J Virol 89(20): 10219-10229

PMC: PMC4580164

PMID: 26223644

RIG-I Mediates an Antiviral Response to Crimean-Congo Hemorrhagic Fever Virus

Christina Spiropoulou

Éric Bergeron

INTRODUCTION

Viral infection of mammalian cells leads to the activation of downstream signaling cascades and the subsequent induction of type I interferon (IFN) and proinflammatory cytokines to restrict viral replication. To initiate the antiviral response, infection must be recognized by the cell via pattern recognition receptors (PRRs). One major class of receptors, often implicated in the recognition of RNA viruses, are the retinoic acid-inducible gene I (RIG-I)-like receptors (RLRs) (1). Major efforts have led to the identification of RLRs involved in the sensing of foreign RNA and of the biochemical and structural determinants for foreign RNA recognition. RLRs are believed to be the core proteins responsible for activating proinflammatory and antiviral cytokine genes, as well as cytokine-stimulated genes, in a broad range of cells in response to RNA virus infection (2).

The RLR family of DexD/H box RNA helicases consists of RIG-I, melanoma differentiation-associated factor 5 (MDA5), and laboratory of genetics and physiology 2 (LGP2). All 3 RLR family members contain a central helicase domain with ATPase activity (3) and a regulatory domain at the C terminus (3, 4). ATPase activity is involved in oligomerizing RIG-I and in allowing MDA5 to release the viral RNA. Upon binding to viral RNA, RIG-I and MDA5 oligomerize on the RNA in a double-stranded RNA (dsRNA) length-dependent manner (5,–7). RNA binding, initiated by the C-terminal RNA binding domain of RIG-I, triggers ATP hydrolysis and the release of caspase activation and recruitment domains (CARD), which contains the signaling module, from its repressed conformation (8, 9). Through their N-terminal CARDs, RIG-I and MDA5 then signal to the adaptor mitochondrial antiviral signaling (MAVS) protein, localized on the outer mitochondrial membrane, to activate transcription factors, including IFN response factor 3 (IRF3), IRF7, and nuclear factor κB (NF-κB), resulting in IFN gene induction (10).

Crimean-Congo hemorrhagic fever virus (CCHFV), a negative-strand RNA virus, is a member of the Nairovirus genus (family Bunyaviridae). Similar to most bunyaviruses, CCHFV's genome consists of 3 segments, large (L), medium (M), and small (S), which encode an RNA-dependent RNA polymerase, surface glycoproteins, and a nucleocapsid (N) protein, respectively. The L segment of CCHFV is almost twice as long as that of other bunyaviruses and contains an ovarian tumor (OTU) domain, which is thought to suppress immune signaling (11, 12). CCHFV is tick-borne and generally circulates in nature in an enzootic tick-vertebrate-tick cycle. CCHFV can infect a wide range of domestic and wild animals; however, CCHF naturally occurs exclusively in humans, often manifesting as severe hemorrhagic disease (13) with case fatality rates upwards of 30%. To date, the immune recognition receptors of CCHFV have not been characterized. The CCHFV genome has a 5′ monophosphate (5′p) end (14) which mimics mammalian ribosomal and tRNA. Since RIG-I is believed to strongly recognize only 5′ppp and 5′pp ends, it has not been implicated in CCHFV recognition.

We investigated putative RLR involvement in CCHFV recognition. Interestingly, while previous evidence suggested the exclusion of RIG-I recognition, we found RIG-I to be a critical mediator of the type I IFN antiviral response to CCHFV. Our work serves as the first description of a pattern recognition receptor activating signaling pathways in response to nairovirus infection.

MATERIALS AND METHODS

Cells and viruses.

Human lung carcinoma cells (A549), African green monkey kidney cells (Vero-E6), human embryonic kidney (HEK-293) cells, HEK-293 cells expressing IFN-dependent luciferase reporter (293T-IFN-β-FF), and all murine embryonic fibroblast (MEF) lines (wild type [WT], RIG-I^{, and RIG-I knockout cells expressing hRIG-I) were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal calf serum, 1%}l-glutamine, and 1% penicillin-streptomycin. Virus infectivity was measured by titration and calculated as the median tissue culture infective dose (TCID₅₀). Rift Valley fever virus (RVFV) rZH501-ΔNSs:GFP-ΔNSm was rescued and propagated in a biosafety level 3 (BSL-3) laboratory as previously described (15). WT CCHFV (IbAr10200 strain) was rescued using reverse genetics in BSR/T7 cells in a BSL-4 laboratory. Recombinant CCHFV stocks were obtained after 2 passages in human adrenal carcinoma cells (SW13) cultured in DMEM supplemented with 10% fetal bovine serum (FBS) and 1% sodium pyruvate. In vitro growth kinetics of recombinant CCHFV were characterized and were undistinguishable from those of the parental virus isolate from Nigeria (16).

ELISA.

For IFN-β enzyme-linked immunosorbent assay (ELISA), supernatant samples were collected at the indicated time points and γ-irradiated prior to subsequent analysis outside the BSL-4 laboratory. Secreted levels of human IFN-β were quantified using the VeriKine or VeriKine-HS human interferon beta ELISA kit (both from PBL Assay Science, Piscataway, NJ) according to the manufacturer's recommendations. The absorbance of individual test samples was converted into picograms per milliliter using a standard curve generated by serially diluting the standard in the same plate.

Antibodies and immunofluorescence.

The following antibodies were used in this study: CCHFV hyperimmune mouse ascitic fluid (HMAF; Cell Signaling Technology, Danvers, MA), diluted 1:500; anti-MAVS/Cardif (C-term; EMD Millipore, Billerica, MA), diluted 1:500; anti-RIG-I D14G6 (Cell Signaling Technology), diluted 1:500; anti-MDA5 D74E4 (Cell Signaling Technology), diluted 1:250; anti-STAT-1 (Tyr701; BD Transduction Laboratories), diluted 1:500; anti-pSTAT1 (pY701; BD Transduction Laboratories, Franklin Lakes, NJ), diluted 1:625; Alexa Fluor 488 goat anti-rabbit IgG (H+L; Life Technologies); and Alexa Fluor 594 goat anti-mouse IgG (H+L; Life Technologies). For MEF immunofluorescence, fixed cells were stained for 2 h at room temperature with anti-CCHFV HMAF, washed with PBS, and stained with Alexa Fluor 488 goat anti-mouse IgG (H+L) diluted 1:333. Cell nuclei were stained with NucBlue live cell stain, and cytoplasm was stained with HCS CellMask red stain (both from Life Technologies) for 30 min at 37°C. Quantification of stained cells was performed using an Operetta high-content imaging system with Harmony high-content imaging and analysis software (PerkinElmer Inc., Waltham, MA).

Quantitative RT-PCR (qRT-PCR).

A549 cells were infected with CCHFV, and cell lysates were collected 1, 8, 12, 24, 48, and 72 hpi. RNA was isolated using the MagMAX-96 total RNA isolation kit (Life Technologies). RNA then was analyzed by a SYBR green-based reverse transcription-PCR (RT-PCR) using the SuperScript III platinum SYBR green one-step quantitative PCR (qPCR) kit with ROX (Life Technologies) on a 7500 real-time PCR system (Applied Biosystems/Life Technologies) and standardized using β-actin as a cell lysate control. Primers for type I IFN genes, proinflammatory cytokines, and RLR were designed as previously described and included hIFN-β (17), hISG56 (18), hMxA (18), hIL-6 (19), hIL-8 (20), hRANTES (18), hRIG-I (21), hMDA5 (22), and hMAVS (IPS-1) (22). To determine CCHFV S segment copy numbers, RNA was isolated and a one-step qRT-PCR was conducted using an Applied Biosystems 7500 real-time PCR system with CCHFV S segment-specific primers (sense, 5′-ATGAACAGGTGGTTTGAAGAGTT-3′; antisense, 5′-TGGCACTGGCCATCTGA-3′) and a fluorescently labeled TaqMan probe (5′-6-carboxy-fluoresceinTGTCCAAATTGGGAACACTCTCGCA-BlackBerry Quencher-3′) (all from TIB Molbiol, Adelphia, NJ). A 124-nucleotide CCHFV fragment was amplified in triplicate for each sample using a SuperScript III platinum one-step qRT-PCR kit with ROX (Life Technologies) according to the manufacturer's instructions.

Western blot analysis.

Cells were lysed in sample buffer (2% SDS, 20 mM Tris, 5 mM NEM, and 1 mM dithiothreitol [DTT]). Prior to analysis, NuPAGE loading buffer and reducing agent (Life Technologies) were added and the samples were boiled briefly. Equal volumes of cell lysate were separated by Bis-Tris polyacrylamide gel electrophoresis in the presence of reducing buffer antioxidant (Life Technologies) and transferred to a nitrocellulose membrane using the iBlot system (Life Technologies). Membranes were processed with target-specific primary antibodies, visualized with species-specific horseradish peroxidase (HRP)-conjugated secondary antibody, and developed using SuperSignal West Dura fast Western blotting kits (Fisher-Thermo Scientific, Grand Island, NY) according to the manufacturer's instructions. Protein imaging and quantification were performed using a FluorChem system and AlphaView software (both from ProteinSimple, San Jose, CA). Protein levels were standardized to those of actin.

siRNA transfections.

A549 cells were cultured to 50% confluence in 6-well plates for virus growth experiments, in 24-well plates for protein analysis, or in 96-well plates for RT-PCR gene expression analysis. Cells were reverse transfected and then retransfected 24 h later using a 1:1 ratio of Lipofectamine RNAiMAX (Life Technologies) and specific short interfering RNA (siRNA) against RIG-I (Santa Cruz Biotechnology, Dallas, TX), IFIH-1/MDA5 (ON-TARGETplus smart siRNA pool from GE Dharmacon), MAVS, or control siRNA (Santa Cruz Biotechnology or Dharmacon, Lafayette, CO). The amount of siRNA used was 60 pmol/well, 10 pmol/well, or 3 pmol/well for 6-, 24-, and 96-well plates, respectively. Forty-eight hours after the initial transfection, cells were infected with CCHFV or control viruses at the indicated MOIs.

Virion RNA and luciferase reporter gene assays.

RNA was isolated from either the cell lysate (total RNA) or the supernatant (virion RNA) of infected cells. SW13 cells or Vero-E6 cells were infected with CCHFV (IbAr10200 strain) or RVFV (rRVFV-ΔNSs:GFP-ΔNSm strain), respectively, and lysates and supernatant were harvested at 48 hpi. Supernatants were pelleted by ultracentrifugation, and samples were extracted in Tripure isolation reagent (Life Technologies). Where indicated, RNA was digested with APex heat-labile alkaline phosphatase (Epicentre Technologies, Madison, WI), RNA 5′ polyphosphatase (Epicentre Technologies), or RNase V1 (Ambion/Life Technologies) according to the manufacturer's directions and purified using phenol and chloroform. 293T-IFN-β-FF-ISRE-RFP dual reporter (293T-IFN-β-FF) cells (48) were transfected with the indicated amounts of RNA using Lipofectamine 2000 (Life Technologies). Cells were lysed 24 h posttransfection, and luciferase activity was measured. Firefly luciferase activity was expressed either as relative luciferase units (RLU) or as percent activity of respective negative-control digests to minimize absolute value variability between independent experiments. Optimization experiments revealed peak immunostimulation with 50 ng of transfected CCHFV virion RNA (vRNA). Increasing CCHFV input RNA up to 5-fold did not increase the IFN response under the conditions used (data not shown).

Coimmunoprecipitation of RIG-I-RNA complexes and RNA extraction.

As previously described (5), HEK-293T cells were transfected with pCAGGS plasmid encoding hemagglutinin (HA)-tagged RIG-I. Cells were lysed 48 h later with polysome lysis buffer consisting of 10 mM HEPES (pH 7.0), 100 mM KCl, 5 mM MgCl₂, 2.5 mM EDTA, and 1% NP-40 (Sigma-Aldrich, St. Louis, MO), supplemented with 50 U/ml Superase-IN RNase inhibitor (Ambion/Life Technologies) and protease inhibitor cocktail (Roche Life Science, Indianapolis, IN). Supernatants from centrifugation-clarified lysates were diluted 1:5 in NT2 buffer containing 50 mM Tris (pH 7.4), 150 mM NaCl, 1 mM MgCl₂, and 0.05% NP-40 supplemented with 50 U/ml Superase-IN and 1 mM DTT and precleared using protein G agarose beads. Precleared samples were incubated with 50 μg of total RNA from CCHFV-, RVFV-, or mock-infected SW13 cells at 4°C for 30 min to allow binding to RIG-I. RIG-I RNPs were immunoprecipitated using anti-HA tag antibody, anti-RIG-I antibody, or anti-GFP antibody (all from Abcam, Cambridge, MA) bound to protein G agarose beads in NT2 buffer supplemented with 50 U/ml Superase-IN and 1 mM DTT. RNA was isolated from the bead-bound RNP complexes with 100 μg/ml proteinase K (Ambion) at 55°C for 30 min. RNA was purified using acid–phenol-chloroform (pH 4.5) solution (Life Technologies), precipitated using ethanol and sodium acetate, and resuspended in 40 μl distilled water. Ten microliters of this RNA was used to transfect 293T-IFN-β-FF cells.

Statistical analysis.

Data were analyzed using GraphPad Prism version 6.0c (GraphPad, LaJolla, CA), by one-way analysis of variance (ANOVA) with a Bonferroni multiple comparison posttest, or by parametric t test with Welch's correction to determine significant differences (P < 0.05) between samples.

Cells and viruses.

ELISA.

Antibodies and immunofluorescence.

Quantitative RT-PCR (qRT-PCR).

Western blot analysis.

siRNA transfections.

Virion RNA and luciferase reporter gene assays.

Coimmunoprecipitation of RIG-I-RNA complexes and RNA extraction.

Statistical analysis.

RESULTS

CCHFV infection induces a type I IFN response.

Currently, the cellular immune-modulating receptors that recognize CCHFV and initiate the IFN-mediated antiviral response are unclear (14, 23, 24). To gain insights into the host antiviral pathways induced in response to CCHFV infection and to identify putative receptors, we examined type I IFN responses and the expression of early IFN-stimulated genes (ISG) in CCHFV-infected human A549 cells. A549 cells are efficient IFN producers, express a spectrum of cytosolic and surface receptors for virus recognition, including RIG-I and MDA5, and are commonly used to study the innate immune response to bunyavirus infections (24,–26). We used quantitative RT-PCR (qRT-PCR) and ELISA to analyze gene and protein expression by CCHFV-infected A549 cells. Higher IFN-β protein secretion was observed in infected cells than in mock-infected cells, with the highest levels observed at 48 h postinfection (Fig. 1A). Increased IFN-β gene expression (MOI, 1.0) was not observed until ≥12 hpi, supporting previous reports of delayed responses (24), and peaked at 48 hpi in infected cells (Fig. 1D).

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430001.jpg

FIG 1

Crimean-Congo hemorrhagic fever virus infection produces type I interferon response. (A) IFN-β is secreted by A549 cells in response to CCHFV infection (multiplicity of infection [MOI], 1). Human IFN-β expression was measured by ELISA in samples collected 24 and 48 h postinfection (hpi). WT indicates cells infected with wild-type CCHFV, while mock indicates mock-infected cells. (B) CCHFV infection (WT; MOI, 1) results in increased total STAT1 and activated (phosphorylated) pSTAT1. Cell lysates were collected from A549 and SW13 cells 24 hpi and analyzed by Western blotting. (C) CCHFV S segment levels increase over time after infection with non-UV-treated virus and correlate with the kinetics and magnitude of the type I IFN response. RNA was isolated from A549 cells infected at an MOI of 0.1 or 1 of CCHFV or UV-treated CCHFV at the indicated time points postinfection and analyzed by quantitative RT-PCR (qRT-PCR) using CCHFV S segment-specific primers. Viral copy numbers were determined based on a standard curve. (D) IFN-β, ISG56, and MxA levels increase in response to CCHFV infection. A549 cells were infected at a CCHFV MOI of 0.1 or 1.0, and RNA samples were collected at the indicated time points. mRNA levels of the indicated genes were assessed by qRT-PCR and normalized to β-actin levels. Data are represented as means ± standard deviations (SD) from a minimum of triplicate samples and are representative of replicate experiments.

The Jak/STAT pathway initiates a signal when IFN binds to the IFN-α/β receptor, resulting in phosphorylation of receptor-associated tyrosine kinases Jak1 and Tyk2, STAT-1 and STAT-2 recruitment and activation via tyrosine phosphorylation, and, finally, activation of IFN-α/β-regulated gene promoters. Thus, we further characterized the IFN response by investigating the effects of CCHFV infection on protein levels of total STAT and activated pSTAT and on expression of ISG56 and MxA. In addition to A549, we tested SW13 cells because they are highly susceptible to CCHFV infection. Increased STAT1 protein expression and phosphorylation was seen in both A549 and SW13 cells infected with CCHFV compared to that of mock-infected cells (Fig. 1B). mRNA levels of ISG56, an early immune-stimulated gene that inhibits translation in virus-infected cells (27, 28), was detected as early as 8 hpi in cells infected with both wild-type (WT) and replication-defective UV-treated CCHFV, and this early response increased with higher multiplicities of infection (MOI) of virus used (Fig. 1D). Notable differences in expression of MxA, an IFN-induced protein previously shown to inhibit CCHFV infection (23), were not observed until 24 hpi at both MOIs tested. Similar to IFN-β expression, a sustained increase in mRNA levels of both ISG56 and MxA was dependent on productive virus replication, as UV-inactivated CCHFV induced these genes poorly (Fig. 1D). CCHFV S-segment RNA levels were evaluated by qRT-PCR. In accordance with previous reports in other cell types (29), we observed that CCHFV infection with a higher MOI (MOI of 1.0 versus 0.1) resulted in increased vRNA synthesis at early time points after infection but in lower relative vRNA levels later in infection (Fig. 1C, compare top and bottom). Higher levels of CCHFV RNA were associated with an increased IFN response (Fig. 1D).

RIG-I and MDA5 expression is upregulated in CCHFV-infected cells.

Our data revealed a classical type I IFN response to CCHFV infection. Based on these observations, we began investigating putative cytosolic receptors by characterizing indicative changes to RLRs, as RLRs are essential for the immune recognition of and response to most RNA viruses (30). As RIG-I and MDA5 have never been investigated in the context of CCHFV infection, we began by analyzing gene and protein expression of RIG-I and MDA5 and found that both RLRs were upregulated in infected A549 cells (Fig. 2). Increased gene expression of RIG-I and MDA5 mRNA was sustained through 72 hpi when using CCHFV at an MOI of 0.1; at an MOI of 1.0, expression plateaued at 48 hpi (Fig. 2A). At 12 hpi, increased RLR protein expression was seen in cells infected with either WT or UV-treated CCHFV (MOI of 5.0). However, at 24 hpi, protein levels continued to increase only in cells infected with WT CCHFV (Fig. 2B). RIG-I protein levels in mock-infected cells were found to be relatively consistent over time; subjective changes in depicted baseline levels likely are due to a cross-well overflow artifact (Fig. 2B, 48 hpi, mock and WT).

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430002.jpg

FIG 2

RIG-I and MDA5 expression is upregulated in CCHFV-infected cells. (A) A549 cells were infected with CCHFV (MOI, 0.1 or 1.0), and samples were collected for RNA isolation and analysis at the indicated time points. RIG-I, MDA5, and MAVS gene expression levels were quantified by qRT-PCR, standardized to β-actin, and expressed as the fold change from the levels of mock-infected cells (±SD from triplicate samples and representative of replicate experiments). (B) A549 cells were infected with CCHFV or UV-treated CCHFV (MOI, 5.0). Cell extracts obtained 12, 24, or 48 hpi were analyzed on a denaturing gel followed by Western blotting with anti-CCHFV N hyperimmune mouse ascitic fluid (HMAF) or with anti-RIG-I, anti-MDA5, or anti-MAVS antibody.

RLRs bind viral RNA and initiate the antiviral immune response by interacting with the adaptor molecule MAVS, which contains a CARD, a proline-rich region, and a transmembrane domain. MAVS forms functional polymers on the mitochondria after viral infection (31,–33) and is a key component of most RIG-I-dependent signaling (34). Therefore, we investigated gene expression and protein levels of MAVS in CCHFV-infected cells. Expression of RIG-I and MDA5, RLRs that signal through MAVS, increased after CCHFV infection. RNA and protein levels of MAVS remained unchanged (Fig. 2A and andB).B). These data provide details of the magnitude of receptor and adaptor molecule expression specifically in the context of CCHFV infection to aid in moving forward with studies investigating the effects of disrupting gene and protein expression.

RIG-I is a key mediator in the interferon-stimulated gene response to CCHFV.

RIG-I and MDA5 recognize RNA viruses differently (35, 36). Some viruses are recognized by both RIG-I and MDA5 (1, 37, 38), but most are recognized predominantly by one or the other. RIG-I and MDA5 both induce type I IFN responses and are also ISGs; their gene expression is upregulated by viral infection and dsRNA (3, 39,–42). Therefore, increased expression of RIG-I and MDA5 in the context of CCHFV infection (Fig. 2) is consistent with induction of a type I IFN response (Fig. 1). To determine if RIG-I or MDA5 contributes to mediating the observed IFN response, we used siRNA to knock down endogenous expression of each receptor and the adaptor molecule MAVS, and we assessed the effects of these knockdowns on downstream ISG expression. Targeted siRNA reduced MDA5 RNA expression by ∼30% and that of RIG-I by over 70% in mock-infected cells. In CCHFV-infected cells, RIG-I, MDA5, and MAVS RNA expression levels were reduced by 99%, 75%, and 58%, respectively, compared to infected cells transfected with control siRNA. Decreased transcript levels from targeted siRNA resulted in reduced protein expression (RIG-I, 93% reduction; MDA5, 95%; and MAVS, 42%) after CCHFV infection compared to cells transfected with control siRNA (Fig. 3A). Decreased expression of RIG-I or MAVS, but not MDA5, significantly reduced both ISG56 and MxA mRNA expression (Fig. 3B) and IFN-β secretion (Fig. 3C) in CCHFV-infected A549 cells.

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430003.jpg

FIG 3

RIG-I is a key mediator of the type I IFN response to CCHFV. (A) RIG-I, MDA5, and MAVS knockdown by siRNA transfection in A549 cells with or without CCHFV or RVFV infection. A549 cells were reverse transfected with RIG-I, MDA5, MAVS, or nontargeted control (ct) siRNA and retransfected 24 h after initial transfection. After 24 h, cells were infected with CCHFV (MOI, 1.0); 24 h later, cells were harvested either for total cellular RNA isolation and qRT-PCR (top) or for Western blot (bottom) analysis. (B) Knockdown of RIG-I and its adaptor protein MAVS significantly (P < 0.05) reduced expression of ISG56 and MxA, while knocking down MDA5 levels did not affect subsequent responses, as measured by qRT-PCR. (C) Decreased RIG-I expression reduced secreted IFN-β in A549 cells. A549 cells were transfected with RIG-I-specific siRNA or nontargeted control (ct) siRNA. At 24 h posttransfection, cells were mock infected or infected with CCHFV (MOI, 1.0), and supernatants were analyzed by ELISA specific for IFN-β. Data are presented as picograms of IFN-β per milliliter of supernatant.

The RIG-I-mediated response controls CCHFV replication.

Previous reports suggest that RIG-I is not involved in recognition of CCHFV due to characteristics of the 5′ termini of the genome (14, 43). As our data support that RIG-I is involved in early immune signaling in CCHFV infection, mediating increased ISG expression and IFN-β production, we were specifically interested in further investigations of RIG-I recognition in CCHFV infection. To investigate the effects of RIG-I on virus replication, we determined CCHFV titers in cells treated with siRNA against RIG-I. When RIG-I expression was decreased by siRNA in A549 cells, mean viral titers increased by 3.95-fold and 3.75-fold compared to those of infected cells without knockdown at 2 and 3 dpi, respectively (Fig. 4A), indicating a direct effect of RIG-I-mediated immune responses on viral replication. To further investigate this effect, we used a series of mouse embryonic fibroblast (MEF) cell lines: WT, RIG-I knockout (RIG-I KO), and a RIG-I KO cell line that endogenously expresses human RIG-I (hRIG-I). As seen in A549 cells, increased viral titers were seen upon CCHFV infection in cells lacking RIG-I expression compared to that of WT cells (Fig. 4B). Interestingly, MEF expressing hRIG-I had viral titers higher than those of RIG-I KO cells but lower than those of WT cells. This result could be due to species differences in RIG-I or to lower efficiency of RIG-I expression in reconstituted MEFs.

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430004.jpg

FIG 4

RIG-I mediated response controls CCHFV replication. (A) RIG-I controls CCHFV replication in A549 cells. A549 cells were transfected with RIG-I specific siRNA or control (ct) siRNA. At 24 h posttransfection, cells were infected with CCHFV (MOI, 1.0); 1, 2, and 3 dpi, supernatants were collected for TCID₅₀ determination. Data are presented as means ± standard errors of the means (SEM) from two replicate experiments. (B) RIG-I controls CCHFV replication in MEF cells. WT, RIG-I knockout (RIG-I^{), and RIG-I knockout MEF cells reconstituted with human RIG-I (hRIG-I) were infected with CCHFV (MOI, 0.1). At the indicated time points postinfection, supernatants were collected for TCID}₅₀ determination. (C) Immunofluorescence-based quantification of the percentage of WT, RIG-I^{, or hRIG-I-reconstituted (recon) MEF lines infected at 24 and 48 hpi with the indicated doses of CCHFV. MEF cells were fixed with 10% formalin for 20 min and rinsed with PBS. All samples were γ-cell irradiated prior to subsequent staining and handling outside the biosafety level 4 laboratory. Data are represented as means ± SD from quadruplicate samples.}

In independent experiments, we used immunofluorescence to assess the effects of RIG-I expression levels on viral replication. Supporting our viral titration data in previous experiments, we found that CCHFV infected significantly more RIG-I KO cells than WT or hRIG-I reconstituted cells (Fig. 4C). At 48 hpi, all MOI of CCHFV infected over 45% of RIG-I KO cells (46.9% to 59.6%) versus WT (0.2% to 5.3%) or hRIG-I-reconstituted MEFs (1.8% to 12%).

CCHFV virion RNA is sufficient to stimulate a RIG-I-mediated IFN response.

In contrast to many negative-strand RNA viruses, CCHFV vRNA has a terminal 5′p instead of the 5′ppp group that is commonly involved in RIG-I recognition (14). IFN responses to incoming RNA virus nucleocapsids occur only if the genome contains a 5′-triphosphate (43). Thus, a 5′p RNA genome is thought to be a strategy to evade or delay host antiviral responses. Early investigation suggested that CCHFV RNA does not elicit an IFN response, which was attributed to the presence of 5′p RNA (14). As our data supported a direct role of RIG-I in the response to CCHFV, using a reporter cell line expressing firefly luciferase driven by an IFN-β promoter (293T-IFN-β-FF), we investigated the identity of the pathogen-associated molecular patterns (PAMPs) sensed by RIG-I in CCHFV infection.

We measured the IFN-dependent luciferase reporter activity 24 and 48 h after CCHFV infection (Fig. 5A). Increased luciferase reporter activity was observed upon infection, as shown in earlier experiments using IFN-β ELISA in A549 cells (Fig. 1A). Luciferase activity then was measured to investigate the nature of the PAMP synthesized during CCHFV replication. As CCHFV 5′p genomic RNA was not thought to be a RIG-I agonist, we initially aimed to investigate whether RNA generated in the cell versus RNA associated with virus particles could be involved in the observed immunostimulation. First, we determined the immunostimulatory effect of total RNA from CCHFV-infected cells in the 293T-IFN-β-FF reporter cell line compared to that of mock-treated cells or 293T-IFN-β-FF cells given total RNA from cells infected with Rift Valley fever virus (RVFV; a bunyavirus with a 5′ppp terminal group and a known RIG-I activator [14, 44]). Total RNA from CCHFV-infected cells induced an IFN-β and ISG56 response, while RNA from uninfected cells (mock) did not. As expected, RNA from RVFV-infected cells stimulated a very robust response (Fig. 5B). To determine if immunostimulatory RNA also was present in CCHFV particles isolated from supernatants of infected cells, we repeated the studies using vRNA. Prior to transfection, the identity and integrity of bunyavirus vRNAs were confirmed using denaturing formaldehyde agarose gel electrophoresis. Bands corresponding in size to CCHFV and RVFV segments confirmed vRNA enrichment and relative purity of these preparations (Fig. 5C). Transfection of CCHFV vRNA resulted in significantly increased IFN-β promoter activity over that of mock-transfected cells (Fig. 5D). This increase was abolished by RIG-I siRNA knockdown, demonstrating that RIG-I is required in the response to vRNA (Fig. 5E).

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430005.jpg

FIG 5

CCHFV 5′-monophosphate vRNA triggers IFN response. (A) 293T-IFN-β-FF reporter cells were infected with CCHFV (MOI, 0.1 or 1.0) and analyzed for reporter activity 1 and 2 dpi. RLU, relative light units. Equal amounts of purified total RNA isolated 24 hpi from the cell lysates of CCHFV- and RVFV-infected A549 cells (B) or virion RNA (vRNA) isolated 48 hpi from supernatants of CCHFV- and RVFV-infected cells (D) were transfected into 293T-IFN-β-FF reporter cells. Luciferase activity was analyzed 1 dpi. (C) Purified CCHFV and RVFV vRNAs were separated on a denaturing formaldehyde agarose gel. (E) 293T-IFN-β-FF reporter cells were transfected with RIG-I-specific siRNA or control (ct) siRNA. After 24 h, cells were transfected with 50 ng of vRNA, and reporter activity was assessed 24 h later. All quantitative data are presented as means ± SEM from 3 independent experiments. A single asterisk indicates a significant difference from mock-infected control cells (P < 0.05); 2 asterisks indicate a significant difference from infected cells transfected with control siRNA (P < 0.05).

Finally, using RVFV RNA as a positive control because of its robust IFN-β induction, we determined if immunostimulatory CCHFV RNA could be isolated by RIG-I coimmunoprecipitation. Total RNA from CCHFV- or RVFV-infected cells was incubated with recombinant RIG-I. RNA complexes were coimmunoprecipitated with anti-RIG-I antibody (RIG-I in Fig. 6A) or control antibody (IgG). The pulldown efficiency of RIG-I was confirmed by Western blotting of the immunoprecipitation samples (Fig. 6B). RNA coimmunoprecipitated in a RIG-I specific manner activated an IFN response more potently than RNA coimmunoprecipitated with IgG, showing that immunostimulatory CCHFV RNA synthesized during the course of CCHFV infection is selectively enriched by RIG-I coimmunoprecipitation (Fig. 6C).

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430006.jpg

FIG 6

CCHFV RNA coimmunoprecipitates with RIG-I. (A) Lysates from HEK-293T cells expressing HA-tagged WT RIG-I were incubated with 50 μg of the indicated RNA. RIG-I-RNA complexes were coimmunoprecipitated with anti-RIG-I antibody or IgG isotype control, and RNA and protein fractions were isolated. RNA was transfected into 293T-IFN-β-FF-Luc cells; 36 h later, IFN-β promoter-driven luciferase activity was measured by luciferase assays. A single asterisk indicates a significant difference (P < 0.05) from control. (B) RIG-I coimmunoprecipitation protein fractions were subjected to immunoblotting using an anti-RIG-I antibody to assess the pulldown efficiency. IP, immunoprecipitate; IB, immunoblot. (C) CCHFV and RVFV RNA levels in the RNA fraction of the coimmunoprecipitation determined by qRT-PCR. The ratios of viral RNA (virus) or control cellular mRNA (B2M) are represented as the abundance of each RNA target in RIG-I-IP over GFP-IP.

Investigation into biochemical nature of RIG-I inducers in CCHFV-infected cells supports contribution of 5′p RNA to immunostimulation.

Immunostimulation with CCHFV RNA induced a RIG-I-mediated type I IFN response. The genome of CCHFV has been characterized previously as 5′p (14). To confirm the biochemical nature of RIG-I inducers in CCHFV-infected cells and to investigate the potential contribution of genomes with 5′p processing, we transfected IFN-β-FF reporter cells with virion RNA, stripped of the nucleoprotein, digested with phosphatases and RNases (Fig. 7A). Alkaline phosphatase was used to remove phosphate groups from RNA and evaluate the potential contribution of CCHFV 5′p or 3′ phosphate in eliciting an IFN response. Removing all phosphate groups abolished the IFN-β promoter activity induced by both CCHF and RVF virion RNA preparation. To confirm that CCHFV immunostimulation was due to 5′p and not 5′pp or 5′ppp RNA possibly present in CCHF virions, we digested virion RNA with 5′-polyphosphatase, an enzyme that sequentially removes the γ- and β-phosphates but does not affect the α-phosphate group. Therefore, adding 5′-polyphosphatase will convert a 5′ppp RNA genome to 5′p, while 5′p RNA will remain unchanged. 5′-Polyphosphatase digestion did not significantly reduce immunostimulation by CCHFV RNA, confirming that the CCHFV genome contains the 5′p moiety and further suggesting that CCHFV IFN induction is independent of 5′ppp or 5′pp moieties. Meanwhile, 5′-polyphosphatase significantly reduced the relative IFN stimulation by RVFV RNA, confirming that the digestion worked. However, when RVFV RNA was digested to 5′p groups, IFN induction was lower than induction by CCHFV RNA (Fig. 7B). Since canonical RIG-I ligands also are dsRNA, we investigated whether immunostimulatory properties of CCHFV RNA were dependent on dsRNA. Virion RNA digestion with dsRNA-specific RNase V1 abolished the luciferase induction seen with RVFV RNA, while 11% of CCHFV activity was retained (Fig. 7A).

An external file that holds a picture, illustration, etc.
Object name is zjv9990908430007.jpg

FIG 7

Characterizing the immunostimulatory RNA of CCHFV. (A) Effects of enzymatic digestion of vRNA on IFN-promoter activity. Fifty nanograms of vRNA digested with APex heat-labile alkaline phosphatase, 5′-polyphophatase, or RNase V1 was transfected into 293T-IFN-FF reporter cells. (+), added enzyme; (−), no enzyme added or enzyme was heat inactivated in the case of APex-treated RNA. To reduce interexperiment absolute value variation, IFN promoter activity was calculated as a percentage of luciferase activity in cells given control RNA incubated with heat-inactivated enzyme (APex) or the respective enzyme buffers alone. Data are presented as standardized means ± SEM from 4 independent experiments done in triplicate or quadruplicate. A single asterisk indicates a significant difference from virus-specific untreated RNA (P < 0.05). (B) RLU values in cells transfected with RNA digested with 5′-polyphosphatase from experiments standardized as shown in panel A.

Investigation into biochemical nature of RIG-I inducers in CCHFV-infected cells supports contribution of 5′p RNA to immunostimulation.

FIG 7

DISCUSSION

Viral infection often is dependent on effective abrogation of the host immune response. As a result, many viruses encode proteins that actively modulate the immune response. Alternatively, viruses can evade the antiviral response by avoiding detection by immune receptors that mediate immune responses, abrogating the initiation of immune signaling. Viral production of 5′p RNA, which mimics cellular ribosomal and tRNA, was proposed as a strategy by which viruses can avoid recognition by RIG-I, an RLR involved in the detection of viruses producing 5′ppp or 5′pp RNA. Members of the family Bunyaviridae (nairoviruses and hantaviruses) and Bornaviridae employ this strategy by processing the 5′ termini of their genome RNA to generate 5′-monophosphorylated RNA (14). Here, we investigated CCHFV, a nairovirus with a 5′p RNA genome, and found that, despite previous assumptions about 5′p recognition, RIG-I and MAVS mediated the upregulation of type-I IFN and ISGs in response to virus infection (Fig. 3).

A PRR for nairoviruses has not been identified to date. Our data support the role of RIG-I in CCHFV antiviral signaling (Fig. 3 to to6).6). While our data strongly support RIG-I-mediated antiviral responses against CCHFV, without further investigation we cannot definitively say whether this phenomenon is genus wide. However, if the ligand is viral RNA, then we would expect the RIG-I response to be comparable. Interestingly, a recent review alluded to IFN immunostimulation by Nairobi sheep disease vRNA, contrasting with the previously published data on CCHFV vRNA (45), suggesting that this observation can be seen in other nairoviruses. Hantaviruses, also Bunyaviridae family members, have a 5′p genome and poorly induce the early innate immune response, yet knocking down RIG-I expression increased viral titers of Hantaan virus (46). While limited to RIG-I knockdown experiments and not PAMP identification, these data, along with our studies, suggest the need to verify RIG-I signaling during infection with viruses previously thought to completely elude the RIG-I-mediated responses by 5′p genome processing.

The mechanism of 5′ terminus processing for nairoviruses has not been described yet and may help explain our findings. Genome processing of hantaviruses has been proposed to involve an endonuclease (potentially the same endonuclease as that involved in cap snatching) that removes the 5′ terminal extension of the genome, leaving the 5′ pU at position +1 (47). Although our data support a putative role of 5′p genomes in immunostimulation (Fig. 7), it does not exclude the possibility of other contributing ligands in CCHFV infection, as production of 5′p termini by CCHFV may involve the generation of 5′ppp intermediates or truncated defective interfering (DI) particles shown to be potent immunostimulatory ligands in the context of other viral infections (48). These 5′ppp by-products, as potent RIG-I ligands, may stimulate RIG-I as we saw in our studies, while the transient nature or scarcity of these products would keep the RIG-I response low compared to that of 5′ppp viruses. Alternatively, complete processing of termini without the presence of immunostimulatory intermediates also may occur, suggesting that while processing the 5′ terminus to a monophosphate likely dampens cellular immune responses to CCHFV, this processing may not be sufficient to fully abrogate signaling through RIG-I, as previously suggested (14).

Incomplete evasion of RIG-I by CCHFV supports the need for a virus-encoded antagonist(s) to actively represses RIG-I-dependent IFN production, as previously suggested by the delayed nuclear translocation of IRF3 observed after infection with live versus UV-inactivated CCHFV (30). The L-protein cysteine protease of some nairoviruses, including CCHFV, contains the OTU domain, which possesses dual deubiquitinase and deISGylase activities (11). The CCHFV OTU would be a logical candidate for suppressing the immune response, as catalytically active OTUs of various origins can block signaling by RIG-I and MAVS, cellular factors involved in the antiviral response to CCHFV infection (Fig. 3). The nairovirus OTU-containing deubiquitinases have been shown to target activated RIG-I to control innate immune signaling (12). However, in these studies the OTU was overexpressed from plasmids and a constitutively active form of mutant RIG-I was used as a signaling agonist. Apparently, under these conditions the OTU nonselectively removes ubiquitin chains from any cellular substrate. Given the importance of K63 polyubiquitin in RIG-I signaling (49), it is not surprising that CCHFV OTU overexpression in this way can block RIG-I signaling. However, it remains to be seen if CCHFV infection can induce a general reduction of protein ubiquitination levels. Future studies should investigate whether CCHFV OTU blocks RIG-I signaling by acting on specific targets of the RIG-I pathway.

Our report identifies RIG-I as a PRR for CCHFV. RIG-I recognizes CCHFV RNA, resulting in downstream type I IFN antiviral signaling and subsequent ISG stimulation. Identification of a PRR for CCHFV infection will aid in determining key antiviral signaling pathways in the control of virus replication. In addition, our studies highlight the need for further investigation into genomic processing of 5′p producing viruses and the potential for 5′p genomes to serve as RIG-I ligands.

^{Viral Special Pathogens Branch, Division of High Consequence Pathogens and Pathology, Centers for Disease Control and Prevention, Atlanta, Georgia, USA}

^{Department of Microbiology, Icahn School of Medicine at Mount Sinai, New York, New York, USA}

^{Global Health and Emerging Pathogens Institute, Icahn School of Medicine at Mount Sinai, New York, New York, USA}

^{Department of Medicine, Division of Infectious Diseases, Icahn School of Medicine at Mount Sinai, New York, New York, USA}

^{Corresponding author.}

Address correspondence to Éric Bergeron, vog.cdc@noregrebe.

Citation Spengler JR, Patel JR, Chakrabarti AK, Zivcec M, García-Sastre A, Spiropoulou CF, Bergeron É. 2015. RIG-I mediates an antiviral response to Crimean-Congo hemorrhagic fever virus. J Virol 89:10219–10229. doi:10.1128/JVI.01643-15.

Received 2015 Jun 30; Accepted 2015 Jul 21.

ABSTRACT

In the cytoplasm, the retinoic acid-inducible gene I (RIG-I) senses the RNA genomes of several RNA viruses. RIG-I binds to viral RNA, eliciting an antiviral response via the cellular adaptor MAVS. Crimean-Congo hemorrhagic fever virus (CCHFV), a negative-sense RNA virus with a 5′-monophosphorylated genome, is a highly pathogenic zoonotic agent with significant public health implications. We found that, during CCHFV infection, RIG-I mediated a type I interferon (IFN) response via MAVS. Interfering with RIG-I signaling reduced IFN production and IFN-stimulated gene expression and increased viral replication. Immunostimulatory RNA was isolated from CCHFV-infected cells and from virion preparations, and RIG-I coimmunoprecipitation of infected cell lysates isolated immunostimulatory CCHFV RNA. This report serves as the first description of a pattern recognition receptor for CCHFV and highlights a critical signaling pathway in the antiviral response to CCHFV.

IMPORTANCE CCHFV is a tick-borne virus with a significant public health impact. In order for cells to respond to virus infection, they must recognize the virus as foreign and initiate antiviral signaling. To date, the receptors involved in immune recognition of CCHFV are not known. Here, we investigate and identify RIG-I as a receptor involved in initiating an antiviral response to CCHFV. This receptor initially was not expected to play a role in CCHFV recognition because of characteristics of the viral genome. These findings are important in understanding the antiviral response to CCHFV and support continued investigation into the spectrum of potential viruses recognized by RIG-I.

ABSTRACT

ACKNOWLEDGMENTS

We thank Stuart Nichol, Laura McMullan, Tatyana Klimova, and Friedemann Weber for their critical reviews of the manuscript and César Albariño for helpful scientific discussions. We thank Maria T. Sanchez-Aparacio for helpful discussions and Richard Cadagan and Osman Lizardo for excellent technical assistance at the Icahn School of Medicine at Mount Sinai.

These studies were supported by the CDC and a CDC foundation project funded by NIAID grant R01AI109008. This research was supported in part by an appointment to the ASM/CDC Postdoctoral Research Fellowship Program and to the Research Participation Program at the Centers for Disease Control and Prevention, administered by the Oak Ridge Institute for Science and Education through an interagency agreement between the U.S. Department of Energy and the CDC (to J.R.S.), and by a National Institutes of Health Loan Repayment Award (to J.R.S.).

The findings and conclusions in this report are those of the authors and do not necessarily represent the official position of the Centers for Disease Control and Prevention.

ACKNOWLEDGMENTS

REFERENCES

References

1. Jensen S, Thomsen AR. 2012 Sensing of RNA viruses: a review of innate immune receptors involved in recognizing RNA virus invasion. J Virol86:2900–2910. doi:10.1128/JVI.05738-11. ] [[Google Scholar]
2. Ng CS, Kato H, Fujita T. 2012 Recognition of viruses in the cytoplasm by RLRs and other helicases–how conformational changes, mitochondrial dynamics and ubiquitination control innate immune responses. Int Immunol24:739–749. doi:10.1093/intimm/dxs099. [] [[PubMed][Google Scholar]
3. Yoneyama M, Kikuchi M, Natsukawa T, Shinobu N, Imaizumi T, Miyagishi M, Taira K, Akira S, Fujita T. 2004 The RNA helicase RIG-I has an essential function in double-stranded RNA-induced innate antiviral responses. Nat Immunol5:730–737. doi:10.1038/ni1087. [] [[PubMed][Google Scholar]
4. Saito T, Hirai R, Loo Y-M, Owen D, Johnson CL, Sinha SC, Akira S, Fujita T, Gale M. 2007 Regulation of innate antiviral defenses through a shared repressor domain in RIG-I and LGP2. Proc Natl Acad Sci U S A104:582–587. doi:10.1073/pnas.0606699104. ] [[Google Scholar]
5. Patel JR, Jain A, Chou Y-Y, Baum A, Ha T, García-Sastre A. 2013 ATPase-driven oligomerization of RIG-I on RNA allows optimal activation of type-I interferon. EMBO Rep14:780–787. doi:10.1038/embor.2013.102. ] [[Google Scholar]
6. Peisley A, Lin C, Wu B, Orme-Johnson M, Liu M, Walz T, Hur S. 2011 Cooperative assembly and dynamic disassembly of MDA5 filaments for viral dsRNA recognition. Proc Natl Acad Sci U S A108:21010–21015. doi:10.1073/pnas.1113651108. ] [[Google Scholar]
7. Peisley A, Wu B, Yao H, Walz T, Hur S. 2013 RIG-I forms signaling-competent filaments in an ATP-dependent, ubiquitin-independent manner. Mol Cell51:573–583. doi:10.1016/j.molcel.2013.07.024. [] [[PubMed][Google Scholar]
8. Leung DW, Amarasinghe GK. 2012 Structural insights into RNA recognition and activation of RIG-I-like receptors. Curr Opin Struct Biol22:297–303. doi:10.1016/j.sbi.2012.03.011. ] [[Google Scholar]
9. Kowalinski E, Lunardi T, McCarthy AA, Louber J, Brunel J, Grigorov B, Gerlier D, Cusack S. 2011 Structural basis for the activation of innate immune pattern-recognition receptor RIG-I by viral RNA. Cell147:423–435. doi:10.1016/j.cell.2011.09.039. [] [[PubMed][Google Scholar]
10. Rehwinkel J, Reis e Sousa C. 2010 RIGorous detection: exposing virus through RNA sensing. Science327:284–286. doi:10.1126/science.1185068. [] [[PubMed][Google Scholar]
11. Frias-Staheli N, Giannakopoulos NV, Kikkert M, Shannon L, Bridgen A, Paragas J, Richt JA, Rowland RR, Schmaljohn CS, Lenschow DJ, Snijder EJ, García-Sastre A, Taylor SL, Virgin HW. 2007 Ovarian tumor domain-containing viral proteases evade ubiquitin- and ISG15-dependent innate immune responses. Cell Host Microbe2:404–416. doi:10.1016/j.chom.2007.09.014. ] [[Google Scholar]
12. Van Kasteren PB, Beugeling C, Ninaber DK, Frias-Staheli N, van Boheemen S, García-Sastre A, Snijder EJ, Kikkert M. 2012 Arterivirus and nairovirus ovarian tumor domain-containing deubiquitinases target activated RIG-I to control innate immune signaling. J Virol86:773–785. doi:10.1128/JVI.06277-11. ] [[Google Scholar]
13. Whitehouse CA. 2004 Crimean-Congo hemorrhagic fever. Antiviral Res64:145–160. doi:10.1016/j.antiviral.2004.08.001. [] [[PubMed][Google Scholar]
14. Habjan M, Andersson I, Klingström J, Schümann M, Martin A, Zimmermann P, Wagner V, Pichlmair A, Schneider U, Mühlberger E, Mirazimi A, Weber F. 2008 Processing of genome 5′ termini as a strategy of negative-strand RNA viruses to avoid RIG-I-dependent interferon induction. PLoS One3:e2032. doi:10.1371/journal.pone.0002032. ] [[Google Scholar]
15. Bird BH, Albariño CG, Nichol ST, Hartman AL, Erickson BR, Ksiazek TG. 2008 Rift valley fever virus lacking the NSs and NSm genes is highly attenuated, confers protective immunity from virulent virus challenge, and allows for differential identification of infected and vaccinated animals. J Virol82:2681–2691. doi:10.1128/JVI.02501-07. ] [[Google Scholar]
16. Bergeron É, Zivcec M, Chakrabarti AK, Nichol ST, Albariño CG, Spiropoulou CF. 2015 Recovery of recombinant Crimean Congo hemorrhagic fever virus reveals a function for non-structural glycoproteins cleavage by furin. PLoS Pathog11:e1004879. doi:10.1371/journal.ppat.1004879. ] [[Google Scholar]
17. Nasirudeen AMA, Wong HH, Thien P, Xu S, Lam K, Xiang D, Liu DX. 2011 RIG-I, MDA5 and TLR3 synergistically play an important role in restriction of dengue virus infection. PLoS Negl Trop Dis5:e926. doi:10.1371/journal.pntd.0000926. ] [[Google Scholar]
18. Prescott J, Ye C, Sen G, Hjelle B. 2005 Induction of innate immune response genes by Sin Nombre hantavirus does not require viral replication. J Virol79:15007–15015. doi:10.1128/JVI.79.24.15007-15015.2005. ] [[Google Scholar]
19. Steensberg A, Keller C, Starkie RL, Osada T, Febbraio MA, Pedersen BK. 2002 IL-6 and TNF-alpha expression in, and release from, contracting human skeletal muscle. Am J Physiol Endocrinol Metab283:E1272–E1278. doi:10.1152/ajpendo.00255.2002. [] [[PubMed][Google Scholar]
20. Okamura S, Fujiwara H, Yoneda M, Furutani A, Todo M, Ikai A, Tada H, Okamura H, Umehara S, Shiozaki A, Kubota T, Komatsu S, Ichikawa D, Okamoto K, Ochiai T, Otsuji E. 2013 Overexpression of IL-6 by gene transfer stimulates IL-8-mediated invasiveness of KYSE170 esophageal carcinoma cells. Anticancer Res33:1483–1489. [[PubMed][Google Scholar]
21. Prescott JB, Hall PR, Bondu-Hawkins VS, Ye C, Hjelle B. 2007 Early innate immune responses to Sin Nombre hantavirus occur independently of IFN regulatory factor 3, and viral entry. J Immunol179:1796–1802. doi:10.4049/jimmunol.179.3.1796. [] [[PubMed][Google Scholar]
22. Schulte M, Sorkin M, Al-Benna S, Stupka J, Hirsch T, Daigeler A, Kesting M, Steinau H-U, Jacobsen F, Steinstraesser L. 2013 Innate immune response after adenoviral gene delivery into skin is mediated by AIM2, NALP3, DAI and mda5. Springerplus2:234. doi:10.1186/2193-1801-2-234. ] [[Google Scholar]
23. Andersson I, Bladh L, Mousavi-Jazi M, Magnusson K, Lundkvist Å, Haller O, Mirazimi A, Preparedness M, Control D, Solna S, Irol JV. 2004 Human MxA protein inhibits the replication of Crimean-Congo hemorrhagic fever virus. J Virol78:4323–4329. doi:10.1128/JVI.78.8.4323-4329.2004. ] [[Google Scholar]
24. Andersson I, Karlberg H, Mousavi-Jazi M, Martı L, Weber F, Mirazimi A. 2008 Crimean-Congo hemorrhagic fever virus delays activation of the innate immune response. J Med Virol1404:1397–1404. [[PubMed][Google Scholar]
25. Handke W, Oelschlegel R, Franke R, Krüger DH, Rang A. 2009 Hantaan virus triggers TLR3-dependent innate immune responses. J Immunol182:2849–2858. doi:10.4049/jimmunol.0802893. [] [[PubMed][Google Scholar]
26. Levine JR, Prescott J, Brown KS, Best SM, Ebihara H, Feldmann H, Best M. 2010 Antagonism of type I interferon responses by new world hantaviruses. J Virol84:11790–11801. doi:10.1128/JVI.00916-10. ] [[Google Scholar]
27. Guo J, Peters KL, Sen GC. 2000 Induction of the human protein P56 by interferon, double-stranded RNA, or virus infection. Virology267:209–219. doi:10.1006/viro.1999.0135. [] [[PubMed][Google Scholar]
28. Hui DJ, Bhasker CR, Merrick WC, Sen GC. 2003 Viral stress-inducible protein p56 inhibits translation by blocking the interaction of eIF3 with the ternary complex eIF2.GTP.Met-tRNAi. J Biol Chem278:39477–39482. doi:10.1074/jbc.M305038200. [] [[PubMed][Google Scholar]
29. Rodrigues R, Paranhos-Baccalà G, Vernet G, Peyrefitte CN. 2012 Crimean-Congo hemorrhagic fever virus-infected hepatocytes induce ER-stress and apoptosis crosstalk. PLoS One7:e29712. doi:10.1371/journal.pone.0029712. ] [[Google Scholar]
30. Schlee M. 2013 Master sensors of pathogenic RNA–RIG-I like receptors. Immunobiology218:1322–1335. doi:10.1016/j.imbio.2013.06.007. [] [[PubMed][Google Scholar]
31. Papatriantafyllou M. 2011 Innate immunity: MAVS build-ups for defence. Nat Rev Immunol11:570. doi:10.1038/nri3050. [] [[PubMed][Google Scholar]
32. Hou F, Sun L, Zheng H, Skaug B, Jiang Q-X, Chen ZJ. 2011 MAVS forms functional prion-like aggregates to activate and propagate antiviral innate immune response. Cell146:448–461. doi:10.1016/j.cell.2011.06.041. ] [[Google Scholar]
33. Tang ED, Wang C-Y. 2009 MAVS self-association mediates antiviral innate immune signaling. J Virol83:3420–3428. doi:10.1128/JVI.02623-08. ] [[Google Scholar]
34. Dixit E, Kagan JC. 2013 Intracellular pathogen detection by RIG-I-like receptors. Adv Immunol117:99–125. doi:10.1016/B978-0-12-410524-9.00004-9. ] [[Google Scholar]
35. Gitlin L, Barchet W, Gilfillan S, Cella M, Beutler B, Flavell RA, Diamond MS, Colonna M. 2006 Essential role of mda-5 in type I IFN responses to polyriboinosinic:polyribocytidylic acid and encephalomyocarditis picornavirus. Proc Natl Acad Sci U S A103:8459–8464. doi:10.1073/pnas.0603082103. ] [[Google Scholar]
36. Kato H, Takeuchi O, Sato S, Yoneyama M, Yamamoto M, Matsui K, Uematsu S, Jung A, Kawai T, Ishii KJ, Yamaguchi O, Otsu K, Tsujimura T, Koh C-S, Reis e Sousa C, Matsuura Y, Fujita T, Akira S. 2006 Differential roles of MDA5 and RIG-I helicases in the recognition of RNA viruses. Nature441:101–105. doi:10.1038/nature04734. [] [[PubMed][Google Scholar]
37. Li J, Liu Y, Zhang X. 2010 Murine coronavirus induces type I interferon in oligodendrocytes through recognition by RIG-I and MDA5. J Virol84:6472–6482. doi:10.1128/JVI.00016-10. ] [[Google Scholar]
38. Chauveau E, Doceul V, Lara E, Adam M, Breard E, Sailleau C, Viarouge C, Desprat A, Meyer G, Schwartz-Cornil I, Ruscanu S, Charley B, Zientara S, Vitour D. 2012 Sensing and control of bluetongue virus infection in epithelial cells via RIG-I and MDA5 helicases. J Virol86:11789–11799. doi:10.1128/JVI.00430-12. ] [[Google Scholar]
39. Kim M-J, Hwang S-Y, Imaizumi T, Yoo J-Y. 2008 Negative feedback regulation of RIG-I-mediated antiviral signaling by interferon-induced ISG15 conjugation. J Virol82:1474–1483. doi:10.1128/JVI.01650-07. ] [[Google Scholar]
40. Su Z-Z, Sarkar D, Emdad L, Barral PM, Fisher PB. 2007 Central role of interferon regulatory factor-1 (IRF-1) in controlling retinoic acid expression. J Cell Physiol1:502–510. [[PubMed][Google Scholar]
41. Kang D, Gopalkrishnan RV, Wu Q, Jankowsky E, Pyle AM, Fisher PB. 2002 mda-5: an interferon-inducible putative RNA helicase with double-stranded RNA-dependent ATPase activity and melanoma growth-suppressive properties. Proc Natl Acad Sci U S A99:637–642. doi:10.1073/pnas.022637199. ] [[Google Scholar]
42. Kang D-C, Gopalkrishnan RV, Lin L, Randolph A, Valerie K, Pestka S, Fisher PB. 2004 Expression analysis and genomic characterization of human melanoma differentiation associated gene-5, mda-5: a novel type I interferon-responsive apoptosis-inducing gene. Oncogene23:1789–1800. doi:10.1038/sj.onc.1207300. [] [[PubMed][Google Scholar]
43. Weber M, Gawanbacht A, Habjan M, Rang A, Borner C, Schmidt AMM, Veitinger S, Jacob R, Devignot S, Kochs G, García-Sastre A, Weber F. 2013 Incoming RNA virus nucleocapsids containing a 5′-triphosphorylated genome activate RIG-I and antiviral signaling. Cell Host Microbe13:336–346. doi:10.1016/j.chom.2013.01.012. ] [[Google Scholar]
44. Weber F, Mirazimi A. 2008 Interferon and cytokine responses to Crimean Congo hemorrhagic fever virus; an emerging and neglected viral zoonosis. Cytokine Growth Factor Rev19:395–404. doi:10.1016/j.cytogfr.2008.11.001. [] [[PubMed][Google Scholar]
45. Lasecka L, Baron MD. 2014 The molecular biology of nairoviruses, an emerging group of tick-borne arboviruses. Arch Virol159:1249–1265. doi:10.1007/s00705-013-1940-z. [] [[PubMed][Google Scholar]
46. Lee M-H, Lalwani P, Raftery MJ, Matthaei M, Lütteke N, Kirsanovs S, Binder M, Ulrich RG, Giese T, Wolff T, Krüger DH, Schönrich G. 2011 RNA helicase retinoic acid-inducible gene I as a sensor of Hantaan virus replication. J Gen Virol92:2191–2200. doi:10.1099/vir.0.032367-0. [] [[PubMed][Google Scholar]
47. Garcin D, Lezzi M, Dobbs M, Elliott RM, Schmaljohn C, Kang CY, Kolakofsky D. 1995 The 5′ ends of Hantaan virus (Bunyaviridae) RNAs suggest a prime-and-realign mechanism for the initiation of RNA synthesis. J Virol69:5754–5762. [Google Scholar]
48. Baum A, Sachidanandam R, García-Sastre A. 2010 Preference of RIG-I for short viral RNA molecules in infected cells revealed by next-generation sequencing. Proc Natl Acad Sci U S A107:16303–16308. doi:10.1073/pnas.1005077107. ] [[Google Scholar]
49. Zeng W, Xu M, Liu S, Sun L, Chen ZJ. 2009 Key role of Ubc5 and lysine-63 polyubiquitination in viral activation of IRF3. Mol Cell36:315–325. doi:10.1016/j.molcel.2009.09.037. ] [[Google Scholar]