FAO Animal Health Manual No.

EPIDEMIOLOGY, DIAGNOSIS
AND CONTROL OF
HELMINTH PARASITES
OF SWINE
Allan Roepstorff
Peter Nansen
Danish Centre for Experimental Parasitology
The Royal Veterinary and Agricultural University
Copenhagen, Denmark

FOOD AND AGRICULTURE ORGANIZATION OF THE UNITED NATIONS

Rome, 1998
 The designations employed and the presentation of material in this
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M-27
ISBN 92-5-104220-9

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0 FAO 1998
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

FOREWORD

Helminth parasites of swine are ubiquitous and although no precise

information is available on the economic impact of these to the pig
producers, there is little doubt that they are important causing
reduced feed conversion efficiency and slower weight gain. Both
the commercial and the small-scale farmer continuously rank worm
infections high among the health problems. The difficulties in
substantiating the losses are associated with the fact that the
infections are chronic and less dramatic than other diseases of
swine which make these infections among the most neglected areas
of veterinary care in developing countries. In addition, a few
helminths are important in veterinary public health, as transmission
to humans is possible through ingestion of raw- or under-cooked
meat. This handbook is written to assist animal health staff in
prevention and control of these infections.

The handbook, in a simple style, reviews the epidemiology of

economically important helminth parasites of swine and present
procedures and techniques for their diagnosis, survey and control.
The book is designed for routine use in all types of animal health
institutions, including universities, research institutes and field
laboratories where diagnostic parasitology is performed. It is hoped
that it will help to improve and standardize diagnostic capabilities as
well as contribute to the collection and use of basic epidemiological
data, the foundation for effective disease control programmes.

Jorgen W. Hansen, D.V.M.; PhD

Senior Officer, Parasitology
Animal Production and Health Division
FAO, Rome
iv The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

TABLE OF CONTENTS

1 THE MAJOR HELMINTH SPECIES - AN INTRODUCTION 1

1.1 INTRODUCTION 1

1.2 HELMINTH GROUPS 2

1.2.1 Nematodes 2
1.2.2 Trematodes 2
1.2.3 Cestodes 3
.1.2.4 Acanthocephala 3
1.3 LOCATION IN ORGANS/TISSUES OF THE HOST 3
1.3.1 Alimentary tract 4
1.3.2 Liver 4
1.3.3 Lungs 5
1.3.4 Other organs and tissues 6

LIFE CYCLES AND EPIDEMIOLOGY OF HELMINTH PARASITES . . . 7 .

2.1 INTRODUCTION 7
2.2 NEMATODES OF THE DIGESTIVE TRACT 7
2.2.1 Hyostrongylus rubidus (red stomach worm) 7
2.2.2 Ascarops strongylina and Physocephalus sexalatus
9
2.2.3 Ascaris suum (large roundworm) 11
2.2.4 Strongyloides ransomi (pig threadworm) 13
2.2.5 Oesophagostomum spp. (nodular worms) 15
2.2.6 Trichuris suis (whipworm) 17
2.2.7 Other nematodes of the digestive tract . . . 19
2.3 NEMATODES OF THE LUNGS 19
2.4 NEMATODES IN OTHER ORGANS 21
2.4.1 Stephanurus dentatus (kidney worm) 21
2.4.2 Trichinella spiralis 23
2.5 TREMATODES 25
2.6 CESTODES 27
2.7 ACANTHOCEPHALA 31

FAECAL EXAMINATIONS FOR PARASITES 35

3.1 INTRODUCTION 35
3.2 COLLECTION OF FAECAL SAMPLES 36
3.3 QUALITATIVE TECHNIQUES FOR FAECAL
EXAMINATIONS 37
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

3.3.1 Test tube flotation 38

3.3.2 Simple flotation 41
3.3.3 Sedimentation (Trematode eggs) 44
3.4 QUANTIVATIVE TECHNIQUES FOR FAECAL EXAMINATIONS
47
3.4.1 Simple McMaster technique 47
3.4.2 Concentration McMaster technique 51
3.4.3 Counting the McMaster chamber 55
3.5 FAECAL CULTURES 56
3.6 IDENTIFICATION OF EGGS AND LARVAE 61
3.7 INTERPRETATION OF FAECAL EGG COUNTS 61
3.7.1 False negative and false positive egg counts 66
3.7.2 The relationship between egg counts and
worm burdens 68

4 POST-MORTEM DIFFERENTIAL WORM COUNTS 69

4.1 INTRODUCTION 69
4.2 THE CONTENTS OF THE STOMACH 70
4.3 THE STOMACH WALL 75
4.3.1 Incubation in physiological saline 76
4.3.2 Pepsin-HCI digestion 79
4.4 THE CONTENTS OF THE SMALL INTESTINE 83
4.5 THE CONTENTS OF THE LARGE INTESTINE 85
4.6 THE LARGE INTESTINAL WALL 86
4.7 THE LIVER AND OMENTUM 87
4.8 THE LUNGS 91
4.9 THE KIDNEYS 94
4.10 THE MUSCLES 95
4.10.1 The compression method 96
4.10.2 The pepsin-HCI digestion method 97
4.11 IDENTIFICATION OF HELMINTHS 100
4.12 INTERPRETATION OF WORM COUNTS 101
4.12.1 The helminth species 101
4.12.2 Specific host-parasite relationships 107
4.12.3 Management systems 107
4.13 GENERAL COMMENTS 108
4.13.1 The subsample technique 108
4.13.2 Occupational hazards 109

5 EXAMINATIONS FOR INFECTIVE LARVAE AND EGGS IN HERBAGE AND

vi The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

SOIL 111
5.1 INTRODUCTION 111
5.2 COLLECTION OF HERBAGE AND SOIL SAMPLES 111
5.3 ISOLATION OF INFECTIVE LARVAE 113
5.3.1 Isolation of infective larvae from herbage 113
5.3.2 Isolation of infective larvae from soil . . . 118
5.4 ISOLATION OF INFECTIVE EGGS FROM SOIL 121
5.5 IDENTIFICATION OF LARVAE AND EGGS 128

6 INVESTIGATING HELMINTH OCCURRENCE AND EPIDEMIOLOGY

IN A PIG POPULATION 131
6.1 INTRODUCTION 131
6.2 HELMINTH OCCURRENCE IN A HERD/FLOCK 132
6.3 LONG-TERM MONITORING OF A HERD/FLOCK 134
6.4 PLOT EXPERIMENTS 137

7 CONTROL OF HELMINTHS IN PIGS 141

7.1 GENERAL PRINCIPLES OF CONTROL 141
7.2 CONTROL OF NEMATODES 143
7.2.1 Stocking rate 143
7.2.2 Grazing management 143
7.2.3 Mixed or alternate grazing 143
7.2.4 Hygiene of pens 144
7.2.5 Dose and move 145
7.2.6 Routine deworming 145
7.2.7 Nose-ringing of sows 146
7.2.8 Adequate nutritional level 146
7.2.9 Genetic resistance 146
7.2.10 The 'gilt-only' system for control of
Stephanurus dentatus 146
7.2.11 Control of Trichinella spiralis 147
7.3 CONTROL OF TREMATODES 147
7.4 CONTROL OF CESTODES 147
7.5 ANTHELMINTICS 148
7.5.1 Definition 148
7.5.2 Characteristics of an ideal drug 149
7.5.3 Dosing methods 150
7.5.4 Anthelmintic classes 150
7.5.5 Which drug to use 7 152

7.6 ANTHELMINTIC RESISTANCE 152

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine vii

7.6.1 Definition and underlying mechanism 152

7.6.2 Detection of anthelmintic resistance 153
Risk factors for development of
anthelmintic resistance 155
7.6.4 Prevention of anthelmintic resistance . . . 157

BIBLIOGRAPHY 159
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1

CHAPTER 1

THE MAJOR HELMINTH SPECIES - AN INTRODUCTION

1.1 INTRODUCTION

As a background for evaluating prevalence and impact of

helminthosis in swine, identification of the species present in a herd,
area, country or region should be investigated. Different species have
different pathogenic effects and bionomics. Some species are
particularly important in the young animal, while others seem to
accumulate in the older age categories. Furthermore, some species
are relatively easy to control, while others are highly persistent in the
environment and difficult to combat, unless more drastical measures
are taken.

Unless the helminth spectrum in a given herd or area is already

defined, a status of the helminth situation should be made initially.
In the first run this should be kept simple, having the primary
purpose to encircle the major species present. With this information
at hand more detailed investigations should be designed to cover the
major, relevant species in the particular herd or area.

Helminths in the pig are relatively easy to identify if one is aware of

their size and gross morphology and the tissues and organs in which
they are normally located. Macroscopically visible lesions typical for
some species may provide supportive information.

Identification of the parasites may be based on post-mortem

examination at slaughter or when animals have died from disease.
Identification may also be based on examination of eggs excreted
with faeces. Only rarely will clinical symptoms be sufficiently specific
to point to a single, responsible species.
2 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

The helminths may be grouped according to their systematics

(Section 1.2) or according to their location in the pig host (Section
1.3).

1.2 HELMINTH GROUPS

The most important helminth species, classified into major groups,

are listed below.

1.2.1 Nematodes

Hyostrongylus (redstomach worm)

Gnathostoma
Ascaris (large roundworm)
Strongyloides (threadworm)
Globocephalus (hookworm)
Trichostrongylus
Oesophagostomum (nodular worm)
Trichuris (whipworm)
Metastrongylus (lungworm)
Stephanurus (kidney worm)
Trichinella

1.2.2 Trematodes

Fasciolopsis (intestinal fluke)

Gastrodiscus
Opistorchis
Fasciola (liver fluke)
Schistosoma (blood fluke)
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 3

1.2.3 Cestodes

Cysticercus cellulosae
Cysticercus tenuicollis
Echinococcus granulosus

1.2.4 Acanthocephala

Macracanthorhynchus hirudinaceus

1.3 LOCATION IN ORGANS/TISSUES OF THE HOST

Helminths usually have typical host locations that may help isolation
and identification. Adult worms in particular are confined to distinct
organs or parts of these, whereas larvae of many species may be
encountered in several organs or tissues in connection with their
migration.

For parasite identification attention should be paid to the young

animal, and animals which have not recently received anthelmintic
treatment. Usually pigs in the age category 2-6 months have the
largest worm burdens and may excrete many eggs with faeces.
However, for some species like Oesophagostomum spp. and
Hyostrongylus worm populations tend to accumulate in older
animals. One must also bear in mind that some species have rather
long prepatent periods, i.e. time interval between infection and start
of egg excretion, for which reason faecal egg counts may be
negative for a long period of time even in the presence of high
parasite burdens in the host. Stephanurus, the kidney worm, has a
prepatent period as long as 6-11 months.
4 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

1.3.1 Alimentary tract

The majority of worm species develops and lives in distinct,

characteristic sites of the alimentary tract. In some situations the
infections are accompanied by macroscopical gut lesions, but in
some cases there are no 'overt lesions even at high parasite burdens.
Larvae, which are less easily identifiable, may in many instances be
more pathogenic than adults. Table 1.1 lists the location and effects
of the most common helminths.

TABLE 1.1 Helminths in different sections of the alimentary tract

Site Helminth Effects

Stomach Hyostrongylus Mucosal damage

Ascarops Mucosal damage
Physocephalus Mucosal damage
Gnathostoma Mucosa! damage

Small intestine Ascaris Adults: mucosal changes

Larvae: liver and lung lesions
Strongyloides Mucosal damage
Globocephalus Blood sucking
Trichostrongyfus Mucosal damage
Macracanthorhynchus Mucosal damage
Fasciolopsis Mucosal damage

Large intestine Oesophagostomum Mucosal damage, nodules

Trichuris Blood sucking
Gastrodiscus Mucosal damage

1.3.2 Liver

Parasites in the liver usually produce macroscopical lesions indicative

of their migration and presence. Large transparent cysts attached to
the visceral surface of the liver will most likely show to be
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 5

Cysticercus tenuicollis, the metacestode of Taenia hydatigina (a dog

tapeworm). Deeper, partially superficial cysts may be metacestodes -
socalled hydatids of Echinococcus granulosis, also a dog tape worm.
Migratory lesions (tracts) in the parenchyma may be caused by
immature Fasciola and Cysticercus tenuicollis larvae. Ascaris and
Schistosoma produce fibrotic or granulamatous lesions, typical for
each of these helminths. Following incision adult Fasciola may be
observed and collected from the bile ducts.

TABLE 1.2 Helminths in the liver

Helminth Effect

Ascaris (larvae) Fibrotic lesions: "white" or "milk"

spots

Schistosoma (eggs) Fibrosis, granulomas

Fasciola (larvae, adults) Fibrosis, bile duct enlargement. Blood

sucking

Cysticercus tenuicollis (metacestodes) Tissue damage, fibrosis

Echinoccocus granulosus Pressure atrophy

(metacestodes: hydatid cysts)

1.3.3 Lungs

The most important helminths that invade the lung are the pig
lungworms, of which there are three common species belonging to
Metastrongylus. These are primarily found in the bronchioli. In
addition, Ascaris larvae pass the lung as part of their host migration -
and may produce traumatic lesions. Hydatid cysts may occasionally
I observed in the lung parenchyma.
6 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

TABLE 1.3 Helminths in the lung

Helmmth Effect

Metastrongylus Bronchitis, pneumonia

Ascaris Traumatic lesions

Hydatid cysts Tissue atrophy

1.3.4 Other organs and tissues

Helminths may be found in a number of tissues outside the

alimentary tract, liver and lungs. Accidental findings may simply
reflect aberrant migration of parasites belonging to the above organs.
However, other parasites have primary, natural predilection sites,
e.g. in muscle tissue and in the kidney, see Table 1 .4 below.

TABLE 1.4 Helminths in various organs

Site Helminth Effects

Muscle Trichinella (larvae) Minimal

Cysticercus cellulosae Minimal

Kidney Stephanurus Damage, moderate

Blood vessels Schistosoma Intestinal and hepatic

damage
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 7

CHAPTER 2

LIFE CYCLES AND EPIDEMIOLOGY OF HELMINTH PARASITES

2.1 INTRODUCTION

This chapter primarily describes life cycles, epidemiology and

pathogenecity of those helminths having highest prevalences and
greatest economic impacts. The description therefore mainly
comprises the helminths listed in Chapter 1 (Sections 1 .2.1-1 .2.3,
Tables 1 .1-1 .4). A more comprehensive list of helminths in pigs, with
focus on life cycles, transmission and host relationships will be found
at the end of the present chapter (Tables 2.1 and 2.2).

2.2 NEMATODES OF THE DIGESTIVE TRACT

With a few exceptions, these nematodes have direct life cycles, i.e.
transmission from host to host without development in intermediate
hosts. The adult worms are located in different parts of the
gastrointestinal tract, where females after mating produce eggs that
are passed out in the faeces. In some species the eggs embryonate
and hatch into larvae which after two moults develop into the
infective third-stage larvae (L3). In other species like Ascaris suum
and Trichuris suis eggs do not hatch and the infective stage is
reached within the egg. In the following the major species will be
dealt with separately, starting with species located in the anterior
part of the alimentary tract.

2.2.1 Hyostrongylus rubklus (red stomach worm)

Fig. 2.1 illustrates the life cycle of this nematode, which is confined
to the pig. The eggs, which are of the typical "strongyle"-type, indi-
8 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Exsheated L3- Young 1.5-larvae reenter the

larvae invade the stomach lumen and become
stomach wall patent at day 18-21

Ingestion of
infective
larvae, which
pass to the
stomach

Infective
L3-larva Unembryonated
egg in faeces

L2-larva

Illustration by Wm P Hamilton CM)

FIGURE 2.1 Hyostrongylus rubidus life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 9

stinguishable from those of e.g. Oesophagostomum species (see

below), are passed with faeces. Here they hatch within a few days,
and infective third stage larvae are developed within 1-2 weeks at
optimal temperatures (15-25°C). They are very motile and are not
only localized to faecal material but also to surrounding soil and
herbage (see Chapter 5). Pigs are infected by ingestion of the
infective larvae, developing into fourth stage larvae which inhabit the
stomach mucosa. The adult worms establish on the mucosal surface
and in the stomach lumen, and start egg production already after 18-
21 days (viz. prepatent period). Egg excretion per female worm is
generally low, compared with other nematodes (Chapter 3). Under
certain circumstances in temperate regions, the larvae may be
arrested in their development in the mucosa (hypobiosis) for periods
up to several months.

During the prepatent period there may be severe damage to the

gastric glands, leading to lowered acidity, mucosal hyperplasia,
nodule formation and haemorrhage. Clinical signs include
inappetance, loss of condition and anaemia, but usually not
diarrhoea.

The infections are mainly confined to outdoor reared pigs due to the
biological requirements for larval development. In some temperate
regions it is practice to mainly keep breeding stock, gilts and sows,
on pasture, which is the reason why these age categories are most
severely affected. The so-called "poor sow syndrome" is usually
attributed to Hyostrongylus rubidus often together with
Oesophagostomum spp. infection.

2.2.2 Ascarops strongylina and Physocephalus sexalatus

Fig. 2.2 shows the principle life cycle of these species, belonging to
the spiruid nematodes. The relatively small eggs already contain well-
developed embryos when passed in the faeces. For further
development the eggs are swallowed by coprophagous beetles. In
these, the larvae develop up to the infective larvae (L3) and pigs be-
10 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Larvae invade the mucosa of the stomach

and infections become patent at week
4 (A. strongyfina) and 6 (P. sexalatus)

Ingestion of I
dung beetles I
with infective
larvae
I

Infective L3- Embryo-

nated egg
larvae encyst
in the dung in faeces
beetles
Faeces with eggs,
containing L1-larvae,
are ingested by dung
beetles

Illustration by Wm P Hamilton CMI

FIGURE 2.2 Ascarops strongylina and Physocephalus sexalatus

life cycles
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 11

come infected when ingesting infected dung beetles. After uptake of

beetles, the larvae are released and develop further to adults in or on
the gastric mucosa. Egg excretion is initiated after approximately 1-2
months.

Heavily infected animals have severe gastric inflammation with

ulcerations. Symptoms are more or less comparable with those
caused by Hyostrongylus rubicius.

Acquisition of these spiruid nematodes almost exclusively takes place

outdoors. A relatively wide range of beetles may serve as
intermediate hosts in tropical and subtropical areas, but infections do
also occur in some temperate regions of North America and Europe.
Infections are likely to be more severe where pigs are fed at a
restricted level and hence more dependent on scavenging.

2.2.3 Ascaris suum (large roundworm)

Fig. 2.3 illustrates the life cycle of this highly prevalent nematode.
The eggs which pass with faeces, usually in high numbers,
immediately start embryonating at temperatures above 1 5°C in the
dung or soil, and may reach the infective stage within 1 to 3 months
(or more) depending on temperature. Following uptake by the pig
host, the L3-larvae hatch in the small and large intestine, and
penetrate the large intestinal wall, from where they travel to the liver.
There they pass in the bloodstream to the lungs and hence to the
small intestine via the bronchi and trachea. The final moults take
place in the intestine, and patency (egg production) is reached after
6-8 weeks.

The initial penetration of the gut wall is apparently harmless, but the
migration in the liver causes local lesions ("white spots", "milk
spots"), which may cause condemnation of livers at slaughter. Yet,
the liver lesions are usually not rather pathogenic. However, the
arrival of the larvae to the lungs may lead to transient pneumonia
even with clinical symptoms if larval challenges are high. The imma-
12 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Larvae migrate up Larvae establish in the small

the bronchi and intestine and become patent
are swallowed at week 6-8

Ingestion of
infective egg
Hatched larvae penetrate
the large intestinali wall
and migrate via the liver
to the lungs

Infective egg Unembryonated

with L3-larva egg in faeces

Illustration by Wm P Hamilton CMI

FIGURE 2.3 Ascaris suum life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 13

ture and adult worms in the small intestine may lead to intestinal
disturbances, depending on the worm loads. Poor feed conversion
and slower weight gains resulting in an extension of the fattening
period have been recorded.

Infections with Ascaris suum are associated with both outdoor and
indoor production. Among all swine helminths, Ascaris is possibly
the most persistant and the most resistant to adverse environmental
conditions, due to its thick and resistant egg shell, which protects
against adverse environmental factors, desiccation and chemicals. In
the most modern and hygienic production enterprises, worm
problems are often left with this species only. In the pasture
environment eggs may maintain infectivity for up to 6-7 years.
Infections usually stimulate the development of a rather strong
protective immune reaction, which under practical conditions means
that older animals have less worms and excrete fewer eggs than the
younger ones. This has important implications for the targeting of
control programmes.

Ascaris suum may occasionally infect other hosts, in the form of

larval migratory lesions in liver and lung (ruminants), or even
resulting in patent infections (man and ruminants). The zoonotic
nature of Ascaris suum is as yet not fully defined.

2.2.4 Strongyloides ransomi (pig threadworm)

Fig. 2.4 illustrates the rather complex life cycle of this parasite,
which includes free-living generations of adult males and females,
and parasitic parthenogenetic females in the small intestine of the
pig. The female worms in the small intestine produce larvated eggs
by parthenogenesis, and these are excreted with faeces and will
soon hatch. After hatching, the larvae may develop into free-living
adult male and female worms, and this may lead to successions of
free-living generations. But under certain temperature and moisture
conditions the third stage larvae (L3) may become infective to the pig
by oral ingestion (penetration of oral mucosa) or skin penetration.
14 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

infective L3-larvae Larvae develop to partheno-

penetrate the skin or genetic females in the small
the mucosa of the oral intestine at day 7-9
cavity (after ingestion)
and migrate to the
udder or to the small
intestine via the lungs

Larvae are arrested in the udder N

until farrowing, whereafter they,
via colostrum, are transferred to
the piglets in which they develop
L3-larvae to parthenogenetic females in
may either the small intestine at day 4-5
infect pigs or Embryonated
develop to
free-living 4 egg excreted
in pig faeces
adults, which 41r. or laid by free-
eventually L2-larva L1-larva living females
produce
infective L3-
larvae

Free-living males and females

Illustration by Wm. P Hamilton CM!

FIGURE 2.4 Strongyloides ransomi life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 15

In the pig, the larvae migrate via the venous system, lungs, and
trachea to mature into adult females in the intestine. In addition,
piglets may acquire parasites immediately after birth from the
mobilisation of tissue-dwelling larvae in the sow which are
subsequently excreted in the milk (colostrum). The time from
infection to egg excretion, i.e. the prepatent period, is only 7-9 days
(or 4-5 days after lactogenic transmission to piglets).

Large intestinal burdens of Strongyloides ransomi may cause

inflammation with erosion of the epithelium. Severe diarrhoea in the
neonatal animal leads to weight loss, dehydration and in some cases
death.

Conditions with high temperature and humidity in connection with

poor hygiene favour development and accumulation of large numbers
of larvae that may severely affect young animals. In addition, the
neonatal pig may suffer from larvae transmitted through suckling.
Fatal cases may occur before eggs are excreted with faeces. The
infection induces strong immunity which explains that older age
categories are usually not clinically affected.

2.2.5 Oesophagostomum spp. (nodular worms)

Fig. 2.5 shows the life cycle of the different species of this parasite
located as adults in the large intestine. These are commonly
represented by two co-existing worms in the large intestine, i.e.
Oesophagostomum dentatum and 0.quadrispinulatum. Species like
0.brevicaudum may also occur. Adult females produce large numbers
of eggs of the "strongyle"-type, similar in morphology to those of
e.g. Hyostrongylus rubidus. The eggs hatch within a few days after
deposition in faeces, and through the first and second stage larvae
the third and infective stage (L3) may be reached within 10 days,
provided high temperature and humidity prevail. These larvae have
a low motility and tend to be localized in faeces and surrounding soil,
to a lesser extent in herbage. Pigs ingest larvae from their
environment, and the infective stages enter the mucosa of particular-
16 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Exsheated L3-larvae invade L4-larvae reenter the

the colon wall and nodules colon lumen and become
are formed around them patent at week 3-7

Ingestion of
infective
larvae, which
pass to the
intestine

Unembryonated
Infective egg in faeces
L3-larva

Illustration by Wm PHamitton CMI

FIGURE 2.5 Oesophagostomum spp. life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 17

ly the large intestine where they become enclosed in nodules and

moult. Then they emerge to the surface of colon and caecum and
develop to adult, egg-producing worms within 3 to 8 weeks
(different prepatent periods for different strains and regions are listed
in the literature).

The nodular worms cause enteritis and nodular formations in the gut
wall. Hyperplasia and ulcerations are prominent features at high
infection pressure. Since the parasite is only moderately
immunogenic, worm loads tend to accumulate in older age
categories, and clinical disease is most often encountered in the sow,
showing inappetence, weight loss and reduced milk production, only
rarely accompanied by diarrhoea. The "poor sow syndrome" is a
clinical condition usually associated with Hyostrongylus rubidus and
Oesophagostomum spp. (see Section 2.2.1).

Transmission of the infection is favoured by the high egg excretion

and humid and unhygienic conditions. Besides Ascaris suum this is
one of the most difficult worms to control, even under modern
conditions with strict control measures, including use of
anthelmintics. Some investigations have shown that sows may
sometimes exhibit a periparturient egg-rise coincident with farrowing,
a phenomenon which will favour transmission to the offspring.

2.2.6 Trichuris suis (whipworm)

Fig. 2.6 shows the life cycle of Trichuns suis. The adult worms,
located in caecum and in the anterior colon, produce eggs that are
excreted with faeces. The larva embryonates and develops within the
thick-shelled egg to the first stage, which is infective. As with
Ascaris suum, eggs do not hatch. After ingestion by the pig host, the
larva is released and penetrates the caecal mucosa and moults.
Subsequently, the remaining moults up to the adult stage take place
on mucosal surfaces of caecum and colon. Egg production occurs
after approximately 6 weeks.
18 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

The hatched larvae The larvae reenter the

invade the colon large intestine and become
wall and moult once patent at week 6-8

I
I
I
Ingestion I
of infec-
tive eggs

Infective egg
with Ll -larva Unembryonated
egg in faeces

Illustration by Wm P Hamilton CMI

FIGURE 2.6 Trichuris suis life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 19

Many infections are subclinical, but high worm burdens in young

animals may cause blood-stained diarrhoea due to the inflammatory
mucosal reactions, and the blood sucking activities of the worms.
Severe infections may lead to weight loss, dehydration and in some
cases death.

Infections with Trichuris suis are mainly associated with outdoor

rearing of pigs. The eggs of this helminth species have much in
common with eggs of Ascaris suum, as they are highly resistant and
may remain infective for years (up to 11 years). A relatively strong
immunity develops, and older animals are usually not carrying high
worm burdens, and hence they contaminate the environment only to
a minor degree.

2.2.7 Other nematodes of the digestive tract

These are briefly listed in Tables 2.1 and 2.2 with information on
type of life cycle, transmission and host relationship.

2.3 NEMATODES OF THE LUNGS

Fig. 2.7 presents the life cycle of the common lungworms in swine,
i.e. Metastrongylus spp. There are three widely distributed species,
M.apri, M.salmi and M.pudendotectus, which often occur
simultaneously in the same individual. The adults are located in the
small bronchi and bronchioles. They produce eggs that pass trachea,
are swallowed, and then excreted with faeces. These eggs may
hatch immediately after being ingested by earthworms which serve
as intermediate hosts. In the earthworm the development to the
infective (L3) takes place within 1-2 weeks at optimal temperatures
of 22-26°C. The longevity of the L3 in the earthworm may be several
years. The pigs become infected by eating earthworms, and the L3
are released and migrate to the mesenteric lymph nodes and moult,
and from there they migrate via the lymphatic-route to the lungs. The
20 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

L3-larvae penetrate the Larvae develop to patency

small intestinal wall and in the bronchi at week 4
migrate to the lungs

Ingestion
of earth-
worms
with
infective
larvae

Embryonated
egg in faeces

Faeces with
Infective L3- eggs, con-
larvae develop taining Ll-
within the larvae, are
earthworrns ingested by
earthworms

Illustration by Wm P Hamilton CM!

FIGURE 2.7 Metastrongylus spp. life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 21

prepatent period is approximately 4 weeks.

The lungworms may cause traumatic lesions during their migration

in the lungs in the form of peribronchial lymphoid hyperplasia, and
when the worms mature, chronic bronchitis and emphysema
develop. In heavy infections coughing is marked, accompanied by
nasal discharge and dyspnoea. Fatal cases do occasionally occur.

Lungworm infections in swine seem to be most severe in young pigs

4-6 months old. Older age categories are normally protected from
clinical attacks due to acquisition of immunity, although they may
suffer subclinically. In a given herd the exposure to infection may be
long-lasting due to the environmental earthworm reservoir.

2.4 NEMATODES IN OTHER ORGANS

2.4.1 Stephanurus dentatus (kidney worm)

Fig. 2.8 shows the life cycle of the kidney worm, which occurs in
tropical and subtropical regions of all continents. Eggs are passed
with the urine, and develop into third-stage infective larvae (L3) in the
environment. Thereafter there are three modes of transmission to the
pig: by ingestion of the free L3, ingestion of earthworms carrying L3,
and percutaneous infection with free L3. Irrespective of mode of
infection the larvae develop further in the liver parenchyma, and
thereafter they migrate in the peritoneal cavity to the perirenal region.
There, as adults they become enclosed in a cyst which they
communicate with the ureter, allowing worm eggs to be excreted in
the urine. The prepatent period ranges from 6-1 1 months. Quite
many worms migrate erratic in the body of the pig.

The pathological lesions due to the migration of the parasite are most
severe in the liver, where hepatic failure may occur in very serious
22 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Larvae migrate to perirenal

tissue and become patent at
month 6-19

mb;

Ingestion of
free-living L3-
larvae or
infected
earthworms.
The larvae
penetrate the
stomach wall Free-living L3-larvae
and migrate to may penetrate the skin
the liver and migrate to the
liver

Infective L3-larva, which

may be free-living or may
infect an earthworm
(transport host) Unem bryo-
.4110"1" nated egg
in urine
L1 -larva
L2-larva

Illustration by Wm P Hamilton CM

FIGURE 2.8 Stephanurus dentatus life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 23

cases. In general, overt clinical signs are rare, unless animals are
exposed to heavy challenge.

The L3 are highly susceptible to desiccation and are favoured by

damp ground. Since pigs often are lying in wet areas they may be
exposed to percutaneous infection. Infected earthworms ensure
persistence of infection in the environment.

2.4.2 Trichinella spiralis

Fig. 2.9 illustrates the life cycle of this nematode, having a unique
transmission biology. The parasite is able to infect a large range of
mammals, but from the zoonotic aspect (transmission to man) pig is
the most important host. The infective larvae are encysted in striated
muscles with particular high larval densities in Musculus rnasseter
and diaphragma. Development is resumed when the larvae are
ingested by another host as a result of cannibalism,
cryptocannibalism or predation. The larvae are then liberated, and in
the small intestine of the new host it undergoes several moults to
become sexually mature within a few days. The adults produce
larvae which enter the lymphatic vessels of the gut and travel via the
bloodstream to the skeletal muscles, where they become
encapsulated after 3-4 weeks.

Infection in pigs is normally not associated with clinical signs,

opposite the infection in man who may suffer severe illness. Fatal
human cases are not rarely reported from endemic areas.

The epidemiology of trichinellosis is determined by one animal eating

another (or offal from this). There are sylvatic and arctic Trichinella
life cycles, where animals of the wild fauna maintain transmission
through predation, cannibalism, carrion feeding etc. Usually these
cycles involve other species than T.spiralis, e.g. T.britovi, T.nelsoni,
T.nativa and others. In the domestic pig cycle, transmission is
maintained by feeding pigs on food waste containing flesh of
infected pig. Rats in piggeries can also maintain a secondary cycle
24 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Females invade the small

intestinal wall within 4-5 days

Larvae establish in the L1-larvae migrate to the striated

small intestine, molt 4 times muscles and encapsulate. The
and become adults larvae may be diagnosed after
17-21 days

Ingestion of
raw meat
(cannibalism,
cryptocanni-
balism or wild
animals)

Encysted larva in
striated muscle

Illustration by Wm P Hamilton CM'

FIGURE 2.9 Trichinella spiralis life cycle

The Epidemiology, Diagnosis and Control of Helminth Paras tes of Swine 25

which on occasions may pass to pigs or vice versa. Trichinella

species from the wild fauna may accidentally be introduced to pig
herds, but it is at present unknown to which extent they are able to
maintain a continuous transmission within the pig population. Until
now, there is overwhelming evidence that Trichinella spiral's is the
predominant species in the pig industries. The zoonotic importance
of this species is furthermore emphasized by the observation that it
may possibly be more pathogenic to man than the other species.

2.5 TR EIVIATO DES

There are several trematodes that may infect pig, as it will appear
from Tables 2.1 and 2.2 in this chapter, e.g. Fasciolopsis buski in the
small intestine, Fasciola hepatica, Opistorchis noverca and
Dicrocoelium dendriticum in the liver, and Schistosoma japonicum
(and other schistosome species) in the blood vessels. They have in
common that they have a broad final host range, among which man
may be an important and vulnerable host. In this way pig may serve
as a zoonotic reservoir host, where Schistosoma japonicum and
Fasciolopsis buski are the most important examples.

Fig. 2.10 shows the principle life cycle of Schistosoma japonicum,

which has a high prevalence in large areas of South-East Asia. The
relatively small eggs pass out with the faeces, and hatch in the
presence of suitable light and temperature, when they come in
contact with water. The short-lived miracidium, a free-swimming
larva, swims around in search of an Oncomelania snail, which it
penetrates and infects. Two generations of sporocysts develop, and
finally fork-tailed cercariae are formed. They break out of the snail
tissue into fresh water, approximately 4-8 weeks after the snail has
become infected. They rise to the surface and "hang" for some days
in the surface film. This exposes them to the pig skin, and they
quickly penetrate this. They reach the circulatory system and are
carried to the lungs from where they migrate to the liver where they
mature. Young adult pairs subsequently migrate in the hepatic portal
26 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Pairs of adults move to the mesenteric

veins of the large intestine and eggs
may be found in faeces at week 7-8

Cercariae
penetrate the
skin or
mucous
membranes
in the mouth
and migrate
to the liver via
the heart and
the lungs

Cercarla e
emerge into
the water

Egg in faeces
(contains a
miracidium)
Two generations of sporocysts The miracidium hatches in
develop in the snail host fresh water and penetrates a
snail host

Illustration by Wm P Hamilton CM

FIGURE 2.10 Schistosoma japonicum life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 27

system and settle primarily in the mesenteric veins of the large

intestine, in close distance to the gut wall. Here they produce eggs,
which penetrate the wall to the intestinal lumen, and are passed with
faeces. It usually takes 7-8 weeks from infection until eggs appear
in the faeces.

Acute disease, characterized by anorexia, dullness and diarrhoea,

occasionally bloody, may occur 7-10 weeks after heavy infection due
to inflammatory reactions related to the passage of eggs through the
gut wall. With time there appears to be a shift away from intestinal
reactions to hepatic lesions caused by the entrapment of eggs that
accidentally are released from the worms and pass via the hepatic
portal system to the liver. This phenomenon is particularly prominent
in the human.

The epidemiology is totally influenced by water as a medium for the

free-swimming larvae and the intermediate host snail. Percutaneous
infection implies that the potential hosts, e.g. pig and man, must
have close water contact for shorter or longer periods of time.
Fertilization with human or animal sludge to crops that are grown
under humid, wet conditions (rice, certain vegetables) favour heavy
transmission. The relative importance of human or animal excreta to
infection of respectively humans and animals (e.g. pigs) is poorly
determined.

2.6 CESTODES

The principal larval cestodes (metacestodes) found in pigs are those

of Taenia solium (Cysticercus cellulosae), Taenia hydatigena
(Cysticercus tenuicollis) and Echinococcus granulosus (hydatids), cf.
Table 2.2. The most important cestode specifically related to the pig
is Taenia soliumICysticercus cellulosae.

Fig. 2.11 shows the life cycle of Taenia solium, which is highly
prevalent in Latin America, India, and many parts of South-East Asia.
28 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Man is infected by eating

raw or inadequately Cysticerci also may establish in
cooked meat. The man, in particular in the central
adult tapeworm nervous system
develops in the
small intestine
at month 2-3
Gravid segments,
containing infective
eggs, in human faeces

An infective
egg, containing
an onchosphere

Ingestion
of eggs
Cysticerci develop in striated muscles,
especially in the heart, tongue,
diaphragm and masseter
Illustration by Wm. P Hamilton CM!

FIGURE 2.11 Taenia solium life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 29

Man as the final host harbours the adult worm in the small intestine.
The tapeworm segments containing infective eggs, are excreted with
faeces. The eggs are ingested by pigs, and onchospheres hatch in
the small intestine. They penetrate the gut wall and via the blood
they are distributed all over the body, but the metacestodes
(cysticerci) almost exclusively develop in the striated muscles, in
particular in the heart, tongue, masseters and diaphragm. They
appear as fluid-filled cysts, 6-8 mm, containing an invaginated
scolex. Man becomes infected through consumption of raw or
inadequately cooked pork meat, and the adult tapeworm is developed
in the small intestine after 2-3 months. A peculiar and severe feature
is occasional establishment in man of metacestodes, in the form of
cerebral cysticercosis, a condition caused by the parasite in the
central nervous system. This may occur either from the ingestion of
tapeworm eggs, or possibly more likely from adult worms in the gut,
which release onchospheres to the stomach by reverse peristalsis.
This may be called autoinfection.

Clinical disease is inapparent in pigs harbouring the metacestodes

and in man with adult tapeworms. However, if man is infected with
metacestodes, e.g. in the form of cerebral cysticercosis a range of
severe neurological disorders may follow.

The epidemiology depends primarily on the close association of rural

pigs with man, in particular pigs with unrestricted access to human
faeces. As mentioned, man may acquire cerebral cysticercosis from
ingesting eggs, e.g. adhering to vegetables fertilized with night soil
or sewage sludge.

Among the other tapeworm infections, Echinococcus granulosis

established as hydatid cysts in liver and lungs, has great zoonotic
importance and severe implications for human health. Usually
infections in ruminants play a more important role than infections in
the pig (see Hansen and Perry, 1994).
30 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Cystacanths develop to
maturity in the small
intestine at month 2-3

Ingestion of
infected dung
beetles

..4.00, egg
Em bryonated
in faeces
Cystacaths,
which are (contains a
infective to fully infective
pigs, develop acanthor)
within the dung
beetle Faeces with eggs
are ingested by a
dung beetle larva
Illustration by Wm P Hamilton CMI

FIGURE 2.12 Macracanthorhynchus hirudinaceus life cycle

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 31

2.7 ACANTHOCEPHALA

Fig. 2.1 2 shows the life cycle of Macracanthorhynchus hiruchnaceus,

the only important species of this phylum. It is found worldwide
except for certain temperate areas, e.g. Western Europe. This large
worm (up to 65 cm) lives in the small intestine. Embryonated eggs
are passed with faeces, and after ingestion by dung beetles an
infective stage, the cystacanth, is developed in approximately 3
months. Pigs become infected by eating dung beetles, and the
cystacanth develops in the small intestine to the adult stage in 2-3
months.

Infections are usually subclinical, but heavy infections may cause gut
lesions, and in rare cases penetration of the gut wall, and fatal
peritoneitis.

The epidemiology is determined by pigs' contact with beetles and

biotic factors favouring beetles. Eggs may survive for many years in
the faeces/soil environment. Pig is the only final host.
 32 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

TABLE 2.1 An overview of gastro-intestinal helminth species in pigs

Organ Helm nth Life cycle Transmission Final hosts

Oeso- Gongylonema pulchrum indirect I: dung beetles etc. wide host

phagus range

Stomach Trichostrongylus axei d rect L3 orally ruminants +

horse

Hyostrongylus rubiclus direct L3 orally pig

011ulanus tricuspis direct? 'vomiting'? Felids and

canids

Ascarops strongylina
Physocephalus sexalatus indirect I: dung beetles mainly pig
Simondsia paradoxa

Gongylonema pulchrum indirect I: dung beetles etc. very wide host

range

Gnasthostoma hispidum indirect I-1: aqua. crustaceans wide range of

G.doloresi 1-2: small vertebrates domestic
animals

Small Trichostrongylus vitrinus direct L3 orally primarily

Intestine T.colubritorrnis ruminants

Globocephalus direct L3 orally domestic

urosubulatus animals
G.Iongemucronatus

Strongyloides ransomi direct L3 orally, colostral, Pg

percutan

Ascaris suum direct L3 in eggs mainly pig

Trichinella spiralis indirect L1 in striated muscles wide host range

Fasciolopsis buski indirect I: fresh water snail man (primarily)

Macracanthorhynchus indirect I: dung beetles pig

hirudinaceus

Large Oesophagostomum
Intestine dentatum' direct L3 orally Pig
0.quadrispinulatum 2
0.brevicaudum

Trichuris suis direct Ll in eggs pig

0.granatensis and 0.georgianum regarded as variant forms of 0.dentatum
0.longicaudum synonymous with 0.quadrispinulatum
I: Intermediate host
 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 33

TABLE 2.2 An overview of non-gasto-intestinal helminths in pigs

Organ Helminth Life cycle Transmission Final hosts

Lungs Metastrongylus apri' ind rect I: earthworms (L3) Pig

M.salmi
M.pudendotectus

Blood system Schistosoma japonicuni indirect 1: freshwater snails large variety of

S.mansoni mammals
Sincognitum

Kidneys Stephanurus dentatus direct L3: oral, percutan pigs

(transport:earthworm)

Liver Taenia hydatigena indirect eggs orally can ds

(metacestodes) 2

Echinococcus indirect eggs orally canids

granulosus (meta-
cestodes, hydatids)

Dicrocoelium indirect 1-1: landsnail primarily

dendriticum 1-2: ants ruminants

Fasciola hepatica indirect I: freshwater snails large variety of

mammals

Opisthorchis 110 verca indirect 1-1: aquatic sna Is fish eating

1-2: fish mammals

Ascarid migrating larvae direct L3 in eggs pigs

Schistosoma spp. (eggs) indirect I: freshvvater snails large variety of

mammals

Çonnective Taenia hydatigena indirect eggs orally canids

tissue (cysticercs)2

Muscles Trichinella spiralis indirect Ll in striated muscles very w de host

(larvae) range

Taenia solium indirect eggs orally man

(metacestodes)3
M.elongatus synonymous with M.apri
The metacestodes of T.hydatigena are called Cysticercus tenuicollis
The metacestodes of T.solium are called Cysticercus cellulosae
I: Intermediate host
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 35

CHAPTER 3

FAECAL EXAMINATIONS FOR PARASITES

3.1 INTRODUCTION

Examination of faecal samples for helminth eggs is an easy way to

diagnose many helminth infections and to get an impression of the
infection level. All helminths which use the pig as a final host, must
find a way for their eggs to become available for the next host. Most
often, the eggs simply pass with the faeces, and this applies to the
large majority of helminths in the gastro-intestinal tract and
associated organs, such as the liver and the lungs, and even
schistosomes living in the mesenteric veins use the same outlet to
the surrounding world.

However, few helminths in the pig do not make use of faeces for egg
transport. Thus, the eggs of Stephanurus dentatus pass with the
urine, while the larvae of Trichinella spiralis migrate directly to the
striated muscles to become encysted there, and infective larvae of
011ulanus tricupis are apparently transmitted in vomit.

Newly deposited pig faeces may contain unembryonated eggs or

eggs with well-developed embryos (e.g. Metastrongylus spp. and
Strongyloides ransom), but no hatched larvae. Therefore, a complete
examination of fresh porcine faeces does not include the Baermann
technique for isolation of parasite larvae.

This chapter presents laboratory techniques suitable for identifying

and quantifying parasite infections on the basis of examination of
faecal material.
36 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

3.2 COLLECTION OF FAECAL SAMPLES

It is important that the examinations are carried out on fresh faeces,

as eggs of many helminths hatch within 24 hours (e.g.
Oesophagostomum spp.) or eyen after 8 hrs (Strongyloides ransom')
at room temperature, and the young larvae will not be detected by
the standard laboratory techniques.

Equipment

Plastic gloves (cheap plastic gloves are often preferable for

the more expensive latex gloves)

Marking pen (waterproof)

Plastic bags

Insulated cooling box (storage temperature: 0-8°C), if the

transport time to the laboratory exceeds 1-2 hrs

3% formalin and plastic containers with tight lids, if a long

transport time is expected, and a cooling box is not available

Procedure

Faecal samples are preferably collected directly from the rectum, to

ensure that they are completely fresh. This will also allow for
registration of the pigs age, sex, reproductive status etc., and
repeated samples from the same individual may be avoided.

If rectal samples cannot be obtained, freshly deposited faeces may

be collected from the pens/pastures.

The samples may be stored in the plastic gloves by turning their

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 37

inside out. Each sample should be unambiguously labelled with

animal identification, date and localization in waterproof ink directly
on the plastic glove. The amount of faecal matter required depends
on the analyses, but at least 4 g is needed for most egg count
procedures, and >20 g is preferred if more than one kind of analysis
is needed, or if an unexpected result must be confirmed by repeated
analyses.

The samples are gathered in larger plastic bags. If the transport time
to the laboratory is expected to exceed 1-2 hours (depending on
temperature), the samples should be packed in a cooling box to avoid
hatching of the eggs. The storage temperature should be 0-8°C, and
care should be taken to avoid freezing, as this may damage the eggs
and invalidate later results. If larval cultures are to be done, the
faecal samples may not be cooled, as even 24 hours at 5°C may
interfere with larval development.

When a cooling box is not available, the samples may be placed in

plastic containers with tight lids, and 3% formalin should be admixed
to the faeces (approx. 1 ml formalin to 4 g faeces). This will preserve
the sample and the parasite eggs, but it should be noted that
quantitative egg counts will not be completely correct, and that
formalin-fixed faeces cannot be used for faecal cultures.

In the laboratory, the samples for egg counts should immediately be

placed in a refrigerator (approx. 4°C) until they are processed.
Samples may be stored at this temperature for more than 3 weeks
without significant changes in egg counts. If faecal cultures are to be
set up, a storage in a refrigerator cannot be recommended at all, but
the cultures should be set up immediately. Note again that faecal
samples should never be kept in a freezer.

3.3 QUALITATIVE TECHNIQUES FOR FAECAL EXAMINATIONS

A large number of different procedures is available for demonstrating

 38 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

eggs in faeces of pigs. Three methods will be described below, all of

them providing results that are only qualitative (or, at the highest,
semiquantitative), because the egg recovery may be rather low and
highly variable.

The most widely used principle for concentration of parasite eggs is

flotation. As most nematode eggs (and coccidia oocysts) have a
specific gravity which is lower than that of plant residues in the
faeces, the eggs may be separated from other faecal particles by
mixing the faeces with a fluid in which the eggs flotate, while the
plant particles sink.

Unfortunately, the specific gravity of helminth eggs varies. While

most nematode eggs will flotate in saturated NaCI, some nematode
species have eggs which will flotate only in fluids with higher
specific gravities, such as saturated MgSO4 or saturated
NaCI + glucose (as used below). Among porcine helminths, this
applies to, for example, Metastrongylus spp. and to some degree
Trichuris suis, and therefore it is recommended to use only one of
the heavy flotation fluids.

Trematode eggs, on the other hand, are in general so heavy that the
flotation principle does not work at all. Therefore, these eggs are
concentrated by different sedimentation techniques, which, however,
are not fully efficient and highly variable.

3.3.1 Test tube flotation

This is a simple qualitative flotation technique for the detection of

nematode eggs (and coccidia oocysts) in the faeces.

Equipment

2 beakers or plastic containers (disposable or recycling)

. Balance or a precalibrated teaspoon (3 g)
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 39

Flotation fluid: Saturated NaCI with 500 g glucose per litre

Measuring cylinder or another container graded by volume
Stirring device (fork, tongue depressor)
Nylon tea strainer or a single layer of cotton gauze
10-12 ml test tube
Test tube rack
* Coverslips and microscope slides
*
Microscope with 40-100x magnification

Procedure

The Test Tube Flotation procedure is illustrated in Fig. 3.1.

Transfer approximately 3 g faeces (weigh out or measure with

precalibrated teaspoon) to plastic container 1.

Pour 50 ml flotation fluid into plastic container 1 by means of the

measuring cylinder.

Mix faeces and flotation fluid thoroughly with a stirring device.

Immediately after stirring, pour the faecal suspension through a tea

strainer or a single layer of cotton gauze into plastic container 2.

Discard the retained faecal debris, and immediately pour the sieved
faecal suspension from plastic container 2 into a test tube, which is
placed in a precisely vertical position in a test tube rack.

The test tube should be topped up with the faecal suspension, so

that it has a convex meniscus at the top. Place a coverslip on the top
of the test tube.
 3 g Faeces
50 ml Flotation fluid

Discard
retained
Stir thoroughly debris

A single
layer of
gauze

Carefully lift off

the coverslip and afiL
transfer it to a
microscope slide

Immediately pour the

suspension into a test tube

Leave for
Examine at 40-100x
20 min. with
magnification a coverslip
I44--
CX0.1

FIGURE 3.1 Test Tube Flotation

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 41

Leave the test tube for about 20 minutes. The helminth eggs will
flotate and thus accumulate just beneath the coverslip.

Lift off the coverslip vertically from the tube together with the
adhering flotation fluid. Some of the accumulated helminth eggs will
now be within the adhering fluid, and the transfer of the coverslip
must be done very carefully in order to retain as many eggs as
possible. Place the coverslip on a microscope slide, and examine the
sample at 40-100x magnification in a microscope.

Note: this method is qualitative and not quantitative.

3.3.2 Simple flotation

The principle for simple flotation is identical to that of the above test
tube flotation. The only difference is that the flotation takes place in
a beaker.

Equipment

2 beakers or plastic containers (disposable or recycling)

Balance or a precalibrated teaspoon (3 g)
Flotation fluid: Saturated NaCI with 500 g glucose per litre
Measuring cylinder
Stirring device (fork, tongue depressor)
Nylon tea strainer or a single layer of cotton gauze
Test tube (dry)
Coverslips and microscope slides
Microscope with 40-100x magn fication
 3 g Faeces
50 mi Flotation fluid

Discard
retained
debris
Stir thoroughly

Lift the test tube and

i A single
layer of
gauze

allow adhering drops to

be deposited on the
slide Press a test tube
Air' to the buttom

i
Coverslip
on
*--
Examine at 40-100x Leave
magnification for 15 min.

FIGURE 3.2 Simple Flotation

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 43

Procedure

The Simple Flotation procedure is illustrated in Fig. 3.2.

Transfer approximately 3 g faeces (weigh out or measure with

precalibrated teaspoon) to plastic container 1.

Pour 50 ml flotation fluid into plastic container 1 by means of the

measuring cylinder.

Mix faeces and flotation fluid thoroughly with a stirring device.

Immediately after stirring, pour the faecal suspension through a tea

strainer or a single layer of cotton gauze into plastic container 2.

Discard the retained faecal debris and leave the container undisturbed
on the table for 10-15 minutes, during which helminth eggs will
flotate and thus accumulate in the surface layer.

Press a dry test tube to the bottom of the faecal suspension, while
a microscope slide is ready for use. Some of the helminth eggs,
accumulated in the surface layer of the suspension, will now be
adhering to the test tube.

In one movement, the test tube is carefully lifted out of the fluid, and
adhering drops of faecal suspension are transferred to the
microscope slide. The bottom end of the test tube must rest on the
slide for several seconds for the drops to run off.

Place a coverslip on the microscope slide, and examine the sample

at 40-100x magnification in a microscope.

Note: this method is qualitative and not quantitative.

44 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

3.3.3 Sedimentation (Trematode eggs)

As mentioned previously, trematodes eggs have high specific

gravities, and therefore the eggs do not flotate in common flotation
fluids, but they may be concentrated by sedimentation. Among
porcine trematodes, this applies to Fasciola hepatica, Dicrocoelium
dendriticum, Fasciolopsis buski and Opisthorchis noverca, while
special techniques are elaborated for Schistosoma spp. (not dealt
with in the present guidelines).

The technique described below is merely a combination of washing

and sieving of faeces, to remove the smallest and the largest faecal
particles. The technique utilizes the high gravity of the eggs, which
facilitates their sedimentation in beakers with steeply sloping sides.

Equipment

1 or 2 beakers or plastic containers (disposable or recycling)

Balance or a precalibrated teaspoon (3 g)
Measuring cylinder
Stirring device (fork, tongue depressor)
Nylon tea strainer or a single layer of cotton gauze
Test tubes and test tube racks. These may preferably be
replaced by a conic sedimentation beaker of glass, but these
are rather expensive (the conic beakers are also very useful
in the Baermann technique described later in this chapter).
Fig.3.3 illustrates the technique using conical beakers
Methylene Blue (1 % solution) or Malachite Green (1 %
solution)
Coverslips and microscope slides
Microscope with 40-100x magnification
The Epidemiology, Diagnosis and Control of Helm nth Parasites of Swine 45

Procedure

The Sedimentation Procedure is illustrated in Fig. 3.3.

Transfer approximately 3 g faeces (weigh out or measure with
precalibrated teaspoon) to plastic container 1.
Pour 50 ml tap water into plastic container 1 by means of the
measuring cylinder.
Mix faeces and tap water thoroughly with a stirring device.
Immediately after stirring, pour the faecal suspension through a tea
strainer or a single layer of cotton gauze into a conic sedimentation
beaker, and fill up the beaker with tap water. Alternatively: pour the
faecal suspension through a tea strainer or a single layer of cotton
gauze into plastic container 2, and transfer approx. 10 ml of the
filtered suspension into a test tube which is placed in a test tube
rack.

Allow the faecal particles, including the trematode eggs, to sediment

for 10 minutes.
Remove the supernatant carefully in one steady movement (conic
sedimentation beakers) or with a pipette (test tube sedimentation).
Care should be taken not to resuspend the sediment during the
process. The supernatant is discarded.
Resuspend the sediment in tap water. The sedimentation beaker
should be almost filled up (or alternatively: the test tube should be
almost filled up).
Allow the faecal particles, including the trematode eggs, to sediment
for 10 minutes.
Remove the supernatant carefully in one steady movement (conical
sedimentation beakers) or with a pipette (test tube sedimentation).
Care should be taken not to resuspend the sediment during the
process. The supernatant is discarded.
 Add extra Discard
tap water retained
debris

3 g Faeces
50 ml tap water
A single
layer of
gauze

Stir thoroughly

Leave for 10 min.

Transfer a few
drops to each of Remove Leave for Remove supematant and
a series of micro- supematant 10 min. resuspend In tap water
scope slides

Add dye
4--
Examine at 40-100x
magnification

FIGURE 3.3 Sedimentation

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 47

Add 1-2 drops of Methylene Blue or Malachite Green. Both dyes will
stain the faecal particles deeply blue/green, while the trematode eggs
remain unstained. This contrast staining allow the brownish eggs to
be discovered more easily. Use a weaker dye solution if the staining
is too heavy.

Transfer a few drops of the stained sediment to a microscope slide

with a pipette, place a coverslip on the microscope slide, and
examine the sample at 40-100x magnification in a microscope.

Repeat the last step until all the sediment has been examined. If
nematode eggs are present in the faecal sample, some of them may
be found in the sediment, but the recovery rate is very low, and
sedimentation cannot replace flotation where nematodes are
concerned.

3.4 QUANTI VATIVE TECHNIQUES FOR FAECAL EXAMINATIONS

The qualitative flotation techniques above, which are used for

nematode eggs (and coccidia oocysts), have been elaborated to
become quantitative, when the eggs are allowed to flotate in a
special counting chamber, called the McMaster chamber. Many
modifications exist, and a Simple McMaster Technique and slightly
more elaborated Concentration McMaster Technique will be
presented in the following.

3.4.1 Simple McMaster technique

No concentration of eggs is carried out in this procedure, and the

sensitivity is 50 eggs per gram faeces.
48 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Equipment

2 beakers or plastic containers (disposable or recycling)

Balance

Measuring cylinder
Flotation fluid: Saturated NaC1 with 500 g glucose per litre
Stirring device (fork, tongue depressor)
Nylon tea strainer or a single layer of cotton gauze
Pasteur pipettes and rubber teats
McMaster counting chamber. Several designs exist, but the
traditional design with two counting fields and a
permanently fixed, solid upper glass with a counting grid on
the inside is recommended. This design may be found in
expensive models of glass and cheap models of plastic. If
many samples are to be examined by skilled personnel, the
glass chambers are recommended, as they do not I3ecome
scratched so easily, and as the visual fields in general are
more clear. On the other hand, the plastic chambers are
recommended when only a low number of samples is to be
examined, or when students and trainees are learning the
technique. The plastic chambers will quickly become
scratched, but they are so cheap that they may then be
discarded; or, alternatively, the scratched outer surface may
be polished (e.g. by a watchmaker)
Filtering paper cut into approx. 1 cm wide strips
Microscope with 40-100x magnification

Procedure

The Simple McMaster Technique is illustrated in Fig. 3.4.

4.0 g Faeces
56 ml Flotation fluid Discard
retained
debris

Stir thoroughly

A single
layer of
gauze

.A1111011111V
Examine at 40-100x Fill a McMaster chamber
magnification in both compartments
and leave for 3-5 min. Immediately take
out a subsample
C rape

FIGURE 3.4 Simple McMaster Technique

50 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Weigh out 4.0 g faeces and transfer it to container 1. The container

should be unambiguously labelled (disposable containers may be
labelled in waterproof marking ink).

Add 56 ml flotation fluid by means of the measuring cylinder. If

another amount of faeces is weighed in the former step (i.e. more or
less than 4.0 g), the volume of flotation fluid should be adjusted
correspondingly (ratio: 14 ml flotation fluid to 1.0 g faeces). This
ratio ensures that 15 ml of the resulting faecal suspension
correspond to 1.0 g faeces.

Mix faeces and flotation fluid thoroughly with a stirring device.

Pour the faecal suspension through a tea strainer or a single layer of

cotton gauze into container 2, immediately after stirring. The retained
debris is discarded. If disposable containers are used, container 2
may be placed into container 1, which is still labelled.

A subsample is taken with a pasteur pipette immediately after the

filtering procedure, when the suspension is still well mixed.

Fill both sides of the McMaster counting chamber with the faecal
suspension. Be careful to avoid air bubbles.

Leave the filled McMaster chamber to rest on the table for 3-5
minutes before counting (minimum 3 minutes to allow all eggs to
flotate, and maximum 10 minutes, as some eggs may be distorted in
the flotation fluid).

Count the number of eggs in both counting fields, and calculate the
number of eggs per gram of faeces by multiplying the number of
eggs by 50 (see Section 3.4.3 Counting the McMaster chamber).

After counting, the McMaster chamber should be washed under a

stream of tap water, shaken to remove most of the water, and dried
with a cotton cloth on the outside and with a strip of filter paper
inside the chamber.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 51

3.4.2 Concentration McMaster technique

This technique is slightly more complicated than the Simple

McMaster Technique, but the recovery of eggs is higher and the
sensitivity is better (20 eggs per gram of faeces). This technique is,
therefore recommended if a centrifuge is available. Furthermore, the
procedure may be more flexible when many samples are handled
simultaneously.

Equipment

2 beakers or plastic containers (disposable or recycling)

Balance

Measuring cylinder
Stirring device (fork, tongue depressor)
Nylon tea strainer or a single layer of cotton gauze
Test tube with 4 ml and 10 ml marks
Test tube stopper
Test tube rack
Centrifuge
Flotation fluid: Saturated NaCl with 500 g glucose per litre
Pasteur pipettes and rubber teats
McMaster counting chamber
Filtering paper cut into approx. 1 cm wide strips
Microscope with 40-100x magnification
52 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Procedure

The Concentration McMaster Technique is illustrated in Fig. 3.5.

Weigh out 4.0 g faeces and transfer it to container 1. The container

should be unambiguously labelled (disposable containers may be
labelled in waterproof marking ink).

Add 56 ml tap water by means of the measuring cylinder.

Some people prefer to weigh out between 4.0 and 6.0 g faeces in
the first step, and then add the corresponding volume of tap water
(the ratio should always be 14 ml tap water to 1.0 g faeces. This
ratio ensures that 15 ml of the resulting faecal suspension
corresponds to 1.0 gram of faeces).

Mix the faeces and the tap water thoroughly with a stirring device.

Allow the faecal suspension to rest for 30 minutes at room

temperature, and again mix the faeces and the tap water thoroughly
with a stirring device.

The 30 minutes' soaking and the repeated mixing ensure that even
firmer pieces of faeces will be completely dissolved, but the step
may not be necessary if the faeces are soft diarrheia. A similar
soaking step cannot be used in the previously mentioned flotation
techniques, as faeces were dissolved directly in the flotation fluid,
which may distort the eggs during a prolonged exposure.

Pour the faecal suspension through a tea strainer or a single layer of

cotton gauze into container 2, immediately after stirring, and discard
the retained debris. If disposable containers are used, container 2
may be placed into container 1, which is still labelled.
 4.0 g Faeces
56 ml Tap water
Discard
retained
debris

Rest for 30 min.

/-----. 4L
). A single
layer of
gauze
Stir thoroughly
ill
Stir thoroughly

Resuspend Remove Immediately pour

sediment in supernatant 10 mi of the sus-
flotation fluid pension into a test
(total volume tube
=4m1) t
Possible
/Fill a McMaster chamber storage
in both compartments
at 5°C /Centrifuge at
and leave for 3-5 min.
1200 RPM
for 5-7 min.
A111211V <k-
Examine at 40-100x
magnification
C rapt

FIGURE 3.5 Concentration McMaster Technique

 54 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Immediately after the filtering procedure, pour faecal suspension into

a test tube to the 10 ml mark. As 15 ml faecal suspension
represents 1 g faeces, the 10 ml suspension will represent 2/3 g.

Centrifuge the test tube for 5-7 minutes at 1200 RPM (revolutions
per minute).

Remove the supernatant with a pipette or another vacuum device,

but be careful not to resuspend the sediment. Correctly done, the
sediment still represents 2/3 g faeces.

At this step it is possible to interrupt the procedure by stoppering the

tube and storing it in a refrigerator (approx. 4C) for up to 7 days
without any significant reduction in the egg counts. If many samples
are to be handled simultaneously, this possibility for storage makes
the laboratory work more flexible and rational, as 50-100 samples
may be sieved and centrifuged in one step, whereafter they are
stored until they are counted one by one.

A similar storage is not possible in the simple McMaster technique,

as the faeces are dissolved directly in the flotation fluid.

Shortly before counting, flotation fluid is added to the 4 ml mark (i.e.

the total volume of faecal sediment and flotation fluid is 4.0 ml).
These 4 ml now represent 2/3 g faeces.

Resuspend the sediment very carefully, using a pasteur pipette

several times. Avoid making bubbles in the suspension, as these will
make the egg counts less reliable.

Fill both sides of the McMaster counting chamber with the faecal
suspension, immediately after resuspension of the sediment. Be
careful to avoid air bubbles.

Leave the filled McMaster chamber to rest on the table for 3-5
minutes before counting (minimum 3 minutes to allow all eggs to
flotate, and maximum 10 minutes, as some eggs may be distorted in
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 55

the flotation fluid). Count the number of eggs in both counting fields
and calculate the number of eggs per gram of faeces by multiplying
the number of eggs by 20 (see Section 3.4.3 Counting the McMaster
chamber).

After counting, the McMaster chamber should be washed under

running tap water, shaken to remove most of the water, and dried
with a cotton cloth on the outside and with a strip of filter paper
inside the chamber.

3.4.3 Counting the McMaster chamber

The filled McMaster chamber must rest on the table for at least 3-5
minutes to allow all eggs to flotate, i.e. accumulate just below the
upper glass of the chamber. It is important that the eggs have
enough time to flotate in order to avoid underestimating the egg
count. On the other hand, the sample becomes less clear, and some
egg types may be distorted and sink, if the sample rests for too long
(15-20 minutes) in the chamber before microscopical examination.
This can be prevented by keeping the filled McMaster slides in the
refrigerator prior to counting.

Focus on the counting grid (or parasite eggs) and count the different
nematode eggs within the engraved area of both sides of the
chamber. Skilled personnel often prefer a 4x10 magnification, while
all others are recommended to use a 10x10 magnification until they
are completely familiar with all types of eggs. If coccidia oocysts are
counted, a 10x10 magnification should always be used, as porcine
coccidia have rather small oocysts (as small as 12 pm, i.e. much
smaller than the oocysts of most ruminants).

When counting the engraved areas, the general rules for counting
should be followed: all eggs inside the grid should be counted plus
all eggs touching two sides of the grid (e.g. the upper and the left
borderlines), while excluding all eggs touching the two other sides of
the grid (e.g. the lower and the right borderlines).
56 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Every type of nematode egg should be counted separately.

The distance between the upper and the lower glass of the McMaster
chamber is 0.15 cm, and the two counting fields each measure 1x1
cm. Therefore, the faecal suspension under the two counting fields
has a volume of 2 x 0.15 ml = 0.3 ml.

In the Simple McMaster Technique, 15 ml faecal suspension

represents g faeces, and therefore 0.3 ml represents 1/50 g
1

faeces. The number of eggs per gram of faeces can now be

calculated as follows: The total number of eggs in both sides of the
chamber should be multiplied by 50. This gives the number of 'Eggs
Per Gram of faeces', usually abbreviated EPG.

In the Concentration McMaster Technique, 4 ml of the final faecal

suspension in the test tube represents 2/3 g faeces, and therefore
the counted volume of 0.3 ml faecal suspension represents 1/20 g
faeces. The number of eggs per gram of faeces (EPG) can now be
calculated by multiplying the total number of eggs in both sides of
the chamber by 20.

Example: 18 eggs are counted in side 1 of the chamber, and 22 eggs

are counted in side 2.
If the Simple McMaster Technique has been used, EPG =
(18 + 22)x50 = 2000.
If the Concentration McMaster Technique was used, EPG
(18 + 22)x20 = 800.

3.5 FAECAL CULTURES

Helminth eggs of the strongylid type (see Section 3.6 Identification

of eggs and larvae) cannot be identified to species. They may belong
to Oesophagostomum spp., Hyostrongylus rubidus or, very rarely,
Trichostrongylus spp. In order to differentiate between these
helminths, it is necessary to make faecal cultures in which the eggs
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 57

will hatch and the larvae develop to the infective third stage (L3).
These L3 larvae may then be identified to genus by microscopical
examination (see Section 3.6 Identification of eggs and larvae).

Several simple techniques for culturing L3 larvae exist. The only

precautions to be taken are that the physico-chemical conditions
should be so favourable that eggs of all species are able to develop
into larvae, and that a large number of larvae should be identified in
order to detect all species present. The latter precaution is necessary
for the detection of Hyostrongylus rubidus, since pigs infected with
H.rubidus most often also harbour Oesophagostomum spp., which
has a much higher fecundity.

It is important that the faecal samples are collected rectally, as

faeces picked up from the ground is very often contaminated with
free-living nematodes. The latter will multiply extremely rapidly in the
cultures and totally outnumber the parasite larvae, which may then
be difficult to find. Furthermore, the faecal samples should be very
fresh, as even a few days of storage in a refrigerator may
significantly reduce the larval development of some species.

One simple technique for faecal cultures and for harvest of larvae
(Baermann technique) is described below.

Equipment

1 beaker or plastic container (disposable or reusable)

Balance

Stirring device (fork, tongue depressor)

Vermiculite (an inert absorbing material which provides the
cultures with a fine structure)
Tap water (not chlorinated)
58 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Humidity chamber (e.g. a plastic tray within a large plastic

bag, or a tray with a lid)
Little stick and a rubber band
Nylon tea strainer or a double layer of cotton gauze
Conical sedimentation beaker (alternatively, other kinds of
beakers may be used, but they do not concentrate the larvae
as well because of the larger bottom area)
Pipettes (e.g. 2 ml)
Microscope slides. The best slides are large and equipped
with a paraffin bank surrounding the sample area
Microscope with 10-40x magnification

Procedure

The Faecal Culture technique is illustrated in Fig. 3.6.

Weigh out 10 g faeces and transfer it to a plastic container.

Add 4 g vermiculite (vermiculite is very light, so its volume is

considerably larger than that of the faeces) and stir thoroughly.

Add tap water if necessary, and then stir thoroughly. The volume of
water depends on the consistency of the culture. Dry faeces needs
more water than normal faeces, and if the faeces is like diarrhoea, it
may be necessary to add more vermiculite to absorb the water
instead of adding water. Eventually, the culture should end up with
a consistency where it does not appear wet, but still has a 100%
relative humidity.

If non-chlorinated tap water is not available, it is necessary to use

deionized or distilled water.
 Positive McMaster-test

Faeces (10 g)
Vermiculite (4 g) Stir thoroughly
Tap water (without chlorine)
Incubate in a humidity
chamber for 10-14 days at 20°C
Stir every 1-2 days

+
Transfer to a
microscope slide Ak--------\ 2 layers of gauze
with a paraffin mounted on sticks
bank

Leave to
Baermannize
for 24 hours
C irvd

FIGURE 3.6 Faecal Culture

60 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

The beaker is placed in a humidity chamber in which the bottom is

covered with water, to ensure a relative humidity of 100%.

The humidity chamber is stored at room temperature (20-21°C) or in

an incubator at similar temperature for 10-14 days, by which time
the larvae should have reached the L3 stage. If the cultures are kept
at lower temperatures (min. temperature 15°C), the incubation period
should be considerably increased.

Stir the cultures every 1-2 days, and add water if they have become
too dry. Stirring may help to break up fungal hyphae and thereby
keep fungal growth at an acceptable level.

After the incubation period, the larvae are recovered by a Baermann

technique. First, place the culture on a double layer of gauze.

Wrap up the culture and fasten it to a small stick of wood by means

of a rubber band.

Submerge the culture in tap water in a conical sedimentation beaker

and leave it there until the next day. During the night, the large
majority of larvae will move out of the culture and sink to the bottom
of the beaker.

Harvest the larvae with a 2 ml pipette by placing your fingertip firmly

at the end of the pipette, move the tip of the pipette to the bottom
of the beaker, relieve the fingertip pressure from the pipette and
allow the larvae to be sucked up into the pipette. Do not suck by
mouth, as larvae may accidentally be sucked into your mouth.

Transfer the larvae to one or more microscope slides equipped with

a paraffin bank. At least 100 parasite L3 larvae (if available) should
be counted and differentiated under a microscope.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 61

3.6 IDENTIFICATION OF EGGS AND LARVAE

Helminth eggs found in pig faeces may have a characteristic

appearance, which permits an unambiguous identification, or they
may be of the strongylid type which cannot be identified before the
infective larvae have been developed in cultures.

Some of the most common eggs, and two types of infective larvae
developed from eggs of the strongylid type, are shown below (Figs
3.7, 3.8 and 3.9 a + b). As coccidia are quite common in pigs, two
types of oocysts are also shown (Fig. 3.9 c + d). Furthermore, some
characteristics are briefly listed in Table 3.1 .

Note that all photos are of the same scale and therefore may be
compared directly.

3.7 INTERPRETATION OF FAECAL EGG COUNTS

It is difficult to present guidelines for the interpretation of egg

counts. First of all, one should be aware of the possibility of false
positive and false negative egg counts. Secondly, the egg counts are
not clearly correlated with the worm burden. And finally, even if it
were possible to predict the worm burdens from the egg counts, one
would still have the problem of how to interpret this worm burden.

The latter topic is discussed in Chapter 4, where it is seen that the

interpretation of estimated worm burdens depends on the helminth
species and its properties, the specific host-parasite relationship (e.g.
the effect of host age immunity), and the management. Therefore,
only the false negative and positive results, and the relationship
between egg counts and worm burdens, will be discussed below.
62 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

FIGURE 3.7 Some of the most common 'strongylid' eggs in pig

faeces. The two upper figures show Oesophagostomum sp. (left) and
Hyostrongylus rubidus (right) which cannot be discriminated. The
two lower figures show the larvated, small egg of Strongyloides
ransomi (left) and the big egg of Stephanurus dentatus (right).
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 63

FIGURE 3.8 Some characteristic thickshelled eggs in pig faeces.

The two upper figures show eggs of Ascaris suum (left) and Trichuris
suis ( right) that are unembryonated when excreted, while the two
lower figures show newly excreted, embryonated eggs of
Metastrongylus sp. (left) and Macracanthorhynchus hirudinaceus
(right).
 The Epiclemiolow, I of H si tes of Swit

FIGURE 3.9 Two types of infective L3-larvae developed from eggs

of the strongylid type: the thick, sluggish, long-tailed larva of
Oesophagostomum sp. (upper, left) and the slender, 'swimming',
short-tailed Hyostrongylus larva. The two lower figures show the
small oocysts of Eimeria sp. (left), typically found in older pigs, and
the even smaller oocysts of lsospora suis (right), typically found in
piglets.
The Epidemiology, Diagnosis and Control of Helm n h Parasites of Swine 65

TABLE 3.1 A list of characteristics of the most common helminth eggs, L3-larvae
(developed from 'strongylid' eggs), and coccidia oocysts (to be continued)

Parasite width x length contents of the fresh egg morphological character st cs

(P)

EGGS

Gong ylonema 30-34 x 57-59 well-developed embryo elliptical egg, smooth surface
pulchrum
Hyostrongylus 31-38 x 60-76 32-64 cleavage cells thin-shelled, transparent egg,
rubidus smooth surf ace (strongylid')
Ascarops 22-26 x 41-45 well-developed embryo small, elliptical egg, slightly
strongylina flatte-ned at each pole,
smooth surface
Physocephalus 22-26 x 41-45 vvell-developed embryo small, elliptical egg, slightly
sexala tus flatte-ned at each pole,
smooth surface
Simondsia 15 x20-30 well-developed embryo oval or ellipsoidal
para doxa

Gnathostoma 39-42 x 72-74 one cell - early embryo thick-shelled egg,

spp.

Trichostrongylus 30-55 x 70-125 16-32 cells thin-shelled, transparent egg,

spp. smooth surface ('strongylid')
Globo cephalus 40 x 68-72 2 cells oval, thin-shelled, transparent
sPP. egg, smooth surface
Strongyloides 30-34 x 53-57 well-developed embryo small, thin-shelled,
ransomi transparent egg, smooth
surface
Ascaris 50-76 x 68-84 one cell (with granula) thick-shelled, rounded or
SUUM elliptical egg with brown
sculptured surface'
Macracantho- 65 x 110 acanthor larva with thick-shelled, oval egg with
rhynchus hook and spines brown colour
Oesophagostornurn 38-53 x 61-83 16-32 cleavage cells thin-shelled, transparent egg,
sop. smooth surface ('strongylid')
Trichuris 28-31 x 60-68 one cell (with granula) thick-shelled, barrel-shaped
suis egg, brown colour, a clear
knob protruding at each pole
Motastrongylus 38-45 x 43-64 well-developed embryo thick-shelled, elliptical egg,
sup. rough surface (small
mammillations)
Stephanurus 53-65 x 91-114 32-64 cleavage cells large, thin-shelled,
denta tus transparent egg in the urine
Dicrocoelium 30 x 45 a miracidium (some srnall, dark brown, operculate
dendriticum structures visible) egg, one side flattened
Fasciola hepatica 90 x 150 a miracidium (granular large elliptical yellow-brown,
mass) operculate egg
66 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Table 3.1 Continued

Parasite width x length (p) contents of the morphological

fresh oocyst characteristics

L3-larvae

Hyostrongylus 22 x 715-735' long, slender L3, the

rubidus posterior end of L., with a
small digitiform process, tail
of sheath short (moves
quickly and can swim)
Oesophagostomum 26 x 500-5322 shorter, thick L, no
spp. digitiform process, tail of the
sheath long and filamentous
(sluggish, cannot swim)

Coccidia oocysts
lsospora 17 x 20 1-2 cells small oval, transparent
suis oocyst

&merle 12-23 x 13-32 one cell spherical-elliptical

spp. transparent oocysts
1 Ascaris suum eggs may be atypical, as sterile eggs are much longer than wide, or
as they may be immature without the outer sculptured, brown layer. The latter eggs
are typically seen when mature females are expelled from the pig due to anthelmintic
treatment or self-cure.
2 Widths and lengths of the L, larva inside the sheath.

3.7.1 False negative and false positive egg counts

The detection of a helminth infection by examination of faecal

samples depends on the egg production of the parasites. If a pig has
recently been moved from a clean to a contaminated area, it may
harbour a heavy worm burden of young immatures which do not
produce eggs, and the faecal examination will be false negative. It
is evident that the likelihood of false negative results increases with
the duration of the prepatent period, e.g. Strongyloides ransomi
(prepatent period: 4-7 days) < Oesophagostomum spp. and
Hyostrongylus rubidus (18-21 days) < Ascaris suum and Trichuris
suis (7-8 weeks) < Stephanurus dentatus (9 months) (see Chapter
2 for prepatent periods).
The Epidemiology, Diagnosis and Control of Helminth Parasites of Sw ne 67

When the transmission rate is high, the pigs will respond

immunologically to the parasites, and a depression in egg output may
take place. This is particularly common when the helminth species is
strongly immunogenic. Ascaris suum is a good example, as often
only 20-40% of the growing pigs (and a much lower percentage of
older animals) harbour patent infections, even if all pigs are
continuously exposed to infective eggs, and their livers may be badly
affected by migrating larvae. But also lightly immunogenic species,
such as Oesophagostomum spp., may at extreme transmission rates
have ceased producing eggs temporarily, despite the fact that a
substantial number of adult (stunted) worms may be found in the
intestine.

False negative egg counts may also be found when a few adult
worms are all either males or females (unisex infections, common
when worm burdens in general are low, such as Ascaris suum), or
when adult worms have a low fecundity (e.g. Hyostrongylus
rubidus), or when the test used is not sufficiently effective.

The phenomenon of false negative egg counts is generally accepted,

whereas the opposite phenomenon, 'false positive egg counts', is
more or less overlooked. False positive egg counts may be found
when unembryonated helminth eggs are eaten by an uninfected host
and then passed with faeces. For such passage to occur, the eggs
must remain unhatched in the environment and during the intestinal
passage, and the host should eat faeces. Pigs are exceptionally good
candidates for false positive samples, as they may eat significant
quantities of faeces or contaminated soil, and as especially Ascaris
suum and Trichuns suis produce huge numbers of eggs which remain
unembryonated for considerable periods of time. As a rule of thumb,
Ascaris EPG <200 may be regarded as false positive (one single
mature female normally produces 400-800 EPG in growing pigs,
although considerable variations exist).
68 The Epidenniology, Diagnosis and Control of Helminth Parasites of Swine

3.7.2 The relationship between egg counts and worm burdens

When evaluating faecal egg counts, one should be aware of the fact
that EPG nearly always fluctuates considerably over time (weeks,
days), and that even in one large faecal sample (from one day)
countings on subsamples vary considerably, indicating that the eggs
are not evenly distributed in the faeces. Also the faecal consistency
may have some influence, as dry, hard faeces with relatively little
content of water will generally have higher EPG values than softer
and more watery faeces.

As mentioned above, pigs respond immunologically to helminth

infections, and the fecundity of the females seems to be the most
sensitive target. Thus, Ascaris females seem to have the highest egg
output when only one single pair is present, while the individual
fecundity decreases with increasing worm burdens. For less
immunogenic species, such as Oesophagostomum spp., there seems
to be a reasonably good correlation between EPG and the size of the
worm burden as long as this is below a certain level, whereas at
higher infection levels the individual egg laying becomes reduced or
may even stop completely.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 69

CHAPTER 4

POST-MORTEM DIFFERENTIAL WORM COUNTS

4.1 INTRODUCTION

Post-mortem worm counts provide a much more precise assessment

of parasite worm burdens than parasite egg counts or counts of
larval cultures, because eggs and to a certain degree larvae may not
always be identified to species, because there is no clear correlation
between egg output and worm burden, and because results from
counts of eggs and larvae may be false positive or false negative.

For worm counts in pigs, the following organs are required: The
gastro-intestinal tract from oesophagus to anus, the liver, the
kidneys, the lungs, a piece of the diaphragm and/or a piece of
musculus masseter. Frozen organs may be used for most analyses,
but fresh organs are much more convenient to handle, and isolation
of parasite larvae by incubation or baermannization can only be
carried out on fresh organs.

First of all, it is important to record all abnormalities and lesions,

bearing in mind that alternative causes of illness or death may occur.
Thereafter, adult and larval nematodes should be isolated, counted
and identified as described below. Light helminth infections in pigs
occur very often, and these infections do not in general -cause clinical
symptoms. Presence of several species at the same time
(polyparasitism) may precipitate disease due to additive or even
synergistic effects. Therefore it is important not only to identify the
species present, but also to assess the numbers of all species.

Suitable methods for differential worm counting under field or

laboratory conditions, using simple, easily obtainable and inexpensive
equipment, are described below.

Counts of gastro-intestinal helminths are most conveniently done first

by examination of the organs from the outside to count eventual
70 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

metacestodes, and thereafter by examination of the stomach, the

small intestine, the large intestine, the muscles, the lungs, the liver,
and the kidneys, separately.

4.2 THE CONTENTS OF THE STOMACH

The stomach may be infected by a number of species belonging to

Trichostrongyloidea (Hyostrongylus rubidus, Trichostrongylus axei,
011ulanus tricuspis) and Spiruroidea (e.g. Ascarops strongylina,
Physocephalus sexalatus, and others). Nearly all species are small,
slender worms (10-30 mm), and the larvae, and for some species the
adults, live in the gastric wall, e.g. in gastric nodules. Therefore,
careful observations should be carried out on the stomach wall,
mucus should be scraped off when examining the gastric contents,
and the gastric mucosa should be scraped off and digested if deeply
imbedded larvae are to be isolated and counted.

Equipment

One or two trays of about 40 x 60 x 15 cm. The precise

size is not important. Suitable plastic trays are easily
procurable. Rectangular trays are easier to pour from
Blunt knife
Pair of scissors

Two wide-mouthed plastic buckets with a capacity of at

least 12 litres. These are used to collect and mix the
contents of the different parts of the intestine before
subsamples are taken. The inner walls of the buckets
should be calibrated in litres
500 ml spoon for taking subsamples. It may be home-made
by mounting a beaker with a handle - the beaker should
contain exactly 500 ml when completely full
Sieve of 100-220 pm. A nylon net or another mesh of
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 71

equivalent size may be mounted with appropriate sides

Physiological saline, 0.9% (9 g of NaCI in 1 litre of tap
water)
Wash bottle
Jet stream of water. It may be a water pistol mounted on
a hose
Tight 500 ml containers, some with a calibrated scale on
the outside
Petri dishes, about 9 cm in diameter
Aqueous solution of iodine to stain the samples. A strong
solution may for example be: 80 g 12 and 400 g KI
dissolved in distilled water to a total volume of 1 litre
Aqueous solution of sodium thiosulphate to decolourize the
faecal matter, while the worms remain stained. The solution
could for example be 30%, i.e. 300 g sodium thiosulphate
crystals dissolved in water to a total volume of 1 litre
Light table/box. This may be a commercially available lamp
or a home-made construction. In principle, an electric bulb
is mounted inside a wide shallow box. The top of the box
is made of translucent white plastic or ground glass. When
samples in clear petri dishes are placed on top, the diffuse
white light shining up through the petri dishes provides a
strong contrast for the stained worms. Be careful to
construct the light board so that it is to some degree
waterproof
Dissection microscope
Needles or fine forceps to handle worms during counting
Bucket for handling of waste water and one bucket for
discarded organs may be useful
70% ethanol with 10% glycerol for eventual storage of
isolated worms
 72 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

. Small tubes (glass or plastic) with tight lids for eventual

storage of isolated worms

Procedure

The handling of Stomach contents is illustrated in Fig. 4.1.

Separate the stomach from the oesophagus and the duodenum. If

necessary, ligate the stomach from the duodenum with a string.

Place the stomach in a tray and open it along the greater curvature
with scissors, so that all contents fall into the tray. Empty the tray
into the 12-litre bucket.

Wash the empty stomach thoroughly with water, preferably a jet

stream, carefully cleaning between the folds of the mucous
membrane.

Pour the washing into the 12-litre bucket with stomach contents.

Scrape off the mucus of the stomach wall with a blunt knife.

Repeat the washing of the stomach wall one more time.

The mucus and the washing water is poured into the 1 2-litre bucket
with the stomach contents.

Open the oesophagus and wash it twice. Pour the washing water
into the stomach bucket with the stomach contents.

Total the volume of stomach contents in the bucket up to 10 litres

with water.

Stir vigorously until all food materials, mucus and water are well
mixed, and transfer one spoonful (500 ml, equal to 5% of the
volume) to the 100-220 pm sieve.
Cut open the ventricle and oesophagus Make the volume up to 10 L,
mix thouroughly, and take out
a spoonful (500 ml)
Stomach contents

Lf Mucus and washing water

Scrape off the mucus of
the stomach wall and
wash both organs twice
10 L mark

Possible Wash with a soft

storage Jet stream of water
Examine at a light table

Mix the solution Add iodine Flush the sample to

and transfer a sub- solution a 500 m.
sample to a petri and leave by means of
Add a few drops dish for 1 hour physiological saline
of thiosulphate in a wash bottle
solution C

FIGURE 4.1 Handling of Stomach Contents

74 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Wash the contents of the sieve well with a stream of water. It should
not be a very hard jet stream, as some worms may be damaged.

Pour the sample from the sieve into a 500 ml container by means of
physiological saline in the wash bottle. Be careful not to use
instruments and not even fingers to remove the sample from the
sieve, as the mesh is very susceptible to mechanical damage.

Repeat the last 3 steps one or more times if larger subsamples are
needed. For example, 10% or 20% subsamples are obtained when
2 or 4 spoonfuls, respectively, are transferred to the sieve and
washed.

Note: The best wash is obtained if the spoonfuls are sieved

separately, and correctly washed samples are much easier to count
than poorly washed samples.

Iodine solution is added to the sample in the container, and the

stomach contents are stirred well in order to secure that all particles,
including the worms, are stained adequately. The colour of the
sample should be dark brown. For a limited period of time the iodine
will work as a fixative. The sample should be allowed to stay in the
iodine for at least one hour before proceeding to the next step. If the
samples are stored for more than a few days, their colour should be
checked and more iodine added, if necessary.

At counting, a little volume of the sample should be poured into a

petri dish.

Add a few drops of sodium thiosulphate. This will decolourize the

faecal particles, while all large worms and to a certain degree
parasite larvae will remain dark. This decolouring process constitutes
a delicate balance: If a large surplus of sodium thiosulphate is added
and the parasites are small (many fourth-stage larvae), then the
worms will decolourize rather quickly, and thus loose their contrast
and 'disappear'. Therefore, the amount of sodium thiosulphate added
should be just enough to decolourize the sample (i.e. no surplus), and
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 75

the sample should be counted immediately.

Count the number of each nematode species present in the petri dish
placed on the light box, and repeat the two latter steps until the
entire sample is counted. Needles or forceps are convenient tools for
moving particles and worms around.

Note: The counting will be much safer if only small volumes of the
sample are poured into the petri dish, and the counting will proceed
faster, even if the number of petri dishes to be counted is increased.

In some cases it will be necessary to place the petri dish under a

stereo microscope to identify the worms to species.

Finally, the total worm count of the sample should be multiplied by

an appropriate factor to obtain the total worm burden in the
stomach. For instance, when the water suspension of the stomach
contents had a volume of 10 litres, and the subsample comprised 4
spoonfuls (each of 500 ml), then 2 litres out of 10 litres, equal to
20%, were counted, and thus the worm count of the subsample
should be multiplied by 5 in order to obtain the total worm burden in
the stomach.

4.3 THE STOMACH WALL

The isolation of helminth larvae from the stomach wall may be

carried out by one of two alternative methods: Incubation of the
stomach wall in physiological saline, or Digestion of the scraped-off
mucosa in pepsin-HCI. If the appropriate laboratory facilities are
available, the digestion method should be preferred. Each of these
methods may be carried out in conjunction with the isolation of adult
parasites from the stomach contents, and both can be used to
determine the numbers of immatures and hence the ratio of
immatures to adults. Furthermore, the number of inhibited larvae
(Hyostrongylus) may be estimated precisely if the pig has been
76 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

isolated from reinfection for at least 3 weeks before slaughter, as this

allows non-inhibited immatures to complete their development.

4.3.1 Incubation in physiological saline

Equipment

One or two trays of about 40 x 60 x 15 cm. The precise

size is not important. Suitable plastic trays are easily
procurable. Rectangular trays are easier to pour from
Sieve of 20-30 pm. A nylon net or another mesh of
equivalent size may be mounted with appropriate sides
Physiological saline, 0.9% (9 g of NaC1 in 1 litre of water)
Wash bottle
Available jet stream of water. It may be a water pistol
mounted on a hose
Tight 500 ml containers, some with a calibrated scale on
the outside
Thermostat or a space with approx. 38 C
Petri dishes, about 9 cm in diameter
Microslides/coverslips
Pasteur pipettes
Aqueous solution of iodine to stain the samples. A strong
solution may for example be: 80 g 12 and 400 g K1
dissolved in distilled water to a total volume of 1 litre
Aqueous solution of sodium thiosulphate to decolourize the
faecal matter, while the worms remain stained. The solution
could for example be 30%, i.e. 300 g sodium thiosulphate
crystals dissolved in water to a total volume of 1 litre
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 77

Dissection microscope
Microscope
Needles or fine forceps to handle worms during counting
70% ethanol with 10% glycerol for storage of isolated
worms, if necessary
Small tubes (glass or plastic) with tight lids for eventual
storage of isolated worms, if necessary

Procedure

The Incubation of the Stomach Wall is illustrated in Fig. 4.2.

The opened and washed stomach is placed with the mucosa

downwards in a tray containing physiological saline. The saline
should be lukewarm (38-40"C).

Incubate the stomach wall overnight at approx. 38'C.

Remove the stomach and wash it well with a jet stream of water.

Pour the incubation fluid and the washing water through a fine-
meshed sieve or nylon net (20-30 pm), and wash the sieve/net with
a jet stream of water.

Flush off the larvae from the sieve/nylon net into a container using
the wash bottle, and total the volume up to 200 ml.

Add some iodine solution so that the whole sample is deeply stained,
and allow the larvae to be coloured for at least one hour.

After thorough mixing, subsamples are transferred to petri dishes and

decolourized with a few drops of sodium thiosulphate immediately
before counting. To ensure that the larvae are easily visible, the
sample in the petri dish should not be totally decoulorized, but a faint
Cut the stomach Filter the incubation
open and wash it and washing fluid
through a 20-30pm
sieve
+
\ ............................. , /
Place the stomach in
Incubate
physiological saline with
the mucosa downwards overnight
at 38°C Flush off the
larvae from the
sieve and make
the volume up to
\ , i 200 ml

Wash the stomach with a

jet stream of water
'V
iodine solution
and leave for 1 hour
rAdd
Di sse cti on
microscope
Microscopical exami- if
nation and possible
storage in glycerol- Mix the solution
ethanol and transfer a
Add a few drops known sub-sample
1.....J to a petri dish
of thiosulphate *--
solution

FIGURE 4.2 Incubation of the Stomach Wall

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 79

brown nuance should remain (see Section 4.13.2: General comments

on occupational hazards).

It is necessary to place the petri dish under a stereo microscope

during counting. Note again: The counting will be much safer if only
small volumes of the sample are poured into the petri dishes.

To identify the parasite species, a certain number (e.g. 100-200) of

randomly selected larvae should be transferred to microslides by
means of a Pasteur pipette, and the larvae should be identified to
species under the microscope.

Finally, the total worm count of the sample should be multiplied by

an appropriate factor to obtain the total worm burden in the
stomach.

If necessary, specimens may be stored in glycerol-ethanol for later

observations.

4.3.2 Pepsin-HCI digestion

Equipment

Blunt knife
Pair of scissors
Sieve of 20-30 pm. A nylon net or another mesh of
equivalent size may be mounted with appropriate sides
Freshly prepared Pepsin-HCI solution (10 g pepsin (3,000
i.u. per mg) dissolved in 10 ml concentrated hydrochloric
acid and filled up to 1 litre with water (approx. pH =2))
Wash bottle
Jet stream of water. It may be a water pistol mounted on
a hose
 80 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Tight 500 ml containers, some with a calibrated scale on

the outside
Stomacher apparatus with thermostat. Alternatively, regular
mechanical stirring at approx. 38C conditions
Plastic bags for stomacher apparatus, if this equipment is
available

Thermostat or a space with approx. 38"C (not necessary if

a heated stomacher is available)
Petri dishes, about 9 cm in diameter
Microslides/coverslips
Pasteur pipettes

Aqueous solution of iodine to stain the samples. A strong

solution may for example be: 80 g 12 and 400 g KI
dissolved in distilled water to a total volume of 1 litre
Aqueous solution of sodium thiosulphate to decolourize the
faecal matter, while the worms remain stained. The solution
could for example be 30%, i.e. 300 g sodium thiosulphate
crystals dissolved in water to a total volume of 1 litre
Dissection microscope
Microscope

Needles or fine forceps to handle worms during counting

70% ethanol with 10% glycerol for storage of isolated
worms
Small tubes (glass or plastic) with tight lids for storage of
isolated worms

Procedure

The Pepsin-HCI digestion is illustrated in Fig. 4.3.

100g mucosa

500 ml digestive fluid Flush off the larvae

from the sieve and
make the volume up
to 200 ml

Filter the digested

solution through a
20 p sieve and
wash with a jet
stream of water Add iodine solution
and leave for 1 hour

Mix the solution

and transfer a
known sub-sample
to a petri dish
00 Dissectial
MiCrOSCOPe
1:=1

Heated magnetic stirrer Add a few drops

or of thiosulphate
solution
stomacher apparatus
37-40°C for 30 min.

C Xer,s1

FIGURE 4.3 Pepsin-HCI digestion of the Stomach mucosa

82 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

The opened and washed stomach is divided into 4-6 pieces which
are placed with the mucosa upwards, and the mucosa is then
scraped off by means of a blunt knife.

The mucosa is transferred to a container/plastic bag containing

pepsin-HCI (max. 100 g mucosa to 500 ml pepsin-HCI).

The mucosa is allowed to be digested at approx. 38C until the

digestion is completed. During digestion the suspension should be
stirred regularly. This step may be carried out by means of a
stomacher apparatus, if available, in which the sample is placed in a
solid plastic bag which is kept warm and stirred during incubation.
In a stomacher the mucosa will be completely digested in 30
minutes.

After total digestion of the mucosa, the digestion fluid is poured

through a 20-30 pm sieve and washed with a jet stream of water.

Flush off the larvae from the sieve/nylon net into a container using
the wash bottle, and total the volume up to 200 ml.

Add some iodine solution, so that the whole sample. is deeply

stained, and allow the larvae to be coloured for at least one hour.

After thorough mixing, subsamples are transferred to petri dishes and

decolourized with a few drops of sodium thiosulphate immediately
before counting. To ensure that the larvae are easily visible, the
sample in the petri dish should not be totally decolourized, but a faint
brown nuance should remain (see Section 4.13.2: General comments
on occupational hazards).

FinanY, the larvae should be counted under a stereo microscope, and

identified to species under a microscope. The total worm burden
should be calculated.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 83

4.4 THE CONTENTS OF THE SMALL INTESTINE

The principles and application of the methods for differential parasite

counts of the small intestine are largely the same as those for
parasite counts of the contents of the stomach. There is no strong
need for incubation or digestion of the intestinal wall. The parasites
of the small intestine vary considerably, from Macracanthorhynchus
hirudinaceus and Ascaris suum, measuring up to 65 cm and 40 cm,
respectively, to small, very slender parasites such as Strongyloides
ransomi and adult Trichinella spp., measuring 10 mm and 1-3 mm,
respectively.

Equipment

The same equipment as mentioned under 'contents of the

stomach'
. Mucosa scraper, i.e. two smooth pieces of wood/bamboo
(approx. 10 cm long) held together at one end by a strong
rubber band

Large-meshed sieve (500-1000 pm), a common household

sieve may be used

Procedure

The handling of the Small Intestinal Contents is illustrated in Fig. 4.4.

Ligate the small intestine at pylorus and close to caecum if

necessary, and cut the small intestine free. Remove fatty tissue from
the outer surface of the small intestine (this will facilitate mucosa
scraping, see below).

Open the small intestine so that all its contents fall into a tray.
 Remove fatty tissue from small Pour the remaining
intestine and cut it open suspension through
Make the volume a large meshed
up to 10 L and A sieve for isolation of
Intestinal contents
mix thouroughly large parasites
Mucus and washing water
Scrape off the Iir
mucus wash twice

10 L mark
J
Take out a
spoonful
(500 ml)

Possible Wash with a soft

storage jet stream of water
Examine at a light table

ii
Ask
i
Mix the solution Add iodine
1 Flush the sample to
and transfer a solution a 500 ml container
subsample to a and leave by means of
Add a few drops petri dish for 1 hour physiological salina
of thiosulphate In a wash bottle
solution 41(--- 4--- C reed

FIGURE 4.4 Handling of the Small Intestinal Contents

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 85

It is important to scrape off the mucous membrane in some manner

in order to recover the smallest parasites, e.g. Strongyloides and
adult Trichinella. This may be done by pulling the opened intestinal
wall through the two pieces of wood of the mucosa scraper. This
procedure should be repeated once.

The procedure for washing, subsampling, staining and counting is

just like that previously described for parasites in the stomach
contents, with the exception of counts for Ascaris and
Macracanthorhynchus.

All contents, and not merely a subsample, should be examined for

Ascaris, Macracanthorhynchus, and Fasciolopsis, as these large
parasites are usually present in rather low numbers. Therefore, after
subsampling from the bucket, a special 'large parasite examination'
may easily be carried out by washing the remainder of the sample in
the bucket through a large-meshed sieve by means of a strong jet
stream of water.

Pour the contents of the sieve into a tray with some water. The large
worms are then easily recognized in the water.

4.5 THE CONTENTS OF THE LARGE INTESTINE

The principles and application of the methods for differential parasite

counts of the large intestine are largely the same as those for
parasite counts of the contents of the stomach and incubation of the
stomach wall. The large intestinal walls should not be digested with
pepsin-HCI, as Oesophagostomum larvae will also be digested, and
therefore cannot be found. Lumen-dwelling parasites of the large
intestine vary in size from 3-4 mm (immature Oesophagostomum sp.)
to 4-5 cm (adult Trichuris suis).
86 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Equipment

The same equipment as previously mentioned under

Contents of the Stomach (Section 4.2)

Procedure

The same procedure as previously mentioned under Contents of the

Stomach (Section 4.2 and Fig. 4.1).

4.6 THE LARGE INTESTINAL WALL

The large intestinal wall may be infected with tissue stages of

Oesophagostomum spp. (L3 and L4). Furthermore, mature and
especially immature Trichuris suis may be so firmly attached to the
mucosa that they are difficult to wash off the mucosa with water.
These worms may be isolated by incubation of the large intestinal
wall in physiological saline at 38°C overnight, as mentioned for the
stomach wall.

Note: the pepsin-HCI digestion procedure is not applicable to the

large intestinal wall, as many of the worms will be digested together
with the mucosa.

Equipment

The same equipment as previously mentioned under

Incubation of the Stomach Wall
An extra bucket for each intestine
Small lengths of s-shaped non-corrosive steel wire
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 87

Procedure

The Incubation of the Large Intestinal Wall is illustrated in Fig. 4.5.

The same procedure as previously mentioned under Incubation of the

Stomach Wall is used, except that the intestinal wall may most easily
be incubated in a bucket in which the intestinal wall is held in
position by small lengths of steel wire.

4.7 THE LIVER AND OMENTUM

Many different.parasitic infections may cause similar reactions in the

liver. Thus, migrating larvae of Ascaris suum and other ascarids (e.g.
Toxocara sp.), immature Fasciola hepatica, and eggs of Schistosoma
japonicum may all cause accumulation of lymphocytes and increased
amount of connective tissue, i.e. 'white spots' or 'milk spots'. When
'white spots' are found in the liver, they indicate a recent passage or
entrapment of parasites in the liver. Enlarged, thick-walled bile ducts
may indicate presence of Fasciola hepatica or Dicrocoelium
dendriticum.

The liver and the omentum should be examined for the presence of
metacestodes (e.g. Taenia hydatigena up to 8 cm, Echinococcus
granulosus up to 20 cm or more), which may be clearly visible as
bladders or pronounced cysts on the surface of the liver, embedded
in the liver tissue, or in the omentum or in other internal organs. Care
should be taken to discriminate between white spots of the
lymphonodular type in the liver and small Taenia hydatigena cysts,
and in cases of doubt, microscopical examination may be necessary
to reveal the scolex.

When examining for the presence of liver flukes (Fasciola hepatica up

to 35 mm, Dicrocoelium dendriticum up to 10 mm), the following
method can be followed.
 open the
large intestine
ifCut
and wash it Filter the
incubation Flush off the
and washing larvae from the
fluid through sieve and make
Distend the the volume 200 ml
intestine with steel a 20-30pm
wire and place it sieve
in physiological
saline

Incubate
overnight Add iodine solution
V at 38°C and leave for 1 hour

intaesshtintrwith
Mix the solution
\ of water and transfer a
known sub-sample
to a petri dish

Di ssecti on
microscope

Possible storage in Add a few drops

glycerol-ethanol of thiosulphate
solution

FIGURE 4.5 Incubation of the Large Intestinal Wall

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 89

Equipment

Sharp knife
Tray of about 40 x 60 x 15 cm or smaller. The precise size
is not important. Suitable plastic trays are easily procurable.
Rectangular trays are easier to pour from
Physiological saline, 0.9% (9 g of NaCI in 1 litre water)
Sieve of 100-220 pm. A nylon net or another mesh of
equivalent size may be mounted with appropriate sides
Shallow tray or a few large petri dishes
Petri dishes, about 9 cm in diameter
Dissection microscope
70% ethanol with 10% glycerol for storage of isolated
WOM1S

Procedure

The handling of the Liver is illustrated in Fig. 4.6.

The liver and omentum are examined macroscopically for

metacestodes (see Fig. 4.7).

The liver is examined for superficially visible white spots (see Fig.
4.7).

Make three cuts in the liver (see illustration) and examine the
exposed tissue and bile ducts.

Transfer the liver into a tray with some litres of physiological saline
and squeeze and tear the liver manually (press the soft liver tissue
 -
Squeeze and tear the liver
Cut off the lobes of the manually and cut open the
liver and examine tissue gall bladder in a tray
and bile ducts containing physiological
saline

Pour tissue juice and

blood through a 100-220
um sieve and wash
gently

<-
Further identification Transfer the Pour and flush the
of small and immature parasites to a sample into a big petri
specimens by small petri dish. dish
microscope Count and identify

FIGURE 4.6 Handling the Liver

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 91

FIGURE 4.7 Liver with a big cyst of Taenia hydatigena

(-Cysticercus tenuicollis, left) and liver with numerous 'white spots'
or 'milk spots', caused by recently migrating Ascaris suum larvae
(right)

between the fingers until all pieces are max. 10 mm in size) in order
to get the flukes out of the bile ducts. Cut the gall bladder open.

Tissue juice and blood are poured through a 100-220 pm sieve and
washed with a soft stream of water. The contents are poured into a
tray and examined directly.

The washed sample is poured into a tray or a large petri dish, and
parasites are transferred to a small petri dish, identified and counted.
Microscopical examination may be necessary when immature
parasites are to be found.

4.8 THE LUNGS

The lungs may be infected with several species of long slender

lungworms, Metastrongylus spp. (up to 6 cm long), located in the
 92 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

bronchi and the bronchioles. These worms may cause gross

pathological changes and lesions, but as several different
microbiological infections may cause pathological lesions as well, it
is necessary to isolate the worms directly.

Equipment

Tray of about 40 x 60 x 15 cm. The precise size is not

important. Suitable plastic trays are easily procurable.
Rectangular trays are easier to pour from
Pair of scissors, which should be pointed and sharp
Physiological saline, 0.9% (9 g of NaCI in 1 litre of water)
Wash bottle
Needles and fine forceps to handle worms during
preparation and counting
Sieve of 200-300 pm. A nylon net or another mesh of
equivalent size may be mounted with appropriate sides
Petri dishes about 9 cm in diameter
Slides and coverslips
Microscope

Procedure

The handling of The Lungs is illustrated in Fig. 4.8.

The bronchi and the bronchioles are opened carefully with a pair of
fine scissors, and the lumen is washed with saline (washing water is
collected in the tray). Special observations are made on the presence
of mucus, fluid, bubbles and other signs of infection.
 Cut open the bronchi and
bronchioles with a pair of
fine scissors. Wash the opened lungs in a
tray containing physiological
Transfer visible worms to
physiological saline saline

1=1

Pour tissue juice and

blood through a 200-
fr 300 1.1M sieve and wash
gently with water

Microscopical
identification
and possible
storage in .4--
glycerol-ethanol Pour and flush the
Count the
parasites sample into petri dishes

FIGURE 4.8 Handling of the Lungs

94 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Macroscopically visible lungworms are transferred directly to a petri

dish with physiological saline.

The lungs, with opened bronchi and bronchioles, are washed with
physiological saline.

Tissue juice, blood and saline are poured through a 200-300 pm

sieve and washed with water (as the worms are v.ery susceptible to
mechanical damage, the procedure should be rather gentle).

The contents of the sieve are flushed off with physiological saline
into petri dishes, and the parasites are counted.

Identification to species level is carried out by placing male

specimens on some slides, with coverslips on top, and examining the
spicules under a microscope.

Isolated specimens may be preserved in 70% ethanol (9 parts) with

glycerol (1 part).

4.9 THE KIDNEYS

The kidneys and the perirenal fatty tissue may be infected with
Stephanurus dentatus, which is a stout worm, approx. 45 mm long.
The examination includes general observations on macroscopic
pathological changes, followed by direct search for macroscopically
visible worms.

Equipment

Sharp knife
Physiological saline, 0.9% (9 g of NaCl in 1 litre of water)
Wash bottle
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 95

Needles and fine forceps to handle worms during

preparation and counting
Petri dishes, about 9 cm in diameter
Dissection microscope

Procedure

The perirenal tissue is examined, and the kidneys are opened with a
knife. Special observations are made on the presence of pathological
lesions and worms embedded in capsules which may contain
greenish pus.

Any worms are freed with saline water in a wash bottle and fine
forceps.

The parasites are transferred to a petri dish with physiological saline,

identified and counted.

4.10 THE MUSCLES

Muscle-dwelling larvae of Trichinella may be found in many striated

muscles all over the body, but the number of larvae per gram, and
thus the possibility of finding the larvae, varies considerably from
muscle to muscle. The predilection sites of the larvae, i.e. the most
heavily infected muscles in the pig, are musculus masseter and
diaphragm, which is rather convenient, as the commercial value of
these muscles is low.

The traditional way of analyzing meat for trichinae is to use a

compressorium. This method is easy to perform, as well as
inexpensive, but it is commonly replaced by a pepsin-HCI digestion
method, which has a much higher sensitivity. Both methods will be
mentioned below, while the ELISA method for detecting circulating
96 The Epidemiology, Diagnosis and Control of Helminth Parasites of Sw ne

antibodies is omitted, as it is more expensive and has higher

technological demands.

4.10.1 The compression method

This method is very simple and has been used for many years for
routine screening of pork. It is also a very inexpensive method,
because the principle is to firmly press a little piece of meat between
two thick pieces of glass mounted with screws (a compressorium),
and then to examine for muscle larvae directly under a dissection
microscope. Unfortunately, this method has a rather low sensitivity
(the detection limit is approx. 3-5 larvae per gram), and the recovery
rate is to a large degree dependent on highly skilled technicians.

Equipment

Pair of scissors for cutting the meat into small pieces

Compressorium
Dissection microscope

Procedure

The Compression Method is illustrated in Fig. 4.9.

One gram of diaphragm or m. masseter is cut into small pieces.

After being placed between the glass plates of the compressorium,

the plates are screwed tightly together, so the meat is pressed into
a thin, transparent layer.

The compressorium is examined directly in a dissection microscope.

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 97

Cut grain sized pieces of

muscle (diaphragms or m.
masseter) and place them
in the compressorium IOW

2.;2 ZirAWIS P/P:0"/

4111111K-1111111111111111111P
.,CreIrd2reW2722;(2-27(-2,74%

Screw the plates firmly

together until the muscle
tissue is transparent

,C,tn7,5,PrWeo:4;?,"
Ailit4rNme=0"imir

Rep!

FIGURE 4.9 The Compression Method

4.10.2 The pepsin-HCI digestion method

Muscle larvae of trichinae are easy to isolate by removing the
surrounding host tissue by means of pepsin-HCI. When digestion
succeeds perfectly, only very few muscle fibres are left, and the
stained larvae are clearly visible and easy to enumerate. Therefore,
this method has a rather high sensitivity. As trichinae may cause
serious zoonotic infections and therefore have impact on human
health, it is often recommended to use a detection limit of 0.1 larvae
per gram of meat, i.e. 10 grams of muscle tissue is examined.

Equipment

Pair of scissors for cutting the meat into small pieces

98 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Freshly prepared Pepsin-HCI solution (30 g pepsin (3,000

i.u. per mg) dissolved in 10 ml concentrated hydrochloric
acid and filled up to 1 litre with water (approx. pH =2))
Stomacher apparatus with thermostat. Alternatively, regular
mechanical stirring at approx. 38'C conditions
Plastic bags for stomacher apparatus, if this equipment is
available

Sieve of 400-500 pm and one sieve of 20-30 pm. Nylon

nets or other meshes of equivalent sizes may be mounted
with appropriate sides
Jet stream of water. It may be a water pistol mounted on
a hose. Alternatively, a vacuum pump and vacuum devices
may be used to get the digested fluid through the sieves
(see Fig. 4.10)
2000 ml conical container
Aqueous solution of iodine to stain the samples. A strong
solution may for example be: 20 g 12 and 40 g KI dissolved
in distilled water to a total volume of 1 litre
Aqueous solution of sodium thiosulphate to decolourize the
other matter, while the larvae remain stained. The solution
could for example be 30%, i.e. 300 g sodium thiosulphate
crystals dissolved in water to a total volume of 1 litre
Petri dishes, about 9 cm in diameter
Dissection microscope

Procedure

The Pepsin-HC1 digestion is illustrated in Fig. 4.10.

10 g muscle tissue
+
500 ml digestive fluid
Transfer the 20 p l Leave in the
filter to a petri iodine solution
dish with iodine for 3 min

*
Transfer to thio-
sulphate solution
and leave for 4
il.LJ sec. to remove
access dye

+
Let dry on
filterpaper
\ f for 1 min

Vacuum
00
I=1 1:=3

Heated magnetic stirrer

or
stomacher apparatus
37-40°C for 30-60 min.

C tereel

FIGURE 4.10 Pepsin HCI-digestion of muscle

 1 00 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Ten grams of diaphragma or m. masseter are cut into small pieces.

The muscle tissue is transferred to a beaker/plastic bag containing

pepsin-HCI (10 g muscle to 500 ml pepsin-HCI solution).

The meat is allowed to be digested at 37'C until the digestion is

completed (approx. IA hour). During digestion, the suspension should
be stirred regularly. This step may be carried out by means of a
stomacher apparatus, if available, in which the sample is placed in a
solid plastic bag which is continuously heated and stirred during
incubation.

After total digestion of the muscle tissue, the digestion fluid is

poured through a 400-500 pm sieve and then through a 20-30 pm
sieve. The digestion fluid is forced through the fine sieve by vacuum.

Place the fine sieve in the iodine solution, so that the whole sample
is deeply stained, and allow the larvae to be coloured for 3 minutes
(4 min. for frozen muscle tissue).

Next, place the fine sieve in sodium thiosulphate for 4 sec. to

decolourize any other tissue than the larvae.

Finally, the larvae are counted under a stereo microscope.

Note: Care should be taken to count only trichinae. Larvae of

migrating nematodes may occasionally be found in muscle tissue,
and these larvae should not be counted as trichinae. Therefore, the
larvae should be viewed at a higher magnification if identification is
doubtful.

4.11 IDENTIFICATION OF HELMINTHS

Table 4.1 provides simple descriptions of the most common gastro-

intestinal helminths of pigs, and Table 4.2 describes the helminths
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 101

found outside the gastro-intestinal tract. These tables do not

represent any elaborate identification key for porcine helminth
species and not even genera, as this is beyond the scope of the
present guidelines, but the tables are intended merely as an overview
of the most commonly found genera and species. For exact
identification of nematodes to genus, the reader may consult the CIH
Keys to the Nematode Parasites of Vertebrates (Anderson, Chabaud
and Willmott, 1974-1983).

4.12 INTERPRETATION OF WORM COUNTS

It is difficult to present a guideline for the interpretation of worm

counts, as a complexity of factors concerning the intrinsic properties
of the helminth species (pathogenicity to pigs as well as man), the
specific host-parasite relationship (transmission route; acquired
immunity - and thus age group of pigs), and the management system
(e.g. risk of transmission; use of anthelmintics) should be considered
before classifying a worm burden as being light or heavy.

4.12.1 The helminth species

For some highly pathogenic helminth species, a single specimen is

one too many, as they may constitute a hazard to public health. This
applies to the most deleterious zoonotic species, such as Taenia
solium and Echinococcus sp., and Trichinella spiralis.

For non-zoonotic helminths, the primary effect is exaggerated in the

pig, and here a fixed number of e.g. 50-100 specimens of liver flukes
(Fasciola hepatica, Dicrocoelium dendriticum), lungworms (Meta-
strongylus spp.), or adult Ascaris suum or Macracanthorhynchus
hirudinaceus may cause so much harm that they constitute an
unacceptably high infection, whereas a similar number of less
pathogenic species (e.g. Oesophagostomum spp., Hyostrongylus
rubidus and the spiruroids) is normally regarded as a light infection.
1 02 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

But even within the same species, one should distinguish between
stages; e.g. 200 small immatures of Ascaris suum in the small
intestine may not be alarming, as the large majority of these are
expected to be expelled rapidly, while the same number of fully
grown adults may severely affect the growth rate and may indicate
that the regulation of the parasite population is out of function.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 103

TABLE 4.1 Characteristics of gastro-intestinal helminth species in pigs (to be

continued)

Organ Helminth Max. Appearance Characteristics of

Length the parasite

Oeso- Gong ylonema 15 cm long, slender worms longitudinal rows of

phagus pulchrum (embedded in the cuticular plaques in
mucosa) the anterior body
region

Stomach Trichostrongylus 7 mm hairlike no buccal capsule,

axei distinct excretory
notch in
oesophageal region

Hyostrongylus 8 mm small, s ender worms reddish, small

rubidus cephalic vesicle

011ulanus 1 mm very tiny worms the head is spiral

tricuspis coiled
(microscopical
examination)

Ascarops 22 mm small, slender worms single cervical ala

strongylina on the left side,
pharynx with 2-3
spiral chitinous
thickenings

Physocephalus 20 mm small, slender worms 3 pairs of cervical

sexala tus alae, pharynx with
1 spiral chitinous
thickening

Simondsia small, slender a (free in 2 cervical Mae, a

paradoxa lumen). Gravid with dorsal and a ventral
sack-formed posterior tooth in buccal
end (in mucosa crypts) cavity

Gnathostoma 4 cm thick-bodied worms in swollen anterior

hispidum gastric nodules ends with rows of
G .doloresi small hooks

Gong ylonema 9 cm long, slender worms longitudinal rows of

pulchrum (embedded in the cuticular plaques in
mucosa) the anterior body
region
104 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

TABLE 4.1 Continued

Organ Helminth Max. Appearance Characteristics of the

Length parasite

Small Trichostrongylus 7 mm hairlike no buccal capsule, distinct

intestine colubriformis excretory notch in
T.vitrinus oesophageal region

Globocephalus 9 mm small, robust worms spherical buccal capsule

urosubula tus without leaf crown, 9-tail
G.Iongemucronatus with terminal spine

Strongyloides 10 mm hairlike only 99, blunt tail, long

ransomi oesophagus

Ascaris 40 cm long, stout, white may be confused only

SLAW) WOM1S with
Macracanthorhynchus
(see below)

Trichinella 3 mm tiny worms, rarely long oesophagus, o": 2

spiralis found as they are cloacal flaps, but no
short-lived spicule, 9: larvae in uterus

FascioIopsis 40 mm large, thick bodied intestinal caeca without

buski flukes side branching
10 mm width

Macracanthorhynchus 65 cm large, stout, white the spiny proboscis is

hirudinaceus worms protruded when placed in
water

Large Oesophagostomum
Intestine dentatum' 15 mm small, stout, white small buccal capsule,
0.quadrispinulatum 2 WOMIS inflated cuticular cephalic
O.brevicaudum vesicle

Trichuris suis 6 cm whip-like, white long, filamentous anterior

worms (the anterior end, thick posterior end
end embedded in
the mucosa)
0.granatensis and 0.georgianum are regarded as variant forms of 0.dentatum
alongicaudum is synonymous with 0.quadrispinulatum
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 105

TABLE 4.2 Characteristics of non-gastrointestinal heirninths in pigs

(to be continued)

Organ Helminth Max. Appearance Characteristics of the

Length parasite

Liver Taenia 8 cm small, thin walled, white the single scolex may be
hydatigena cysts - large, fluid-filled recognized after
(cysticercs)' bladders dissection

Echinococcus 20 cm large, thick walled, numerous scolices in

granulosus white hydatids each hydatid
E.multilocularis
(hydatids)

Dicrocoelium 10 mm lanceolate, semi- uterus filled with dark

dendriticum transparent flukes brown eggs, testes in
the anterior part of the
body

Fasciala hepatica 35 mm grey-brownish, 10 mm immatures: lanceo/ated,

width in parenchyma
adults: leaf-shaped,
conical anterior end with
distinct shoulders,
intestinal caeca with
numerous branches

Opisthorchis 8 mm small, lanceolate flukes grossly resembling D.

noverca dendriticum, the suckers
are smaller, the eggs are
light brown, and testes
are located in the
posteriorly part of the
body

Ascarid migrating 1 mm small, robust larvae Larvae: no

larvae milk spots of varying characteristics
appearance Milk spots: non-specific
host response

Schistosoma spp. milk spots of varying non-specific host

(eggs) appearance response
106 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

TABLE 4.2 Continued

Organ Helminth Max. Length Appearance Characteristics of the

parasite

Lungs Metastrongylus 6 cm long, slender, white the host, the site and the
apri2 worms long slender form are
M.salmi sufficient for generic
M.pudendotectus identification

Blood Schistosoma 2 cm males and females o' : broad, flat with a

system japonicum in permanent gynaecophoric canal,
S.mansoni copulation carrying the slender 9
S.incognitum

Kidneys Stephanurus 45 mm large, stout worms prominent buccal capsule,

dentatus transparent cuticle

Connect. Taenia 8 cm small, white cysts - the single scolex may be

tissue hydatigena large, fluid-filled recognized after
(cysticercs)1 bladders dissection

Muscles Trichinella 1 mm L1 encapsulated in the coiled, encapsulated

spiralis (larvae) striated muscles, larvae within striated
old capsules may be muscles are sufficient for
calcified generic identification

Taenia solium 10 mm elongated, white within the cyst is a scolex

(cysticercs)3 cysts with 4 suckers and a
rostellum bearing 2
concentric rows of hooks
The cysticerc of T.hydatigena may be called Cysticercus tenuicollis
M.elongatus is synonymous with M.apri
The cysticerc of T.solium may be called Cysticercus cellulosae
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 107

4.12.2 Specific host-parasite relationships

Most helminths have a characteristic infection pattern related to the

age of their hosts. This pattern may be influenced by the risk of
being infected, but more often the immunogenicity is the most crucial
factor. Thus, for example the highly immunogenic Strongyloides
ransomi, which is transmitted to newborn piglets via colostrum,
normally has the highest intensity of infection in very young pigs.
Growing pigs, 3-6 months of age, may be the age group most
heavily infected with Ascaris suum and Trichuris suis, as these
parasites are highly immunogenic, while they are not transmitted so
effectively to the piglets. Finally, helminths such as
Oesophagostomum spp. and Hyostrongylus rubidus seem to have
relatively lower immunogenicity and they are gradually accumulated
with age and thus dominate in adult pigs.

Therefore, the age of the infected pigs should be considered when

evaluating a recorded worm burden. lf, for example, young pigs
harbour many Oesophagostomum spp. or Hyostrongylus rubidus, this
indicates an unusually high transmission, while the same worm
burdens may be acceptable in sows. On the other hand, many
Ascaris suum or Trichuris suis, or even low numbers of Strongyloides
ransomi in sows may indicate that these hosts have not had
substantial experience with these infections during their growth, and
therefore the herd's overall transmission level with these helminths
may be low.

4.12.3 Management systems

The management system has been shown to have a strong influence

on helminth prevalences. If pigs are reared in an extensive outdoor
production system without any helminth control, it is not surprising
if the worm burdens are heavy. On the other hand, only few helminth
species (perhaps only Ascaris suum) are expected to be found in a
highly intensive production system, and therefore even moderate
worm burdens with unexpected helminths in such a system should
1 08 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

call for attention. lf, for example, moderate numbers of Taenia

hydatigena or Oesophagostomum spp. are found in a highly
industrialized herd with routine anthelmintic prophylaxis, the
veterinary advisers should look for either control with Taenia-infected
dogs, or for particular niches in which Oesophagostomum larvae are
able to develop. In the latter case, one should also be aware of
anthelmintic resistance.

4.13 GENERAL COMMENTS

4.13.1 The subsample technique

The principle of subsampling is easily understandable, and

subsampling is a convenient way to reduce the workload. But in
practice subsampling is not so easy to carry out in a correct way. If
the mixing of the total volume is not appropriate, the counts of the
subsamples may either overestimate or underestimate the total worm
burden. Therefore, the total sample should be mixed very well either
until immediately before each subsample is taken or while it is being
taken.

For small samples in a container (e.g. a container with a total of 200

ml), a commercially available air pump used for non-professional
aquaria may be a good solution. The air may be led through a plastic
tube mounted with a glass Pasteur pipette to the bottom of the
sample, and the mixing effect of the bubbles should be allowed to
continue during the subsampling.

Subsampling techniques may never be totally adequate. One way to

overcome the inaccuracy is to use exactly the same procedure every
time, in order to make the error systematical and thus to reduce the
between-samples variation. Furthermore, this variation should be
checked by counting a series of subsamples from the same source.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 109

4.13.2 Occupational hazards

When using iodine or other chemicals which may be harmful to

human health, one should be aware of the risks and know some
ways to reduce them. Iodine will evaporate from stained samples
when they are counted, i.e. when people are bending over the petri
dish. The risk may be reduced by adding sodium thiosulphate, which
reduces the amount of free iodine, and by sitting in a well-ventilated
place or outdoors. If iodine samples are counted routinely in the
laboratory, a local ventilation outlet located close to the petri dish
may be a good permanent solution, and it may for example be
constructed of a commercial vacuum cleaner with an outlet through
the window.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 111

CHAPTER 5

EXAMINATIONS FOR INFECTIVE LARVAE AND

EGGS IN HERBAGE AND SOIL

5.1 INTRODUCTION

This chapter deals with supplementary diagnostic procedures for

isolation of infective stages of parasites in the environment of the
pigs. In contrast to most other domestic animals, the pig will eat
both herbage and contaminated soil. Infective third-stage larvae
(Oesophagostomum spp. and Hyostrongylus rubidus) may be found
both in grass and soil, while infective eggs (e.g. of Ascaris suum and
Trichuris suis) may primarily be found in soil, and it may be relevant
to make estimates for both types of infective stages. However, even
if herbage and soil have been examined thoroughly, one should be
aware that the results make sense only for helminth species with an
uncomplicated direct life cycle, such as those mentioned above.

The techniques all have variable efficiencies, and the numbers of

eggs/larvae within the samples may vary considerably; therefore the
procedures described below should all be regarded as qualitative and
semiquantitative.

5.2 COLLECTION OF HERBAGE AND SOIL SAMPLES

The principles for collection of herbage and soil are similar. The
samples should be as representative as possible, and therefore one
sample should be collected as many small subsamples, picked up
from all over the area concerned, and this sampling should be
repeated, so two identical samples are examined (i.e. duplicate
sampling). This precaution is important, because the eggs/larvae are
expected to be unevenly distributed with 'hot spots' of high numbers
of infective stages scattered in otherwise low-infective areas.
112 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

The collection of herbage samples may in practice be performed by

walking along two W-routes covering the whole area (Fig. 5.1), and
for every 10 steps picking up a little amount of herbage (without soil,
it may be necessary to use scissors) and placing it in a plastic bag
(total amount for the whole area: 300-600 g). If the pasture is small,
herbage should be sampled more often than for every 10 steps to get
at least 30 subsamples. As pigs may destroy the vegetation by
rooting, only limited areas with vegetation may be available for
sampling. Herbage contaminated with faeces should be avoided.

Similarly, the collection of soil samples may be performed by

following two W-routes covering the entire area, picking up a little
amount of soil between 3 fingers (without vegetation and roots), and
then placing it in a plastic bag (total amount for the whole area: 200-
300 g). Soil samples cannot be collected in areas with very dense
vegetation. Direct sampling of faeces should be avoided.

Sampling area

FIGUR 5.1 Two schematic W-routes for collection of herbage (or

soil) samples. At least 30 subsamples should be collected from each
W-route.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 11 3

5.3 ISOLATION OF INFECTIVE LARVAE

Infective third-stage larvae may be found both on the vegetation and

in the soil. Unlike e.g. the ruminants, pigs both ingest herbage and
pick up significant amounts of soil when rooting for earthworms,
insects, roots etc. Furthermore, Oesophagostomum larvae seem to
be much less motile than Hyostrongylus larvae, and they do not
migrate upwards on the herbage to the same extent. Therefore it
may be relevant to examine herbage as well as soil for infective
parasite larvae.

5.3.1 Isolation .of infective larvae from herbage

The procedure is a simple Baernnann technique.

Equipment
1-2 buckets
Double-layer cotton gauze
Balance

Domestic detergent
Conical sedimentation beaker
Measuring cylinder (1 litre) and some kind of vacuum device may
be useful
Pipette (e.g. 5 ml)
Test tube
Test tube rack

Aqueous iodine solution (IIK). Either Lugols solution (10g 12 and

30 g KI dissolved in distilled water to a total volume of 1 litre) or
114 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

a stronger solution (e.g. 80 g 12 and 400 g KI dissolved in

distilled water to a total volume of 1 litre)
Sodium thiosulphate solution
Pasteur pipette
Microscope slides, preferably large, with a paraffin bank
surrounding the sample area.
Microscope with 10-40x magnification

Procedure

The method for Isolation of infective larvae from herbage is illustrated

in Fig. 5.2.

Place the collected grass sample on a large piece of double-layer

cotton gauze.

Form the gauze into a bag and immerse the bag in a bucket filled
with tap water to which a few drops of detergent have been added.

During the first 3-4 hours, move the bag gently up and down the
bucket several times to agitate the sample.

Leave the bag in the water at room temperature overnight.

Next morning, remove the bag and run fresh tap water over it and
into the bucket. Leave the contents of the bucket to sediment for at
least 1 hour.

The bag of grass should be dried (sun/oven/incubator), and the net

dry weight (gross dry weight minus dry weight of the gauze) is
measured when the sample is completely dry.
 Wrap the grass jt Lift and allow the Keep the gause bag
4r, sample in a sample to drip off and grass for
double layer of before retuming ir determination of
gauze and it to the bucket grass dry weight
immerse it in tap (repeat several
water with added times)
detergent
Leave to sediment
for 1 hour and
--». remove supematant
Leave
ovemight

Transfer a subsample
to a microscope slide
with a paraffin bank
and examine at 40-
100x magnification
Add a weak iodine
and count, or add
a stmng iodine
and store 4e--\

Suspend the
sediment and pour
Leave ovemight into a conical beaker
Add a few
drops of for sedimentation
thiosulphate of larvae

FIGURE 5.2 Isolation of Infective Larvae from Herbage

116 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Remove the supernatant carefully (decantation, vacuum), avoiding

resuspension of the sediment. Leave Y2-1 litre containing all
sediment.

If the volume is too large, the sedimentation may be repeated in a

high, narrow glass (e.g. a 1-litre measuring cylinder), whereafter the
supernatant again is discarded after 1 hour.

Suspend all the sediment, pour it into a conical sedimentation beaker,

and leave it at room temperature overnight. Parasite larvae will then
accumulate at the bottom.

Harvest the larvae with a 5 ml pipette, by placing your fingertip at

the end of the pipette, move the tip of the pipette to the bottom of
the beaker, let the larvae be sucked up into the pipette, and transfer
the larvae to a test tube. This step may be repeated if some larvae
remain in the beaker.

If any large particles clog the pipette opening, the pipette may simply
be turned around, so the larvae are sucked into the wide end. Do not
use mouth suction, as larvae may then be accidentally sucked into
your mouth.

An alternative to the conical sedimentation beaker is a large glass

funnel (20 cm diameter), fixed in a stand, and fitted with a flexible,
transparent tubing. The tubing should carry 2 screw clamps placed
10 cm apart. The lower clamp is fastened, while the upper clamp is
open, allowing the larvae to accumulate just above the lower clamp
overnight. Harvest the larvae by closing the top clamp and collecting
the trapped sediment with about 10 ml of fluid in a test tube by
opening the bottom clamp.

The samples may be stored for max. 7 days in a refrigerator (4-5°C)

before examination.

The samples may now be stained either with a weak iodine solution
(LugoIs solution) or with a strong iodine solution followed by
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 11 7

decolouring with sodium thiosulphate.

When using LugoIs solution, some drops of the samples are

transfered to the microscope slide with a paraffin bank/ring, a drop
of LugoIs solution is added, and the larvae should be counted
immediately (within 15 minutes) under a microscope (10-40x
magnification). There will usually be many more non-parasitic
nematodes present than parasite larvae, and the counting may
therefore be rather laborious. But the parasitic larvae will tend to
obtain the brown iodine colour rather slowly, while free-living
nematodes will obtain the brown colour immediately. As the parasitic
larvae are uncoloured when counted, their morphological
characteristics are clearly visible.

If iodine is to be used for preservation, some drops of the strong

iodine solution should be added to the samples, which then may be
stored for a considerable period of time before counting. After
transfer of some drops of the sample to the microscope slide with a
paraffin bank/ring, 1-2 drops of sodium thiosulphate are added (as
little volume as possible, but the aqueous solution should loose its
iodine colour) and the larvae are immediately identified and counted
under a microscope. When using this counter-staining, the parasitic
nematodes will tend to retain the brown iodine colour for 15 minutes
after the decolouring with sodium thiosulphate, while the free-living
nematodes will loose the brown colour immediately. As the parasitic
larvae are brown when counted, their morphological characteristics
are difficult to observe.

Both alternative methods help to distinguish between free-living and

parasitic nematodes. Nevertheless, the cuticle sheath with the
prolonged tail is the most important key character, and the colour
should be used only as a help.

As the net dry weight of the sample is known, the results may now
be expressed as numbers of larvae per kg dry weight, although it
should be emphasized that this method is only semiquantitative.
118 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

5.3.2 Isolation of infective larvae from soil

The procedure is a simple Baermann technique. The first steps are

different from the handling of grass samples, as the soil contains a
lot of small particles which should be retained when the
baermannization occurs.

Equipment

Beaker (e.g. 500 ml)

* Balance

Tray. The precise size is not important, but it should be bigger

than the toy sieve (see below). Suitable plastic trays are easily
procurable. A rectangular shape facilitates pouring from it

Plastic sieve with flat bottom. A cheap toy sieve is suitable, but
a sieve like a letter tray may have a larger surface

4 small rubber stoppers (for test tubes), or other objects of

similar size

Sheets of fine, loosely woven laboratory paper ('kleenex')

Conical sedimentation beaker

Pipette (e.g. 5 ml)

Test tube

Test tube rack

Aqueous iodine solution (II K). Either LugoIs solution (10 g 12 and
30 g KI dissolved in distilled water to a total volume of 1 litre) or
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 11 9

a stronger solution (e.g. 80 g 12 and 400 g KI dissolved in

distilled water to a total volume of 1 litre)

* Sodium thiosulphate solution

Pasteur pipette
* Microscope slides, preferably large, with a paraffin bank
surrounding the sample area

Microscope with 10-40x magnification

Procedure

The Isolation of infective larvae from sod is illustrated in Fig. 5.3.

Mix the soil sample thoroughly in a beaker and take a subsample of

50 g by means of the balance. The subsample should be collected
from several different places in the soil mixture.

Place the toy sieve or another large-meshed plastic device on the 4

stoppers in a tray.

Place a single layer of kleenex inside the sieve, and add tap water
until the water level just touches the kleenex; if too much water is
added, the sample tends to be more dirty. Ensure that the kleenex is
not torn, as any holes will allow soil particles to pass.

Distribute the 50 g soil sample in the sieve. Continue to be careful

not to tear the kleenex.

Leave the sample in the sieve for one or two nights at room
temperature, or 37°C if an incubator is available. Now the larvae will
migrate through the loosely woven kleenex, while soil particles are
retained. Carried out correctly, the resulting larval suspension is
rather clean.
 Mix the soil sample
thoroughly and take
two 50 g subsamples Incubate at
20-37°C for
1-2 days
Carefully,
remove the
perforated
tray
Determination Place the soil on one Place the tray in a humidity
of dry weight layer of kleenex in a chamber. The water level
perforated tray or should just reach the
plastic sieve buttom af the tray or sieve

Transfer a subsample
to a microscope slide
with a paraffin bank
and examine at 40-
100x magnification
Add a weak iodine
and count, or add
a strong iodine
and store

Suspend the
sediment and pour
into a conic beaker

Add a few Leave ovemight

drops of for sedimentation
thlosulphate of larvae

FIGURE 5.3 Isolation of Infective Larvae from Soil

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 21

When harvesting, remove the sieve carefully, pour the contents of

the tray directly into a conical sedimentation beaker, and leave it at
room temperature overnight.

Parasite larvae will now accumulate at the bottom of the

sedimentation beaker. If the sample volume is too large for the
sedimentation beaker, sedimentation for 1 hour in a high, narrow
glass (e.g. a 1 -litre measuring cylinder) may be inserted. Again, an
alternative to the conical sedimentation beaker is a large glass funnel,
as described in Section 5.3.1.

Now the remaining part of the procedure is identical to the procedure

previously described in Section 5.3.1: Isolation of infective larvae
from herbage. The results may be recalculated as numbers of larvae
per kg soil, although it must be emphasized that this method is only
semiquantitative.

5.4 ISOLATION OF INFECTIVE EGGS FROM SOIL

The helminth eggs, which do not hatch outside a host, will remain
in the faeces until they are spread to the soil by rain, earthworms,
insects etc. Thus, it is relevant to look for infective eggs in soil
samples and not in herbage. This method may be useful for common
helminths like Ascaris suum and Trichuris suis, but it may also be
used for detection of Metastrongylus spp., even though infected
earthworms may complicate the interpretation of the results.

Equipment

Beaker (e.g. 500 ml)

Pair of scissors
Stirring device (fork, tongue depressor)
1 22 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Balance

Test tube, 50 ml (with a 50 ml mark)

Test tube stopper/lid
Test tube rack (for 50 ml tubes)
0.5 Molar NaOH
Centrifuge with holders for 50 ml tubes
Pasteur pipettes and rubber teats
Vacuum device, if available
Flotation fluid: Saturated NaCl with 500 g glucose per litre
Laboratory sticks
Erhlenmeyer flask (min. 250 ml), preferably having a cork with
two holes, mounted with small glass tubes and rubber tubes, if
a vacuum device is available
Sieve, 200-300 pm (e.g. 212 pm)
Sieve, 20 pm. The mesh size is critical, as the small eggs of
Trichuris, having a width of 21-31 pm, must not be able to
pass. On the other hand, the mesh size should be as large as
possible to avoid blocking
Jet stream of water. It may be a water pistol mounted on a hose
Wash bottle
McMaster counting chamber
Filtering paper cut into approx. 1 cm wide strips
Microscope with 40-100x magnification
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 23

Procedure

The Isolation of infective eggs from soil is illustrated in Fig. 5.4.

Cut any roots into small pieces with a pair of scissors, and mix the
soil sample thoroughly in a beaker, using a stirring device.

Weigh out a subsample of 10 g in a 50 ml test tube on a balance.

The subsample should be collected from several different places in
the soil mixture.

Weigh out a similar subsample of 10 g on a petri dish. This sample

should be dried (sun/oven/incubator), and the net dry weight (gross
dry weight minus the weight of the petri dish) is measured when the
sample is completely dry.

Add 0.5 M NaOH to the 50 ml mark on the test tube and mix the soil
suspension by means of a small applicator stick. Close the test tube
with a stopper/lid, and let the soil sample soak overnight at room
temperature or in a refrigerator.

The NaOH makes the eggs less sticky and breaks down the texture
of the soil particles.

Next day, centrifuge the sample at 1200 RPM (revolutions per

minute) for 7 minutes.

Remove the supernatant with a pipette or another vacuum device,

and discard the supernatant. Be careful not to resuspend the
sediment.

Add 20-30 ml flotation fluid (saturated NaCI with glucose) and

resuspend the sediment by stirring with a small applicator stick. After
thorough mixing, flotation fluid is added up to the 50 ml mark.
 Mix the soil sample Fill up with NaOH to Gradually mix Centrifuge at 1200
thoroughly and take the 50 ml mark, and fill up with RPM for 7 min and
two 10 g subsamples close wfth a stopper flotation fluid to transfer 40 ml
and leave overnight the 50 ml mark supernatant to an
Erhlenmeyer flask
4
Determination
of dry weight Centrifuge at
LAsb-1 1200 RPM
for 7 min and
reMOVe
supematant

Flush the
Resuspend sample on the
Transfer the whole sample to two or
the pellet in 20pm sieve by
more McMaster slides. Fill In all flotation fluid tap water to a 50
compartments and leave for 3-5 and mix well ml test tube
min before counting all eggs, both Wash thoroughly
inside and outside the grids through a double
sieve
r
sandEPI *-
Centrifuge at
1
AlliMEW 1200 RPM for I. 400 pm

Examine at 40x 7 min and

remove 1 20 pm
magnification
supernatant

FIGURE 5.4 Isolation of Infective Eggs from Soil

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 25

Centrifuge the sample for 7 minutes at 1200 RPM. Many helminth

eggs will now flotate.

Suck approx. 40 ml supernatant into a 250 ml Erhlenmeyer flask. If

a vacuum device is available, this may be most easily done by means
of a cork with two holes, one for inlet of the supernatant and one for
outlet of air. It is advisable to suck a small amount of water through
the system after the flotation fluid in order to avoid crystallization of
salt and glucose.

Be careful not to suck sediment from the test tube, as any sediment
particles will easily stop the fine sieve later in the procedure (see
below).

Repeat the last 3 steps, i.e. add flotation fluid, centrifuge and
transfer the supernatant to the Erhlenmeyer flask, three times. Each
time the supernatant should be collected in the same Erhlenmeyer
flask. The repetitions are necessary to increase the recovery of
helminth eggs.

Discard the washed sediment in the test tube, and wash the test
tube with water. The test tube may now be reused after the
supernatant has been sieved.

Pour the supernatant from the Erhlenmeyer flask into a double sieve
(a large-meshed sieve, 200-300 pm, placed over a fine-meshed sieve,
20 pm).

Wash the sample thoroughly with a jet stream of water. Now the
coarse particles will be trapped in the large-meshed sieve, while fine
particles, including any helminth eggs, will accumulate on the fine
sieve.

Separate the two sieves and wash the fine sieve once more with
water, while accumulating the particles in one side of the sieve. Use
a washing bottle with tap water for the final accumulation of
particles and for their transfer to the 50 ml test tube.
1 26 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Centrifuge the test tube for 7 minutes at 1200 RPM. As the eggs are
suspended in water, they will now be spun down.

Suck off and discard the supernatant.

Add a volume of flotation fluid (saturated NaCI with glucose),

corresponding to at least 4-5 times the volume of the sediment. Very
often the total volume of flotation fluid and sediment may be less
than 2.0 ml.

Resuspend the sediment by sucking slovvly up and down in a pasteur

pipette, and transfer the total volume to 1-3 McMaster counting
chambers, the number of counting slides depending on the sample
volume.

Add a little more flotation fluid to the test tube and transfer this to
the counting chambers to wash out the test tube and the pipette.

As the entire sample is to be counted, the McMaster chambers

should be filled restrictively, with no surplus of sample along the
edges of the chambers, i.e. the total amount of flotation fluid should
be placed underneath the upper glass, allowing all helminth eggs
present to accumulate under this glass.

Count all helminth eggs in the counting chambers, not only inside the
normal counting area, but also outside the counting area. Count all
McMaster chambers. A magnification of approx. 40x is suitable.

Any helminth eggs should be identified to species, and at a higher

magnification it is clearly visible whether they contain developed
embryos (see Section 5.5).

After counting, the McMaster chamber should be washed under a

stream of tap water, shaken to remove most of the water, and dried
with a cotton cloth on the outside and with a strip of filter paper
inside the chamber.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 27

As the total sample represents 10 g soil, the counts may be

expressed as number of helminth eggs per gram of soil by multiplying
by a factor of 0.1.

Example: 1 5 Ascaris suum eggs are found in the entire sample. Then
the number of eggs per gram of soil is 1 5x0.1 = 1.5 eggs per gram.

FIGURE 5.5 A fila riform oesophagus of a infective L3-larva (left) and

a rhabditiform oesophagus of a free-living nematode (right)
1 28 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

5.5 IDENTIFICATION OF LARVAE AND EGGS

The infective third-stage larvae may be distinguished from free-living

nematodes, being located inside a cuticular sheath, i.e. the cuticle of
the L2 larvae. Normally, this sheath is clearly visible at good
magnification. Furthermore, the L3 larvae have a filariform
oesophagus, while soil nematodes (and L1 and L2 larvae of parasitic
nematodes) have a bulbous rhabditiform oesophagus (Fig. 5.5). Very
often herbage samples contain a number of infective larvae of non-
porcine helminths, e.g. ruminant genera such as Ostertagia,
Cooperia, Trichostrongylus etc., and these larvae should be omitted
when making research on pig helminths.

FIGURE 5.6 Infective eggs of Ascaris suum (left, the thickshelled

egg contains a slender infective L3-larva) and Trichuris suis (right,
the thickshelled egg contains a slender infective Ll -larva).
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 29

The characteristics of Oesophagostomum sPp. and Hyostrongylus

rubidus are presented in Section 3.6, Identification of eggs and
larvae, while some characteristics of L3 larvae of non-porcine
helminths may be seen in the FAO guidelines on Helminth Parasites
in Ruminants (Hansen & Perry, 1994).

The infective helminth eggs which may be found in soil samples, are
identical to the corresponding eggs found in freshly deposited faeces
(see Section 3.6, Identification of eggs and larvae), except that they
may contain larvae, which may be clearly visible in a good
microscope (Fig. 5.6).
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 31

CHAPTER 6

INVESTIGATING HELMINTH OCCURRENCE AND EPIDEMIOLOGY

IN A PIG POPULATION

6.1 INTRODUCTION

This chapter deals with strategies for investigating helminth

occurrence in a pig herd or flock and helminth epidemiology, using
the methods described in Chapters 3-5. Investigations may be
divided into 3 parts:

Estimation of the helminth occurrence in a herd/flock

Long-term monitoring of helminth status in a herd/flock or

of a control programme

Plot experiments

While a) in general is the first step of an investigation, b) and c) are

two different ways of throwing light on the more dynamic
epidemiology of the helminth species, b) may give information on
the changes in prevalence rates and intensities of the infection in the
pig population over time, and c) may provide knowledge about the
development, dissemination and survival of the infective free-living
stages.

The unit of investigation is the herd or flock. Where groups of herds

are grazed communally or have close contact with each other, the
whole group should ideally be the unit of investigation. Helminth
infections generally involve an entire herd or flock, and to be
effective, diagnosis, treatment and control measures should be
directed at the entire unit.

When investigating a herd of pigs, one should be aware that many

1 32 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

factors may exert an influence on the helminth infections (number of

species, prevalence rates, infection intensities). One of the most
important is the age of the pigs combined with the immunogenicity
of the helminth species concerned (see Chapter 4.12). Other
important factors are management (housing facilities, hygiene) and
local seasonal climatic variations, which may have immense influence
on the transmission rates. Also, the reproductive status of the host
(dry or lactating sows), the nutritional status of the hosts, and any
control measures (management and anthelmintic routines, especially
the time span since the latest anthelmintic treatment) may closely
determine to the infection rates.

6.2 HELMINTH OCCURRENCE IN A HERD/FLOCK

To reveal the significance of the helminth infections within a

population of pigs, faecal samples (see Chapter 3) should be taken
from a representative number of live animals, preferably belonging to
selected age groups. Furthermore, as many dead (moribund or
sacrificed) animals as possible should be examined using the post-
mortem procedures (see Chapter 4).

Live animals

It is necessary to collect faecal samples from at least the following

three groups of pigs, to increase the chance for diagnosing the
parasite species present in the population, and in order to measure
the infection levels in those age groups in which the parasite species
are prevailing and potentially causing problems (Table 6.1 , where
only the most common parasites are mentioned).

Any animals with obvious clinical disease suggestive of parasitism

should be sampled. However, most helminth infections in pigs are
subclinical, and samples from diseased animals should never replace
the sampling from a representative number of randomly selected ani-
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 133

Table 6.1 Suggested age groups of pigs to sample, together with

some of their most prevalent parasites.
Age group of pigs to sample Most prevalent parasite species

piglets (2-3 weeks of age) Strongyloides ransomi, lsospora suis

fatteners (5-6 months of age) Ascaris suum, Trichuris suis

sows Oesophagostomum spp., Hyostrongylus

rubidus, Stephanurus den tatas, Eimeria
spp.

mals. The latter sampling is necessary for a full understanding of the

herd/flock problem.

There is no magic number of samples; but in general, the more

animals sampled, the greater the validity of the results and the better
the understanding of the infection patterns. The following suggested
numbers are based on both general principles and practical/logistical
constraints (Table 6.2), however, factors like expected prevalence
rates, variability in the egg counts, and accuracy of the estimations
should also be taken into account.

Table 6.2 Suggested sample sizes for given total numbers of pigs in
an age group.
Number of animals in Number (percentage) of animals to sample
the age group

1-10 all animals (100%)

11-25 10-15 animals (90%-60%)
26-100 15-30 animals (58%-30%)
101-200 30-40 animals (30%-20%)
201-500 40-50 animals (20%-10%)
>500 50- .. animals (10%)
1 34 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Dead (moribund or sacrificed) animals

As many as possible of the animals that die should be examined (see

Chapter 4 for post-mortem examinations). If a considerable number
within the flock is affected, one or two very ill or moribund animals
should be sacrificed for examination. However, care should be taken
with respect to the interpretation of results originating from only a
few animals, as some helminths (e.g. Ascaris suum) are heavily
overdispersed within the host population, and therei ore a f ew
negative post-mortem examinations do not exclude presence of
parasites.

6.3 LONG-TERM MONITORING OF A HERD/FLOCK

The initial estimation of helminth occurrence in a herd/flock is

important, but it does not provide full understanding of the
epidemiology. Therefore, a long-term monitoring of parasites in the
herds/flocks should be carried out, if possible.

Faecal samples (see Chapter 3) from living animals will provide the
most important information, but they may be supplemented with
post-mortem examinations (Chapter 4) on dead (moribund or
sacrificed) pigs or tracer (sentinel) pigs, and examination of soil/grass
(Chapter 5) may also be valuable. Furthermore, climatic data should
be recorded.

Live animals

If the flock/herd is large, and the sows are farrowing all the year
round, it may be possible to sample only the age groups specified in
Table 6.1. However, in most cases cohorts of piglets must be
selected (preferably several cohorts, starting at different times during
the year in order to reveal seasonal variation), and then identified
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 135

animals should be sampled repeatedly at specified intervals (see

below). With respect to the sows, a number of individuals should be
randomly selected at the beginning of the monitoring period, and
these individuals may then be sampled at the same specified
intervals.

Again, there is no magic number of samples, but the above

mentioned sample sizes (Table 6.2) is suggested.

Ideally, the sampling should be carried out every 2 weeks during

periods with suspected high transmission rates (rainy season,
summer season), continuing for approx. every month into the more
unfavourable season (dry season, winter season). For the remaining
dry/winter season, samplings every 4-6 weeks will be appropriate.
The samplings should optimally cover 3 calendar years to ensure the
recording of an average situation and provide an impression of the
year-to-year variation.

In practice, this ideal sampling may not be feasible. However, it is

acceptable to sample once a month during the rainy/summer season
(including the first month of the dry/winter season), and once every
2 months during the remaining part of the dry season/winter.

Dead (slaughtered, moribund or sacrificed) animals

Every opportunity should be taken to sample animals that die from

whatever reason, but sacrificing animals are not advocated for long-
term monitoring programmes unless special parasite problems are
suspected. Instead, pigs are slaughtered regularly (slaughter weight
depends on local traditions) and representative numbers of randomly
selected fatteners and sows should be examined at the various
seasons to monitor the worm burden.
1 36 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Tracer (sentinel) animals

Tracer pigs are intended to provide direct information on the

bioavailability of infective larvae/eggs in the environment, i.e. not
only a number of infective stages in a handful of grass or soil
selected by the researcher, but the number of parasites which in
practice are established in a pig grazing and rooting area.

Tracers should be parasite-naive. The principle is to place these

animals on the pasture for a predetermined short period of time (e.g.
2 weeks), during which they will pick up infective parasite stages.
The tracers are then moved to a completely helminth-free
environment for the following 4 weeks, during which the young
parasites will grow up so that they will be easy to identify post-
mortem. The pigs are then slaughtered and subjected to post-mortem
examinations. Ideally, 2 or more tracer pigs should be introduced to
the pasture once a month over at least one calendar year.

The suggested tracer protocol will work well for many helminths,
however it should be noticed that the large majority of worms of very
immunogenic species, like Ascaris suurn, may already have been
expelled by the tracer pigs, when these are slaughtered 4 weeks
after the last exposure.

The tracer principle has proven to be very valuable in epidemiological

research on ruminant helminths. However, there are several practical
problems associated with this principle in pig herds. Firstly, it may be
very difficult or impossible to rear helminth-free and non-immune
animals unless very intensively managed indoor herds are available.
Secondly, good animal housing facilities are required for the tracer
pigs after they have picked up the infections, as they should not be
further infected in these pens, and they should not be able to re-
infect themselves. Finally, if the tracer pigs are introduced into a
population of pigs with an established hierarchy, they will be
regarded as strangers in the flock and be more or less suppressed
and stressed, and as a consequence they may not graze and root
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 37

normally. The latter situation may, however, be eliminated either at

very low stocking rates, or by having the tracer pigs fenced off in a
separate area.

Sampling of grass/soil

When carrying out a long-term monitoring programme, it may be very

valuable to obtain measures on the numbers of infective stages in
soil and grass, even though the measures are not identical to the
bioavailability measured by tracer pigs (see above).

In practice, the sampling of soil and grass should follow the same
intervals as the animal sampling mentioned above (see Live animals
in this section). The samples should be taken at the same time of the
day on each occasion. Furthermore, every sampling should cover the
same area of the pasture. This should at least be the 'general'
grazing area, but very often it will be a good idea to include separate
samplings from suspected high-risk areas (e.g. muddy pools or other
wet areas).

6.4 PLOT EXPERIMENTS

Useful epidemiological information can be obtained by studying the

seasonal development, dissemination and survival of helminth stages
in or around pig faeces. Faecal material, containing a known number
of eggs, should be deposited on a pasture typical for the geographic
area, and the deposition should be carried out at regular intervals
throughout a minimUm period of one year.

The following is a simple experimental design, which will still provide

good information on transmission characteristics of helminths with
a direct life cycle.
1 38 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Identify a parasite-free grass-covered area representative of pastures

with pigs. As especially Ascaris and Trkhuris have eggs with a very
long potential survival time, it may be difficult to find a completely
helminth-free area, but then the background contamination should be
monitored before the start of the experiment.

Select at least 12 plots of approx. 1 m2 each. Fence these plots to

prevent grazing, and cover the plots with wide-meshed nets to keep
birds away. Care should also be taken to prevent mice, voles etc.
from disturbing the plots.

Cut the grass to a height similar to that of grazed paddocks, and

keep the grass at this level throughout the experiment, but without
removing the cut-off.

Identify a source of infected animals for continuous supply of faecal

material. If the origin of faeces is a commercial farm, fixed
arrangements should be made to exclude anthelmintic treatment of
the donor animals. If no naturally infected pigs are available, a group
of pigs should be experimentally infected.

Collect as much faeces as possible from the donor animals, but keep
the faecal clumps separate until faecal examinations have revealed
which of them contain the highest concentrations of eggs (or any
eggs at all).

Mix approx. 6 kg faeces so that all helminth species are present with
as high numbers of eggs as possible. If the faeces is too dry, water
may be added. Care should be taken to mix the faeces thoroughly.

Determine the egg concentrations in the final faecal mixture by

running 10 McMaster tests (preferably the concentration McMaster
technique; see Chapter 3) on 10 samples collected at different sites
in the material (if the variation between egg counts is too great, the
mixing and the McMasters should be repeated), and calculate the
mean EPGs.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 39

Deposit 5 kg faeces in small clumps of 10-50 g evenly on one plot.

Collect samples (small amounts of grass, soil beneath the faeces

clumps, and faecal material, as long as the faeces is not totally
decomposed) by taking a number of small subsamples at different
places on the plot, and combine these subsamples into 3 large
samples (i.e. grass, soil and faeces, respectively).

Analyze for eggs in faeces and in soil (EPG, determine species and
developmental stage), and for larvae in faeces, soil and grass (larvae
per gram, determine species). All methods are described in Chapter
5. All quantitative measures should be related to the total number of
eggs of each species deposited on the plot.

Samples should be collected frequently (e.g. 1 week intervals) shortly

after the deposition and with larger intervals later on. The exact
sampling protocol may, however, depend on the helminth species
present (slow or fast embryonation), the climate and the region. It is
recommended that the sampling continues for at least one year.

Repeat the deposition and thus the sampling schedule every month
for at least one year.

Monitor at least ambient temperature and rainfall daily, during the

entire period.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 141

CHAPTER 7

CONTROL OF HELMINTHS IN PIGS

7.1 GENERAL PRINCIPLES OF CONTROL

The purpose of a helminth control strategy is to keep the parasitic

challenge (especially to young pigs and lactating sows) at a minimum
rate to avoid clinical symptoms and production losses. Total
eradication from a geographical region is unlikely for most parasites,
due to the immense numbers of eggs shed and the high persistence
of the infective stages in the environment.

Before choosing and starting any control programme, it is necessary

to have a detailed knowledge about the helminth infections in the pig
population(s), i.e. helminth species present, prevalence rates and
transmission patterns. These characteristics may differ between
geographical regions, local herd management traditions etc. If this
knowledge is missing or is only scarce, any control programme
should, therefore, start with an investigation of the helminth
occurrence and epidemiology, as described in detail in Chapter 6.

The success of helminth transmission is to a large extent depending

on herd factors. This is visualized in Table 7.1, where a rough
overview of helminth occurrence in different production systems is
presented for a number of the most important helminths.

It is clearly seen that while extensively outdoor-reared pigs may

theoretically harbour all existing helminth species, intensive outdoor
management may eliminate those helminths whose transmission
depends on contact with human faeces, dog faeces or fresh water.
Furthermore, helminths which depend on intermediate hosts may be
more or less eliminated if the outdoor-reared pigs are confined to
concrete yards. All helminths with an indirect life cycle are eradicated
in indoor production systems, perhaps with the exception of Trichi-
1 42 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Table 7.1 An overview of anticipated helminth occurrence in

different production systems.

Helminth Summary of Management system

transmission
characteristics outdoor outdoor indoor indoor
extensive intensive extensive intensive

Ascaris eggs
Oesophagostomum larvae
Trichuris eggs
Strongyloides milkflarvae
Hyostrongylus larvae
Stephanurus larvae
Trichinae. musc les
Gongylonema dung beetles
Ascarops dung beetles
Physocephalus dung beetles
Metastrongylus earthworms
Echinococcus dog faeces
Fascia/a aquatic snails
Taenia solium human faeces

nella sp.. When indoor pig rearing develops from extensive to

intensive management, the number of helminths with a direct life
cycle will gradually decrease, Ascaris suum, and to a lesser degree
Oesophagostomum sp., being the most persistent of them all.

From this simple table it is evident that the most efficient way to
control porcine helminths is to improve the management and hygiene
of the herd. First of all, such improvement will eliminate some
helminth species, but additionally the WOTM burdens of the remaining
species may be reduced to more acceptable levels. Eradication of
helminth species is more or less impossible to obtain merely by
routine anthelmintic treatment programmes, which is the other major
control principle. Yet, it is often practically impossible to improve
management sufficiently, and therefore helminth control programmes
normally include both management and the use of anthelmintics.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 43

7.2 CONTROL OF NEMATODES

Apart from the beneficial results of improving management (see

above), control of nematodes may be achieved as described below.

7.2.1 Stocking rate

The density of pigs (stocking rate) in extensive outdoor pig rearing

should not be too high. Overstocking will force the pigs to a closer
contact with faecal material and may result in the consumption of a
higher number of infective nematode stages.

7.2.2 Grazing management

Grazing management may be used to minimize the uptake of infective

eggs/larvae and to create safe pastures. Alternating plant crops with
pig rearing will reduce the contamination on a pasture considerably,
although it should be recognized that infective eggs of especially
Ascaris and Trichuris may survive for years under favourable
conditions. The development of such grazing programmes requires
a thorough knowledge of the parasites seasonal development and
survival in the particular area. As an example, in the temperate
regions, the eggs of Oesophagostomum, Ascaris, and Trichuris
cannot embryonate and develop to infectivity during the winter (i.e.
below 10-1 5°C), but the two latter may accumulate and then
develop en bloc when favourable conditions turn up in the early
summer. Under these circumstances it may be beneficial to move the
pigs to a safe pasture just before the temperature increases in the
spring, and then eventually use the contaminated areas for crops.

7.2.3 Mixed or alternate grazing

As pigs have only few helminths in common with ruminants, grazing

 144 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

management may include mixed grazing (pigs grazing together with

ruminants) or alternate grazing (pigs alternating with ruminants on
the same pasture). It may be necessary to provide the sows with
nose-rings to avoid rooting and other serious damage to the
vegetation (see below). As seen in Tables 2.1 and 2.2, a few
helminths may infect both pigs and ruminants, and therefore local
identification of the helminth species present are recommended
before starting this regimen. Some of the species (e.g. the flukes
Fasciola hepatica and Schistosoma japonicum) may be controlled
simply by avoiding contact with freshwater, but Trichostrongylus
axei and Dicrocoelium den driticum may still be present. Furthermore,
when ruminants are using a pig contaminated area (outdoor, indoor),
attention should be paid to the risk of liver and lung lesions caused
by migrating Ascaris suum.

7.2.4 Hygiene of pens

When pigs are kept in concrete pens (outdoors, indoors), the dung
should be removed daily to reduce the large majority of nematode
eggs before they become infective. Furthermore, the floor should be
kept as dry as possible, as free-living stages of all helminths (even
Ascaris suum) require nearly 100% relative humidity to develop. The
draining capacity, and thus the dry microclimate at floor level, may
be the main reason why slatted floors in some intensive systems
seem to be rather effective in reducing helminth transmission
indoors.

It is often recommended to wash concrete floors (e.g. using high-

pressure devices or steam cleaning), but this recommendation is
questionable, as water may improve the general conditions for
egg/larval development and survival (it is impossible to remove all
eggs) and furthermore may help spreading the infective stages from
developmental foci (sheltered crevices etc.) to all over the pens.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 145

7.2.5 Dose and move

Before animals are moved to safe areas (outdoor, indoor), they may
be dosed with an anthelmintic to remove any worms present, in
order to keep the environment free of contamination for as long as
possible (dose and move). This principle has been shown to be rather
effective, although it unfortunately also increases the risk of
development of anthelmintic resistance (see Section 7.6).

7.2.6 Routine deworming

Routine deworming programmes often appeal to farmers for reasons

of convenience, and as a result worm treatments generally is the only
control measure carried out. However, the effect of each treatment
will be rather transitory if the pigs are re-infected continuously, while
the effect is considerably prolonged if the transmission rate is low.
Each treatment with a drug will increase the selection pressure for
development of anthelmintic resistance, and therefore helminth
control programmes should reduce the number of treatments to a
minimum and rather increase other control measures. Nevertheless,
some kinds of routine anthelmintic treatment are relevant in the
control of nematodes in most managements systems.

Several programmes for deworming of pigs have been worked out,

and most are adjusted to the age or the reproduction cycle. The
standard procedure is treatment of sows shortly before farrowing,
before a move to clean farrowing units. The objective is to eliminate
the worms from the sows in order to prevent contamination of the
environment of the newborn piglets. In dirty herds this treatment
may be supplemented by an additional treatment of the sows at
breeding (and treatment of boars and gilts). To reduce the infection
rates of the offspring, it is often recommended to treat piglets at
weaning and growing pigs once or twice during the fattening period,
although it should be realized that in some intensive systems piglets
and growing pigs do not need any routine drug treatment at all.
146 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

The choice of drug should partly depend on the worm species

present. Some drugs have a broader spectrum of activity than others,
some nematodes may be controlled only by certain drugs; some
drugs are more expensive than others etc. (see Section 7.5).
Furthermore, it is important to alternate between drugs with different
modes of action in order to reduce the risk of developing anthelmintic
resistance (see Section 7.6), and to avoid drugs against which
resistance has already developed.

7.2.7 Nose-ringing of sows

In some geographical areas it is common to fix an iron ring in the

snouts of sows kept on pasture, to reduce their rooting activity. As
this is an effective way to keep the grass cover on the pasture, it
may also reduce the uptake of soil-transmitted nematodes.

7.2.8 Adequate nutritional level

The overall effect of helminth infections may be reduced by ensuring

an adequate level of nutrition (especially proteins), although this
should be no substitute for a sound parasite control programme.

7.2.9 Genetic resistance

Very little is known about genetic resistance to helminth infections

in pigs, although a clear difference in infection levels between two
breeds has been described. Especially in sheep, genetic differences
in susceptibility have been documented within and between breeds,
and it is likely that such differences may exist in pigs as well.

7.2.10 The 'gilt-only' system for control of Stephanurus dentatus

The kidney worm Stephanurus dentatus has a very long prepatent

The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 147

period (6 to 19 months), and therefore the mature worms are

generally found in brood sows older than 2 years. The 'gilt-only'
system simply means that all breeding animals are consistently
slaughtered after having produced their first litter, and this
management practice has proven effective in the eradication of
kidney worms from a pig population.

7.2.11 Control of Trichinella spiralis

Pigs become infected with Trichinella by eating infected meat (wild

or domestic animals). Therefore, prevention of infection will depend
on the prevention of cannibalism, by avoiding any animal tissue being
fed to pigs without adequate cooking (boiling for 30 minutes), and
control of rodents.

7.3 CONTROL OF TREMATODES

Most digeneans which infect pigs include freshwater snails in their

life cycles (Fasciola hepatica, Fasciolopsis buski, Schistosoma spp.,
Opisthorchis noverca, see Tables 2.1 and 2.2, Chapter 2), and these
infections may be controlled simply by avoiding contact between pigs
and freshwater reservoirs (even small ponds and temporary pools),
or by preventing pigs from eating raw fish (Opisthorchis).

The only trematode which is not controlled in this way, is

Dicrocoelium dendriticum, as this fluke is transmitted terrestrially via
slugs and ants. However, Dicrocoelium may be controlled by keeping
the pigs in concrete yards or indoors.

7.4 CONTROL OF CESTODES

All cestodes infecting pigs have carnivorous mammalians as final

 1 48 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

hosts, and the metacestodes in the internal organs of pigs are

difficult to eliminate with drugs. Therefore, treatment for cestodes in
pigs is not included in any control programme.

All control strategies rely on the prevention of contact between pigs

and infected faecal matter. In the case of the zoonotic Taenia solium,
community sanitation is the ultimate control measure, preferably
supplemented by drug treatment of infected human. Similarly, T.
hydatigena in pigs may be controlled by preventing contact between
pigs and faeces of dogs (or other canids), although this parasite
seldom becomes a problem calling for special control activities.
Finally, Echinococcus spp. will never be regarded as merely a pig
infection, as it is clearly a community health problem, involving
especially dogs (and wild canids) as the main target for control
efforts. A well-coordinated eradication programme includes
information of the public in order to obtain routine drug treatment of
all dogs and to prevent re-infection of the dogs (and wild canids) by
inactivating all animal tissues/organs (thorough boiling or freezing)
when slaughtering domestic animals, including pigs. Furthermore,
eradication programmes must include control of wild canids if these
contribute significantly to the completion of the life cycle of the
parasite in the area.

7.5 ANTHELMINTICS

7.5.1 Definition

An anthelmintic is a compound which destroys or causes helminths

to be removed from the gastro-intestinal tract or other organs and
tissues they may occupy in their hosts.

Currently a series of safe anthelmintics are available, some with

broad spectrum activity and others with activity against specific
helminth infections. Many modern anthelmintics are effective against
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 149

both adults and larval stages, including dormant larvae.

Due to their cost, their tendency to delay or interfere with natural

host immunity mechanisms, and not least the rapidly increasing
prevalence of anthelmintic resistance, anthelmintics may not be the
most desirable method of managing helminth problems. However, in
many circumstances the sensible use of anthelmintic drugs is likely
to be an inevitable and often the only available method of controlling
helminth parasites. But, they should not be used indiscriminately.

7.5.2 Characteristics of an ideal drug

An ideal drug should have a broad spectrum activity against adult

and larval helminth parasites. A number of factors influence the
efficacy of an anthelmintic drug. The individual pigs often harbour
several different helminth species, which do not have the same
sensitivity to a given anthelmintic. In addition, there is usually a
difference in sensitivity between adult and larval stages, with
immature stages and especially dormant larvae being less sensitive
than the adult parasites. Furthermore, recent observations indicate
that the concentration of a drug in situ may depend profoundly on
the composition of the diet and the feeding regimen, with restricted
feeding increasing the concentration, and thus the efficacy, of some
orally administered drugs in the gastro-intestinal tract.

The ideal drug should also be metabolized rapidly in order to avoid

metabolic residues in pigs slaughtered for human consumption, and
thus to reduce the slaughter withdrawal period. Furthermore, the
long-lasting presence of subtherapeutic concentrations of a drug may
constitute a severe risk factor for the development of anthelmintic
resistance.

A good drug has low toxicity to the host, and the ratio of the
therapeutic dose to the maximum tolerated dose of pigs should be as
large as possible.
1 50 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

There should be no unpleasant side-effects to the pigs, the operator

or to the environment. Some drugs may cause inappetence or pain
at the injection site.

The selected drugs should be competitively priced and ready to use

correctly in an easy way. Furthermore, they should be stable and not
loose activity on exposure to normal ranges of temperature, light and
humidity.

7.5.3 Dosing methods

Oral dosing is by far the most common and easiest way of

administration of anthelmintics to pigs, because pigs, in contrast to
e.g. ruminants on pastures, depend on daily feeding in troughs.
Thus, many drugs are simply admixed to fodder, while oral dosing of
individual animals, as commonly done in ruminants, is not necessary.
This implies that pigs often are group-treated which unfortunately
sometimes results in a subtherapeutic dose in individual animals,
when the drug is only administered once. When drugs are
administered over several days, this risk is less. Furthermore, the
efficiency of a drug (especially Class I and Ill drugs, see below) may
be considerably increased by low dosing for several days.

A number of anthelmintics are available for injection. In order to

avoid local reactions (such as abscess formation at the injection site)
the highest possible hygienic standards should be maintained.

No anthelmintics are until now available in a formulation for external

application ("pour on" preparations) to pigs.

7.5.4 Anthelmintic classes

On the basis of their mode of action, anthelmintic drugs can be

subdivided into 5 classes.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 51

Class I anthelmintics: Benzimidazoles and pro-benzimidazoles. These

drugs exert their action on the intracellular polymerization of the
tubulin molecules to microtubules. As the cellular functions are
disrupted, the worms die. Examples of Class I compounds are
thiabendazole, fenbendazole, parbendazole, flubendazole, febantel,
and thiophanate.

Class II anthelmintics: lmidazothiazoles and tetrahydropyrimidines.

These drugs act on the acetylcholine receptor in the neuromuscular
system of the worms, causing a persistent depolarization of muscle
cells and a spastic paralysis of the worms, which are then removed
by gut motility. Examples of Class II drugs are levamisole, pyrantel,
and morantel.

Class Ill anthelmintics: Avermectins and milbemycins. The

compounds act on the nervous system of the worms, causing flaccid
paralysis and removal by gut motility. Class Ill consists of the
avermectins (ivermectin, doramectin) and the milbemycins
(moxidectin) that also have some effect against ectoparasites, e.g.
mange mites.

Class IV anthelmintics: Salicylanilids and substituted nitrophenols.

These drugs are typically used against bloodsucking parasites and are
not important in pigs.

Class V anthelmintics: Acetylcholine esterase antagonists. The Class

V drugs are organophosphorous compounds, which are only used to
a limited extent. Examples are dichlorvos and neguvon.

Piperazines have previously been classified as Class III anthelmintics.

These drugs act on the GABA receptors, causing flaccid paralysis of
the worms. However, recent knowledge indicates that their mode of
action is different from that of avermectins and milbymicins, and
cross resistance has not been documented.
1 52 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

7.5.5 Which drug to use ?

It is important first to identify the nature of the parasitic problem in

order to select the appropriate drug. As an example, control of
Strongyloides ransomi, which is transmitted to piglets via colostrum,
may be exerted by administration of avermectins to the sows, and
this may also be the drug of choice if both helminths and mange
(Sarcoptes spp.) should be controlled simultaneously. However, as
it will be emphasized below it is important to change regularly
between drugs of different anthelmintic classes, in order to delay
development of anthelmintic resistance, and if anthelmintic resistance
against a given drug has already developed, all drugs belonging to
this anthelmintic class should be completely avoided.

7.6 ANTHELMINTIC RESISTANCE

7.6.1 Definition and underlying mechanism

Anthelmintic resistance is defined as a significant increase in the

ability of individuals within a strain of parasites to tolerate doses of
a compound which would prove lethal to the majority of individuals
in a normal population of the same species.

Anthelmintic resistance constitutes a widespread and rapidly

increasing problem in helminth control programmes. The mechanism
behind anthelmintic resistance is simple selection. No drugs are able
to remove 1 00 % of the parasites exposed to the drug, and a few
worm individuals (the least susceptible) will survive, while the large
majority (the most susceptible) will be eliminated. When selection
continues repeatedly, the resistance genes will accumulate in the
worm population and the drug will loose its effect. It has been shown
that the general fitness of resistant worm populations is high.
Therefore, once field isolates have developed a solid anthelmintic
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 53

resistance, the likelihood of reversion to susceptibility is low, and

worm populations remain resistant for many years, even without
further selection.

Until recently, anthelmintic resistance was primarily confined to

trichostrongyle/strongyle nematodes of grazing small ruminants and
horses. In these host species, resistance to one, two or more classes
of anthelmintics (see Section 7.5) is now so widespread that several
farmers are left without any means of drug control of helminths of
grazing animals. The situation regarding helminths in pigs does not
seem to be quite so serious yet, but recent European reports indicate
that anthelmintic resistance may be rather common among
Oesophagostomum spp. in housed pigs.

7.6.2 Detection of anthelmintic resistance

Many cases of anthelmintic resistance, including resistant

Oesophagostomum strains in pigs, have been diagnosed after
specific investigations rather than after experiencing a breakdown of
control at farm level. This is probably attributable to the subclinical
course of most helminth infections.

The Faecal Egg Count Reduction Test (FECRT) is the most important
test to be used under field conditions, as it is applicable for all types
of anthelmintics and all species of helminths in which eggs are shed
in faeces. FECRT is simple to carry out:

Collect faecal samples from 20-30 identified animals which

have not been treated for at least 2-3 months (if
anthelmintic resistance is suspected for Ascaris or Trichuris
or other helminths with long prepatent periods, the pigs
should have been left untreated for considerably longer).

In the laboratory, the faecal samples are subjected to a

McMaster egg count procedure (Chapter 3.4).
1 54 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Distribute the animals by faecal egg counts, if practicable,

into 2 groups of at least 10 animals each.

The animals of one group should be carefully weighed and

dosed according to weight with the drug under suspicion
(the pigs of the other group are left untreated).

10-14 days post treatment, new faecal samples are

collected from the same individuals. Care should be taken
to reduce the risk of false positive egg counts (see Section
3.7.1) by housing the pigs in clean pens.

The faecal samples are subjected toa McMaster egg count

procedure, and larval cultures are set up in order to
differentiate between species which have eggs of the
strongylid type (Chapter 3.5).

Calculate the post-treatment arithmetric mean for egg

counts of the treated (5-et) and the control group (Rc) and
calculate the 95% confidence interval. The Faecal Egg
Count Reduction (FECR) is 100 (1 - 5-et/Re).

An anthelmintic is regarded as efficient if FECR>95%.

Resistance is present if the FECR<95% and the 95%
confidence level is less than 90%. If only one of the two
criteria is met, resistance is suspected.

Larval cultures will reveal which species are present after

treatment, and possibly indicate which species that might
be resistant.

If more than one drug (class of anthelmintic) is suspected to have

reduced efficiency, additional treatment group(s) must be included in
the trial. It is necessary to determine to which drug (class of
anthelmintics) the parasites are susceptible and immediately change
to an efficient drug. To ultimatively confirm the presence of
anthelmintic resistance, groups of pigs may be experimentally
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 55

infected with the isolate and subsequently subjected to treatment,

slaughter and worm counts. Two standard experimental designs
called Controlled slaughter assay and Critical slaughter assay exist,
but both are expensive and time-consuming. Additionally, a number
of in vitro procedures have been elaborated to detect anthelmintic
resistance by incubating isolated trichostrongyle/strongyle eggs or
larvae in serial concentrations of drugs and thereafter measure the
hatching, motility or survival of the parasites (Egg hatch test, Larval
development assay, etc.), or by using advanced biochemical
techniques (e.g. Tubulin binding assay). These techniques require
much experience and special laboratory equipment. None of these
have gained widespread application in the field.

A description of the in vivo methods for detection of anthelmintic

resistance is found in the recommendations from the World
Association for the Advancement of Veterinary Parasitology (Coles
et al. 1992, Veterinary Parasitology 4, 35-44).

7.6.3 Risk factors for development of anthelmintic resistance

Theoretically, a series of risk factors for development of anthelmintic

resistance has been recognized, and many of them have proven to
be important in practice. The most essential risk factors are listed
below.

Frequency of anthelmintic treatment. A number of surveys on

anthelmintic resistance unanimously conclude that the more
frequently parasitized animals are treated with anthelmintics, the
higher the risk for development of anthelmintic resistance. If the
intervals between treatments approach the prepatent period,
development of resistance may be rapid, as only individuals surviving
consecutive treatments will mate and produce more resistant
offspring. This is probably the main reason why the problem is so
widespread in especially horses, sheep, and goats, as these animal
species are often treated 5-12 times a year. In comparison, pigs are
normally treated 2-4 times a year, or less.
1 56 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Use of the same class of drugs for extended periods. As pointed out
in Section 7.5, there are many drugs and trademarks, but actually
only five classes of anthelmintics exist. As anthelmintics within the
same class have an identical mode of action, anthelmintic resistance
developed against one drug means that the resistant worm
population is also more or less resistant to the other drugs of the
class (side resistance). Therefore, treatments with drugs of the same
class for extended periods of time expose the worm population to a
consistent high selection pressure. This will evidently result in a more
rapid accumulation of resistant genes than if there had been a
systematic alternation between drugs with different modes of action.

Time of treatment. If only repressive treatments are used, i.e. the

infected pigs are treated while they remain in a heavily contaminated
environment (e.g. permanent pasture), resistant worms surviving the
treatment will produce resistant progeny, but due to the high number
of external infective stages, the resistant genes will quickly be
'diluted' with sensitive genes. However, most integrated control
programmes (e.g. strategic treatment at turnout and the dose and
move system) include treatment(s) before the animals are moved to
a clean pasture/pen, and consequently only survivors of the
treatment will contribute to the following generations, and the
frequency of resistant genes will increase more rapidly.

Dose size. Until recently, a correctly administered drug had to

eliminate only 80-90% of the worm population in order to be
recognized as an efficient anthelmintic. Now there is a general
agreement that all worms of an anthelmintic sensitive population
should be eliminated by a correct treatment. This tightening of the
criteria for an efficient drug is clearly based on the above-mentioned
fact that those few worms which are able to survive a treatment
constitute the basis for the development of anthelmintic resistance.
Similarly, underdosing has been shown to be a potential risk factor.
Common reasons for underdosing are that the farmer does not know
the weight of his animals, that he uses an average dose for all
animals in the flock (including the heaviest individuals), or that he
uses mass treatment by mixing the drug into the fodder or drinking
water, whereby some individuals may get too low doses.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 57

Pharmacokinetic behaviour of the drug. After administration,

anthelmintics show varying pharmacokinetic behaviour, i.e. when and
for how long time the drug concentrations are above the therapeutic
level, and for how long time a subtherapeutic, but still selective,
concentration persists. Furthermore, drugs will not reach identical
high concentrations everywhere in the body of the host, and hence
also the anthelmintic resistance selection pressure will differ with
parasite species.

Spread of resistant strains. The most important way of geographical

spread of anthelmintic resistance is by transport of host animals
harbouring resistant worm populations.

7.6.4 Prevention of anthelmintic resistance

There is an urgent need for development and adoption of strategies

to prevent anthelmintic resistance from being developed in pig
helminths, and to prevent the spread of already developed (but often
undiscovered) anthelmintic resistance.

Even though anthelmintic resistance in pig helminths is apparently

not a widespread problem at the present, it should be recognized that
new anthelmintics with a novel mode of action may possibly not be
expected on the market within the next decades. Hence, the source
of anthelmintics in the near future is the already existing one, and it
is very important to increase the life span of these anthelmintics by
reducing the risk of development of resistance.

Knowledge of risk factors (see above) provides veterinary advisers

with several practical recommendations which may delay the
development of anthelmintic resistance in pigs.

Reduce the dosing frequency/include afternative methods of control.

Anthelmintics should be used only when necessary, and should be
based on parasitological data and information about management and
hygiene. In the large majority of cases anthelmintic intervention may
be justified, but it should be combined with improved grazing
1 58 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

management, pen hygiene etc. in order to reduce the number of

treatments.

Use correct doses. When anthelmintic treatment is recommended,

care should be taken that the animals receive at least the full
recommended dose according to live weight. If the pigs are treated
individually, and not all of them are weighed, then the pigs should be
treated with a dose corresponding to the live weight of the heaviest
animal. A special problem arises when the pigs are treated flockwise
with a drug mixed up in the fodder or the drinking water. Here it may
be suspected that some pigs will be underdosed if the drug is
administered for only one day, while the risk of underdosing may be
reduced by administering the drug over several days in succession.
Many formulations of anthelmintics are easily adulterated.
Furthermore, it is strongly recommended that only registered drugs
from authorized sources should be purchased.

Rotate between anthelmin tic classes. When anthelmintic treatment

is suggested, present information recommends that the anthelmintics
from different classes (different modes of action) should be used in
a rotation scheme on a yearly basis. Such a programme could start
with the use of an anthelmintic from class I the first year, then a
compound from class II the following year, and thereafter a drug
from class Ill the third year. In the 4th year a benzimidazole (class I)
could be used again, etc. If resistance against one class has been
recorded, all drugs belonging to this class should, of course, be
abandoned from the rotation scheme.

Treat new animals effectively and establish quarantine. When new

animals, e.g. breeding animals, are to be introduced to the herd, they
should be kept separate from the rest of the herd for the first 3-7
days. It is wise to treat them a few days before arrival and a few
days after arrival, when they are still in quarantine. They should be
treated with anthelmintics, possibly with two or three drug classes,
each at the recommended dose.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 1 59

BIBLIOGRAPHY

Anderson, R.C., Chabaud, A.G., Willmott, S. 1974-1983. CIH Keys

to the Nematode Parasites of Vertebrates. No. 1-10. CAB.

Anonymous. Manual of Veterinary Parasitological Laboratory

Techniques. Reference Book 418. Ministry of Agricufture, Fisheries
and Food. London. 1987.

Armour, J. 1980. The epidemiology of helminth disease in farm

animals. Vet. Par. 6: 7-46.

Bjorn, H., Roepstorff, A. Waller, P.J. and Nansen, P. 1990.

Resistance to levamisole and cross resistance between pyrantel and
levamisole in Oesophagostomum quadrispinulatum and
Oesophagostomum dentatum of pigs. Vet. Par. 37: 21-30.

Coles, G.C., Taylor, M.A., Bauer, C., Borgsteede, F.H.M., Geerts, S.,
Klei, T.R., Waller, P.J. 1992. Methods for the detection of
anthelmintic resistance in nematodes of veterinary importance. Vet.
Par. 44: 357-359.

Connan, R.M. 1967. Observations on the epidemiology of parasitic

gastro-enteritis due to Oesophagostomum spp. and Hyostrongylus
rubidus in the pig. Vet. Rec. 80: 424-429.

Eriksen, L. 1981. Host parasite relations in Ascaris suum infection in

pigs and mice. Ph.D. Thesis, Royal Veterinary and Agricultural
University, Copenhagen, 193 pp.

Hansen, J. and Perry B. 1994. The Epidemiology, Diagnosis and

Control of Helminth Parasites of Ruminants. International Laboratory
for Research on Animal Diseases. Nairobi/FAO Rome.

Jacobs, D.E. and Dunn, A.M. 1969. Helminths of Scottish pigs:

occurrence, age incidences and seasonal variations. J. Helminth. 43:
327-340.
1 60 The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine

Levine, N.D. 1980. Nematode Parasites of Domestic Animals and of

Man. Burgess, Minneapolis, MN.

Murrell, K.D. 1986. Epidemiology, pathogenesis, and control of major

swine helminth parasites. Vet. Clin. N. Am.: Food Anim. Pract. 2:
439-454.

Nilsson, 0. 1982. Ascariasis in the pig. An epizootiological and

clinical study. Acta Vet. Scand. SuppL 79: 1-108.

Pattison, H.D., Smith, W.C. and Thomas, R.J. 1979. The effect of
sub-clinical nematode parasitism on reproductive performance in the
sow. Anim. Prod. 29: 321-326.

Pattison, H.D., Thomas, R.J. and Smith, W.C. 1980a. The effect of
subclinical nematode parasitism on digestion and performance in
growing pigs. Anim. Prod. 30: 285-294.

Pattison, H.D., Thomas, R.J. and Smith, W.C. 1980b. A survey of

gastrointestinal parasitism in pigs. Vet. Rec.107: 415-418.

Roepstorff, A. 1991. Transmission of intestinal parasites in Danish

sow herds. Vet. Par. 39: 149-160.

Roepstorff, A. and Jorsal, S.E. 1990. Relationship of the prevalence

of swine helminths to management practices and anthelmintic
treatment in Danish sow herds. Vet. Par. 36: 245-257.

Roepstorff, A. and Nansen, P. 1994. Epidemiology and control of

helminth infections in pigs under intensive and non-intensive
production systems. Vet. Par. 54: 69-85.

Roepstorff, A., Nilsson, 0., Oksanen, A., Gjerde, B., Richter, S.H.,
Örtenberg, E., Christensson, D., Martinsson, K.B., Bartlett, P.C.,
Nansen, P., Eriksen, L. HeIle, 0., Nikander, S. and Larsen, K. 1998.
Intestinal parasites in swine in the Nordic countries: prevalence and
geographical distribution. Vet. Par. 76: 305-319.
The Epidemiology, Diagnosis and Control of Helminth Parasites of Swine 161

Skerman, K.D. and Hillard, J.J. 1966. A Handbook for Studies of

Helminth Parasites of Ruminants. Near East Animal Health Institute,
Teheran. Handbook No. 2. Rome: Food and Agricultural Organization
of the United Nations.

Soulsby, E.J.L. 1982. Helminths, Arthropods and Protozoa of

Domesticated Animals, 7th edn. Lea and Febiger, Philadelphia, PA.

Stewart, T.B., Hale, O.M. and Andrews, J.S. 1964. Eradication of

the swine kidney worm, Stephanurus dentatus, from experimental
pastures by herd management. Am. J. Vet. Res. 25: 1141-1150.

Urquhart, G.M., Armour, J., Duncan, J.L., Dunn, A.M. and Jennings,
F.W. 1987. Veterinary Parasitology, Longman Scientific & Technical,
Essex.
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