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Journal of Medical Entomology logoLink to Journal of Medical Entomology
. 2017 Feb 8;54(3):559–567. doi: 10.1093/jme/tjw225

Functional Validation of Apoptosis Genes IAP1 and DRONC in Midgut Tissue of the Biting Midge Culicoides sonorensis (Diptera: Ceratopogonidae) by RNAi

M K Mills 1, D Nayduch 2, D S McVey 2, K Michel 1,3
PMCID: PMC5502902  PMID: 28399198

Abstract

Culicoides biting midges transmit multiple ruminant viruses, including bluetongue virus and epizootic hemorrhagic disease virus, causing significant economic burden worldwide. To further enhance current control techniques, understanding vector–virus interactions within the midge is critical. We developed previously a double-stranded RNA (dsRNA) delivery method to induce RNA interference (RNAi) for targeted gene knockdown in adult Culicoides sonorensis Wirth & Jones. Here, we confirm the C. sonorensis inhibitor of apoptosis 1 (CsIAP1) as an anti-apoptotic functional ortholog of IAP1 in Drosophila, identify the ortholog of the Drosophila initiator caspase DRONC (CsDRONC), and demonstrate that injection of dsRNA into the hemocoel can be used for targeted knockdown in the midgut in C. sonorensis. We observed CsIAP1 transcript reduction in whole midges, with highest transcript reduction in midgut tissues. IAP1knockdown (kd) resulted in pro-apoptotic caspase activation in midgut tissues. In IAP1kd midges, midgut tissue integrity and size were severely compromised. This phenotype, as well as reduced longevity, was partially reverted by co-RNAi suppression of CsDRONC and CsIAP1. Therefore, RNAi can be directed to the midgut of C. sonorensis, the initial site of virus infection, using dsRNA injection into the hemocoel. In addition, we provide evidence that the core apoptosis pathway is conserved in C. sonorensis and can be experimentally activated in the midgut to reduce longevity in C. sonorensis. This study thus paves the way for future reverse genetic analyses of midgut–virus interactions in C. sonorensis, including the putative antiviral properties of RNAi and apoptosis pathways.

Keywords: RNAi, apoptosis, innate immunity, IAP1, DRONC

Culicoides midges are hematophagous insects, able to biologically transmit arboviruses of international importance, such as bluetongue virus (BTV) and epizootic hemorrhagic disease virus (EHDV). Both viruses belong to the genus Orbivirus with segmented dsRNA genomes and cause hemorrhagic fevers with significant economic impact worldwide, owing to livestock production loss and trade restrictions, as well as negatively impacting wildlife populations (Purse et al. 2015). Current control methods against these diseases focus on vaccines, which is currently only available for BTV; livestock management; and vector control (USDA-ARS workshop report 2013). Over the past few years, knowledge of host–virus–vector interactions for orbiviruses has increased significantly (Coetzee and Venter 2015). However, significant knowledge gaps remain, especially with regards to virus–vector interactions and molecular underpinnings that determine Culicoides vector competence to orbiviruses (Drolet et al. 2015).

Besides abiotic factors such as temperature, transmission of arboviruses is influenced by biotic factors within their insect vector (Hardy et al. 1983, Carpenter et al. 2015, Franz et al. 2015). Both EHDV and BTV must successfully infect and escape the midgut, ultimately infect the salivary glands, and escape into the saliva to be passed horizontally to the next host (Mellor 1990, Sieburth et al. 1991, Fu et al. 1999). The midgut is the first physical barrier to infection against ingested pathogens and was confirmed as the major barrier to infection and dissemination against BTV in the principal North American vector, Culicoides sonorensis Wirth & Jones (Diptera: Ceratopogonidae) (Fu et al. 1999).

In insects, innate cellular antiviral mechanisms exist, including RNA interference (RNAi) and programmed cell death (Kingsolver et al. 2013, Lamiable and Imler 2014). RNAi functions as an antiviral pathway by detecting viral double-stranded RNA (dsRNA), cleaving it into 21 = 23 nucleotide short interfering RNA (siRNA), which are then used as templates to detect and degrade viral transcripts by base complementarity (Wilson and Doudna 2013). Not surprisingly, several viral mechanisms exist to counter RNAi, including expression of RNAi suppressors and/or sequestration of viral dsRNA to prevent detection (Zhou and Rossi 2011). Several studies indicate that BTV utilizes the latter strategy, as its dsRNA genome is sequestered to its inner viral capsid and only positive-sense capped RNA transcripts are released into the cytosol (Mohl and Roy 2014). Furthermore, the BTV core has a high affinity for dsRNA, effectively shielding the BTV genome from host detection (Diprose et al. 2002). Nevertheless, a recent study demonstrated that RNAi in cell lines can limit BTV infection when cells are provided with an external RNAi trigger of dsRNA specific to BTV nonstructural protein 1 (Schnettler et al. 2013). Therefore, if RNAi triggers can target viral transcripts in the Culicoides midgut, this immune response could be boosted experimentally to strengthen the initial infection barrier against BTV and possibly EHDV and ultimately integrated into control strategies to reduce BTV and EHDV burden.

Apoptosis, programed cell death, has also been demonstrated to act as an antiviral mechanism in several insect species, including vectors of arboviruses, such as Aedes aegypti (L.) and Culex quinquefasciatus Say (most recently reviewed in Clem 2016). Apoptosis is controlled by a conserved and continuously expressed regulatory network (Zmasek et al. 2013), which upon activation initiates a core proteolytic cascade consistent of cysteine-dependent aspartate-directed proteases of the caspase family. Apoptosis is primarily regulated through the anti-apoptotic Inhibitor of Apoptosis 1 (IAP1) protein, which binds to inactive caspase zymogens, preventing activation and subsequent cell death. Inhibition of IAP1–caspase binding results in the dimerization causing auto-activation of initiator caspases, such as DrosophilaNedd2-like caspase (DRONC; Liu and Clem 2011) and the death-related ced-3/Nedd2-like gene (DREDD; Chen et al. 1998). Initiator caspases then activate executioner caspases through proteolytic cleavage and subsequent dimerization. Executioner caspases mediate proteolysis of multiple cellular proteins and ultimately cause cell death (Zmasek et al. 2013). Putative orthologs of the core apoptosis pathway were detected in the C. sonorensis reference transcriptome (Nayduch et al. 2014), suggesting that the pathway is intact. Apoptosis in animals is correlated with viral infection but not necessarily antiviral (Richard and Tulasne 2012). However, recent work established apoptosis as an antiviral response to arbovirus infection in mosquitoes (Clem 2016); for instance, RNAi silencing of initiator caspase Dronc increased infection prevalence of dengue virus in Ae.aegypti mosquitoes (Ocampo et al. 2013).

The role of apoptosis in insect vectors during orbivirus infection has yet to be elucidated. While BTV and EHDV infections cause apoptosis that contributes to the pathology in ruminant hosts (Mortola et al. 2004, Li et al. 2007, Schwartz-Cornil et al. 2008, Shai et al. 2013), there was no detection of apoptosis induction after BTV infection of three insect cell lines, including the hemocyte-like embryonic KC cell line from C. sonorensis (Mortola et al. 2004). However, comparable in vivo data from infected midges have yet to be performed.

To enable molecular studies on the intrinsic factors underlying vector competence in Culicoides midges, we previously established a dsRNA delivery method for targeted gene knockdown (kd) by RNAi in C. sonorensis (Mills et al. 2015). Injection of dsRNA specific to the putative ortholog of IAP1 (CsIAP1) into the thorax of adult midges resulted in significantly reduced target transcripts and caused increased mortality rates. However, the level of RNAi penetrance in different Culicoides tissues and whether the induction of apoptosis underlies the observed mortality phenotype remained unclear. Here we demonstrate that dsRNA does indeed induce kd in the target midgut tissue. We also confirm that both CsIAP1 and CsDRONC function as anti-apoptotic and pro-apoptotic orthologs of Drosophila IAP1 and DRONC in C. sonorensis.

Materials and Methods

Insect Rearing and Injection of Adult Female C. sonorensis

The C. sonorensis AK strain was reared and injected as described previously (Mills et al. 2015). In all experiments, female midges were injected with 50 nl of total dsRNA (4 µg/µl in RNAse-free water). For single-kd experiments, females were injected only with either dsGFP or dsCsIAP1, while midges used for co-injection experiments were injected with a 1:1 dsRNA ratio with combinations of dsGFP, dsIAP1, and dsDRONC. Throughout all experiments, all midges were fed sugar ad libitum and were not provided a bloodmeal.

DsRNA Synthesis

All dsRNA products were synthesized as previously described. DsGFP (154 bp in length, bp 107–261 in the enhanced GFP gene sequence) and dsCsIAP1 (343 bp in length, bp 381–724 in GAWM01009039) were synthesized using published primers (An et al. 2011, Mills et al. 2015). DsCsDRONC (283 bp in length, bp 394–658 in GAWM01016707) was synthesized using the following primers for first round of PCR: dsCsDRONC _F 5′-TAATACGACTCACTATAGGGGGCAGAGGTTGATAAAGAGA GA-3′; dsCsDRONC _R 5′-TAATACGACTCACTATAGGGGTTT ACCAATCAAATGTGT-3′. Purified dsRNA was resuspended in RNase-free water at a final concentration of 4 μg/μl. CsIAP sequences can also be viewed as previously described (Mills et al. 2015).

Sample Collection, Total RNA Extraction, and cDNA Synthesis

Whole midges were frozen and stored at −80 °C prior to RNA extraction. For midgut and remaining carcass collection, midguts were dissected in ice-cold phosphate-buffered saline (PBS) on ice, immediately flash-frozen in liquid nitrogen, and stored at −80 °C at 2 and 5 d post injection (dpi). For single-kd experiments, RNA from frozen midges and/or midguts and carcasses (n = 20–40 per treatment and replicate) was extracted using the RNeasy Micro Kit standard protocol (Qiagen, Valencia, CA, USA) and eluted in 10 µl RNAse-free water (Fisher Scientific, Waltham, MA, USA). For all co-injection experiments, RNA was extracted from frozen midges (n = 20) using a phenol:chloroform protocol as described previously (Mills et al. 2015).

Culicoides sonorensis cDNA was synthesized from 100 ng of purified total RNA with the iScript cDNA synthesis kit (Bio-Rad, Hercules, CA, USA).

RT-qPCR

For single-kd experiments, female midges and/or midguts (n = 20–40 per treatment and replicate) were collected 2 and 5 dpi, and RT-qPCR was performed as described previously using elongation factor 1b as the reference gene (Mills et al. 2015). Females (n = 20 per treatment group) were collected at 5 dpi for double-kd (dkd) experiments, and RT-qPCR analyses were performed using published CsIAP1 primers (Mills et al. 2015) and the following CsDRONC primers: CsDRONC_F 5′-CTGAACATGCACATGATACC-3′; CsDRONC_R 5′-CTGAGCAAACTTGTTTGTATTC-3′.

RT-qPCR analyses for co-injection experiments were performed using iQ SYBR Green Supermix (Bio-Rad) according to the manufacturer’s protocol with 6 μl undiluted cDNA as template for each 20 μl volume reaction. All RT-qPCR analyses used the same amplification protocol (Mills et al., 2015) and were performed with three technical replicates. Biological replicates (n) for kd (2 dpi, n = 4; 5 dpi, n = 3) and dkd (n = 3) experiments were performed.

Fold changes between treatments were calculated using the Pfaffl method (Pfaffl 2001), with a primer efficiency of E = 1.79 for CsDRONC (Supp. Fig. 1 [online only]), using Elongation factor 1b as the reference gene and untreated (UT) controls as the calibrator condition (Mills et al. 2015). Expression data between kd treatments and control were compared statistically using unpaired t-tests, while co-injection expression data were analyzed using one-way ANOVA, followed by Tukey’s multiple comparison post-test. All statistical analyses were performed using GraphPad Prism software version 6 (GraphPad Software Inc., La Jolla, CA, USA).

Immunostaining

Midgut tissues were dissected into 1× PBS and immediately fixed in 4% paraformaldehyde in PBS for 30 min at room temperature (RT). Tissues were washed in PBS with 0.3% Triton X-100 (PBST) four times at RT. Midguts were transferred into blocking buffer (1% bovine serum albumin in PBST) and incubated for 30 min at RT. Tissues were incubated with monoclonal rabbit anti-cleaved caspase-3 (Asp175) (D3E9) primary antibody (1:500 in PBST, Cell Signaling Technology, Catalog# 9579, Danvers, MA, USA) overnight at 4 °C. Midguts were washed four times for 30 min each in PBST at RT and subsequently incubated with Alexa594-conjugated goat anti-rabbit secondary antibodies (1:1000 in PBST, Invitrogen, Carlsbad, CA, USA) for 2 h at RT. Tissues were washed thrice for 30 min in PBST and then incubated with DAPI (5 ng/ml in PBST) for 30 min at RT. Midguts were mounted in Vectashield mounting medium containing DAPI (Vector Laboratories, Burlingame, CA, USA), sealed with nail polish, and stored at 4 °C until imaging.

Confocal Microscopy and Measurement of Midgut Area

Representative images were acquired using a LSM700 Confocal Microscope (Zeiss, Oberkochen, Germany) with identical laser and microscope settings between samples. Images were processed in ZEN 2010 software (Zeiss, Oberkochen, Germany), and figures were prepared using Photoshop and Illustrator software (Adobe Systems, San Jose, CA, USA). The area of each midgut (n = 6 per treatment) was determined by using the contour tool (Supp. Fig. 2 [online only]) of the ZEN 2010 software (Zeiss, Jena, Germany). Data passed the KS normality test and were analyzed statistically using two-way ANOVA (time, treatment), followed by Tukey’s multiple comparison post-test. All statistical analyses were performed using GraphPad Prism software version 6 (GraphPad Software Inc., La Jolla, CA, USA).

Caspase Activity Assay

Assays were performed as described previously (Wang et al. 2012). Midges were collected 2 dpi, flash-frozen, and stored at −80°C until further analysis. Frozen midges were homogenized with a motorized pestle in 30 μl lysis buffer (25 mM Tris-HCl, pH 8, 50 mM NaCl) and subjected to four freeze–thaw cycles using liquid nitrogen to free cellular contents. Resulting homogenates were centrifuged for 10 min at 16,000 × g in 4 °C. Supernatants were transferred to new tubes and stored at −80°C until further analyses. Protein concentration was determined using the Pierce BCA Assay Protein Assay Kit standard protocol (Fisher Scientific, Waltham, MA, USA) and adjusted to 50 μg protein per sample in a total volume of 48 μl lysis buffer. Samples were added to 50 μl reaction buffer (100 mM HEPES buffer, pH 7.4, 2 mM DTT, 0.1% CHAPS, 1% sucrose) with 200 μM Ac-DEVD-AFC caspase substrate (MP Biomedical, Santa Ana, CA, USA) and incubated at 37 °C for 15 min. Caspase activity was monitored for 15 min to determine caspase activity (Δ relative fluorescent units per minute). Experiments were performed in four biological replicates using n = 10 midges per replicate, and caspase activity for each sample was measured in duplicate. Data were analyzed statistically using Kruskal–Wallis, followed by Dunn’s multiple comparison post-test.

Midgut Classification

Midgut morphology was deemed “altered” if dissections revealed one or more of the following morphological changes: 1) no clear midgut was identifiable, 2) midgut size was visibly reduced by at least one-third compared with the average size of midguts observed in the control group, and 3) loss of posterior to anterior tapering of the midgut. If none of the morphological changes was observed, midguts were classified as “normal.” Experiments were performed with four biological replicates using 30 to 40 midguts per replicate. Data were analyzed statistically using the contingency table chi-square test, followed by Tukey’s multiple comparison post-test.

Sequence Alignment and Phylogenetic Analysis

Amino acid sequence alignments were performed in MEGA 6.0 (Tamura et al. 2013) using Clustal W (Larkin et al. 2007) with default settings. To reconstruct the phylogenetic relationships, maximum likelihood trees were generated using MEGA 6.0, with the following settings: bootstrap method with 1,000 iterations, Jones–Taylor–Thornton substitution model, complete gap deletions, and nearest-neighbor-interchange.

Survival Analysis

After dsRNA injection, survival was monitored every 24 h until all midges within the experiment were dead. Survival experiments were performed with four biological replicates using 100 midges per replicate. Resulting data were analyzed and graphed using Kaplan–Meier and compared using the Log-rank (Mantel–Cox) test and hazard ratios. Median survival data were evaluated statistically with the Kruskal–Wallis test, followed by Dunn’s multiple comparison post-test. Percent daily mortality was defined as the percentage of dead midges out of the total population between two consecutive days and evaluated statistically using Kruskal–Wallis, followed by Dunn’s multiple comparison post-test.

Results

Delineating RNAi Knockdown Persistence of CsIAP1 in Midgut and Carcass Tissues

To determine the efficacy and longevity of RNAi kd following hemocoel injection of dsRNA, we examined the time period and midgut penetrance of transcript knockdown (kd). Adult female midges were injected with 80 ng of dsRNA specific to CsIAP1 (dsCsIAP1). Whole body or dissected midguts and remaining carcasses were collected for reverse transcription (RT)-quantitative(q)PCR analysis at 2 and 5 dpi (Fig. 1). Similar to previous findings (Mills et al. 2015), CsIAP1 transcripts were significantly reduced by 30% in whole dsCsIAP1 kd (IAP1kd) midges at both time points when compared with dsGFP-injected controls (Unpaired t-test; day 2: df = 4, t = 3.098, P = 0.021; day 5: df = 4, t = 4.301, P = 0.013; Fig. 1A and D). Transcript reduction levels were even higher when midge midguts were examined. Midguts dissected from IAP1kd midges exhibited a statistically significantly transcript reduction of 50% at 2 dpi, while the corresponding carcasses showed statistically significant kd levels of 45% (Unpaired t-test, midguts t = 6.132, df = 5, P = 0.002; carcasses t = 3,568, df = 5, P = 0.016; Fig. 1B and C). In addition, IAP1kd midguts at 5 dpi showed a 66% transcript reduction, which was not statistically significantly different compared with dsGFP-treated midguts, owing to high variation between the biological replicates (Fig. 1F; Unpaired t-test, t = 2.55, df = 4, P = 0.063). Moreover, the small 20% CsIAP1 transcript reduction in carcasses from dsCsIAP1-injected midges at 5 dpi was also not statistically significant (Fig. 1F; Unpaired t-test, t = 1.418, df = 4, P = 0.229).

Fig. 1.

Fig. 1

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Temporospatial analysis of CsIAP1 transcript levels after dsRNA injection. Graphs depict mean transcript levels at 2 (A–C) and 5 (D–E) dpi. Graphs show relative expression in whole midges (A and D), carcass (B and E), and corresponding midguts (C and F). All data were calibrated relative to UT controls (dotted line), and quantitative RT-PCR results were calculated using elongation factor 1b as the reference gene. Data are presented as mean ± SEM from multiple biological replicates (2 dpi, N = 4; 5 dpi, N = 3). Statistically significant differences between samples from dsIAP1 and dsGFP-treated midges are denoted by different letters (Unpaired t-test, P < 0.05).

Overall, dsRNAs injected into the hemocoel of C. sonorensis females function as an efficient trigger of RNAi as early as 2 dpi and persist for at least 5 dpi. These data further demonstrate that uptake of the RNAi trigger is successful in midgut tissues, as targeted transcript reduction was even more efficient in the midgut as compared with the remainder of the midge body.

IAP1kd Induces Increased Caspase Activity and Apoptosis

Based on phylogenetic evidence supporting CsIAP1 as an evolutionarily conserved inhibitor of initiator and executioner caspases in the apoptosis pathway (Hay et al. 1995, Tenev et al. 2007, Mills et al. 2015), we hypothesized that IAP1kd would cause an increase in overall caspase activity and, consequently, apoptosis. To test this hypothesis, caspase activity assays were performed on whole midges at 2 dpi using the substrate of human executioner caspase 3, which can be cleaved by insect caspases DRONC and DrICE (Hawkins et al. 2000, Wang et al. 2012). Extracts from IAP1kd midges showed a statistically significant increase in caspase activity when compared with untreated (UT) and dsGFP-injected controls (Kruskal–Wallis: H (2) = 7.423, P = 0.013; Dunn’s multiple comparisons post-test, P < 0.01; Fig. 2).

Fig. 2.

Fig. 2

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In vitro caspase activity of whole female midge extracts. Midges were collected at 2 dpi and pooled (n = 10 per biological replicate) for caspase activity assessment. Caspase activity was determined using the caspase substrate Ac-DEVD-AFC. Statistical significance is denoted by asterisks (Kruskal–Wallis, followed by Dunn’s multiple comparisons test, P < 0.05). Data are presented as the median with interquartile range from four biological replicates.

Given IAP1kd was most pronounced in the midgut, we used an independent experimental method to demonstrate that increased caspase activity could be observed specifically in this tissue. Midguts were dissected at 2 dpi and stained with anti-cleaved human executioner caspase-3 antibody (Fig. 3) that specifically binds activated caspase-3 in evolutionarily distant dipteran species (Buchon et al. 2009, Heerman et al. 2015). IAP1kd resulted in strong staining of midgut tissues with anti-cleaved caspase-3 antibodies (Fig. 3C and D), while midguts dissected from dsGFP-injected (Fig. 3G and H) and UT (Supp. Fig. 3 [online only]) midges showed virtually no staining. This staining signal was specific to the anti-cleaved caspase-3 antibody, as no staining was observed in midguts from all treatments that were only incubated with the secondary antibody (Supp. Fig. 4 [online only]). In addition to strong anti-cleaved caspase-3 staining, several epithelial cells in IAP1kd-midguts also contained fragmented nuclei (Fig. 3B and Supp. Fig. 5 [online only]), which are a hallmark of apoptosis (Umansky 1982). While antibody staining and nucleus fragmentation varied in intensity and region between midguts isolated from different midges, we did not identify specific areas of the midgut that were targeted preferentially. Nucleus fragmentation was not observed in midguts dissected from dsGFP-injected or UT midges. In summary, these results demonstrate that CsIAP1 functions as a caspase inhibitor and attenuates apoptosis in C. sonorensis.

Fig. 3.

Fig. 3

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Active caspase staining of midge midguts. (A and E) Representative images of whole midge midgut at 2 dpi via DAPI stain (blue). Scale bar, 100 nm. (B–D) DsCsIAP1 injection led to increased fluorescence related to anti-cleaved caspase-3 antibody staining (red) compared with UT (Additional file 1: Suppl. Fig. 1 [online only]) and dsGFP (F–H) controls. Images (B–D and F–H) were taken from midgut location denoted by white squares. Scale bar, 20 nm.

Increased Caspase Activity Due to IAP1kd Negatively Affects Midgut Morphology

Next, we investigated the possible influence of increased caspase activity and apoptosis after dsIAP1 injection on the overall morphology of the midgut. Visual examination of midguts from IAP1kd midges at 2 and 5 dpi revealed strong size differences between the treatment group and controls (Fig. 4A–D). To quantify these changes, we determined average midgut size by measuring the area of individual midguts from each treatment group by microscopy at 2 dpi (Supp. Fig. 2 [online only]). The average area of midguts was significantly affected by IAP1kd and did not vary according to time (two-way ANOVA, Treatment: F(2, 34) = 19.07, P < 0.0001; Time: F(1, 34) = 0.138, P = 0.712: Fig. 4E). On average, midgut size of IAP1kd midges was reduced by more than 50% when compared with UT and dsGFP-injected controls at 2 dpi (Tukey’s post-test, P < 0.01, Fig. 4E).

Fig. 4.

Fig. 4

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Altered midgut morphologies in IAP1kd midges. (A–D) Representative images depicting midge midguts with DAPI stain (blue). UT and dsGFP-injected midguts (A and B) are considered to have “normal” morphology, whereas dsCsIAP1-injected midguts (C and D) are deemed “altered,” based on criteria detailed in Materials and Methods section (E) The mean area per midgut from UT and dsGFP-injected controls were compared with IAP1kd midges at 2 and 5 dpi. Midgut areas from UT and dsGFP-injected midges are shown for 2 dpi only (for numbers at 5 dpi, see Additional file 5: Supp. Table 1 [online only]). Statistically significant differences in midgut area are denoted by different letters (Two-way ANOVA; Tukey’s post-test, P < 0.05). Data are presented as the mean ± SEM (n = 6 midguts per treatment). (F) Graph depicts the number of midguts observed with either the normal or altered morphology at indicated time points. Data are presented as the combined number of midguts from five biological replicates. Scale bar, 100 nm.

In addition to the significant size decrease, we frequently observed additional morphological changes decreasing overall midgut tissue integrity. Midguts from IAP1kd midges were more fragile, and some midguts lacked the typical distinction between narrow anterior and wider posterior midgut sections (Fig. 4C and D). Furthermore, in some cases, the alimentary canal could only be identified through the presence of a crop and Malpighian tubules and seemingly did not contain a discernable midgut. To quantify these changes after IAP1kd, we determined the frequency of altered midgut morphologies in the treatment versus control groups. Any of the altered morphologies described above were only observed in IAP1kd midges, and nearly two-thirds of all IAP1kd midguts exhibited one or more morphological alterations (Fig. 2F; Supp. Table 1 [online only]; Contingency table, χ2 = 426.4, df = 4, P < 0.0001). The frequency of altered midgut morphology in the IAP1kd midges remained equally high at 2 and 5 dpi (Chi-square test; χ2 = 1.914, df = 1, P = 0.1666). These results show that the presence of IAP1kd in the midgut of C. sonorensis leads to altered tissue morphology that disturbs and ultimately destroys its structural integrity.

The Initiator Caspase DRONC is Required for IAP1kd-Induced Phenotypic Changes

To determine whether initiator caspases are conserved and can be targeted in C. sonorensis, we next looked for an ortholog of the initiator caspase DRONC and tested if it was required for the IAP1kd-induced phenotypes. Specifically, we hypothesized that co-injection of two dsRNAs targeting CsDRONC and CsIAP1 would revert the IAP1kd-induced phenotypes.

To identify putative orthologs of initiator caspase(s), we mined the C. sonorensis transcriptome data (Nayduch et al. 2014) by Blast analysis using reference sequences from ImmunoDB (http://cegg.unige.ch/Insecta/immunodb, Supp. Fig. 6 [online only]). One sequence with accession number GAWM01016707 was identified by best reciprocal Blast hit using Ae. aegypti DRONC (AAEL011562, Supp. Fig. 6 [online only]). Sequence alignment of the deduced amino acid sequence of GAWM01016707 with known initiator caspases from Ae. aegypti, Anopheles gambiae Giles, and Drosophilamelanogaster confirmed the presence of the critical residues required for caspase function (Fuentes-Prior and Salvesen 2004, Bryant et al. 2010, Supp. Fig. 7 [online only]). Next, we used the sequence alignment to reconstruct the phylogenetic relationship of GAWM01016707 with the known Dipteran initiator caspases by maximum likelihood. The tree topology of the initiator caspases mirrored the phylogenetic relationships of these Dipteran species (Wiegmann et al. 2011), and identified GAWM01016707 as the ortholog of the initiator caspase DRONC and not DREDD in C. sonorensis (Supp. Fig. 8 [online only]). We therefore named this gene CsDRONC.

To functionally confirm CsDRONC as an initiator caspase, we performed experiments, where midges were co-injected with 40 ng of each dsRNA specific to CsIAP1 and CsDRONC (dsIAP1/dsDRONC), which is half the dose previously used for IAP1kd. For single-dsRNA treatment controls, either dsRNA was replaced with equal amount of dsGFP (dsGFP/dsIAP1, dsGFP/dsDRONC). Knockdown levels were assessed for each transcript using RT-qPCR. Average transcript reduction in whole midges was specific to the dsRNA injected (Supp. Fig. 9 [online only]), and reduction levels were similar between dsIAP1/dsDRONC-injected midges and single-dsRNA treatment controls, respectively. However, transcript reduction of neither IAP1 nor DRONC was statistically significant as compared with dsGFP-treated controls (One-way ANOVA: CsDRONC, P = 0.325; CsIAP1, P = 0.4661), and we observed substantial variation of CsIAP1 and CsDRONC transcript levels observed in the dsGFP-injected controls. Interestingly, using a dose of 40 ng of dsCsIAP1 led to an average transcript reduction of 25% in whole midges at 5 dpi, as compared with the 40% reduction observed previously using an 80-ng dsRNA dose (Supp. Fig. 9 [online only], Fig. 1A).

We next assessed the effect of dsIAP1/dsDRONC injection on decreased tissue integrity of midguts induced by IAP1kd. Nearly three-quarters of all midguts from dsGFP/dsIAP1-injected midges showed one or more signs of altered morphology (Fig. 5, Supp. Table 2 [online only]). Midgut morphology was partially restored in dsIAP1/dsDRONC-injected midges (Contingency table, df = 1, χ2 = 9.889, P = 0.0017), demonstrating that midgut damage due to IAP1kd is at least partially dependent on CsDRONC.

Fig. 5.

Fig. 5

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CsDRONC is required for IAP1kd-mediated changes in midgut morphology. Graph depicts the number of midguts observed with either normal or altered morphology at 5 dpi. Data are presented as the total midgut number combined from six biological replicates. Statistically significantly different treatment groups are denoted by P value. (Contingency table, P = 0.0017).

We previously reported that IAP1kd significantly accelerated mortality in adult female midges (Mills et al. 2015). We confirmed our previous results and also observed this phenotype after dsGFP/dsCsIAP1 injection (Fig. 6, Supp. Fig. 10 [online only]), with median survival reduced by 50% (Kruskal–Wallis, H(4) = 11.68, P = 0.0199; Dunn’s post-test, P < 0.05 when comparing dsGFP/dsCsIAP1 and dsGFP/dsGFP-injected treatments). In addition, daily mortality doubled 2 d after dsGFP/dsCsIAP1 as compared with dsGFP/dsGFP-injected controls (Kruskal–Wallis, H(4) = 15.15, P = 0.004; Dunn’s post-test, P < 0.05; Fig. 6B and C, Supp. Fig. 10 [online only]). However, the survival phenotype observed in these midges injected with 40 ng of dsIAP1 was dampened as compared with our previous results, which used a dsIAP1 dose of 70 ng (Supp. Fig. 11 [online only]).

Fig. 6.

Fig. 6

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CsDRONC partially mediates IAP1kd-induced lethality. (A) Survival curves of midges after injection of dsRNA specific to one gene of interest (IAP1kd or DRONCkd), both genes of interest (IAP1/dsDRONCdkd), and dsGFP controls. Additional graphs depict corresponding median survival (B) and daily mortality rates at 2 dpi (C). Data were combined from four biological replicates (shown individually in Supp. Fig. 10 [online only]). Different letters represent statistically significant differences (Kruskal–Wallis, Dunn’s multiple comparison post-test, P < 0.05). Median survival and daily mortality rates are represented as median with interquartile range.

Injection of dsIAP1/dsDRONC partially reverted the dsGFP/dsIAP1-induced mortality phenotype (Fig. 5A): Median survival of dsIAP1/dsDRONC-injected midges was intermediate to dsGFP/dsIAP1 and dsGFP/dsGFP-injected controls, respectively, and not statistically significantly different from either of these treatment groups (Kruskal–Wallis, H(4) = 11.68, P = 0.0199; Dunn’s post-test, P < 0.05). Phenotypic rescue was especially pronounced early in the time course, as the dsGFP/dsIAP1-dependent mortality at 2 dpi was fully reverted by dsIAP1/dsDRONC injection (Kruskal–Wallis, H(4) = 15.15, P = 0.004; Dunn’s post-test, P < 0.05; Fig. 6B and C, Supp. Fig. 10 [online only]). Thus, both IAP1kd phenotypes decreased midgut tissue integrity, and increased mortality rates seem dependent on the initiator caspase CsDRONC.

Discussion

The overarching hypothesis tested by this study was that apoptosis is inducible in the midgut of adult female C. sonorensis by injecting an RNAi trigger into the hemocoel. Our data demonstrate that injection of long dsRNAs against IAP1 into the thorax indeed triggers apoptosis and ablates the midgut tissue. Our work also extends the use of RNAi in this important vector species, as we demonstrate that simultaneous injection of dsIAP1 and dsDRONC partially reverts the phenotype induced by IAP1kd.

We previously reported that dsCsIAP1 injection resulted in increased mortality of C. sonorensis (Mills et al. 2015). This study now provides clear evidence that this shortened life span is due to increased apoptosis and tissue damage, especially in the midgut. Multiple methods to detect apoptosis are available, including the presence of nucleus fragmentation (Umansky 1982). In dipterans, caspase activity assays (Wang and Clem 2011, Wang et al. 2012) and immunofluorescence analyses with anti-cleaved caspase-3 antibody (Buchon et al. 2009, Heerman et al. 2015, Dong et al. 2016) were utilized previously to detect apoptosis. Indeed, we observed caspase-mediated apoptosis after IAP1kd, by all three methods, verifying CsIAP1 (GAWM01009039) as the ortholog of IAP1 in C. sonorensis.

CsIAP1 transcript reduction was pronounced in midge midguts and presumably the cause for the observed morphological changes. This midgut damage was linked to increased caspase-dependent apoptosis, as observed through the presence of fragmented nuclei and anti-cleaved caspase-3 staining, indicating a large number of midgut cells undergoing apoptosis at 2 dpi. The increase in apoptosis resulted in the loss of midgut epithelial cells, causing reduced midgut size and increased fragility observed in IAP1kd midguts. Similar to results in D. melanogaster with the kd of hand (Lo et al. 2007), these altered midgut morphologies were linked to increased mortality, as these alterations very likely prevent the uptake of essential nutrients, resulting in starvation.

In addition to confirming the functional ortholog of IAP1, we identified the initiator caspase DRONC in C. sonorensis. As activation of DRONC was previously demonstrated to be essential for apoptosis (Daish et al 2004), we hypothesized the simultaneous kd of CsDRONC and CsIAP1 would inhibit IAP1kd-phenotypes. We mined the C. sonorensis transcriptome (Nayduch et al. 2014) and, based on phylogenetic analyses, identified GAWM01016707 as the ortholog of the initiator caspase DRONC. Co-injection of dsIAP1 and dsDRONC required us to lower the dose for each dsRNA, which led to transcript kd levels just outside of statistical significance. In addition, the protein half-life of DRONC is about 4.3-fold higher than that of IAP1 in D. melanogaster, which may also contribute to the incomplete rescue of IAP1kd-phenotypes (Yoo et al. 2002). Future experiments are needed to test whether these limitations can be overcome using sequential injections of individual RNAi triggers. Nevertheless, the results from the reverse genetic experiments strongly suggest that CsDRONC is required for IAP1kd-phenotypes, as simultaneous injection of dsIAP1 and dsDRONC partially rescued the IAP1kd-induced mortality and altered midgut morphology phenotypes. Together, these data demonstrate CsDRONC (GAWM01016707) as an initiator caspase in the midge and the conserved nature of the core apoptosis pathway in C. sonorensis.

In addition to further characterizing apoptosis in C. sonorensis, this study extends our understanding of RNAi in this species. First, our data demonstrate that long dsRNAs injected into the hemocoel provide the trigger for RNAi in multiple tissues, as transcript reduction was not only detected in dissected midguts but also in the remaining carcasses of adult females. Second, the results from the co-injection experiments show that the expression of at least two genes can be targeted simultaneously. Transcript reduction was specific to the dsRNA injected, as injection of dsCsDRONC reduced transcript levels of its target by up to 30%, while not affecting CsIAP1 transcript levels, and vice versa. Although transcript level reduction was not statistically significantly different from control levels, the partial rescue of IAP1kd-mortality and midgut phenotypes provides strong evidence that Culicoides midges are able to utilize multiple dsRNA triggers for simultaneous transcript kd.

Finally, the analysis of transcript levels after dsRNA injection revealed that RNA interference in C. sonorensis mainly, if not solely, relies on environmental RNAi. The hallmark of environmental RNAi is that the RNAi trigger is taken up by the cells individually and, in contrast to systemic RNAi, is not amplified or propagated between cells (Huvenne and Smagghe 2010). As a result, kd levels can be titrated, where RNAi trigger dose correlates positively with transcript kd levels (Whyard et al. 2009). We observed similar results in our co-injection experiments. Midges in these experiments were injected with half the dsRNA dose per target transcript than in single-kd experiments owing to viscosity and injection volume limitations. This lowered dsRNA dose resulted in less pronounced transcript reduction and lessened severity of the mortality phenotype in dsGFP/dsIAP1-injected midges when compared with our previous single-kd experiments (Mills et al. 2015). Similar results were obtained in other Dipteran species, suggesting that environmental RNAi is a trait common to all Dipterans, and the ability to propagate the RNAi trigger was lost at the base of this insect order (Scott et al. 2013).

RNAi has revolutionized our ability to study vector–pathogen interactions in non-model Dipteran insects, including mosquitoes (Blandin et al. 2002, Xi et al. 2008), tsetse flies (Wang et al. 2009), and sand flies (Sant’anna et al. 2009, Coutinho-Abreu and Ramalho-Ortigao 2010). Similar to these vector species, the results presented here pave the way for future reverse genetic analyses of midgut–virus interactions within the biting midge C. sonorensis, including the putative antiviral properties of RNAi and apoptosis pathways.

Supplementary Material

Supplementary Data
Click here for additional data file. (608.3KB, pdf)

Acknowledgments

Thanks go to Dr Lee Cohnsteadt, Mr James A. Kempert, and Mr William E. Yarnell at the Arthropod-Borne Animal Disease Research Unit, USDA-ARS, Manhattan, KS, for provision of all adult C. sonorensis midges and housing materials used in this study. Additional thanks go to Matthew Herman and Dr Marcelo Ramalho-Ortigao at Kansas State University, Manhattan, KS, for the use of the anti-cleaved caspase-3 antibodies and the Confocal Microscopy and Molecular Biology Cores funded by KSU-CVM for use of their facilities. Research reported in this publication was supported by NIAID of the National Institutes of Health under award number R01 AI095842, which is an NIH grant from the Institute of Allergy and Infectious Diseases (K.M.) and by Specific Cooperative Agreements 58-5430-1360 and 58-5430-4-022 (K.M.) from the USDA Agricultural Research Service. The content is solely the responsibility of the authors and does not necessarily represent the official views of the funding agencies. This is contribution no. 16-312-J from the Kansas Agricultural Experiment Station.

Supplementary Data

Supplementary data are available at Journal of Medical Entomology online.

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Supplementary Materials

Supplementary Data
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