Research ArticleImmunology

IRE1α promotes viral infection by conferring resistance to apoptosis

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Sci. Signal.  06 Jun 2017:
Vol. 10, Issue 482, eaai7814
DOI: 10.1126/scisignal.aai7814

Resisting death by virus

The unfolded protein response (UPR) alleviates the cellular stress caused by the accumulation of proteins, such as that occurring during viral infection. During the UPR, processing of Xbp1 mRNA by the nuclease IRE1α generates the transcription factor XBP1, which drives the expression of genes encoding type I interferons (IFNs). Fink et al. found that Xbp1-deficient cells had defective antiviral responses to infection by hepatitis C virus (HCV); however, the cells were not compromised in type I IFN production or responses. Instead, these cells showed increased activation of IRE1α, which cleaved the proapoptotic microRNA miR-125a, leading to decreased apoptosis and increased viral replication. Liver biopsies from HCV-infected patients also showed increased IRE1α activation and decreased miR-125a abundance. Together, these data suggest that IRE1α functions to enhance cell survival in response to viral infection and may provide a potential therapeutic target.

Abstract

The unfolded protein response (UPR) is an ancient cellular pathway that detects and alleviates protein-folding stresses. The UPR components X-box binding protein 1 (XBP1) and inositol-requiring enzyme 1α (IRE1α) promote type I interferon (IFN) responses. We found that Xbp1-deficient mouse embryonic fibroblasts and macrophages had impaired antiviral resistance. However, this was not because of a defect in type I IFN responses but rather an inability of Xbp1-deficient cells to undergo viral-induced apoptosis. The ability to undergo apoptosis limited infection in wild-type cells. Xbp1-deficient cells were generally resistant to the intrinsic pathway of apoptosis through an indirect mechanism involving activation of the nuclease IRE1α. We observed an IRE1α-dependent reduction in the abundance of the proapoptotic microRNA miR-125a and a corresponding increase in the amounts of the members of the antiapoptotic Bcl-2 family. The activation of IRE1α by the hepatitis C virus (HCV) protein NS4B in XBP1-proficient cells also conferred apoptosis resistance and promoted viral replication. Furthermore, we found evidence of IRE1α activation and decreased miR-125a abundance in liver biopsies from patients infected with HCV compared to those in the livers of healthy controls. Our results reveal a prosurvival role for IRE1α in virally infected cells and suggest a possible target for IFN-independent antiviral therapy.

INTRODUCTION

Great advances have been made in our understanding of the molecular definitions of pattern recognition receptors (PRRs) and the pathogen-associated molecular patterns (PAMPs) that cells use to distinguish viruses from self (1, 2). PRR engagement results in the transcription of the genes that encode the type I interferons (IFNs) IFN-α and IFN-β, which bind to the IFN-α/β receptor to induce the expression of hundreds of IFN-stimulated genes (ISGs). ISGs act in concert to block further viral replication and spread, as well as to support the activation of adaptive antiviral immunity (3). However, many viruses have evolved evasion mechanisms to limit PRR recognition and signal transduction, and PRR-independent mechanisms for innate sensing of viral infections remain unclear.

Endoplasmic reticulum (ER) stress occurs during infection by various viruses, presumably due to the overwhelming synthesis of viral proteins (4). The unfolded protein response (UPR) is a ubiquitous cellular pathway to detect and alleviate ER stress. The UPR is initiated by three sensors that reside within the ER: protein kinase receptor–like ER kinase (PERK), activating transcription factor 6 (ATF6), and inositol-requiring enzyme 1 (IRE1) (57). IRE1, a highly conserved UPR sensor, oligomerizes and autophosphorylates in response to ER stress, which activates its cytosolic ribonuclease (RNase) domain and initiates a nonconventional mRNA splicing reaction of Xbp1 mRNA (8). Once processed, the spliced Xbp1 mRNA encodes a transcription factor, which controls the expression of target genes. IRE1α targets other specific mRNAs, leading to their degradation in a process termed regulated IRE1-dependent decay (RIDD) (9, 10). ER stress synergistically enhances cytokine and IFN responses to PRR engagement through IRE1α and X-box binding protein 1 (XBP1) (1113). Specific activation of IRE1α also occurs during innate immune recognition of PAMPs by Toll-like receptors (TLRs) (11). In this setting, XBP1 promotes the production of inflammatory cytokines and IFN-β. Moreover, IRE1α generates ligands for RIG-I–like receptors (RLRs) during the UPR (14), which are degraded by SKIV2L RNA exosomes to prevent inappropriate activation of type I IFN responses (15). These observations prompted us to investigate the possible role of XBP1 in innate immune responses to viral infections, with the hypothesis that XBP1 could promote IFN-mediated viral resistance.

Here, we describe an unexpected role for XBP1 in antiviral resistance, not through enhancement of the IFN response but rather by modulating susceptibility to host cell apoptosis. Xbp1-deficient cells were resistant to apoptosis during infection with vesicular stomatitis virus (VSV) and herpes simplex virus (HSV), and failure to undergo cell death resulted in increased viral replication. Xbp1 deficiency results in activation of its upstream enzyme IRE1α, which degrades specific cytosolic RNA targets (16, 17). We found that apoptosis resistance in the Xbp1-deficient cells required IRE1α and its degradation of microRNA (miRNA), miR-125a. Notably, the hepatitis C virus (HCV) nonstructural protein 4B (NS4B), which stimulates IRE1α activation (18), promoted the survival of infected cells and viral replication. Moreover, liver biopsies from patients infected with HCV showed IRE1α activation and reduced miR-125a abundance compared to healthy controls. These findings highlight the role of UPR effectors in regulating IFN-independent mechanisms of innate antiviral resistance through the induction of apoptosis to limit viral infection.

RESULTS

Xbp1 deficiency impairs control of viral infection

To determine the effect of Xbp1 deficiency on host defense against viral replication, we infected Xbp1−/− mouse embryonic fibroblasts (MEFs) with an RNA virus, VSV, and a DNA virus, HSV. XBP1 deficiency was achieved through an insertion of the neomycin resistance gene into parts of exons 1 and 2, as well as the intervening intron (19). This insertion still enables Xbp1 mRNA splicing but results in a frameshift of the remaining amino acids to prevent protein production. Compared to wild-type (WT) MEFs, VSV replication was enhanced in Xbp1−/− MEFs as determined by measuring VSV-G–green fluorescent protein (GFP) relative abundance by flow cytometry (Fig. 1, A and B) and by plaque assays of released virus from MEF culture medium (Fig. 1C). Similarly, Xbp1−/− MEFs also supported increased replication of HSV-1–GFP as indicated by measurement of GFP abundance (Fig. 1, D and E) and viral titer in the supernatant (Fig. 1F).

Fig. 1 Xbp1 deficiency enhances the susceptibility of MEFs to HSV and VSV.

(A to F) WT and Xbp1−/− MEFs were infected with VSV-GFP at a multiplicity of infection (MOI) of 1 [(A) to (C)] or with HSV-1–GFP at an MOI of 10 [(D) to (F)]. Twenty-four hours later, the extent of infection was determined by measuring the relative abundance of GFP by flow cytometry. Data are from one experiment representative of three independent experiments (A and D). The mean fluorescence intensity (MFI) of GFP in the indicated cells was then determined. Data are means ± SD of three independent experiments (B and E). Viral titers in the cell culture medium were measured by plaque assay at 48 (C) and 72 (F) hours after infection. PFU, plaque-forming units. Data are means ± SD of three independent experiments (C and F). *P < 0.05; ***P < 0.001 compared to WT, unpaired t test.

To determine whether the impaired viral control in Xbp1−/− MEFs resulted from deficient IFN responses, we measured the expression of genes encoding type I IFNs and of an ISG, Mx1, in MEFs infected with VSV. Unexpectedly, we observed enhanced induction of Ifna4 and Ifnb1 in Xbp1−/− MEFs infected with VSV compared to WT MEFs (fig. S1, A and B). Induction of Mx1 also increased in Xbp1−/− MEFs (fig. S1C). Moreover, Xbp1−/− MEFs were not impaired in IFN responsiveness because pretreatment with IFN-β prevented VSV replication in Xbp1−/− MEFs (fig. S1D). In contrast and consistent with previous reports (12, 13), the Xbp1-deficient MEF response to transfection with polyinosinic:polycytidylic acid [poly(I:C)] [an MDA5 (melanoma differentiation–associated protein 5) agonist] was impaired (fig. S1E), suggesting that the enhanced IFN response to VSV was specific to replicating virus. Together, these findings suggest that Xbp1 contributes to protective antiviral responses independently of type I IFNs.

Xbp1 deficiency confers resistance to virus-triggered cell death

Viral infection often culminates in the death of infected host cells. To determine whether the phenotype we observed in the Xbp1-deficient MEFs was due to a difference in the death of infected cells, we evaluated cell death and the abundance of virally encoded GFP after infection. During VSV infection, we found that most of the WT cells (~85%) were dead 24 hours after infection. In contrast, most Xbp1−/− MEFs (~74%) were resistant to cell death and accumulated higher amounts of viral protein as determined by measurement of GFP abundance (Fig. 2, A and B). Similarly, whereas a large proportion of HSV-infected WT cells underwent cell death, infected Xbp1−/− MEFs were resistant to death (Fig. 2, C and D). To determine whether acute ablation of Xbp1 expression would have a similar effect, we treated WT MEFs with small interfering RNA (siRNA) targeting Xbp1. Xbp1 knockdown strongly suppressed VSV-induced cell death and enhanced production of virally encoded GFP (fig. S2, A and B), consistent with the results of experiments with Xbp1−/− MEFs. Reconstitution of Xbp1−/− MEFs with plasmid-encoded Xbp1 restored VSV-induced cell death and restricted the production of virally encoded GFP (fig. S2C). To determine whether these findings extended to additional cell types, we cultured bone marrow–derived macrophages (BMDMs) from mice with a tamoxifen-inducible conditional Xbp1 deletion (Xbp1flox/flox ESR Cre) (20). Xbp1Δ BMDMs were resistant to death during VSV infection (Fig. 2, E and F, and fig. S2D), indicating that Xbp1 genetic deficiency results in protection from cell death in fibroblasts and macrophages. These results suggest that there should not be an Xbp1-dependent antiviral phenotype against viruses that do not trigger the death of infected cells. We found that infection with a VSV-G pseudotyped lentivirus encoding GFP did not result in host cell death (fig. S3A). In this case, we did not observe enhanced production of virally encoded GFP in Xbp1−/− MEFs (fig. S3, A and B). These findings suggest that impaired control of VSV and HSV infection by Xbp1−/− MEFs was directly related to their resistance to cell death.

Fig. 2 Xbp1-deficient cells are resistant to cell death during infection with VSV and HSV.

(A to D) WT and Xbp1−/− MEFs were left uninfected (mock) or were infected with VSV-GFP [(A) and (B)] or HSV-1–GFP [(C) and (D)] for 24 hours. Cell death was then assessed with a membrane-impermeant, amine-reactive fluorescent dye, which was measured by flow cytometry. Data are from one experiment and are representative of three experiments (A and C). The percentages of dead cells were then determined. Data are means ± SD of three independent experiments (B and D). (E and F) BMDMs were cultured from Xbp1flox/flox ESR Cre+ (Xbp1Δ) or Cre littermate (WT) mice in the presence of tamoxifen. Cells were infected with VSV-GFP at the indicated MOI for 24 hours. Viability was then assessed by measuring 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) reduction. Data are means ± SD of three replicates and are representative of three experiments (E and F). **P < 0.01 compared to WT, unpaired t test.

Apoptosis limits the replication of VSV and HSV

The end point of cell death can result from numerous upstream signaling pathways. VSV infection induces apoptotic cell death in Jurkat cells (21) and MCF-7 breast adenocarcinoma cells (22). WT MEFs infected with VSV demonstrated active caspase-3 (Fig. 3A), indicating apoptotic cell death in these cells. In contrast, Xbp1−/− MEFs failed to activate caspase-3 during VSV infection (Fig. 3A). Similarly, HSV-infected WT MEFs contained active caspase-3, and Xbp1−/− MEFs were resistant to the activation of this apoptotic effector (Fig. 3B). These findings were not limited to MEFs because the activation of caspase-3 and caspase-7 also occurred in VSV-infected BMDMs and Xbp1Δ BMDMs were resistant to VSV-induced caspase-3 activation (Fig. 3C).

Fig. 3 Apoptosis induced by VSV and HSV limits viral infection.

(A to B) WT and Xbp1−/− MEFs were left uninfected (mock) or were infected with VSV-GFP (A) or HSV-1–GFP (B) for 24 hours. Cells were then stained with an antibody specific for active (cleaved) caspase-3, which was measured by flow cytometry. Data are from one experiment and are representative of three experiments. (C) BMDMs were cultured from Xbp1flox/flox ESR Cre+ (Xbp1Δ) or Cre littermate (WT) mice in the presence of tamoxifen. Cells were infected with VSV-GFP at the indicated MOI for 7 hours. Caspase-3 activity was then assessed by measuring fluorometric substrate cleavage and is shown relative to that in uninfected cells. Data are means ± SD of three replicates and are representative of three experiments. (D to F) MEFs were infected in the presence of zVAD to inhibit caspase activity. Twenty-four hours after infection, cell death was assessed with a membrane-impermeant, amine-reactive fluorescent dye, which was measured by flow cytometry. The extent of infection was determined by measuring the relative abundance of GFP by flow cytometry. Data are from one experiment and are representative of three independent experiments. *P < 0.01 compared to WT, unpaired t test.

Some viruses induce apoptosis as a means of viral transmission and avoidance of the immune system (23). In other cases, apoptosis is beneficial for the host and limits viral replication. We observed decreased abundance of virally encoded GFP in the population of dead cells during HSV infection of WT MEFs (Fig. 2C), suggesting that apoptosis may limit viral replication. To test this hypothesis, we added a caspase inhibitor, carbobenzoxy-valyl-alanyl-aspartyl-[O-methyl]- fluoromethylketone (zVAD), to infected cells. We found that zVAD prevented the death of VSV-infected MEFs (Fig. 3D) and led to the increased abundance of virally encoded GFP (Fig. 3, D and E), phenocopying the result obtained from experiments with Xbp1−/− MEFs. Similarly, inhibition of apoptosis with zVAD increased the abundance of virally encoded GFP in HSV-infected MEFs, to an extent similar to that attained in Xbp1−/− MEFs (Fig. 3F). In addition, overproduction of the antiapoptotic protein BCL2 limited the death of VSV-infected cells and led to the increased abundance of virally encoded GFP (fig. S3C). Together, these findings suggest that Xbp1-deficient cells are resistant to virus-induced apoptosis and that apoptosis directly limits viral replication.

Resistance to virally induced apoptosis in Xpb1−/− cells is independent of Beclin 1 and CHOP

ER stress is associated with autophagy, which regulates cell survival (24). In particular, XBP1 promotes transcription of the gene encoding the autophagy component, Beclin 1 (25). Consistent with these data, we found decreased Beclin 1 in Xbp1-deficient cells (fig. S4A). However, knockdown of Beclin 1 with Beclin 1–specific siRNA did not affect VSV infection or the induction of cell death in either WT or Xbp1-deficient MEFs (fig. S4B). Our finding that Xbp1-deficient cells were resistant to VSV- and HSV-induced apoptosis suggests the possibility that apoptosis during viral infection may directly result from XBP1-mediated transcriptional activity. During ER stress, XBP1 partially contributes to the production of the apoptosis mediator CHOP (C/EBP homologous protein) (26), an induction process that may play a role in virus-induced apoptosis. As a functional control, transfection with Chop-specific siRNA prevented the death of MEFs treated with the ER stress–inducing agents tunicamycin and thapsigargin (fig. S5, A and B). In contrast, Chop knockdown did not prevent the death of VSV- or HSV-infected MEFs (fig. S5B). Further arguing against a direct role for XBP1-mediated transcriptional activity in virally induced cell death, we did not observe Xbp1 splicing (fig. S6A) or induction of the expression of the UPR-responsive genes Hspa5 [encoding BiP (binding immunoglobulin protein)] and Chop during VSV infection (fig. S6, B and C), consistent with published observations for VSV (27) and HSV (28, 29).

Xbp1-deficient cells are resistant to the intrinsic pathway of apoptosis

Because we did not find evidence for XBP1-mediated transcriptional activity in promoting apoptosis during infection, we hypothesized that Xbp1-deficient cells may be inherently resistant to apoptosis in general. We therefore treated Xbp1−/− MEFs with a panel of apoptosis-inducing stimuli. Staurosporine and gliotoxin stimulate the intrinsic or mitochondrial pathway of apoptosis (3033), whereas ligation of the tumor necrosis factor (TNF) and Fas receptors initiates the extrinsic apoptotic pathway (34). Xbp1−/− MEFs were specifically resistant to stimuli that induced the intrinsic pathway of apoptosis, both as demonstrated by increased viability (Fig. 4A) and impaired caspase-3 activation (Fig. 4B). Consistent with previous studies demonstrating a protective role of Xbp1 during ER stress (26, 3537), Xbp1−/− MEFs were slightly more susceptible to death induced by tunicamycin (Fig. 4A). In contrast, there was no difference between WT and Xbp1−/− MEFs in necrotic death induced by a high concentration of cycloheximide (Fig. 4A). These findings were further verified in experiments with Xbp1Δ BMDMs, which demonstrated resistance to staurosporine and the selective Bcl-2 inhibitor ABT-737 (38) but not to TNF-induced apoptosis (Fig. 4C). Thus, Xbp1 genetic deficiency results in specific protection from the intrinsic pathway of apoptosis.

Fig. 4 Xbp1-deficient cells are resistant to the intrinsic pathway of apoptosis.

(A and B) WT and Xbp1−/− MEFs were treated with staurosporine (sts), gliotoxin (glio), TNF and low-dose cycloheximide (TNF), Fas antibody and low-dose cycloheximide (Fas), and tunicamycin (TM) to induce the UPR, or high-dose cycloheximide (CHX). Twenty-four hours later, viability was assessed by measuring MTS reduction (A). Seven hours after treatment, caspase-3 activity was assessed by measuring fluorometric substrate cleavage and is shown relative to WT cells (B). Data are means ± SD of three replicates and are representative of three experiments. (C) BMDMs were cultured from Xbp1flox/flox ESR Cre+ (Xbp1Δ) or Cre littermate (WT) mice in the presence of tamoxifen and treated with the indicated inducers of cell death as described in (A). Twenty-four hours later, viability was assessed by measuring MTS reduction. Data are means ± SD of three replicates and are representative of three experiments. *P < 0.05; **P < 0.001 compared to WT, unpaired t test.

The resistance of Xbp1-deficient cells to apoptosis results from the activation of IRE1α

Xbp1 deficiency results in activation of its upstream enzyme IRE1α, which degrades specific cytosolic RNA targets (16, 17). Although they cannot make XBP1s protein, Xbp1-deficient cells transcribe mRNA that contains the IRE1α cleavage sites. Consistent with previous reports, we observed IRE1α activation in Xbp1-deficient cells, indicated by Xbp1 mRNA splicing (fig. S7A). The magnitude of IRE1α activation in Xbp1-deficient cells was not as robust as the response to canonical UPR stimulation with thapsigargin, nor were classical RIDD substrates diminished (fig. S7B), consistent with other studies (39).

To determine whether IRE1α was involved in resistance to apoptosis, we knocked down Ire1α with siRNA (fig. S7C). As a functional control, Ire1α-specific siRNA efficiently prevented Xbp1 mRNA splicing (fig. S7D). IRE1α knockdown in Xbp1−/− MEFs reversed resistance to VSV-induced cell death (Fig. 5A and fig. S7E). Expression of human IRE1α, which was resistant to mouse Ire1α-specific siRNA, prevented this reversal (fig. S7F). IRE1α knockdown alone was minimally cytotoxic but restored caspase-3 activation in Xbp1−/− MEFs in response to staurosporine (Fig. 5B) and sensitized these cells to staurosporine-induced cell death (Fig. 5C). To determine whether the RNase activity of IRE1α mediated resistance to apoptosis, we treated Xbp1-deficient cells with the selective IRE1α nuclease inhibitor 4μ8C (8-formyl-7-hydroxy-4-methylcoumarin) (40). We found that 4μ8C reversed the resistance of Xbp1-deficient cells to apoptosis both during infection with VSV and during treatment with staurosporine (Fig. 5D). As a negative control, the structurally similar compound AMC (7-amino-4-methylcoumarin) had no effect at an equimolar concentration (fig. S2D). These findings indicate that the RNase activity of IRE1α contributes to resistance to the intrinsic pathway of apoptosis observed in the setting of Xbp1 deficiency. We consistently observed Xbp1 mRNA splicing to a small extent in WT cells (fig. S7, A and C), suggesting that IRE1α has some basal activity (41), which could regulate apoptotic responses in WT cells. Consistent with this, IRE1α knockdown in WT MEFs was not toxic alone (Fig. 5, B, C, and E) but sensitized cells to VSV-induced cell death and limited viral replication (Fig. 5, E and F).

Fig. 5 The resistance of Xbp1-deficient cells to apoptosis results from the activation of IRE1α.

(A to F) Analysis of the effects of IRE1α knockdown on cell death. (A, E, and F) WT and Xbp1−/− MEFs were transfected with siRNA targeting Ire1α or control siRNA (ctrl siRNA). Cells were then left uninfected (mock) or infected with VSV-GFP for 24 hours. Cell death was then assessed with a membrane-impermeant, amine-reactive fluorescent dye, and the relative abundance of GFP was measured by flow cytometry. Data are from one experiment and are representative of three (A) or two (E and F) independent experiments. (B and C) The indicated siRNA-transfected MEFs were left untreated (mock) or treated with staurosporine. Seven hours later, caspase-3 activity was assessed by measuring fluorometric substrate cleavage and is shown relative to that in untreated WT cells (B). Twenty-four hours after treatment, viability was assessed by measuring MTS reduction (C). (D) BMDMs were cultured from Xbp1flox/flox ESR Cre+ (Xbp1Δ) or Cre littermate (WT) mice in the presence of tamoxifen and the IRE1α inhibitor 4μ8C. Cells were then infected with VSV-GFP at an MOI of 2 or were treated with staurosporine. Viability was assessed 24 hours later by measuring MTS reduction. Data are means ± SD of three replicates and are representative of three experiments (B, C, and D). *P < 0.01; **P < 0.001, unpaired t test.

IRE1α targets the proapoptotic miRNA miR-125a

Studies of coding genes targeted by IRE1α for RIDD have not revealed obvious candidates to explain our observed IRE1α-mediated resistance to apoptosis (9, 42). Thus, we focused on miRNAs, which are also targeted by RIDD (43). To this end, we performed miRNA profiling of Xbp1-deficient cells and found four miRNAs that were decreased in abundance in Xbp1-deficient cells with active IRE1α (Fig. 6A). The miRNA miR-125a was represented twice among these differentially expressed miRNAs. Quantitative polymerase chain reaction (PCR) analysis confirmed the decrease in miR-125a abundance in Xbp1-deficient cells, which was restored by reconstitution of the cells with plasmid-encoded Xbp1 (fig. S8A). The miRNA miR-125a sensitizes cells to apoptosis and is thought to negatively regulate antiapoptotic Bcl-2 family members, including B cell lymphoma–extra large (Bcl-xL) and myeloid cell leukemia 1 (Mcl-1) (4446). In accordance with decreased miR-125a, we found an increase in the abundances of antiapoptotic Bcl-xL and Mcl-1 proteins in Xbp1-deficient cells (Fig. 6B). To test whether these effects were dependent on IRE1α, we crossed Xbp1flox/flox × ESR Cre+ mice to Ern1flox/flox mice. BMDMs obtained from Xbp1flox/flox × Ern1flox/flox × ESR Cre+ mice were treated with tamoxifen to generate XBP1Δ IRE1αΔ cells. We found that the protein amounts of both Bcl-xL and Mcl-1 in XBP1Δ IRE1αΔ BMDMs were reduced compared to those in XBP1Δ cells, albeit not to the amount observed in the WT cells (Fig. 6B). These results indicated that antiapoptotic Bcl-xL and Mcl-1 proteins were increased in abundance in Xbp1-deficient cells, in a manner largely dependent on IRE1α. Consistent with our observation of sensitization to apoptosis by IRE1α knockdown in WT cells (Fig. 5E), IRE1αΔ cells had increased miR-125a abundance (fig. S8B). Inhibition of the RNase activity of IRE1α with the selective IRE1α nuclease inhibitor 4μ8C was sufficient to increase the abundance of miR-125a (fig. S8B).

Fig. 6 IRE1α mediates reduction in proapoptotic miR-125a.

(A) BMDMs were cultured from Xbp1flox/flox ESR Cre+ (Xbp1Δ) or Cre littermate (WT) mice in the presence of tamoxifen. Volcano plot demonstrating the distribution of miRNAs between WT and Xbp1Δ BMDMs measured using the NanoString nCounter assay. Data are from one experiment with quadruplicate samples. (B) BMDMs were cultured from Xbp1flox/flox ESR Cre+ (Xbp1Δ), Xbp1flox/flox Ern1 flox/flox ESR Cre+ (Xbp1Δ Ire1αΔ), or Cre littermate (WT) mice in the presence of tamoxifen. The relative abundances of Bcl-xL, Mcl-1, and β-actin in the cell lysates were determined by Western blotting and densitometry. The ratio of Bcl-xL or Mcl-1 to β-actin is shown, normalized to that in WT cells. Data are means ± SD of three independent experiments. a.u., arbitrary units. (C and D) WT and Xbp1−/− MEFs were transfected with negative control miRNA mimetic (miR-ctrl) or a miR-125a mimetic. Cells were left untreated (mock) or treated with staurosporine. Seven hours later, caspase-3 activity was assessed by measuring fluorometric substrate cleavage and is shown relative to that in untreated WT cells (C). Twenty-four hours after treatment, viability was assessed by measuring MTS reduction (D). Data are means ± SD of three replicates and are representative of two experiments. (E) The indicated miRNA-transfected MEFs were infected with VSV-GFP for 24 hours. Cell death was then assessed with a membrane-impermeant, amine-reactive fluorescent dye, which was measured by flow cytometry. The extent of infection was determined by measuring the relative abundance of GFP by flow cytometry. Data are from one experiment and are representative of two independent experiments. *P < 0.01; **P < 0.001, unpaired t test.

Finally, we wished to examine the extent to which miR-125a degradation by IRE1α was responsible for the prosurvival phenotype of the Xbp1-deficient cells. Reconstitution of miR-125a with an miRNA mimetic was sufficient to restore caspase-3 activation in Xbp1−/− MEFs in response to staurosporine (Fig. 6C) and to sensitize Xbp1-deficient cells to the intrinsic pathway of apoptosis (Fig. 6D and fig. S8C). Reconstitution with miR-125a also reversed the resistance of Xbp1−/− MEFs to VSV-induced cell death (Fig. 6E). Finally, neutralizing miR-125a in WT cells with an miRNA hairpin inhibitor resulted in an antiapoptotic state resembling that observed in Xbp1-deficient cells (fig. S8D). Together, these findings suggest that the IRE1α-dependent decrease in miR-125a abundance contributes to the resistance to apoptosis.

IRE1α activation during HCV infection mediates resistance to apoptosis

Some viruses encode genes that promote IRE1α activation. HCV NS4B activates IRE1α (18), and IRE1α activation is also seen in HCV-infected cells (47). Curiously, in cells expressing NS4B, IRE1α splices Xbp1 mRNA, but XBP1 targets are not transcribed (18), suggesting that the virus uses IRE1α for another reason. Furthermore, HCV is suggested to cause resistance to the intrinsic pathway of apoptosis (48, 49), although the mechanism of this effect remains unknown.

Consistent with previous reports (18), we detected IRE1α activation as indicated by spliced XBP1 mRNA in cells transiently transfected with a plasmid encoding NS4B (Fig. 7A). The abundance of miR-125a was decreased (fig. S8E), and NS4B expression induced the IRE1α-dependent resistance to staurosporine (Fig. 7, B and C). We infected Huh-7.5 human hepatoma cells with trans-packaged HCV replicons encoding Gaussia luciferase (Gluc), which enabled us to quantitate HCV replication over time (50). IRE1α inhibition alone was not cytotoxic, but it sensitized HCV-infected cells to death (Fig. 7D). Furthermore, inhibition of IRE1α decreased the secretion of virally encoded luciferase, a marker of viral replication (Fig. 7E). These findings suggest that IRE1α activation during HCV infection may promote viral replication by inhibiting the death of the infected cells. To determine whether these findings extended to human HCV infection, we quantified spliced XBP1 mRNA in the human liver tissue of patients infected with HCV. We detected HCV-associated IRE1α activation, as indicated by an increase in spliced XBP1 mRNA abundance in liver tissue from HCV-infected patients compared to that in the liver tissue of HCV-negative controls (Fig. 7F). In addition to IRE1α activation, HCV-infected patients also exhibited statistically significantly reduced miR-125a abundance (Fig. 7G), suggesting that IRE1α activation during human HCV infection may confer resistance to apoptosis.

Fig. 7 IRE1α-mediated apoptosis resistance induced by HCV NS4B.

(A) HeLa cells were transfected with constructs expressing GFP alone (vector) or together with HCV NS4B. RNA was isolated 72 hours later. XBP1 mRNA maturation from the unspliced (u) to spliced (s) form was analyzed by reverse transcription PCR (RT-PCR). Data are from one experiment and are representative of two independent experiments. (B and C) Transfected cells were treated with IRE1α inhibitor 2 for 24 hours, and staurosporine was then added. Twenty hours later, cell death was assessed with a membrane-impermeant, amine-reactive fluorescent dye, which was measured by flow cytometry. Data are from one experiment and are representative of three independent experiments (B). The percentage of dead cells among transfected GFP-positive cells was calculated (C). Data are means ± SD of three independent experiments. (D and E) Huh-7.5 cells were infected with trans-packaged HCV encoding luciferase. Forty-eight hours later, viability was determined by measuring cellular adenosine triphosphate (D). Secretion of virally encoded luciferase was measured 24 and 48 hours after infection (E). Data are means ± SD of three replicates and are representative of two experiments. (F and G) RNA was isolated from the liver tissue of HCV-infected patients (n = 11) and HCV-negative controls (n = 6). Expression of spliced XBP1 and miR-125a (relative to an internal control) was determined by quantitative RT-PCR. Data are means ± SEM. *P < 0.05; **P < 0.01, unpaired t test. ***P < 0.01, Mann-Whitney test.

DISCUSSION

Here, we examined the IRE1α-XBP1 branch of the UPR in innate antiviral defense. We uncovered an unexpected role for Xbp1 and IRE1α in modulating susceptibility to the intrinsic pathway of apoptosis, which is induced during VSV and HSV infection and plays a critical role in limiting viral replication. Xbp1−/− MEFs were protected from virally induced cell death and, as a consequence, sustained more viral replication despite an increased IFN response compared to that of WT cells. In Xbp1−/− cells with active IRE1α, the abundance of the proapoptotic miRNA miR-125a was decreased, conferring resistance to the intrinsic apoptotic pathway. IRE1α activation by HCV NS4B in WT cells also conferred resistance to apoptosis and promoted viral replication. Therefore, Xbp1-deficient cells with active IRE1α gained apoptosis resistance, which suggests that IRE1α knockdown or inhibition is a model for the loss of apoptosis resistance. Finally, we observed IRE1α activation and substantially less miR-125a abundance in liver biopsies from HCV-infected patients, suggesting the in vivo relevance of the survival strategy used by HCV. These results highlight a previously unappreciated role of the IRE1α-XBP1 axis in the regulation of apoptosis and its consequences in viral susceptibility.

Previous studies showed that after the engagement of TLRs and RLRs, XBP1 plays an important role in enhancing cytokine and IFN production in macrophages and dendritic cells (1113). Given the observation that transfection with poly(I:C) induced less IFN production in Xbp1−/− MEFs compared to that in WT MEFs, our results are consistent with these previous findings that Xbp1 promotes RLR signaling for IFN production in MEFs. However, despite this impairment in IFN production downstream of RLRs, infected Xbp1-deficient MEFs produced large amounts of IFN. We speculate that the enhanced IFN response observed in Xbp1−/− MEFs may result from both prolonged cellular survival and an accumulation of viral PAMPs. Apoptotic caspases can cleave and inactivate signaling proteins important for the IFN response, suggesting that the apoptotic process directly antagonizes the IFN response (5154). In addition, IRE1α can cleave host RNA for RLR stimulation (15). Therefore, our results highlight a distinct consequence of IRE1α activation, whereby Xbp1 deficiency results in a robust prosurvival response, leading to prolonged RLR stimulation that mitigates impairment in RLR signaling to generate enhanced IFN responses. The prosurvival signals induced through IRE1α activation are so dominant that they overcome ISG-mediated antiviral functions and enable virus replication.

In addition to protection from virus-induced death, we found that Xbp1 deficiency conferred resistance to the intrinsic pathway of apoptosis stimulated by various chemical inducers. IRE1α activation in Xbp1-deficient cells contributed to this apoptosis resistance because siRNA-mediated knockdown of Ire1α expression or inhibition of IRE1α nuclease function rendered the Xbp1-deficient cells susceptible to apoptosis. RIDD targets both coding and noncoding RNAs, including miRNAs (42). We found that IRE1α reduced the amount of miR-125a, leading to the enhanced expression of its target genes. The targets include genes encoding prosurvival members of the Bcl-2 family (4446). miRNA controls target genes at the transcriptional and translational levels (55). We found that the prosurvival proteins Bcl-xL and Mcl-1 were present in increased amounts. The biological targets and functions of the other miRNAs identified in this study, namely miR-1224 and miR-804, have not yet been well described. The relevance of these other miRNAs in IRE1α-dependent phenotypes will be investigated in future studies.

Our study revealed an unexpected role for IRE1α in controlling apoptosis. Many studies of the UPR have been performed with high concentrations of pharmacological inducers of ER stress, such as tunicamycin and thapsigargin, which inevitably lead to cell death. In these experimental settings of irremediable ER stress, various mechanisms have been proposed for inducing apoptosis (56). The net effect of IRE1α activity in promoting cell death compared to cell survival has been controversial, with some studies suggesting that IRE1α promotes cell survival (5759), and others suggesting that IRE1α promotes cell death (6062). Specifically, IRE1α was proposed to induce apoptosis by degrading miRNAs that lead to increased caspase-2 abundance (60), although there are conflicting data challenging this observation (63). IRE1α has been linked to apoptosis in cells irreversibly damaged by the activation of the JNK (c-Jun N-terminal kinase)–ASK1 (apoptosis signal–regulating kinase 1) pathway (64, 65). IRE1α appears to have opposing roles in regulating apoptosis: prosurvival in the absence of irreversible UPR but proapoptotic under irreversible UPR. This double-edged nature of IRE1α may be important to consider for the use of IRE1α inhibitors in the treatment of various human diseases (66).

We speculate that IRE1α may represent an ancient form of cell protection that has been subverted by some viruses for their own replicative advantage. HCV and other members of the Flaviviridae family induce ER stress and activate IRE1α. Our results suggest that HCV NS4B induces IRE1α-dependent protection from apoptosis, which may favor the development of chronic infection and hepatocellular carcinoma. In the setting of avian coronavirus infection, IRE1α also promotes cell survival in association with JNK and Akt regulation (67). Influenza virus induces only the IRE1α arm of the UPR (not ATF6 or PERK), and inhibitors of IRE1α reduce viral replication (68). Therefore, our findings predict that the IRE1α-RIDD pathway could be exploited as a novel target of intervention against viral infections.

MATERIALS AND METHODS

Cells and viruses

Xbp1+/+ and Xbp1−/− MEFs were gifts from L. H. Glimcher (Weill Cornell Medical College, New York, NY). H1-HeLa cells stably overexpressing the antiapoptotic protein BCL2 have been previously described (69). MEFs and H1-HeLa cells were propagated in high-glucose Dulbecco’s modified Eagle’s medium (DMEM) (Gibco) and supplemented with 10% heat-inactivated fetal bovine serum (FBS), 1% Hepes, and penicillin (100 U/ml) and streptomycin (100 μg/ml) (Gibco). Huh-7.5 cells were propagated in high-glucose DMEM with 10% heat-inactivated FBS and 1 mM nonessential amino acids (Invitrogen). Xbp1flox/flox (20) (a gift from L. H. Glimcher, Weill Cornell Medical College), Ern1flox/flox (70) [RIKEN BioResource Center (BRC), Japan], and CAGGCre-ERTM (the Jackson Laboratory) mice were bred in the Yale animal facility. All procedures performed in this study complied with the federal guidelines and institutional policies set by Yale Animal Care and Use Committee. BMDMs were prepared from 6- to 12-week-old male and female mice according to a previously described method (71) and cultured in 0.2 μM 4-hydroxytamoxifen (Sigma) during days 2 to 4 of differentiation to induce Cre-mediated recombination. The following genotype combinations of BMDMs were treated with tamoxifen to generate the cells described in Fig. 6: Xbp1flox/flox × ESR Cre+ (XBP1Δ), Xbp1flox/flox × Ern1flox/flox × ESR Cre+ (XBP1Δ IRE1αΔ), or Xbp1flox/flox × ESR Cre (WT). VSV-G–GFP was a gift from J. Rose (Yale University, New Haven, CT) and A. Geballe (Fred Hutchinson Cancer Research Center, Seattle, WA) and was maintained and titered in baby hamster kidney cells. HSV-1–GFP (72) was a gift from P. Desai and S. Person (Johns Hopkins University, Baltimore, MD) and was maintained and titered in Vero cells. VSV-G pseudotyped lentivirus was made by harvesting culture supernatants of human embryonic kidney 293T cells transfected with plasmids encoding VSV-G, GFP, and Gag-Pol. Trans-packaged HCV replicons were prepared by transfecting the JFH/Gluc replicon into Huh-7.5[core-NS2] cells as previously described (50)

Infection and stimulation of cells

Cells were incubated for 1 hour with VSV or HSV-1 in serum-free medium or pseudotyped virus in phosphate-buffered saline 0.1% bovine serum albumin, and then the inoculum was removed, and incubation continued in complete medium. Cells were incubated for 4 hours with trans-packaged HCV, and then the inoculum was removed, and incubation continued in complete medium containing 60 μM IRE1 Inhibitor II (Calbiochem). zVAD (Invivogen) was added at 20 to 100 μM after removal of the viral inoculum. Poly(I:C) (1 μg/ml) was delivered complexed to Lipofectamine 2000 (Invitrogen). Cells were treated with 0.1 to 1 μM staurosporine (Enzo Life Sciences), 1 μM gliotoxin (Sigma), 10 μM ABT-737 (Santa Cruz Biotechnology), TNF (50 ng/ml) + cycloheximide (0.1 μg/ml) (Abcam), antimouse CD95 (2 μg/ml) (BD Pharmingen), Fas-activating antibody + cycloheximide (0.1 μg/ml), tunicamycin (10 to 100 μg/ml) (Sigma), 1 μM thapsigargin (Calbiochem), or cycloheximide (100 μg/ml) (high-dose CHX). Cells were treated with the IRE1 inhibitor 4μ8C (Calbiochem) or the structurally similar compound AMC (Sigma) at 25 μM for 3 days before infection or apoptosis induction or 40 μM IRE1 Inhibitor II (Calbiochem) for 24 hours before apoptosis induction.

Plasmids

pFLAG.XBP1u.CMV2 (Addgene plasmid #21832, from D. Ron) (73), the empty vector control c-Flag pcDNA3 (Addgene plasmid #20011, from S. Smale) (74), hIRE1a.pcD (Addgene plasmid #21892, from R. Kaufman), and hIRE1a wt (Addgene plasmid #20744, from F. Urano) (75) were used. MEFs were transfected using TransIT-2020 (Mirus Bio). HCV NS4B was expressed with an authentic N-terminal Ala residue by fusing a human ubiquitin gene to a codon-optimized NS4B gene (76): Ubiquitin was amplified by using primers YO-0905 (TTA ATT AAC GAG GAT CCC GCC ACC ATG CAG ATC TTC GTG AAG AC) and YO-0928 (TCG ATC AGG GCT GCT CTG CTG GCT CCA CCG CGG AGA CGC AGC ACC); codon-optimized NS4B was amplified by using primers YO-0927 (GGT GCT GCG TCT CCG CGG TGG AGC CAG CAG AGC AGC CCT GAT CGA) and YO-0931 (GTT TAA ACT TAA CAA GGG ATG GGG CAG TCC T). Ubi-NS4B was then assembled in secondary PCRs with primers YO-0905 and YO-0931, cloned into pCR2.1 (Invitrogen) for sequencing, and then subcloned into pIRES2-EGFP (Clontech) by using the common Sac I and Pst I restriction sites. H1-HeLa cells were transfected using TransIT-HeLa (Mirus Bio).

Expression analysis

RNA isolated using the RNeasy kit (Qiagen) was used to synthesize complementary DNA (cDNA) using the iScript cDNA Synthesis Kit (Bio-Rad), and quantitative PCR was performed on a Stratagene Mx3000P or Bio-Rad CFX Connect using SYBR Green (Bio-Rad) with the following primers (all primers listed in the 5′ to 3′ orientation): Ifna4, CTG CTA CTT GGA ATG CAA CTC (forward) and CAG TCT TGC CAG CAA GTT GG (reverse); Ifnb1, GCA CTG GGT GGA ATG AGA CTA TTG (forward) and TTC TGA GGC ATC AAC TGA CAG GTC (reverse); Mx1, AGT CCT TTC CAC AGG CAG AA (forward) and CAT TGA GAG AAA CTC ACC TAA GAA C (reverse); Xbp1s, GAG TCC GCA GCA GGT (forward) and GTG TCA GAG TCC ATG GGA (reverse); Hspa5 (Bip), TCA TCG GAC GCA CTT GGA (forward) and CAA CCA CCT TGA ATG GCA AGA (reverse); Chop, GTC CCT AGC TTG GCT GAC AGA (forward) and TGG AGA GCG AGG GCT TTG (reverse); Blos1, CAA GGA GCT GCA GGA GAA GA (forward) and GCC TGG TTG AAG TTC TCC AC (reverse); Pdgfrb, AAC CCC CTT ACA GCT GTC CT (forward) and TAA TCC CGT CAG CAT CTT CC (reverse); and human spliced XBP1, TGC TGA GTC CGC AGC AGG TG (forward) and GCT GGC AGG CTC TGG GGA AG (reverse). For analysis of Xbp1-splicing, primers flanking the spliced sequences in Xbp1 mRNA [ACA CGC TTG GGA ATG GAC AC (forward) and CCA TGG GAA GAT GTT CTG GG (reverse)] were used for PCR amplification, and products were separated by electrophoresis through a 3% agarose gel and visualized by ethidium bromide staining.

miRNA expression analysis

RNA isolated using the miRNeasy Mini Kit (Qiagen) was subjected to miRNA profiling using the nCounter mouse miRNA expression assay version 1.5 (NanoString) according to the manufacturer’s protocol. Data were analyzed by using the nSolver software with normalization to the geometric mean of the top 100 miRNAs as recommended by the manufacturer. Quantitative RT-PCR with the miRCURY Universal RT microRNA qPCR system (Exiqon) was used to measure miR-125a-5p. Expression was calculated relative to the manufacturer’s suggested endogenous control (miR-103a-3p), with equivalent results also obtained relative to miR-16-5p.

Assessment of cell death

Cells were stained with the LIVE/DEAD Fixable Far Red Dead Cell Stain Kit (Molecular Probes) and analyzed by flow cytometry on a BD FACSCalibur or BD LSRFortessa. Viability was also assessed by measuring MTS reduction using the CellTiter 96 AQueous One Solution Cell Proliferation Assay (Promega). Cells were stained with the Caspase-3, Active Form, Apoptosis Kit (BD Pharmingen) and analyzed by flow cytometry on a BD FACSCalibur. Caspase-3/7 activity was measured using the SensoLyte Homogeneous Rh110 Caspase-3/7 Assay Kit (AnaSpec).

Western blotting

Cell pellets were lysed in SDS sample buffer (Cell Signaling Technology). Proteins were separated by SDS–polyacrylamide gel electrophoresis, transferred to polyvinylidene difluoride membranes, and incubated with antibodies against Mcl-1 (BioLegend, 613601), Bcl-xL (Cell Signaling Technology, 54H6), or β-actin (Cell Signaling Technology, 13E5).

Intracellular staining

Cells were fixed in Cytofix/Cytoperm solution (BD) on ice, washed with Perm/Wash buffer (BD), and stained with DyLight 550–conjugated antibody against Beclin 1 (Novus Biologicals, NB110-87318R) or an equal concentration of isotype control antibody. Washed cells were analyzed by flow cytometry on a BD LSRFortessa flow cytometer.

siRNA, miR mimetics, and inhibitors

Gene-specific siGENOME siRNA or siGENOME Non-Targeting siRNA #4 (which targets firefly luciferase mRNA and has at least four mismatches to all mouse genes) obtained from Dharmacon (Thermo Fisher Scientific) was delivered complexed to Lipofectamine RNAiMAX (Invitrogen). After incubation for 48 hours, cells were replated at equal density before infection or apoptosis induction. miR mimetics, miRIDIAN microRNA hairpin inhibitor negative control #1, and miRIDIAN miR-125a hairpin inhibitor were obtained from Dharmacon (Thermo Fisher Scientific) and delivered complexed to Lipofectamine RNAiMAX (Invitrogen).

Luciferase assay

Conditioned cell culture medium was collected at 24 or 48 hours after infection, clarified by centrifugation, mixed with 1/4 volume 5× lysis buffer (New England Biolabs), and assayed for luciferase activity (New England Biolabs).

Liver specimens

Liver samples from percutaneous biopsies of liver transplant recipients chronically infected with HCV or from HCV-negative control liver transplant recipients were obtained with the approval of the University of Washington Institutional Review Board. Tissue was archived in RNAlater and stored at −80°C. RNA was isolated using the miRNeasy Mini Kit (Qiagen).

Statistical analyses

Sample size was chosen according to previous experience in similar experiments; all samples were included in analysis. The unpaired Student’s t test or the Mann-Whitney test was used for comparisons between two groups. P values of less than 0.05 were considered statistically significant.

SUPPLEMENTARY MATERIALS

www.sciencesignaling.org/cgi/content/full/10/482/eaai7814/DC1

Fig. S1. Increased IFN and ISG expression during VSV infection of Xbp1-deficient MEFs.

Fig. S2. Knockdown of Xbp1 with siRNA mimics Xbp1 deficiency, and reconstitution with Xbp1 reverses resistance to cell death.

Fig. S3. Xbp1 deficiency does not increase susceptibility to a noncytotoxic virus and mimics BCL2 overexpression.

Fig. S4. Resistance to virally induced apoptosis in Xpb1-deficient cells is independent of Beclin 1.

Fig. S5. Death of VSV- and HSV-infected cells does not require Chop.

Fig. S6. VSV infection does not activate the UPR.

Fig. S7. The resistance of Xbp1-deficient cells to apoptosis results from the activation of IRE1α.

Fig. S8. The miRNA miR-125a regulates resistance to apoptosis.

REFERENCES AND NOTES

Acknowledgments: We thank H. Dong and K. Hayashi for technical support; B. Cookson and N. Kaminski for sharing laboratory facilities and equipment; and L. Glimcher, J. Rose, A. Geballe, P. Desai, S. Person, D. Ron, S. Smale, R. Kaufman, and F. Urano for providing us with various reagents. Funding: This work was supported by funding from the Howard Hughes Medical Institute and NIH R01 AI054359 and AI064705 (to A.I.) and K08 AI119142 (to S.L.F.). S.L.F. was supported by NIH award T32 HL007974. B.D.L. was funded by R01 AI087925. Author contributions: S.L.F., T.R.J., R.D.M., T.I., C.S.L., B.D.L., and A.I. designed the experiments. S.L.F., T.R.J., R.D.M., and B.D.L. performed the experiments. S.L.F., T.R.J., R.D.M., and A.I. analyzed the data. S.L.F and A.I. wrote the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: The Xbp1+/+ and Xbp1−/− MEFs and Xbp1flox/flox mice require a materials transfer agreement (MTA) from Harvard University. The Ern1flox/flox mice require an MTA from RIKEN BRC. The following plasmids require an MTA from Addgene: pFLAG.XBP1u.CMV2, c-Flag pcDNA3, hIRE1a.pcD, and hIRE1a wt. The JFH-1 HCV replicon requires MTAs from Rockefeller University and Apath LLC (in the United States) or from the Tokyo Metropolitan Organization for Medical Research and Toray Industries (elsewhere). The replicon was packaged by using the HCV strain J6 structural genes, which were obtained under an MTA with the NIH. The Huh-7.5 cell line requires an MTA from Washington University and Apath LLC.
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