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Review

The Insect Microbiome Modulates Vector Competence for Arboviruses

by
Natapong Jupatanakul
1,
Shuzhen Sim
2 and
George Dimopoulos
1,*
1
Harry Feinstone Department of Molecular Microbiology and Immunology, Bloomberg School of Public Health, Johns Hopkins University, 615 N. Wolfe Street, Baltimore, MD 21205, USA
2
Genome Institute of Singapore, 60 Biopolis Street, #02-01 Genome, Singapore 138672, Singapore
*
Author to whom correspondence should be addressed.
Viruses 2014, 6(11), 4294-4313; https://doi.org/10.3390/v6114294
Submission received: 28 August 2014 / Revised: 31 October 2014 / Accepted: 3 November 2014 / Published: 11 November 2014
(This article belongs to the Special Issue Impact of the Insect Microbiome on Arbovirus Transmission)
Versions Notes

Abstract

:
Diseases caused by arthropod-borne viruses (arboviruses), such as Dengue, West Nile, and Chikungunya, constitute a major global health burden and are increasing in incidence and geographic range. The natural microbiota of insect vectors influences various aspects of host biology, such as nutrition, reproduction, metabolism, and immunity, and recent studies have highlighted the ability of insect-associated bacteria to reduce vector competence for arboviruses and other pathogens. This reduction can occur through mechanisms, such as immune response activation, resource competition, or the production of anti-viral molecules. Studying the interactions between insect vectors and their microbiota is an important step toward developing alternative strategies for arbovirus transmission control.

1. Introduction

Over 130 arthropod-borne viruses (arboviruses) in the families Togaviridae, Flaviviridae, Bunyaviridae, Reoviridae, and Orthomyxoviridae can cause disease in humans [1]. Among these viruses, Dengue virus (DENV), West Nile virus (WNV), and Chikungunya virus (CHIKV) have become major global public health concerns, with increasing incidence in recent decades as a result of the expansion of the vectors’ geographic range, global transport, unplanned urbanization, and climate change [1,2,3,4,5,6].
Arboviruses are maintained in endemic areas by horizontal transmission between vertebrates and blood-feeding insect vectors. While arboviruses can cause serious pathology in humans, they have minimal impact on insect mortality. The insect immune system can control, but not clear, arbovirus infection; for this reason, infected insects can be vectors for life [7].
The replication cycle of arboviruses in insects has been extensively characterized; for example, DENV replication is well characterized in the Aedes aegypti mosquito [7]. After the mosquito ingests an infectious blood meal, the virus has to pass through various infection barriers [8]. It has to infect and replicate in the midgut epithelium (midgut infection barrier), then escape from the midgut to spread throughout the insect body and infect other tissues (midgut escape barrier). In order to transmit the disease, the virus then has to infect and replicate in the salivary glands and disseminate into mosquito saliva (salivary gland infection and escape barriers) [8]. The extrinsic incubation period (EIP), i.e., the time from virus ingestion until its dissemination in mosquito saliva, where it can be transmitted to naïve humans, can vary depending on conditions such as mosquito strain, virus strain, and temperature, but it generally ranges from 7–14 days [7,9,10,11,12,13,14,15,16,17,18,19].
Insects constantly acquire microorganisms such as bacteria and fungi from their natural habitats and may also vertically acquire some species from their parents [20,21,22]. These diverse microbial communities affect multiple aspects of insect biology, such as nutrition, digestion, metabolism, development, and immunity, and, therefore, have great potential to alter vector competence for arboviruses [23,24,25,26,27].
Several studies of the microbiomes of the major mosquito vectors of arboviruses, Ae. aegypti, Ae. albopictus, and Culex quinquefasciatus have been performed along with analyses of anopheline microbiomes.
This review will summarize and discuss recent work on the interactions between the insect gut microbiota, insect host biology, and arboviruses, and how these studies may lead towards the development of alternative methods for arbovirus control.

2. Insect Microbiomes: Source, Dynamics, and Composition

Several studies have characterized the microbiomes of field mosquitoes using either culture-dependent or -independent methods [22,28,29,30,31,32]. The composition of the mosquito microbiome can vary depending on factors such as the species, sex, and life-stage of the mosquito, its geographical origin and feeding behavior, and the organ surveyed [22,28,29,30,31,32]. The relationship between these microbes and insects is complex and can range from pathogenesis to commensalism or mutualism [25,33].

2.1. The Mosquito Microbiome from Larvae to Adult

The mosquito life cycle consists of aquatic larval and pupal stages and a terrestrial adult stage. Because of these completely different habitats, the mosquito microbiome in different developmental stages can be distinct; this is particularly true for the gut. Mosquito gut contents are usually cleared when the insect undergoes metamorphosis and molting during the larvae-to-pupae and pupae-to-adult transitions [34], and the midgut microbiome in adult mosquitoes thus has to be repopulated. In the Anopheles gambiae mosquito, guts of aquatic stages have been found to be predominantly populated by Cyanobacteria, which serve as the larval diet [35,36,37]. On the contrary, adult An. gambiae guts are predominantly populated by Proteobacteria and Bacteroidetes picked up from the environment and ingested food after emergence [37]. There is also evidence that the gut microbiota is important for larval development. A recent study showed that when gut bacteria were depleted, mosquito larvae failed to molt and develop to the next stage [23]. Larval development could be restored by supplementing the breeding water with certain bacteria.
The adult mosquito gut microbiome has been the most extensively studied. While different mosquito species from the same geographical area share several core bacterial taxa, the composition of individual guts is highly variable [29]. The most common bacteria among different mosquito species from Kenya were Gammaproteobacteria (such as Aeromonas), Flavobacteria (such as Chrysobacterium), and Alphaproteobacteria (such as Asaia) [29]. A study of Culex quinquefasciatus, a mosquito vector for WNV, from India identified Proteobacteria (such as Enterobacter, Pseudomonas, Pantoea, and Proteus), Firmicutes (such as Bacillus), and Actinobacteria (such as Acinetobacter) as gut microbiota [38]. Studies in other mosquito species, such as Ae. aegypti and Ae. albopictus, insect vectors for DENV, CHIKV, and yellow fever virus (YFV), also identified Actinobacteria (such as Streptomyces, Microbacterium, and Micrococcus), Firmicutes (such as Bacillus), and Proteobacteria (such as Asaia, Chromobacterium, Enterobacter, Pantoea, Pseudomonas, and Serratia) [22,24,31,32,34,39,40].
The influence of the mosquito’s diet is reflected in the differences between male and female Aedes microbiomes [22,31]. Male mosquitoes acquire soil- and water-associated Actinobacteria through nectar feeding [22]. In female mosquitoes, however, bacteria in the phylum Proteobacteria, especially the family Enterobacteriaceae, which can tolerate redox stress from blood-meal digestion, are the main components of the midgut microbiome [22,37].
Although many studies have treated insects as holobionts, a few have tried to characterize the microbiota associated with individual organs, such as salivary glands, reproductive organs, and hemocoel [20,41,42,43,44]. Some microbes identified in non-gut tissues include intracellular bacteria such as Wolbachia (reproductive organs, salivary glands, head, muscle, and Malpighian tubules) and Spiroplasma (hemolymph, hemocytes, thoracic flight muscle, and nerve cells) [20,40,42]. These studies provide additional insight into how the microbiome can influence mosquito biology and vector competence. For example, Wolbachia in the salivary glands provides Ae. albopictus with resistance to DENV infection [45], and bacteria residing in reproductive organs have the potential for vertical transmission and will facilitate the administration of these microorganisms in the field [46,47,48]. Spiroplasma, a maternally-inherited endosymbiont extensively studied in Drosophila, has been found to cause pathology, influence insect reproduction, and alter the susceptibility of Drosophila to certain pathogens [49,50,51]. Pathogenicity of Spiroplasma in mosquitoes has also been documented [42,52,53,54,55]; however, the role of Spiroplasma in mosquito vector competence for arboviruses is as yet poorly studied.

2.2. Wolbachia and Cytoplasmic Incompatibility (CI)

Bacteria of the genus Wolbachia are maternally inherited, obligate intracellular symbionts that have been estimated to infect 66% of insects [56]. Several arbovirus vectors such as Culex quinquefasciatus and Ae. albopictus are naturally infected with Wolbachia, but not Ae. aegypti [43,57,58,59]. Recent research, however, has shown that stable transinfection of Wolbachia from Drosophila and Ae. albopictus into Ae. aegypti is possible [41,60,61] and, in fact, has great potential as an arboviral control strategy (described below in Section 3).
Wolbachia spreads quickly through populations because of its ability to alter insect reproduction through mechanisms such as feminization, parthenogenesis, and cytoplasmic incompatibility (CI), which increase the reproductive success of infected insects [62]. In mosquitoes, CI ensures that offspring will be infected by Wolbachia because uninfected eggs fertilized with sperm from infected males will not survive [63,64]. This phenomenon, which is maintained in stably trans-infected Ae. aegypti, is useful for the dissemination of Wolbachia in field mosquito populations [47].

2.3. The Insect Eukaryotic Microbiome

In addition to bacterial microbiota, studies have also isolated eukaryotic microorganisms such as fungi and yeast using culture-dependent methods. An early study identified 18 non-pathogenic yeast isolates in the genera Candida, Yarrowia, Rhodotorula, and Cryptococcus from larval and adult stages of Aedes, Culex, and Anopheles mosquitoes [65]. A later study isolated Candida and Pichia yeast from Ae. aegypti midguts [66]. Wickerhamomyces anomalus yeast has also been found in the midgut and reproductive organs of various mosquito species, suggesting a complex eukaryotic microbiome in various tissues [67,68]. These findings are not limited to mosquitoes. For example, 39 fungi were isolated from the cuticle and midgut of five sandfly species, suggesting that eukaryotic microbiota might be common among insects [69].
The eukaryotic microbiota has been much less well-studied than the bacterial microbiota, especially with metagenomic sequencing methods, and further work is required in order for us to fully understand its impact on insect biology and arbovirus transmission.
Paratransgenesis, which involves the genetic modification of insect microbiota to inhibit human pathogens, is considered a promising novel disease control approach [70,71]. The eukaryotic microbiome, especially yeast, have a high potential for paratransgenesis due to their safety, large scale production systems, and available genetic manipulation tools [72,73,74]. Yeasts can be genetically modified to inhibit arboviruses through secretion of antiviral anti-microbial peptides (AMP), such as a cecropin-like peptide possessing anti-DENV and anti-CHIKV activity [75]. Paratransgenesis can also be applied to entomopathogenic fungi, thus maximizing disease control potential through a combination of vector killing and reduction of vector competence. This approach has been employed with the fungus Metarhizium anisopliae, by engineering it to express the SM1 peptide, which inhibits Plasmodium development in Anopheles mosquitoes [76].

3. Microbiota-Driven Mechanisms Affecting Vector Competence

Insect microbiomes have long been co-evolving with their hosts. Early studies of insect symbionts suggested beneficial roles in nutrition; for example, the gut microbiota of termites greatly facilitate cellulose digestion [77,78]. Recent studies in medically important insect vectors also indicate the importance of microbiota for nutrient digestion, metabolism, egg production, development, and immune responses [23,25,26,27,79,80,81,82]. In other cases, endosymbionts, such as Wolbachia and Spiroplasma, may require nutrients from the host for efficient replication [83,84]. These interactions have great potential to influence vector competence for pathogens, since arboviruses require host factors and cellular machinery for their replication and are also controlled by insect immune responses. In addition to these indirect effects, microbiota may also directly interact with arboviruses, since some bacteria species are known to secrete anti-viral compounds [85,86,87,88]. A novel Chromobacterium sp. (Csp_P) species isolated from field-caught Ae. aegypti can reduce mosquito susceptibility to DENV infection when introduced to the mosquito midgut tissue [89]. Certain microbiota can also increase vector competence for arbovirus infection [90,91].

3.1. Immune System Modulation

The insect immune system relies mainly on innate immune responses, which recognize pathogen associated molecular patterns (PAMPs) through pattern recognition receptors (PRRs). Pathogen recognition activates immune signaling pathways such as the Toll pathway, the immune deficiency (IMD) pathway, and the Janus kinase/signal transducers and activators of transcription (JAK-STAT) pathway [92,93,94]. Activation of these immune signaling pathways triggers immune defense mechanisms, such as melanization, encapsulation, phagocytosis, apoptosis, and production of AMPs [95,96,97,98,99,100,101,102].
Each of these immune signaling pathways can be activated by a wide spectrum of microorganisms and viruses. The Toll pathway is activated in response to Gram-positive bacteria, fungi, and DENV [16,103,104,105,106,107,108,109]. The IMD pathway controls immune responses to bacteria, DENV, and the human Plasmodium parasite P. falciparum [16,75,110,111,112,113]. The JAK-STAT pathway is a cytokine-induced signaling pathway that plays important roles in insect anti-viral (DENV and WNV) immunity as well as immune responses to bacteria, fungi, and Plasmodium parasites [16,114,115,116,117,118,119]. Given the overlapping and broad-spectrum nature of immune signaling cascades, microbiota can activate insect immune responses and indirectly affect insect vector competence for arboviruses [39,109,119].
The role of the microbiome on mosquito immunity and vector competence was first characterized in Anopheles mosquitoes and Plasmodium parasites [120]. Transcriptomic comparison between septic and aseptic An. gambiae using microarrays identified a number of immune-related genes up-regulated in the presence of midgut microbiota, which subsequently resulted in lower susceptibility to Plasmodium infection in septic mosquitoes. This study provided a fundamental basis for subsequent studies concerning the effect of the mosquito microbiome on arbovirus infection.
The effect of the microbiome on insect vector competence for arboviruses has been studied in DENV and Ae. aegypti [39,109]. Removal of mosquito gut microbiota by treatment with antibiotics results in higher midgut DENV titers [109], and gene expression analysis has revealed that aseptic Ae. aegypti have lower levels of AMP gene expression (attacin, cecropin, defensin, and gambicin), suggesting a lower level of immune activation [109]. DENV infection of Ae. aegypti salivary glands induces the Toll and IMD pathways and results in the expression of a putative cecropin-like peptide with antibacterial, anti-DENV, and anti-CHIKV activity [75]. Subsequently, the bacterium Proteus sp. (Prsp_P), derived from the gut of field mosquitoes, has been shown to up-regulate AMP gene expression and confer increased resistance to DENV infection of the mosquito gut [39]. These results emphasize the overlap between antibacterial and antiviral insect immune responses.
Wolbachia contributes to Drosophila’s resistance to virus infection, and trans-infection of Wolbachia from Drosophila to Ae. aegypti also increases the mosquitoes’ resistance to DENV, CHIKV, YFV, and Plasmodium infection [57,61,121,122,123]. Introducing Wolbachia into a new insect host can elicit immune responses, as shown in the trans-infection of wMel and wMelPop from Drosophila to Ae. aegypti [124]. It has, however, also been shown that Wolbachia provides protection against DENV infection in Drosophila without activating Drosophila’s immune response [124], suggesting that Wolbachia provides protection against arbovirus infection through both immunity-dependent and -independent mechanisms, depending on the combination of Wolbachia strain and insect host. Wolbachia strain wAlbB from Ae. albopictus has also been trans-infected into Ae. aegypti and shown to contribute to DENV resistance [41,125,126]. Gene expression analysis of Ae. aegypti infected with wAlbB has revealed wAlbB-induced production of reactive oxygen species (ROS), which in turn induce the activation of the Toll pathway [126].
In addition to immune signaling, RNA interference (RNAi) is another major insect anti-viral mechanism. In the canonical exogenous small interfering (siRNA) pathway, viral genomes are recognized and degraded based on sequence complementarity, through the action of Dicer2 (Dcr2) and the RNA-induced silencing complex (RISC) [127,128,129]. The components of the exogenous siRNA pathway are constitutively expressed in the cytoplasm, and there is to date no evidence that this mechanism can be activated by microorganisms other than viruses. However, insects also rely on other small RNA pathways, such as the Piwi-interacting RNA (piRNA) [130] and microRNA (miRNA) pathways [131,132], to restrict arbovirus infection. Recent studies have shown that Wolbachia wMelPop-CLA can alter the mosquito's miRNA profile [133], and can also alter mosquito gene expression through the induction of host microRNAs (miRNAs) [134,135]. The induction of miRNA aae-miR-12 promotes the growth of Wolbachia through a down-regulation of DNA replication licensing factor (MCM6) and the monocarboxylate transporter (MCT1) genes; the increased Wolbachia growth then reduces vector competence for DENV in a density dependent manner [134]. Aae-miR-2940, another miRNA induced by Wolbachia, suppresses Ae. aegypti DNA methyltransferase (AaDnmt2) gene expression. The down-regulation of AaDnmt2 again promotes Wolbachia replication but reduces DENV titers in mosquito cells [135].
The impact of microbiota on immune activity and vector competence in insects has also been documented in insects other than arbovirus vectors. The tsetse fly symbiont, Wigglesworthia glossinidia, activates the IMD pathway and inhibits trypanosome parasite infection [136]. Studies of W. glossinidia in tsetse suggest the importance of the microbiota in the larval stages for immune maturation in adult insects. Wild-type flies that lack W. glossinidia during larval development appear to have compromised immune responses such as AMP expression, prophenol-oxidase activity, melanization, and increased hemocyte number [81,137]. These results emphasize the importance of certain microbiota in particular developmental stages for maturation of the insect immune system in adults; however, this phenomenon is yet to be studied in arbovirus vectors.
In some cases, the insect microbiota does not confer resistance to arbovirus infection but instead increases the insects’ susceptibility to arbovirus infection. For example, trans-infection of wAlbB Wolbachia to Culex tarsalis increases the susceptibility of the mosquitoes to WNV infection [90]. Gene expression analysis has revealed down-regulation of Rel1, a transcription factor responsible for activating Toll pathway-dependent effectors, suggesting that Wolbachia can suppress insect immune responses [90]. The presence of Serratia odorifera in the Ae. aegypti midgut increases the mosquitoes’ susceptibility to DENV infection, possibly through a suppression of immune responses via the binding of prohibitin [91]; however, this possibility has yet to be experimentally confirmed. Other than arboviruses, recent studies have shown that Wolbachia increases mosquito susceptibility to Plasmodium parasite infection [138,139]. One Wolbachia strain can result in either an increase or a decrease of Plasmodium infection, for example, wAlbB reduces P. falciparum infection but increases infection of An. gambiae with P. berghei [139,140]. Environmental factors such as temperature can also affect the outcome of Plasmodium infection when mosquitoes are infected with Wolbachia [141]. For example, Wolbachia wAlbB reduces P. yoelii infection at 28 °C, but increase parasite load at 20 °C. These observations emphasize that the relationship between insect vector, insect microbiota, and human pathogens is far more complex than anticipated, and environmental factor can influence these interaction.

3.2. Resource Competition

Certain combinations of Wolbachia strain-insect species do not result in the elicitation of insect immune responses, as reported in D. simulans and Ae. albopictus [142] or in infections of Drosophila with the wAu and wMel strains [143], suggesting that immune activation is not the only mechanism affecting vector competence. Both arboviruses and insect microbiota, especially endosymbionts, such as Wolbachia and Spiroplasma, require nutrients and host factors for efficient replication. The anti‑viral effect of Wolbachia in an Ae. albopictus cell line has been shown to be density-dependent [144,145], suggesting that high densities of Wolbachia competing for limited resources can affect vector competence.
Lipids, for example, are required by both the microbiota and arboviruses. Several arboviruses such as DENV and WNV use receptor-mediated endocytosis for cell entry in both vertebrate and invertebrate hosts, a process that involves remodeling of lipid membranes [146,147,148,149,150]. After cell entry, viruses modify intracellular compartments of the host to facilitate protein processing and virus replication and assembly [151,152]. DENV influences expression of genes involved in lipid synthesis to alter the host’s lipid composition, lipid homeostasis, and intracellular membrane trafficking [153,154,155]. Wolbachia also uses lipids from host cells for replication and therefore competes with and inhibits DENV and CHIKV replication [61,156].
In the Drosophila and honeybee models, Spiroplasma replication requires lipid and vitamins from its insect host [83,84]. Its requirements in mosquitoes have not been studied, but if similar to those of Drosophila and honeybees, it is plausible that the bacteria may also be able to influence vector competence through Wolbachia-like resource competition mechanisms.

3.3. Secondary Metabolite Production

Actinomycetes, bacteria commonly found in mosquito gut, have long been known to secrete secondary metabolites with anti-bacterial, anti-fungal, and anti-viral activity [88,157]. Another bacterium commonly found in soil and water, Chromobacterium violaceum, has also been studied for its anti-viral activity [86,158]. A recently characterized Chromobacterium sp. (Csp_P) isolated from field mosquito guts, has shown a promising potential as vector-borne disease control tool. Csp_P blocks infection of An. gambiae and Ae. aegypti with Plasmodium and dengue virus, respectively, and exerts entomopathogenic activity against larval and adult stages, likely though the production of secondary metabolites [89]. Bacteria isolated from the Ae. albopictus midgut, such as Pseudomonas rhodesiae, Enterobacter ludwigii, and Vagococcus salmoninarium, have been shown to directly inhibit La Crosse virus independently of the mosquito, suggesting that these bacteria may produce anti-viral molecules [159].
These discoveries suggest that certain species of disease vector’s natural gut microbiome directly influences arbovirus infection through natural products. Isolation of these bacteria and anti-pathogen molecules may open an interesting avenue for the discovery and development of novel therapeutic drugs.

4. Field Applications of Insect Microbiota for Arbovirus Transmission Control

The concept of insect microbiota as an arbovirus control tool has great potential, but it also raises numerous practical and safety concerns. In addition to exploring and characterizing anti-viral mechanisms, studies to address the applicability of these microorganisms to the field should be pursued. For example, the microbial composition of field mosquito guts can be far more complex than in those of mosquitoes in laboratory settings, and this complexity may interfere with the proposed arbovirus transmission-blocking strategy. This complexity was addressed in a recent study of the effect of the mosquito microbiome on the ability of Wolbachia to establish itself in a new insect host [160]. Interactions between the microbiota and Wolbachia inhibited transmission of Wolbachia to the next generation, and also resulted in mosquito mortality.
To date, the most advanced field application of insect microbiota for controlling arbovirus transmission is the Eliminate Dengue program in Australia, which has released Wolbachia-infected Ae. aegypti to control DENV transmission [47,60,161]. In this case, Wolbachia successfully invaded the natural mosquito population, and a follow-up study has found that field-caught Wolbachia-infected Ae. aegypti still maintain their refractoriness to DENV [162]. This program has since been expanded to other countries, including China, Vietnam, Indonesia, Colombia, and Brazil [163].
For gut bacteria which, unlike Wolbachia, are unable to drive themselves into a population, achieving sustained delivery to mosquitoes in nature remains an important and understudied practical issue. Existing measures for mosquito population control such as oviposition traps, spraying of toxins or insect pathogens, and artificial nectar bait [164,165,166,167,168,169] can be adapted as dissemination strategies; however, continued release may be required to maintain these microbiota in the mosquito population.
Paratransgenic approaches, while not explored in the area of arbovirus control, have the potential to reduce arbovirus transmission in a number of ways. Microbiota can be engineered to (1) have enhanced entomopathogenic activity [170,171], (2) secrete anti-pathogen molecules (extensively studied for arthropod-borne parasites) [70,71]), or (3) secrete molecules that activate insect immune responses against the pathogen. Research to identify candidate genes and molecules that increase entomopathogenic activity, inhibit arboviruses, and activate insect immune responses is required to move the field forward.
Collectively, research that will pave the way for the use of insect-derived bacteria as an alternative arbovirus control strategy is still at an early stage. Extensive studies are required to ensure safety and effectiveness prior to a field release of insect microbiota as an arbovirus control strategy.

5. Conclusion

The global burden of arboviral diseases has been rapidly increasing in recent decades. Studies of insect-associated microbial species suggest that they can alter vector competence by modulating host immune responses, competing with arboviruses for resources, and secreting anti-viral factors. Understanding the tripartite relationships between the insect, its microbiome, and the arboviral pathogens it harbors will allow us to develop alternative strategies to reduce the burden of arboviral diseases.

Acknowledgments

This work has been supported by National Institutes of Health/National Institute of Allergy and Infectious Disease grants R21AI090188, R01AI101431 and a fellowship from the Royal Thai Government to NJ.

Conflicts of Interest

The authors declare no conflict of interest.

References and Notes

  1. Cleton, N.; Koopmans, M.; Reimerink, J.; Godeke, G.-J.; Reusken, C. Come fly with me: Review of clinically important arboviruses for global travelers. J. Clin. Virol. 2012, 55, 191–203. [Google Scholar] [CrossRef] [PubMed]
  2. Reiter, P. Yellow fever and dengue: A threat to Europe? Euro. Surveill. 2010, 15, 19509. [Google Scholar]
  3. Soverow, J.E.; Wellenius, G.A.; Fisman, D.N.; Mittleman, M.A. Infectious disease in a warming world: How weather influenced West Nile virus in the United States (2001–2005). Environ. Health Perspect. 2009, 117, 1049–1052. [Google Scholar] [CrossRef] [PubMed]
  4. Weaver, S.C.; Reisen, W.K. Present and future arboviral threats. Antivir. Res. 2010, 85, 328–345. [Google Scholar] [CrossRef] [PubMed]
  5. Weaver, S.C. Urbanization and geographic expansion of zoonotic arboviral diseases: Mechanisms and potential strategies for prevention. Trends Microbiol. 2013, 21, 360–363. [Google Scholar] [CrossRef] [PubMed]
  6. Sutherst, R.W. Global change and human vulnerability to vector-borne diseases. Clin. Microbiol. Rev. 2004, 17, 136–173. [Google Scholar] [CrossRef] [PubMed]
  7. Salazar, M.I.; Richardson, J.H.; Sánchez-Vargas, I.; Olson, K.E.; Beaty, B.J. Dengue virus type 2: Replication and tropisms in orally infected Aedes aegypti mosquitoes. BMC Microbiol 2007, 7, 9. [Google Scholar] [CrossRef] [PubMed]
  8. Black, W.C.; Bennett, K.E.; Gorrochótegui-Escalante, N.; Barillas-Mury, C.V.; Fernández-Salas, I.; de Lourdes Muñoz, M.; Farfán-Alé, J.A.; Olson, K.E.; Beaty, B.J. Flavivirus susceptibility in Aedes aegypti. Arch. Med. Res. 2002, 33, 379–388. [Google Scholar] [CrossRef] [PubMed]
  9. Alto, B.W.; Lounibos, L.P.; Mores, C.N.; Reiskind, M.H. Larval competition alters susceptibility of adult Aedes mosquitoes to dengue infection. Proc. Biol. Sci. 2008, 275, 463–471. [Google Scholar] [CrossRef] [PubMed]
  10. Alto, B.W.; Bettinardi, D. Temperature and dengue virus infection in mosquitoes: Independent effects on the immature and adult stages. Am. J. Trop. Med. Hyg. 2013, 88, 497–505. [Google Scholar] [CrossRef] [PubMed]
  11. Turell, M.J. Effect of environmental temperature on the vector competence of Aedes taeniorhynchus for Rift Valley fever and Venezuelan equine encephalitis viruses. Am. J. Trop. Med. Hyg. 1993, 49, 672–676. [Google Scholar] [PubMed]
  12. Murdock, C.C.; Paaijmans, K.P.; Cox-Foster, D.; Read, A.F.; Thomas, M.B. Rethinking vector immunology: The role of environmental temperature in shaping resistance. Nat. Rev. Microbiol. 2012, 10, 869–876. [Google Scholar] [CrossRef] [PubMed]
  13. Dohm, D.J.; O’Guinn, M.L.; Turell, M.J. Effect of environmental temperature on the ability of Culex pipiens (Diptera: Culicidae) to transmit West Nile virus. J. Med. Entomol. 2002, 39, 221–225. [Google Scholar] [CrossRef] [PubMed]
  14. Reisen, W.K.; Fang, Y.; Martinez, V.M. Effects of temperature on the transmission of west nile virus by Culex tarsalis (Diptera: Culicidae). J. Med. Entomol. 2006, 43, 309–317. [Google Scholar] [CrossRef] [PubMed]
  15. Richards, S.L.; Mores, C.N.; Lord, C.C.; Tabachnick, W.J. Impact of extrinsic incubation temperature and virus exposure on vector competence of Culex pipiens quinquefasciatus say (Diptera: Culicidae) for West Nile virus. Vector-Borne Zoonotic Dis. 2007, 7, 629–636. [Google Scholar] [CrossRef] [PubMed]
  16. Sim, S.; Jupatanakul, N.; Ramirez, J.L.; Kang, S.; Romero-Vivas, C.M.; Mohammed, H.; Dimopoulos, G. Transcriptomic profiling of diverse Aedes aegypti strains reveals increased basal-level immune activation in dengue virus-refractory populations and identifies novel virus-vector molecular interactions. PLoS Neglect. Trop. Dis. 2013, 7, e2295. [Google Scholar] [CrossRef]
  17. Ocampo, C.B.; Caicedo, P.A.; Jaramillo, G.; Ursic Bedoya, R.; Baron, O.; Serrato, I.M.; Cooper, D.M.; Lowenberger, C. Differential expression of apoptosis related genes in selected strains of Aedes aegypti with different susceptibilities to dengue virus. PLoS One 2013, 8, e61187. [Google Scholar] [CrossRef] [PubMed]
  18. Barón, O.L.; Ursic-Bedoya, R.J.; Lowenberger, C.A.; Ocampo, C.B. Differential gene expression from midguts of refractory and susceptible lines of the mosquito, Aedes aegypti, infected with Dengue-2 virus. J. Insect Sci. 2010, 10, 41. [Google Scholar] [CrossRef] [PubMed]
  19. Tjaden, N.B.; Thomas, S.M.; Fischer, D.; Beierkuhnlein, C. Extrinsic incubation period of dengue: Knowledge, backlog, and applications of temperature dependence. PLoS Neglect. Trop. Dis. 2013, 7, e2207. [Google Scholar] [CrossRef]
  20. Werren, J.H. Biology of Wolbachia. Annu. Rev. Entomol. 1997, 42, 587–609. [Google Scholar] [CrossRef] [PubMed]
  21. Colman, D.R.; Toolson, E.C.; Takacs-Vesbach, C.D. Do diet and taxonomy influence insect gut bacterial communities? Mol. Ecol. 2012, 21, 5124–5137. [Google Scholar]
  22. Valiente Moro, C.; Tran, F.H.; Raharimalala, F.N.; Ravelonandro, P.; Mavingui, P. Diversity of culturable bacteria including Pantoea in wild mosquito Aedes albopictus. BMC Microbiol. 2013, 13, 70. [Google Scholar] [CrossRef] [PubMed]
  23. Coon, K.L.; Vogel, K.J.; Brown, M.R.; Strand, M.R. Mosquitoes rely on their gut microbiota for development. Mol. Ecol. 2014, 23, 2727–2739. [Google Scholar] [CrossRef] [PubMed]
  24. Minard, G.; Mavingui, P.; Moro, C.V. Diversity and function of bacterial microbiota in the mosquito holobiont. Parasit. Vectors 2013, 6, 146. [Google Scholar] [CrossRef] [PubMed]
  25. Dillon, R.J.; Dillon, V.M. The gut bacteria of insects: Nonpathogenic interactions. Annu. Rev. Entomol. 2004, 49, 71–92. [Google Scholar] [CrossRef] [PubMed]
  26. Charroux, B.; Royet, J. Drosophila immune response: From systemic antimicrobial peptide production in fat body cells to local defense in the intestinal tract. Fly (Austin) 2010, 4, 40–47. [Google Scholar] [CrossRef]
  27. Ryu, J.H.; Kim, S.H.; Lee, H.Y.; Bai, J.Y.; Nam, Y.D.; Bae, J.W.; Lee, D.G.; Shin, S.C.; Ha, E.M.; Lee, W.J. Innate immune homeostasis by the homeobox gene caudal and commensal-gut mutualism in Drosophila. Science 2008, 319, 777–782. [Google Scholar] [CrossRef] [PubMed]
  28. Boissière, A.; Tchioffo, M.T.; Bachar, D.; Abate, L.; Marie, A.; Nsango, S.E.; Shahbazkia, H.R.; Awono-Ambene, P.H.; Levashina, E.A.; Christen, R.; et al. Midgut microbiota of the malaria mosquito vector Anopheles gambiae and interactions with Plasmodium falciparum infection. PLoS Pathog. 2012, 8, e1002742. [Google Scholar] [CrossRef] [PubMed]
  29. Osei-Poku, J.; Mbogo, C.M.; Palmer, W.J.; Jiggins, F.M. Deep sequencing reveals extensive variation in the gut microbiota of wild mosquitoes from Kenya. Mol. Ecol. 2012, 21, 5138–5150. [Google Scholar] [CrossRef] [PubMed]
  30. Zouache, K.; Raharimalala, F.N.; Raquin, V.; Tran-Van, V.; Raveloson, L.H.R.; Ravelonandro, P.; Mavingui, P. Bacterial diversity of field-caught mosquitoes, Aedes albopictus and Aedes aegypti, from different geographic regions of Madagascar. FEMS Microbiol. Ecol. 2011, 75, 377–389. [Google Scholar] [CrossRef] [PubMed]
  31. Terenius, O.; Lindh, J.M.; Eriksson-Gonzales, K.; Bussière, L.; Laugen, A.T.; Bergquist, H.; Titanji, K.; Faye, I. Midgut bacterial dynamics in Aedes aegypti. FEMS Microbiol. Ecol. 2012, 80, 556–565. [Google Scholar] [CrossRef] [PubMed]
  32. Chouaia, B.; Rossi, P.; Montagna, M.; Ricci, I.; Crotti, E.; Damiani, C.; Epis, S.; Faye, I.; Sagnon, N.; Alma, A.; et al. Molecular evidence for multiple infections as revealed by typing of Asaia bacterial symbionts of four mosquito species. Appl. Environ. Microbiol. 2010, 76, 7444–7450. [Google Scholar] [CrossRef] [PubMed]
  33. Dharne, M.; Patole, M.; Shouche, Y.S. Microbiology of the insect gut: Tales from mosquitoes and bees. J. Biosci. 2006, 31, 293–295. [Google Scholar] [CrossRef] [PubMed]
  34. Moll, R.M.; Romoser, W.S.; Modrzakowski, M.C.; Moncayo, A.C.; Lerdthusnee, K. Meconial peritrophic membranes and the fate of midgut bacteria during mosquito (Diptera: Culicidae) metamorphosis. J. Med. Entomol. 2001, 38, 29–32. [Google Scholar] [CrossRef] [PubMed]
  35. Damiani, C.; Ricci, I.; Crotti, E.; Rossi, P.; Rizzi, A.; Scuppa, P.; Capone, A.; Ulissi, U.; Epis, S.; Genchi, M.; et al. Mosquito-bacteria symbiosis: the case of Anopheles gambiae and Asaia. Microb. Ecol. 2010, 60, 644–654. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  36. Thiery, I.; Nicolas, L.; Rippka, R.; Tandeau de Marsac, N. Selection of cyanobacteria isolated from mosquito breeding sites as a potential food source for mosquito larvae. Appl. Environ. Microbiol. 1991, 57, 1354–1359. [Google Scholar] [PubMed]
  37. Wang, Y.; Gilbreath, T.M.; Kukutla, P.; Yan, G.; Xu, J. Dynamic gut microbiome across life history of the malaria mosquito Anopheles gambiae in Kenya. PLoS One 2011, 6, e24767. [Google Scholar] [CrossRef] [PubMed]
  38. Chandel, K.; Mendki, M.J.; Parikh, R.Y.; Kulkarni, G.; Tikar, S.N.; Sukumaran, D.; Prakash, S.; Parashar, B.D.; Shouche, Y.S.; Veer, V. Midgut microbial community of Culex quinquefasciatus mosquito populations from India. PLoS One 2013, 8, e80453. [Google Scholar] [CrossRef] [PubMed]
  39. Ramirez, J.L.; Souza-Neto, J.; Torres Cosme, R.; Rovira, J.; Ortiz, A.; Pascale, J.M.; Dimopoulos, G. Reciprocal tripartite interactions between the Aedes aegypti midgut microbiota, innate immune system and dengue virus influences vector competence. PLoS Neglect. Trop. Dis. 2012, 6, e1561. [Google Scholar] [CrossRef]
  40. Minard, G.; Tran, F.H.; Dubost, A.; Tran-Van, V.; Mavingui, P.; Moro, C.V. Pyrosequencing 16S rRNA genes of bacteria associated with wild tiger mosquito Aedes albopictus: A pilot study. Front. Cell. Infect. Microbiol. 2014, 4, 59. [Google Scholar] [PubMed]
  41. Bian, G.; Xu, Y.; Lu, P.; Xie, Y.; Xi, Z. The endosymbiotic bacterium Wolbachia induces resistance to dengue virus in Aedes aegypti. PLoS Pathog. 2010, 6, e1000833. [Google Scholar] [CrossRef] [PubMed]
  42. Abalain-Colloc, M.L.; Rosen, L.; Tully, J.G.; Bove, J.M.; Chastel, C.; Williamson, D.L. Spiroplasma taiwanense sp. nov. from Culex tritaeniorhynchus mosquitoes collected in Taiwan. Int. J. Syst. Bacteriol. 1988, 38, 103–107. [Google Scholar] [CrossRef]
  43. Ricci, I.; Cancrini, G.; Gabrielli, S.; D'Amelio, S.; Favi, G. Searching for Wolbachia (Rickettsiales: Rickettsiaceae) in mosquitoes (Diptera: Culicidae): Large polymerase chain reaction survey and new identifications. J. Med. Entomol. 2002, 39, 562–567. [Google Scholar] [CrossRef] [PubMed]
  44. Eleftherianos, I.; Atri, J.; Accetta, J.; Castillo, J.C. Endosymbiotic bacteria in insects: Guardians of the immune system? Front. Physiol. 2013, 4, 46. [Google Scholar]
  45. Mousson, L.; Zouache, K.; Arias-Goeta, C.; Raquin, V.; Mavingui, P.; Failloux, A.-B. The native Wolbachia symbionts limit transmission of dengue virus in Aedes albopictus. PLoS Neglect. Trop. Dis. 2012, 6, e1989. [Google Scholar] [CrossRef]
  46. Herren, J.K.; Paredes, J.C.; Schüpfer, F.; Lemaitre, B. Vertical transmission of a Drosophila endosymbiont via cooption of the yolk transport and internalization machinery. MBio 2013, 4. [Google Scholar] [CrossRef]
  47. Hoffmann, A.A.; Montgomery, B.L.; Popovici, J.; Iturbe-Ormaetxe, I.; Johnson, P.H.; Muzzi, F.; Greenfield, M.; Durkan, M.; Leong, Y.S.; Dong, Y.; et al. Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature 2011, 476, 454–457. [Google Scholar] [CrossRef] [PubMed]
  48. Ricci, I.; Damiani, C.; Rossi, P.; Capone, A.; Scuppa, P.; Cappelli, A.; Ulissi, U.; Mosca, M.; Valzano, M.; Epis, S. Mosquito symbioses: From basic research to the paratransgenic control of mosquito-borne diseases. J. Appl. Entomol. 2011, 135, 487–493. [Google Scholar] [CrossRef]
  49. Herren, J.K.; Lemaitre, B. Spiroplasma and host immunity: Activation of humoral immune responses increases endosymbiont load and susceptibility to certain Gram-negative bacterial pathogens in Drosophila melanogaster. Cell. Microbiol. 2011, 13, 1385–1396. [Google Scholar] [CrossRef] [PubMed]
  50. Haselkorn, T.S.; Watts, T.D.; Markow, T.A. Density dynamics of diverse Spiroplasma strains naturally infecting different species of Drosophila. Fly (Austin) 2013, 7, 204–210. [Google Scholar] [CrossRef]
  51. Anbutsu, H.; Fukatsu, T. Evasion, suppression and tolerance of Drosophila innate immunity by a male-killing Spiroplasma endosymbiont. Insect Mol. Biol. 2010, 19, 481–488. [Google Scholar] [PubMed]
  52. Williamson, D.L.; Tully, J.G.; Rosen, L.; Rose, D.L.; Whitcomb, R.F.; Abalain-Colloc, M.L.; Carle, P.; Bove, J.M.; Smyth, J. Spiroplasma diminutum sp. nov., from Culex annulus mosquitoes collected in Taiwan. Int. J. Syst. Bacteriol. 1996, 46, 229–233. [Google Scholar] [CrossRef] [PubMed]
  53. Humphery Smith, I.; Grulet, O.; Le Goff, F.; Chastel, C. Spiroplasma (Mollicutes: Spiroplasmataceae) pathogenic for Aedes aegypti and Anopheles stephensi (Diptera: Culicidae). J. Med. Entomol. 1991, 28, 219–222. [Google Scholar] [PubMed]
  54. Humphery Smith, I.; Grulet, O.; Chastel, C. Pathogenicity of Spiroplasma taiwanense for larval Aedes aegypti mosquitoes. Med. Vet. Entomol. 1991, 5, 229–232. [Google Scholar] [CrossRef] [PubMed]
  55. Humphery Smith, I.; Goff, F.L.; Robaux, P.; Chastel, C. Biosafety of an Experimentally Proven Mosquito Vector Pathogen, Spiroplasma taiwanense. Biocontr. Sci. Technol. 1993, 3, 73–78. [Google Scholar] [CrossRef]
  56. Hilgenboecker, K.; Hammerstein, P.; Schlattmann, P.; Telschow, A.; Werren, J.H. How many species are infected with Wolbachia?-A statistical analysis of current data. FEMS Microbiol. Lett. 2008, 281, 215–220. [Google Scholar] [CrossRef] [PubMed]
  57. Iturbe-Ormaetxe, I.; Walker, T.; O' Neill, S.L. Wolbachia and the biological control of mosquito-borne disease. EMBO Rep. 2011, 12, 508–518. [Google Scholar] [CrossRef] [PubMed]
  58. Kittayapong, P.; Baisley, K.J.; Baimai, V.; O'Neill, S.L. Distribution and diversity of Wolbachia infections in Southeast Asian mosquitoes (Diptera: Culicidae). J. Med. Entomol. 2000, 37, 340–345. [Google Scholar] [CrossRef] [PubMed]
  59. Sinkins, S.P.; Braig, H.R.; O'Neill, S.L. Wolbachia pipientis: bacterial density and unidirectional cytoplasmic incompatibility between infected populations of Aedes albopictus. Exp. Parasitol. 1995, 81, 284–291. [Google Scholar] [CrossRef] [PubMed]
  60. Walker, T.; Johnson, P.H.; Moreira, L.A.; Iturbe-Ormaetxe, I.; Frentiu, F.D.; McMeniman, C.J.; Leong, Y.S.; Dong, Y.; Axford, J.; Kriesner, P.; et al. The wMel Wolbachia strain blocks dengue and invades caged Aedes aegypti populations. Nature 2011, 476, 450–453. [Google Scholar] [CrossRef] [PubMed]
  61. Moreira, L.A.; Iturbe-Ormaetxe, I.; Jeffery, J.A.; Lu, G.; Pyke, A.T.; Hedges, L.M.; Rocha, B.C.; Hall-Mendelin, S.; Day, A.; Riegler, M.; et al. A Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya, and Plasmodium. Cell 2009, 139, 1268–1278. [Google Scholar] [CrossRef] [PubMed]
  62. Ma, W.-J.; Vavre, F.; Beukeboom, L.W. Manipulation of arthropod sex determination by endosymbionts: diversity and molecular mechanisms. Sex Dev. 2014, 8, 59–73. [Google Scholar] [CrossRef] [PubMed]
  63. Yen, J.H.; Barr, A.R. New hypothesis of the cause of cytoplasmic incompatibility in Culex pipiens L. Nature 1971, 232, 657–658. [Google Scholar] [CrossRef] [PubMed]
  64. Dobson, S.L.; Fox, C.W.; Jiggins, F.M. The effect of Wolbachia-induced cytoplasmic incompatibility on host population size in natural and manipulated systems. Proc. Biol. Sci. 2002, 269, 437–445. [Google Scholar] [CrossRef] [PubMed]
  65. Ignatova, E.A.; Nagornaia, S.S.; Povazhnaia, T.N.; Ianishevskaia, G.S. The yeast flora of blood-sucking mosquitoes. Mikrobiol. Z. 1996, 58, 12–15. [Google Scholar] [PubMed]
  66. Gusmão, D.S.; Santos, A.V.; Marini, D.C.; Bacci, M.; Berbert-Molina, M.A.; Lemos, F.J. A. Culture-dependent and culture-independent characterization of microorganisms associated with Aedes aegypti (Diptera: Culicidae) (L.) and dynamics of bacterial colonization in the midgut. Acta Trop. 2010, 115, 275–281. [Google Scholar] [CrossRef] [PubMed]
  67. Ricci, I.; Mosca, M.; Valzano, M.; Damiani, C.; Scuppa, P.; Rossi, P.; Crotti, E.; Cappelli, A.; Ulissi, U.; Capone, A.; et al. Different mosquito species host Wickerhamomyces anomalus (Pichia anomala): perspectives on vector-borne diseases symbiotic control. Antonie van Leeuwenhoek 2011, 99, 43–50. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  68. Ricci, I.; Damiani, C.; Scuppa, P.; Mosca, M.; Crotti, E.; Rossi, P.; Rizzi, A.; Capone, A.; Gonella, E.; Ballarini, P.; et al. The yeast Wickerhamomyces anomalus (Pichia anomala) inhabits the midgut and reproductive system of the Asian malaria vector Anopheles stephensi. Environ. Microbiol. 2011, 13, 911–921. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  69. Akhoundi, M.; Bakhtiari, R.; Guillard, T.; Baghaei, A.; Tolouei, R.; Sereno, D.; Toubas, D.; Depaquit, J.; Abyaneh, M.R. Diversity of the bacterial and fungal microflora from the midgut and cuticle of Phlebotomine sand flies collected in North-Western Iran. PLoS One 2012, 7, e50259. [Google Scholar] [CrossRef] [PubMed]
  70. Wang, S.; Ghosh, A.K.; Bongio, N.; Stebbings, K.A.; Lampe, D.J.; Jacobs-Lorena, M. Fighting malaria with engineered symbiotic bacteria from vector mosquitoes. Proc. Natl. Acad. Sci. USA 2012, 109, 12734–12739. [Google Scholar] [CrossRef] [PubMed]
  71. Hurwitz, I.; Fieck, A.; Read, A.; Hillesland, H.; Klein, N.; Kang, A.; Durvasula, R. Paratransgenic control of vector borne diseases. Int. J. Biol. Sci. 2011, 7, 1334–1344. [Google Scholar] [CrossRef] [PubMed]
  72. Macauley-Patrick, S.; Fazenda, M.L.; McNeil, B.; Harvey, L.M. Heterologous protein production using the Pichia pastoris expression system. Yeast 2005, 22, 249–270. [Google Scholar] [CrossRef] [PubMed]
  73. Nicaud, J.-M.; Madzak, C.; Broek, P.; Gysler, C.; Duboc, P.; Niederberger, P.; Gaillardin, C. Protein expression and secretion in the yeast Yarrowia lipolytica. FEMS Yeast Res. 2002, 2, 371–379. [Google Scholar] [PubMed]
  74. Madzak, C.; Gaillardin, C.; Beckerich, J.-M. Heterologous protein expression and secretion in the non-conventional yeast Yarrowia lipolytica: A review. J. Biotechnol. 2004, 109, 63–81. [Google Scholar] [CrossRef] [PubMed]
  75. Luplertlop, N.; Surasombatpattana, P.; Patramool, S.; Dumas, E.; Wasinpiyamongkol, L.; Saune, L.; Hamel, R.; Bernard, E.; Sereno, D.; Thomas, F.; et al. Induction of a peptide with activity against a broad spectrum of pathogens in the Aedes aegypti salivary gland, following infection with dengue virus. PLoS Pathog. 2011, 7, e1001252. [Google Scholar] [CrossRef] [PubMed]
  76. Fang, W.; Vega-Rodriguez, J.; Ghosh, A.K.; Jacobs-Lorena, M.; Kang, A.; St Leger, R.J. Development of transgenic fungi that kill human malaria parasites in mosquitoes. Science 2011, 331, 1074–1077. [Google Scholar] [CrossRef] [PubMed]
  77. Ohkuma, M. Termite symbiotic systems: Efficient bio-recycling of lignocellulose. Appl. Microbiol. Biotechnol. 2003, 61, 1–9. [Google Scholar] [CrossRef] [PubMed]
  78. Brune, A. Termite guts: the world's smallest bioreactors. Trends Biotechnol. 1998, 16, 16–21. [Google Scholar] [CrossRef]
  79. De Gaio, A.O.; Gusmão, D.S.; Santos, A.V.; Berbert-Molina, M.A.; Pimenta, P.F.; Lemos, F.J. Contribution of midgut bacteria to blood digestion and egg production in Aedes aegypti (Diptera: Culicidae) (L.). Parasites Vectors 2011, 4, 105. [Google Scholar] [CrossRef] [PubMed]
  80. Lee, W.J.; Brey, P.T. How microbiomes influence metazoan development: Insights from history and Drosophila modeling of gut-microbe interactions. Annu. Rev. Cell Dev. Biol. 2013, 29, 571–592. [Google Scholar] [CrossRef] [PubMed]
  81. Weiss, B.L.; Wang, J.; Aksoy, S. Tsetse immune system maturation requires the presence of obligate symbionts in larvae. PLoS Biol. 2011, 9, e1000619. [Google Scholar] [CrossRef]
  82. Dale, C.; Welburn, S.C. The endosymbionts of tsetse flies: Manipulating host-parasite interactions. Int. J. Parasitol. 2001, 31, 628–631. [Google Scholar] [CrossRef] [PubMed]
  83. Chang, C.J. Vitamin requirements of three Spiroplasmas. J. Bacteriol. 1984, 160, 488–490. [Google Scholar] [PubMed]
  84. Herren, J.K.; Paredes, J.C.; Schupfer, F.; Arafah, K.; Bulet, P.; Lemaitre, B. Insect endosymbiont proliferation is limited by lipid availability. eLife 2014, e02964. [Google Scholar]
  85. Běhal, V. Nontraditional microbial bioactive metabolites. Folia Microbiol. 2001, 46, 363–370. [Google Scholar] [CrossRef]
  86. Andrighetti-Fröhner, C.R.; Antonio, R.V.; Creczynski-Pasa, T.B.; Barardi, C.R.M.; Simões, C.M.O. Cytotoxicity and potential antiviral evaluation of violacein produced by Chromobacterium violaceum. Mem. Inst. Oswaldo Cruz 2003, 98, 843–848. [Google Scholar] [CrossRef] [PubMed]
  87. Habib, E.S.; Yokomizo, K.; Nagao, K.; Harada, S.; Uyeda, M. Antiviral activity of fattiviracin FV-8 against human immunodeficiency virus type 1 (HIV-1). Biosci. Biotechnol. Biochem. 2001, 65, 683–685. [Google Scholar] [CrossRef] [PubMed]
  88. Chaudhary, H.S.; Soni, B.; Shrivastava, A.R.; Shrivastava, S. Diversity and versatility of Actinomycetes and its role in antibiotic production. J. Appl. Pharm. Sci. Vol. 2013, 3, S83–S94. [Google Scholar]
  89. Ramirez, J.L.; Short, S.M.; Bahia, A.C.; Saraiva, R.G.; Dong, Y.; Kang, S.; Tropathi, A.; Mlambo, G.; Dimopoulos, G. Chromobacterium Csp_P reduces malaria and dengue infection in vector mosquitoes and has entomopathogenic and in vitro anti-pathogen activities. PLoS Pathog. 2014, e1004398. [Google Scholar] [CrossRef]
  90. Dodson, B.L.; Hughes, G.L.; Paul, O.; Matacchiero, A.C.; Kramer, L.D.; Rasgon, J.L. Wolbachia enhances west nile virus (WNV) infection in the mosquito Culex tarsalis. PLoS Neglect. Trop. Dis. 2014, 8, e2965. [Google Scholar] [CrossRef]
  91. Apte-Deshpande, A.; Paingankar, M.; Gokhale, M.D.; Deobagkar, D.N. Serratia odorifera a midgut inhabitant of Aedes aegypti mosquito enhances its susceptibility to dengue-2 virus. PLoS One 2012, 7, e40401. [Google Scholar] [CrossRef] [PubMed]
  92. Levashina, E.A. Immune responses in Anopheles gambiae. Insect Biochem. Mol. Biol. 2004, 34, 673–678. [Google Scholar] [CrossRef] [PubMed]
  93. Fragkoudis, R.; Attarzadeh-Yazdi, G.; Nash, A.A.; Fazakerley, J.K.; Kohl, A. Advances in dissecting mosquito innate immune responses to arbovirus infection. J. Gen. Virol. 2009, 90, 2061–2072. [Google Scholar] [CrossRef] [PubMed]
  94. Kingsolver, M.B.; Huang, Z.; Hardy, R.W. Insect antiviral innate immunity: Pathways, effectors, and connections. J. Mol. Biol. 2013, 425, 4921–4936. [Google Scholar] [CrossRef] [PubMed]
  95. Dimopoulos, G. Insect immunity and its implication in mosquito-malaria interactions. Cell. Microbiol. 2003, 5, 3–14. [Google Scholar] [CrossRef] [PubMed]
  96. Lavine, M.D.; Strand, M.R. Insect hemocytes and their role in immunity. Insect Biochem. Mol. Biol. 2002, 32, 1295–1309. [Google Scholar] [CrossRef] [PubMed]
  97. Blandin, S.; Shiao, S.-H.; Moita, L.F.; Janse, C.J.; Waters, A.P.; Kafatos, F.C.; Levashina, E.A. Complement-like protein TEP1 is a determinant of vectorial capacity in the malaria vector Anopheles gambiae. Cell 2004, 116, 661–670. [Google Scholar] [CrossRef] [PubMed]
  98. Cerenius, L.; Lee, B.L.; Söderhäll, K. The proPO-system: Pros and cons for its role in invertebrate immunity. Trends Immunol. 2008, 29, 263–271. [Google Scholar] [CrossRef] [PubMed]
  99. Cerenius, L.; Söderhäll, K. The prophenoloxidase-activating system in invertebrates. Immunol. Rev. 2004, 198, 116–126. [Google Scholar] [CrossRef] [PubMed]
  100. Dimarcq, J.L.; Hoffmann, D.; Meister, M.; Bulet, P.; Lanot, R.; Reichhart, J.M.; Hoffmann, J.A. Characterization and transcriptional profiles of a Drosophila gene encoding an insect defensin. Eur. J. Biochem. 1994, 221, 201–209. [Google Scholar] [CrossRef] [PubMed]
  101. Blandin, S.A.; Levashina, E.A. Phagocytosis in mosquito immune responses. Immunol. Rev. 2007, 219, 8–16. [Google Scholar] [CrossRef] [PubMed]
  102. Irving, P.; Ubeda, J.-M.; Doucet, D.; Troxler, L.; Lagueux, M.; Zachary, D.; Hoffmann, J.A.; Hetru, C.; Meister, M. New insights into Drosophila larval haemocyte functions through genome-wide analysis. Cell. Microbiol. 2005, 7, 335–350. [Google Scholar] [CrossRef] [PubMed]
  103. Belvin, M.P.; Anderson, K.V. A conserved signaling pathway: The Drosophila toll-dorsal pathway. Annu. Rev. Cell Dev. Biol. 1996, 12, 393–416. [Google Scholar] [CrossRef] [PubMed]
  104. Kim, T.; Kim, Y.-J. Overview of innate immunity in Drosophila. J. Biochem. Mol. Biol. 2005, 38, 121–127. [Google Scholar] [CrossRef] [PubMed]
  105. Lemaitre, B.; Nicolas, E.; Michaut, L.; Reichhart, J.M.; Hoffmann, J.A. The dorsoventral regulatory gene cassette spätzle/Toll/cactus controls the potent antifungal response in Drosophila adults. Cell 1996, 86, 973–983. [Google Scholar] [CrossRef] [PubMed]
  106. Rutschmann, S.; Kilinc, A.; Ferrandon, D. Cutting edge: The toll pathway is required for resistance to gram-positive bacterial infections in Drosophila. J. Immunol. 2002, 168, 1542–1546. [Google Scholar] [CrossRef] [PubMed]
  107. Valanne, S.; Wang, J.-H.; Rämet, M. The Drosophila Toll signaling pathway. J. Immunol. 2011, 186, 649–656. [Google Scholar] [CrossRef] [PubMed]
  108. Zambon, R.A.; Nandakumar, M.; Vakharia, V.N.; Wu, L.P. The Toll pathway is important for an antiviral response in Drosophila. Proc. Natl. Acad. Sci. USA 2005, 102, 7257–7262. [Google Scholar] [CrossRef] [PubMed]
  109. Xi, Z.; Ramirez, J.L.; Dimopoulos, G. The Aedes aegypti toll pathway controls dengue virus infection. PLoS Pathog. 2008, 4, e1000098. [Google Scholar] [CrossRef] [PubMed]
  110. Kaneko, T.; Silverman, N. Bacterial recognition and signalling by the Drosophila IMD pathway. Cell. Microbiol. 2005, 7, 461–469. [Google Scholar] [CrossRef] [PubMed]
  111. Choe, K.-M.; Werner, T.; Stöven, S.; Hultmark, D.; Anderson, K.V. Requirement for a peptidoglycan recognition protein (PGRP) in Relish activation and antibacterial immune responses in Drosophila. Science 2002, 296, 359–362. [Google Scholar] [CrossRef] [PubMed]
  112. Avadhanula, V.; Weasner, B.P.; Hardy, G.G.; Kumar, J.P.; Hardy, R.W. A novel system for the launch of alphavirus RNA synthesis reveals a role for the Imd pathway in arthropod antiviral response. PLoS Pathog. 2009, 5, e1000582. [Google Scholar] [CrossRef] [PubMed]
  113. Garver, L.S.; Bahia, A.C.; Das, S.; Souza-Neto, J.A.; Shiao, J.; Dong, Y.; Dimopoulos, G. Anopheles Imd pathway factors and effectors in infection intensity-dependent anti-Plasmodium action. PLoS Pathog. 2012, 8, e1002737. [Google Scholar] [CrossRef] [PubMed]
  114. Paradkar, P.N.; Trinidad, L.; Voysey, R.; Duchemin, J.-B.; Walker, P.J. Secreted Vago restricts West Nile virus infection in Culex mosquito cells by activating the Jak-STAT pathway. Proc. Natl. Acad. Sci. USA 2012, 109, 18915–18920. [Google Scholar] [CrossRef] [PubMed]
  115. Bahia, A.C.; Kubota, M.S.; Tempone, A.J.; Araújo, H.R.C.; Guedes, B.A.M.; Orfanó, A.S.; Tadei, W.P.; Ríos-Velásquez, C.M.; Han, Y.S.; Secundino, N.F.C.; et al. The JAK-STAT pathway controls Plasmodium vivax load in early stages of Anopheles aquasalis infection. PLoS Neglect. Trop. Dis. 2011, 5, e1317. [Google Scholar] [CrossRef]
  116. Souza-Neto, J.A.; Sim, S.; Dimopoulos, G. An evolutionary conserved function of the JAK-STAT pathway in anti-dengue defense. Proc. Natl. Acad. Sci. USA 2009, 106, 17841–17846. [Google Scholar] [CrossRef] [PubMed]
  117. Wang, L.; Ligoxygakis, P. Pathogen recognition and signalling in the Drosophila innate immune response. Immunobiology 2006, 211, 251–261. [Google Scholar] [CrossRef] [PubMed]
  118. Bina, S.; Zeidler, M. JAK/STAT Pathway Signalling in Drosophila Melanogaster; Landes Bioscience: Austin, TX, USA, 2009; pp. 24–42. [Google Scholar]
  119. Dong, Y.; Morton, J.C.; Ramirez, J.L.; Souza-Neto, J.A.; Dimopoulos, G. The entomopathogenic fungus Beauveria bassiana activate toll and JAK-STAT pathway-controlled effector genes and anti-dengue activity in Aedes aegypti. Insect Biochem. Mol. Biol. 2012, 42, 126–132. [Google Scholar] [CrossRef] [PubMed]
  120. Dong, Y.; Manfredini, F.; Dimopoulos, G. Implication of the mosquito midgut microbiota in the defense against malaria parasites. PLoS Pathog. 2009, 5, e1000423. [Google Scholar] [CrossRef] [PubMed]
  121. Van den Hurk, A.F.; Hall-Mendelin, S.; Pyke, A.T.; Frentiu, F.D.; McElroy, K.; Day, A.; Higgs, S.; O'Neill, S.L. Impact of Wolbachia on infection with chikungunya and yellow fever viruses in the mosquito vector Aedes aegypti. PLoS Neglect. Trop. Dis. 2012, 6, e1892. [Google Scholar] [CrossRef] [Green Version]
  122. Kambris, Z.; Blagborough, A.M.; Pinto, S.B.; Blagrove, M.S.C.; Godfray, H.C.J.; Sinden, R.E.; Sinkins, S.P. Wolbachia stimulates immune gene expression and inhibits Plasmodium development in Anopheles gambiae. PLoS Pathog. 2010, 6, e1001143. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  123. Cirimotich, C.M.; Ramirez, J.L.; Dimopoulos, G. Native microbiota shape insect vector competence for human pathogens. Cell Host Microbe 2011, 10, 307–310. [Google Scholar] [CrossRef] [PubMed]
  124. Rancès, E.; Ye, Y.H.; Woolfit, M.; McGraw, E.A.; O'Neill, S.L. The relative importance of innate immune priming in Wolbachia-mediated dengue interference. PLoS Pathog. 2012, 8, e1002548. [Google Scholar] [CrossRef] [PubMed]
  125. Xi, Z.; Khoo, C.C.H.; Dobson, S.L. Wolbachia establishment and invasion in an Aedes aegypti laboratory population. Science 2005, 310, 326–328. [Google Scholar] [CrossRef] [PubMed]
  126. Pan, X.; Zhou, G.; Wu, J.; Bian, G.; Lu, P.; Raikhel, A.S.; Xi, Z. Wolbachia induces reactive oxygen species (ROS)-dependent activation of the Toll pathway to control dengue virus in the mosquito Aedes aegypti. Proc. Natl. Acad. Sci. USA 2012, 109, E23–E31. [Google Scholar] [CrossRef] [PubMed]
  127. McFarlane, M.; Arias-Goeta, C.; Martin, E.; O'Hara, Z.; Lulla, A.; Mousson, L.; Rainey, S.M.; Misbah, S.; Schnettler, E.; Donald, C.L.; et al. Characterization of Aedes aegypti Innate-Immune Pathways that Limit Chikungunya Virus Replication. PLoS Neglect. Trop. Dis. 2014, 8, e2994. [Google Scholar] [CrossRef]
  128. Blair, C.D. Mosquito RNAi is the major innate immune pathway controlling arbovirus infection and transmission. Future Microbiol. 2011, 6, 265–277. [Google Scholar] [CrossRef] [PubMed]
  129. Franz, A.W.E.; Sánchez-Vargas, I.; Adelman, Z.N.; Blair, C.D.; Beaty, B.J.; James, A.A.; Olson, K.E. Engineering RNA interference-based resistance to dengue virus type 2 in genetically modified Aedes aegypti. Proc. Natl. Acad. Sci. USA 2006, 103, 4198–4203. [Google Scholar] [CrossRef] [PubMed]
  130. Hess, A.M.; Prasad, A.N.; Ptitsyn, A.; Ebel, G.D.; Olson, K.E.; Barbacioru, C.; Monighetti, C.; Campbell, C.L. Small RNA profiling of Dengue virus-mosquito interactions implicates the PIWI RNA pathway in anti-viral defense. BMC Microbiol. 2011, 11, 45. [Google Scholar] [CrossRef] [PubMed]
  131. Yan, H.; Zhou, Y.; Liu, Y.; Deng, Y.; Puthiyakunnon, S.; Chen, X. miR-252 of the Asian tiger mosquito Aedes albopictus regulates dengue virus replication by suppressing the expression of the dengue virus envelope protein. J. Med. Virol. 2014, 86, 1428–1436. [Google Scholar] [CrossRef] [PubMed]
  132. Slonchak, A.; Hussain, M.; Torres, S.; Asgari, S.; Khromykh, A.A. Expression of mosquito microRNA Aae-miR-2940-5p is downregulated in response to West Nile virus infection to restrict viral replication. J Virol. 2014, 88, 8457–8467. [Google Scholar] [CrossRef] [PubMed]
  133. Mayoral, J.G.; Etebari, K.; Hussain, M.; Khromykh, A.A.; Asgari, S. Wolbachia infection modifies the profile, shuttling and structure of microRNAs in a mosquito cell line. PLoS One 2014, 9, e96107. [Google Scholar] [CrossRef] [PubMed]
  134. Osei-Amo, S.; Hussain, M.; O’Neill, S.L.; Asgari, S. Wolbachia-Induced aae-miR-12 miRNA negatively regulates the expression of MCT1 and MCM6 genes in Wolbachia-infected mosquito cell line. PLoS One 2012, 7, e50049. [Google Scholar] [CrossRef] [PubMed]
  135. Zhang, G.; Hussain, M.; O'Neill, S.L.; Asgari, S. Wolbachia uses a host microRNA to regulate transcripts of a methyltransferase, contributing to dengue virus inhibition in Aedes aegypti. Proc. Natl. Acad. Sci. USA 2013, 110, 10276–10281. [Google Scholar] [CrossRef] [PubMed]
  136. Wang, J.; Wu, Y.; Yang, G.; Aksoy, S. Interactions between mutualist Wigglesworthia and tsetse peptidoglycan recognition protein (PGRP-LB) influence trypanosome transmission. Proc. Natl. Acad. Sci. USA 2009, 106, 12133–12138. [Google Scholar] [CrossRef] [PubMed]
  137. Pais, R.; Lohs, C.; Wu, Y.; Wang, J.; Aksoy, S. The obligate mutualist Wigglesworthia glossinidia influences reproduction, digestion, and immunity processes of its host, the tsetse fly. Appl. Environ. Microbiol. 2008, 74, 5965–5974. [Google Scholar] [CrossRef] [PubMed]
  138. Hughes, G.L.; Rivero, A.; Rasgon, J.L. Wolbachia can enhance Plasmodium infection in mosquitoes: Implications for malaria control? PLoS Pathog. 2014, 10, e1004182. [Google Scholar] [CrossRef] [PubMed]
  139. Hughes, G.L.; Vega-Rodriguez, J.; Xue, P.; Rasgon, J.L. Wolbachia strain wAlbB enhances infection by the rodent malaria parasite Plasmodium berghei in Anopheles gambiae mosquitoes. Appl. Environ. Microbiol. 2012, 78, 1491–1495. [Google Scholar] [CrossRef] [PubMed]
  140. Hughes, G.L.; Koga, R.; Xue, P.; Fukatsu, T.; Rasgon, J.L. Wolbachia infections are virulent and inhibit the human malaria parasite Plasmodium falciparum in Anopheles gambiae. PLoS Pathog. 2011, 7, e1002043. [Google Scholar] [CrossRef] [PubMed]
  141. Murdock, C.C.; Blanford, S.; Hughes, G.L.; Rasgon, J.L.; Thomas, M.B. Temperature alters Plasmodium blocking by Wolbachia. Sci Rep 2014, 4, 3932. [Google Scholar] [CrossRef] [PubMed]
  142. Bourtzis, K.; Pettigrew, M.M.; O'Neill, S.L. Wolbachia neither induces nor suppresses transcripts encoding antimicrobial peptides. Insect Mol. Biol. 2000, 9, 635–639. [Google Scholar] [CrossRef] [PubMed]
  143. Chrostek, E.; Marialva, M.S.P.; Yamada, R.; O'Neill, S.L.; Teixeira, L. High anti-viral protection without immune upregulation after interspecies Wolbachia transfer. PLoS One 2014, 9, e99025. [Google Scholar] [CrossRef] [PubMed]
  144. Lu, P.; Bian, G.; Pan, X.; Xi, Z. Wolbachia induces density-dependent inhibition to dengue virus in mosquito cells. PLoS Neglect. Trop. Dis. 2012, 6, e1754. [Google Scholar] [CrossRef]
  145. Frentiu, F.D.; Robinson, J.; Young, P.R.; McGraw, E.A.; O'Neill, S.L. Wolbachia-mediated resistance to dengue virus infection and death at the cellular level. PLoS One 2010, 5, e13398. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  146. Chu, J.J.H.; Leong, P.W.H.; Ng, M.L. Analysis of the endocytic pathway mediating the infectious entry of mosquito-borne flavivirus West Nile into Aedes albopictus mosquito (C6/36) cells. Virology 2006, 349, 463–475. [Google Scholar] [CrossRef] [PubMed]
  147. Chu, J.J.H.; Ng, M.L. Infectious entry of West Nile virus occurs through a clathrin-mediated endocytic pathway. J. Virol. 2004, 78, 10543–10555. [Google Scholar] [CrossRef] [PubMed]
  148. van der Schaar, H.M.; Rust, M.J.; Waarts, B.-L.; van der Ende-Metselaar, H.; Kuhn, R.J.; Wilschut, J.; Zhuang, X.; Smit, J.M. Characterization of the early events in dengue virus cell entry by biochemical assays and single-virus tracking. J. Virol. 2007, 81, 12019–12028. [Google Scholar] [CrossRef] [PubMed]
  149. Van der Schaar, H.M.; Rust, M.J.; Chen, C.; van der Ende-Metselaar, H.; Wilschut, J.; Zhuang, X.; Smit, J.M. Dissecting the cell entry pathway of dengue virus by single-particle tracking in living cells. PLoS Pathog. 2008, 4, e1000244. [Google Scholar] [CrossRef] [PubMed]
  150. Acosta, E.G.; Castilla, V.; Damonte, E.B. Functional entry of dengue virus into Aedes albopictus mosquito cells is dependent on clathrin-mediated endocytosis. J. Gen. Virol. 2008, 89, 474–484. [Google Scholar] [CrossRef] [PubMed]
  151. Junjhon, J.; Pennington, J.G.; Edwards, T.J.; Perera, R.; Lanman, J.; Kuhn, R.J. Ultrastructural characterization and three-dimensional architecture of replication sites in dengue virus-infected mosquito cells. J. Virol. 2014, 88, 4687–4697. [Google Scholar] [CrossRef] [PubMed]
  152. Martín-Acebes, M.A.; Blázquez, A.-B.; Jiménez de Oya, N.; Escribano-Romero, E.; Saiz, J.-C. West Nile virus replication requires fatty acid synthesis but is independent on phosphatidylinositol-4-phosphate lipids. PLoS One 2011, 6, e24970. [Google Scholar] [CrossRef] [PubMed]
  153. Perera, R.; Riley, C.; Isaac, G.; Hopf-Jannasch, A.S.; Moore, R.J.; Weitz, K.W.; Pasa-Tolic, L.; Metz, T.O.; Adamec, J.; Kuhn, R.J. Dengue virus infection perturbs lipid homeostasis in infected mosquito cells. PLoS Pathog. 2012, 8, e1002584. [Google Scholar] [CrossRef] [PubMed]
  154. Jupatanakul, N.; Sim, S.; Dimopoulos, G. Aedes aegypti ML and Niemann-Pick type C family members are agonists of dengue virus infection. Dev. Comp. Immunol. 2013, 43, 1–9. [Google Scholar] [CrossRef] [PubMed]
  155. Heaton, N.S.; Perera, R.; Berger, K.L.; Khadka, S.; LaCount, D.J.; Kuhn, R.J.; Randall, G. Dengue virus nonstructural protein 3 redistributes fatty acid synthase to sites of viral replication and increases cellular fatty acid synthesis. Proc. Natl. Acad. Sci. USA 2010, 107, 17345–17350. [Google Scholar] [CrossRef] [PubMed]
  156. Sinkins, S.P. Wolbachia and arbovirus inhibition in mosquitoes. Future Microbiol. 2013, 8, 1249–1256. [Google Scholar] [CrossRef] [PubMed]
  157. Sacramento, D.R.; Coelho, R.; Wigg, M.D.; De Toledo Luna Linhares, L.F.; Matos dos Santos, M.G.; De Azevedo Soares Semêdo, L.T.; Ribeiro da Silva, A.J. Antimicrobial and antiviral activities of an actinomycete (Streptomyces sp.) isolated from a Brazilian tropical forest soil. World J. Microbiol. Biotechnol. 2004, 20, 225–229. [Google Scholar] [CrossRef]
  158. Durán, N.; Menck, C.F.M. Chromobacterium violaceum: A review of pharmacological and industiral perspectives. Crit. Rev. Microbiol. 2001, 27, 201–222. [Google Scholar] [CrossRef] [PubMed]
  159. Joyce, J.D.; Nogueira, J.R.; Bales, A.A.; Pittman, K.E.; Anderson, J.R. Interactions between la crosse virus and bacteria isolated from the digestive tract of Aedes albopictus (diptera: culicidae). J. Med. Entomol. 2011, 48, 389–394. [Google Scholar] [CrossRef] [PubMed]
  160. Hughes, G.L.; Dodson, B.L.; Johnson, R.M.; Murdock, C.C.; Tsujimoto, H.; Suzuki, Y.; Patt, A.A.; Cui, L.; Nossa, C.W.; Barry, R.M.; et al. Native microbiome impedes vertical transmission of Wolbachia in Anopheles mosquitoes. Proc. Natl. Acad. Sci. USA 2014, 111, 12498–12503. [Google Scholar] [CrossRef] [PubMed]
  161. Rasgon, J.L. Dengue fever: Mosquitoes attacked from within. Nature 2011, 476, 407–408. [Google Scholar] [CrossRef] [PubMed]
  162. Frentiu, F.D.; Zakir, T.; Walker, T.; Popovici, J.; Pyke, A.T.; van den Hurk, A.; McGraw, E.A.; O'Neill, S.L. Limited dengue virus replication in field-collected Aedes aegypti mosquitoes infected with Wolbachia. PLoS Neglect. Trop. Dis. 2014, 8, e2688. [Google Scholar] [CrossRef] [Green Version]
  163. Eliminate Dengue Program. Available online: http://www.eliminatedengue.com/program (accessed on 21 July 2014).
  164. Müller, G.C.; Beier, J.C.; Traore, S.F.; Toure, M.B.; Traore, M.M.; Bah, S.; Doumbia, S.; Schlein, Y. Successful field trial of attractive toxic sugar bait (ATSB) plant-spraying methods against malaria vectors in the Anopheles gambiae complex in Mali, West Africa. Malar. J. 2010, 9, 210. [Google Scholar] [CrossRef] [PubMed]
  165. Müller, G.C.; Junnila, A.; Schlein, Y. Effective control of adult Culex pipiens by spraying an attractive toxic sugar bait solution in the vegetation near larval habitats. J. Med. Entomol. 2010, 47, 63–66. [Google Scholar] [CrossRef] [PubMed]
  166. Stewart, Z.P.; Oxborough, R.M.; Tungu, P.K.; Kirby, M.J.; Rowland, M.W.; Irish, S.R. Indoor application of attractive toxic sugar bait (ATSB) in combination with mosquito nets for control of pyrethroid-resistant mosquitoes. PLoS One 2013, 8, e84168. [Google Scholar] [CrossRef] [PubMed]
  167. Qualls, W.A.; Müller, G.C.; Revay, E.E.; Allan, S.A.; Arheart, K.L.; Beier, J.C.; Smith, M.L.; Scott, J.M.; Kravchenko, V.D.; Hausmann, A.; et al. Evaluation of attractive toxic sugar bait (ATSB)-Barrier for control of vector and nuisance mosquitoes and its effect on non-target organisms in sub-tropical environments in Florida. Acta Trop. 2014, 131, 104–110. [Google Scholar] [CrossRef] [PubMed]
  168. Barbosa, R.M.R.; Souto, A.; Eiras, A.E.; Regis, L. Laboratory and field evaluation of an oviposition trap for Culex quinquefasciatus (Diptera: Culicidae). Mem. Inst. Oswaldo Cruz 2007, 102, 523–529. [Google Scholar] [CrossRef] [PubMed]
  169. Honório, N.A.; de Barros, F.S.M.; Tsouris, P.; Rosa-Freitas, M.G. Occurrence of Toxorhynchites guadeloupensis (Dyar Knab) in oviposition trap of Aedes aegypti (L.) (Diptera: Culicidae). Neotrop. Entomol. 2007, 36, 809–811. [Google Scholar] [CrossRef] [PubMed]
  170. Wang, S.; Fang, W.; Wang, C.; St Leger, R.J. Insertion of an esterase gene into a specific locust pathogen (Metarhizium acridum) enables it to infect caterpillars. PLoS Pathog. 2011, 7, e1002097. [Google Scholar] [CrossRef] [PubMed]
  171. Fang, W.; Azimzadeh, P.; St Leger, R.J. Strain improvement of fungal insecticides for controlling insect pests and vector-borne diseases. Curr. Opin. Microbiol. 2012, 15, 232–238. [Google Scholar] [CrossRef] [PubMed]

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Jupatanakul, N.; Sim, S.; Dimopoulos, G. The Insect Microbiome Modulates Vector Competence for Arboviruses. Viruses 2014, 6, 4294-4313. https://doi.org/10.3390/v6114294

AMA Style

Jupatanakul N, Sim S, Dimopoulos G. The Insect Microbiome Modulates Vector Competence for Arboviruses. Viruses. 2014; 6(11):4294-4313. https://doi.org/10.3390/v6114294

Chicago/Turabian Style

Jupatanakul, Natapong, Shuzhen Sim, and George Dimopoulos. 2014. "The Insect Microbiome Modulates Vector Competence for Arboviruses" Viruses 6, no. 11: 4294-4313. https://doi.org/10.3390/v6114294

APA Style

Jupatanakul, N., Sim, S., & Dimopoulos, G. (2014). The Insect Microbiome Modulates Vector Competence for Arboviruses. Viruses, 6(11), 4294-4313. https://doi.org/10.3390/v6114294

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