Introduction Lymphatic filariasis is a disease caused by filarial nematodes including Wuchereria bancrofti and Brugia malayi, transmitted through the bite of infected mosquitoes. These parasites perpetuate socioeconomic instability in developing countries by inflicting crippling morbidity and debilitating stigmatization. The impact of this disease is vast - over 120 million people are infected in 81 endemic countries [1]. In an effort to alleviate morbidity and eliminate transmission of this disease, the Global Program for the Elimination of Lymphatic Filariasis (GPELF) has orchestrated a systematic mass drug administration (MDA) program centered on the repeated dosing of either diethylcarbamazine citrate (DEC) and albendazole or albendazole and ivermectin in areas where the other filarial parasites, Onchocerca volvulus and Loa loa are co-endemic. This strategy has reduced prevalence in many areas [2] but lymphatic filariasis remains a significant global health concern. Many factors contribute to continued transmission, but central is the inadequate portfolio of effective drugs; none of the MDA drugs are effective against all life stages of the parasite with notable inefficacy against adult worms [3]–[5]. This means MDA must be provided annually for the duration of the lifespan of adult parasites. This situation is compounded by gaps in our understanding of mechanisms of drug action and pharmacology – the site of action of DEC is unknown despite being the drug of choice for lymphatic filariasis control for decades, and the filaricidal mechanism of ivermectin at therapeutic concentrations is also equivocal. There is a very real and significant need for additional and more effective antifilarial drugs, and a better understanding of the mode of action of existing drugs [6]. A major obstacle to the rational development of such drugs is the experimental intractability of parasitic nematodes. An example of this complication is RNA interference (RNAi), a reverse genetic tool that allows researchers to rapidly and specifically ‘turn off’ genes of interest. RNAi has fast become a standard tool in rational drug discovery for the identification and validation of potential new drug targets [7], [8]. By suppressing specific genes and examining the resulting phenotype, it is possible to delineate gene function and appraise the potential value of encoded proteins as drug targets. Successful applications of present RNAi protocols to parasitic nematodes have been sporadically reported, limited in their effectiveness and seldom repeated [9]. Some success has been achieved with Nippostrongylus brasiliensis [10], Ascaris suum [11], Trichostrongylus colubriformis [12], Ostertagia ostertagi [13] and Haemonchus contortus [14], [15]. Germane to the study of filarial worms, RNAi has been described in B. malayi [16], [17], Onchocerca volvulus [18], [19] and Litomosoides sigmodontis [20]. The conclusion has been reached, however, that successful RNAi “only works on a limited number of genes, and in some cases the effect is small and difficult to reproduce” [14]. The inability to depend on present RNAi protocols with parasitic nematodes has proved a major stumbling block to the identification and validation of new drug targets, to a better understanding of anthelmintic mode of action, and to advancing our comprehension of parasite biology. The recalcitrance of animal parasitic nematodes to RNAi is perplexing, given that Caenorhabditis elegans, a free-living nematode, and plant parasitic nematodes are readily susceptible to the technique [21]–[28]. One hypothesis advanced to explain this recalcitrance is that because present RNAi protocols employ in vitro approaches including soaking nematodes in an RNAi trigger, feeding nematodes bacteria producing the trigger, or electroporating of the trigger into the parasite, the RNAi trigger is not provided in a manner conducive to systemic gene suppression [29]. Implicit in the use of these protocols is the removal of a parasite from the host and its maintenance in a liquid culture. Therefore these protocols have distinct drawbacks such as difficulty maintaining healthy, viable worms that behave normally in vitro, limitation of use of parasites or life stages for which in vitro culture is defined, and poor efficacy in RNAi trigger delivery methods that can prove lethal to the parasite [30]. The aim of this study was to develop an innovative in vivo approach to RNAi in parasitic nematodes that overcomes the drawbacks associated with present in vitro experimental paradigms. Our approach is based on the filarial nematode B. malayi. We establish a B. malayi infection in an intermediate host, the mosquito Aedes aegypti, and then initiate suppression of parasite genes by injecting an RNAi trigger directly into the mosquito. The mosquito acts as an ideal culture and delivery system, ensuring the RNAi trigger is exposed to healthy, developing parasites. Using this approach we have effectively and quantifiably suppressed expression of Bm-cpl-1, a B. malayi gene encoding a cathepsin L-like cysteine protease. Dramatic aberrant phenotypes accompany this suppression, including a marked retardation of motility, an inhibition of normal parasite migration behavior within the mosquito and impaired parasite growth and development. Suppression is specific; non-target RNAi has no effect on nematode viability or behavior, and the level of gene suppression and extent of the resultant phenotypes suggest this new protocol is more effective than previous methods. The development of an in vivo RNAi protocol to reliably suppress gene expression in filarial worms has great potential for the identification and validation of novel drug targets, and more broadly, to explore parasitic nematode biology and host-parasite interactions. Results A Brugia RNAi trigger rapidly disseminates throughout the mosquito host Our hypothesis is that mosquitoes provide an optimal culture and delivery system for an RNAi trigger targeted to developing Brugia malayi parasites. Healthy, viable, developing parasites are subjected to the RNAi trigger because the parasites undergo growth and development in the mosquito intermediate host. In order to test the extent of dissemination of the RNAi trigger from the site of intrathoracic injection, 150 ng of an equimolar mix of four 3′ Cy 3-labelled Bm-cpl-1 siRNAs was injected into adult Aedes aegypti mosquitoes as described. The dissemination of this RNAi trigger through the mosquito was tracked over 15 d post-injection by periodic microdissection of the mosquito and evaluation of internal fluorescence compared to saline injected controls. The labeled siRNA mix spread rapidly from the site of injection and maximal fluorescence was observed 24 h post-injection (Fig. 1). The intensity of fluorescence slowly decreased until reaching basal levels at five d post-injection after which fluorescence intensity was not appreciably different from control mosquitoes. Our observations closely parallel those of a previous report that describes the spread of 140 ng AlexaFluor 555-labeled siRNA in the mosquito Anopheles gambiae from an injection site to the midgut and pericardial cells 36 h post-injection [31]. Systemic dispersion and persistence of RNAi signal from the site of injection suggests B. malayi larvae are likely to be exposed to the RNAi trigger in our experimental model. 10.1371/journal.ppat.1001239.g001 Figure 1 Dissemination and persistence of intrathoracically injected Cy 3-labelled Brugia malayi Cathepsin-L1 siRNAs in Aedes aegypti. Midgut and Malpighian tubule tissues are shown in light (upper panel) and fluorescence (lower panel) micrographs from 1 to 9 days post-injection (scale bar 100 µm). Brugia gene suppression in vivo is potent and specific Recently it has been shown that B. malayi genes encoding cathepsin L-like enzymes can be suppressed in vitro by soaking adult parasites in culture media containing siRNA [17]. We tested the capacity of our novel methodology to suppress larval stage B. malayi gene expression in vivo by injecting mixed siRNAs specific to the cathepsin L-like Bm-cpl-1 gene directly into Ae. aegypti mosquitoes harboring L3 stage B. malayi parasites. Gene suppression was assayed 48 h post-injection using a semi-quantitative RT-PCR in which the intensity of Bm-cpl-1 amplification in the linear phase of the reaction was compared to an internal B. malayi reference gene (Bm-flp-14) that is expressed stably and at comparable levels to Bm-cpl-1. Control mosquitoes were injected with equal volumes of Aedes physiologic saline. This methodology was optimized to amplify Bm-cpl-1 from a heterogeneous mosquito/parasite total RNA preparation from a single mosquito. Suppression was concentration-dependent because injection of 0.15 ng siRNA did not appear to reduce Bm-cpl-1 transcript levels. However, injection of 15 ng or 1.5 ng of siRNA decreased transcript levels, and injection of 150 ng mixed siRNA into mosquitoes profoundly suppressed Bm-cpl-1 expression; the target parasite gene could not be amplified (Fig. 2). This suppression was also specific; expression of the Bm-flp-14 reference gene was unaffected by siRNA injection and target gene expression was normal in saline-injected controls. 10.1371/journal.ppat.1001239.g002 Figure 2 Concentration-dependent, in vivo suppression of Brugia malayi Cathepsin-L1 (Bm-cpl-1) using siRNA (Top) or dsRNA (Bottom) RNAi triggers. Micrograph shows ethidium bromide stained agarose gel electrophoresis of relative RT-PCR analysis of individual, B. malayi-infected mosquitoes 48 h post-injection of RNAi trigger at 10 d post-infection. Amplified product for the target gene, Bm-cpl-1, is shown above a neuropeptide reference gene (Bm-flp-14). Application of dsRNA is the commonly used method for triggering RNAi in parasitic nematodes and has advantages over siRNA; dsRNA can be generated in-house more quickly than commercially produced siRNAs at lower cost. B. malayi-infected mosquitoes were also subjected to treatment with dsRNA as an RNAi trigger. The effect of dsRNA was concentration-dependent such that injection of 15 ng dsRNA results in Bm-cpl-1 suppression but 1.5 ng dsRNA had no appreciable effect. Injection of 150 ng of dsRNA potently suppressed Bm-cpl-1 transcript abundance and suppression appeared specific, with Bm-flp-14 expression unaffected by dsRNA (Fig. 2). RT-qPCR was used to quantify the level of Bm-cpl-1 suppression relative to two reference genes (Bm-flp-14 and Bm-tph-1) using the efficiency-corrected (EΔΔCq) relative quantification method [32]. PREXCEL-Q software was used to optimize the performance of the RT-qPCR assay; and important data pertinent to PCR efficiency, linear dynamic range and normalization of the assay are documented in Table 1. Bm-tph-1 showed stable Cq values across the experiment and therefore was the most appropriate reference gene for these studies, as shown previously [33]. The suppressive effects of both RNAi treatments were almost identical; injection of 150 ng siRNA reduced Bm-cpl-1 transcript by 83% compared to saline-injected controls (P 95%). Establishing Brugia infection B. malayi microfilaria (mf) infected cat blood was obtained from the University of Georgia NIH/NIAID Filariasis Research Reagent Resource Center. To establish a consistent and repeatable parasitemia, mf were first purified using a filtration protocol [60]. Blood containing the parasites was diluted with phosphate buffered saline (1∶5 ratio, blood:PBS) then syringe filtered through a 0.45 µm Millipore filter. Captured mf were washed three to five times with PBS then a further three to five times with Aedes physiologic saline [61] before centrifugation at 6,800× g for five min. Supernatant was removed and the pelleted mf resuspended in fresh Aedes saline to a concentration of 40 worms per µL. To inoculate mosquitoes, 20 mf were injected as described. Microdissection of the mosquitoes throughout a 14 dpi period confirmed this method established a controlled infection that progressed in a predictable and consistent manner. We also tried a blood feeding approach to establish infection but this produced an inconsistent worm burden that is too variable to reliably assess subsequent gene suppression experiments. siRNA and dsRNA generation and injection Short interfering RNAs (siRNA) targeting a B. malayi cathepsin L-like gene (Bm-cpl-1 AF331035 [34]) were generated commercially (Qiagen, CA) and modified with a 3′-Cy3 fluorophore on the sense strand. The location of each siRNA was optimized using a proprietary algorithm and the sequence of each siRNA is as follows: BmCL1-1, AAGGCTTAGTTTCTTATACAA; BmCL1-2, CCGAATGGAAAGATTATGTAA; BmCL1-3, CAGAAGTGCATTGAAGGAATA; and BmCL1-4, CCGGTATTTACTCCAGTAATA. Equimolar amounts of each siRNA were combined and this mix was used for injection and gene suppression experiments. dsRNA duplexes were generated in-house using a T7 transcription-based approach. A 410 base pair transcription template was polymerase chain reaction (PCR) amplified from a B. malayi L3 stage cDNA library (kindly provided by Dr. S. Williams, Smith College, MA) using gene specific oligonucleotides designed to incorporate a T7 promoter sequence (TAATACGACTCACTATAGGGTACT) at both the 5′ and 3′ ends of the amplicon. For the Bm-cpl-1 template, oligonucleotide sequence was: L1T7dsRNAF 5′ TAATACGACTCACTATAGGGTACTACGGTTACCAAATTC 3′ and L1T7dsRNAR 5′ TAATACGACTCACTATAGGGTACTCGACAACAACAGGTC 3′. The location of this transcription template was carefully chosen so as to exclude the pro region of Bm-cpl-1, a domain with high sequence homology to other cathepsin L family genes, and consequently increase the specificity of this dsRNA duplex. Transcription templates were gel purified and dsRNA duplexes synthesized using the MEGAscript RNAi Kit (Ambion, TX) according to manufacturer's protocols. dsRNA species were quantified with a NanoVue spectrophotometer (GE Healthcare, NJ) prior to use. The timing of siRNA or dsRNA injection into B. malayi-infected mosquitoes coincided with the presence of the parasite stage of interest: to target second larval stage (L2) parasites siRNA or dsRNA were injected five to eight dpi; to target third larval stage (L3) parasites siRNA or dsRNA were injected nine to 12 dpi (and for the lifespan of mosquito) [35]. The mosquitoes were processed to confirm suppression of the target gene, as described below, 48 h post-injection of siRNA or dsRNA. Relative quantitative RT-PCR Brugia infected, RNA-treated and control mosquitoes were cold-anesthetized on ice. Total RNA was extracted from individual mosquitoes using RNAqueous Kit (Ambion, TX) before DNase treatment using the TURBO DNA-free Kit (Ambion, TX) in thin-walled PCR tubes. The RNA was stabilized with RNase Out Inhibitor (Invitrogen, CA) and stored in RNase-free microcentrifuge tubes at 4°C. This RNA was used as a template for a relative semi-quantitative multiplex RT-PCR using the SuperScript III One-Step RT-PCR System with Platinum Taq DNA Polymerase (Invitrogen, CA). The principle of this reaction is to amplify a target gene of interest and compare its intensity with a multiplexed and normalized internal standard during the linear phase of product amplification. A putative neuropeptide encoding gene, Bm-flp-14 (Accession number AI508026) served this role. This gene was chosen as we had previously determined its stable transcript production during the B. malayi L3 stage by PCR (C. Song, unpublished). The oligonucleotide primers used to amplify Bm-cpl-1 were: CPL-1 F 5′ ACAGGGCAATATGACGAGAC 3′ and CPL-1 R 5′ ATCGAAGCAACGTGGCACAT 3′. These primer locations flank the region of Bm-cpl-1 homologous to the dsRNA construct. The oligonucleotide primers used to amplify the Bm-flp-14 internal standard were: FLP-14 F 5′ CTCGTCCACTCTTATCACTG 3′ and FLP-14 R 5′ ACCGCAATGATATACAACATATA 3′. The profile for this PCR was: cDNA synthesis at 50°C for 30 minutes; an initial denaturation phase of 94°C for 2 min; 38 cycles of 94°C for 30 s, 60°C for 30 s, 68°C for 1 min and a final extension phase of 68°C for 5 min. Reactions were visualized on a 1.2% agarose gel containing ethidium bromide. Quantitative RT-PCR Total RNA was extracted from individual mosquitoes and DNase-treated as described above for three replicated RNAi experiments and before addition of RNase Out Inhibitor and storage, each RNA sample was quantified spectrophometrically per a previous report [62]. This RNA served as a template in our RT-qPCR assays using the qScript One-Step Fast RT-PCR Kit with ROX (Quanta BioSciences, MD). Establishing PREXCEL-Q parameters PREXCEL-Q, a qPCR assay development and project management software, was used to establish our RT-qPCR parameters and to determine valid working ranges for all of our samples per target and reference genes. A mixture of the RNA samples was used to determine the optimum template to use while avoiding RT-qPCR inhibition for each of the three targets at concentration ranges of between 0.01 and 0.08 ng/µL for the siRNA experiments and between 0.02 to 0.14 ng/µL for the dsRNA experiments. For subsequent quantitative assessment of transcript abundance, each RNA sample was diluted to 0.06 ng/µL for the siRNA RT-qPCR study and 0.11 ng/µL for the dsRNA RT-qPCR study, with 6 µL of sample used per 25 µL reaction. Primers and probes The target sequence under evaluation in the RT-qPCR study was the B. malayi cathepsin L-like transcript previously described. Two reference genes were used, a neuropeptide-encoding gene Bm-flp-14 also previously described, and Bm-tph-1 (Accession number U80971), a tumor protein homolog-encoding gene that is a proven reference gene for qPCR of Brugia development in mosquitoes [33]. TaqMan minor groove binding (MGB) probes were used in this study to facilitate the use of shorter gene specific primer-probe sets. All probes and primers were designed using Primer Express v. 2.0 software (Applied Biosystems, CA) and synthesized by Applied Biosystems. The primer and probe sequences used are shown in Table 2. 10.1371/journal.ppat.1001239.t002 Table 2 Primer and TaqMan probe sequences for RT-qPCR experiment. Gene Primer sequence (5′ - 3′) Amplicon Size (bp) Bm-cpl-1 ForwardReverseProbe GGTTACGGAACGCATCGAA TGGGTTCCCCAGCTATTTTTAA6FAM-TCACGGTGATTACTGGAT-MGBNFQ 62 Bm-flp-14 ForwardReverseProbe TGGGAAGAGGAAGCATGAATACTT TGCAGCGGGAACTTTGATC6FAM-AGATTTGGTCGTAAGTAGTTG-MGBNFQ 66 Bm-tph-1 ForwardReverseProbe TTGCAACGATATGTTGATCTTCAA ACGAGTCCGACGCAAGCT6FAM-ATGCATTCACAGATGAC-MGBNFQ 62 TaqMan RT-qPCR 25 µL volume reactions were prepared in duplicate for each RNA sample, and 20 µL of this reaction mixture is applied per well on a 96-well plate (using white-well reaction plates, Eppendorf, NY). Individual components of each RT-qPCR reaction were as follows: 6 µL prediluted RNA (as determined by PREXCEL-Q), 6.25 µL 4X One-step Fast Master Mix with ROX, 1.25 µL qScript One-Step Fast RT, 775 nM each primer, 150 nM probe, nuclease-free water to 25 µL. Cycling conditions included an initial cDNA synthesis step of 50°C for 5 min followed by an RT denaturation/Taq activation phase of 95°C for 30 s then 45 cycles of 95°C for 3 s and 58°C for 30 s. Four point standard curves were created for each target (within the ng/µL ranges already specified above) by diluting the RNA sample mixture in each case according to precise, PREXCEL-Q-determined parameters (eight-fold dilution from highest to lowest concentration). No-template control reactions substituted nuclease-free water for RNA, and thermocycling was performed on an ABI GeneAmp 5700 SDS (Applied Biosystems). Quantification cycle (Cq) values were obtained at an appropriate threshold per each target (∼0.1 DRn in all cases), and data were processed using custom Excel files by the efficiency-corrected (EΔΔCq) relative quantification method [32]. Phenotype analysis After confirmation of Bm-cpl-1 suppression, multiple assays were performed to describe worm phenotypes. Each phenotypic assay was performed 14 dpi and at either four or seven d post-injection. Mosquitoes were cold-anesthetized then the wings and legs removed and discarded using a dissecting microscope. The head, thorax and abdomen were partitioned and further dissected to release the parasites. The following characteristics of dissected parasites were observed: (1) Parasite location. In order to be successfully transmitted, these parasites have to actively migrate to the head of the mosquito and vigorously writhe free of the proboscis. Parasite migration through the mosquito was recorded and measured according to escape point from the mosquito body (abdomen, thorax or head). (2) Worm motility. A scoring schema of: one (immobile), two (compromised motility, immobile for stretches of time), three (sluggish, partial movement), four (in motion, some straight segments), or five (all parts of the worm in constant motion) was used to quantify parasite movement in a blind fashion by an independent evaluator. Additional observations of aberrant motility included knotting at one or both ends, paralysis of caudal region and presence of a distinct angular kink were also recorded. (3) Parasite growth and development. Digital images of RNAi and control worms were captured so that length and diameter could be calculated using NIS Elements D 2.30 software (Nikon, NY). (4) Parasite viability. The number of parasites that survived to the infectious stage was recorded so that infection prevalence and mean intensity could be calculated. (5) Mosquito viability. We documented the number of mosquitoes that survived through the development of parasites to the infectious stage because these parasites inflict significant pathology and decrease mosquito survival. Microscopy Nikon Eclipse 50i fluorescence microscope under UV light (EXFO, ON) equipped with a Hy-Q FITC filter set (Chroma, VT). Images were captured using a Digital Sight DS-2Mv camera and NIS Elements D 2.3 software (Nikon, NY). Statistical analysis t-tests were used to analyze the effect of RNAi treatment on gene expression in the RT-qPCR experiments and parasite size, and ANOVA to analyze the effect of RNAi treatment on worm motility based on our one through five blind-scoring schema. Chi square tests were used to analyze the effect of RNAi treatment on all other worm and mosquito behaviors assayed. In all tests, P values ≤0.05 were considered statistically significant.