77

7
Genetic approaches in Aedes aegypti for control of dengue:
an overview
Ken E. Olson
1
, Luke Alphey
2
, Jonathan O. Carlson
3
, Anthony A. James
4
Abstract
The mosquito-borne dengue viruses (DV) cause an estimated 50 million human
infections annually. The incidence of severe dengue disease in Southeast Asia and
Latin America is increasing at an alarming rate. There are currently no vaccines or
anti-viral therapies available to mitigate dengue disease. Current methodologies for
controlling the principal vector, Aedes aegypti, are inadequate and ineffective. A
potential solution to this growing human-health crisis is to develop new genetics-
based vector control (GVC) approaches as part of an integrated control strategy. GVC
includes both population reduction and population replacement strategies and
represents a broad spectrum of genetic mechanisms at various stages in their
development for field-testing. To realize the full potentials of these GVC strategies it
is critical that we investigate, evaluate and, where appropriate, develop these
strategies to the point where they can be deployed at field sites in one or more
disease-endemic countries (DECs).
Keywords: dengue; Aedes aegypti; genetic-based vector control
Introduction
Dengue fever (DF) and its more serious form, dengue hemorrhagic fever (DHF)
and dengue shock syndrome (DHF/DSS) are caused by four closely related but
antigenically distinct, single-strand RNA viruses transmitted by mosquitoes to
humans. DVs cause more human morbidity and mortality than any other vector-borne
viral disease with 2.5-3.0 billion people at risk of infection and 50-100 million DF and
250,000-500,000 DHF/DSS annual cases (Gubler 1996; 1998). All four DV serotypes
cause disease and case-fatality rates for untreated DHF/DSS can be 30-40%. The risk
of DHF/DSS is highest in areas where two or more DV serotypes are transmitted
(Halstead 1988; Monath 1994; Rigau-Perez et al. 1998). At this time, there is no
licensed vaccine and no clinical cure for the disease.
1
Arthropod-borne and Infectious Diseases Laboratory, 3185 Rampart Road, Mail Delivery 1692,
Foothills Research Campus, Department of Microbiology, Immunology and Pathology, Colorado State
University, Fort Collins, CO 80523, USA. E-mail: kolson@colostate.edu
2
Department of Zoology, Wellcome Trust Center, University of Oxford, Oxford OX1 3PS, UK.
E-mail: luke.alphey@zoology.oxford.ac.uk
3
Department of Microbiology, Immunology, and Pathology, Colorado State University, Fort Collins,
CO 80521-1692, USA. E-mail: jonathan.carlson@colostate.edu
4
Department of Microbiology & Molecular Genetics, Department of Molecular Biology &
Biochemistry, University of California, Irvine CA 92697-3900 USA. E-mail: aajames@uci.edu
Chapter 7
78
Ae. aegypti is by far the most important and efficient vector of DV because of its
affinity for humans (Gubler 1998). Dengue control currently depends on reduction or
elimination of Ae. aegypti. In the 1940-1960s most tropical American countries used
integrated programmes of environmental management and insecticides to eliminate
mosquitoes (Gubler 1998), but many of these were abandoned in the early 1970s
(Reiter and Gubler 1997). Ae. aegypti re-infested countries where it had been
eliminated and dengue epidemics renewed. In 2004, Ae. aegypti is distributed more
widely than it was before eradication began, and is now in large urban areas where a
greater number of people than in the past are at risk (Gubler 2004). Remarkably,
despite the successes of the past, current dengue vector control programmes are often
nonexistent or ineffective (Reiter and Gubler 1997). Rather than maintaining
integrated programmes that specifically target Ae. aegypti, ministries of health merged
all mosquito control and relied on outdoor applications of aerosol insecticides to kill
adult mosquitoes. This costly approach is ineffective in most cases because the
majority of females rest indoors where they avoid insecticide contact (Reiter and
Gubler 1997). Furthermore, many insecticides are useless due to the spread of
resistance (Hemingway, Field and Vontas 2002).
Several GVC strategies for reducing DV transmission have been identified as
potential dengue disease control methods and are designed either to reduce the overall
population of DV-transmitting vectors or to replace existing vector populations with
populations that cannot transmit the virus. Two vector population reduction
approaches are currently being investigated and are in early laboratory cage trials. The
first population reduction strategy is the development and use of natural or genetically
engineered densoviruses that are pathogenic to Ae. aegypti (Carlson, Afanasiev and
Suchman 2000). The second population reduction strategy is the development and use
of insects carrying dominant lethal mutations (RIDL, see below, Thomas et al. 2000).
This approach would require mating of genetically modified vectors (GMV)-RIDL
males with local vector populations producing offspring that die prior to becoming
adults. Both approaches are designed to reduce transmission of DVs by reducing the
vector population. Approaches designed to replace populations of vectors are more
long-term, but could have significant consequences for dengue disease control in the
future (J ames 2000). In these approaches, an effector gene, such as an anti-DV gene,
is appropriately expressed to block transmission by the vector. GVC approaches
require identification of tissue-specific promoters, anti-pathogen effector genes, and
genetic drive mechanisms such as synthetic transposable elements (TE) to introgress
the effector gene into the population, eliminating vector competence. Successful GVC
strategies will require knowledge of vector ecology in DECs and large cage trials in
DECs prior to release of biocontrol agents or GMVs.
Current state of the art
Genetic approaches leading to vector population reduction
Mosquito densoviruses as tools for population reduction and transduction
The Aedes densonucleosis virus (AeDNV; family Parvoviridae) is mosquito-
specific and does not infect vertebrates or non-target invertebrates. Larvae are
infected in oviposition sites and die in a dose-dependent manner depending on viral
titre and stage of infection. AeDNV is maintained through metamorphosis and is
transmitted vertically to offspring (Barreau, J ousset and Bergoin 1997). Infected
female mosquitoes deliver viruses to multiple breeding sites and viral concentrations
Olson et al.
79
increase as larvae become infected and shed, thus increasing horizontal transmission
to other larvae. Survival of infected adult females also decreases significantly in a
dose-dependent manner (Kuznetsova and Butchasky 1988, Suchman and Carlson,
unpublished). Shortening the female adult lifespan would reduce vectorial capacity
since a significant proportion of females would not survive the extrinsic DV
incubation period. Recently, a number of other densoviruses have been discovered
that also may be adapted as biocontrol and transducing agents (Kittayapong, Baisley
and O'Neill 1999).
AeDNV research has the most immediate potential to deliver products for an
effective field trial once a field site is selected and more extensive cage experiments
completed. Prototype population cage experiments testing the ability of AeDNV to
persist, spread and reduce mosquito populations have already been performed and are
encouraging: a relatively low inoculum of virus in a larval rearing site replicates to
levels that reduce the mosquito population, and female mosquitoes originally from
the site inoculate virus into new sites.
Critical laboratory needs and challenges for using densoviruses as biocontrol
agents
Optimize densovirus preparations and use of AeDNV in cage experiments
Laboratory-based cage experiments need to be performed to determine 1) if other
densoviruses persist and spread more efficiently than AeDNV; 2) if mosquitoes from
DEC field sites are susceptible to AeDNV; 3) if different strains of mosquitoes vary in
their susceptibility to other mosquito DNVs; 4) if large-scale production and use of
the AeDNV bio-control agent is feasible; and 5) whether recombinant viruses
expressing anti-vector effectors (such as RNAi interference targeting the expression
of critical vector genes) can enhance lethality of the virus for Aedes aegypti larvae.
Large-scale cage trials to assess densovirus potential for persistence and spread
Large-scale cages can be used to replicate laboratory experiments with natural
populations under field conditions. Ae. aegypti from long-term cage experiments need
to be compared with the local populations outside the cage at regular intervals by
genetic analyses to look for genetic effects of the virus on populations
(Gorrochotegui-Escalante et al. 2002; Garcia-Franco et al. 2002; Root et al. 2003).
These studies should yield valuable data on the ability of the virus to persist and
spread in a wild mosquito population, and to control mosquito populations in the field.
These studies will also help refine the experimental design for cage experiments for a
number of GVC strategies.
Development and use of Release-of-Insects-carrying-a-Dominant-Lethal (RIDL),
a GMV-based development of Sterile-Insect Technique (SIT).
Drosophila melanogaster has been engineered with the basic genetic properties of
an RIDL strain (Thomas et al. 2000; Heinrich and Scott 2000), using a repressible
gene expression system (tet-off) based on the tetracycline-repressible transactivator
tTA (Baron and Bujard 2000; Gossen and Bujard 1992). Mathematical modelling
suggests that for insects with strong density-dependent regulation of population size, a
RIDL system imposing lethality at a larval or pupal stage has major advantages over
conventional SIT and will provide a simple and effective dengue control method.
Preliminary studies in Ae. aegypti in which tTA is expressed under tetO/hsp control,
Chapter 7
80
produced >95% lethality in larvae in the absence of tetracycline (Alphey et al.,
unpublished data). Two transgenic lines are currently being evaluated in cage trials.
These lines are being introgressed into the genetic background of local transmitting
strains of Ae. aegypti to study fitness issues, release parameters, and population-
dynamics overtime.
Critical laboratory needs and challenges for using RIDL to reduce vector
populations
Increase the penetrance of RIDL-induced lethality
An ideal RIDL system would kill 100% of the individuals supposed to be affected.
This is not essential for population suppression or to prevent the spread of the
transgene within the target population, and indeed is not provided by current
radiation-based SIT programmes for other insects. However, the system can in
principle be refined in this regard by using alternative RIDL effectors, such as pro-
apoptotic genes (Heinrich and Scott 2000), generating and testing more strains with
the current constructs, or combining more than one insertion or construct to give a
more highly penetrant and redundant system.
Construct a female-specific RIDL system
Most known female-specific promoters from Aedes are induced after the uptake of
the blood meal. A RIDL system could potentially be developed around such a
promoter to cause females to die soon after biting. Such females would be unable to
transmit DV, which have a 10-14 day extrinsic incubation period. Alternatively, it
should be possible to identify the sex-specific elements of Aedes Actin-4, a gene that
expresses an actin in the female pupal developing flight muscles (Muoz et al. 2004),
to drive pre-adult lethality and thereby prevent biting-female development. Such a
system would also avoid the need for physical sexing of the release generation and
potentially allow the release of any of a wide range of developmental stages.
Determine key parameters for eventual use of RIDL technology in the field
These parameters include the economic and fitness costs of mass rearing of GMV-
RIDL strains, the effect of the release ratio (GMV-RIDL / wild-type) of release into
cage populations for optimal (most cost-effective) population reduction, and the
ability of GMV-RIDL to compete with local mosquitoes for mating and resources.
These parameters will feed into a suitable combined epidemiological and
entomological model of dengue transmission, the development of which is another
key requirement. This will provide a realistic estimate of the cost-effectiveness of
RIDL, and a rational method for comparing this to other approaches, applied singly or
in combination, in different transmission regimes.
Genetic approaches leading to vector population replacement
Much work has focused on developing GMVs that are refractory for DV
transmission by developing germ-line-transformed Ae. aegypti that appropriately
express an anti-pathogen effector gene. By targeting the pathogen, rather than the
vector, expression of the effector gene should have minimal impact on the
reproductive fitness of the GMV. The long-term goal is to replace existing
transmission-competent vector populations with GMV populations that are no longer
permissive for DV transmission. Replacement of Ae. aegypti populations to block DV
transmission may be a real alternative to current vector control strategies. Ae. aegypti
Olson et al.
81
is responsible for most of the severe dengue epidemics, it is relatively easy to
manipulate genetically and maintain in the laboratory, and the vectors continuously
exchange genes locally and appear to have few gene flow barriers within 150 km
(Gorrochotegui-Escalante et al. 2002). At least three genetic-transformation systems
have been described and used successfully in Ae. aegypti to generate GMVs. These
transformation systems are based on the Class II TEs Mos1 (Mariner), Hermes and
piggyBac (J asinskiene et al. 1998; Coates et al. 1998; Kokoza et al. 2000). Mos1 and
piggyBac are the most commonly used TEs for generating GMVs.
Anti-dengue virus effector genes RNAi
During the last three years, considerable progress has been made toward
identifying effector genes that can profoundly reduce Ae. aegypti competence for DV
transmission (Adelman et al. 2001; 2002; Olson et al. 2002; Tavanty et al. 2004). The
major thrust of research has been to design and express double stranded RNAs
(dsRNAs) that make DV-susceptible cells non-permissive for virus replication. This
strategy is based on RNA interference (RNAi), an ancient potent, innate immune
response in insects and a related response termed post-transcriptional gene silencing
in plants (Tijsterman, Ketting and Plasterk 2002).
We now know that Drosophila melanogaster, Caenorhabditis elegans, humans and
plants have the RNAi pathway, which is triggered by the presence of intracellular
double-stranded RNA (dsRNA). The presence of dsRNA in cells is an early warning
signal of RNA-virus invasion that directs an innate response resulting in destruction
of any mRNA having sequence identity with the dsRNA. Many RNA viruses generate
dsRNA in infected cells as a byproduct of replication and these replicative
intermediates serve as potent recognition patterns for inducing the RNAi intracellular
response. If RNA viruses trigger RNAi, why are mosquitoes such efficient vectors of
arboviruses? We do not know for sure, but DV may escape the antiviral effects of
RNAi in competent mosquitoes either by failing to present the threshold concentration
of dsRNA molecules required for triggering the response or by encoding a viral
protein that suppresses the RNAi response. Currently, there is no evidence for a DV
RNAi suppressor protein. However, Uchil and Satchidanandam (2003) have recently
shown that the dsRNA replicative form (RF) of DVs is sequestered in double-
membrane structures in the cytoplasm of infected cells which may limit RF exposure
to the RNAi pathway.
RNAi is activated by dsRNA and results in a reduced steady-state level of specific
RNA molecules with sequence similarity to the dsRNA (Cogoni and Macino 1997;
Vaucheret et al. 1998). The mechanism of RNAi has been studied in some detail in
Drosophila melanogaster. In the fruitfly, the RNase III enzyme Dicer is responsible
for digesting dsRNA into 21-23 bp small interfering RNAs (siRNAs). The siRNAs are
then unwound into single-stranded siRNAs in an ATP-dependent step and
incorporated into an enzyme complex termed the RNA-induced silencing complex
(RISC). The single-stranded siRNAs guide RISC to the target mRNA and the
complex cleaves the message or inhibits its translation (Schwarz et al. 2002). This
strategy has been used in transgenic plants to develop resistance to a number of RNA-
virus pathogens. Several groups now have evidence that mosquito species such as Ae.
aegypti, Anopheles stephensi and An. gambiae develop an RNAi response very similar
to that found in D. melanogaster. These vectors are capable of silencing endogenous
gene expression or virus replication after introduction of dsRNA targeted to a specific
gene (Adelman et al. 2002; Travanty et al. 2004; Brown et al. 2003). Replication of
several arboviruses appears to trigger the RNAi response in mosquito cells and we
Chapter 7
82
now have evidence for the genes involved in the An. gambiae antiviral response to the
arbovirus Onyong-nyong alphavirus (Togaviridae) (Sanchez-Vargas et al. 2004;
Keene et al. 2004).
RNAi maybe an Achilles heel for replication of RNA viruses and we should be
able to induce a robust RNAi response to DVs in the midgut or other relevant tissues
of a transgenic mosquito by expressing DV-specific dsRNA. This strategy would
sensitize the cells to the presence of the RNA virus leading to the destruction of the
virus genome either as the virus uncoats or following virus transcription in the cell.
The midgut is a likely target for mounting this line of defence because it is the first
tissue the virus encounters in the vector and is the major determinant of vector
competence in the mosquito. In addition, oral infection of midguts with high
concentrations of virus begin with relatively few foci of infection of epithelial cells
that spread throughout the gut over a 5-7 day period prior to dissemination. A virus-
specific dsRNA should be able to suppress DV replication during that time frame.
Both Ae. aegypti midgut and salivary-gland promoters are available to test whether
RNAi can be used to promote resistance to DVs in the vector (J ames et al. 1991;
Moreira et al. 2000).
The RNAi approach of developing resistance in Ae. aegypti has the following
advantages: 1) RNAi does not require expression of a potentially antigenic protein; 2)
the strategy utilizes the machinery of a natural innate immune response that is present
in the mosquito (Sanchez-Vargas et al. 2004); 3) a number of anti-DV dsRNA
effector sequences have already been identified that cause profound resistance in
mosquito cell culture and in adult mosquitoes (Adelman et al. 2001); 4) the anti-DV
dsRNA effector sequence (500-600 bp) should be less prone to the effects of single-
point virus mutations and selection since the active units of RNAi activity are 21-23
bp siRNA blocks formed from the dsRNA trigger (Travanty and Olson, unpublished
data, Blair, Adelman and Olson 2000); 5) transgenic lines that express dsRNAs from
several non-Ae. aegypti promoters have now been generated (Travanty et al. 2004); 6)
DV-2 pathogenesis studies of virus in Ae. aegypti have been performed to determine
the temporal and spatial infection patterns of the virus after oral infection (Sanchez-
Vargas and Olson, unpublished data); 7) DV challenge protocols for assessing
resistance in transgenic mosquitoes are available (Sanchez-Vargas and Olson,
unpublished data).
Critical laboratory short-term needs and challenges for using RNAi-based
disease control strategies and other effector gene strategies
Identify Ae. aegypti midgut and salivary gland promoters that can be utilized to
deliver anti-DV at the correct time and place in the mosquito tissue.
We are currently evaluating the Ae. aegypti ferritin heavy chain, carboxypeptidase,
GFAT and glutamine synthetase midgut promoters and the D7 and apyrase salivary-
gland promoters for gene-expression potential. To test both RNAi and promoter
activity we are developing transgenics that express GAL4 and transgenics with anti-
DV dsRNA expression under UAS control (Brand and Perrimon 1993). The two lines
can be crossed and offspring evaluated for RNAi efficacy. Identifying suitable
promoters is a key to this strategy. It is apparent that the siRNA 23-nucleotide signal
is not amplified in insects as it is in plants and C. elegans therefore RNAi probably
does not spread from cell to cell in mosquitoes (Hoa et al. 2003). This makes it critical
that the antiDV dsRNA is expressed in the same vector cells that are critical for DV
infection and replication.
Olson et al.
83
Identify the most efficient construct format for delivering the dsRNA
Currently we are designing effector RNAs that comprise 300 bases of DV target
sequence in a sense orientation followed by Ae. aegypti intron sequence and an exact
antisense complement of the sense RNA (Adelman et al. 2002; Travanty et al. 2004).
There may be a need to develop new constructs for expression in mosquitoes that
form larger dsRNAs in the 500-600 bp range. Does the intron size matter, since it is
ultimately cleaved? What untranslated sequences are needed to stabilize expression of
the effector gene in target tissues?
Identify the specificity of an effector dsRNA based on DV2 sequence
Will it protect the mosquito from infection with other DV2 genotypes or other DV
serotypes? There is indication that it is possible to target multiple serotypes by
carefully choosing DV-specific target sequences (Sanchez-Vargas et al. 2004). Will
this approach drive selection of DV with altered infection characteristics?
Develop a recombinant/reporter virus to rapidly assess RNAi in transgenic
mosquitoes
Researchers have considerable experience developing infectious cDNA clones of
flaviviruses and alphaviruses and have developed alphaviruses that express eGFP as a
marker of infection (Foy et al. 2004; Keene et al. 2004). The development of a DV-
expressing GFP as a marker would greatly facilitate identification and
characterization of transgenic lines for virus resistance.
Development of protein-based effector genes
A number of effector-gene strategies will most likely need to be developed to
engineer resistance effectively into vector populations. Ito et al. (2002) showed that
peptides recognizing mosquito-tissue surface proteins block entry of a malaria
sporozoite into the salivary glands of a transgenic mosquito. The challenge here is to
identify effector proteins that block DV transmission yet can be effective against a
rapidly evolving RNA virus. These peptide-based effectors could take the form of
single-chain antibodies (Cappuro et al. 2000??) that bind to and neutralize DV or
mimic the envelope glycoprotein domain-III region of DVs (Hung et al. 2004).
Long-term research challenges for GVC-replacement technology
Development of an efficient anti-DV effector gene is only the first step towards the
long-term goal of using genetically manipulated insects to control DV. We also need
to demonstrate that transposon-mediated systems or other genetic drive systems will
successfully invade field populations. The first step in this process is to evaluate
transposon-mediated drive of genes through mosquito cage populations. In D.
melanogaster, studies with autonomous (self-mobilizing constructs that carry a copy
of their transposase within the transposon) and non-autonomous (stable constructs
mobilized only by externally supplied transposase) TEs carrying marker genes have
shown that elements will increase the frequency of the marker gene when introduced
into cage population of flies (Carareto et al. 1997). This mobility was characterized by
a tight linkage of the transposon with an active marker gene for as many as 40
generations. However, stability of the marker gene varied inversely with the size of
the final, loaded, autonomous element. Researchers need to conduct cage
experiments to evaluate the mobility and stability of loaded autonomous TEs as they
spread through cage populations of mosquitoes; maintenance of the integrity of the
loaded TE during population replacement and beyond is one of several major
Chapter 7
84
challenges to the development of usable gene drive systems. Obviously serious
discussions must take place to identify potential field sites for evaluation of control,
especially those strategies involving vector replacement strategies.
Future directions for research and capacity/partnership building
Discussion of other laboratory and field research that will need to be performed to
realize GVC approaches fully is found elsewhere in this book. Critical research needs
include the development and the characterization of genetic drive mechanisms, the
development of a much more complete understanding of the ecology of dengue
disease transmission in DECs, and the formation of full and meaningful partnerships
with DECs to evaluate GVC approaches. To realize the full potentials of GVC
strategies it is critical that we investigate, evaluate and, where appropriate, develop
GVC strategies to the point where they can be deployed at field sites in one or more
DECs. A number of gaps in knowledge have slowed or prevented the development of
genetic control methods. These gaps exist between the state-of-the-art laboratory
development of novel anti-DV tools and knowledge of field properties of mosquitoes
that will affect their use, and between scientists in the developed world and the DEC
scientists who would be responsible for implementing the technology. Further gaps
exist among scientists and the agencies that would be responsible for the deployment
of any genetic control strategy, and in policies and procedures for evaluating how
genetic control methods fit into the overall strategy of existing or planned control
programmes; these problems have become acute as the tools have now been
developed to allow implementation of some methods. Finally, gaps exist between the
enthusiasm of scientists for these genetic methods and the level of awareness of
potential end-users of the risks and benefits of using them for controlling dengue
transmission.
References
Adelman, Z. N., Blair, C. D., Carlson, J . O., et al., 2001. Sindbis virus-induced
silencing of dengue viruses in mosquitoes. Insect Molecular Biology, 10 (3),
265-273.
Adelman, Z.N., Sanchez-Vargas, I., Travanty, E.A., et al., 2002. RNA silencing of
dengue virus type 2 replication in transformed C6/36 mosquito cells
transcribing an inverted-repeat RNA derived from the virus genome. Journal
of Virology, 76 (24), 12925-12933.
Baron, U. and Bujard, H., 2000. Tet repressor-based system for regulated gene
expression in eukaryotic cells: principles and advances. Methods in
Enzymology, 327, 401-21.
Barreau, C., J ousset, F.X. and Bergoin, M., 1997. Venereal and vertical transmission
of the Aedes albopictus parvovirus in Aedes aegypti mosquitoes. American
Journal of Tropical and Medical Hygiene, 57 (2), 126-131.
Blair, C.D., Adelman, Z.N. and Olson, K.E., 2000. Molecular strategies for
interrupting arthropod-borne virus transmission by mosquitoes. Clininical
Microbiology Reviews, 13 (4), 651-661.
Brand, A.H. and Perrimon, N., 1993. Targeted gene expression as a means of altering
cell fates and generating dominant phenotypes. Development, 118 (2), 401-
415.
Olson et al.
85
Brown, A.E., Bugeon, L., Crisanti, A., et al., 2003. Stable and heritable gene silencing
in the malaria vector Anopheles stephensi. Nucleic Acids Research, 31 (15),
e85/1-6.
Carareto, C. M. A., Kim, W., Wojciechowski, M. F., et al., 1997. Testing transposable
elements as genetic drive mechanisms using Drosophila P element constructs
as a model system. Genetica, 101 (1), 13-33.
Carlson, J ., Afanasiev, B. and Suchman, E., 2000. Densoviruses as transducing
vectors for insects. In: Handler, A.M. and J ames, A.A. eds. Insect
transgenesis: methods and applications. 319-333, Boca Raton, 139-159.
Coates, C. J ., J asinskiene, N., Miyashiro, L., et al., 1998. Mariner transposition and
transformation of the yellow fever mosquito, Aedes aegypti. Proceedings of
the National Academy of Sciences of the United States of America, 95 (7),
3748-3751.
Cogoni, C. and Macino, G., 1997. Isolation of quelling-defective (qde) mutants
impaired in posttranscriptional transgene-induced gene silencing in
Neurospora crassa. Proceedings of the National Academy of Sciences of the
United States of America, 94 (19), 10233-10238.
Foy, B.D., Myles, K.M., Pierro, D.J ., et al., 2004. Development of a new Sindbis virus
transducing system and its characterization in three Culicine mosquitoes and
twoLepidopteran species. Insect Molecular Biology, 13 (1), 89-100.
Garcia-Franco, F., Munoz Mde, L., Lozano-Fuentes, S., et al., 2002. Large genetic
distances among Aedes aegypti populations along the South Pacific coast of
Mexico. American Journal of Tropical and Medical Hygiene, 66 (5), 594-598.
Gorrochotegui-Escalante, N., Gomez-Machorro, C., Lozano-Fuentes, S., et al., 2002.
Breeding structure of Aedes aegypti populations in Mexico varies by region.
American Journal of Tropical and Medical Hygiene, 66 (2), 213-222.
Gossen, M. and Bujard, H., 1992. Tight control of gene expression in mammalian
cells by tetracycline-responsive promoters. Proceedings of the National
Academy of Sciences of the United States of America, 89 (12), 5547-5551.
Gubler, D.J ., 1996. The global resurgence of arboviral diseases. Transactions of the
Royal Society of Tropical Medicine and Hygiene, 90 (5), 449-451.
Gubler, D.J ., 1998. Dengue and dengue hemorrhagic fever. Clininical Microbiology
Reviews, 11 (3), 480-496.
Gubler, D.J ., 2004. Cities spawn epidemic dengue viruses. Nature Medicine, 10 (2),
129-130.
Halstead, S.B., 1988. Pathogenesis of dengue: challenges to molecular biology.
Science, 239 (4839), 476-481.
Heinrich, J .C. and Scott, M.J ., 2000. A repressible female-specific lethal genetic
system for making transgenic insect strains suitable for a sterile-release
program. Proceedings of the National Academy of Sciences of the United
States of America, 97 (15), 8229-8232.
Hemingway, J ., Field, L. and Vontas, J ., 2002. An overview of insecticide resistance.
Science, 298 (5591), 96-97.
Hoa, N.T., Keene, K.M., Olson, K.E., et al., 2003. Characterization of RNA
interference in an Anopheles gambiae cell line. Insect Biochemistry and
Molecular Biology, 33 (9), 949-957.
Hung, J .J ., Hsieh, M.T., Young, M.J ., et al., 2004. An external loop region of domain
III of dengue virus type 2 envelope protein is involved in serotype-specific
binding to mosquito but not mammalian cells. Journal of Virology, 78 (1),
378-388.
Chapter 7
86
Ito, J ., Ghosh, A., Moreira, L.A., et al., 2002. Transgenic anopheline mosquitoes
impaired in transmission of a malaria parasite. Nature, 417 (6887), 452-455.
J ames, A.A., Blackmer, K., Marinotti, O., et al., 1991. Isolation and characterization
of the gene expressing the major salivary gland protein of the female
mosquito, Aedes aegypti. Molecular and Biochemical Parasitology, 44 (2),
245-253.
J ames, A.J ., 2000. Control of disease transmission through genetic modification of
mosquitoes. In: Handler, A.M. and J ames, A.A. eds. Insect transgenesis :
methods and applications. CRC Press, Boca Raton, 319-333.
J asinskiene, N., Coates, C. J ., Benedict, M. Q., et al., 1998. Stable transformation of
the yellow fever mosquito, Aedes aegypti, with the Hermes element from the
housefly. Proceedings of the National Academy of Sciences of the United
States of America, 95 (7), 3743-3747.
Keene, K.M., Foy, B.D., Sanchez-Vargas, I., et al., 2004. RNA interference acts as a
natural innate immune response to O'nyong-nyong virus (Alphavirus;
Togaviridae) infection of Anopheles gambiae mosquitoes. Proceedings of the
National Academy of Sciences of the United States of America, 101 (49),
17240-17245.
Kittayapong, P., Baisley, K.J . and O'Neill, S.L., 1999. A mosquito densovirus
infecting Aedes aegypti and Aedes albopictus from Thailand. American
Journal of Tropical and Medical Hygiene, 61 (4), 612-617.
Kokoza, V., Ahmed, A., Cho, W.L., et al., 2000. Engineering blood meal-activated
systemic immunity in the yellow fever mosquito, Aedes aegypti. Proceedings
of the National Academy of Sciences of the United States of America, 97 (16),
9144-9149.
Kuznetsova, M.A. and Butchasky, L.P., 1988. Effect of the viral drug Viroden on
Aedes aegypti: experimental study. Meditsinskaya Parazitologiya i
Parazitarnye Bolezni, 3, 52-54.
Monath, T.P., 1994. Dengue: the risk to developed and developing countries.
Proceedings of the National Academy of Sciences of the United States of
America, 91 (7), 2395-2400.
Moreira, L.A., Edwards, M.J ., Adhami, F., et al., 2000. Robust gut-specific gene
expression in transgenic Aedes aegypti mosquitoes. Proceedings of the
National Academy of Sciences of the United States of America, 97 (20),
10895-10898.
Muoz, D., J imenez, A., Marinotti, O., et al., 2004. The AeAct-4 gene is expressed in
the developing flight muscles of female Aedes aegypti. Insect Molecular
Biology, 13 (5), 563-568.
Olson, K.E., Adelman, Z.N., Travanty, E.A., et al., 2002. Developing arbovirus
resistance in mosquitoes. Insect Biochemistry and Molecular Biology, 32 (10),
1333-1343.
Reiter, P. and Gubler, D.J ., 1997. Surveillance and control of urban dengue vectors.
In: Gubler, D.J . and Kuno, G. eds. Dengue and dengue hemorrhagic fever.
CAB International, New York, 425-462.
Rigau-Perez, J .G., Clark, G.G., Gubler, D.J ., et al., 1998. Dengue and dengue
haemorrhagic fever. Lancet, 352 (9132), 971-977.
Root, J .J ., Black, W.C.Th., Calisher, C.H., et al., 2003. Analyses of gene flow among
populations of deer mice (Peromyscus maniculatus) at sites near hantavirus
pulmonary syndrome case-patient residences. Journal of Wildlife Diseases, 39
(2), 287-298.
Olson et al.
87
Sanchez-Vargas, I., Travanty, E.A., Keene, K.M., et al., 2004. RNA interference,
arthropod-borne viruses, and mosquitoes. Virus Research, 102 (1), 65-74.
Schwarz, D.S., Hutvagner, G., Haley, B., et al., 2002. Evidence that siRNAs function
as guides, not primers, in the Drosophila and human RNAi pathways.
Molecular Cell, 10 (3), 537-548.
Thomas, D.D., Donnelly, C.A., Wood, R.J ., et al., 2000. Insect population control
using a dominant, repressible, lethal genetic system. Science, 287 (5462),
2474-2476.
Tijsterman, M., Ketting, R.F. and Plasterk, R.H., 2002. The genetics of RNA
silencing. Annual Review of Genetics, 36, 489-519.
Travanty, E.A., Adelman, Z.N., Franz, A.W., et al., 2004. Using RNA interference to
develop dengue virus resistance in genetically modified Aedes aegypti. Insect
Biochemistry and Molecular Biology, 34 (7), 607-613.
Uchil, P.D. and Satchidanandam, V., 2003. Characterization of RNA synthesis,
replication mechanism, and in vitro RNA-dependent RNA polymerase activity
of J apanese encephalitis virus. Virology, 307 (2), 358-371.
Vaucheret, H., Beclin, C., Elmayan, T., et al., 1998. Transgene-induced gene silencing
in plants. Plant Journal, 16 (6), 651-659.