RNA interference (RNAi) was first described in plants as a mechanism for “post-transcriptional gene silencing” [17], and later, “virus-induced gene silencing” [18], two phenomena which, at the time, were seemingly unrelated. Several years after these initial observations, double-stranded RNA (dsRNA) was found to be the trigger for RNAi in Caenorhabditis elegans [19] and Drosophila melanogaster [20]. In invertebrates and plants, exo-RNAi is induced by cellular recognition of long dsRNAs as pathogen-associated molecular patterns (PAMPs), which naturally occur as viral genome replication intermediates and genomic RNA secondary structures in the case of RNA viruses, and as convergent transcripts in DNA viruses. These dsRNAs are recognized and cleaved by Dicer-2 (DCR2), a cytoplasmic RNase III enzyme, resulting in 19–23 base pair (bp) fragments (predominately 21 bps) termed siRNAs. siRNA duplexes produced in this manner exhibit 5' monophosphates and 3' hydroxyls, as well as two-nucleotide (nt) overhangs on their 3' termini. These siRNAs are then loaded into the Argonaute-2 (AGO2)-containing RNA-induced silencing complex (RISC) through association with a DCR2/R2D2 heterodimer [21]. After the duplex is unwound a single-stranded RNA known as the guide strand remains associated with the RISC, and is 2'-O methylated by the methyltransferase DmHEN1 [22,23], and the complimentary strand, known as the passenger strand, is discarded. The RISC then recognizes cognate mRNA (in this case, virus genomic RNA) by sequence complementarity with the guide strand. Degradation of the target occurs through the Slicer endonuclease activity of AGO2 [24]. Unlike miRNAs, where mismatches between the guide strand and target are tolerated, even a single mismatch in complementarity between a siRNA and its target can result in diminished or abolished silencing [25,26]. In this way, the siRNA pathway acts as a highly potent antiviral pathway in controlling arbovirus infection.

The role of the siRNA pathway in antiviral defense in arthropods has been the subject of intense investigation in recent years. In Drosophila, numerous studies have demonstrated that RNAi inhibits virus replication [27,28,29]. Notably, Drosophila with a null mutant DCR2 enzyme exhibit ~70% mortality and dramatically higher virus titers when inoculated with Sindbis virus (SINV, Togaviridae), as compared to wild-type controls [28]. In mosquitoes, evidence of the involvement of the siRNA pathway during arbovirus infection has been observed in several virus/arthropod pairings, including o'nyong-nyong virus (ONNV, Togaviridae) in Anopheles gambiae [30], SINV in Aedes aegypti [31], and DENV in Ae. aegypti [32]. viRNAs produced in response to WNV had been detected in Drosophila S2 cells, but not Ae. albopictus C6/36 cells [33] due to a dysfunctional siRNA pathway [34] resulting from a single nucleotide deletion introducing a premature stop codon within the open reading frame (ORF) of DCR-2 [35]. One potential pitfall in interpreting the results of these studies is the utilization of non-natural virus/vector pairings and/or infection routes (i.e., intrathoracic inoculation), with the exception of Sanchez-Vargas et al. [32]. Brackney et al. utilized next-generation sequencing (NGS) to profile the antiviral RNAi response to WNV in its natural vector, Cx. quinquefasciatus mosquitoes, following peroral infection, and found viRNAs produced in the midgut of mosquitoes at 7 and 14 days post-infection (dpi) [36]. viRNAs produced in this manner were primarily 21 nts in length (indicative of DCR2 processing), and were asymmetrically distributed along the length of the virus genome.

Given the requirement for high target sequence complementarity in siRNAs, it comes as no surprise that RNAi can drive viral diversity and evolution through the generation of RNAi-escape mutants that differ sufficiently from the master sequence. Viral escape from one or a few transfected siRNAs has been observed in a variety of different systems, including hepatitis C virus (HCV, Flaviviridae) [37], human immunodeficiency virus-1 (HIV-1, Retroviridae) [26], turnip mosaic virus (TuMV, Potyviridae) [38], and poliovirus (PV, Picornaviridae) [39]. Based on the observation that WNV population structure was more complex in mosquitoes than in birds [40], it was hypothesized that the mosquito RNAi pathway may serve as a potent selective pressure on the virus to favor generation and maintenance of rare mutants. Indeed, a correlation between nucleotide targeting and increased likelihood for corresponding point mutations has been observed [36], though it is important to note that this observation was associative, and does not demonstrate causation. Taken together, these studies highlight the role this pathway may play in mosquito innate antiviral immunity, and shed light on how it may influence virus diversification and evolution. Moreover, the error rate of the virus replicase complex may serve in part as an evolutionary mechanism for circumventing the mosquito siRNA-based antiviral response through the generation of rare mutants which differ sufficiently from the master sequence. This is circumstantially supported by the observation that a chikungunya virus (CHIKV, Togaviridae) mutant expressing a high fidelity RNA-dependent RNA polymerase (RdRP) exhibited lower infection and dissemination titers in mosquitoes as compared to the wild-type virus [41]. However, it should be noted that the molecular basis for this observation was not investigated, and that immunological factors other than RNAi could potentially influence the fitness of a genetically homogenous virus population.

Systemic RNAi, first described in plants and worms, is the process by which the siRNA response spreads beyond the site of initiation into surrounding cells and tissues (see [42] for a review). The mechanism of spread differs between plants and animals, with short-distance transport of siRNAs in plants occurring through plasmodesmal junctions connecting cells, and long-distance transport being mediated by the vascular system. In C. elegans, spread of the RNAi signal is mediated by members of the SID family of transmembrane transporters [43,44]. Evidence suggests that this process may occur in dipterans as well, and has broad implications in understanding RNAi-based antiviral immunity in these systems. Studies in flies have revealed a systemic RNAi pathway [45,46], and cell-to-cell spread of viRNAs produced in response to Semliki Forest virus (SFV, Togaviridae) infection has been demonstrated in mosquito cell culture [47]. In C. elegans, primary viRNAs are amplified in a target dependent manner by the endogenous RdRP RRF-1, which contributes to systemic RNAi spread and subsequent maintenance of long-term silencing [48]. Secondary viRNAs produced in this manner are composed entirely of antisense polarity, and exhibit 5' di- or triphosphates, making them structurally unique, and thus a distinct class of small RNA molecules [49,50]. While an RdRP capable of amplifying viRNAs has not been conclusively identified in dipterans, a recent publication suggests that viral RNA produced during flock house virus (FHV, Nodaviridae) in Drosophila can be reverse-transcribed into viral cDNAs mediated by the reverse transcriptase activity of endogenous long terminal repeat (LTR)-retrotransposons [51]. Viral cDNA produced in this manner may then be integrated into the host cell genome, or circularized into stable, extrachromasomal DNA which can be efficiently transcribed into dsRNAs that can be fed back into the siRNA pathway, leading to a primed immune response, and allowing for a persistent infection to develop. Additionally, viRNAs produced in response FHV infection in C. elegans have been observed to be transgenerationally inherited from mother to offspring in successive generations [52], raising the intriguing possibility that similar mechanisms of amplification and non-Mendelian, extrachromosomal inheritance of small RNAs may exist in mosquitoes as well, though it should be noted that to date, there is a lack of experimental data supporting this. Given the aforementioned importance of the RNAi pathway in mosquito innate immunity to viral infection, inheritance of viRNAs might be expected to influence mosquito vector competence and arbovirus populations in nature.

Cross-talk between SRRPs and other innate immune pathways is an emerging feature of mosquito antiviral defense against arboviruses. DCR2 belongs to the same family of DExD/H-box helicases as the RIG-I-like receptors, which are involved in the induction of the IFN response in mammalian systems. Deddouche et al. reported that the secreted peptide vago is induced in D. melanogaster in response to Drosophila C virus (DCV, Dicistroviridae) and SINV infection in the fat bodies of flies, and that induction of vago was reliant on the amino terminal DExD/H-box domain of DCR2 [53]. Notably, infection by FHV did not induce vago expression, likely due to the FHV encoded viral suppressor of RNAi, B2, which binds dsRNA thereby interfering with downstream signaling by DCR2. While vago was shown to control virus infection in this study, the mechanism by which it did so was unclear. Pradakar et al. further explored vago’s role in antiviral immunity using cultured mosquito cells and WNV [54]. It was found that vago was effective in limiting WNV infection in Cx. quinquefasciatus-derived Hsu cells, and that induction of vago resulted in activation of Jak/STAT signaling, leading to the induction of the STAT-dependent virus inducible gene vir-1, thereby restricting WNV replication. Although the identity of the cellular receptor for vago is currently not clear, studies published to date suggest that components of the RNAi pathway can have diverse, multifunctional roles in controlling arbovirus infections in mosquitoes.

Recent evidence suggests that a second class of small RNAs with distinct biogenesis may also be induced in arthropods following virus infection. The p-element induced wimpy testes (PIWI) class of Argonaute proteins were first discovered in 1997 by Lin et al. and shown to be potent regulators of spermatogenesis in Drosophila [55]. In 2006, several studies were published indicating that PIWI proteins interact with a unique class of small RNAs, named PIWI-interacting RNAs (piRNAs; the nomenclature at the time was variable, and they were also referred to as repeat-associated small interfering RNAs [rasiRNA], which are now considered a distinct subclass of piRNAs) [56,57,58,59,60]. piRNAs exhibit some unique features that distinguish them from miRNAs and siRNAs. First, piRNAs are considerably larger than most miRNAs or siRNAs, the latter two ranging from 20–23 nts in length, the former 24–30 nts. Secondly, piRNAs are produced from single stranded precursor molecules independently of Dicer processing. Like siRNAs, but not miRNAs, piRNAs are modified by DmHEN1, which results in 2'-O-methylation at the 3' terminus of the RNA [23,61]. Additionally, expression of piRNAs shows tissue specificity, with gonadal tissue being highly enriched for this species and associated proteins. Endogenously transcribed and processed piRNAs have been shown to be important repressors of transposable elements (TEs) in these tissues. However, expression of piRNAs has been detected in somatic tissue as well [62,63].

Biogenesis of piRNAs is proposed to occur through two pathways: the primary pathway and the ping-pong dependent amplification loop. In the primary pathway, piRNAs are processed from single-stranded precursor molecules transcribed from genomic loci (piRNA clusters). In flies, primary piRNAs are associated with PIWI and Aubergine (AUB) [64], and are typically antisense to TEs [59]. Primary piRNAs produced in this manner exhibit a strong bias for a uridine residue at the 5'-terminus of the transcript (U₁). These primary piRNAs are then fed into the ping-pong dependent amplification cycle, whereby after binding to their target transcript, cleavage occurs 10 nts upstream from the 5' terminus of the primary piRNA [65,66]. Thus, Argonaute 3 (AGO3)-associated secondary piRNAs exhibit an adenine residue in the 10 position (A₁₀). Secondary piRNAs subsequently bind complementary transcripts, resulting in cleavage at the A₁₀-U basepairing, producing piRNAs corresponding to the sequence of the initial primary piRNA they were derived from, thereby restarting the cycle.

Recently, the piRNA pathway has been implicated in antiviral immunity in invertebrates. A possible hypothesis is that the piRNA pathway can act in a compensatory manner, such as when the siRNA pathway is overburdened or suppressed. Indeed, in Drosophila, flies with a null mutant PIWI protein exhibited significantly higher titers of WNV in comparison to wild-type controls [33]. Silencing of AGO3 in An. gambiae has been shown to result in increased dissemination of ONNV [30], and dsRNA-knockdown of piRNA-pathway component proteins has been shown to result in increased titers of SFV in Ae. aegypti-derived Aag2 cells, with knockdown of PIWI-4 in particular showing considerable effect in this regard [67]. Additionally, sequencing of DENV2-infected Ae. albopictus C6/36 cells, which, as previously mentioned, have a dysfunctional siRNA pathway [34,35], revealed a shift from the stereotypically predominant distribution of siRNAs (19–23 nts, no significant strand bias) to products consistent with the piRNA pathway (24–30 nts, predominately positive strand, A₁₀ bias) [68]. Wu et al. described a population of virus-derived small RNAs in their sequencing of persistently-infected Drosophila ovary somatic sheet (OSS) cells that held hallmarks of piRNAs; specifically, being between 24–30 nts in length, exhibiting a strong (95%) strand bias, and a preference for a 5' uracil (though, notably, no bias for an A₁₀ was seen) [69]. Similarly, 24–30 nt small RNAs produced in response to DENV2 in Ae. aegypti mosquitoes have been found by deep-sequencing [70]. Interestingly, piRNA-like small RNAs sequenced in the latter study exhibited no preference for a 5' uracil, and only a slight bias for an A₁₀ residue.

Recently, piRNA-like small RNAs exhibiting characteristics of ping-pong dependent amplification have been sequenced in mosquitoes and mosquito cells after virus infection. Morazzani et al. found that piRNA-like small RNAs are produced in the soma (non-ovarial tissue) of both Ae. aegypti and Ae. albopictus after infection with CHIKV, and that, unlike endogenously transcribed piRNAs, dsRNA was likely the biogenic precursor for virus-derived piRNAs [35]. Likewise, small RNAs from Aag2 and Ae. albopictus-derived U4.4 cells, infected with SINV, shared these same characteristics [71]. In the same study, sequencing data from C6/36 cells infected with La Crosse virus (LACV, Bunyaviridae) revealed ping-pong dependent signatures in the piRNA-like viral small RNA population. Similar results from small RNA deep-sequencing of virus infected invertebrate hosts has also been seen in Rift Valley fever virus (RVFV, Bunyaviridae) infected mosquito cells [72], as well as Schmallenberg virus (SBV, Bunyaviridae)-infected Culicoides sonorensis-derived KC cells and Aag2 cells, and blue-tongue virus (BTV, Reoviridae)-infected Culicoides and mosquito cells [73] (though notably, 24–30 nt small RNAs sequenced from BTV-infected cells did not exhibit signatures of ping-pong dependent amplification). The disparity between different virus/host pairings producing piRNAs either possessing or lacking signatures of ping-pong dependent amplification suggests that the piRNA response may be differentially modulated in response to infection by diverse viruses. While no studies profiling the piRNA response in a Culex mosquito/WNV pairing have been published, the existing data in these numerous other infection models suggest that it too may play a significant role in this system.

Most arboviruses cause persistent infections within their arthropod vectors. This has led to speculation as to how these viruses maintain infection in the face of a robust RNAi response. In addition to the evolutionary mechanisms (i.e., the viral replicase error rate) described above, many plant and insect-specific viruses have developed molecular mechanisms for subverting the host RNAi response. For example, the previously mentioned FHV encodes a protein, B2, which binds to dsRNA and inhibits the function of DCR2, effectively rendering the siRNA pathway inert [74]. Plasmid-expressed LACV NSs protein has been shown to inhibit IFN and RNAi in mammalian cells and mice [75,76]. However, NSs fails to show any RNAi-suppressive effect in LACV-infected C6/36 cells or in NSs plasmid-transfected U4.4 cells infected with SFV [75]. A recent publication demonstrated that DENV NS4b functions as a VSR in human Huh7 cells via inhibition of dsRNA processing by Dicer [77]. However, whether NS4b behaves similarly in mosquitoes has not been investigated, and to date, no VSR activity has been described for an arbovirus protein during mosquito infection. Thus, it is currently not clear how arboviruses establish and maintain persistent infection of arthropods.

Recently, intriguing evidence has been presented that viral subgenomic RNAs may function as VSRs in some cases. All flaviviruses studied thus far produce a sub-genomic RNA product (sfRNA) from the 3' UTR of the virus genome [78]. In WNV, this RNA comprises the last 525 nts of the virus genome. sfRNA is produced by incomplete degradation of the viral genome by the cellular 5'-to-3' exoribonuclease XRN1, which stalls on the conserved pseudoknot-like structures present at the 5'-terminus of the 3' UTR, resulting in large amounts of sfRNA accumulating within infected cells [79,80]. Recent evidence using fluorescent reporter assays suggests that the WNV sfRNA may act as a VSR by interfering with DCR2 in a concentration-dependent manner [81], though this type of assay alone may be prone to misinterpretation, and should be corroborated by genetic rescue experiments [82]. The mechanism through which sfRNA exerts this effect is unclear, but it is hypothesized that it acts as a decoy substrate for DCR2. Interestingly, sfRNA has also been shown to act in a negative feedback loop by suppressing XRN1 activity due to the enzyme stalling at the 3' UTR [83], illustrating that multiple antiviral pathways can be manipulated by this decay product.

The Toll pathway was originally described in Drosophila as an evolutionarily conserved signaling cascade involved in the establishment of the dorso-ventral axis as well as in many other developmental processes [85,86]. It has since been characterized as having a significant role in innate immunity to gram-positive bacteria and fungi [87,88]. More recently, some studies suggest that the Toll pathway may serve an important role in antiviral immunity. In 2005, Zambon et al. observed a significant increase in the expression of the Toll regulated AMPs, Drosomycin and Metchnikowin, upon Drosophila X virus (DXV; Birnaviridae) challenge of Drosophila [89]. It was further shown that flies lacking a functional Toll pathway were significantly more susceptible to DXV challenge. Similarly, it was determined that Toll pathway components and AMPs were significantly up-regulated upon DENV infection of Ae. aegypti mosquitoes [90]. Further, suppression of Cactus (a negative regulator of Toll signaling) or MyD88 (a Toll signaling adapter protein) resulted in modest but significant reductions and accumulations of infectious DENV particles in the mosquito midguts, respectively [91]. These findings were confirmed by another study in which Toll pathway components were up-regulated upon DENV infection of Ae. aegypti salivary glands [92]. In addition, a modest increase in Dif (a Toll inducible NF-κB transcription factor) was recognized during early stages of SINV infection of Ae. aegypti [93]. Together, these data suggest that the Toll pathway plays a role in antiviral immunity in insects; however, others have observed conflicting results [94,95,96].

Colpitts et al. observed significant reductions in the expression levels of the Ae. aegypti ortholog of Drosophila Spӓtzle 5 (an upstream signaling peptide of the Toll pathway) upon Ae. aegypti infection with three flaviviruses; WNV, yellow fever virus (YFV, Flaviviridae) and DENV [94]. In addition, transcript levels of the Ae. aegypti ortholog to Drosophila Toll was reduced upon YFV infection. These findings challenge those previously published by Xi et al. [90]. Many confounding factors may account for this difference, including but not limited to experimental design, mosquito strains, virus strains and/ or environmental conditions. The authors of this study did not directly address these discrepancies, but rather suggested that viral-associated reductions potentially indicate an evolved mechanism by which arboviruses suppress antiviral pathways. However, there is currently no evidence supporting this. Using the SFV—Aedes U4.4 cell system, Fragkoudis et al. observed a down-regulation of Toll pathway component transcript levels upon infection by SFV. Furthermore, prior activation of the Toll pathway did not seem to adversely affect SFV replication in U4.4 cells [95]. From these studies it is difficult to determine with confidence the contribution of the Toll pathway to antiviral immunity of insects as a whole. However, these findings may indicate that Toll-mediated antiviral activity is specific to each virus-insect pairing. With regards to WNV, little is known about the role of this pathway during infection of insects except for the apparent down-regulation of the Spӓtzle5-like cytokine during infection of Ae. aegypti mosquitoes and that none of the canonical Toll pathway genes and/or associated AMPs were significantly altered during WNV infection of Cx. quinquefasciatus [96]. Further research will be needed to fully elucidate the possible contributions of Toll pathway mediated antiviral immunity during WNV infection of mosquitoes.

The Imd pathway is another immune signaling cascade of insects and bears striking similarities to the Tumor Necrosis Factor (TNF) pathway of mammals. It was initially described after the identification of a mutant Drosophila line that had significantly decreased levels of numerous AMPs, yet maintained normal levels of another AMP, Drosomycin [87,97,98]. These findings indicated that the expression of AMPs was controlled by two or more regulatory cascades. It was later determined that imd mutant flies were highly susceptible to infection with gram-negative bacteria yet maintain resistance to gram-positive bacteria and fungi [85]. Recently, the Imd pathway has been implicated in antiviral immunity in insects. The first indication of the potential significance of the Imd pathway during viral infections was observed in the Drosophila, cricket paralysis virus (CrPV; Dicistroviridae) model [99]. Somewhat paradoxically, the authors found that while AMPs were not up-regulated during infection with CrPV, suggesting that neither the Imd or Toll pathways were responsive to CrPV infection, mutant flies lacking components of the Imd pathway were more susceptible to viral infection resulting in shortened lifespan and increased viral replication. Together, these data suggest that Imd activation is uncoupled from AMP induction during infection with CrPV, and implies that induction of the Imd pathway results in transcription of several other genes with presently unknown roles in immunity [100]. The role of the Imd pathway in antiviral immunity is further supported by the observations of Avadhanula et al. and Huang et al. Using a novel SINV replicon transgenic Drosophila line, the authors observed increased AMP expression as well as a modest but significant increase in susceptibility of Imd-deficient lines to SINV challenge [101,102]. Additionally, Sigma virus (SIGMAV; Rhabdoviridae) infection of Drosophila induced the expression of numerous Imd controlled AMPs; however, the significance of these observations on SIGMAV replicative fitness or fly survivorship were not assessed [103]. While there have been several studies implicating the role of Imd during antiviral immunity, there have been just as many suggesting otherwise. The induction of almost all AMPs were observed during DXV infection of Drosophila, yet mutant flies lacking a functional Rel (an Imd inducible transcription factor) were no more susceptible to DXV challenge than the controls [89]. Further, a lack of AMP induction was observed during DCV infection of Drosophila [53,104]. As with the Toll pathway, it is difficult to discern the significance of the Imd pathway in antiviral immunity based on these studies. This uncertainty is further confounded by the fact that the majority of these viruses do not naturally infect Drosophila, with the exception of SIGMAV and DCV.

These ambiguities in the literature also encompass mosquito-virus interactions. Recently, a cecropin-like peptide presumed to be under the control of the Imd signaling cascade was found to be significantly induced upon DENV infection of Ae. aegypti salivary glands [92]. Characterization of this peptide revealed that it could potently inhibit DENV and CHIKV replication in mosquito cell culture. However, the significance of its antiviral effect has not been confirmed with in vivo knock-down studies. In addition, other studies have observed either down-regulation or insignificant differences in the expression of Imd-controlled AMPs or pathway components during SFV infection of Aedes U4.4 cells or ONNV infection of An. gambiae [95,105], although in the former study, decreased SFV replication in Imd and Jak/STAT activated U4.4 cells was observed. The biological relevance of these findings is difficult to determine at this time because functional assays assessing their significance during natural infections of adult mosquitoes have not been performed. Studies investigating the role of Imd during WNV infection of mosquitoes are lacking, and it remains to be seen what if any effect this pathway has on shaping antiviral immunity to WNV in relevant mosquito vectors; however, it was demonstrated that Imd gene transcripts and/or associated AMP transcripts were unaltered during WNV infection of Cx. quinquefasciatus [96].

The Jak/STAT pathway is an evolutionarily conserved pathway first described for its role in embryonic segmentation in Drosophila [106]. Subsequently it was determined that it has an important role in antibacterial defense. It is comprised of the three major components, the receptor Domeless, the Janus Kinase (Jak) Hopscotch, and the transcription factor STAT [106,107,108]. Unlike the Toll and Imd pathways, a well characterized subset of inducible effector AMPs have not been associated with this pathway; however, a handful of inducible genes containing a STAT binding site in their promoters have been identified, with some of these gene products appearing to have antiviral effects.

The initial findings implicating a role of Jak/STAT in antiviral immunity in insects were observed in the Drosophila—DCV model. By performing microarray analysis on bacterial, fungal, and DCV challenged flies, the authors identified a subset of gene products that were up-regulated during DCV infection but not during fungal or bacterial challenge [104]. Upon closer inspection the authors identified a gene, vir-1, that contained the STAT binding site within its promoter and was strongly induced during DCV and FHV challenge. Furthermore, it was demonstrated that Jak/STAT deficient flies were more susceptible to DCV challenge as determined by increased DCV replication and increased mortality. These results were confirmed by studies performed by Kemp et al. who also demonstrated that the Jak/STAT pathway was important in Drosophila antiviral immunity to CrPV [109]. Interestingly, when Jak/STAT deficient flies were challenged with five other evolutionarily divergent viruses, including SINV, DXV and FHV, the authors found no effect on the survivorship of the flies. These results indicate that Jak/STAT involvement in antiviral immunity may be specific to each virus–insect pairing and not broadly applicable to all systems.

A role for Jak/STAT involvement in mosquito immunity to arboviruses has been described. Specifically, components of the Jak/STAT pathway as well as Jak/STAT inducible gene products were found to be up-regulated upon DENV infection of Ae. aegypti mosquitoes [90,110]. Included in these findings were two novel Jak/STAT inducible genes termed Dengue Virus Restriction Factors 1 and 2 (DVRF1-2). Further, it was demonstrated that suppression of these two genes resulted in 2.5- and 2.2-fold increases in DENV-2 replication in mosquitoes, respectively [110]. In addition, a recent study demonstrated that WNV induces the expression of vago in Hsu cells. Subsequent silencing of vago increased WNV titers. The antiviral effect of vago expression was determined to arise from downstream vir-1 activation via vago-induced Jak/STAT signaling. Studies in mosquitoes will be needed to validate the in vivo role of vago in controlling arbovirus infection. As with the Toll and Imd pathways others have observed conflicting data on the role of Jak/STAT pathway in antiviral immunity. In these studies there were no indications of Jak/STAT up-regulation upon infection in four separate virus-vector models (ONNV/An. gambiae, SFV/Aedes U4.4 cells, WNV-DENV-YFV/Ae. aegypti, WNV/Cx. quinquefasciatus) [94,95,96,105]. However, as previously mentioned, activation of the Jak/STAT pathway prior to SFV infection resulted in reduced viral replication in U4.4 cells. From these studies it is difficult to accurately assess the importance of the Jak/STAT pathway in antiviral immunity. Future studies assessing the importance of this pathway on arboviral infection, dissemination and transmission rates within mosquitoes will help clarify its significance. In the context of WNV and mosquitoes, this holds true especially considering the systems utilized; however, it is difficult to determine the relevance considering that one study was limited to cell culture and the other in an ancillary vector. Additional research into the role of Jak/STAT in antiviral immunity in primary mosquito vectors is clearly required in order to fully understand its possible influence on mosquito antiviral responses.

Among arthropods, the PO cascade is an evolutionarily conserved extracellular pathway responsible for wound healing and melanization of bacterial and parasitic pathogens [84]. This pathway can be induced by cuticular damage or upon recognition of PAMPs. This in turn activates a serine protease cascade ultimately resulting in the activation of the prophenoloxidase activating enzyme (PPAE). Active PPAE cleaves the prophenoloxidase zymogen to produce PO, which catalyzes the conversion of mono- and diphenolic substrates to quinones, which is then converted to melanin [84]. While direct melanization of viruses has not been observed, there is evidence that by-products of the pathway may have antiviral effects. It was demonstrated that plasma from the tobacco budworm has virucidal effects on Helicoverpa zea single capsid nucleopolyhedrovirus (HzSNPV; Baculoviridae) and chemical inhibition of PO resulted in increased viral titers [111,112]. It was also determined that 5,6-dihydroxyindole, a byproduct of the PO cascade, could almost completely inactivate Autographa californica nucleopolyhedrovirus (AcMNPV; Baculoviridae) in vitro [113]. Similar antiviral activity has been observed in mosquitoes. Tamang et al. observed increased SINV titers in Armigeres subalbatus mosquitoes after suppression of prophenoloxidase I [114]. Such observations were also found during SFV infection of U4.4 cells. Specifically, recombinant SFV over-expressing an inhibitor of the PO cascade was able to replicate to significantly higher titers than the control virus. It was further determined that inhibition of the PO cascade could decrease the survivorship of SFV infected Ae. aegypti [115]. The authors went on to demonstrate that the specific effector molecules involved in the antiviral effects were the pathway intermediate quinones. The specific nature of their antiviral effect remains to be determined. Together, these studies highlight the potential role of the PO cascade in insect immunity to arboviruses. These are the first studies to assess the role of the PO cascade in antiviral immunity in mosquitoes and it will be interesting to see if this antiviral effect is conserved among other virus-vector pairings. Future research into the involvement PO cascade during WNV infection of Culex spp. mosquitoes is warranted.

Autophagy is an evolutionarily conserved pathway that serves an important role in maintaining cellular homeostasis and cell survival [121]. Induction of autophagy results in the formation of double-phospholipid membrane vesicles termed autophagosomes which sequester the targeted organelles and proteins. Subsequently, the autophagosomes fuse with lysosomes forming autolysosomes which mediate degradation of the contents [122]. During normal growth conditions autophagy maintains cellular homeostasis by degrading unwanted or damaged organelles and protein aggregates. In times of cellular stress, autophagy catabolizes these cellular components thereby generating a pool of energy and macromolecules that maintain crucial cellular functions until favorable growth conditions return [121]. In addition, it appears to play a significant role in antiviral immunity in invertebrates. Shelly et al. demonstrated that deletion of autophagy-related genes (Atg’s) increased vesicular stomatitis virus (VSV; Rhabdoviridae) replication and decreased Drosophila survivorship [123]. It was later demonstrated that pre-formed PAMPs in the context of UV-inactivated virus appear to interact with Toll-7 at the plasma membrane which leads to the activation of autophagy independently of the canonical Toll pathway [124]. These studies were the first to examine the antiviral role of autophagy to an arbovirus in the context of an invertebrate host. Additional studies are required in order to clarify whether these findings can be replicated in natural virus-vector pairings.

Apoptosis is a conserved mechanism of programmed cell death in multicellular organisms [125] that is critical to a variety of biological processes including embryonic development, maintenance of homeostasis, and lysis of virus-infected cells by cytotoxic T-lymphocyes [119]. In vertebrates, apoptosis may be induced through two known pathways: the extrinsic, or death-receptor pathway, and the intrinsic, or mitochondrial pathway [126], though in dipterans only the intrinsic pathway is known to exist [127].

Many viruses from diverse families encode anti-apoptotic genes, such as inhibitors of apoptosis (IAPs), p53-binding proteins, and bcl-2 homologs. This has led to the hypothesis that apoptosis functions as an innate immune pathway in response to viral infection [120]. FHV strongly induces apoptotic events in Drosophila DL-1 cells through depletion of the Drosophila inhibitor-of-apoptosis-1 (DIAP1), a negative regulator of initiator caspase DRONC and effector caspase DrICE; however replication of FHV is not negatively impacted by apoptosis [128]. Similar depletion of host cell IAPs were observed in DL-1 cells [129] and Spodoptera frugiperda (order Lepidoptera) SF21 cells [129,130] infected with AcMNPV, which encodes its own viral IAP, p35, with depletion of IAPs in both cell lines being triggered by viral DNA replication [129]. However, p35-deficient mutant viruses demonstrated reduced infectivity of SF21 cells, and reduced infectivity and lethality in Spodoptera frugiperda larvae [131]. Furthermore, studies in flies infected with FHV or AcMNPV reveal that rapid induction of apoptosis by the upstream regulator p53 limits viral gene expression and proliferation [132], further highlighting the importance of apoptosis in controlling virus infection in these systems.

Numerous studies have investigated the role apoptosis plays in arbovirus infection models. Apoptosis is a natural consequence of arbovirus infection in mammalian cells, where cytopathic effect (CPE) is frequently seen; conversely, most arboviruses are thought to cause minimal cytopathology in insect cells, instead resulting in non-lytic, persistent infections [133,134]. However, cell death consistent with apoptosis has been observed in mosquito midgut and salivary gland tissues following infection with a variety of arboviruses [135,136,137,138,139,140,141,142], including WNV. With regards to WNV, apoptosis in the midgut epithelium [138] or salivary glands [136,142] of Cx. pipiens or Cx. quinquefasciatus mosquitoes was associated with a resistance to infection or reduced ability to transmit virus, respectively. Thus, an attractive hypothesis is that apoptosis functions in an antiviral capacity to arbovirus infection in mosquitoes, and that it may be more likely to occur in mosquitoes incapable of vectoring a particular virus rather than permissive hosts [143,144]. Indeed, pro-apoptotic genes in a refractory strain of Ae. aegypti mosquitoes were significantly up-regulated between 24 and 48 h post-infection (hpi) in response to DENV2 infection as compared to a susceptible strain of Ae. aegypti or blood-only fed mosquitoes from the same strain [145]. In the same study, dsRNA-knockdown of the caspase inhibitor AeIAP1 in two susceptible strains converted the phenotype from susceptible to refractory, and knockdown of initiator-caspase AeDronc in the refractory strain resulted in increased permissiveness to DENV2 infection. The same group previously reported up-regulation of pro-apoptotic genes in the DENV2-susceptible strain in comparison to the refractory strain at 48 hpi [146], indicating that apoptotic events controlling virus infection may be induced acutely and rapidly limit the course of infection. However, it is important to note that neither of these studies directly measured apoptotic events in midgut tissue after DENV2 infection, and that the methods used to measure differential gene expression were different between studies (qPCR and suppressive subtractive hybridization, respectively [145,146]). Conversely, modulation of apoptosis by RNAi-mediated silencing of either apoptosis inhibitors or initiators in Ae. Aegypti, subsequently infected with SINV, produced results contrary to the hypothesis that apoptosis acts in an antiviral manner to arbovirus infection in mosquitoes [147]. Specifically, inhibition of apoptosis led to decreased infection and dissemination rates, and induction of apoptosis led to greater infection and dissemination. The authors speculate that experimental systemic induction or inhibition of apoptosis prior to virus infection, in contrast to apoptosis being stimulated in individual cells in response to virus infection, resulting in widespread destruction of structural barriers to viral infection/dissemination, may account for this apparent discrepancy. In summary, the variety of studies in mosquitoes investigating the role of apoptosis in response to arbovirus infection strongly implicate the pathway as contributing to the acute antiviral response; and that either the presence or lack of an effective apoptotic response has important consequences for vector competence in mosquitoes.