Nairovirus RNA Sequences Expressed by a Semliki Fo
http://www.100md.com
病菌学杂志 2005年第14期
Unité de Génétique moléculaire des Bunyaviridés, Institut Pasteur, Paris, France
Laboratoire de virologie, CRSSA, Centre de Recherche Emile Pardé, Grenoble, France
University of Minnesota, Department of Entomology, St. Paul, Minnesota 55108
ABSTRACT
We report the successful infection of the cell line ISE6 derived from Ixodes scapularis tick embryos by the tick-borne Hazara virus (HAZV), a nairovirus in the family Bunyaviridae. Using a recombinant Semliki Forest alphavirus replicon that replicates in these cells, we were able to inhibit replication of HAZV, and we showed that this blockage is mediated by the replication of the Semliki Forest alphavirus replicon; the vector containing the HAZV nucleoprotein gene in sense or antisense orientation efficiently inhibited HAZV replication. Moreover, expression of a distantly related nucleoprotein gene from Crimean-Congo hemorrhagic fever nairovirus failed to induce HAZV silencing, indicating that the inhibition is sequence specific. The resistance of these cells to replicate HAZV correlated with the detection of specific RNase activity and 21- to 24-nucleotide-long small interfering RNAs. Altogether, these results strongly suggest that pathogen-derived resistance can be established in the tick cells via a mechanism of RNA interference.
INTRODUCTION
Pathogen-derived resistance (PDR) is a genetic mechanism that reduces host or cell susceptibility to virus infection and has been illustrated in several viral infection models. It is caused by expression of certain pathogen-derived genes interfering with infection by a closely related pathogen. It was first described by Sanford and Johnston in 1985 (35) and has been shown to be effective against arboviruses with a positive or negative RNA genome like dengue virus, La Crosse virus, or Rift Valley fever virus (7, 14, 30). Resistance was demonstrated after in vitro or in vivo expression of a region of the viral genome. The cells or mosquitoes expressing these viral sequences became resistant to infection with the homologous virus (8, 27). To induce PDR, the Sindbis alphavirus vector was utilized because its genome could be manipulated through an infectious cDNA (28, 32). The replicon of the Semliki Forest virus (SFV), another alphavirus, was also used to interfere with the replication of Rift Valley fever virus in mosquito cells (7). Resistance to infection was dependent on RNA but not protein expression. PDR was also induced by artificial, inverted-repeat RNAs homologous to the targeted dengue virus. Plasmid-transfected mosquito cells generated small interfering RNAs (siRNAs) that triggered silencing of homologous viral gene expression (2, 34).
The concept of PDR and RNA interference (RNAi) has been relatively well documented in mosquitoes and mosquito cells, but very little is known in ticks and tick cells for the tick-borne viruses. Among bunyaviruses, the genus Nairovirus comprises 34 tick-borne viruses classified into seven serogroups. The main representatives are the widely distributed human pathogen Crimean-Congo hemorrhagic fever virus (CCHFV), which causes severe and often fatal hemorrhagic fever in humans (16), and the Nairobi sheep disease virus, which circulates in Africa and Asia (23) and induces acute and fatal hemorrhagic gastroenteritis in sheep and goats. Although nairovirus infection in vertebrate cells has been documented (for reviews, see references 36 and 38), there are only a few reports concerning nairovirus infection in their tick vector or in tick cells (31).
In this study, we used the ISE6 tick cell line, derived from Ixodes scapularis (25), and show that these cells supported the efficient replication of Hazara virus (HAZV), a nairovirus isolated from Ixodes ticks collected in Pakistan (5, 11, 18). Because it is not pathogenic for humans, it was used as a model for the very pathogenic CCHFV, classified in the same serogroup (13). We addressed the question of whether PDR against HAZV can be established in tick cells. This was achieved by expressing HAZV genome sequences via infection with specific recombinant SFV (rSFV). In these cells, PDR correlated with the detection of sequence-specific RNase activity and siRNAs, which strongly suggest the existence of a mechanism of RNAi. Altogether, these results establish the feasibility for the control of tick-borne viruses in their vector.
MATERIALS AND METHODS
Virus and cells. BHK-21 cells were grown in Glasgow minimum essential medium supplemented with 5% fetal calf serum, 10% tryptose phosphate, 10 mM HEPES, and antibiotics. Vero E6 cells were grown in Dulbecco's minimum essential medium supplemented with 10% fetal calf serum and penicillin/streptomycin. Cell line ISE6 was established from embryos of the black-legged tick Ixodes scapularis and grown as previously described (25).
Hazara virus (strain JC280) and Dugbe virus (strain IbAr 1792) were provided by E. Gould (Natural Environment Research Council, Oxford, United Kingdom). Stocks grown by infecting BHK-21 cells at a multiplicity of infection (MOI) of 0.01 reached 106 to 107 focus-forming units (FFU) per ml. Virus was titrated by focus formation in Vero E6 cells. At day 5 postinfection, the agarose overlay was removed, cells were fixed with methanol, and infected foci were visualized after staining with HAZV antibody (dilution, 1:250) revealed by peroxidase-labeled anti-mouse antibody (dilution, 1:1,000).
Recombinant Semliki Forest virus. SFV replicons carrying partial or complete sequences of the HAZV or CCHFV S segment or a partial sequence of the HAZV L segment of Hazara virus were prepared by inserting the viral sequences, amplified by reverse transcription-PCR (primer sequence available upon request), into the unique BamHI site of plasmid pSFV1. Recombinant plasmids were linearized at the unique SpeI site, in vitro transcribed by the SP6 RNA polymerase, and electroporated into BHK-21 cells along with Helper-2 RNA to produce suicide particles, as described previously (22). The sequence of CCHFV was derived from strain IbAr10200 (GenBank accession number U88410).
IFA. For immunofluorescence assay (IFA), cells grown and infected on a glass slide were fixed with 3.7% formaldehyde in phosphate-buffered saline and permeabilized with 0.5% Triton X-100. Mouse hyperimmune ascites raised against HAZV (a generous gift from R. Shope) was used at a dilution of 1:500 followed by fluorescein isothiocyanate-labeled secondary antibody (diluted 1:100) to stain the infected cells. Finally, the cells were stained by Evans' blue and observed under a fluorescent microscope with an appropriate barrier and excitation filters.
Western blot. For Western blotting, infected cells were lysed in TNE (50 mM Tris-HCl [pH 7.4], 100 mM NaCl, 0.5 mM EDTA) buffer containing 0.6% NP-40 and treated with benzonase (Merck) to digest nuclear DNA. Protein concentration was quantified by using a BCA protein assay kit (Pierce). Equal amounts of protein samples were loaded onto a 10% polyacrylamide gel and separated under denaturing conditions at 150 V and 20 mA. Proteins were transferred onto a nitrocellulose membrane (Hybond C extra; Amersham) and revealed using mouse hyperimmune ascites incubated overnight at 4°C at a dilution of 1:5,000 and secondary peroxydase-labeled antibody. Bands were revealed by Super Signal West Pico chemiluminescence (Pierce) on Fuji Medical X-ray films.
RNA analysis. For Northern blots, total RNA was extracted using Trizol and denatured in the presence of 50% formamide, and RNA (3 μg) was electrophoresed in a 0.8% denaturing agarose gel containing 6% formaldehyde (21). The samples were blotted onto Hybond N membranes (Amersham). Prehybridization and hybridization with the riboprobes (see below) were performed at 42°C in a buffer containing 50% formamide, 5x Denhardt's solution, 0.5% sodium dodecyl sulfate (SDS), 5x SSPE (0.75 M NaCl, 50 mM NaH2PO4, 5 mM EDTA, pH 7.4), and 20 μg/ml salmon sperm DNA. The membrane was hybridized overnight and washed successively with 2x SSPE containing 0.1% SDS at room temperature for 20 min, 1x SSPE containing 0.1% SDS for 20 min at 65°C, and 0.1x SSPE containing 0.1% SDS for 20 min at 65°C. The riboprobe complementary to the HAZV S segment was synthesized from full-length S cDNA cloned in pGem-4Z (Promega) downstream from the SP6 promoter. The plasmid was linearized with the appropriate restriction enzyme, and the RNA was synthesized using SP6 polymerase in the presence of [-32P]CTP (Amersham).
Detection of RNase activity in cell extracts was carried out as described previously (17) using 32P-labeled specific and unspecific probes, i.e., in vitro RNA transcripts encompassing, respectively, the first 784 nucleotides (nt) of the mRNA of the HAZV nucleoprotein or the first 501 nt of the mRNA of the nucleoprotein of Rift Valley fever virus and cloned in pGem-4Z (Promega) downstream from the SP6 promoter.
RNase protection assays were performed using the RPA III kit (Ambion) following the manufacturer's instructions. Each assay was carried out with 20 μg of total cellular RNA hybridized to an RNA probe synthetized in vitro using SP6 RNA polymerase. The HAZV S RNA probe is 126 nt long, including a 20-nt sequence from the pGEM 4Z vector, and represents the S viral complementary RNA sequence from positions 1 to 106. The SFV RNA probe is 98 nucleotides long and represents the sequence of the SFV-1 genome from positions 7303 to 7401. The protected RNAs were heat denatured and analyzed by electrophoresis in a 15% polyacrylamide-7 M urea gel together with an RNA ladder.
RESULTS
ISE6 cells support replication of HAZV. HAZV replicated in ISE6 cells without any cytopathic effect, even after 1 to 2 weeks of infection, and the virus was shed into the extracellular medium. Virus titer increased progressively, reaching 5 x 106 to 8 x 106 FFU/ml on day 8 postinfection (p.i.) (Fig. 1A), a value close to that obtained with BHK-21 cells. Viral replication was also monitored by Western blotting and IFA. The nucleoprotein accumulated in the cytoplasm (Fig. 1B and C) as has been shown for bunyaviruses which replicate in the cytoplasm (36). Compared to the mammalian system, the virus grew slowly in ISE6 cells. By 96 h p.i., only 10 to 20% of cells that had been infected at an MOI of 1 reacted with antibody specific to nucleoprotein as observed by IFA (Fig. 1C, day 4). The percentage of positive cells increased with time, reaching approximately 60 to 80% on day 8 p.i. (Fig. 1C).
Recombinant SFV-HAZ infects ISE6 cells and protects against HAZV superinfection. Arthropod cells, and ISE6 cells in particular, are very fragile, and plasmid transfection efficiency is very low in these cells. Therefore, we tested the ability of rSFV to infect tick cells. pSFV1-based plasmids were constructed by inserting the complete HAZV S-segment cDNA in the genomic and antigenomic orientations into the multiple-cloning site, producing the corresponding suicide viruses SFV-HAZ-Sg and SFV-HAZ-Sag, respectively. The suicide viruses infect a wide variety of cells but cannot propagate because the encapsidated replicon lacks the region coding for the structural proteins (37). The recombinant suicide virus SFV-HAZ-Sag containing the message sense sequence expresses the HAZV nucleoprotein after infection of BHK-21 cells (not shown). Nucleoprotein expression was also monitored in ISE6 monolayers by IFA and Western blot. The majority of cells infected at an MOI of 5 were positive for viral nucleoprotein by 24 and 48 h p.i., but the percentage of positive cells (detected by IFA) and the amount of protein (estimated by Western blot) decreased during the following days, being undetectable at 8 to 10 days p.i. (data not shown). Furthermore, in contrast to mammalian cells, no visible cytopathic effect was observed during infection in ISE6 cells.
In an attempt to induce PDR against HAZV in ISE6 cells, we utilized SFV-HAZ-Sag and SFV-HAZ-Sg (i.e., expressing no protein) as well as SFV-HAZ-Lg and SFV-HAZ-Lag, containing part of the L segment in both orientations, and SFV-CCHF-Sag, expressing the nucleoprotein of CCHFV. The latter was used to study the importance of sequence conservation for induction of PDR and RNAi. PDR was evaluated by titration of HAZV produced in the culture medium at day 8 p.i. Since rSFV does not produce infectious viral particles, it does not interfere with the titration of HAZV.
When cells infected with SFV-HAZ-Sg or SFV-HAZ-Sag were superinfected with HAZV on day 2 p.i., the HAZV yield was reduced by 3 to 4 log compared to the titer reached in cells infected either with HAZV alone or with HAZV and SFV1, the recombinant SFV with no insert (Table 1). This indicates that expression of the HAZV S segment induced strong resistance against HAZV, while infection by the SFV1 vector has no inhibitory effect. The interference occurred in cells expressing the N open reading frame as well as the complete HAZV S segment with the sequence in the coding or noncoding orientation, confirming that the mechanism underlying PDR was not protein dependent but was due to viral RNA synthesis. In later experiments, PDR against HAZV was therefore induced using SFV-HAZ-Sg which expresses only RNA but no protein. In cells infected with SFV-HAZ-Sg or SFV-HAZ-Sag, a dramatic reduction of HAZ virus titer was linked to significant reduction in viral proteins and viral RNA as monitored by Western and Northern blotting (Fig. 2). In contrast to SFV expressing the S sequence, recombinant virus expressing the L sequence or the CCHFV S segment had almost no interfering effect. The reason why the L sequence has no effect is not known, but this has also been observed with other bunyaviruses (7, 29, 30). The results with the CCHFV S segment are not unexpected, considering that the HAZV and CCHFV sequences have only 60% identity (24) and that RNA-dependent silencing is highly sequence specific. Similarly, replication of another member of the nairoviruses, Dugbe virus, in ISE6 cells expressing the HAZV sequences was not affected, confirming that sequence specificity is crucial (not shown).
To determine if PDR can persist, cells were infected with SFV-HAZ-Sg and challenged with HAZV at day 0, 1, 2, 3, 4, or 5 p.i. The results summarized in Table 2 indicate that interference occurred almost to the same extent when ISE6 cells infected by rSFV were superinfected with HAZV 1 to 5 days after infection. This shows that the process is rapidly established after HAZV RNA expression and lasted for several days.
HAZV replication can be silenced by SFV-HAZ superinfection. The above-mentioned data showed that HAZV S RNA, when expressed prior to HAZV infection, mediated PDR. To determine whether cells first infected by HAZV could be cured or virus replication could be reduced by superinfection with rSFV, ISE6 cells were first infected with HAZV at an MOI of 1. Two days postinfection, cells were superinfected with SFV-1 or SFV-HAZ-Sg. Four and 8 days after HAZV infection, the replication of HAZV was assayed. SFV-HAZ-Sg superinfection reduced HAZV yield more than 100-fold (Table 3). This shows that prior infection with HAZV does not compromise the ability of the rSFV replicon to silence the targeted virus.
Recombinant SFV-HAZ induces RNAi against HAZV. If PDR was due to RNAi, we assumed that a specific Dicer-like RNase activity specifically targeting HAZV RNA should be detectable, as described in Drosophila melanogaster cells and organisms that exhibit RNA silencing (6). Thus, the presence of RNase activity in extracts of ISE6 cells infected with SFV-HAZ-Sg and harvested 48 h p.i. was assayed after incubation with specific or nonspecific RNA molecules. The probe representing the HAZV S mRNA sequence was rapidly and almost completely degraded after a 30-min incubation (Fig. 3A, compare lanes 1 and 2), whereas the RNA molecules representing the unspecific sequence were only partially degraded after 60 min (lanes 4 to 6). These results strongly suggest the presence of a specific RNase activity as well as a low-activity nonspecific RNase that probably targets any naked RNA.
Because small 21- to 24-nt-long interfering RNAs are a hallmark of RNAi, we attempted to detect specific siRNAs using an RNase protection assay. Total RNA from ISE6 cells infected with SFV-HAZ-Sg at day 2 or 3 p.i. was denatured, hybridized with a 32P-labeled single-stranded RNA (ssRNA) probe representing a region of the HAZV N mRNA, and digested with ssRNA-specific RNase. As shown in Fig. 3, using a probe corresponding to 106 nt at the 5' end of the antigenome of the S segment of HAZV, we were able to detect 21- to 24-nt-long siRNA in cells infected with SFV-HAZ-Sg (Fig. 3B, lanes 3 and 4) but not in uninfected cells (lane 2) or cells infected with SFV1 (not shown). Moreover, cells infected with SFV-HAZ-Sg and collected at 72 h harbored significantly more siRNAs than those collected at 48 h p.i., suggesting that siRNAs accumulated during rSFV replication. Of note, during the time of the experiments (up to 10 days) HAZV by itself did not induce RNAi via expression of siRNAs in infected cells (data not shown). This is in agreement with the fact that the viral nucleoprotein accumulated over time during infection (Fig. 1).
Moreover, siRNAs corresponding to a region specific for the replicon were also detected (Fig. 3B, lanes 6 and 7), but as expected, they were found in cells infected with SFV-HAZ-Sg as well as with SFV1 (Fig. 3B, lanes 10 and 11). This implies that once SFV infection is initiated, its expression is downregulated. This is exactly what we observed: expression of HAZV nucleoprotein by SFV-HAZ-Sag was visible 24 and 48 h p.i. but decreased progressively afterwards. Interestingly, SFV downregulation correlated with the presence of siRNAs. We should point out that cells infected with SFV-HAZ-Sag on day 3 p.i. or later and which expressed low levels of N RNA nevertheless efficiently silenced HAZV replication. Evidently, the level of siRNAs was high enough to block multiplication of both the SFV replicon and HAZV.
DISCUSSION
Ticks are vectors for many viral, bacterial, and protozoan pathogens affecting humans and animals (e.g., tick-borne encephalitis virus, Crimean-Congo hemorrhagic fever virus, Lyme borreliae, and babesias). A major challenge for the control of arboviruses is to find means to interfere with the replication of the virus in its vector. We report here the use of recombinant Semliki Forest virus to induce RNA silencing against the nairovirus HAZV in tick cells expressing the nucleoprotein gene. The SFV replicon is a useful vector infecting a wide range of vertebrate and invertebrate cells. Mosquitoes are the natural vector of SFV, and the SFV replicon was shown to inhibit replication of Rift Valley fever virus in mosquito cells via an RNA-mediated mechanism (7). Similarly, recombinant Sindbis viruses can induce silencing against Dengue and La Crosse viruses in mosquitoes and mosquito cells (1, 27). To our knowledge, this is the first report concerning the silencing of a tick-borne virus in I. scapularis tick cells via RNAi. Similar to results obtained previously by others (7, 14, 30), we found the nucleoprotein gene to be a potent inducer of RNAi against bunyaviruses, whereas the L gene was much less efficient. It seems that not every sequence is able to induce RNAi. Indeed, Caplen et al. found that when RNAi was induced against SFV by synthetic double-stranded RNAs (dsRNAs), dsRNA representing a sequence from the nsp-1 gene failed to trigger silencing, whereas dsRNA from nsp-2 and nsp-4 were highly active (10). It should be noted that rSFV allows induction of a wide range of siRNAs, which reduces the risk for generation of viral escape mutants due to nucleotide substitutions in the sequence targeted by the siRNAs, as has been shown to occur with synthetic siRNAs against poliovirus and human immunodeficiency virus (9, 15).
One hallmark of RNAi is the presence of 21- to 24-nt siRNAs which are produced through the specific cleavage of dsRNA by an RNase III-like enzyme named Dicer (6). The siRNAs associate with various proteins forming the enzymatic complex RISC (RNA-induced silencing complex) including the Argonaute protein, which harbors a helicase and targets mRNAs (17). The antisense strand of the siRNA guides RISC to the mRNA target, and the nuclease component cleaves the RNA in a sequence-specific manner. The mechanism was demonstrated in plants, Drosophila melanogaster, Caenorhabditis elegans, fungi, mammals, and mosquitoes. Less is known from ticks, but the experiments described here indicate the existence of such a mechanism: (i) the phenomenon is sequence specific, SFV-CCHF-Sag does not interfere with HAZV replication, and Dugbe virus is not silenced by SFV-HAZ-Sg (Table 1); (ii) the RNase activity preferentially degrades the HAZV probe (Fig. 3A); and (iii) specific 21- to 24-nt-long siRNAs are present in ISE6 cells infected with recombinant SFV (Fig. 3B). The siRNAs detected with the SFV-specific probe were observed in ISE6 cells infected with SFV-HAZ-Sg as well as with SFV1, indicating that SFV is at the same time the inducer and the target for RNA silencing, the silencing being directed towards the viral sequence expressed by the replicon and toward the replicon itself. This implies that SFV expression is self-inhibited, as evidenced by the fact that expression of the N protein is optimal at 48 h p.i. and declines progressively thereafter. This decrease did not affect silencing of HAZV but seems to correlate with increasing production of siRNAs over time (Fig. 3B, compare lanes 3 and 4 and 6 and 7). Similarly, declining protein or RNA expression in alphavirus-infected insects and cell lines did not interfere with efficient silencing that occurred even with low levels of effector RNAs (1, 34). The alphavirus O'nyong-nyong replicated to much higher titers in mosquitoes when the RNAi pathway was blocked by silencing the Argonaute2 gene (part of the RISC). This provides further evidence that RNAi functions as an antiviral response in insects (20). Our results suggest that a similar mechanism is active in tick cells, and it is likely that an Argonaute homolog is involved in RISC formation.
RNAi is being used as a tool for functional gene analysis in invertebrates including ticks and is now being used as a therapeutic tool in invertebrates and vertebrates. In ticks, various genes coding for proteins secreted by the salivary glands, i.e., the histamine binding protein, the anticoagulant Salp-14, or synaptobrevin (3, 4, 19, 26), have been specifically silenced using this approach. These reports not only showed that dsRNA-induced gene silencing can occur in ticks but suggest that RNA interference can spread from the injection site to the salivary glands. A similar mechanism, involving the SID-1 protein (12, 39), exists in plants and in C. elegans but seems absent in D. melanogaster (33).
In summary, if applicable to the whole tick, our results suggest that nairovirus replication could potentially be inhibited via expression of the nucleoprotein RNA sequence in the tick vector, ablating virus transmission to the vertebrate hosts. This could be of special interest for the highly pathogenic CCHFV. With the recent development of transgenic mosquitoes, it may soon be possible to apply the technology to other invertebrates. Transgenic ticks expressing pathogen-specific siRNAs could potentially be used to disrupt the natural transmission cycle of many human and animal diseases.
ACKNOWLEDGMENTS
We thank C. Antoniewski for helpful discussions and advice on setting up RNase protection assays.
S.G. was the recipient of a Ph.D. fellowship from the Délégation Générale de l'Armenent.
REFERENCES
Adelman, Z. N., C. D. Blair, J. O. Carlson, B. J. Beaty, and K. E. Olson. 2001. Sindbis virus-induced silencing of dengue viruses in mosquitoes. Insect Mol. Biol. 10:265-273.
Adelman, Z. N., I. Sanchez-Vargas, E. A. Travanty, J. O. Carlson, B. J. Beaty, C. D. Blair, and K. E. Olson. 2002. RNA silencing of dengue virus type 2 replication in transformed C6/36 mosquito cells transcribing an inverted-repeat RNA derived from the virus genome. J. Virol. 76:12925-12933.
Aljamali, M. N., A. D. Bior, J. R. Sauer, and R. C. Essenberg. 2003. RNA interference in ticks: a study using histamine binding protein dsRNA in the female tick Amblyomma americanum. Insect Mol. Biol. 12:299-305.
Aljamali, M. N., J. R. Sauer, and R. C. Essenberg. 2002. RNA interference: applicability in tick research. Exp. Appl. Acarol. 28:89-96.
Begum, F., C. L. Wisseman, Jr., and J. Casals. 1970. Tick-borne viruses of West Pakistan. II. Hazara virus, a new agent isolated from Ixodes redikorzevi ticks from the Kaghan Valley, W. Pakistan. Am. J. Epidemiol. 92:192-194.
Bernstein, E., A. A. Caudy, S. M. Hammond, and G. J. Hannon. 2001. Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409:363-366.
Billecocq, A., M. Vazeille-Falcoz, F. Rodhain, and M. Bouloy. 2000. Pathogen-specific resistance to Rift Valley fever virus infection is induced in mosquito cells by expression of the recombinant nucleoprotein but not NSs non-structural protein sequences. J. Gen. Virol. 81:2161-2166.
Blair, C. D., Z. N. Adelman, and K. E. Olson. 2000. Molecular strategies for interrupting arthropod-borne virus transmission by mosquitoes. Clin. Microbiol. Rev. 13:651-661.
Boden, D., O. Pusch, F. Lee, L. Tucker, and B. Ramratnam. 2003. Human immunodeficiency virus type 1 escape from RNA interference. J. Virol. 77:11531-11535.
Caplen, N. J., Z. Zheng, B. Falgout, and R. A. Morgan. 2002. Inhibition of viral gene expression and replication in mosquito cells by dsRNA-triggered RNA interference. Mol. Ther. 6:243-251.
Darwish, M. A., H. Hoogstraal, T. J. Roberts, R. Ghazi, and T. Amer. 1983. A sero-epidemiological survey for Bunyaviridae and certain other arboviruses in Pakistan. Trans. R. Soc. Trop. Med. Hyg. 77:446-450.
Feinberg, E. H., and C. P. Hunter. 2003. Transport of dsRNA into cells by the transmembrane protein SID-1. Science 301:1545-1547.
Foulke, R. S., R. R. Rosato, and G. R. French. 1981. Structural polypeptides of Hazara virus. J. Gen. Virol. 53:169-172.
Gaines, P. J., K. E. Olson, S. Higgs, A. M. Powers, B. J. Beaty, and C. D. Blair. 1996. Pathogen-derived resistance to dengue type 2 virus in mosquito cells by expression of the premembrane coding region of the viral genome. J. Virol. 70:2132-2137.
Gitlin, L., S. Karelsky, and R. Andino. 2002. Short interfering RNA confers intracellular antiviral immunity in human cells. Nature 418:430-434.
Gonzalez-Scarano, F., M. J. Endres, and N. Nathanson. 1991. Bunyaviridae: pathogenesis. Curr. Top. Microbiol. Immunol. 169:217-249.
Hammond, S. M., E. Bernstein, D. Beach, and G. J. Hannon. 2000. An RNA-directed nuclease mediates post-transcriptional gene silencing in Drosophila cells. Nature 404:293-296.
Honig, J. E., J. C. Osborne, and S. T. Nichol. 2004. The high genetic variation of viruses of the genus Nairovirus reflects the diversity of their predominant tick hosts. Virology 318:10-16.
Karim, S., V. G. Ramakrishnan, J. S. Tucker, R. C. Essenberg, and J. R. Sauer. 2004. Amblyomma americanum salivary glands: double-stranded RNA-mediated gene silencing of synaptobrevin homologue and inhibition of PGE2 stimulated protein secretion. Insect Biochem. Mol. Biol. 34:407-413.
Keene, K. M., B. D. Foy, I. Sanchez-Vargas, B. J. Beaty, C. D. Blair, and K. E. Olson. 2004. RNA interference acts as a natural antiviral response to O'nyong-nyong virus (Alphavirus; Togaviridae) infection of Anopheles gambiae. Proc. Natl. Acad. Sci. USA 101:17240-17245.
Lehrach, H., D. Diamond, J. M. Wozney, and H. Boedtker. 1977. RNA molecular weight determinations by gel electrophoresis under denaturing conditions, a critical reexamination. Biochemistry 16:4743-4751.
Liljestrom, P., and H. Garoff. 1991. A new generation of animal cell expression vectors based on the Semliki Forest virus replicon. Biotechnology (New York) 9:1356-1361.
Marczinke, B. I., and S. T. Nichol. 2002. Nairobi sheep disease virus, an important tick-borne pathogen of sheep and goats in Africa, is also present in Asia. Virology 303:146-151.
Marriott, A. C., and P. A. Nuttall. 1992. Comparison of the S RNA segments and nucleoprotein sequences of Crimean-Congo hemorrhagic fever, Hazara, and Dugbe viruses. Virology 189:795-799.
Munderloh, U. G., Y. Liu, M. Wang, C. Chen, and T. J. Kurtti. 1994. Establishment, maintenance and description of cell lines from the tick Ixodes scapularis. J. Parasitol. 80:533-543.
Narasimhan, S., R. R. Montgomery, K. DePonte, C. Tschudi, N. Marcantonio, J. F. Anderson, J. R. Sauer, M. Cappello, F. S. Kantor, and E. Fikrig. 2004. Disruption of Ixodes scapularis anticoagulation by using RNA interference. Proc. Natl. Acad. Sci. USA 101:1141-1146.
Olson, K. E., Z. N. Adelman, E. A. Travanty, I. Sanchez-Vargas, B. J. Beaty, and C. D. Blair. 2002. Developing arbovirus resistance in mosquitoes. Insect Biochem. Mol. Biol. 32:1333-1343.
Olson, K. E., S. Higgs, P. J. Gaines, A. M. Powers, B. S. Davis, K. I. Kamrud, J. O. Carlson, C. D. Blair, and B. J. Beaty. 1996. Genetically engineered resistance to dengue-2 virus transmission in mosquitoes. Science 272:884-886.
Powers, A. M., K. I. Kamrud, K. E. Olson, S. Higgs, J. O. Carlson, and B. J. Beaty. 1996. Molecularly engineered resistance to California serogroup virus replication in mosquito cells and mosquitoes. Proc. Natl. Acad. Sci. USA 93:4187-4191.
Powers, A. M., K. E. Olson, S. Higgs, J. O. Carlson, and B. J. Beaty. 1994. Intracellular immunization of mosquito cells to LaCrosse virus using a recombinant Sindbis virus vector. Virus Res. 32:57-67.
Pudney, M., C. J. Leake, and S. M. Buckley. 1982. Replication of arboviruses in arthropod in vitro systems: an overview, p. 159-194. In K. Maramorosch and J. Mitsuhashi (ed.), Invertebrate cell culture application. Academic Press, New York, NY.
Rayms-Keller, A., A. M. Powers, S. Higgs, K. E. Olson, K. I. Kamrud, J. O. Carlson, and B. J. Beaty. 1995. Replication and expression of a recombinant Sindbis virus in mosquitoes. Insect Mol. Biol. 4:245-251.
Roignant, J. Y., C. Carre, B. Mugat, D. Szymczak, J. A. Lepesant, and C. Antoniewski. 2003. Absence of transitive and systemic pathways allows cell-specific and isoform-specific RNAi in Drosophila. RNA 9:299-308.
Sanchez-Vargas, I., E. A. Travanty, K. M. Keene, A. W. Franz, B. J. Beaty, C. D. Blair, and K. E. Olson. 2004. RNA interference, arthropod-borne viruses, and mosquitoes. Virus Res. 102:65-74.
Sanford, J. C., and S. A. Johnston. 1985. The concept of parasite-derived resistance-deriving resistance genes from the parasite's own genome. J. Theor. Biol. 113:395-405.
Schmaljohn, C., and J. W. Hooper. 2001. Bunyaviridae: the viruses and their replication, vol. 2. Lippincott Williams & Wilkins, Philadelphia, Pa.
Smerdou, C., and P. Liljestrom. 1999. Two-helper RNA system for production of recombinant Semliki Forest virus particles. J. Virol. 73:1092-1098.
Whitehouse, C. A. 2004. Crimean-Congo hemorrhagic fever. Antivir. Res. 64:145-160.
Winston, W. M., C. Molodowitch, and C. P. Hunter. 2002. Systemic RNAi in C. elegans requires the putative transmembrane protein SID-1. Science 295:2456-2459.(Stephan Garcia, Agnès Bil)
Laboratoire de virologie, CRSSA, Centre de Recherche Emile Pardé, Grenoble, France
University of Minnesota, Department of Entomology, St. Paul, Minnesota 55108
ABSTRACT
We report the successful infection of the cell line ISE6 derived from Ixodes scapularis tick embryos by the tick-borne Hazara virus (HAZV), a nairovirus in the family Bunyaviridae. Using a recombinant Semliki Forest alphavirus replicon that replicates in these cells, we were able to inhibit replication of HAZV, and we showed that this blockage is mediated by the replication of the Semliki Forest alphavirus replicon; the vector containing the HAZV nucleoprotein gene in sense or antisense orientation efficiently inhibited HAZV replication. Moreover, expression of a distantly related nucleoprotein gene from Crimean-Congo hemorrhagic fever nairovirus failed to induce HAZV silencing, indicating that the inhibition is sequence specific. The resistance of these cells to replicate HAZV correlated with the detection of specific RNase activity and 21- to 24-nucleotide-long small interfering RNAs. Altogether, these results strongly suggest that pathogen-derived resistance can be established in the tick cells via a mechanism of RNA interference.
INTRODUCTION
Pathogen-derived resistance (PDR) is a genetic mechanism that reduces host or cell susceptibility to virus infection and has been illustrated in several viral infection models. It is caused by expression of certain pathogen-derived genes interfering with infection by a closely related pathogen. It was first described by Sanford and Johnston in 1985 (35) and has been shown to be effective against arboviruses with a positive or negative RNA genome like dengue virus, La Crosse virus, or Rift Valley fever virus (7, 14, 30). Resistance was demonstrated after in vitro or in vivo expression of a region of the viral genome. The cells or mosquitoes expressing these viral sequences became resistant to infection with the homologous virus (8, 27). To induce PDR, the Sindbis alphavirus vector was utilized because its genome could be manipulated through an infectious cDNA (28, 32). The replicon of the Semliki Forest virus (SFV), another alphavirus, was also used to interfere with the replication of Rift Valley fever virus in mosquito cells (7). Resistance to infection was dependent on RNA but not protein expression. PDR was also induced by artificial, inverted-repeat RNAs homologous to the targeted dengue virus. Plasmid-transfected mosquito cells generated small interfering RNAs (siRNAs) that triggered silencing of homologous viral gene expression (2, 34).
The concept of PDR and RNA interference (RNAi) has been relatively well documented in mosquitoes and mosquito cells, but very little is known in ticks and tick cells for the tick-borne viruses. Among bunyaviruses, the genus Nairovirus comprises 34 tick-borne viruses classified into seven serogroups. The main representatives are the widely distributed human pathogen Crimean-Congo hemorrhagic fever virus (CCHFV), which causes severe and often fatal hemorrhagic fever in humans (16), and the Nairobi sheep disease virus, which circulates in Africa and Asia (23) and induces acute and fatal hemorrhagic gastroenteritis in sheep and goats. Although nairovirus infection in vertebrate cells has been documented (for reviews, see references 36 and 38), there are only a few reports concerning nairovirus infection in their tick vector or in tick cells (31).
In this study, we used the ISE6 tick cell line, derived from Ixodes scapularis (25), and show that these cells supported the efficient replication of Hazara virus (HAZV), a nairovirus isolated from Ixodes ticks collected in Pakistan (5, 11, 18). Because it is not pathogenic for humans, it was used as a model for the very pathogenic CCHFV, classified in the same serogroup (13). We addressed the question of whether PDR against HAZV can be established in tick cells. This was achieved by expressing HAZV genome sequences via infection with specific recombinant SFV (rSFV). In these cells, PDR correlated with the detection of sequence-specific RNase activity and siRNAs, which strongly suggest the existence of a mechanism of RNAi. Altogether, these results establish the feasibility for the control of tick-borne viruses in their vector.
MATERIALS AND METHODS
Virus and cells. BHK-21 cells were grown in Glasgow minimum essential medium supplemented with 5% fetal calf serum, 10% tryptose phosphate, 10 mM HEPES, and antibiotics. Vero E6 cells were grown in Dulbecco's minimum essential medium supplemented with 10% fetal calf serum and penicillin/streptomycin. Cell line ISE6 was established from embryos of the black-legged tick Ixodes scapularis and grown as previously described (25).
Hazara virus (strain JC280) and Dugbe virus (strain IbAr 1792) were provided by E. Gould (Natural Environment Research Council, Oxford, United Kingdom). Stocks grown by infecting BHK-21 cells at a multiplicity of infection (MOI) of 0.01 reached 106 to 107 focus-forming units (FFU) per ml. Virus was titrated by focus formation in Vero E6 cells. At day 5 postinfection, the agarose overlay was removed, cells were fixed with methanol, and infected foci were visualized after staining with HAZV antibody (dilution, 1:250) revealed by peroxidase-labeled anti-mouse antibody (dilution, 1:1,000).
Recombinant Semliki Forest virus. SFV replicons carrying partial or complete sequences of the HAZV or CCHFV S segment or a partial sequence of the HAZV L segment of Hazara virus were prepared by inserting the viral sequences, amplified by reverse transcription-PCR (primer sequence available upon request), into the unique BamHI site of plasmid pSFV1. Recombinant plasmids were linearized at the unique SpeI site, in vitro transcribed by the SP6 RNA polymerase, and electroporated into BHK-21 cells along with Helper-2 RNA to produce suicide particles, as described previously (22). The sequence of CCHFV was derived from strain IbAr10200 (GenBank accession number U88410).
IFA. For immunofluorescence assay (IFA), cells grown and infected on a glass slide were fixed with 3.7% formaldehyde in phosphate-buffered saline and permeabilized with 0.5% Triton X-100. Mouse hyperimmune ascites raised against HAZV (a generous gift from R. Shope) was used at a dilution of 1:500 followed by fluorescein isothiocyanate-labeled secondary antibody (diluted 1:100) to stain the infected cells. Finally, the cells were stained by Evans' blue and observed under a fluorescent microscope with an appropriate barrier and excitation filters.
Western blot. For Western blotting, infected cells were lysed in TNE (50 mM Tris-HCl [pH 7.4], 100 mM NaCl, 0.5 mM EDTA) buffer containing 0.6% NP-40 and treated with benzonase (Merck) to digest nuclear DNA. Protein concentration was quantified by using a BCA protein assay kit (Pierce). Equal amounts of protein samples were loaded onto a 10% polyacrylamide gel and separated under denaturing conditions at 150 V and 20 mA. Proteins were transferred onto a nitrocellulose membrane (Hybond C extra; Amersham) and revealed using mouse hyperimmune ascites incubated overnight at 4°C at a dilution of 1:5,000 and secondary peroxydase-labeled antibody. Bands were revealed by Super Signal West Pico chemiluminescence (Pierce) on Fuji Medical X-ray films.
RNA analysis. For Northern blots, total RNA was extracted using Trizol and denatured in the presence of 50% formamide, and RNA (3 μg) was electrophoresed in a 0.8% denaturing agarose gel containing 6% formaldehyde (21). The samples were blotted onto Hybond N membranes (Amersham). Prehybridization and hybridization with the riboprobes (see below) were performed at 42°C in a buffer containing 50% formamide, 5x Denhardt's solution, 0.5% sodium dodecyl sulfate (SDS), 5x SSPE (0.75 M NaCl, 50 mM NaH2PO4, 5 mM EDTA, pH 7.4), and 20 μg/ml salmon sperm DNA. The membrane was hybridized overnight and washed successively with 2x SSPE containing 0.1% SDS at room temperature for 20 min, 1x SSPE containing 0.1% SDS for 20 min at 65°C, and 0.1x SSPE containing 0.1% SDS for 20 min at 65°C. The riboprobe complementary to the HAZV S segment was synthesized from full-length S cDNA cloned in pGem-4Z (Promega) downstream from the SP6 promoter. The plasmid was linearized with the appropriate restriction enzyme, and the RNA was synthesized using SP6 polymerase in the presence of [-32P]CTP (Amersham).
Detection of RNase activity in cell extracts was carried out as described previously (17) using 32P-labeled specific and unspecific probes, i.e., in vitro RNA transcripts encompassing, respectively, the first 784 nucleotides (nt) of the mRNA of the HAZV nucleoprotein or the first 501 nt of the mRNA of the nucleoprotein of Rift Valley fever virus and cloned in pGem-4Z (Promega) downstream from the SP6 promoter.
RNase protection assays were performed using the RPA III kit (Ambion) following the manufacturer's instructions. Each assay was carried out with 20 μg of total cellular RNA hybridized to an RNA probe synthetized in vitro using SP6 RNA polymerase. The HAZV S RNA probe is 126 nt long, including a 20-nt sequence from the pGEM 4Z vector, and represents the S viral complementary RNA sequence from positions 1 to 106. The SFV RNA probe is 98 nucleotides long and represents the sequence of the SFV-1 genome from positions 7303 to 7401. The protected RNAs were heat denatured and analyzed by electrophoresis in a 15% polyacrylamide-7 M urea gel together with an RNA ladder.
RESULTS
ISE6 cells support replication of HAZV. HAZV replicated in ISE6 cells without any cytopathic effect, even after 1 to 2 weeks of infection, and the virus was shed into the extracellular medium. Virus titer increased progressively, reaching 5 x 106 to 8 x 106 FFU/ml on day 8 postinfection (p.i.) (Fig. 1A), a value close to that obtained with BHK-21 cells. Viral replication was also monitored by Western blotting and IFA. The nucleoprotein accumulated in the cytoplasm (Fig. 1B and C) as has been shown for bunyaviruses which replicate in the cytoplasm (36). Compared to the mammalian system, the virus grew slowly in ISE6 cells. By 96 h p.i., only 10 to 20% of cells that had been infected at an MOI of 1 reacted with antibody specific to nucleoprotein as observed by IFA (Fig. 1C, day 4). The percentage of positive cells increased with time, reaching approximately 60 to 80% on day 8 p.i. (Fig. 1C).
Recombinant SFV-HAZ infects ISE6 cells and protects against HAZV superinfection. Arthropod cells, and ISE6 cells in particular, are very fragile, and plasmid transfection efficiency is very low in these cells. Therefore, we tested the ability of rSFV to infect tick cells. pSFV1-based plasmids were constructed by inserting the complete HAZV S-segment cDNA in the genomic and antigenomic orientations into the multiple-cloning site, producing the corresponding suicide viruses SFV-HAZ-Sg and SFV-HAZ-Sag, respectively. The suicide viruses infect a wide variety of cells but cannot propagate because the encapsidated replicon lacks the region coding for the structural proteins (37). The recombinant suicide virus SFV-HAZ-Sag containing the message sense sequence expresses the HAZV nucleoprotein after infection of BHK-21 cells (not shown). Nucleoprotein expression was also monitored in ISE6 monolayers by IFA and Western blot. The majority of cells infected at an MOI of 5 were positive for viral nucleoprotein by 24 and 48 h p.i., but the percentage of positive cells (detected by IFA) and the amount of protein (estimated by Western blot) decreased during the following days, being undetectable at 8 to 10 days p.i. (data not shown). Furthermore, in contrast to mammalian cells, no visible cytopathic effect was observed during infection in ISE6 cells.
In an attempt to induce PDR against HAZV in ISE6 cells, we utilized SFV-HAZ-Sag and SFV-HAZ-Sg (i.e., expressing no protein) as well as SFV-HAZ-Lg and SFV-HAZ-Lag, containing part of the L segment in both orientations, and SFV-CCHF-Sag, expressing the nucleoprotein of CCHFV. The latter was used to study the importance of sequence conservation for induction of PDR and RNAi. PDR was evaluated by titration of HAZV produced in the culture medium at day 8 p.i. Since rSFV does not produce infectious viral particles, it does not interfere with the titration of HAZV.
When cells infected with SFV-HAZ-Sg or SFV-HAZ-Sag were superinfected with HAZV on day 2 p.i., the HAZV yield was reduced by 3 to 4 log compared to the titer reached in cells infected either with HAZV alone or with HAZV and SFV1, the recombinant SFV with no insert (Table 1). This indicates that expression of the HAZV S segment induced strong resistance against HAZV, while infection by the SFV1 vector has no inhibitory effect. The interference occurred in cells expressing the N open reading frame as well as the complete HAZV S segment with the sequence in the coding or noncoding orientation, confirming that the mechanism underlying PDR was not protein dependent but was due to viral RNA synthesis. In later experiments, PDR against HAZV was therefore induced using SFV-HAZ-Sg which expresses only RNA but no protein. In cells infected with SFV-HAZ-Sg or SFV-HAZ-Sag, a dramatic reduction of HAZ virus titer was linked to significant reduction in viral proteins and viral RNA as monitored by Western and Northern blotting (Fig. 2). In contrast to SFV expressing the S sequence, recombinant virus expressing the L sequence or the CCHFV S segment had almost no interfering effect. The reason why the L sequence has no effect is not known, but this has also been observed with other bunyaviruses (7, 29, 30). The results with the CCHFV S segment are not unexpected, considering that the HAZV and CCHFV sequences have only 60% identity (24) and that RNA-dependent silencing is highly sequence specific. Similarly, replication of another member of the nairoviruses, Dugbe virus, in ISE6 cells expressing the HAZV sequences was not affected, confirming that sequence specificity is crucial (not shown).
To determine if PDR can persist, cells were infected with SFV-HAZ-Sg and challenged with HAZV at day 0, 1, 2, 3, 4, or 5 p.i. The results summarized in Table 2 indicate that interference occurred almost to the same extent when ISE6 cells infected by rSFV were superinfected with HAZV 1 to 5 days after infection. This shows that the process is rapidly established after HAZV RNA expression and lasted for several days.
HAZV replication can be silenced by SFV-HAZ superinfection. The above-mentioned data showed that HAZV S RNA, when expressed prior to HAZV infection, mediated PDR. To determine whether cells first infected by HAZV could be cured or virus replication could be reduced by superinfection with rSFV, ISE6 cells were first infected with HAZV at an MOI of 1. Two days postinfection, cells were superinfected with SFV-1 or SFV-HAZ-Sg. Four and 8 days after HAZV infection, the replication of HAZV was assayed. SFV-HAZ-Sg superinfection reduced HAZV yield more than 100-fold (Table 3). This shows that prior infection with HAZV does not compromise the ability of the rSFV replicon to silence the targeted virus.
Recombinant SFV-HAZ induces RNAi against HAZV. If PDR was due to RNAi, we assumed that a specific Dicer-like RNase activity specifically targeting HAZV RNA should be detectable, as described in Drosophila melanogaster cells and organisms that exhibit RNA silencing (6). Thus, the presence of RNase activity in extracts of ISE6 cells infected with SFV-HAZ-Sg and harvested 48 h p.i. was assayed after incubation with specific or nonspecific RNA molecules. The probe representing the HAZV S mRNA sequence was rapidly and almost completely degraded after a 30-min incubation (Fig. 3A, compare lanes 1 and 2), whereas the RNA molecules representing the unspecific sequence were only partially degraded after 60 min (lanes 4 to 6). These results strongly suggest the presence of a specific RNase activity as well as a low-activity nonspecific RNase that probably targets any naked RNA.
Because small 21- to 24-nt-long interfering RNAs are a hallmark of RNAi, we attempted to detect specific siRNAs using an RNase protection assay. Total RNA from ISE6 cells infected with SFV-HAZ-Sg at day 2 or 3 p.i. was denatured, hybridized with a 32P-labeled single-stranded RNA (ssRNA) probe representing a region of the HAZV N mRNA, and digested with ssRNA-specific RNase. As shown in Fig. 3, using a probe corresponding to 106 nt at the 5' end of the antigenome of the S segment of HAZV, we were able to detect 21- to 24-nt-long siRNA in cells infected with SFV-HAZ-Sg (Fig. 3B, lanes 3 and 4) but not in uninfected cells (lane 2) or cells infected with SFV1 (not shown). Moreover, cells infected with SFV-HAZ-Sg and collected at 72 h harbored significantly more siRNAs than those collected at 48 h p.i., suggesting that siRNAs accumulated during rSFV replication. Of note, during the time of the experiments (up to 10 days) HAZV by itself did not induce RNAi via expression of siRNAs in infected cells (data not shown). This is in agreement with the fact that the viral nucleoprotein accumulated over time during infection (Fig. 1).
Moreover, siRNAs corresponding to a region specific for the replicon were also detected (Fig. 3B, lanes 6 and 7), but as expected, they were found in cells infected with SFV-HAZ-Sg as well as with SFV1 (Fig. 3B, lanes 10 and 11). This implies that once SFV infection is initiated, its expression is downregulated. This is exactly what we observed: expression of HAZV nucleoprotein by SFV-HAZ-Sag was visible 24 and 48 h p.i. but decreased progressively afterwards. Interestingly, SFV downregulation correlated with the presence of siRNAs. We should point out that cells infected with SFV-HAZ-Sag on day 3 p.i. or later and which expressed low levels of N RNA nevertheless efficiently silenced HAZV replication. Evidently, the level of siRNAs was high enough to block multiplication of both the SFV replicon and HAZV.
DISCUSSION
Ticks are vectors for many viral, bacterial, and protozoan pathogens affecting humans and animals (e.g., tick-borne encephalitis virus, Crimean-Congo hemorrhagic fever virus, Lyme borreliae, and babesias). A major challenge for the control of arboviruses is to find means to interfere with the replication of the virus in its vector. We report here the use of recombinant Semliki Forest virus to induce RNA silencing against the nairovirus HAZV in tick cells expressing the nucleoprotein gene. The SFV replicon is a useful vector infecting a wide range of vertebrate and invertebrate cells. Mosquitoes are the natural vector of SFV, and the SFV replicon was shown to inhibit replication of Rift Valley fever virus in mosquito cells via an RNA-mediated mechanism (7). Similarly, recombinant Sindbis viruses can induce silencing against Dengue and La Crosse viruses in mosquitoes and mosquito cells (1, 27). To our knowledge, this is the first report concerning the silencing of a tick-borne virus in I. scapularis tick cells via RNAi. Similar to results obtained previously by others (7, 14, 30), we found the nucleoprotein gene to be a potent inducer of RNAi against bunyaviruses, whereas the L gene was much less efficient. It seems that not every sequence is able to induce RNAi. Indeed, Caplen et al. found that when RNAi was induced against SFV by synthetic double-stranded RNAs (dsRNAs), dsRNA representing a sequence from the nsp-1 gene failed to trigger silencing, whereas dsRNA from nsp-2 and nsp-4 were highly active (10). It should be noted that rSFV allows induction of a wide range of siRNAs, which reduces the risk for generation of viral escape mutants due to nucleotide substitutions in the sequence targeted by the siRNAs, as has been shown to occur with synthetic siRNAs against poliovirus and human immunodeficiency virus (9, 15).
One hallmark of RNAi is the presence of 21- to 24-nt siRNAs which are produced through the specific cleavage of dsRNA by an RNase III-like enzyme named Dicer (6). The siRNAs associate with various proteins forming the enzymatic complex RISC (RNA-induced silencing complex) including the Argonaute protein, which harbors a helicase and targets mRNAs (17). The antisense strand of the siRNA guides RISC to the mRNA target, and the nuclease component cleaves the RNA in a sequence-specific manner. The mechanism was demonstrated in plants, Drosophila melanogaster, Caenorhabditis elegans, fungi, mammals, and mosquitoes. Less is known from ticks, but the experiments described here indicate the existence of such a mechanism: (i) the phenomenon is sequence specific, SFV-CCHF-Sag does not interfere with HAZV replication, and Dugbe virus is not silenced by SFV-HAZ-Sg (Table 1); (ii) the RNase activity preferentially degrades the HAZV probe (Fig. 3A); and (iii) specific 21- to 24-nt-long siRNAs are present in ISE6 cells infected with recombinant SFV (Fig. 3B). The siRNAs detected with the SFV-specific probe were observed in ISE6 cells infected with SFV-HAZ-Sg as well as with SFV1, indicating that SFV is at the same time the inducer and the target for RNA silencing, the silencing being directed towards the viral sequence expressed by the replicon and toward the replicon itself. This implies that SFV expression is self-inhibited, as evidenced by the fact that expression of the N protein is optimal at 48 h p.i. and declines progressively thereafter. This decrease did not affect silencing of HAZV but seems to correlate with increasing production of siRNAs over time (Fig. 3B, compare lanes 3 and 4 and 6 and 7). Similarly, declining protein or RNA expression in alphavirus-infected insects and cell lines did not interfere with efficient silencing that occurred even with low levels of effector RNAs (1, 34). The alphavirus O'nyong-nyong replicated to much higher titers in mosquitoes when the RNAi pathway was blocked by silencing the Argonaute2 gene (part of the RISC). This provides further evidence that RNAi functions as an antiviral response in insects (20). Our results suggest that a similar mechanism is active in tick cells, and it is likely that an Argonaute homolog is involved in RISC formation.
RNAi is being used as a tool for functional gene analysis in invertebrates including ticks and is now being used as a therapeutic tool in invertebrates and vertebrates. In ticks, various genes coding for proteins secreted by the salivary glands, i.e., the histamine binding protein, the anticoagulant Salp-14, or synaptobrevin (3, 4, 19, 26), have been specifically silenced using this approach. These reports not only showed that dsRNA-induced gene silencing can occur in ticks but suggest that RNA interference can spread from the injection site to the salivary glands. A similar mechanism, involving the SID-1 protein (12, 39), exists in plants and in C. elegans but seems absent in D. melanogaster (33).
In summary, if applicable to the whole tick, our results suggest that nairovirus replication could potentially be inhibited via expression of the nucleoprotein RNA sequence in the tick vector, ablating virus transmission to the vertebrate hosts. This could be of special interest for the highly pathogenic CCHFV. With the recent development of transgenic mosquitoes, it may soon be possible to apply the technology to other invertebrates. Transgenic ticks expressing pathogen-specific siRNAs could potentially be used to disrupt the natural transmission cycle of many human and animal diseases.
ACKNOWLEDGMENTS
We thank C. Antoniewski for helpful discussions and advice on setting up RNase protection assays.
S.G. was the recipient of a Ph.D. fellowship from the Délégation Générale de l'Armenent.
REFERENCES
Adelman, Z. N., C. D. Blair, J. O. Carlson, B. J. Beaty, and K. E. Olson. 2001. Sindbis virus-induced silencing of dengue viruses in mosquitoes. Insect Mol. Biol. 10:265-273.
Adelman, Z. N., I. Sanchez-Vargas, E. A. Travanty, J. O. Carlson, B. J. Beaty, C. D. Blair, and K. E. Olson. 2002. RNA silencing of dengue virus type 2 replication in transformed C6/36 mosquito cells transcribing an inverted-repeat RNA derived from the virus genome. J. Virol. 76:12925-12933.
Aljamali, M. N., A. D. Bior, J. R. Sauer, and R. C. Essenberg. 2003. RNA interference in ticks: a study using histamine binding protein dsRNA in the female tick Amblyomma americanum. Insect Mol. Biol. 12:299-305.
Aljamali, M. N., J. R. Sauer, and R. C. Essenberg. 2002. RNA interference: applicability in tick research. Exp. Appl. Acarol. 28:89-96.
Begum, F., C. L. Wisseman, Jr., and J. Casals. 1970. Tick-borne viruses of West Pakistan. II. Hazara virus, a new agent isolated from Ixodes redikorzevi ticks from the Kaghan Valley, W. Pakistan. Am. J. Epidemiol. 92:192-194.
Bernstein, E., A. A. Caudy, S. M. Hammond, and G. J. Hannon. 2001. Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409:363-366.
Billecocq, A., M. Vazeille-Falcoz, F. Rodhain, and M. Bouloy. 2000. Pathogen-specific resistance to Rift Valley fever virus infection is induced in mosquito cells by expression of the recombinant nucleoprotein but not NSs non-structural protein sequences. J. Gen. Virol. 81:2161-2166.
Blair, C. D., Z. N. Adelman, and K. E. Olson. 2000. Molecular strategies for interrupting arthropod-borne virus transmission by mosquitoes. Clin. Microbiol. Rev. 13:651-661.
Boden, D., O. Pusch, F. Lee, L. Tucker, and B. Ramratnam. 2003. Human immunodeficiency virus type 1 escape from RNA interference. J. Virol. 77:11531-11535.
Caplen, N. J., Z. Zheng, B. Falgout, and R. A. Morgan. 2002. Inhibition of viral gene expression and replication in mosquito cells by dsRNA-triggered RNA interference. Mol. Ther. 6:243-251.
Darwish, M. A., H. Hoogstraal, T. J. Roberts, R. Ghazi, and T. Amer. 1983. A sero-epidemiological survey for Bunyaviridae and certain other arboviruses in Pakistan. Trans. R. Soc. Trop. Med. Hyg. 77:446-450.
Feinberg, E. H., and C. P. Hunter. 2003. Transport of dsRNA into cells by the transmembrane protein SID-1. Science 301:1545-1547.
Foulke, R. S., R. R. Rosato, and G. R. French. 1981. Structural polypeptides of Hazara virus. J. Gen. Virol. 53:169-172.
Gaines, P. J., K. E. Olson, S. Higgs, A. M. Powers, B. J. Beaty, and C. D. Blair. 1996. Pathogen-derived resistance to dengue type 2 virus in mosquito cells by expression of the premembrane coding region of the viral genome. J. Virol. 70:2132-2137.
Gitlin, L., S. Karelsky, and R. Andino. 2002. Short interfering RNA confers intracellular antiviral immunity in human cells. Nature 418:430-434.
Gonzalez-Scarano, F., M. J. Endres, and N. Nathanson. 1991. Bunyaviridae: pathogenesis. Curr. Top. Microbiol. Immunol. 169:217-249.
Hammond, S. M., E. Bernstein, D. Beach, and G. J. Hannon. 2000. An RNA-directed nuclease mediates post-transcriptional gene silencing in Drosophila cells. Nature 404:293-296.
Honig, J. E., J. C. Osborne, and S. T. Nichol. 2004. The high genetic variation of viruses of the genus Nairovirus reflects the diversity of their predominant tick hosts. Virology 318:10-16.
Karim, S., V. G. Ramakrishnan, J. S. Tucker, R. C. Essenberg, and J. R. Sauer. 2004. Amblyomma americanum salivary glands: double-stranded RNA-mediated gene silencing of synaptobrevin homologue and inhibition of PGE2 stimulated protein secretion. Insect Biochem. Mol. Biol. 34:407-413.
Keene, K. M., B. D. Foy, I. Sanchez-Vargas, B. J. Beaty, C. D. Blair, and K. E. Olson. 2004. RNA interference acts as a natural antiviral response to O'nyong-nyong virus (Alphavirus; Togaviridae) infection of Anopheles gambiae. Proc. Natl. Acad. Sci. USA 101:17240-17245.
Lehrach, H., D. Diamond, J. M. Wozney, and H. Boedtker. 1977. RNA molecular weight determinations by gel electrophoresis under denaturing conditions, a critical reexamination. Biochemistry 16:4743-4751.
Liljestrom, P., and H. Garoff. 1991. A new generation of animal cell expression vectors based on the Semliki Forest virus replicon. Biotechnology (New York) 9:1356-1361.
Marczinke, B. I., and S. T. Nichol. 2002. Nairobi sheep disease virus, an important tick-borne pathogen of sheep and goats in Africa, is also present in Asia. Virology 303:146-151.
Marriott, A. C., and P. A. Nuttall. 1992. Comparison of the S RNA segments and nucleoprotein sequences of Crimean-Congo hemorrhagic fever, Hazara, and Dugbe viruses. Virology 189:795-799.
Munderloh, U. G., Y. Liu, M. Wang, C. Chen, and T. J. Kurtti. 1994. Establishment, maintenance and description of cell lines from the tick Ixodes scapularis. J. Parasitol. 80:533-543.
Narasimhan, S., R. R. Montgomery, K. DePonte, C. Tschudi, N. Marcantonio, J. F. Anderson, J. R. Sauer, M. Cappello, F. S. Kantor, and E. Fikrig. 2004. Disruption of Ixodes scapularis anticoagulation by using RNA interference. Proc. Natl. Acad. Sci. USA 101:1141-1146.
Olson, K. E., Z. N. Adelman, E. A. Travanty, I. Sanchez-Vargas, B. J. Beaty, and C. D. Blair. 2002. Developing arbovirus resistance in mosquitoes. Insect Biochem. Mol. Biol. 32:1333-1343.
Olson, K. E., S. Higgs, P. J. Gaines, A. M. Powers, B. S. Davis, K. I. Kamrud, J. O. Carlson, C. D. Blair, and B. J. Beaty. 1996. Genetically engineered resistance to dengue-2 virus transmission in mosquitoes. Science 272:884-886.
Powers, A. M., K. I. Kamrud, K. E. Olson, S. Higgs, J. O. Carlson, and B. J. Beaty. 1996. Molecularly engineered resistance to California serogroup virus replication in mosquito cells and mosquitoes. Proc. Natl. Acad. Sci. USA 93:4187-4191.
Powers, A. M., K. E. Olson, S. Higgs, J. O. Carlson, and B. J. Beaty. 1994. Intracellular immunization of mosquito cells to LaCrosse virus using a recombinant Sindbis virus vector. Virus Res. 32:57-67.
Pudney, M., C. J. Leake, and S. M. Buckley. 1982. Replication of arboviruses in arthropod in vitro systems: an overview, p. 159-194. In K. Maramorosch and J. Mitsuhashi (ed.), Invertebrate cell culture application. Academic Press, New York, NY.
Rayms-Keller, A., A. M. Powers, S. Higgs, K. E. Olson, K. I. Kamrud, J. O. Carlson, and B. J. Beaty. 1995. Replication and expression of a recombinant Sindbis virus in mosquitoes. Insect Mol. Biol. 4:245-251.
Roignant, J. Y., C. Carre, B. Mugat, D. Szymczak, J. A. Lepesant, and C. Antoniewski. 2003. Absence of transitive and systemic pathways allows cell-specific and isoform-specific RNAi in Drosophila. RNA 9:299-308.
Sanchez-Vargas, I., E. A. Travanty, K. M. Keene, A. W. Franz, B. J. Beaty, C. D. Blair, and K. E. Olson. 2004. RNA interference, arthropod-borne viruses, and mosquitoes. Virus Res. 102:65-74.
Sanford, J. C., and S. A. Johnston. 1985. The concept of parasite-derived resistance-deriving resistance genes from the parasite's own genome. J. Theor. Biol. 113:395-405.
Schmaljohn, C., and J. W. Hooper. 2001. Bunyaviridae: the viruses and their replication, vol. 2. Lippincott Williams & Wilkins, Philadelphia, Pa.
Smerdou, C., and P. Liljestrom. 1999. Two-helper RNA system for production of recombinant Semliki Forest virus particles. J. Virol. 73:1092-1098.
Whitehouse, C. A. 2004. Crimean-Congo hemorrhagic fever. Antivir. Res. 64:145-160.
Winston, W. M., C. Molodowitch, and C. P. Hunter. 2002. Systemic RNAi in C. elegans requires the putative transmembrane protein SID-1. Science 295:2456-2459.(Stephan Garcia, Agnès Bil)