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Journal of Virology, November 2008, p. 10349-10358, Vol. 82, No. 21
0022-538X/08/$08.00+0     doi:10.1128/JVI.00935-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Toll-Like Receptor 3 Has a Protective Role against West Nile Virus Infection

Stephane Daffis,¹ Melanie A. Samuel,² Mehul S. Suthar,⁴ Michael Gale Jr.,⁴ and Michael S. Diamond^1,2,3*

Departments of Medicine,¹ Molecular Microbiology,² Pathology and Immunology, Washington University School of Medicine, St. Louis, Missouri 63110,³ Department of Immunology, University of Washington School of Medicine, Seattle, Washington 98195-7650⁴

Received 5 May 2008/ Accepted 13 August 2008

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Protection against West Nile virus (WNV) infection requires rapid viral sensing and the generation of an interferon (IFN) response. Mice lacking IFN regulatory factor 3 (IRF-3) show increased vulnerability to WNV infection with enhanced viral replication and blunted IFN-stimulated gene (ISG) responses. IRF-3 functions downstream of several viral sensors, including Toll-like receptor 3 (TLR3), RIG-I, and MDA5. Cell culture studies suggest that host recognizes WNV in part, through the cytoplasmic helicase RIG-I and to a lesser extent, MDA5, both of which activate ISG expression through IRF-3. However, the role of TLR3 in vivo in recognizing viral RNA and activating antiviral defense pathways has remained controversial. We show here that an absence of TLR3 enhances WNV mortality in mice and increases viral burden in the brain. Compared to congenic wild-type controls, TLR3^–/– mice showed relatively modest changes in peripheral viral loads. Consistent with this, little difference in multistep viral growth kinetics or IFN-/β induction was observed between wild-type and TLR3^–/– fibroblasts, macrophages, and dendritic cells. In contrast, a deficiency of TLR3 was associated with enhanced viral replication in primary cortical neuron cultures and greater WNV infection in central nervous system neurons after intracranial inoculation. Taken together, our data suggest that TLR3 serves a protective role against WNV in part, by restricting replication in neurons.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Recent studies have begun to delineate the molecular mechanisms that regulate alpha interferon (IFN-) and IFN-β induction after infection by RNA and DNA viruses (reviewed in references 13, 24, 25, and 57). Pattern recognition receptors (PRR) sense conserved structural microbial elements identified as pathogen-associated molecular patterns. PRR involved in the recognition of RNA viruses can be divided into two classes. Toll-like receptors (TLR) on the cell surface or within endosomes recognize single- and double-stranded RNA and signal through the adaptor molecules MyD88 and TRIF. In comparison, RIG-I and MDA5 helicases recognize single- and double-stranded RNA in the cytosol and signal through the adaptor protein IPS-1 (also known as MAVS, Cardif, and VISA) (reviewed in references 28, 42, and 55). Recognition by viral RNA sensors results in the downstream activation and nuclear translocation of IRF-3 and IRF-7, which transcriptionally activate IFN- and -β gene promoters (46).

Cell culture experiments defined TLR3 as a PRR that recognizes double-stranded RNA, activates IRF-3 and NF-B transcriptional pathways, and induces type I IFN (2). The role of TLR3 in vivo in protection against viral infections has been less clear (reviewed in references 5, 36, and 58). Experiments in TLR3^–/– mice infected with lymphocytic choriomeningitis virus, vesicular stomatitis virus, and reovirus failed to show increased mortality or altered viral burden phenotypes (12). In contrast, TLR3 restricts replication, regulates cytokine production, and protects against infection by mouse cytomegalovirus and encephalomyocarditis virus (EMCV) (21, 54). Indeed, a deficiency in TLR3 in humans was identified as a predisposing genetic risk factor for herpes simplex virus (HSV) encephalitis (61) and influenza A virus-induced encephalopathy (23). An adverse role for TLR3 also has been proposed since TLR3^–/– mice infected with influenza, punta toro, and vaccinia viruses showed improved survival and decreased production of inflammatory cytokines (19, 26, 33). Analogously, conflicting results have been observed with respect to the role of TLR3 in inducing IFN after infection with the encephalitic flavivirus West Nile virus (WNV). TLR3 was largely dispensable for WNV recognition and induction of IFN responses in vitro (15, 47), whereas in mice, an absence of TLR3 protected mice from lethal infection (59). TLR3^–/– mice infected via intraperitoneal injection with WNV showed decreased systemic tumor necrosis factor alpha (TNF-) and interleukin-6 (IL-6) production, blood-brain barrier (BBB) permeability, and infection in the brain.

Given this apparent conflict and our previous studies demonstration of an essential role for IRF-3 in controlling WNV infection (6), we reexamined the pathogenesis of virulent WNV infection in TLR3^–/– mice. Remarkably, we observed increased susceptibility of TLR3^–/– mice to WNV infection. TLR3 had a modest effect on WNV infection in peripheral tissues and was not required for the induction of a systemic IFN-/β response. The increased mortality was associated with elevated viral infection in neurons in the brain. Our experiments suggest that TLR3 has a protective role against WNV and restricts infection in neurons in the central nervous system (CNS).

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Viruses. The WNV strain (3000.0259) was isolated in New York in 2000 (11) and passaged once in C6/36 Aedes albopictus cells to generate a insect cell-derived stock (5 x 10⁷ PFU/ml) and one additional time in Vero T-144 cells to generate a mammalian-cell-derived stock (2 x 10⁷ PFU/ml).

Mouse experiments and quantitation of viral burden. C57BL/6 wild-type inbred mice were commercially obtained (Jackson Laboratories, Bar Harbor, ME). Congenic, backcrossed TLR3^–/– mice have been previously described and were obtained from R. Flavell (Yale University, New Haven, CT). All mice were genotyped and bred in the animal facilities of the Washington University School of Medicine, and experiments were performed in accordance with Washington University Animal Studies guidelines. Matched 8- to 12-week-old mice were used for all in vivo studies. WNV was diluted in Hanks balanced salt solution (HBSS) supplemented with 1% heat-inactivated fetal bovine serum (FBS) and inoculated by footpad or intraperitoneal injection in a volume of 50 or 100 µl, respectively. Intracranial (i.c.) inoculation was performed by injecting 10¹ PFU of WNV diluted in 10 µl of HBSS with 1% heat-inactivated FBS.

Quantification of tissue viral burden and viremia. To monitor viral spread in vivo, mice were infected with 10² PFU of insect cell-derived WNV by footpad inoculation and sacrificed at specific time points after inoculation. In other experiments, mice were infected with 10¹ PFU of insect cell-derived WNV by the i.c. route. After extensive cardiac perfusion with phosphate-buffered saline (PBS), organs were harvested, weighed, and homogenized, and virus was titrated by using a standard plaque assay as described previously (9). Viremia and viral burden in the lymph nodes were also measured by analyzing levels of positive-strand viral RNA levels using fluorogenic quantitative reverse transcription-PCR (qRT-PCR) as described previously (45) with the following primers and probe specific for the E gene of WNV: forward primer, 5'-TCAGCGATCTCTCCACCAAAG-3'; reverse primer 5'-GGGTCAGCACGTTTGTCATTG-3'; and probe, 5'-FAM-TGCCCGACCATGGGAGAAGCTC-3'-TAMRA (32). Lymph node viral RNA was normalized by measuring in parallel levels of 18S r RNA by qRT-PCR using commercially available primers and probes (Applied Biosystems, Foster City, CA).

Quantification of IFN activity. (i) L929 bioassay. Relative levels of biologically active IFN in serum were determined by using an EMCV L929 cytopathic effect bioassay as described previously (6). The concentrations of type I IFN were expressed as international units (IU) of IFN per ml and calculated by using a standard curve of recombinant IFN- (PBL Biomedical Laboratories) run in parallel in the bioassay.

(ii) IFN- and IFN-β mRNA by qRT-PCR. RNA was isolated from primary cells by using an RNeasy kit (Qiagen). IFN- and -β mRNA were measured by qRT-PCR using previously published primer sets (43). To analyze the relative fold induction of IFN- and β mRNA, 18S rRNA expression levels were also determined for normalization by using the C_T method (34).

(iii) IFN- and -β ELISA. A commercial capture enzyme-linked immunosorbent assay (ELISA) kit was used to measure levels of secreted IFN- and -β protein in cell supernatants (PBL Biomedical Laboratories).

WNV-specific antibody and CD8⁺ T-cell responses. The levels of WNV-specific immunoglobulin M (IgM) and IgG were determined by using an ELISA against purified WNV E protein (10). Intracellular IFN- staining was performed on day-7-infected splenocytes using a D^b-restricted NS4B peptide restimulation assay as previously described (41, 53). Samples were processed by two-color flow cytometry.

Leukocyte isolation from the CNS. Quantification of infiltrating CNS lymphocytes was performed by flow cytometry as previously described (50, 53). Briefly, wild-type and TLR3^–/– mouse brains were harvested on day 9 after infection, dispersed into a single-cell suspension with a cell strainer, and digested with 0.05% collagenase D, 0.1 µg of trypsin inhibitor TLCK/ml, 10 µg of DNase I/ml, and 10 mM HEPES (Life Technologies) in HBSS for 1 h. Viable cells were separated by discontinuous Percoll-gradient (70/37/30%) centrifugation for 30 min (850 x g at 4°C). Cells were counted and stained for CD4, CD8, CD45, and CD11b with directly conjugated antibodies (BD Pharmingen) for 30 min at 4°C and then fixed with 1% paraformaldehyde. The data collection and analysis were performed with a FACSArray flow cytometer and FloJo software.

Evaluation of BBB permeability. BBB permeability changes after WNV infection were determined by measuring Evans Blue diffusion into the CNS using a published protocol (20, 59). Briefly, mice were injected intraperitoneally with 800 µl of 1% (wt/vol) solution of Evans blue dye and perfused via intracardiac puncture with PBS 1 h later. Brains were subsequently removed, weighed, and stored at –80°C until further use. For Evans blue quantification, brains were homogenized in 1 ml of PBS, and proteins were precipitated with 1 ml of 100% trichloroacetic acid. The mixture was vortexed for 2 min and cooled for 30 min at 4°C. After centrifugation (30 min at 4,000 x g), the optical density of the supernatants were measured at 620 nm by using a spectrophotometer. The levels of Evans blue were expressed as micrograms of dye per gram of brain tissue by using a standard curve.

Primary cell culture and infection. (i) BM-M and BM-DC. Bone marrow-derived macrophages (BM-M) and bone-marrow-derived dendritic cells (BM-DC) were generated as described previously (45). Briefly, cells were isolated from the bone marrow of wild-type and TLR3^–/– mice and cultured for 7 days either in the presence of 40 ng/ml of macrophage colony-stimulating factor (PeproTech, Inc., Rocky Hill, NJ) to generate BM-M or in the presence of 20 ng/ml of granulocyte-macrophage colony-stimulating factor and 20 ng/ml of IL-4 (PeproTech) to generate BM-DC. Multistep virus growth curves were performed after infection at a multiplicity of infection (MOI) of 0.01 for BM-M or of 0.001 for BM-DC. Supernatants were titrated by plaque assay on BHK21-15 cells.

(ii) Fibroblasts. Mouse embryonic fibroblasts (MEFs) were generated from wild-type and TLR3^–/– 14-day-old embryos and maintained in Dulbecco modified Eagle medium supplemented with 10% FBS. Cells were used between passages 2 and 4 for all experiments. Multistep virus growth curves and IFN- and -β ELISA were performed after infection at MOIs of 0.001 and 0.1, respectively.

(iii) Cortical neurons. Primary cortical neurons were prepared from wild-type and TLR3^–/– mouse 15-day-old embryos as described previously (45). Cortical neurons were seeded in 24-well poly-D-lysine/laminin-coated plates in Dulbecco modified Eagle medium containing 5% heat-inactivated FBS and 5% horse serum for 24 h and then cultured for 4 days with Neurobasal medium containing B27 and L-glutamine (Invitrogen). Multistep virus growth curves and IFN- and -β protein quantitation were performed after infection at an MOI of 0.001.

Immunohistochemistry and confocal microscopy. Mice were infected with 10¹ PFU of WNV i.c. and sacrificed at day 4 postinfection. After perfusion with 10 ml of PBS and 10 ml of 4% paraformaldehyde, brains were harvested and fixed in 4% paraformaldehyde overnight at 4°C. Tissues were cryoprotected in 30% sucrose, and frozen sections were cut. Tissue staining was performed as previously described (30). Briefly, frozen brain sections were hydrated in PBS containing 10% normal goat serum and permeabilized with 0.1% Triton X-100. Staining was performed by incubating sections overnight at 4°C with the following primary antibodies: anti-WNV (rat immune serum or a mixture of mouse monoclonal antibodies E16 and E60 specific for the WNV E protein) (38), MAP-2 (microtubule associated protein 2; Chemicon), GFAP (glial fibrillary acidic protein; Dako), and CD11b (BD Biosciences). Primary antibodies were detected with secondary fluorescein isothiocyanate- or Cy3-conjugated goat anti-mouse, anti-rat, or anti-rabbit IgG (Dako). Nuclei were counterstained with TO-PRO3 (Molecular Probes). Fluorescence staining was visualized with a Zeiss 510 Meta LSM confocal microscope.

Statistical analysis. For in vitro experiments, an unpaired two-tailed Student t test was used to determine statistically significant differences. For viral burden analysis, differences in log titers were analyzed by the Mann-Whitney test. Kaplan-Meier survival curves were analyzed by the log rank test. All data were analyzed by using Prism software (GraphPadPrism4, San Diego, CA).

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
TLR3 prevents lethal WNV infection. Mice lacking IRF-3 show increased vulnerability to WNV infection due to enhanced viral replication in peripheral and CNS tissues (6). To begin to assess how the host recognizes WNV in vivo and triggers a protective IRF-3-dependent immune response, we examined the role of TLR3, one of the PRR upstream of IRF-3. TLR3^–/– and congenic wild-type C57BL/6 mice were infected via footpad inoculation with 10² PFU of a highly pathogenic New York WNV strain generated in C6/36 Aedes albopictus mosquito cells. Notably, TLR3^–/– mice were more vulnerable to WNV infection, with a 33% survival rate, compared to age-matched congenic wild-type mice, which had a 71% survival rate (P < 0.0001, Fig. 1A). Thus, an absence of TLR3 resulted in enhanced susceptibility to WNV. However, these results contrast with a previous study in which survival of WNV-infected TLR3^–/– mice was improved compared to wild-type mice (59). Since the disparity between studies could be explained by experimental differences, including the route of inoculation (subcutaneous versus intraperitoneal), the passage history of the virus (mammalian versus insect-cell derived), and/or the virus dose (10² PFU versus 10³ PFU), we pursued this discrepancy further. To assess the impact of the route of inoculation on mortality, wild-type and TLR3^–/– mice were infected with 10³ PFU of C6/36-derived virus via an intraperitoneal route; a higher WNV dose was used in accordance with the original TLR3 study (59). Notably, the susceptibility of TLR3^–/– mice to WNV after intraperitoneal infection also was enhanced with a 44% survival rate compared to wild-type mice, which had a 70% survival rate (P < 0.0001, Fig. 1B).

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FIG. 1. Survival analysis of wild-type and TLR3–/– C57BL/6 mice. (A and B) Eight- to twelve-week-old mice (n = 10 to 20) were inoculated with either 102 PFU of C6/36- or Vero-derived WNV via the subcutaneous route (footpad injection) (A) or with 103 PFU of C6/36- or Vero-derived WNV via the intraperitoneal route (B) and monitored for mortality for 21 days. Survival curves for TLR3–/– and wild-type mice were statistically significant independently of the virus stock or the route of infection (P < 0.0001).

To test the effect of passage history of the virus, we infected wild-type and TLR3^–/– mice subcutaneously with 10² PFU of WNV derived from monkey Vero cells. Infection with Vero cell-derived WNV did not significantly modify the survival pattern of wild-type or TLR3^–/– mice compared to that observed with C6/36 insect cell-derived WNV, since the increased susceptibility of TLR3^–/– mice was sustained (Fig. 1A). Similar results were observed with a higher dose (10³ PFU) of Vero cell-derived WNV administered through an intraperitoneal route (Fig. 1B). To exclude a dosing effect, wild-type mice were infected with higher doses (e.g., 10⁴ PFU) of insect cell or mammalian cell-derived WNV through each route, and yet still no significant difference in mortality was observed (data not shown). Thus, in contrast to what was seen previously (59), we observed a more vulnerable phenotype in TLR3^–/– mice after WNV infection by either subcutaneous or intraperitoneal inoculation routes. Taken together, our mortality data suggest that TLR3 is required for protection against lethal WNV infection.

TLR3 has a modest effect on WNV replication in peripheral tissues. Because TLR3 contributes to induction of the innate immune responses against viruses in cell culture and in vivo (54), we hypothesized that a deficiency in TLR3 in vivo would allow greater WNV replication. To evaluate this, mice were infected subcutaneously with 10² PFU WNV, and the viral burden was measured by fluorogenic qRT-PCR or viral plaque assay at days 1, 2, 4, 6, 8, and 10 in blood, peripheral organs (draining lymph nodes, spleen and kidney), and the CNS tissues (brain and spinal cord).

Although statistical significance was not reached, the levels of viral RNA in sera from TLR3^–/– mice trended slightly higher at days 2 and 4 after infection than in wild-type animals (Fig. 2A, P = 0.2). In lymph nodes, however, no difference in viral burden was observed at any time points (Fig. 2B, P > 0.5). Similar small differences in WNV RNA levels were detected systemically in TLR3^–/– mice after intraperitoneal infection with WNV (59).

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FIG. 2. Virological analysis of wild-type and TLR3–/– C57BL/6 mice. (A to F) Viral burden in peripheral and CNS tissues of wild-type and TLR3–/– C57BL/6 mice infected with 102 PFU of C6/36-derived viruses administered subcutaneously. WNV RNA in serum (A) and draining lymph node (B) and infectious virus in the spleen (C), kidney (D), brain (E), and spinal cord (F) were determined from samples harvested on days 2, 4, 6, 8, and 10 by qRT-PCR (A and B) or viral plaque assay (C to F). The data are shown as viral RNA equivalents or PFU per gram of tissue for 10 to 12 mice per time point. For viral load data, the solid line represents the median PFU per gram at the indicated time point, and the dotted line represents the limit of sensitivity of the assay. Asterisks indicate values that are statistically significant (*, P < 0.05; **, P < 0.005; ***, P < 0.0001) compared to wild-type mice.

A deficiency of TLR3 modified the kinetics of WNV infection in the spleen since modestly higher virus titers were observed at days 4 (6-fold, P < 0.005) and 6 (4-fold, P < 0.005) (Fig. 2C). By days 8 and 10, WNV was cleared from the majority of TLR3^–/– mice although, in contrast to wild-type mice, a few still had detectable infectious virus. Analogously, a modest but statistically significant increase of viral load in the spleen was observed previously in TLR3^–/– mice at day 6 after infection (59).

In wild-type C57BL/6 mice, little or no replication is detected in the kidney after WNV infection. In contrast, IFN-/βR^–/– and IRF-3^–/– mice sustain virus infection in the kidney, suggesting that innate immunity restricts WNV tissue tropism (6, 43). Although not statistically significant, a subset (5 of 10) of TLR3^–/– mice showed evidence of modest viral replication in the kidney at day 4 (Fig. 2D), with lower levels detected sporadically at days 6, 8, and 10 after infection. Thus, TLR3 appears to be partially responsible for the IRF-3-dependent restriction of WNV infection in the kidney.

WNV spread more rapidly to the CNS in TLR3^–/– mice. Low but detectable levels of infectious WNV were observed in the brains of approximately half (5 of 9) of the TLR3^–/– mice at day 4 after infection, whereas virus was not detected in the brains from wild-type mice at this time. By day 6, significantly higher (30-fold, P < 0.0001) WNV titers were measured in brains from TLR3^–/– mice than in brains from wild-type mice (Fig. 2E). Enhanced viral replication was observed in the brains of TLR3^–/– mice at days 8 and 10, although this did not attain statistical significance. A similar pattern was observed in the spinal cord, where earlier entry was detected in TLR3^–/– mice at day 6 (Fig. 2F). A trend toward enhanced viral replication was also observed at days 8 and 10 in the spinal cord of TLR3^–/– mice. Collectively, these data suggest that a deficiency of TLR3 results in increased WNV burden in peripheral and CNS tissues, although more modestly and variably than that observed in IRF-3^–/– mice (6).

Antibody and CD8⁺ T-cell responses are unchanged in TLR3^–/– mice after WNV infection. TLR3 stimulation has been shown to promote B-cell activation (2) and cross-priming of virus-specific T cells (17, 22, 48). Since a depressed antiviral antibody response leads to rapid and sustained dissemination of WNV into the CNS (9, 10), we evaluated whether a deficiency of TLR3 modulated the levels of WNV-specific IgM and IgG. Notably, similar levels of WNV-specific antibodies were detected in wild-type and TLR3^–/– mice at two time points after infection (Fig. 3A and B). Thus, the phenotype observed in TLR3^–/– mice is unlikely to be due to a defect in B-cell priming. Since CD8⁺ T cells are also required for the control and clearance of WNV in the CNS (50), we evaluated whether a deficiency in TLR3 altered WNV-specific CD8⁺ T-cell priming. Splenocytes from WNV-infected wild-type or TLR3^–/– mice were harvested at day 7 after infection and restimulated ex vivo specifically with a D^b-restricted immunodominant NS4B peptide (41, 53) or with phorbol ester (phorbol myristate acetate [PMA]) and ionomycin. Activation was measured by intracellular staining of IFN- in CD8⁺ T cells using flow cytometry. Restimulation with the WNV-specific peptide or PMA and ionomycin resulted in a similar percentage and absolute number of CD8⁺ T cells expressing IFN- in both wild-type and TLR3^–/– mice (Fig. 3C, P > 0.4). Thus, the absence of TLR3 did not significantly affect WNV-specific CD8⁺ T-cell activation, and the higher viral burden in the CNS of TLR3^–/– mice is not a consequence of inadequate priming of adaptive immune responses.

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FIG. 3. WNV-specific antibody responses and CD8+ T-cell activation in wild-type and TLR3–/– mice. (A and B) Wild-type and TLR3–/– mice were infected with WNV, and serum was collected at the indicated time points. The development of specific IgM (A) or IgG (B) against WNV was determined by ELISA using purified WNV E protein. The data are the average of at least five mice per time point. (C) Wild-type and TLR3–/– mice were infected with WNV, and splenocytes were harvested at day 7 after infection. The percentage of IFN--producing CD8+ T cells after ex vivo restimulation with a WNV-specific NS4B peptide or PMA and ionomycin was determined by flow cytometry. n.d, values that were below the limit of detection of the assay; ns, differences that were not statistically different. (D) Trafficking of inflammatory cells into the CNS after WNV infection. Wild-type and TLR3–/– mice were infected with 102 PFU of insect cell-derived WNV, and brains were harvested after extensive perfusion at day 9 after infection. CNS leukocytes were isolated after by Percoll centrifugation and analyzed by flow cytometry. The absolute number of specific inflammatory cells (CD4+ T, CD8+ T, CD45+, and CD11b+ cells) in the brains of wild-type and TLR3–/– mice after WNV infection were calculated and reflect an average of five mice per group.

TLR3^–/– mice show no defect in accumulation of inflammatory cells in the CNS. Although CD8⁺ T-cell priming in the spleen was normal in TLR3^–/– mice, because the viral burden accumulated at a late stage in pathogenesis, we speculated there might be a leukocyte trafficking defect into the CNS. Inefficient migration of leukocytes in the CNS results in sustained WNV burden in the brain and vulnerability to lethal infection in several genetically deficient mice (18, 30, 53). To address this, leukocytes were recovered from the brains of wild-type and TLR3^–/– mice at day 9 after extensive perfusion and analyzed by flow cytometry. Notably, similar numbers of CD45^low/CD11b^high activated microglia, CD45^high/CD11b^high macrophages, CD4⁺ T cells, and CD8⁺ T cells were observed in the brains of WNV-infected wild-type and TLR3^–/– mice (Fig. 3D, P > 0.2). Thus, a deficiency of TLR3 in vivo did not significantly alter leukocyte recruitment and activation in the brain and is unlikely to explain the enhanced viral load in the CNS of TLR3^–/– mice.

A deficiency of TLR3 does not alter BBB permeability. Since the previous study with WNV infection in TLR3^–/– C57BL/6 mice showed altered BBB permeability leading to enhanced CNS spread (59), we assessed whether in our model with insect cell-derived virus, a deficiency of TLR3 modulated BBB permeability and, possibly, earlier entry in the CNS. Wild-type and TLR3^–/– mice were infected subcutaneously with 10² PFU of insect cell-derived WNV and administered Evans blue dye intraperitoneally at various time points after infection. BBB permeability was determined by quantifying the relative level of extravasated dye in brains after tissue homogenization and spectrophotometry. Somewhat surprisingly, in wild-type mice, little change in BBB permeability was observed at days 3, 5, and 7 after WNV infection (Fig. 4), results that agree with a recent study showing no correlation between increased BBB permeability and WNV-induced lethality for wild-type C57BL/6 and BALB/c mice (37). Similarly, we observed no change in BBB permeability in TLR3^–/– after WNV infection. In our experimental model, altered BBB permeability after WNV infection between wild-type and TLR3^–/– mice was not readily apparent and likely does not explain the enhanced WNV infection phenotype in the CNS of TLR3^–/– mice.

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FIG. 4. Effect of TLR3 on BBB permeability upon WNV infection. Wild-type and TLR3–/– mice were infected with 102 PFU of insect cell-derived WNV via subcutaneous injection and administered a 1% Evans blue solution at the indicated time points. The levels of Evans blue in whole brains were quantified by measuring the absorbance at 620 nm by spectrophotometry after tissue homogenization and precipitation. The data are the average of five mice per time point.

A deficiency of TLR3 does not affect IFN induction in the draining lymph node or in circulation. In vitro studies suggest that TLR3, in concert with RIG-I and MDA5, recognizes viral RNA and induces IFN through the activation of the transcriptional regulator IRF-3. Since IRF-3 is required for the generation of a normal IFN response in draining lymph nodes after WNV infection (6), we tested whether TLR3 signaling contributes to this. Notably, the levels of IFN- and β mRNA gradually and equivalently increased after WNV infection in both wild-type and TLR3^–/– mice (Fig. 5A and B). Thus, TLR3 appears to be dispensable for induction of the type I IFN response in the draining lymph node following WNV infection.

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FIG. 5. IFN induction in draining lymph node and serum of wild-type and TLR3–/– mice infected with WNV. (A and B) Mice were inoculated with 102 PFU of WNV by footpad injection and sacrificed at the indicated times. Total RNA from the draining lymph was analyzed for IFN- (A) and IFN-β (B) mRNA expression by qRT-PCR. The data are normalized to 18S rRNA and are expressed as the relative fold increase over normalized RNA from uninfected controls. Average values are from 5 to 12 mice per time point, and error bars indicate the standard deviations. (C) IFN activity was determined from serum collected on days 1 to 4 after infection by an EMCV protection bioassay in L929 cells. Serum was added to L929 cells and, after subsequent infection with EMCV, the cytopathic effect was measured. The data reflect the average of serum samples harvested from 5 to 10 mice per time point, and the data are expressed as IU of IFN-/ml. The specificity of the assay was confirmed with an anti-IFN-/β receptor neutralizing MAb (data not shown).

Since a prior study with WNV demonstrated that TLR3 signaling augments IFN-β mRNA and TNF- protein production in blood (59), we assessed the systemic levels in wild-type and TLR3^–/– mice after subcutaneous infection with 10² PFU of WNV. Using an EMCV bioassay, we observed similar levels of type I IFN activity in the serum of wild-type or TLR3^–/– mice after infection (Fig. 5C). Moreover, no differences in TNF- serum levels were observed in wild-type or TLR3^–/– mice, since all samples clustered near the limit of detection (25 pg/ml) by ELISA or cytokine bead array assays (data not shown); these relatively low values of TNF- in serum after WNV infection also are consistent with previous studies (18, 51). Collectively, a deficiency of TLR3 did not modify systemic production of type I IFN and TNF- in mice in this model.

TLR3 does not modulate WNV replication and IFN induction in primary myeloid and fibroblast cells. Diverse cell populations can differently utilize PRR to induce innate immune responses (reviewed in references 1 and 39). Since these more subtle effects may not be apparent in whole-tissue viral burden experiments, we assessed whether an absence of TLR3 affected WNV replication and/or IFN induction in different primary cells. MEFs, BM-M, and myeloid dendritic cells (BM-DC) were generated from wild-type and TLR3^–/– mice and infected with WNV. Although IRF-3^–/– cells supported increased replication (6), no difference in viral growth was observed between wild-type and TLR3^–/– MEFs, BM-DC, and BM-M (Fig. 6A, D, and G). Consistent with this, IFN-/β protein secretion by WNV-infected MEF and BM-DC was virtually identical in WNV-infected wild-type and TLR3^–/– cells (Fig. 6B, C, E and F). Because IFN-/β protein production by WNV-infected BM-M was below the level of detection, we measured IFN-/β mRNA levels in these cells. Similar levels of IFN-/β mRNA were detected in wild-type and TLR3^–/– cells (data not shown). Thus, a deficiency of TLR3 does not compromise IFN production in MEF, BM-M, and BM-DC after WNV infection.

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FIG. 6. Effect of TLR3 on WNV infection and IFN induction in primary fibroblasts, dendritic cells, and macrophages. MEFs (A), BM-DC (D), and BM-M (G) generated from wild-type or TLR3–/– mice were infected with WNV, and virus production was evaluated at the indicated times postinfection by plaque assay. IFN- and β levels were quantified by a capture ELISA from the supernatants of WNV-infected MEFs (B and C) and BM-DC (E and F). Values are an average of quadruplicate samples generated from at least three independent experiments.

Effect of TLR3 on WNV infection and IFN induction in primary cortical neurons. Infection of neurons by several RNA viruses, including WNV, induces type I IFN that could serve an antiviral role (6, 8). Previous studies revealed an IRF-3-dependent restriction of WNV infection and regulation of IFN production in neurons (6). To evaluate whether this occurred through a TLR3-dependent recognition pathway, we analyzed viral growth kinetics and IFN production in WNV-infected primary cortical neurons cultures from wild-type and TLR3^–/– mice (Fig. 7). Consistent with what was observed for IRF-3^–/– neurons (6), small, yet statistically significant differences in viral growth were observed at 24 h (1.5-fold increase, P < 0.05) and 48 h (3-fold increase, P < 0.0001) after WNV infection. In contrast, no significant decrease in IFN- and -β protein production was observed in either wild-type or TLR3^–/– neurons (P > 0.3); indeed, IFN-β production was modestly enhanced in TLR3^–/– cells at 24 h (3-fold increase, P = 0.0003). Thus, in primary cortical neurons TLR3 is not essential for regulating IFN gene induction but, nevertheless, exerts a subtle restriction of WNV replication.

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FIG. 7. Role of TLR3 on WNV infection and IFN production in primary cortical neurons. (A) Primary cortical neurons generated from wild-type or TLR3–/– mice were infected at an MOI of 0.001, and virus production was evaluated at the indicated times by plaque assay. (B and C) The production of IFN- (B) and IFN-β (C) proteins by WNV-infected cortical neurons was analyzed by ELISA. Values are an average of triplicate samples generated from three independent experiments. Asterisks indicate values that are statistically significant (*, P < 0.05; **, P < 0.005; ***, P < 0.0001).

TLR3 regulates WNV replication in the CNS. Although virus titers in the CNS after peripheral WNV infection in TLR3^–/– mice were elevated, it was unclear whether this effect was due directly to an increased permissiveness of neurons or indirectly through earlier CNS dissemination. To determine whether TLR-3 was required by neurons to restrict WNV in vivo, we directly evaluated WNV replication in CNS tissues. Wild-type and TLR3^–/– mice were infected with 10¹ PFU of WNV via an i.c. route, and the viral burden in the brain and spinal cord was measured on days 2 and 4. Although no significant difference was observed at day 2, TLR3^–/– mice developed a 40-fold-higher viral burden in the brain (10^7.2 and 10^5.6 PFU/g for TLR3^–/– and wild-type mice, respectively; P < 0.0001) and a 6-fold higher viral burden in the spinal cord (10^4.8 and 10^4.0 PFU/g for TLR3^–/– and wild-type mice, respectively; P < 0.05) (Fig. 8). Thus, TLR3 limits WNV infection in the brain and spinal cord after i.c. infection.

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FIG. 8. TLR3 controls viral replication in the CNS after i.c. WNV infection. Wild-type or TLR3–/– mice were inoculated with 101 PFU of WNV by i.c. inoculation. Brains (A) and spinal cord (B)s were harvested at the indicated time points, and the virus titers were determined as described in Fig. 2 (*, P < 0.05; ***, P < 0.0001).

To further define the role of TLR3 in controlling viral replication in the CNS, we examined the cellular distribution of WNV antigen in the brain by immunohistochemistry and confocal microscopy using antibodies against WNV and specific CNS cell types. Significantly higher numbers of WNV-infected cells were observed in the cerebral cortex and hippocampus in TLR3^–/– mice (Fig. 9). The majority of infected cells were neurons, since prominent costaining was observed between MAP-2 and WNV antigen (Fig. 9A). In contrast, no costaining was detected between GFAP⁺ astrocytes in either wild-type or TLR3^–/– mice (Fig. 9B). Finally, a small subset of CD11b⁺ myeloid cells costained positive with WNV antigen in wild-type mice, although there was no apparent increase in number in TLR3^–/– mice (Fig. 9C). Taken together, WNV has a primary tropism for neurons, and a deficiency of TLR3 enhances its ability to infect susceptible neuron populations.

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FIG. 9. TLR3 controls WNV replication in the CNS by restricting infection of neurons. (A to C) Brains sections from wild-type and TLR3–/– mice that were infected i.c. with WNV were costained for WNV antigen (red), the nuclear stain TO-PRO3 (blue) and the neuronal marker MAP-2 (green) (A), the astrocyte-specific marker GFAP (green) (B), or the microglial/macrophage cell-specific marker CD11b (green) (C). White arrows indicate double-positive cells. The data are representative of sections from at least five wild-type or TLR3–/– mice.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Protection against WNV infection requires rapid detection of the virus and induction of an IFN response. Studies in MEFs suggested that host cells can sense WNV infection via the cytosolic helicase RIG-I and to a lesser extent through MDA5 (15, 16). The role of TLR3 in inducing protective antiviral responses against WNV and other viruses has remained less clear. Our experiments demonstrate that TLR3 contributes to protection against lethal WNV infection in mice, although the phenotype is more subtle than observed with RIG-I^–/– (M. Suthar and M. Gale, unpublished results) or IRF-3^–/– (6) mice. TLR3^–/– mice showed increased viral burden in the CNS after subcutaneous inoculation, although only modest differences were observed in peripheral lymphoid, intravascular, and visceral compartments. Cell culture studies also showed little difference in viral growth kinetics between wild-type and TLR3^–/– MEF, BM-M, and BM-DC. Despite its position upstream of the transcriptional regulator IRF-3, an absence of TLR3 did not alter the induction of type I IFN after WNV infection in cells or in vivo, which is consistent with a previous report (21). A lack of TLR3, however, enhanced viral replication in neurons in culture and in vivo. Thus, TLR3 limits WNV infection in a cell type-restricted manner.

Some of our results contrast with a previous study in TLR3^–/– mice that used a mammalian cell-passaged WNV. In that report, TLR3 signaling had immunopathological consequences since WNV-infected TLR3^–/– mice showed reduced cytokine (e.g., TNF- and IL-6) responses, BBB permeability, neuroinvasion, and mortality compared to wild-type mice (59). Nonetheless, some similarities among models were observed; in both studies, the absence of TLR3 resulted in modest increases in viral burden in peripheral tissues. The primary difference among models was the distinct systemic cytokine response and its effects on BBB penetration by WNV. An additional difference reflected the baseline mortality after WNV infection of similarly aged C57BL/6 mice: Wang et al. (59) reported 100% mortality after intraperitoneal infection compared to the substantially lower (ca. 20 to 30%) mortality seen in our model after subcutaneous or intraperitoneal infection. Since the two studies used North American WNV strains that were virtually identical (99.9% at the amino acid level) (3, 11), the disparity of outcome in TLR3^–/– mice could in theory be explained by the distinct route of inoculation (subcutaneous versus intraperitoneal), passage history of the virus (mammalian Vero cell versus insect-cell derived), and/or the virus dose (10² PFU versus 10³ PFU).

The route of peripheral administration could influence the pathogenicity of WNV infection. Natural infection of vertebrates is usually acquired through peripheral inoculation by the bite of an infected mosquito. This subcutaneous infection may stimulate a local immune response in the skin and draining lymph node, possibly via activation of DC and T cells, leading to an early type I and II IFN response that restricts dissemination (44, 51). In contrast, intraperitoneal or intravenous infection may lead to rapid dissemination and systemic inflammation. Nevertheless, we did not observe any significant effect of the peripheral inoculation route on mortality since wild-type mice infected via intraperitoneal and subcutaneous routes showed similar sublethal phenotypes.

Recent data have also suggested that the cell type used to prepare the viral stock may directly modulate the intrinsic innate immune response of a cell. Type I IFN was more potently induced in myeloid dendritic cells after infection with alphaviruses produced in mammalian cell compared to mosquito cell-derived virus (49). Similarly, mosquito cell-derived WNV or envelope protein, which displays N-linked carbohydrates in a high mannose form, inhibited the poly(I-C)-induced production of TNF- and IFN-/β, whereas Vero cell-derived WNV, which displays complex processed sugars, did not (4). Analogously, Vero but not insect cell-derived WNV induced IFN- and the inflammatory chemokine CXCL10 in plasmacytoid dendritic cells (52). Cell-specific differences in infectious particle-to-antigen ratios or the content of RNA-containing defective viral particles also could independently affect the induction of innate immune responses. For example, under certain virus propagation conditions, Vero cell-passaged virus has a high particle-to-PFU ratio, presumably because of the presence of large numbers of noninfectious or defective viral particles (7, 31); this may have an adjuvant-like effect in stimulating a cytokine response. Alternatively, differences in NS1 content of the virus stocks could also affect immune responses, since NS1 has been recently suggested to inhibit TLR3 signal transduction (60). WNV- and Japanese encephalitis virus-infected Vero cells but not insect cells secrete large amounts of NS1 (35, 52). Thus, WNV that is propagated in different host cells may induce distinct inflammatory profiles, which alter pathogenesis in vivo. However, in our experiments, infection of wild-type mice with either insect cell- or mammalian cell-derived virus did not alter mortality, and the enhanced susceptibility of TLR3^–/– mice for WNV was sustained with both viruses. Using our virus propagation conditions, WNV derived from mosquito or mammalian cells did not induce an inflammatory response that exceeded a pathological threshold resulting in early neuroinvasion. Consistent with this, we observed no difference in TNF- levels or BBB permeability between wild-type and TLR3^–/– mice. However, this does not rule out the possibility that viral stocks propagated in insect cell or mammalian cells according to different protocols (time to harvest, serial passage, content of defective particles and cellular debris) could induce pathological levels of inflammation.

Although TLR3 appears to be less important for controlling WNV in most peripheral tissues and cells, our studies point to a more unique role in neurons. Although other PRRs, including RIG-I and MDA5, may play a dominant role in triggering the innate immune response and ISG expression in some cells during WNV infection (16), in neurons, TLR3 appears to be important for suppressing WNV replication. TLR3 is expressed constitutively in several CNS cell types, including neurons, astrocytes, and microglia (14, 40, 56), and is upregulated in vivo in human Purkinje cells following rabies virus and HSV infections (27). Moreover, human neurons activate TLR3-dependent antiviral responses after exposure to rabies virus and HSV infections (40). Although many CNS cells express TLR3, it may be required in neurons for virus control: enhanced WNV infection of susceptible neuronal populations rather than a broader cell tropism was seen in brains from TLR3^–/– mice. Interestingly, cultures of primary TLR3^–/– cortical neurons did not show impairment of an IFN-/β response despite modestly increased viral replication. This contrasts with microglia (56) and astrocytes (29), which show blunted cytokine responses after poly(I-C) treatment of TLR3^–/– cells. Although more detailed WNV infection studies with additional CNS cells types are necessary, our data suggest that TLR3 in neurons is necessary to restrict WNV infection. The exact mechanism(s) by which TLR3 is engaged by WNV to exert this control is currently under investigation.

In summary, TLR3 is essential for restriction of WNV infection in neurons and protection from lethal encephalitis. Cell-specific TLR3 responses regulate immune responses in vivo against WNV and possibly other viruses. A better understanding of the mechanisms that govern induction of protective immune responses may provide novel therapeutic strategies against viral pathogens.

 
We thank R. Flavell for the TLR3^–/– mice and G. Sen for the ISG54 antibody.

This study was supported by a Predoctoral Fellowship from the Howard Hughes Medical Institute (M.A.S.), the NIH (AI057568 [M.G.]), and the State of Washington Endowment (M.G.).

 
* Corresponding author. Mailing address: Departments of Medicine, Molecular Microbiology, and Pathology and Immunology, Washington University School of Medicine, 660 South Euclid Avenue, Box 8051, St. Louis, MO 63110. Phone: (314) 362-2842. Fax: (314) 362-9230. E-mail: diamond{at}borcim.wustl.edu

Published ahead of print on 20 August 2008.

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Journal of Virology, November 2008, p. 10349-10358, Vol. 82, No. 21
0022-538X/08/$08.00+0     doi:10.1128/JVI.00935-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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