nuptial gifts function in a much broader biological context.
While I review the more well-known aspects of nuptial gift giving, I also point out areas that
are currently begging for exploration.

B. PREY ITEMS AND REGURGITANTS

Males of some mecopteran, dipteran, hymenopteran, and heteropteran species present a prey
item to the female for her feeding during cop~lation.~* The male sometimes may feed on the prey
item himself before finding a mate. In the most well-described cases in Mecoptera, bittacid males
capture an insect prey, dangle in the vegetation, and call females by means of a pheromone; once
a female approaches, the prey is presented to the female for feeding during ~opulation.5~~"'~ If
the prey is small or has been largely eaten, females may prematurely terminate copulation.5,17
Otherwise,the male and female may fight over the prey item once males terminate ~opulation,5.'~
with the male retaining the prey most of the time. Prey may then be reused by the male in another
courtship. In at least one species, Harpobittacus similis, males with larger prey did not call as
much as males with small prey, but still attracted females.I7
Dance flies (Diptera: Empididae) exhibit a variant on this theme. Males catch prey, but the
two sexes meet either at malei8 or female7 leks. Prey items in Empis borealis can include
con specific^.^ Other empidids have altered nuptial gift giving in a stereotyped ay.^^-'^ While
male E. borealis simply present prey to females to feed on during mating, in more advanced
species prey are wrapped in silk (Hilara species) or covered with a frothy balloon (Empis
species). In yet other species, the silk or balloon surround inedible dried prey fragments;
finally, in some species the male presents the female only with an empty wrapping. Females
do not feed on this package, but only manipulate it during mating. By my definition, neither
of these last two types of gifts qualify as nuptial gifts.
Phenotypic Consequences and Evolutionary Implications 219

D. BODY PARTS AND HEMOLYMPH

The most extreme form of male nuptial gifts is cannibalism of the male by the female. This
occurs in three insect orders,12 including the mantids and ceratopogonine flies. In three
ceratopogonine tribes, females pierce the mating male's head or thorax and completely
consume him during mating.' It is not known if males are always eaten. Numbers of matings
obtained by females and effect of the male meal on female foraging and reproduction are not
known for these species. However, since cannibalism ends further male reproduction, selec-
tion pressures operating on this behavior should be quite large. Buskirk et argue that a low
expected lifetime number of matings for males (independent of cannibalism) and a significant
increase in the male's reproductive success with cannibalism are the two factors selecting for
sexual cannibalism.
A similar, but less drastic, form of nutrient donation occurs in some Orthoptera. Within the
Haglidae, female Cyphoderris buckelli chew on a male's fleshy hind wings, ingesting both
tissue and hem~lymph.~.~' Females begin feeding before copulation. The male's "gin trap", or
pinching organ which holds the female by the venter, restricts females from leaving before
copulation. Feeding is terminated by the male. Females may eat as much as one third of a
male's wings at a mating and also consume a sizeable spermatophylax. Other orthopterans
feed on hemolymph from wounds inflicted on males. Included in this group are A. fasciatus
(Gryllidae) females, who open a male's tibial spur with a mandible and feed on the
Finally, some orthopteran females feed from secretions from male metanotal glands. Included
here are female Oecanthus (Gryllidae) and some Eneopterinae and Gryllinae.3.42

E. SPERM
Sperm serve as a male nutrient donation to the female in insect species with haemocoelic
insemination, including the bedbug families Nabidae, Cimicidae, Plokiophilidae, and
Polyctenidae (Hemiptera: C i m i c ~ i d e a )In
. ~primitive
~ groups, the sperm and accessory fluids
are injected into the female's haemocoele and some sperm are digested. In other groups, the
sperm are injected into a specialized tissue, the mesospermalege, and absorbed either by free
phagocytes or by specialized cells in the mesospermalege. H i n t ~ nargued ~ ~ that haemocoelic
insemination was advantageous because females got a meal thereby, although no data exist on
the effects of sperm phagocytosis on female fitness.
Some groups, such as Lepidoptera, produce both apyrene (anucleate) and eupyrene (nucle-
ated) sperm. Apyrene sperm have been proposed to function as nutrient resources for the
female or the eupyrene sperm. However, there is no evidence for this, and apyrene sperm do
not contain much energy.(j3

11. OVERVIEW OF ADAPTIVE FUNCTIONS

OF MALE NUPTIAL GIFTS
As background to examination of the effects of male nuptial gifts on insect physiology,
ecology, fitness, and evolution, we need to understand the potential adaptive functions of the
nutrient donations. Male nuptial gifts are part of male reproductive effort. This effort can have
two potentially adaptive functions: investment in the female's resource budget andtor in
obtaining or guarding a mate.12sMBoth forms of effort can increase male reproductive success.
Contributions of resources used by females can increase female fecundity directly through an
increase in nutrients available to make eggs, or potentially indirectly through an increase in
nutrients available to support female survival. Use of resources as bribes to obtain a mate or
to prevent or delay female remating can also increase a male's reproductive success through
increasing the number of eggs that he fertilizes. These two forms of effort, contribution to a
female's resource budget and control of mating opportunities, are not mutually exclusive.
Contributions that influence mating opportunities may also be used by the female to make eggs.
 Insect Reproduction

111. PHYSIOLOGICAL COSTS AND

BENEFITS OF MALE NUPTIAL GIFTS:
EFFECTS ON RESOURCE AND TIME BUDGETS
Nuptial gifts are associated with physiological costs andlor benefits, affecting the resource
and time budgets. Changes in resource and time budgets in turn may affect foraging, survival,
and reproductive strategies, and thence fitness. I deal with specific impacts on fecundity and
survival below under demographic fitness component consequences of nuptial gifts; here, I
focus on underlying physiological effects.

A. FEMALE PERSPECTIVE
The benefit gained by females from a nuptial gift will depend on the usable donation size
relative to the female's total resource pool. Types of nutrients that are limiting to females
should have more effect than those available in abundance.65-66
Male-derived nutrients can be allocated as are nutrients derived from other sources: to egg
production, to maintenance, to defense, and to foraging activity. The specific allocation
pattern of male-derived nutrients should depend on the type of nutrients donated by males, the
female's other available reserves, and state of ovarian de~elopment.~~ If males donate com-
pounds that can be used directly in egg production with minimal processing and mating occurs
during or just prior to a peak of vitellogenic activity in the ovaries, then such compounds
should be more likely to be used in egg production. For example, in X. hamata, females feed
on male-produced urates after mating, and the timing of mating during a reproductive cycle
corresponds to the period when uptake of uric acid by the developing eggs is at a peak. As
noted earlier, large amounts of male-derived uric acid may be incorporated into eggs in this
species.52
Male-derived nutrients can reduce the need for females to forage on their own for food; this
occurs in Hylobittacusapicalis, E. borealis, Heliconiuscharitonius and Heliconius ~ y d n o . ~ . ~ , ' ~ , ~ ~
I have suggested that reduction in foraging by females with increasing male-derived nutrients
is most likely to occur when non-nutritional factors limit egg production such that females
cannot increase fecundity by maintaining foraging levels.66Such factors may include mortality
risk to the female while foraging, time restrictions for oviposition, or body size limitations on
the number of oocytes that can be matured at once.
Male nuptial gifts fed on by the female, such as prey items, external spermatophores, and
male body parts, will be incorporated into the female resource budget at the same rates as any
other similar quality food item to be used, stored or excreted. As noted above, however, rates
and targets of allocation of these items could be affected by the male if males have control over
the composition of the gift. Internal spermatophoresand male accessory gland compounds that
are absorbed by the female may be treated differently. The donation is initially present within
the female in a form of storage. Donation usage rates should depend on the balance of draw
down of other forms of storage such as fat body, use of free nutrients from newly absorbed
food, and use of the "stored" male accessory gland products. The dynamics and priority of use
of various types of nutrients from these sources in reproduction, survival, and foraging remain
to be examined. Timing and amount of reproduction, feeding sources for the adult and juvenile
female, effects of storage on risk of predation, etc., may also influence allocation patterns.
Such studies, in combination with information on patterns of paternity, will give us an
understanding of the effect of the nuptial gift on the female's resource budget and on the
reproductive success of each sex. This in turn will provide a mechanistic basis for answers to
questions about the evolution of specific forms of nuptial gifts as outlined below.
We have initial information on absorption rates of internal spermatophores and on the
usage rates in oogenesis of materials from both internal and external donations. Most of these
data come from radiolabel experiments. Males are labeled with radioactive amino acids and
Phenotypic Consequences and Evolutionary Implications 221

mated with females; then eggs andlor females are tested for radioactivity at intervals after
mating. In both an orthopteran, D. verrucivorus, and two lepidopterans, D. plexippus and H.
hecale, the amount and timing of incorporation of labeled compounds into eggs is independent
of the previous female mating hist~ry.~.~' In all Orthoptera and Lepidoptera examined to date,
label is incorporated into the next eggs laid by a female although the peak of incorporation
may be delayed by several days.s.bl.66-70 Timing of the peak of incorporation varies from
immediately after mating in Colias eurytheme,'j9to 10 days after mating in D. verr~civorus.~~
Elevated levels of label may be found in eggs for only 5 to 6 days as in C. e ~ r y t h e r n efor
,~~
7 to 10 days as in Dryas juliabl or D. plexippu~,~ or for up to 15 to 20 days as in Heliconius
er at^,^ H. c h a r i t o n i ~ s or
,~~D.~ verr~civorus.~~
~ This timing agrees with data on rates of
decrease in size of internal spermatophores: spermatophores of D. plexippus reach a baseline
size within 7 to 10 days,7' spermatophores of Pontia protodice, with a biology similar to C.
eurytheme, reach a baseline size within 5 days,72and spermatophores of H. charitonius are
absorbed within about 14 days.64In Acanthoscelides obtectus (Coleoptera), radiolabel de-
clines within 48 hours in the spermatophore, peaks in eggs laid at 36 hours, and declines by
48 hours.73In essentially all studies, all eggs laid by females after mating with a labeled male
contain at least a small amount of radioactivity. This would be expected if nutrients are
incorporated into eggs at all stages of development, such that eggs that are in late development
stages and incorporating large amounts of nutrients are heavily labeled, whereas eggs in early
stages of development, incorporating small amounts of nutrients, are lightly labeled.
The differences among species in temporal pattern of radiolabel incorporation into eggs
should depend on: (1) species-specific timing of incorporation of specific nutrient types into
eggs; (2) the particular compounds that are labeled; (3) pool sizes of those compounds; (4) the
rate at which male compounds are absorbed through the gut or reproductive walls; (5) whether
more than one day's batch of eggs is matured at once; (6) the number of eggs laid per day;
and (7) the total usable size of the male investment relative to mass of individual eggs or
clutches of eggs. These factors have been explored to some extent in B. germanica,s3 where
timing of uptake of uric acid into developing eggs is known, as is timing of ingestion of male-
derived urates. However, for the most part we know little about factors affecting different
patterns observed among species. It would also be interesting to expand the studies on
physiology of female use of male-derived nutrients to Trichoptera, as K l ~ a l i f areports~~
differences among species in the size and timing of absorption of the spermatophore.
The interaction of nuptial gifts with female time budgets is also not well understood.
Mating may not interact with the foraging or reproductive time budgets in some species. For
example, monarch butterflies (D. plexippus) can mate ~vernight?~ a time when the female
would not be actively engaged in foraging or searching for oviposition sites anyway. How-
ever, for many species in which females mate more than once, this lack of impact on the time
budget is likely to be the exception rather than the rule. Conflicts over the time spent mating
could affect the evolution and maintenance of nuptial gift-giving behavior, particularly if
mating is prolonged to allow male nutrient donations. In such cases, the time cost and
nutritional rewards of foraging by a female will be balanced against the costs and rewards of
male nuptial gifts.
Females may pass up opportunities to use male nuptial gifts altogether. Although Melanoplus
sanguinipes (Orthoptera: Acrididae) females incorporate protein derived from male accessory
gland products into eggs,26the amount of material transferred into the female's spermatheca
is small, about 5 pg, while the amount of protein in eggs laid between matings is large, on the
order of 100 mg.25However, the spermatophore, which could represent a larger male invest-
ment, was observed to be rubbed off the female's ovipositor and discarded in this species.25
Why females should bypass cheaply available nutrients is unknown. The answer could depend
on the composition of the spennatophore, or the conditions under which observations were
made.
222 Insect Reproduction

B. MALE PERSPECTIVE
Nuptial gifts affect the male's resource and time budgets. Resources spent on large
ejaculates or capturing prey are not available for other uses; time spent mating, replenishing
accessory glands, or capturing nuptial prey is not available for other uses.
Impacts of nuptial gifts on the energy and resource budget may in turn affect ability to
attract mates. Cyphoderris strepitans males produce an external spermatophore, eaten by the
female, and allow females to eat part of their hind wings and hemolymph during mating.76
Virgin males of this species call for significantly longer than recently mated males, suggesting
that energy reserves needed for calling have been reduced by mating andlor that intensity of
calling depends on distension of the male's accessory glands. Similarly, Requena verticalis
males on low-protein diets maintain spermatophore mass, but reduce calling, probably as a
result of energy li~nitation.'~
Nuptial gifts may result in matings lost because of a refractory period while accessory
glands are replenished or fresh prey are obtained. The form of this cost differs among insect
orders. As G ~ y n n points e ~ ~ out, Lepidoptera will mate before they have replenished the
accessory glands; matings simply take longer, presumably to include time for glands to be
refilled and an ejaculate formed. Even so, smaller spermatophores are formed if remating
occurs rapidly,30.62.78-79 which could affect male reproductive success. Further, in at least one
lepidopteran, recently mated males' courtship persistence time was an order of magnitude
lower than males that had not mated recently,80which will also affect male success. Orthoptera,
Megaloptera, and Coleoptera, however, have a male refractory period during which time
males will not attempt mating and accessory glands are replenished. The length of this
refractory period depends on the relative size of the spermatophore: in two species without
spermatophylaxes, Gryllus b i m a c ~ l a t u sand ~ ~ Gryllus v e l e t i ~male
, ~ ~ refractory periods are 1
hour and 30 minutes, respectively; in a species with a relatively small spermatophylax,
~ - ~ ~refractory period is 3 hours; and in a species with a relatively
Gryllodes s i g i l l a t ~ s , 8male
large spermatophylax, R. ~erticalis?~ the male refractory period is about 3 days. Megaloptera
exhibit a similar pattern. Protohermes immaculatus males have smaller spermatophores
relative to male body weight than do P, grandis males; P. immaculatus males can remate daily,
whereas P. grandis males have a refractory period of about 2 days.38Differences among
species in the length of the refractory period may also result from differences in the normal diet.
Within Meloidae (Coleoptera), seed feeding species can mate every 4 hours, whereas species
feeding on relatively protein-poor flower petals have a refractory period of 1 to 2 days.55
Nuptial gifts may also affect survival through impacts on the energy budget. This is
discussed in detail below.
Nuptial gift size should affect the relative costs to the male's resource and time budgets. Size
of the nutrient donation is expected to vary within species with the male resource budget, female
ability to receive and process nutrients, and sperm precedence patterns. First, ejaculate or
spermatophore size is positively correlated with male size (and hence, presumably nutrient
reserves) within a species in many groups, including Lytta magister and Tegrodera alogra
(Coleoptera:Mel~idae)?~ H. charitonius and D.julia (Lepidoptera:N~mphalidae),~ C. eurytheme
(Lepidoptera: Pieridae)?O Papilio machaon (Lepidoptera: Papilionidae),s3Plodia interpuctella
(Lepidoptera: P~ralidae)?~ , ~ ~sigillatus (Ortho-
Ostrinia nubilalis (Lepidoptera: P ~ r a l i d a e )G.
ptera: G ~ y l l i d a e ) , R.
~ ~verticalis
-~~ (Orthoptera: Tettigonidae),s7 Conocephalus nigropleurum
(Orthoptera: Tettigonidae),s8and D. verrucivorus (Orthoptera: T e t t i g ~ n i d a e ) .In ~ ~G.. ~sigillatus
~
and D. verrucivorus, the correlation occurs because the male's investment in the spermatophylax
or whole spermatophore, respectively, is a constant proportion of his body eight;^^.^^ male
investment as a proportion of body weight actually declines with male size in R. verti~alis.~' The
data suggest that absolute spermatophore size is more constrained in R. verticalis than in the
other two species. These differences could be related to differences among species in diet, in
spermatophore size relative to male body size, andor in spermatophore function.
Phenotypic Consequences and Evolutionary Implications 223

In contrast, no relationship was found between male size and ejaculate mass or volume in
P. protodice (Lepidoptera:Pie~idae)'~ or Pararge aegeria (Lepidoptera: Satyridae).goRutowski72
suggests that this may be due to differences in the primary function of the spermatophore
among Lepidoptera, with P. protodice donations functioning more to prevent female remating
than as a nutrient investment. This idea is explored in more detail below.
Among species, Reiss9' argues that the investment per unit time in reproduction by either
sex should scale allometrically with body weight, with a coefficient between 0.5 and 0.9; the
precise predictions for each group will depend on the allometries of energy intake and
nonreproductive expenditure with body weight. Extensive data to test this hypothesis do not
exist for male nutrient donations.
Male ejaculate or spermatophoresize increases with male age in virgin male D. ple~ippus?~
but not in 0. n ~ b i l a l i sThis
. ~ ~suggests that males of some species may need time after eclosion
to reach complete sexual maturity (as measured by filling of the accessory glands) andfor that
adult feeding contributes to the formation of spermatophores.
Spermatophore size decreases with number of previous matings by a male in some
O r t h ~ p t e r aLepid~ptera~O.'~,~~.~~
,~~ and Tri~hoptera.~~ Change in size of spermatophores with
previous mating history is likely to be determined by the opportunity for males to replenish
reserves from feeding, the average and range in number of matings by males, and the intensity
of selective pressure for maintaining a minimum size to guarantee sperm transfer.
Quality of the male's diet and presence of parasites affect the male's resource budget and
hence may affect the size of the nutrient donation. Z Ushowed ~ ~that~the number of spermato-
phores produced in 24 hours decreased with number of gregarine parasitic cysts in the feceae
of G. veletis and G. pennsylvanicus (Orthoptera: Gryllidae), but no relationship existed for
shorter term spermatophore production measures. Further, R. verticalis males infected with a
protozoan gut parasite had lower mating frequencies than uninfected males when fed a poor
diet. However, the effect of parasitic infection disappeared when males were maintained on
a rich diet, indicating that parasites have the effect of lowering experienced diet quality as far
as impact on mating success is concerned?'
Female ability to receive or process the nutrient donation could constrain the size of the
nuptial gift, placing upper limits on size variation. In species with an internal spermatophore,
female size and previous number of matings (if the spermatophoreis not completely absorbed)
may constrain the size of the male's nutrient investment due to space available in the bursa
copulatrix or appendix bursa. Some evidence for this comes from D. julia, where female
winglength interacted with other parameters to affect spermatophore size.64Finally, female
mating status has an effect on spermatophylax and ampulla size in D. verrucivorus; virgin
females obtain larger spermatoph~res.~~

IV. EFFECTS OF NUPTIAL GIFTS ON

DEMOGRAPHIC FITNESS COMPONENTS
Fecundity and survival are components of individual fitness and, as noted earlier, observed
patterns of birth and death may be direct consequences of male nutrient donations because of
the impact of these donations on the time and resource budgets of each sex. In this section,
I explore the translation of effects on time and resource budgets into effects on fecundity and
survival. Explicitly evolutionary relationships between nuptial gifts and fitness components
are dealt with later.

A. AGE-SPECIFIC FECUNDITY PATTERNS

Mating can affect age-specific fecundity patterns either through hormonal mechanisms
stimulating oogenesis ancllor o v i p o ~ i t i o nor
~ ~through
- ~ ~ alteration of the female's resource
budget via nuptial gifts. These two causes can be difficult to separate, especially in species
224 Insect Reproduction

with internal spermatophores. Here, I am not concerned with oogenesis or oviposition stimu-
lating factors, but rather with effects on fecundity via effects on the female's resource budget.
Male nutrient donations may affect fecundity independently of whether nuptial gifts arose
and/or are maintained specifically in the selective context of increasing female reproductive
success (and hence, male success). Rather, whether fecundity is affected by nuptial gifts
should depend on the size and composition of the donation relative to the female's overall
nutrient stores and the timing of the donation relative to ~ o g e n e s i sFecundity
.~~ enhancement
is expected if: (1) at least some oogenesis occurs after mating; and (2) the type of nutrient
donated is a limiting factor in egg production. A nutrient may be limiting either due to lack
of foraging opportunities either in the adult or juvenile stage or due to an evolutionary history
during which females have become dependent on male donations to replace foraging. In these
cases, we expect that egg numbers will increase over the short term after mating, consistent
with timing of incorporation of male nutrients into eggs outlined above. Alternatively, lifetime
fecundity may also be enhanced if male donations allow females to reduce foraging and by
so doing, decrease death rates from predation or increase time available to lay eggs. This
means that species which live either in harsh or dangerous environments or whch use
ephemeral but nutritious resources, are likely to show an effect of male donations on fecund-
ity .13,62
Some support exists for these ideas, as outlined by B ~ g g s Within . ~ ~ Orthoptera,
spermatophylax consumption has an effect on the next few days' fecundity in several species,
including R. v e r t i c a l i ~and
~ ~Chorthippus b r u n n e u ~This
. ~ ~ impact is affected by the quality of
the diet fed to experimental females. More directly, spermatophylax consumption and a
seasonally available high quality diet had equivalent effects on female fecundity in an
unnamed zaprochiline katydid." Not all tettigonids show an effect of male-donated nutrients
on fecundity, however. Even on a restricted diet, no effect was detected for D. verr~civorus.'~
The spermatophylax in this species is just large enough to ensure that all sperm are transferred
to the female after mating,",% and has a relatively low protein ~ontent,~' suggesting little
opportunity for male-donated nutrients to be important in the female's resource budget, and,
hence, to affect fecundity - unless mating were to occur frequently. Within Lepidoptera,
larger spermatophores increased post-mating fecundity in C. eurytheme.lWFemales in this
experiment were fed a relatively poor diet consisting of a 10% sucrose solution; we do not
know how often females are food stressed in the field. Within Diptera, semi-starved D.
subobscura females showed an increase in fecundity the first 2 days after mating if they were
fed by males as compared to not fed.21A comparison of crop sizes in wild and lab-fed or lab-
starved flies showed no difference in crop size between wild and starved flies, but wild flies
had significantly smaller crops than fed flies.21This suggests that females are frequently
nutrient stressed in the field. Hence, male-donated nutrients likely play an important role in
egg production in this species. In Drosophila mojavensis, receipt of a large male donation
increases early fecundity only if females are held without access to yeast.lo1Further, females
of Panorpa spp. feeding on dead arthropods during mating showed increased fecundity
compared to those not feeding.20ThornhillZOfound evidence for interspecific competition for
food in this group, and for significant mortality when individuals forage from spider webs,
suggesting that food was scarce and costly to obtain. Finally, withn Coleoptera, unfed
Caryedon serratus females show higher fecundity if allowed multiple matings instead of only
one mating, although the authors are not convinced that their results were due to a nutritional
rather than hormonal stimulus effect.34

B. SURVIVAL PATTERNS
In cases of sexual cannibalism, male nutrient donation ipso facto reduces male survival
rate. The effect of donations on male survival is seldom this severe for other types of male
donations. Nonetheless, we lack a detailed understanding of the impact of nutrient donations
and mating numbers on male survival. Interesting questions include: Do males whose nutrient
Phenotypic Consequences and Evolutionary Implications 225

donation is a sizeable portion of their body mass suffer a relatively greater survival cost than
males with a relatively smaller donation? Does the survival rate differ depending on whether
specialized male structures are eaten or accessory gland products are donated? If accessory
gland products are involved, does the effect on survival depend on the match between the
composition of the male-donated nutrients and the composition of the adult male diet and
hence the ability to replenish the nutrient pool? Understanding the answers to these questions
will allow us to understand physiological costs of mating to the males and how such costs
translate into fitness-related survival patterns.
Refractory periods found in Orthoptera and Megaloptera, or the decrease in nutrient
donations with frequent matings seen in Lepidoptera, may act to buffer survival rates against
effects from reproductive expenditures by limiting reproduction. Within Lepidoptera, D.
plexippus males given the opportunity to mate every day had the same life span as males with
no opportunity to mate;Io2a similar result was obtained for P. aegeriaw, the orthopterans G.
sigillatus and G. veleti~,~' and the megalopterans Prothemes grandis and P. immac~latus.~~
This lack of effect could stem from a limitation on reproductive expenditure, from compen-
satory feeding by males with more matingsIo3as occurs in H. c h a r i t o n i ~ sor, ~from
~ ~ a lack
of impact of mating expenditures on types of nutrients important in survival.
In an interesting twist, a lack of opportunity to pass nutrients to females can impact male
survival under some conditions in at least one roach. X. hamata males fed on foods with a high
nitrogen content (as are chosen in the lab and field) can die from uric acid toxicity if not
allowed to mate and give some urates to females.52How important this is under field
conditions as a source of mortality is unknown, but if the variance in distribution of mating
numbers among males is at all high, significant numbers of males could be exposed to this
mortality source under appropriate nutritional environments.
Effects of male donations on female survival are somewhat more variable than those seen
to date on male survival. No survival effect of increasing the quantity of male nutrient
donations has been found for D. p l e x i p p ~ s ~or~ , 0.
~ ' nubilalisx4within Lepidoptera, or D.
verrucivorusyxwithin Orthoptera. However, increasing male nutrient donations increased
female survival in C. eurythemelWand Psuedaletia unipunctalo5within Lepidoptera, and G.
sigillatus and G. veletisxlwithin Orthoptera. As for males, the observed results may be affected
by possibilities (in the field or lab) for compensatory feeding if females receive small
donations, or by the type of nutrients donated by males at mating and their importance to
survival processes.hhIn the latter case, it is also possible that a shortage of male-donated
nutrients normally used for survival could cause a reallocation of female-derived resources
away from reproduction, maintaining female survival rates at the expense of reproduction.
Such reallocation driven by decreases in male-derived nutrients has not been explored, but
Speyeria mormonia females (Lepidoptera: Nymphalidae) reallocate resources from reproduc-
tion to survival in the face of adult female food shortages.lo6
Effects on survival may occur not just through impacts on the resource budget, but also
through increases or decreases in exposure to predators because of changes in foraging activity
by either males or females. Through feeding on nuptial gifts rather than foraging on their own,
the incidence of spider predation is apparently reduced in the hanging fly H. apicalis, as
females are found significantly less frequently than males in webs, whereas there is no
difference by sex in a similar species without nuptial prey gifts.16 Conversely, in mormon
crickets and conocephaline katydids, the sex competing for mates (and hence, with a relatively
smaller investment) is more active and suffers more wasp predation.'07
Finally, nuptial gifts can affect offspring survival rates. Increasing the female's supply of
defensive compounds, such as pyrrolizidine alkaloids in ithomiine b ~ t t e r f l i e sor~ ~m0ths,5~
could increase the survival rates of offspring. Offspring survival can also be increased if male
donations increase the female's supply of trace nutrients, including sodium, as seen in
Thymelicus l i n e ~ l aThe
. ~ ~same effect may explain data for R. verticali~:~~
females eating more
or larger spermatophylaxes laid larger eggs; larger eggs have higher over-winter survival
226 Insect Reproduction

rates, and, if male, differ in developmental rate compared to smaller eggs. Improved female
diet alone did not increase egg size in this species.

V. EFFECTS OF NUPTIAL GIFTS ON POPULATION STRUCTURE

Individual fecundity and survival influence fitness relative to the fecundity and survival of
other members of a population. The composition of the population is affected by its age
structure, individual dispersal habits, and effective population size. Male nuptial gifts can have
effects on these three parameters, and hence on the population context within which selection
may occur.

A. AGE STRUCTURE
Adult age structure is affected by development time of juveniles, as well as age-specific
death rates of adults. Male nuptial gifts may interact with juvenile development time through
impacts on the resource budget of each sex. If nutrients gathered during the juvenile stage are
used by males in procuring or making nuptial gifts, then longer development times resulting
in larger male resource stores and consequently larger donations should be favored, all else
being equal.lo8Conversely, females receiving substantial nutrient input from males could have
shorter development times if resources normally obtainable only in the juvenile stage are
provided by males and juvenile mortality rates are high. These ideas could be tested using
selection experiments in a species with a suitably short life cycle. Alternatively, development
times could be compared for closely related species from the same habitat with similar food
sources and sperm precedence patterns, but which differ in male nutrient donations.
Further, sperm precedence patterns favoring protandry, or adult emergence of males before
females, may also affect the size of male nutrient donation^.^^ If sperm from multiple matings
mix, then a male's net fitness increment from mating with a female decreases with increasing
number of previous matings by the female. Males could offset this decline by increasing the
number of sperm transferred, which in Orthoptera can entail a larger nutrient donation to
ensure sperm transfer. However, the net gain from a given increase in number of sperm
transferred diminishes with increasing numbers of previous matings by the female. As
predicted in a protandrous system with sperm mixing and in which the male nutrient donation
is just large enough to ensure complete sperm transfer, D. verrucivorus males make their
largest contribution to virgin females, independent of male mating statu~.~"

B. DISPERSAL
Male nuptial gifts may interact with dispersal in two ways: nutrient donations may provide
necessary resources for successful dispersal by females, or females may view males as a
resource and refrain from leaving areas with males.lW
The first idea, that nuptial gifts allow females to disperse or to migrate, has been explored
in monarch butterflies (D. ple~ippus)."~-"* Overwintering monarchs in Mexico and coastal
California have different mating regimes before the spring migration.Il0In Mexico, nearly a
third of migrating females are virgins, and males and females leave the colony at about the
same time. For virgin females, mating presumably occurs on the trip north, since females
oviposit on milkweeds during the migration. Prior to migration, mating males are more worn
and smaller than the average for the population, whereas mating females are less worn and
larger, and males attempting matings appear to discriminate against mated females.ll' In
coastal California, 95% of females mated between 1 and 7 times before moving north, and
many males may not leave the colony at all. Thus, there are different patterns in these two
populations for the timing of entry of male-derived nutrients into the female resource budget
with respect to the timing of migration. These differences could be affected by sperm
precedence patterns, physical environmental parameters, activity levels of over-wintering
individuals, and distance to the nearest milkweed.'I0
Phenotypic Consequences and Evolutionary Implications 227

Wells et have shown that female lipid content increases once mating begins in the
California monarch aggregations, while male lipid content continues to decline. They attrib-
uted this difference to a shift of nutrients to the female from males at mating. Using measured
energy expenditures by monarchs, the amount of energy estimated to be obtained at each
mating by the female, and the timing of first access to milkweed plants for oviposition in the
spring, they did Monte Carlo simulations examining expected long-term fate of monarch
populations. Their results indicate that multiple mating upon leaving the winter roosting site
is necessary for long-term persistence of the population.
Experimental measurements of effects of male nutrients on either dispersal ability or on
reproductive ability once a new habitat has been reached have not yet been published.
However, several authors have turned the prediction around, arguing that species which are
long-lived migrants should be expected to have large nutrient donations from the male to the
female.10sP. unipuncta is one such case, as female fecundity and life span are increased by
multiple mating in this migrant.lo5
An alternative effect of significant male nutrient donations is for females to refrain from
dispersing and to remain in areas with males, using males as a nutrient resource. Monarch
butterflies in Australia, which do not form overwintering aggregations but breed year-round,
may provide an example of this effect.logFemale density is lower inside dense milkweed
patches than on the edges, and male density shows the reverse pattern, suggesting that male
harrassment may drive females away from the center of the patches. However, if males are
removed from an area, females tend to disperse, whereas if population density is simply
reduced, the effect is not seen. Use of males as a resource, and hence females remaining in
an area with males, need not be the only explanation for this result, but it is certainly a possible
cause. It could be instructive to test this hypothesis by contrasting the female dispersal patterns
in Hylobittacus apicalis, whose females depend on male-provided prey items,5J6with related
species with QO prey gifts and whose females hunt.

C. EFFECTIVE POPULATION SIZE

The effective population size (N,) is a measure of the number of individuals actually
contributing offspring to the next generation, as opposed to the total number of individuals in
a population. Among other things, effective population size governs the power of genetic drift
as a significant evolutionary factor. As the effective population size becomes smaller, drift
may play a more powerful role affecting evolutionary change.
A given population will exhibit a characteristic mean number of matings per individual, but
some individuals will mate more often than others. Male nuptial gifts may affect Ne by
affecting the male refractory period after a mating, as noted for Orthoptera and
The male refractory period in turn affects the size of the pool of males
available to mate by removing males from the pool for a length of time equal to the refractory
period. Thus, the identity of the males in the pool of potential mates is constantly changing,
with some turnover rate dependent on the size of the donation. The larger the pool of males
available for mating relative to the absolute number of males, the greater the chances that the
pool contains males that have recently mated, and thus the greater the chances that a male that
has already mated will secure the next mate, or that a male that has not yet mated will not
secure the next mate. Note that this effect only occurs if the operational sex ratio is male
biased. Otherwise, all males in the pool at any point in time should be able to obtain matings,
since there are more females willing to mate than males available, resulting in all males in the
population obtaining at least one mating.
The practice of males donating nutrients at mating may also affect N, through resource-
based effects on the operational sex ratio (OSR). When the OSR is biased towards one sex,
the possibility exists that not all members of the over-represented sex will obtain matings and
contribute to the effective population size. Hence, the component of Ne determined by mating
success (a first step in representation in the next generation) may be significantly lower than
228 Insect Reproduction

the actual observed population size for the over-represented sex, but close to the actual
observed population size for the under-represented sex. The OSR can be affected by resource
availability in populations with large nutrient donations at mating. If females are in a resource-
poor environment, they attempt to replace nutrients unavailable from the environment with
nutrients from males at mating; conversely, males may experience longer refractory periods
since a given donation is more expensive relative to available resources. The OSR may
become female biased under such conditions. In a rich resource environment, females are less
willing to spend time mating, since nutrients are not scarce and males experience shorter
refractory times. The OSR may become male biased under such conditions, even in the same
population.
Male refractory times are not the only item that may constrain the pool of available males
in a resource-poor environment. In species with nuptial prey, the abundance of prey items in
the environment may affect hunting times, which could in turn affect the number of males with
prey available as mates, affecting male N,. In environments with low prey availability, males
which are better hunters or more able to steal prey items may get more matings, increasing
the number of males with zero lifetime reproductive success and decreasing N,.
Species for which diet affects the male refractory period or the size of the male nuptial gift,
or for which prey availability is important, then, may experience fluctuations in N, as a
function of variation in the nutrient environment, even without fluctuations in observed
numbers of individuals. These fluctuations may have important consequences for the popu-
lation genetic structure, and the role of genetic drift.
Data on effects of donation size on N,do not currently exist; for that matter, population data
on insect lifetime reproductive success in reasonably natural environments are scarce.Il3
Species which alter the size of donations as a function of the food environment would be good
candidates for study.

VI. PATTERNS OF THE EVOLUTION OF NUPTIAL GIFTS

A. ORIGINS: PHYLOGENETIC HISTORY
Nuptial gifts may be a very old trait within Insecta. Kl~alifa'~ and Davey29,43argue that
presence of spermatophores is a primitive trait within this class and has been lost multiple
times during radiation of various groups. Thysanura possess spermatophores, as do the
orthopteroids and neuropterans. DaveyZ9points out that other groups without spermatophores
often retain reduced accessory glands. The antiquity of spermatophores within Insecta means
that the potential for male nutrient donations via the spermatophore or similar accessory gland
secretions is probably at least as old as the class. Although spermatophores are believed to
have arisen in the context of facilitating sperm transfer in a terrestrial e n ~ i r o n m e n t the
,~~
possibility of using the structure as a means to provide females with extra nutrients was
present. Thus, spermatophores may have been exaptations for male nutrient donations.
Within orders, or genera, phylogenetic history can be an important determinant of both the
presence and size of male nutrient donations. Pitnick et a1.Il4found that monophyletic species
groups in Drosophila have similar sized ejaculates and similar levels of incorporation into
ovaries and female soma of radiolabeled male nutrient donations. This is in spite of the fact
that species compared within a species group differed in nutritional ecology and habitat.
Likewise, comparison of D. mojavensis and D. pachea, both cactophilic species endemic to
the Sonoran desert but members of different species groups, showed that these two species
differ dramatically in ejaculate size and incorporation by the female, but were similar to other
members of their species group.
Phylogenetic history undoubtably plays an important role in determining the degree of
male nutrient donation within Orthoptera as well. As noted above, families differ in the
presence or absence of a spermatophylax, which increases the size of the male donation.
B o l d y r e ~ "argues
~ that the ancestral spermatophore was probably similar to a simple ampulla
Phenotypic Consequences and Evolutionary Implications 229

without the spermatophylax; in that case, the spermatophylax could have evolved as an
elaboration in circumstances where increased male nutrient donation was f a v ~ r e dThere
. ~ ~ is
some uniformity of spermatophore structure within families,IL6so the elaboration of the
spermatophore occurred relatively early. Many of the relationships between male nutrient
donations and other aspects of the biology of a given orthopteran species may be constrained
by the phylogenetic history associated with spermatophore production.
Phylogenetic history may also play a role in the development of presentation of prey items
to females at mating or other behaviors associated with male nuptial gifts. Certainly the
variations seen in Empis or Hilara species, ranging from presentation of a prey item in
primitive species to wrapping the prey item in silk or a balloon to presentation of an empty
wrapping in the most derived species, indicate that phylogeny can have significant effects.I8-l9
At a more basic level, a survey of mecopteran species which do and do not have male nuptial
gifts, examining both phylogenetic relationships and habitat or ecological specialization,
could be instructive as to the relative roles of phylogeny and ecology or habitat in influencing
the evolution of presentation of nuptial prey.

B. CURRENT VARIATION AND FUTURE PROSPECTS: HERITABILITY

Male nuptial gifts range from secretions, whose evolution must be based on metabolic
control of resource allocation, to presentation of prey items, whose evolution must be based
on behavioral adaptations. The discussion so far assumes that variation in nuptial gifts has a
genetic basis, and hence selection and evolutionary change is possible. However, the genetics
underlying nuptial gift giving is not yet understood for any species. In fact, in only one case
has heritability of any aspect of the nuptial gift been measured. Based on father-son regres-
sions, Sakaluk demonstrated a -47% heritability for spermatophylax mass/male body mass in
G. s i g i l l a t ~ s . ~
None
~ - ~ of
~ ampulla mass/male body mass, ampullas mass alone, or
spermatophylax mass alone showed significant heritabilities. Sakaluk points out that this
result suggests that the size of the ampulla and spermatophylax are genetically uncoupled.
This makes sense given that the composition of the two are not the same, and hence the two
are likely produced through different metabolic pathways with the potential for differing
control. Further, the fact that percent body mass allocated to spermatophylaxes showed
heritable variation while absolute spermatophylax mass did not suggests that the controlling
factor is an enzyme regulating allocation rate from some finite pool to male accessory glands
andlor gene(s) affecting the relative size of the accessory glands.
Sakaluk suggests that variation for relative spermatophylax size may be maintained by
fluctuating food environments. In a high food environment, a large spermatophylax may not
increase the female's fecundity, but may decrease the number of matings a male obtains due
to a longer male refractory period. The disadvantage of a longer refractory period still holds
in low food environments, but female fecundity may be increased by a large spermatophylax.
Similarly, lack of heritable variation for ampulla mass may have resulted from lack of
variation in selection pressures affecting ampulla mass since the costs or benefits of number
of sperm transferred do not vary with environmental food availability.

C. INTERACTION OF NUPTIAL GIFTS WITH

MATE COMPETITION AND MATE CHOICE
Given the possible adaptive function of nuptial gifts in enhancing female fecundity and
securing matings, we can ask whether or not aspects of the reproductive biology of insects are
coadapted with male nuptial gift giving. That is, are types, compositions, sizes, or timing of
nuptial gifts coadapted with female fecundity, security of paternity, mate competition, or
choice, or mating system? Are there patterns of association among traits which have high
fitness value and could be selected for in natural environments over evolutionary time? I first
consider interactions with sex roles, then with mating systems, then with age-specific fecund-
ity, and then I conclude by pointing out that these interactions themselves are not independent.
230 Insect Reproduction

Coadaptation of nuptial gifts occurs not just with mating systems, but with the combination
of mating systems and age-specific fecundity.

1. Sex Roles and Sexual Selection: Theory

Much of the work on insect nuptial gifts has been done to explore the effects of relative
reproductive investment by each sex on sex roles and the operation of sexual selection. This
work was stimulated by a series of authors, beginning with Darwin. Before examining the
results of experiments relating male nuptial gifts to sexual selection, I will explore the
conceptual context for those experiments.
Trivers1I7built on arguments by Darwin,lt8 Fisher,Il9 Bateman,120 and W i l l i a m ~ , land ~~
argued that differences between the sexes in amount of investment in offspring should control
the operation of sexual selection. Trivers defined this critical parameter, parental investment,
as "any investment by the parent in an individual offspring that increases the offspring's
chance of surviving (and hence reproductive success) at the cost of the parent's ability to
invest in other offspring."117Thus, individual fitness is the currency in which parental
investment is measured; investments must increase the fitness through one offspring's sur-
vival at the cost of ability to realize fitness increases through other offspring. Trivers goes on
to argue that the sex with the larger parental investment will limit the realized reproductive
success of the sex with the smaller investment. Thus, the sex with the smaller investment
should experience intrasexual competition for mates. Further, the sex with the larger invest-
ment has more to lose from a bad choice of mates than does the sex with the smaller
investment. Thus, the sex with the larger investment should exhibit mate choice. Presumably
to avoid circularity, Trivers defined parental investment explicitly to exclude investment in
obtaining mates through "sexual competition for mates".Il7
Trivers' definition has lead to a proliferation of papers on male nuptial gifts as parental
investment vs. mating This distinction is critical to testing Trivers'
hypothesis, since only shifts in relative parental investment are predicted to lead to shifts in
sex roles.
Taking a different approach to the problem of sex roles, Emlen and Oring128examined
environmental and ecological effects on mating systems. They argued that which sex expe-
riences intrasexual competition for mates depends on the Operational Sex Ratio (OSR),
defined as the sex ratio of individuals currently ready to mate. That is, if receptive females are
present in smaller numbers than reproductively active males, females represent a scarce
resource for males, and males should compete among themselves for access to females and
vice versa. As pointed out, the OSR may be driven by parental investment
patterns if the size of the parental investment affects the amount of time spent out of the mating
pool by either sex.
Recently, these arguments have been substantially expanded by Clutton-Brock and Parker.130
They include time in their analysis, build explicit connections between investment and the
OSR, and acknowledge a variety of other factors that may influence the OSR or intrasexual
mate competition. They begin by stating that the OSR determines which sex will compete for
mates, using arguments similar to Emlen and Oring's.12%However, rather than parental
investment, Clutton-Brock and Parker focus on potential reproductive rates of each sex as the
major factor determining the OSR, along with the local adult sex ratio and sexual differences
in survival rates. Potential reproductive rates are in turn affected by investment in offspring,
courtship, and mating, as well as physiological and environmental constraints. In effect,
Clutton-Brock and Parker have interposed the OSR and potential reproductive rates as
intermediary steps between parental investment and mate competition, and have acknowl-
edged other factors that can affect the outcome as to which sex is competitive.
Clutton-Brock and Parker's130 theory does not require the distinction between mating and
parental investment required by Trivers. Each type of investment is important in so far as it
affects the potential reproductive rate. Since a nuptial gift may serve both functions, as for
Phenotypic Consequences and Evolutionaly Implications 231

example nuptial prey in Mecoptera or spermatophores in some Lepidoptera, this theory is

more amenable to experimental test.
There is a remaining problem yet unaddressed. Many authors explicitly or implicitly
assume that where intrasexual mate competition occurs, mate choice must be occurring in the
opposite sex or that conditions leading to mate competition produce mate choice by the
opposite sex. Clutton-Brock and Parker130are obvious exceptions to this pattern, as they only
address mate competition in their analysis. Trivers,l17however, considers factors affecting the
operation of sexual selection in general, and does not examine potential differences in causes
of intrasexual competition and intersexual mate choice.
There is no expectation for mate choice to automatically accompany mate competition or
vice versa, however. Males may compete with each other for access to a female's feeding
territory, or, in some Heliconius, males defend female pupae from each other.l3I In these cases,
the female is not exerting mate choice. Alternatively, in Colias, females choose mates,132-134
but males are not competing with each other for access to females.
Three factors are important prerequisites for the mate choice to be expected. First, more
than one potential mate must be reliably available, to allow a choice to be made. This does
not mean that the operational sex ratio must be skewed towards the nonchoosy sex, although
that certainly helps. Second, fertility (for females) or paternity (for males) must be assured,
unless choice is to be made among potential mates on the basis of which one will assure
genetic representation of the chooser in offspring. Third, one sex must be able to control
resources needed by the AND there must be differences among individuals in
quality or quantity of the resources controlled, in order for a basis for choice to exist. Such
resources may be genetic, nutritional, or portions of habitat. Thus, choice is likely to be seen
in populations with skewed OSRs due to density or lekking behavior, in populations whose
individuals mate multiply andlor show last male sperm precedence, and in species in which
variation in male nutrient donations or female fecundity are discernible due to age, size, or
behavioral differences among differing individuals.

2. Experimental Evidence: Nuptial Gifts, OSR, and Mate Competition

Mate competition can take at least two forms in insects with nuptial gifts. The first is the
standard one of fighting between members of one sex for access to a receptive individual of
the opposite sex. Fighting for access to mates has been documented within O r t h ~ p t e r a ~ ~ . ~ ~ ~ . ~ ~ ~
and some Lepidoptera.I3l Within Lepidoptera, some Heliconius species' males find a female
pupa and sit on it about 24 hours before female emergence. Two males will fit on a pupa, one
on each side. Males may attempt to take over pupae by dislodging resident males in much the
same way they attempt to take over matings in other species. One defending male then mates
with the female either just before or at female eclosion.
The second form of mate competition is theft of nuptial prey items, as occurs in H.
bittacus.16Males can act as "transvestites", approaching a calling male as if for mating; in one
study, calling males actually offered the prey item to the thief 67% of the time.16Males which
gain a prey item may also thereby be able to gain a mate at the expense of delayed mating
possibilities by the male losing the prey item.
Clutton-Brock and Parker'sI3O theory predicts that changes in the potential reproductive
rate of each sex, which may be driven by the relative value of the nuptial gift, affect the
operational sex ratio and hence, determine which sex is the competitive sex. Evidence from
Orthoptera is consistent with the theory. For example, at high food availability, mating
frequency by a zaprochiline katydid female is reduced; at low food availability, female
fecundity is maintained only through spermatophore con~umption.9~ Thus, the fraction of
females in the mating pool at any one time is lower at high food availability such that the
operational sex ratio should be more likely to be male biased at high food availability and
female biased at low food availability. As expected, males compete for mates when adult food
is plentiful, but females compete for mates when adult food is scarce.136In another case,
232 Insect Reproduction

Anabrus simplex occurs in either high or low density in Colorado, U.S.137Males in the high
density site have lighter accessory glands than males in the low density site. Given the
refractory period after a mating while glands are refilled, these data are consistent with the
observation that few males were calling at the high density site. Average number of matings
by females did not differ between sites. If high density led to longer male refractory periods
at the high density site, the OSR may have been biased towards females at that site, and
towards males at the lower density site. This would produce the observed sex role reversal
between the two sites, with competitive females at the high density site only.
The fact that sex role reversals tend to be observed in Orthoptera with relatively large male
investments and multiple matings by females is also consistent with the theory. Larger male
investments lead to increased refractory times (see above), decreasing the number of sexually
active males at any point in time. Further, the possibility of female dependence on male
nutrient donations when other adult food resources are scarce means that the proportion of
sexually active females in the population can increase under some circumstances. Thus, the
OSR can swing from male to female biased depending on food availability, accompanied by
changes in which sex competes for mates.
The interaction between energy needed for male nutrient donations and for mate calling can
also produce a biased OSR, but only if noncalling males are not part of the mating pool. For
example, C. strepitans males' wing pads are fed on by females during mating, impairing the
ability of nonvirgin males to call for mates.76Likewise, R. verticalis males held on low diets
maintain spermatophore size, but reduce calling for mates due to energetic limitation^.'^ Since
female competition has not been observed in the latter species, the theory predicts that male
refractory periods should be relatively unchanged by low quality diets and that noncalling
males really are part of the mating pool - that females are able to find them for mating
purposes.
In species without direct mate competition, sex role reversals may take the form of shifts
in the sex initiating courtship. This has been documented for the pierid butterflies P. p r o t ~ d i c e ' ~ ~
Colias philodice, and C. e ~ r y t h e r n ewhose
, ~ ~ ~ females approached males and elicited courtship
chases once the bursa1 spermatophore contents were largely depleted. This system would be
interesting to examine in the context of interactions among resource availability, potential
reproductive rates, and the operational sex ratio.

3. Mate Choice Based on Nuptial Gifts

Mate choice can be accomplished in several ways. The first way is primary choice through
rejection or acceptance of mating with a prospective mate. Secondary choice by females
occurs if the female rapidly remates with another male (in systems without first male sperm
precedence) or if females control the length of copulation and hence, the amount of sperm
transferred, as occurs in some bittacid~.~ In the orthopteran Gryllus bimaculatus, females use
both approaches: females may remove some males' spermatophores before all sperm are
transferred, but remain near large males, remating several times.140
Above, I predicted that presence of choosiness should depend on reliable availability of
mates, assurance of fertility or paternity, and ability of one sex to control resources needed by
the other combined with distinguishable differences among individuals in ability to provide
those resources. Available data for choice involving nuptial gift donations supports these
predictions. First, food availability can influence choosiness in each sex through effects on
mate availability. For example, in a zaprochiline tettigoniid, calling males were frequent under
conditions of high food availability and females were choosy, rejecting males. However,
females were eager to mate and receive nutrients under low food availability, the OSR was
female biased, and males were choosy.'36
Second, in D. julia (Lepidoptera), females tend to mate on their second and subsequent
matings with males who have had large numbers of matings, but have not mated recently;
these males provide larger spermatophores than males that have mated more recently. This
Phenotypic Consequences and Evolutionary Implications 233

pattern is not observed for the first matings by fema1es.l4' Females in this species can fertilize
all eggs from a single mating if access to males is restricted.142A similar increase in female
discrimination is observed in Colias butterflies.143Thus, female choosiness can increase after
a first mating if sufficient sperm are obtained from one mating to fertilize all of a female's
eggs. In a twist on this idea, D. verrucivorus males contribute a larger spermatophore to virgin
females than to nonvirgins, which constitutes a form of investment choice; males probably
father more offspring of virgins than non virgin^.^^
Both sexes can exercise mate choice simultaneously. For example, in P. protodice, male
courtship duration depends on the type of female: larger, younger females are courted longer
than others.144In the same species, females discriminate among males based on the duration
and intensity of courtship.s0Since choice can be exercised by both sexes simultaneously, then
the identity of the choosy sex(es) cannot be dependent only on the operational sex ratio or on
the relative amount of parental investment by each sex. An examination is needed of threshold
conditions for choice, considering mate availability, fertilitylpaternity assurance, and discern-
ible differences among individuals in resource control.

D. INTERACTION OF NUPTIAL GIFTS WITH

MATING SYSTEMS AND SECURITY OF PATERNITY
Nuptial gifts can function as investments in obtaining a mate or in ensuring complete sperm
transfer. A male either presents a prey or is expected to donate a ~permatophore,~~ which gives
the female sufficient net benefit that she is willing to mate with that male. Nuptial gifts may
thus function as bribes, as an index of male genetic quality, or both. For species which have
external spermatophores or which present arthropod prey for the female to feed on during
mating, the size and quality of the male donation can determine whether the maximum number
of sperm are transferred. For example, in G. sigillatus, the length of time the ampulla remains
attached, allowing sperm to be transferred and stored in the female, depended on the size of
the spermatophylax and how long it takes females to eat it; once they had eaten the
spermatophylax, they removed the ampulla and ate it as Copulation duration in H.
apicalis depended on size of the offered nuptial prey item, with a threshold size above which
copulations lasted long enough for complete sperm tran~fer.~ In these cases, the function of
the donation may differ, depending on the vantage point: for females, it is still an investment
in their resource budget; for males, it is expenditure in obtaining a mate and not necessarily
a direct investment in offspring.
A prediction following from this function is that a threshold investment in the female's
resource budget exists, and males that cross that threshold obtain mates andor complete sperm
transfer. The threshold might shift, depending on the state of the female's resource pool, the
relative time cost of mating, and the current risks associated with foraging herself. Quantita-
tive shifts in thresholds have not been explored in detail yet, except in the extreme case where
females are on very poor diets, which can lead to sex role reversals where females are willing
to accept any male as they compete for males, rather than choosing among males.97.136-137
Male nuptial gifts can also function to prevent females from remating. This can be
accomplished in several ways. First, female refractory periods may be induced through
compounds in the e j a c ~ l a t e . Second,
~ ~ - ~ ~ mating plugs (which might also function as nuptial
gifts if they are absorbed by the female) may be formed in the female's genital tract, which
physically prevent other males from mating with the female.28Third, for some species with
internal spermatophores, the spermatophore triggers a stretch receptor in the bursa, initiating
nonreceptivity; this has been shown for Pieris r a p ~ eThe . ~ decay
~ ~ of female nonreceptivity
in another Lepidoptera, D. plexippus, is correlated with the initial size of the ~permatophore;~~
whether resumption of receptivity is cued by a threshold size of the absorbed spermatophore,
by decay of some factor transferred by the male whose volume is correlated with spermato-
phore size, or by changes in the female's nutritional status as a result of decreasing input of
spermatophore nutrients remains to be determined.
234 Insect Reproduction

Males make a nutrient donation to the female prior to the transfer of sperm in all known
cases, even if female utilization of that donation is delayed.30In Lepidoptera and other groups
with internal spermatophores, the spermatophore is passed to the female prior to sperm
movement; in Orthoptera, the sperm are often deposited in an external ampulla, and the
ampulla, spermatophylax, andor male body parts are available to the female prior to sperm
movement into the female's body; in Mecoptera and Diptera with nuptial prey or salivary gifts,
prey are presented to females prior to the onset of copulation. This uniformity among cases
where the nutrient value of the donation to the female varies suggests that selective pressures
associated with mate acquisition, complete insemination and mate guarding were crucial to the
evolution of nuptial gifts. That is, males that were able to present gifts first obtained matings
from females and also obtained complete inseminations. In some of these groups, then, female
choice of mates with an initial gift may have played an important role in the evolution of the
form and sequence of gift giving.
While mate acquisition, mate guarding, and security of paternity may have played a role
in the evolution of male nutrient donations, male nutrient donations may also function as the
selective context for changes in mating systems and sperm precedence patterns. In short, the
two sets of traits, mating systems and donation qualitylquantity, should be coadapted. In
species whose females mate several times, the timing of remating can be closely linked to the
timing of the decay of use of the previous male's nutrients in egg production. Female C.
eurytheme remate after 4 to 6 days in a field p o p ~ l a t i o nmale
; ~ ~ nutrients are primarily found
in eggs laid during the first 3 to 4 days after a mating.(j9 However, both R. verticalis and D.
verrucivorus females show a peak of incorporation of male nutrients into laid eggs much later
than the end of the female refractory period.70.85-146 Actual timing of female remating in the
field will depend on female access to mates as well as the length of the refractory period.146
Intermating intervals may thus be longer than the refractory period which was observed under
experimental conditions.
Differences not just in nutrient use as outlined above, but also in donation quality or
quantity, are expected for related species that differ in expected number of female matings,
sperm precedence patterns, andfor alternative mechanisms of mate guarding not involving the
male nutrient donation. Extensive data to test this do not yet exist. We do know that
spermatophore size is correlated with the duration of the intermating interval within some
species but not others, and that the species-specific time required to regain the ability to make
a normal sized spermatophore after a mating is correlated with mean number of matings in
some groups.
Time since the last mating by the male is positively correlated with spermatophore size in
several Lepidoptera.30~64~72~83-84-90+92
Further, male lepidopterans remate rapidly, increasing the
time in copula if mating has occured recently, whereas male orthopterans generally have a
post-mating refractory period, with subsequent spermatophore sizes unaffected. This suggests
differences between the two groups in mate acquisition strategies, with lepidopterans remating
whenever possible even if a small spermatophore results, whereas orthopteran males appear
to need the ability to make a minimum investment before mating will occur.78
Differences in the mean number of lifetime matings among lepidopteran species are
correlated with the rate of recovery of ability to produce a "normal" sized ~permatophore.~~
Four species from a diversity of families with an average number of matings greater than 2.0
were able to make a spermatophore equal in mass to that of the first mating within 2 to 3 days
after the first mating; individual males from three species with an average number of matings
less than 2.0 had not recovered the ability to make normal sized spermatophores even 5 days
post-mating. Further, male mating success is highly skewed in at least three species including
one with mean number of matings less than 2.0; in H. cydno, H. charitonius, and D. julia, less
than 15% of the males obtain half the m a t i n g ~ .This
' ~ ~ difference in recovery time, then, means
that a significant number of females which only mate once may be receiving small spermato-
phores without the opportunity to obtain more nutrients from males at a later time.
Phenotypic Consequences and Evolutionary Implications 235

E. INTERACTION OF NUPTIAL GIFTS WITH

AGE-SPECIFIC FECUNDITY: FECUNDITY ENHANCEMENT
Female intake of male-donated food, in whatever form, is a contribution by the male to the
female's resource budget.66Depending on subsequent allocation of that food by the female,
it may also be an investment in offspring; depending on paternity patterns, it may be an
investment in the donating male's offspring. Males can theoretically influence allocation
patterns by the type of nutrients they donate; nutrients which are in short supply for egg
production and which are usable with little modification should be allocated to offspring by
the female. Although the fitness of the male may also be enhanced if "his" nutrients are
allocated to female survival rather than invested in his offspring, this effect has never been
experimentally tested.
From a female perspective, the nuptial gift may be regarded as another food source, with
attendant risks and benefits which differ in detail from those of "ordinary" food sources.
Females should be expected to allocate male-donated resources in a manner consistent with
maximizing their own lifetime reproductive success. Thus, the possibility exists for conflict
in evolutionary time between the sexes over female allocation of male-derived nutrients.I5
Such conflicts may have molded the present-day size, composition and timing of male nutrient
donations, and the consequent role donations have in enhancing female fecundity.
Most work has focused on the end product - number of eggs produced - as the trait
affected by changes in available resources due to male nuptial gifts. However, other aspects
of reproductive resource allocation patterns may be coadapted with nuptial gift giving. That
is, the sources of resources and dynamics of their use in reproduction may be shifted in
expectation of a given level of nutritional intake by the female from the male. This can have
two manifestations: the first is allocation to reproductive reserves, and the second is ovarian
dynamics.
Holometabolous insects in particular have the luxury of reallocating resources during
metamorphosis to suit adult needs. BoggslOs predicted that allocation of larval resources to
body vs. reproductive reserves during metamorphosis should vary among species and sexes
as a function of the expected adult intake of nutrients, including nuptial gifts, and output of
nutrients in reproduction. Three species of heliconiine butterflies fit the predictions. These
species differ in the quality of the spermatophore received by the female and in the number
of matings. Thus, allocation of larval reserves to reproduction accomodates expected nuptial
gifts.
Boggs6Vurther suggests that the nutritional function of the male nuptial gift should be
matched with the female's ovarian dynamics. Females whose eggs are matured after adult
emergence have the opportunity to use male donations in egg production, rather than just for
somatic maintenance. Species with large nutrient donations thus are expected to yolk eggs
after adult eclosion.

F. NONINDEPENDENCE OF ADAPTIVE FUNCTION:

INTERACTIONS BETWEEN FECUNDITY ENHANCEMENT
AND MATING SYSTEMS EFFECTS OF NUPTIAL GIFTS
Nuptial gifts' interactions with mating systems can be affected by the interaction between
nuptial gifts and female reproduction, and vice versa. For example, once male nutrient
donations of whatever form exist, the size and quality of the donation should be affected by
selective pressures related to female and male resource budgets for reproduction, in the
context of security of paternity.1sJ4%e effectiveness of mechanisms delaying remating in
species with sperm precedence will constrain the possible sizelquality of the nutrient donation
through pressure on males not to provide nutrients to make another male's offspring. For
example, complete sperm precedence coupled with rapid female remating in particular should
select for males to donate only small quantities of rapidly utilizable nutrients to females. The
state of male and female resource budgets will depend on foraging efficiencies, food availability,
236 Insect Reproduction

and demands of survival expenses on the resource pool. The fecundity-enhancement function
of male nutrient donations thus should have been most strongly selected for in environments
which are resource poor, because of low prey or high predator densities or low quality diets,
and in species whose males have high confidence of paternity. Note that a species feeding on
a highly dispersed but high quality prey could still be considered to reside in a resource-poor
environment when compared with species feeding on less dispersed prey items. By this
criteria, many parasites, including bedbugs, could be considered to reside in resource-poor
environments. Quality of the resource environment should also put a constraint on the
evolution of the fecundity-enhancement function of male nutrient donations, however. Nutri-
tional environments which are very poor or risky should not be able to support the resource
pools needed by males to make large donations. Tallamy" presents a variant of this hypoth-
esis, including both prezygotic and postzygotic paternal reproductive expenditures. He pro-
vides extensive supporting evidence from a wide variety of arthropods.
Male nuptial gifts may thus have mixed functions, both as resource and mating investments
- but only from the male's perspective. From the female's perspective, the sole role of the
nutrient donation is as investment in her resource budget; what differs is the size of the
investment, its possible uses, and the possibility of exercising choice among males with
different quality gifts.
The observed relative importance of the two functions should depend on the following
factors. Timing of oogenesis relative to mating and certainty of paternity should affect
elaboration of the role of gifts as investment in the female's resource budget (from a male
viewpoint). The ability of the female to terminate mating before sperm transfer is complete,
the alternative behavioral or hormonal methods of mate guarding, the ability of the male to
mate with unwilling females, and the availability of other female mates should all affect
elaboration of the role of gifts as mating investment.
Within Orthoptera, we now know that spermatophores can function primarily as a mating
investment guaranteeing complete sperm transfer, or have an additional primary function as
an investment in the female's resource budget. Conclusions are based on size of the
spermatophylax relative to that needed to ensure complete sperm transfer, protein content of
the spermatophylax, and the sensitivity of its size to male diet.99-'22-124
Similar studies have not
yet been done in other groups.

VII. FUTURE PROSPECTS

Male nuptial gifts have primarily served as a case example to study theories of sexual
selection. The focus has been testing hypotheses concerning the effect of the balance of
investment between the two sexes on sex roles and the operational sex ratio. Work in this area
should continue to expand, with a better understanding of causes of mate competition and mate
choice and inclusion of study of the physiological and genetic underpinnings of observed
behaviors.
Study of male nuptial gifts, however, has broader potential than just illuminating sexual
selection theory. Like many other reproductive traits, male donations have effects at organi-
zational levels ranging from individual physiology through behavior to population demography.
As discussed above, effects of size or frequency of gift donation, for example, can impact the
male's resource budget, affecting courting intensity, the operational sex ratio, sex roles,
survival, and effective population size. Nuptial gifts also have the potential to affect the
evolutionary pathway of traits within a population or of whole phylogenetic groups of
organisms through direct effects on fitness or by interacting with other traits to affect fitness.
Thus, we may expect to see suites of traits associated with a particular pattern of male nuptial
gift giving in the context of a particular resource environment and constraints imposed by
phylogenetic history. The degree of variability in the resource environment, the relative
harshness of the environment, and the level of survival risk associated with obtaining resources
Phenotypic Consequences and Evolutionary Implications 237

should be important environmental descriptors in this case. As a corollary, we should only see
some combinations of traits as a transient state in evolutionary time, if at all. Traits at issue
here include the size and composition of the nuptial gift, the timing of oogenesis, the mean
and variance in number of matings by each sex, mate competition, mate choice, allocation of
male-derived resources by females, time and allocation costs to males of mating, and age at
maturity for each sex. Some of these traits will determine other traits considered above. For
example, the mean and variance in number of matings, combined with total population size,
determines the population effective population size.
In particular, male nuptial gifts provide a nearly unique opportunity to examine emerging
ideas concerning resource allocation. Since male nutrient donations are an allocation to
reproduction by the male, but a meal to the female, they are part of the life history strategy
of one sex, but the foraging strategy, broad sense, of the other. Resource allocation links
foraging and life history allocation; the study of male nutrient donations will allow us to link
not only foraging and allocation, but the life strategies (sensu Gatto et of both sexes.
Other possibilities abound as well. Internal spermatophores are a form of "stored" nutrients
for the female. Questions concerning the use of available vs. stored nutrients as a function of
environmental food availability can be addressed in this system. Questions concerning the
effects of variation in availability of different nutrient types at different times in the life cycle
can be addressed as well. With the expansion of groups in which male nutrient donations have
been studied, we can begin to examine the role of phylogenetic history in constraining
allocations related to male nutrient donations, thus limiting suites of traits observed in nature.
We should also be able to explore circumstances under which convergent evolution is more
or less likely to occur.

ACKNOWLEDGMENTS
I thank P. Grandcolas, D. Gwynne, J. McNeil, D. Mullins, C. Nalepa, K. Oberhauser, S.
Sakaluk, L. Simmons, L. Svard, D. Tallamy, and N. Wedell for preprints or discussion, and
R. Rutowski and W. Watt for comments on the manuscript.

REFERENCES
1. Downes, J. A., Feeding and mating in the insectivorousCeratopogoninae (Diptera),Mem. Entomol. Soc. Can.,
104, 1978.
2. Morris, G. K., Mating systems, paternal investment and aggressive behavior of acoustic Orthoptera, Flu.
Entomol., 62, 9, 1979.
3. Bell, P. D., Opportunistic feeding by the female tree cricket, Oecanthus nigricornis (Orthoptera: Gryllidae),
Can. Entomol., 112, 432, 1980.
4. Bidochka, B. J. and Snedden, W. A., Effect of nuptial feeding on the mating behaviour of female ground
crickets, Can. J. Zool., 63, 207, 1985.
5. Thornhill, R., Sexual selection and nuptial feeding behavior in Bittacus apicalis (Insecta: Mecopten), Am.
Nut., 110, 529, 1976.
6. Thornhill, R. and Alcock, J., The Evolution of Insect Mating Systems, Harvard University Press, Cambridge,
1983.
7. Svensson, B. G. and Petersson, E., Sex-role reversed courtship behaviour, sexual dimorphism and nuptial
gifts in the dance fly, Empis borealis (L.),Ann. Zool. Fennici, 24, 323, 1987.
8. Boggs, C. L. and Gilbert, L. E., Male contribution to egg production in butterflies: evidence for transfer of
nutrients at mating, Science, 206, 83, 1979.
9. Smith, R. L., Evolution of exclusive postcopulatory paternal care in the insects, Flu. Entomol., 63.65, 1980.
10. Nalepa, C. A. and Jones, S. C., Evolution of monogamy in termites, Biol. Rev., 66, 83, 1991.
I I. Tallamy, D. W., Nourishment and the evolution of paternal investment in subsocial arthropods, in Nourisll-
ment and Evolution in Insect Societies, Hunt, J . and Nalepa, C., Eds., Westview Press. Boulder, 1994.
238 Insect Reproduction

12. Thornhill, R., Sexual selection and paternal investment in insects, Am. Nut., 110, 153, 1976.
13. Zeh, D. W. and Smith, R. L., Paternal investment by terrestrial arthropods, Am. Zool., 25, 785, 1985.
14. Ridley, M., Mating frequency and fecundity in insects, Biol. Rev., 63, 509, 1988.
15. Parker, G. A. and Simmons, L. W., Nuptial feeding in insects: theoretical models of male and female
interests, Ethology, 82, 3, 1989.
16. Thornhill, R., Adaptive female-mimicking behavior in a scorpionfly, Science, 205, 412, 1979.
17. Gwynne, D. T., Nuptial feeding behaviour and female choice of mates in Harpobittacus similis (Mecoptera:
Bittacidae), J. Aust. Entomol. Soc., 23, 271, 1984.
18. Downes, J. A., The feeding and mating behaviour of the specialized Empidinae (Diptera): observations on
four species of Rhamphomyia in the high arctic and general discussion, Can. Entomol., 102, 769, 1970.
19. Kessel, E. L., The mating activities of balloon flies, Syst. Zool., 4, 97, 1955.
20. Thornhill, R., Panorpa (Mecopten: Panorpidae) scorpionflies: systems for understanding resource-defense
polygyny and alternative male reproductive efforts, Ann. Rev. Ecol. Syst., 12, 355, 1981.
21. Steele, R. H., Courtship feeding in Drosophila subobscura. I. The nutritional significance of courtship
feeding, Anim. Behav., 34, 1087, 1986.
22. Leopold, R. A., The role of male accessory glands in insect reproduction, Annu. Rev. Entomol., 199, 1976.
23. Chen, P. S., The functional morphology and biochemistry of insect male accessory glands and their secre-
tions, Annu. Rev. Entomol., 29, 233, 1984.
24. Kasuga, H., Aigaki, T. and Osani, M., System for supply of free arginine in the spermatophoreof Bombyx
mori: arginine liberating activities of contents of male reproductive glands, Insect Biochem., 17, 317, 1987.
25. Cheeseman, M. T. and Gillott, C., Long hyaline gland discharge and multiple spermatophore formation by
the male grasshopper, Melanoplus sanguinipes, Physiol. Entomol., 14, 257, 1989.
26. Friedel, T. and Gillott, C., Contribution of male-produced proteins to vitellogenesisin Melanoplus sanguinipes,
J . Insect Physiol., 23, 145, 1977.
27. Osani, M., Aigaki, T., Kasuga, H. and Yonezawa, Y., Role of arginase transferred from the vesicula
serninalis during mating and changes in amino acid pools of the spermatophore after ejaculation in the
silkworm, Bombyx mori, Insect Biochem., 16, 879, 1986.
28. Dickinson, J. L. and Rutowski, R. L. 1989. The function of the mating plug in the chalcedon checkerspot
butterfly, Anim. Behav., 38, 154, 1989.
29. Davey, K. G., Reproduction in the Insects, Freeman, San Francisco, 1965.
30. Rutowski, R. L. and Gilchrist, G. W., Copulation in Colias eurytheme (Lepidoptera: Pieridae): patterns and
frequency, J. Zool. London (A), 209, 115, 1986.
31. Markow, T. A., Drosophila males provide a material contribution to offspring sired by other males, Funct.
Ecol., 2, 77, 1988.
32. Gerber, G. H., Evolution of the methods of spermatophore formation in pterygotan insects, Can. Entomol.,
102,358, 1970.
33. Rutowski, R. L., Newton, M., and Schaefer, J., lnterspecificvariation in the size of the nutrient investment
made by male butterflies during copulation, Evolution, 37, 708, 1983.
34. Boucher, L. and Huignard, J., Transfer of male secretions from the spermatophore to the female insect in
Caryedon serratus (01.): analysis of the possible trophic role of these secretions. J . Insect Physiol., 33,949,
1987.
35. Svard, L. and Wiklund, C., Mass and production rate of ejaculates in relation to monandrylpolyandry in
butterflies, Behav. Ecol. Sociobiol., 24, 395, 1989.
36. Gwynne, D. T., Sexual difference theory: mormon crickets show role reversal in mate choice, Science, 213,
779, 1981.
37. Simmons, L. W., Male size, mating potential and lifetime reproductive succes in the field cricket, Gryllus
bimaculatus (De Geer), Anim. Behav., 36, 372, 1988.
38. Hayashi, F., Male mating costs in two insect species (Protohermes, Megaloptera) that produce large
spermatophores, Anim. Behav., 45, 343, 1993.
39. Alexander, R. D. and Otte, D., The evolution of genitalia and mating behavior in crickets (Gryllidae) and
other Orthoptera, Misc. Publ. Mus. Zool. Univ. Mich., 133, 1967.
40. Sakaluk, S. K. and Cade, W. H., Female mating frequency and progeny production in singly and doubly
mated house and field crickets, Can. J. Zool., 58, 404, 1980.
41. Evans, A. R., Mating systems and reproductive strategiesin three australian gryllid crickets: Bobilla victoriae
Otte, Balamara gidya Otte and Teleogryllus cornmodus (Walker) (Orthopten: Gryllidae: Nemobiinae;
Trigonidiinae; Gryllinae), Ethology, 78, 21, 1988.
42. Mays, D. L., Mating behaviour of Nemobiinae crickets Hygronemobius, Nemobius and Pteronernobius
(Orthoptera: Gryllidae), Flu. Entomol., 54, 13, 1971.
43. Davey, K. G., The evolution of the spermatophore in insects, Proc. R. Entomol. Soc. London Ser. A, 35, 107,
1960.
44. Marshall, L. D., Male nutrient investment in the Lepidoptera: what nutrients should males invest?, Am. Nut.,
120,273, 1980.
Phenotypic Consequences and Evolutionary Implications 239

45. Engebretson, J. A, and Mason, W. H., Transfer of 65Zn at mating in Heliothis virescens, Environ. Entomol.,
9, 119, 1980.
46. Pivnick, K. A. and McNeil, J. N. Puddling in butterflies: sodium affects reproductive success in Thymelicus
lineola, Physiol. Entomol., 12, 461, 1987.
47. Adler, P. H. and Pearson, D. L., Why do male butterflies visit mud puddles? Can. J. 2201.. 60,322, 1982.
48. Arms, K., Feeny, P., and Lederhouse, R. C., Sodium: stimulus for puddling behavior by tiger swallowtail
butterflies, Science, 185, 372, 1974.
49. Boggs, C. L. and Jackson, L. A., Mud puddling by butterflies is not a simple matter, Ecol. Entomol., 16, 123,
1991.
50. Sculley, C. and Boggs, C.L. unpublished manuscript.
51. Mullins, D. E. and Keil, C. B., Paternal investment of urates in cockroaches, Nature, 283, 567, 1980.
52. Schal, C. and Bell, W. J., Ecological correlates of paternal investment of urates in a tropical cockroach,
Science, 218, 170, 1982.
53. Mullins, D. E., Keil, C. B., and White, R. H., Maternal and paternal nitrogen investment in Blattella
germanica (L.) (Dictyoptera; Blattellidae), J. Exp. Biol., 162, 55, 1992.
54. Sierra, J. R., Woogon, W.-D., and Schmid, H., Transfer of cantharidin during copulation from the adult
male to the female Lytta vesicatoria ("Spanish flies"), Experientia, 32, 142, 1976.
55. Alcock, J. and Hadley, N. F., Tests of alternative hypotheses for assortative mating by size: a comparison
of two meloid beetles (Coleoptera: Meloidae), J. Kans. Entomol. Soc., 60, 41, 1987.
56. Achey unpublished, cited in Brower, L. P., New perspective on the migration biology of the monarch
butterfly, Danaus plexippus L. in Migration: Mechanisms and Adaptive Signipcance, Rankin, M . A., Ed.,
Univ. of Texas Contributions in Marine Science, Austin, TX, 1985, 748.
57. Brown, K. S., Adult-obtained pyrrolizidine alkaloids defend ithomiine butterflies against a spider predator,
Nature, 309, 707, 1984.
58. Dussourd, D. E., Ubik, K., Harris, C., Resch, J., Meinwald, J., and Eisner, T., Biparental defensive
endowment of eggs with acquired plant alkaloid in the moth Utethesia ornatrix, Proc. Natl. Acad. Sci. U.S.A.,
85, 992, 1988.
59. Marshall, L. D. and McNeil, J. N., Spermatophore mass as an estimate of male nutrient investment: a closer
look in Pseudaletia unipuncta (Haworth) (Lepidopten: Noctuidae), Funct. Ecol., 3, 605, 1989.
60. Buskirk, R. E., Frohlich, C., and Ross, K. G., The natural selection of sexual cannibalism, Am. Nat., 123,
612, 1984.
61. Morris, G. K., Gwynne, D. T., Klimas, D. E., and Sakaluk, S. K., Virgin male mating advantage in a
primitive acoustic insect (Orthoptera: Haglidae), J. Insect Behav., 2, 173, 1989.
62. Hinton, H. E., Sperm transfer in insects and the evolution of haemocoelic insemination,in Insect Reproduc-
tion. Symposia of the Royal Entomological Society, London, Highnam, K . C., Ed., Royal Ent. Soc. London,
2, 95, 1964.
63. Silbergleid, R. E., Shepherd, J. G., and Dickinson, J. L., Eunuchs: the role of apyrene sperm in Lepi-
dopten?, Am. Nat., 123, 255, 1984.
64. Boggs, C. L., Selection pressures affecting male nutrient investment at mating in heliconiine butterflies,
Evolution, 35, 931, 1981.
65. Gwynne, D. T., Courtship feeding and the fitness of female katydids (Orthoptera: Tettigoniidae),Evolution,
42, 545, 1988.
66. Boggs, C. L., A general model of the role of male-donated nutrients in female insects' reproduction, Am. Nat.,
136,598, 1990.
67. Wedell, N., Mating effort or paternal investment: incorporation rate and cost of male donations in the
wartbiter. Behav. Ecol. Sociobiol., 32, 239, 1993.
68. Greenfield, M. D., The question of paternal investment in Lepidoptera: male-contributed proteins in Plodia
interpunctella, Int. J. Invert. Repro., 5, 323, 1982.
69. Boggs, C. L. and Watt, W. B., Population structure of pierid butterflies. IV. Genetic and physiological
investment in offspring by male Colias, Oecologia, 50, 320, 1981.
70. Bowen, B. J., Codd, C. G., and Gwynne, D. T., The katydid spermatophore (Orthoptem: Tettigoniidae):
male nutritional investment and its fate in the mated female, Aust. J. Zool., 32, 23, 1984.
71. Oberhauser, K. S., Rate of ejaculate breakdown and intermating intervals in monarch butterflies, Behav.
Ecol. Sociobiol., 31, 367, 1992.
72. Rutowski, R. L., Production and use of secretions passed by males at copulation in Pieris protodice
(Lepidopten, Pieridae), Psyche, 91, 141, 1984.
73. Huignard, J., Transfer and fate of male secretions deposited in the spennatophore of females ofAcanthoscelides
obtectus Say (Coleoptera Bruchidae), J. Insect Physiol., 29, 55, 1983.
74. Khalifa, A., Spermatophore production in Trichoptera and some other insects, Trans. Entomol. Soc. Lond.,
100, 18, 1949.
75. Svard, L. and Wiklund, C., Fecundity, egg weight and longevity in relation to multiple matings in females
of the monarch butterfly, Behav. Ecol. Sociobiol., 23, 39, 1988.
240 Insect Reproduction

76. Sakaluk, S. K. and Snedden, W. A., Nightly calling durations of male sagebrush crickets, Cyphoderris
strepirans: size, mating and seasonal effects, Oikos, 57, 153, 1990.
77. Simmons, L. W., Teale, R. J., Maier, M., Standish, R. J., Bailey, W. J., and Wither, P. C., Some costs
of reproduction for male bushcrickets. Requena verticalis (Orthoptera: Tettigoniidae): allocating resources to
mate attraction and nuptial feeding, Behav. Ecol. Sociobiol., 31, 57, 1992.
78. Gwynne, D. T., Testing parental investment and the control of sexual selection in katydids: the operational
sex ratio, Am. Nar., 136, 474, 1990.
79. Oberhauser, K. S., Effects of spermatophores on male and female monarch butterfly reproductive success,
Behav. Ecol. Sociobiol., 25, 237, 1989.
80. Rutowski, R. L., The butterfly as an honest salesman, Anim. Behav., 27, 1269, 1979.
81. Burpee, D. M. and Sakaluk, S. K., Repeated matings offset costs of reproduction in female crickets, Evol.
Ecol., 7, 1, 1993.
82. Sakaluk, S. K., Spermatophore size and its role in the reproductive behaviour of the cricket, Gryllodes
supplicans (Orthoptera: Gryllidae), Can. J. Zool., 63, 1652, 1985.
83. Svard, L. and Wiklund, C., Different ejaculate delivery strategies at first versus subsequent matings in the
swallowtail butterfly, Papilio machaon L., Behav. Ecol. Sociobiol., 18, 325, 1986.
84. Royer, L. and McNeil, J. N., Male investment in the European corn borer, Osrrinia nubilalis (Hiibner)
(Lepidoptera: Pyralidae): Impact on female longevity and reproductive performance, Funct. Ecol., 7, 209,
1993.
85. Sakaluk, S. K. and Smith, R. L., Inheritance of male parental investment in an insect, Am. Nat.. 132, 594,
1988.
86. Sakaluk, S. K., Burpee, D. M., and Smith, R. L., Phenotypic and genetic variation in the stridulatory organs
of male decorated crickets, Gryllodes sigillatus (Orthoptera: Gryllidae), Can. J . Zool., 70, 453, 1992.
87. Gwynne, D. T., Bowen, B. J., and Codd, C. G., The function of the katydid spermatophore and its role in
fecundity and insemination (Orthoptera: Tettigoniidae), Ausr. J. Zool., 32, 15, 1984.
88. Gwynne, D. T., Mate selection by female katydids (Orthoptera: Tettigoniidae, Conocephalus nigropleurum),
Anim. Behav., 30, 734, 1982.
89. Wedell, N., Protandry and mate assessment in the wartbiter Decticus verrucivorus (Orthoptera: Tettigoniidae),
Behav. Ecol. Sociobiol., 31, 301, 1992.
90. Svard, L., Parental investment in a monandrous butterfly, Parargae aegeria, Oikos, 45, 66, 1985.
91. Reiss, M. J., The allometry of reproduction: why larger species invest relatively less in their offspring, J.
Theor. Biol., 113. 529, 1985.
92. Oberhauser, K. S., Male monarch butterfly spermatophore mass and mating strategies, Anim. Behav., 36,
1384, 1988.
93. George, J. A. and Howard, M. G., Insemination without spermatophores in the oriental fruit moth,
Grapholirha molesta (Lepidoptera: Tortricidae), Can. Enromol., 100, 190, 1964.
94. Zuk, M., The effects of gregarine parasites, body size, and time of day on spermatophore production and
sexual selection in field crickets, Behav. Ecol. Sociobiol., 21, 65, 1987.
95. Simmons, L. W., Some constraints on reproduction for male bushcrickets, Requena verticalis (Orthoptera:
Tettigoniidae): diet, size and parasite load, Behav. Ecol. Sociobiol., 32, 135, 1993.
96. Butlin, R. K., Woodhatch, C. W., and Hewitt, G. M., Male spermatophore investment increases female
fecundity in a grasshopper, Evolution, 41, 22 1, 1987.
97. Simmons, L. W. and Bailey, W. J., Resource influenced sex roles of zaprochiline tettigoniids (Orthoptera:
Tettigoniidae), Evolution, 44, 1853, 1990.
98. Wedell, N. and Arak, A., The wartbiter spermatophore and its effect on female reproductive output
(Orthoptera: Tettigoniidae, Decticus verrucivorus), Behav. Ecol. Sociobiol., 24, 117, 1989.
99. Wedell, N., Sperm competition selects for nuptial feeding in a bushcricket, Evolution, 45, 1975, 1991.
100. Rutowski, R. L., Gilchrist, G. W., and Terkanian, B., Female butterflies mated with recently mated males
show reduced reproductive output, Behav. Ecol. Sociobiol., 20, 319, 1987.
101. Markow, T. A., Gallagher, P. D., and Krebs, R. A., Ejaculate-derived nutritional contribution and female
reproductive success in Drosophila mojavensis (Patterson & Crow), Funct. Ecol., 4, 67, 1990.
102. Gwynne, D. T., Male mating effort, confidence of paternity and insect sperm competition, in Sperm
competition and the Evolution ofAnimal Mating Systems, Smith, R. L., Ed.,Academic Press, New York, 117,
1984.
103. Tuomi, J., Hakala, T., and Haukioja, E., Alternative concepts of reproductive effort, costs of reproduction
and selection in life-history evolution, Am. Zool., 23, 25, 1983.
104. Woods, H. A., Boggs, C. L., and Karlsson, B., unpublished manuscript.
105. Svard, L. and McNeil, J. N., Female benefit, male risk: Polyandry in the true armyworm Pseudaletia
unipuncta, Behav. Ecol. Sociobiol. 35, 319, 1994.
106. Boggs, C. L. and Ross, C. L., The effect of adult food limitation on life history traits in Speyeria mormonia
(Lepidoptera: Nymphalidae), Ecology, 74, 433, 1993.
Phenotypic Consequences and Evolutionary Implications 241

107. Gwynne, D. T. and Dodson, G. N., Nonrandom provisioning by the digger wasp, Palmodes laeviventris
(Hymenoptera: Sphecidae), Ann. Entomol. Soc. Am., 76, 434, 1983.
108. Boggs, C. L., Nutritional and life-history determinants of resource allocation in holometabolous insects, Am.
Nat., 117, 692, 1981.
109. Zalucki, M. P., Sex around the milkweed patch - the significance of patches of host plants in monarch
reproduction, in Biology and Conservation of the Monarch Butrerfy. Malcolm, S. B. and Zalucki, M. P,, Eds.,
Natural History Museum of Los Angeles County, Los Angeles, 69, 1993.
110. Brower, L. P., New perspective on the migration biology of the monarch butterfly, Danaus plexippus L. in
Migration: Mechanisms and Adaptive Significance, Rankin, M. A., Ed., University of Texas Contributions in
Marine Science, Austin, TX, 748, 1985.
11 1. van Hook, T., Non-random mating in monarch butterflies overwintering in Mexico, in Biology and Conser-
vation of the Monarch Butterfy, Malcolm, S. B. and Zalucki, M. P,, Eds., Natural History Museum of Los
Angeles County, Los Angeles, 49, 1993.
112. Wells, H., Wells, P. H., and Rogers, S. H., Is multiple mating an adaptive feature of monarch butterfly winter
aggregation?, in Biology and Conservation of the Monarch Butrefly, Malcolm, S. B. and Zalucki, M. P,, Eds.,
Natural History Museum of Los Angeles County, Los Angeles, 61, 1993.
113. Clutton-Brock, T. H., Reproductive Success: Studies of Individual Variation in Contrasting Breeding
Systems, University of Chicago Press, Chicago, 1988.
114. Pitnick, S., Markow, T. A., and Riedy, M. F., Transfer of ejaculate and incorporation of male-derived
substances by females in the Nannoptera species group (Dipten: Drosophilidae), Evolution, 45, 774, 1991.
115. Boldyrev, B. T., Contributions a I'ttude de las structure des spermatophores et des particularites de la
copulation chez Locustodea et Gryllodea, Horae Soc. Entomol. Ross., 41, 1, 1915.
116. Gwynne, D. T., Male nutritional investment and the evolution of sexual differences in the Tettigoniidae and
other Orthoptera, in Orthopteran Mating Systems: Sexual Competition in a Diverse Group of Insects, Gwynne,
D. T. and Mol~is,G., Eds., Westview Press, Boulder, 337, 1983.
117. Trivers, R. L., Parental investment and sexual selection, in Sexual Selection and the Descent of Man,
Campbell, B., Ed., Aldine-Athelton, Chicago, 1972.
118. Darwin, C., The Descent of Man and Selection in Relation to Sex, John Murray, London, 1871.
119. Fisher, R. A., The Genetical Theory of Natural Selection, Oxford University Press, Oxford, 1930.
120. Baternan, A. J., Intra-sexual selection in Drosophila, Heredity, 2, 349, 1948.
121. Williarns, G. C., Adaptation and Natural Selection, Princeton University Press, Princeton, NJ, 1966.
122. Sakaluk, S. K., Male crickets feed females to ensure complete sperm transfer, Science, 223, 609, 1984.
123. Gwynne, D. T., Courtship feeding in katydids (Orthoptera: Tettigoniidae): investment in offspring or in
obtaining fertilizations?, Am. Nar., 128, 342, 1986.
124. Reinhold, K. and Heller, K.-G., The ultimate function of nuptial feeding in the bushcricket Piecilimon
veluchianus (Orthoptera: Tettigoniidae: Phaneropterinae), Behav. Ecol. Sociobiol., 32, 55, 1993.
125. Alexander, R. D. and Borgia, G., On the origin and basis of the male-female phenomenon, in Sexual
Selection and Reproductive Competition in Insects, Blurn, M. S. and Blum, N. A., Eds., Academic Press, New
York, 1979.
126. Low, B. S., Environmental uncertainties and the parental strategies of marsupials and placentals, Am. Not.,
112, 197, 1978.
127. Quinn, J. S. and Sakaluk, S. K., Prezygotic male reproductive effort in insects: why do males provide more
than sperm?, Fla. Entomol., 69, 84, 1986.
128. Emlen, S. T. and Oring, L. W., Ecology, sexual selection, and the evolution of mating systems, Science, 197,
215, 1977.
129. Gwynne, D. T., Role-reversal in katydids: habitat influences reproductive behaviour (Orthoptera: Tettigoniidae,
Metaballus sp.), Behav. Ecol. Sociobiol., 16, 355, 1985.
130. Clutton-Brock, T. H. and Parker, G. A., Potential reproductive ntes and the operation of sexual selection,
Q. Rev. Biol., 67, 437, 1992.
131. Gilbert, L. E., Postmating female odor in Heliconius butterflies: a male-contributed antiaphrodisiac?,
Science, 193, 419, 1976.
132. Rutowski, R. L., The form and function of ascending flights in Colias butterflies, Behav. Ecol. Sociobiol.,
3, 163, 1978.
133. Rutowski, R. L., Evidence for mate choice in a sulphur butterfly (Colias eurytheme), Z. Tierpsychol., 70, 103,
1985.
134. Watt, W. B., Carter, P. A., and Donohue, K., Females' choice of "good genotypes" as mates is promoted
by an insect mating system, Science, 233, 1187, 1986.
135. Borgia, G., Sexual selection and the evolution of mating systems, in Sexual Selection and Reproductive
Competition in Insects, Blum, M . S. and Blum, N. A., Eds., Academic Press, New York, 19, 1979.
136. Gwynne, D. T. and Sirnmons, L. W., Experimental reversal of courtship roles in an insect, Nature, 346, 172,
1990.
242 Insect Reproduction

137. Gwynne, D. T., Sexual selection and sexual differences in mormon crickets (Orthoptera: Tettigoniidae,
Anabrus simplex), Evolulion, 38, 1011, 1984.
138. Rutowski, R. L., Courtship solicitation by females of the checkered white butterfly, Pieris prorodice, Behav.
Ecol. Sociobiol., 7, 113, 1980.
139. Rutowski, R. L., Long, C. E., Marshall, L. D., and Vetter, R. S., Courtship solicitation by Colias females
(Lepidoptera: Pieridae), Am. Midl. Nat., 105, 334, 1981.
140. Simmons, L. W., Female choice in the field cricket Gryllus bimaculatus (De Geer), Anim. Behav., 34, 1463,
1986.
141. Boggs, C. L., Resource allocation and reproductive strategies in several heliconiine species, Ph.D. disserta-
tion, University of Texas at Austin, 1979.
142. Boggs, C. L., personal observation.
143. Watt, W. B., Eggs, enzymes, and evolution: natural genetic variants change insect fecundity, Proc. Nad.
Acad. Sci. U.S.A., 89, 10608, 1992.
144. Rutowski, R. L., Epigarnic selection by males as evidenced by courtship partner preferences in the checkered
white butterfly (Pieris prorodice), Anim. Behuv., 30, 108, 1982.
145. Sugawara, T., Stretch reception in the bursa copulatrix of the butterfly, Pieris rapae crucivora, and its role
in behaviour, J. Comp. Physiol., 130, 191, 1979.
146. Gwynne, D. T., Courtship feeding in katydids benefits the mating male's offspring, Behav. Ecol. Sociobiol.,
23, 373, 1988.
147. Boggs, C. L., unpublished data.
148. Markow, T. A. and Ankney, P. F., Drosophila males contribute to oogenesis in a multiple mating species,
Science, 224, 302, 1984.
149. Gatto, M., Matessi, C., and Slobodkin, L. B., Physiological profiles and demographic rates in relation to
food quantity and predictability: an optimization approach, Evol. Ecol., 3, 1, 1989.
 Anthocoris confusus, 148
Anthonomus grandis, 7
Acanthoscelides obtectus, 47, 221 Anticarsia gemmatalis, 156
Accessory glands, 8-9, 3 6 , 4 8 4 9 Ants, 15
endocrine regulation, 10,38,40-42, 95 Aphididae
FES, 46 gonadal growth, 134-135
mating effects, 42 hormones, 99-101
nonspermatophore secretions, 217 parthenogenesis, 133-1 34
nuptial gifts, 216, 221 sex determination, 73
opaque, 47-48 Aphids. See Aphididae
spermatophores. See Spermatophores Aphis fabae, 100, 146, 157
Acheta domesticus, 10, 41 Apis mellifera, 44, 77, 78
Achieved fecundity, 144, 152 Aplysia californica, 45
Acrididae, 5 Apomictic parthenogenesis, 131, 132-133
Acridinae, 8 Apyrene sperm, 3 9 4 0 , 4 8 , 102, 219
Acyrthosiphon pisum, 133, 155 Aquatic insects, 8
Adenotrophic viviparity, 121 Archaeognatha, 2
Adipokinetic hormone, 22 Arctium minus, 162
Adoxophyes orana, 153 Armyworm. See Mamestra brassicae;
Adult feeding, 147, 151 Pseudaletia; Spodoptera
Adult size, 145-147, 175, 187 Artemia, 132
and larval habitat, 21 1 Asclepias, 160
swarm mating success, 203-21 1 Asparagus aphid, 150
Aedeagus, 36 Astegopteryx styracicola, 137
Aedes Attractants, 120, 121, 159
aegypti, 19, 21 Autocidal control techniques, 110
FES/RIS, 4 5 , 4 6 4 7 Automictic parthenogenesis, 131, 132-133
hormones, 97-98 Autosterilization, 118-1 23
landmarks, 202-203
oocyte development, 20,25
sex determination, 80
SIT, 113 Bacterial endosymbionts, 218
spermatheca, 7 Battus philenor, 164
albimanus, 116 Bedbugs, 71, 219
atropalpus, 98 Bees, 4, 7, 22, 44, 77, 78
punctor, 144 Beetles, 7, 18, 145-146
Aestivation, 151 Behavior, 120, 123
Age, of mating, 151-152 Bembidion, 77
Age structure, 226 Bemisia tabaci, 72
Alary polymorphism, 156, 157 Betula, 151
Alderflies, 14. See also Megaloptera Bibionidae, 199, 201
Allatostatins, 96 Biological insect control, 109-124
Amphorophora idaei, 133 Bird cherry aphid. See Rhopafosiphum padi
Ampulla, 218, 229, 233 Blackflies, 8 1
Anabrus simplex, 232 Black swallowtail butterfly, 153
Anagasra kuhniella, 34, 35 Bfatrella germanica. See also Cockroaches
Anarete pritchardii, 203 chorionization, 26
Anopheles, 80 germ cells, 10
quadrimaculatus, 1 12-1 13 male reproductive system, 41
Antennae, 201 nuptial gifts, 218, 221
Anrheraea polylphemus, 18 oogenesis, 20, 22, 24
Anthocharis cardamines, 188 pheromones, 101
 Insect Reproduction

Blattodea, 5, 68 Chitin synthesis inhibitors, 121, 122

Blowfly, 84, 113, 116-1 18, 21 1 Chorion, 2, 15.25-28
Body weight, 220. See also Adult size Choristonerrra
Bombyx mori fumiferana, 147, 150, 159
male reproductive system, 38, 39, 42, 46, 48 murinana, 148
mate acquisition, 176 Chorthippus
oogenesis, 12 brunneus, 164-1 65, 224
pheromones, 101 curtipennis, 8
sex determination, 78-79 parallelus, 164-1 65
sperm release, 103 Chromosome translocations, as control
Brachycaudus helichrysi, 155 technique, 115
Brachycorynella asparagi, 150 Chrysomela knabi, 145-146
Brevicoryne brassicae, 149, 151 Chrysomya rujifacies, 83, 84
Brine shrimp, l32 Chrysopa perla, 12
Bruchid beetle, 161 Chrysoperla carnea, 148
Bryophenocladius v e m l i s , 206-207 Cimex lectularius, 7 1
Bupalus piniarius, 146 Cinnabar moth, 150, 160
Bursa copulatrix, 7,44, 233 Circadian rhythm, 40, 44
Butterflies. See Lepidoptera Clathrin, 23, 25
Clitumnus extadentatus, 5
C Cluster-laying, 164
Clutch size, 163
Cabbage aphid, 149, 151 Coccinea, 13, 58, 72, 150
Cactoblastis cactorum, 161 Cochliomyia hominivorax, 45, 113-1 15
Cadra cautella, 145, 148 Cockroaches, 5. See also Blattella germanica
Caetifera, 4, 8 accessory glands, 8
Calliphora chorionization, 28
crowding, 21 1 milk glands, 10
erythrocephala, 84 nuptial gifts, 218
rujifacies, 84 oogenesis, 96
vomitoria, 98 oviposition, 28
Callosobruchus maculatus, 161 pheromones, 101
Calpodes ethlius, 26, 34 sex determination, 68
Cannibalism, 216, 219, 224 spermatheca, 7
Carausius morosus, 69 unreceptive stimuli, 44
Carbohydrates, 20 vitellogenin uptake, 23
Caryedon serratus, 224 Codling moth, 102, 147, 153, 155
Cassia mimosoides, 157-1 58 Coleoptera, 2, 6
Cavariella, 137 male reproductive system, 36, 39, 43
CCSF (corpus cardiacum-stimulating factor), nuptial gifts, 217, 218, 222, 224
97 oogenesis, 11, 14, 19, 21, 22
Cerajocera tussilaginis, 162 sex determination, 60, 63, 75-77, 86
Ceratitis spermatheca, 7
capitata, 83, 84, 113, 116 Colias
Chagasia bathana, 80 eurytheme, 221,222,224,232, 234
Chaoboridae, 199 hyale, 188
Chilo partellus, 42, 163 mate selection, 23 1, 233
Chironomidae, 199, 200, 203, 208 palaeno, 188
Chironomus philodice, 232
crowding, 21 I eriphyle, 144, 218
oppositus, 80, 8 1 eurytheme, 147, 148
plumosus, 36 Collateral glands, 36
size and mating success, 207, 208 Collembola, 67
swarm size, 207-2 11 Colleterial glands, 2, 7, 8-9
wing length and muscle mass, 204-205 Colophina, 137
 Competition Defensive compounds, 218
asymmetric, 21 1 Delia
intrasexual, 201, 207, 230-231 antiqua, 83-84
and nuptial gifts, 229-233 radica, 83
between SIT and wild males, 110-1 11, 112 Density-dependent factors, 111
between swarming males, 199-200, 206-207 Derrnaptera, 2, 7
Complex life cycles, 137-138 male reproductive system, 36, 37
Compsothespis, 69 oogenesis, 12-1 3
Conflicts, in protandry, 179-180, 185-188 sex determination, 61, 68, 71
Conocephalus nigropleurum, 222 Deterrents, 159
Cooperation, in protandry, 185-1 88 Development time, 226
Copulation Diadromus pulchellus, 34, 77
duration, 233 Dicranomiya trinotata, 80
frequency, 152-154,234 Dictyoptera, 4, 68
Corn earwoms, 101, 156 Dimorphism, 201, 202, 203
Corpora allata, 10, 22, 29, 38, 95-96 Dioryctria amatella, 159
Corpus cardiaca, 28, 47 Diplacus auranticus, 162
Corpus cardiacum-stimulating factor (CCSF), 97 Diploptera punctata, 10, 96. See also
Corpus luteum, 28 Cockroaches
Corydalidae, 2 Diplura, 67
Costs Diprion pini, 145
autosterilization, 1 18 Diptera, 6, 197
screwworm infestation and eradication, 113, accessory glands, 8
114, 115 behavior, 123
Cotesia rubecula, 155 chorionization, 26
Crataegus monogyna, 151 fecundity and fertility, 144
Creophilus maxillosus, 6 follicle cells, 16, 21
Crickets, 5, 10, 28, 45, 218. See also Acheta germ cell formation, 10, 39
domesticus; Gryllodes; Gryllus hormones, 97-99
Crocidosema plebejana, 160, 161 male reproductive system, 36, 37, 42
Crowding, 156, 157, 211 nuptial gifts, 216, 217, 224, 234
Culex, 80 oogenesis, 22
tritaeniorhynchus, 116, 123 pheromones, 120
Cuticular intima, 6, 7 sex determination, 58, 59,63, 64-65, 79-85, 86
Cyclical parthenogenesis, 134-139 swarm formation, 201
Cydia pomonella, 102, 147, 153, 155 vision, 120
Cyphoderris Dispersal, 156-158, 163, 226227
buckelli, 2 19 Distribution, Ideal Free, 208-210
strepitans, 222, 232 Diuraphis noxia, 155
Cyrtobagous salvinae, 161 Dolichopoda, 70
Cystoblast, l l Dominant Y system, 59, 63, 66, 85
Cytoskeleton, 16 Dragonflies, 204-205
Drepanosiphum platanoidis, 137, 146,
D 150-151, 157
Dricranotropis hamata, 150
Dacus, 120 Drosophila
jarvisi, 164 body size and mating success, 207
tryoni, 164 crowding, 2 11
Danaus, 7 funebris, 42, 45, 46, 48
plexippus, 153, 160, 163, 165, 218 melanogaster
nuptial gifts, 221, 225, 226, 233 female ducts, 10
Dance flies, 216 FESIRIS, 45
Daphnia, 133 follicle cells, 16
Darwin, Charles, 176 germ cell formation, 10-1 1
Dasyllis grossa, 83 hormones, 98
 Insect Reproduction

JH receptor protein, 41 Elymana sulphurella, 150

male reproductive system, 34, 36, 48 Embioptera, sex determination, 68, 70
oogenesis, 11-19, 21, 23, 25 Embryonic development, 10, 1 1, 37-39, 135
ovarian structure, 6 Emergence. See Protandry
ovipositor, 4 Emphis, 208
sex determination, 59, 83, 84 Empis borealis, 2 16, 220
size and fecundity, 146 Encarsia, 132
miranda, 83, 84-85 Encelis incisus, 150
mojavensis, 224 Endocrine regulation, 95-103
nebulosa, 217 accessory glands, 10, 38, 4 1 4 2
persimilis, 85 early oogenesis, 15
pheromones, 120 Endomeiosis, 133
phylogenetic history, 228 Endosomes, 23
pseudobscura, 85 Energids, 11
sechellia, 45 Ensifera, 4, 15
sex determination, 57, 64-67, 70 Environmental factors, 2, 29, 33
subobscura, 217 fecundity and fertility, 154-156
willistonin, 2 17 ovariole number, 145, 157
Dryas julia, 221, 222, 223,232-233,234 parthenogenesis, 132, 133
Ductus ejaculatorius duplex, 40 protandry, l80
Dysdercus seasonal changes in foliar nitrogen, 150-151
fasciatus, 10, 165 seasonal habitat trends, 138
intermedius, 18 sex determination, 58, 69, 70, 73
SIT, 111, 114
Epeorus longimanus, 203, 204
Ephemeroptera, 2, 22
Earwigs, 12-13. See also Dermaptera male reproductive system, 36, 37
Ecdysone, 10,22,40,42,97, 103 sex determination, 67
Ecdysteroid hormones, 6, 37, 38-39, 95 swarm mating, 200
accessory gland regulation, 41 Ephestia, 79
egg laying, 46 kiihniella, 10, 38, 103
oogenesis, 96 Epiphyas postvittana, 146, 147, 153, 156
Ecdysterone, 38 Eradication, 111, 119
Eclosion, 178, 235 Erigeron, 150
Economic aspects ESS (Evolutionary Stable Strategy), 177, 179,
autosterilization, 1 18 207
screwworm infestation and eradications, Esterase 6, 36, 48
113, 114, 115 Estivation, 137
Ectadenia, 36 Estrified phosphate, 20
EDNH (egg development neurosecretory Euceraphis punctipennis, 151
hormone), 97 Eumeas, 164
Effectiveness, of control techniques, 111, 119 Euphydryas editha, 158
Effective population size, 227-228 nutrition, 147, 166
Egg development neurosecretory hormone protandry, 181, 182
(EDNH), 97 Euploea core, 148, 165
Egg hatch, 154, 235 Eupyrene sperm, 39-40,48, 102, 219
Eggs Eurema
distribution, 46-47, 160 brigitta, 157-1 58
germ cells, 10-1 1, 37 herla, 157-1 58
size, 164, 165 European corn borer. See Ostrinia nubilalis
Eggshell, 2, 15, 25-28 Eurosta solidaginis, 162
Ejaculate, 222-223 Eusimulium
Ejaculatory duct, 34, 35, 37 aurum, 80
Electrophoretic current, 18, 19 vernum, 80
Euura lasiolepis, 16 1 Frankliniella occidentalis, 150
Euxoa messoria, 147, 148 Fruit fly. See Dacus; Drosophila
Evolution Fusomes, 6, 11
leks, 201-203
male adornment, 200
mate acquisition, 176-177
nuptial gifts, 228-236 Galerucella lineola, 159
optimal size, 187 Galleria mellonella, 22
parthenogenesis, 132-1 33 Galloisiana nipponensis, 15
sex determination, 85-86 Game theory model, 179
Evolutionary individual, 13 1 Gas exchange, 27-28
Evolutionary Stable Strategy (ESS), 177, 179, Gene dosage, 67, 85
207 Generation-specific reproduction strategies,
Exaptations, 228 136, 138
Eye pigment mutations, 116-1 17 Genetically impaired female technique,
116-1 17
Genetic control, 115-1 18
"Genetic hitchhiking," 60
Fecundity, 144-158,223-224 Genetic sexing systems, 115-1 16
adult longevity, 148 Genic balance systems, 59, 60, 66, 70, 71, 82
adult size, 146 Genital chamber, 7
age in mating, 15 1-152 Genitalia, external, 4, 36
host quality, 149 Germarium, $ 6 , 11, 12, 14
nuptial gifts, 220, 229, 235-236 Germ cells, 10-1 1, 37
population structure, 226 Glandular cells, 34
Fecundity-enhancing substances (FES), 44-47 Glossina, 9-10, 28
Female-killing system, 117 austeni, 116
Female reproductive system, 1-29, 220-221. control, 121-122, 123
See also Parthenogenesis morsitans, 116
Fertility, 144-158, 165 palpalis, 59, 84, 112
Fertilization, 40 SIT, 112
FES (fecundity-enhancing substances), tachinoides, 112
44-47 Gomphocerus rufus, 8
Fiorinia externa, 150 Gonadal growth, 134-1 35
Fire ant, 77, 78 Gonepteryx rhamni, 184, 185
Fitness, 205-206, 223-226, 230, 235 Goniozus nephantidis, 78
of offspring, 148, 158, 159-166 Gonopore, 6-7, 36, 37
and population structure, 226 Grain aphid, 133, 149, 157
Fitness payoff, 208, 209, 210-21 1 Grapholitha funebrana, 155
Fixed distribution models, 177-178 Graphosoma lineatum, 165
Flexible distribution models, 177-178, Grasshoppers, 10, 26, 165.
180-181 See also Melanoplus
Flight, 156-158 Growth rates, gonadal, 134-135
aerobic, 206 Gryllidae, 5
mating success, 203-205 Grylloblattodea, 68
wing muscle autolysis, 157, 162 Gryllodes
Flush feeding, 15 1 sigillatus, 153, 222, 225, 229, 233
Follicle cells, 11, 12, 15-17 veletis, 153, 225
and chorionization, 26 Gryllotalpa fossor, 67
and vitogellin synthesis, 21, 25 Gryllus
Folsomia candida, 150 bimaculatus, 5, 15, 28, 222, 232
Food. See Foraging; Nuptial gifts; Nutrition veletis, 222, 225
Foraging, 220, 225, 236 Gynandromorphs, 58
Formica, 15 Gypsy moth. See Lymantria dispar
 Insect Reproduction

and females, 96-101

juvenile. See Juvenile hormones
Habitat, 138,211,227 and males, 101-103
Habrobracon and oogenesis. See Oogenesis
hebetor, 77, 78 oostatic, 22, 99
juglandis, 12 prothoracicotropic, 38
Haematobia irritans, 1 13 and reproduction, 95-1 03
Haplodiploid sex determination, 63-64, 72, 77, Horn fly, 113
78,82 Host alteration, 137, 139
Harpobittacus similis, 2 16 Host growth phase, 150
Heat-shock proteins, 18 Host location, 157
Heel fly, 113 Host odors, 121
Heliconius Host plant quality, 149-151, 158, 164
charitonius, 144, 153 chemistry, 160-162
nuptial gifts, 220, 221, 222, 225, 234 and female emergence, 179
cydno, 220,234 phenology, 162-163
erato, 221 "Hotshot" model, 201
mate competition, 231 "Hotspot" model, 201
Helicoverpa Housefly. See Musca domestica
armigera, 160 Humidity, 155-156
zea, 101, 156 Hyalophora cecropia, 17, 41, 45, 46, 47
Heliothis Hydrotaea meriodionalis, 83
virescens, 38, 39, 102 20-Hydroxyecdysone, 10, 15,22, 97
fecundity and fertility, 147, 148, 153, chorionization, 28
154 spermatogenesis, 102-103
zea, 145, 156, 165 sperm release, 103
Hemiptera, 2, 4, 6 Hylobittacus apicalis, 220, 225, 227,
fecundity and fertility, 144-158 233
hormones, 99 Hymenoptera, 4
male reproductive system, 43 follicle cells, 16
nuptial gifts, 217, 219 germ cell formation, 11
oogenesis, 13, 22 nuptial gifts, 216
oviposition and larviposition, 158-166 oogenesis, 22
sex determination, 60, 61, 86 parthenogenesis, 132, 133
Heritability, 229 sex determination, 63-64, 72, 77-78
Hermaphroditism, 58, 72 Hyperomyzus lactucae, 155
Hessian fly, 82, 101 Hypodenna lineaturn, 113
Heteropeza pygmaea, 82, 135
Heteroptera
nuptial gifts, 216
sex determination, 7 1 Icerya, 58, 72
spermatheca, 7, 8 Ideal Free Distribution, 208-210
Hibernation, 137 IGRs (insect growth regulators), 121
Hilara, 2 16 Immigration, 1 10-1 11, 112
"Hitchhiking," genetic, 60 Inbreeding, 77-78, 177
Holcus lanatus, 150 Initiatorin, 48
Homoptera, 37 Insect growth regulators (IGRs), 121
hormones, 99-101 Insecticides, 83, 109, 134
sex determination, 63, 72 integrated pest management,
Honeybees, 4, 7, 22,44, 77, 78 109-1 24
Hormones Insemination, hemocoelic, 37
adipokinetic, 22 Integrated pest management, 109-1 24
ecdysteroids, 6, 37, 38-39, 41, 46 Intersexes, 58, 69, 79
Index

Intraclonal variation, 133 Locusta migratoria, 5, 8

Isoptera, 68, 69 chorionization, 26, 27, 28
FESRIS, 45
J follicle cells, 15, 16
Juvenile hormones, 9, 10, 15, 95-101 germ cells, 10
in accessory glands, 41, 95 male reproductive system, 37, 38, 43-44
chorionization, 28 oogenesis, 15, 20, 21, 22, 23
in control, 121 pheromones, 101
oogenesis, 96 Longevity, 148
and RIS, 47 Lopinga achine, 165, 188
spermatogenesis, 38, 40-42 Lovebug, 34, 36, 199, 201, 203
yolk protein production, 21-22 Lucilia, 6
cuprina, 59, 83, 84
sericata, 113, 116-1 18
Lymantria dispar, 78-79, 102, 103, 149,
Kairomones, 120 164
Killing systems, 110 egg size, 165-166
Kogatus modestus, 146 lifespan, 164
Lytta
magister, 222
nuttalli, 34
Lampbrush chromosomes, 15 vesicatoria, 2 18
Landmark, for swarming, 201, 202-203, 206,
21 1
Larval habitat, 21 1
Larviposition, 158-1 66 Macromolecular factor, 102
Lasiomrnata, 165, 188 Macroneurus appendiculatus, 76
Leafhoppers, 150 Malacosoma californicum pluviale, 160
Leaf-mining fly, 161-1 62 Male reproductive systems, 3349,
Leks, 175, 199-203, 21 1. See also Swarms 222-223
Lepidoptera, 7 Mamestra brassicae, 102
chorionization, 26, 27 Manduca sexta, 23, 102, 153, 154
fecundity and fertility, 144-158 Mantis religiosa, 69
male reproductive system, 37.43, 48 Mantodea, 68-69
endocrine regulation, 38-39, 42 Maternal sex determination, 72, 73, 78, 82. See
internal organs, 34-35, 36 also Parthenogenesis
nuptial gifts, 217-218, 219, 234 Mating
female resource budget, 22 1, 225 acquisition, 175-194.229-233.234
male resource budget, 222, 223, 224 age at, 151-152
oogenesis, 20, 21, 22 copulation frequency, 152-154, 234
oviposition and larviposition, 158-1 66 courtship, 232
sex determination, 58, 67, 78-79 disruption, as control, 109-124
Lepisma, 4 effects other than sperm transfer, 44
Leptidea sinapis, 185-1 87, 188 nuptial gifts. See Nuptial gifts
Leucophaea, 16, 20 options, 206-207
Lice, 7 1 success, 203-2 11, 228
Lifespan, 163-164, 178 swarm-based, 199-2 12
Lifetime reproductive success, 205-206, 219, Mating plug, 36, 217, 233
235 Mayetiola destructor, 82, 101
Lipids, 20, 27, 41 Mayflies, 203, 204. See also Ephemeroptera
Liriomyza trifolii, 161-1 62 Mecoptera, 78, 216, 217, 234
Liriope, 80 Mediterranean flour moth, 10, 38, 79, 103
Lithocolletis quercus, 162 Mediterranean fruit fly. See Ceratitis capita
 Insect Reproduction

Megaloptera, 2, 6 Muller's ratchet principle, 59-60, 132

nuptial gifts, 217, 222 "Multiple factor systems," 63, 64, 66,
oogenesis, 14 80-8 1
refractory period, 225, 227 Multiple sex chromosome systems, 61-63, 69,
sex determination, 75-77 82,84
vitellogenesis, 19 Hemiptera, 73-75
Megarcys signata, 146 Lepidoptera, 79
Megaselia scalaris, 83 Orthoptera, 70
Megoura viciae, 100 Siphonaptera, 78
Meiosis Musca, 6, 8
and chromosome translocation, 115-1 16 domestica
oogenesis, 14, 15 autosterilization, 119, 121, 122-123
sex determination, 58, 74 FESIRIS, 45.46
spermatogenesis, 39, 102 genetic control, 116
Melanoplus, 144 hormones, 98
difSerentialis, 10 male reproductive system, 35, 36
sanguinipes, 5, 35, 221 oogenesis, 22
male reproductive system, 41, 42, 43, 46, oviposition, 28
48 pheromones, 101
Melipona compressipes fasciculata, 78 sex determination, 83, 85
Meloe proscarabaeus, 6 Muscle mass, and flight, 203-205
Meroistic ovaries and ovarioles, 17, 18 Mutations, 59-60
Mesadenia, 36 accumulation and parthenogenesis, 132
Mesospermalege, 2 19 control technique, 110, 115
Metamorphosis, 235 eye pigment, 116-1 17
Metopolophiurn Mycalesis perseus, 162
dirhodum, 133, 149 Mycetocytes, 21
persicae, 133 Myrmeleotettix maculatus, 165
M factors, 63 Myzus persicae, 155
Micropinocytosis, 23
Micropyles, 27, 28 N
Midges. See Anarete pritchardii;
Chironomidae Nasonia vitripennis, 78
Migration, 136, 137, 156-158 Natural selection, 176-177, 180, 207, 229. See
Milk glands, 9-10 also Evolution
Milkweed bug. See Oncopeltus fasciatus Nauphoeta cinerea, 23
Models Nemobius sylvestris, 2 17-2 18
distribution, 177-178, 180-18 1 Neodiprion
game theory, 179 nigroscutum, 77
"hotshot," 201 serfifer, 146
Monte Carlo, 227 Neohermes filicornis, 76
optimization, 178, 180 Neo-XY systems, 59, 60-61
of protandry, 177-178, 180-182 beetles, 77
Monarch butterflies. See Danaus plexippus Drosophila, 85
Monte Carlo model, 227 Hemiptera, 72
Morphology Orthoptera, 70
female reproductive system, 2-10 Phasmida, 69
male reproductive system, 33-37 primitive Exopterygota, 67-68
Mosquitos. See Aedes; Anopheles; Culex Neuroptera, 2 17
Moths, 18. See also Antheraea polylphemus; Neuropteroidea, 75-77
Epiphyas postvittana; Hyalophora Neurosecretory factors, 42, 46, 97, 100
cecropia New World screwworm fly, 45, 113-1 15
mRNA, 4 Nezara viridula, 34
Index

Nitrogen, 149-150, 159 nuptial gifts, 23 1, 234

Notoptera, 4 for egg production, 22 1
Nuculaspis tsugae, 150 evolution of, 228-229
Nuptial gifts, 215-235 spermatophores, 2 17, 223
evolution, 228-236 wings, 219
fecundity, 235-236 refractory period, 222, 225, 227
fitness, 223-226 sex determination, 67, 68, 70, 86
mate choice, 232-222 spermatheca, 8
physiological costs and benefits, Ostrinia nubilalis, 103, 155, 222, 225
220-223 Ovaries, 2 , 4 4 9 7
population structure, 226-228 Ovarioles, 2, 3,4, 28
Nurse cells, 2, 5-6 number of, 144-145, 157
oogenesis, 12, 13-14, 17-19 Oviducts, 6, 28
spermatogenesis, 34 Oviposition, 28-29, 46, 158-1 66
Nutrition and nutrition, 147, 158
adult, 147-148, 165 and plant phenology, 162
copulation as nutrient transfer, 153 stimulants, 159
and dispersal, 226-227 Ovipositor, 4
fecundity and fertility, 145-15 1 Owen, Richard, 131
larval, 145-147, 164
and mate choice, 232
transfer by nuptial gifts, 218, 224-225
Nymphs. See Ephemeroptera Paedogenesis, 135
Pales ferruginea, 79-80
Panoistic ovaries and ovarioles, 2, 5
oogenesis, 11, 15, 17, 18
previtellogenesis, 19
Odonata, 2, 4 Panolis jlammea
male reproductive system, 36, 37 attractants, 159
nuptial gifts, 217 egg size, 165
sex determination, 60, 67 fecundity and fertility, 146, 147, 148,
Offspring 151-152, 154
fitness, 148, 158, 159-166 lifespan, 163
size, 164-166 plant selection, 160-1 61, 164
suvival and nuptial gifts, 224-225 sex ratio, 153, 154
Oncopeltus fasciatus, 7, 10, 72, 163 Panorpa, 217, 224
Oocytes, 4-5, 14-15, 135 Panstrongylus megistus, 15
Oogenesis, 11-19,96-101, 236 Papilio
stimulation by nuptial gifts, 220-22 1, machaon, 222
223 polyxenes asrerius, 153
Oosorption, 29 Parapediasia teterella, 164, 165
Oostatic hormone, 22, 99 Pararge aegeria, 148, 223
Operational sex ratio (OSR), 227-228, 230, egg size and weight, 147, 165,
231-232,236 166
Operophtera brumata, 151 nuptial gifts, 223, 225
Optic lobes, 44 protandry, 182-183, 188
Optimization model, 178, 180 Parasites, 223
Opuntia, 161 Paroposis atomaria, 160
Ornamentation, 200 Pars intercerebralis, 8, 15, 44, 99
Orthoptera Parthenogenesis, 69,73,99-100
colleterial glands, 9 cyclical, 134-1 39
male reproductive system, 36, 37, 39.43 evolution of, 132-1 33
mate choice, 232 Partial sterility technique, 115-1 16
 Insect Reproduction

Patency, 23, 99 Plarycotis vitata, 151

Paternity, security of, 233-234, 235 Plecia nearctica, 34, 36, 203
PBAN (pheromone-biosynthesis-activating Plecoptera, 2, 68
peptide), 101 Plodia interpunctella, 153-154, 222
Pedicels, 28 Plutella maculipennis, 155
Pedicularis semibarbata, 158 Pole cells, 10-1 1
Pedogenesis, 82 Polyphenism, 135-137
Pemphigus, 137 Polyploidy, 18
Penetration, of ovipositor, 4 Polytrophic ovaries and ovarioles, 2, 5
"Penis," 36, 37 oogenesis, 13, 17, 18
Periphyllus Pontia
califomiensis, 162 daplidice, 188
testudinaceus, 137 protodice, 221, 223, 232, 233
Periplaneta americana Population size, 227-228
accessory glands, 48 Population structure, 226-228
colleterial glands, 9, 10 Porthetria dispar, 57
follicle cells, 15, 16 Postembryonic development, 38-39
germ cells, 10 Postzygotic investments, 216, 230
oogenesis, 15.96 Potential fecundity, 144, 146, 147
seminal fluid, 44 Precocene II,98
Perla immarginata, 68, 151, 156 Predation, 208, 210-21 1, 225
Perlodes, 68 Previtellogenesis, 19
Pesticides Prey items, 216-217, 220, 228, 229
integrated pest management, 109-124 and copulation duration, 233
residues, 109 male to male, 23 1
resistance to, 83, 109, 134 Pristiphora erichsonii, 145
Phallic lobes, 36, 37 Procladius crassinervis, 206
Phallotreme, 36 Prokelisia marginata, 148
Phasmida, 68.69 Prostaglandin, 46
Pheromone-biosynthesis-activating peptide Prostaglandin synthetase, 45
(PBAN), 101 Protandry
Pheromones, 101, 120 benefits, 178-179,226
Phlebotomus pemiciosus, 80 cooperation/conflicts, 179-1 80, 185-1 88
Phormia, 19 models, 177-178, 180-182,201
regina, 45, 59, 98, 99 Protandry theory, 152, 175-180
Photoperiod, 100, 155 assumptions, 182-1 85
Phratora vitellinae, 159 tests of, 180-188
Phthiraptera, 2 Proteins
Phthorimaea operculella, 147 FESIRIS, 45,46
Phyllocolpa, 160 heat-shock, 18
Phylloxera caryaecaulis, 74, 75 in secretions, 41
Phylogenetic history, of nuptial gifts, yolk, 15, 19-22
228-229,236 Prothoracic glands, 95
Pieris Prothoracicotropic hormone, 38
brassicae, 164, 188 Protogyny, 152
napi, 187-1 88 Protothermes
rapae, 158, 159, 165, 188,233 grandis, 222, 225
Pine beauty moth. See Panolis$ammea immaculatus, 222, 225
Pine looper moth, 146 Prunus padus, 150
Pinus Pseudaletia
nigra, 149 pheromones, 101
sylvestris, 145 punctella, 2 18
Plannipennia, 75-77 spermatogenesis, 102
Planthopper, 148 unipuncta, 225, 227
Pseudoregma, 137 Scarabaeinae, 6
Psocoptera, 2, 4, 37, 71, 217 Schisrocerca gregaria, 5, 9, 10
Psylla male reproductive system, 38, 44
peregrina, 151 Schizaphis graminum, 133
subferruginea, 151 Schoutedenia ralumensis, 73, 74
Publilia reticulara, 151 Sciara coprophila, 82
Pupal weight, 183-1 84 Screwworm fly, 45, 113-1 15
Pyrrhocoris apterus, 57 Scrophularia californica, 162
Secondary metabolites, 151
Secretions
EDNH, 97
Radiation, 115. See also Sterile insect technique neurosecretory factors, 42, 46, 100
(SIT) nonspermatophore, 2 17
Raphidioptera, 2, 6, 14, 75-77 as nuptial gifts, 217, 219
Raspberry aphid, 133 proteins in, 41
Receptivity-inhibitingsubstances (RIS), 36, vitogellin, 22-23
44-47 Security of paternity, 233-234, 235
Recessive-X systems, 59, 66, 72, 76, 84 Seminal fluid, 44
Refractory period, 225, 227, 228, 233, 234 Seminal vesicle, 40
Regurgitants, 2 16-217 Semiochemicals, 120
Reinfestation, 112 Senecia jacobaeae, 150, 159, 160
Reproductive potential, 157 Sensory equipment, dimorphism in, 201,
Reproductive strategies, generation-specific, 202
136, 138 Sesamia nonagriodes, 148, 154, 155
Reproductive success, 205-206, 219, Sex chromatin body, 79
235 Sex determination, 57-86
Reproductive systems, 1-29, 33, 49, 220-221 dosage compensation, 67, 85
Requena verticalis, 222, 223, 234 evolution of, 85-86
Resource budget, 216, 220,226 general aspects, 58-67
female, 217, 218, 224, 233, 235 haplodiploid, 63-64, 72, 77, 78, 82
male, 222, 235 maternal, 72, 73, 78, 82
Rheumaptera hasrata, 149 molecular basis, 64-66, 85
Rhodnius, 7, 8, 19 "multiple factor systems," 63, 64, 66,
male reproductive system, 42, 46, 47 80-8 1
prolixus, 16, 28, 45 multiple sex chromosome systems. See
hormones, 99, 101-102, 103 Multiple sex chromosome systems
Rhopalosiphum padi XWXO systems, 59-61, 62, 67-77,78-79,
fecundity, 145, 146, 155, 156 83, 85
flight, 158 XWXY systems. See XXIXY systems
host plants, 149, 150, 151, 162 ZOIZZ systems, 79
Rhyacionia buoliana, 153 Sex factors, 58
Rhynchaenus fagi, 165 Sex mosaics, 69
RIS (receptivity-inhibiting substances), 36, Sex ratio, 58, 71, 72, 73, 75, 82
44-47 and autosterilization, 118
Rotifers. 132 fecundity and fertility, 153-1 54
host quality, 149
from inbreeding, 77
mate acquisition, 175
Salix, 145-146, 159, 160 Operational Sex Ratio, 227-228, 230,
Salvina molesta, 161 231-232,236
Samea multiplicalis, 161 Sex roles, 230-231, 232, 233
Samia cynthia, 10, 38, 39 Sexual selection, 230-23 1, 235,
Sawfly, 145 236
Scale insects, 13, 58, 72, 150 Shoot-galling sawfly, 161
 Insect Reproduction

Sialis flavilatera, 19 Sycamore aphid, 137, 146, 150-151, 157

Silkmoth. See Bombyx mori Synaptonemal complexes, 14
Silkworm, 20
Simulium vittatum, 26
Siphonaptera, 2
Sitobion avenue, 133, 149, 157 Tegrodera alogra, 222
SIT (sterile insect technique), 110-1 15 Teleogryllus commodus, 44, 45, 46
Size. See Adult size; Clutch size; Eggs, size; Telescoping of generations, 134
Offspring, size; Population size; Telotrophic ovaries and ovarioles, 2, 6
Swarms, size oogenesis, 13, 14-15, 18-19
Smell, 101, 120, 121 Temperature, 154-1 56, 185-1 86
Snakeflies. See Raphidioptera Tenebrio molitor, 7, 10
Solenobia, 79 male reproductive system, 35, 37, 38, 41, 42,
Solenopsis invicta, 77, 78 43,48
Solidago, 150, 162 Tepenoids. See Juvenile hormones
Spanish flies, 218 Tephriris bardanae, 162
Spartina, 151 Terminal filament, 5, 10
Spear-marked black moth, 149 Termites, 68, 69
Sperm, 39,47,219 Territoriality, 175
release, 40, 103 Testes, 34, 35, 39, 102, 103
Sperm activator, 48 Tettigonioidea, 5, 224
Spermalege, 37 Thaumastoptera calceata, 80
Spermatheca and spermathecal accessory Thaumetopoea pityocampa, 149
glands, 2, 7-8, 47, 221 Thelytoky, 72, 73, 74, 78, 79, 132
Spermatodesm, 34.39 Therioaphis trijolii forma maculata, 134
Spermatogenesis, 39-40 Thermobia domestica, 67
hormone regulation, 101-103 Thrips, 150. See also Thysanoptera
sex determination during, 73, 74 Thysanoptera, 4, 36, 37
Spermatophores, 8, 33, 47, 48 sex determination, 63, 74-75
evolution, 226, 228-229 Thysanura, 67, 228
formation, 42-44 Tipula
as nuptial gifts. See Nuptial gifts caesia, 80
Spermatophylax, 218, 219, 224, 229, 236 pruinosa, 80
Speyeria mormonia, 225 Tobacco budworm. See Heliothis virescens
Spodoptera Tobacco homworm, 23, 102, 153, 154
exempta, 147-1 48 Toxicity
frugiperda, 150, 156 mammalian, 121
latifascia, 156 of retained nuptial gifts, 225
litura, 42, 153 Tracheae, for ovarioles, 4
ornithogalli, 165 Transposable element, 58, 66, 8 1, 85
Spruce budworm, 147, 150, 159 Trehalases, 48
Stable fly, 112 Trialeurodes vaporariorum, 72
Staphylinidae, 6 Triatoma infestans, 10
Sterilants, 120-121 Tribolium destructor, 16
Sterile insect technique (SIT), 110-1 15 Trichogramma, 132
Stick insect, 5, 69 Trichoptera, 43
Stimulus pooling, 201-202 nuptial gifts, 217, 221, 223
Stomoxys calcitrans, 112 sex determination, 58, 78
Stonefly, 146 Triflumuron, l22
Strepsiptera, 37, 77 tRNA, 4
Supella longipalpa, 101 Tropharium, 13, 14, 17
Survival patterns, 224-226 Trophic cord, 2, 6
Swarm mating, 199-212,201-203,206. See Trophocytes, 21
also Leks Tsetse flies. See Glossina
Index

Tsuga, 150 Weevils, 161

Tyria jacobaeae, 150, 160 Wing muscle autolysis, 157, 162

Urocytes, 21
Xestoblatta hamata, 218, 220, 225
Uroleucon gravicome, 150
XXIXO systems, 59-61, 62,67-77,78-79, 83,
Uterus, 9
85
Utethesia ornatrix, 2 18
XXJXY systems, 58-59, 61
beetles, 77
v Dermaptera, 7 1
evolution of, 85
Vagina, 7
Musca domestica, 83
Vas deferens, 34, 35, 37, 39,40
Orthoptera, 68, 70
Vas efferens, 34
Panorpoid orders, 78, 84
Veronia noveboracensis, 15 1
primitive Exopterygota, 67
Virginoparin, 100
Viruses, 121
Vision, 120, 201 Y
Vitellarium, 12, 14 Yolk proteins, 15, 19-22
Vitelline membrane, 2, 26-27 Yponomeuta evonymellus, 149, 164
Vitelline membrane bodies, 26
Vitellin (vn), 19-21, 23
Vitellogenesis, 5-6, 15-17, 19-25
Vitellogenin (vg), 4, 19-21, 96, 99, 220
Vitogellin, 21-25 Zeiraphera canadensis, 147
Zoraptera, 70
ZOIZZ systems, 79
W ZWIZZ systems, 58-59,78
Wachtliella persicariae. 82 Zygentoma, 2
Wasps, 12, 77, 78, 225 Zyginidia saitellaris, 150