Nuclear IFI16 induction of IRF-3 signaling during herpesviral infection and degradation of IFI16 by the viral ICP0 protein

Abstract

Innate sensing of microbial components is well documented to occur at many cellular sites, including at the cell surface, in the cytosol, and in intracellular vesicles, but there is limited evidence of nuclear innate signaling. In this study we have defined the mechanisms of interferon regulatory factor-3 (IRF-3) signaling in primary human foreskin fibroblasts (HFF) infected with herpes simplex virus 1 (HSV-1) in the absence of viral gene expression. We found that the interferon inducible protein 16 (IFI16) DNA sensor, which is required for induction of IRF-3 signaling in these cells, is nuclear, and its localization does not change detectably upon HSV-1 d109 infection and induction of IRF-3 signaling. Consistent with the IFI16 sensor being nuclear, conditions that block viral DNA release from incoming capsids inhibit IRF-3 signaling. An unknown factor must be exported from the nucleus to activate IRF-3 through cytoplasmic STING, which is required for IRF-3 activation and signaling. However, when the viral ICP0 protein is expressed in the nucleus, it causes the nuclear relocalization and degradation of IFI16, inhibiting IRF-3 signaling. Therefore, HSV-1 infection is sensed in HFF by nuclear IFI16 upon release of encapsidated viral DNA into the nucleus, and the viral nuclear ICP0 protein can inhibit the process by targeting IFI16 for degradation. Together these results define a pathway for nuclear innate sensing of HSV DNA by IFI16 in infected HFF and document a mechanism by which a virus can block this nuclear innate response.

Viral infection elicits a number of antiviral innate immune responses, including the secretion of type I IFNs, IFN-α and -β, which act in an autocrine or paracrine manner to induce the production of IFN-stimulated genes (ISG) whose protein products mediate the antiviral state (1). In addition to mediating a localized response, IFN bridges the innate and adaptive immune responses to promote immunological memory and clearance of viral infection (2).

The antiviral response is induced by a set of germ-line–encoded pattern-recognition receptors (PRRs) (3) that recognize pathogen-specific moieties, which include viral genomic DNA and RNA species as well as RNA replication intermediates. These PRRs signal through distinct adaptor molecules but converge on the activation of TBK1 (4), a serine/threonine protein kinase that phosphorylates the constitutively expressed IFN-regulatory factor 3 (IRF-3) (5). Normally cytoplasmic, phosphorylated IRF-3 translocates to the nucleus where it promotes the transcription of IFN-responsive genes, including IFNβ (6).

The host cell is known to distinguish between viral and self nucleic acids by sensing both chemical and compartmentalization differences in these molecules. For instance, the membrane-bound Toll-like receptors (TLRs) 3 and 7/8 detect RNA species that accumulate in endosomes (7). In addition, resident phagocytic cells use these receptors to detect foreign pathogens during immune surveillance of tissues (8). Chemical differences between self and viral RNAs also exist and are detected by a class of cytosolic PRRs known as the “RIG-I–like receptors” (RLRs). The eponymous member of this class, RIG-I, recognizes dsRNA that contains 5′ triphosphates (9), which are unique to RNA virus infection. MDA5, another member of this family, also binds viral dsRNA but potentiates signaling based on the length of the RNA species it detects (10).

Although recognition of RNA virus infection is well understood, less is known about how DNA viruses are sensed by the host cell. The first DNA sensor identified was TLR9, which recognizes unmethylated CpG DNA in endosomal compartments and is particularly potent at detecting foreign DNA in plasmacytoid dendritic cells (11, 12). More recently, cytosolic PRRs have been identified that sense DNA virus infection in the cytoplasm; these include DAI, Pol III, and the DEAD/H-box helicase DDX41 (13-16). These sensors are thought to distinguish between cellular and viral DNA because of compartmentalization differences.

An interesting paradox in DNA sensing involves herpesviruses, which constitute a class of large dsDNA viruses that replicate in the nucleus of infected cells. Although DNA from these viruses is a potent activator of IRF-3, it is unclear how they are sensed during infection. Currently, there is little evidence that nuclear DNA sensing can induce a type I IFN response to virus infection. Recently, IFI16, a member of the PYHIN family of proteins, was implicated in the type I IFN response to HSV-1 (17). Although originally reported to be a cytosolic DNA sensor (17), IFI16 is localized in the nucleus of many cell types (18), making it a potential candidate for sensing nuclear HSV-1 DNA.

The importance of the IRF-3 pathway in restricting virus replication has made it a target for virus-mediated inhibition. HSV-1 induces an IRF-3–dependent type I IFN/ISG response in human fibroblasts in the absence of viral gene expression (19–21). This response is potently inhibited by viral gene expression, suggesting that a viral protein inhibits this response. Like many large DNA viruses, HSV-1 encodes multiple mechanisms for inhibiting IFN expression (22). For instance, the late gene product ICP34.5 disrupts phosphorylation of IRF-3 by TBK1, whereas the US11 tegument protein inhibits RIG-I interaction with its downstream adaptor MAVS (23, 24). In addition, the immediate-early ICP0 protein has long been known to inhibit IRF-3 signaling in human fibroblasts; however, the mechanism of its inhibition has not been determined (25). Previously, we have shown that ICP0 causes the relocalization of Sendai virus (SeV)-activated IRF-3 to nuclear foci (26). This observation led us to hypothesize that ICP0 sequesters IRF-3 from cellular promoters to mediate inhibition of signaling. However, because SeV signals through RIG-I, it has remained unclear whether HSV-1 activates IRF-3 in a similar manner.

In this study, we investigated the initial activation of IRF-3 signaling in primary human fibroblasts in response to HSV-1. We observed that type I IFN expression in response to replication-defective HSV-1 requires the accumulation of viral DNA in the nuclear compartment. Furthermore, we identified IFI16 as a nuclear sensor of HSV-1 infection. In addition, we describe an immune-evasion strategy used by ICP0, which inhibits IRF-3 signaling by promoting the degradation of IFI16 during infection.

Results

Induction and Inhibition of IRF-3 Signaling in Human Fibroblasts.

To investigate the mechanisms by which HSV infection induces IFN-β expression and how virus-encoded ICP0 inhibits that process, we defined a system to investigate IRF-3 signaling in human foreskin fibroblasts (HFF) infected with the HSV-1 replication-defective d109 and d106 viruses. The d109 virus contains mutations in the five immediate-early genes ICP0, ICP4, ICP22, ICP27, and ICP47, effectively blocking all viral gene expression during infection (27). In contrast, the d106 virus expresses ICP0 but no additional immediate-early gene products. Consistent with previous results in human embryonic lung cells (25), we found that HFF infected with d109 showed a multiplicity of infection (MOI)-dependent induction of the IRF-3–responsive gene ISG54, whereas d106 virus infection led to much lower levels of expression of this gene (Fig. 1A). Because HSV-1 mutants that lack ICP0 have been shown to have differing particle-to-pfu ratios (28), we also determined whether cells were exposed to the same amount of viral DNA during equal-MOI infections with d109 and d106. This question was particularly important because DNA-sensing pathways play a major role in both detecting HSV-1 and inducing IRF-3 signaling during infection (17, 29). To test this possibility, we used real-time PCR to quantify the amount of viral DNA associated with cells during infection. Interestingly, infection with d109 led to fivefold more viral DNA being associated with cells at 2 h postinfection (hpi) (Fig. 1B). When the amount of infectious virus was adjusted, we observed similar amounts of viral DNA in cells infected with the d109 and d106 viruses (Fig. 1C). These results suggested that d109 has a higher particle-to-pfu ratio than d106; therefore, further experiments were conducted with both equal-pfu and equal-genome infections.

Fig. 1.

Establishment of an HSV-1 infection system to study IRF-3 signaling. (A) Induction of ISG54. HFF were infected with increasing MOIs of d106 or d109 virus, and total cellular RNA was harvested at 6 hpi. ISG54 mRNA levels were normalized to γ-actin levels and further normalized to mock-infected samples. Values are shown as mean ± SEM. (B) Viral genome numbers in infected cells. RAW264.7 cells were infected with d106 or d109 at an MOI of 1. (C) Total cellular DNA was harvested at 2 hpi, and relative viral DNA levels were analyzed by quantitative PCR using primers specific for the ICP8 gene. Levels were normalized to cellular GAPDH gene levels. Shown are ICP8 gene levels from RAW264.7 cells (Left) or HFF (Right) infected with d106 at an MOI of 5 and 50 or with d109 at an MOI of 1 and 10. Values are shown as mean ± SEM (n = 3).

To examine the kinetics of IRF-3–responsive gene expression in response to HSV-1 infection, we infected cells with d109 (MOI 10) or d106 (MOI 10 and 50) and harvested total RNA at 2, 4, and 6 hpi. The relative levels of expression of IFNβ, ISG56, and ISG54 transcripts were measured by RT-PCR and normalized to γ-actin RNA levels. By 4 hpi, d109 infection induced measurable amounts of the responsive genes (Fig. 2 A–C). In contrast, we observed lower levels of expression of these genes in d106 virus-infected cells, confirming that ICP0 was sufficient to inhibit IRF-3 signaling. As a control, ICP0 expression had no effect on levels of expression of the NF-κB–dependent IL-6 gene (Fig. 2D). Similar results for the IRF-3–responsive genes were seen in infected RAW246.7 macrophages (Fig. S1). These results argued that HSV-1 infection induces IRF-3 signaling early during infection and that expression of ICP0 rapidly counteracts this pathway. Therefore, these results defined a system to study the mechanism of activation and inhibition of the IRF-3 pathway.

Fig. 2.

HSV-1 induces IRF3-responsive genes in the absence of viral gene expression. RNA was harvested from HFF infected with d106 (MOI 10 or 50) or d109 (MOI 10) at 2, 4, and 6 hpi. RNA levels for IFNβ (A), ISG54 (B), ISG56 (C), and IL-6 (D) were determined by quantitative RT-PCR. Cellular RNA levels were normalized to γ-actin levels and further normalized to control 6-hpi values. Values are shown as mean ± SEM (n = 4). *P ≤ 0.05, compared with d106-infected cells (Student t test).

Conditions That Block Release of HSV DNA from Capsids Reduce IFN-β and ISG Induction.

The ability of d109 virus to induce IFN-β indicated that HSV-1 infection activates IFN-β before immediate-early gene expression. Therefore, to define the mechanism of induction of IFN-β, we assessed how d109 activates IRF-3 signaling in HFF. Upon entry, HSV-1 capsids rapidly translocate to the nuclear periphery where they dock at nuclear pores and subsequently release their viral DNA into the nucleus. To investigate whether accumulation of viral DNA in the nucleus is required for IRF-3–responsive gene expression, we examined IFN-β expression during infection in the presence of tosyl phenylalanyl chloromethyl ketone (TPCK). This serine/cysteine protease inhibitor inhibits cleavage of VP1-2 during infection and blocks viral DNA release into the nucleus (30). HFF treated with TPCK or control medium were infected with d109 or were treated with the TLR-3 antagonist polyinosinic:polycytidylic acid (poly I:C). Total RNA was harvested at 6 hpi and was analyzed by RT-PCR. TPCK treatment virtually eliminated the expression of IFNβ and ISG54 in response to d109 infection as compared with control cells (Fig. 3 A and B) but had a less dramatic effect on the induction of ISG54 in response to the TLR-3 antagonist poly I:C (Fig. S2A). These results argued for a specific effect of TPCK on HSV-induced signaling as opposed to poly I:C signaling. Although cell viability was not affected by TPCK treatment at the concentration used (5 μM) (Fig. S2B), TPCK could have additional effects on cellular processes, including the chymotrypsin-like activity of the proteasome. Therefore, we also treated cells with MG132 to determine the contribution of the proteasome in sensing d109 infection. Interestingly, MG132 treatment had no effect on ISG54 expression, although it did reduce IFNβ somewhat (Fig. 3 A and B). This differential inhibition by MG132 may be explained by differences in transcription factor recruitment to the IFNβ and ISG54 promoters. Although recruitment of IRF-3 is sufficient to activate the ISG54 promoter, expression of IFNβ requires additional transcription factors, including NF-κB, whose activation is dependent on proteasomal degradation of the IκB inhibitor (31). Consistent with TPCK but not MG132 blocking IRF-3–responsive gene expression, TPCK reduced TBK1 phosphorylation at Ser-172 in d109-infected cells, whereas MG132 showed no effect (Fig. 3C). Phosphorylation of this serine is essential for TBK1 kinase activity and thus for the activation of IRF-3 (32). In total, these results argued that the IRF-3 signaling response to HSV-1 infection requires the release of capsid-associated viral DNA into the nucleus and does not require proteasome activity.

Fig. 3.

Induction of IRF-3–responsive genes requires release of viral DNA from incoming capsids. HFF were pretreated with DMSO, TPCK, or MG132 for 30 min before and throughout infection with d109 at an MOI of 10. (A and B) Cellular RNA was harvested at 6 hpi, and IFNβ (A) and ISG54 (B) levels were determined by quantitative RT-PCR. RNA levels were normalized to 18S rRNA followed by normalization to corresponding mock-treatment values. *P ≤ 0.05, compared with DMSO-treated cells (Student t test). Values are shown as mean ± SEM (n = 3). (C) Western blot analysis of phospho-TBK1 (pTBK1) and tubulin levels in mock- or d109-infected cells not treated or treated with drugs.

IFI16 Is Necessary for HSV-1–Induced IFN-β Expression in HFF.

The IFI16 DNA sensor and its mouse ortholog p204 have been shown to be necessary for IRF-3–responsive gene expression in response to HSV-1 infection in THP-1 monocytes, RAW264.7 macrophages, and corneal epithelial cells (17, 29). To test the importance of IFI16 in HSV-1 activation of IRF-3 signaling in HFF, we examined IFN-β expression during infection in the absence of IFI16. HFF were transfected with control siRNA (siControl) or siRNA targeting IFI16 (siIFI16) for 3 d, followed by infection with d109 or SeV for 6 h. Western blot analysis of the resulting protein lysates showed a greater than 90% knockdown of IFI16 in siIFI16-treated cells compared with cells treated with siControl (Fig. 4A). When mRNA expression was examined in d109-infected cells by RT-PCR, there was a marked reduction in IFN-β expression in siIFI16-treated cells compared with control cells (Fig. 4C). In contrast, knockdown of IFI16 did not have a significant effect on SeV-induced IFN-β expression (Fig. 4D), consistent with previous findings (17). Furthermore, phosphorylation of TBK1 at Ser-172 was diminished in d109-infected IFI16-knockdown cells relative to cells receiving siControl (Fig. 4A). Together these results were consistent with IFI16 being necessary to activate the IRF-3 signaling cascade during HSV-1 infection.

Fig. 4.

IFI16 and STING are required for HSV-1–induced IFN-β induction. HFF were treated with siControl, siIFI16, or siRNA targeting STING (siSTING) for 3 d following infection with d109 (MOI 10) or SeV (100 HAU/8 × 10^5 cells). Cells were harvested for RNA and protein analysis at 6 hpi. (A and B) Western blot analysis of (A) IFI16 and pTBK1 and (B) STING and GAPDH levels in siRNA-treated cells. RNA samples from d109 (C) or SeV (D) infected cells were analyzed by quantitative RT-PCR. IFNβ RNA levels were normalized to 18S rRNA followed by normalization to corresponding mock values. Values are shown as mean ± SEM (n = 3).

STING, a cytoplasmic scaffolding protein, is essential for the IFN-β response to immunostimulatory DNA molecules and HSV-1 infection in mouse embryonic fibroblasts (33). In addition, an immunostimulatory VACV 70mer has been shown to induce an association between STING and IFI16 in THP-1 cells by coimmunoprecipitation (17). To determine whether STING is required for the HSV-1–mediated IFN-β response in HFF, we examined IFN-β mRNA expression in HFF treated with STING siRNA. At 6 hpi we observed a marked reduction in IFN-β expression in the absence of STING in d109-infected cells (Fig. 4C). Treatment with STING siRNA had no significant effect on SeV-induced IFN-β expression (Fig. 3D). Interestingly, although STING protein expression was reduced by only ∼60% upon treatment with STING siRNA (Fig. 4B), its loss had a greater impact on IFN-β expression than IFI16 knockdown in response to d109 infection. In addition, we also observed a decrease in STING protein levels in SeV-infected siControl- and siSTING-treated HFF, consistent with the previously observed RNF5-mediated degradation of STING in response to SeV infection (34). Together, these results argued that STING is important for the induction of IFN-β in response to HSV-1 infection and may play a role downstream of IFI16 in HFF.

IFI16 Is Nuclear in Uninfected and d109-Infected HFF.

The subcellular localization of IFI16 can be either nuclear or cytoplasmic, depending on the cell type (reviewed in ref. 18), and it has been reported to be nuclear in HFF (35). We confirmed that IFI16 is nuclear in HFF by immunofluorescence (Fig. 5A). Nuclear export of IFI16 has been reported during infection of endothelial cells with Kaposi sarcoma-associated herpesvirus (KSHV) and was associated with an induction of inflammasome and NF-κB signaling (36). Interestingly, when we examined IFI16 localization in HFF during infection with d109, we did not observe a change in the localization of IFI16; i.e., IFI16 was nuclear in mock- and d109-infected cells (Fig. 5A). Similar results were obtained using the 1G7 monoclonal antibody (Santa Cruz), which showed nuclear export of IFI16 in a previous study (36). To confirm this phenotype, we examined IFI16 localization by biochemical fractionation. Nuclear and cytoplasmic fractionation was performed on HFF infected with d106 or d109 viruses at 4 hpi. Consistent with our immunofluorescence results, IFI16 was completely nuclear in uninfected and d109-infected cells (Fig. 5B).

Fig. 5.

IFI16 is localized in the nucleus during HSV-1 infection. (A) Fibroblasts infected with d109 virus were fixed and stained with an antibody specific for IFI16 (shown in green) at 4 hpi. Right panel is a higher magnification of a representative cell in the left panel. (Scale bar, 5 μm.) (B) Nuclear and cytoplasmic fractions were prepared from cells infected with d109 and d106 and were analyzed by immunoblot for ICP0, IFI16, and STING localization. Tubulin and lamin A/C represent the fractionation efficiency. (C) Cells were treated with leptomycin B for 30 min before and throughout infection with d109 or SeV for 8 h. Flow cytometry then was used to examine the phosphorylation status of TBK1. Values are shown as mean ± SEM (n = 4).

STING was localized in the cytoplasm of uninfected cells, as shown by fractionation, and it showed no change in localization following d109 infection (Fig. 5B). Although in this experiment cells infected with a higher MOI of d106 showed a slight increase in nuclear-associated STING (Fig. 5B), this increase was not observed in other experiments. The absence of IFI16 in the cytoplasm of d109-infected HFF suggested that IFI16 potentiates signaling from the nucleus to the cytoplasm without major changes in its distribution in response to HSV-1. To test whether nuclear export is required for activation of IRF-3 signaling during HSV-1 infection, we examined the phosphorylation status of TBK1 by flow cytometry when leptomycin B was used to inhibit the nuclear exportation of proteins by the CRM1 pathway. HFF were treated with leptomycin B for 30 min before infection with HSV-1 d109 or, to stimulate through another sensor infection with SeV. Infected cells were fixed and stained at 8 hpi with a specific antibody raised against phospho-TBK1 ser172. Treatment of HFF with leptomycin B decreased the percentage of d109-infected cells that stained positive for TBK1 phosphorylation by ∼70% (Fig. 5C). In contrast, phospo-TBK1 staining was decreased only slightly in leptomycin B -treated, SeV-infected cells compared with control-treated cells. These results suggested that signaling in response to HSV-1 infection requires CRM1-mediated export of a factor(s).

Nuclear ICP0 Inhibits IRF-3 Upstream and Downstream of Activation.

The induction of IFN-β in response to HSV-1 infection is dependent on the IRF-3 transcription factor (37, 38). The viral ICP0 protein has been implicated in inhibiting the IRF-3 pathway; however, the mechanism of inhibition has remained unclear (25). To determine the stage at which IRF-3 activation was inhibited by ICP0, we initially examined IRF-3 localization during HSV-1 infection by nuclear/cytoplasmic fractionation. Nuclear and cytoplasmic fractions were prepared from HFF infected with d106 or d109 at 4, 6, and 8 hpi. The efficiency of fractionation was confirmed by the complete localization of GAPDH in the cytoplasmic fraction and lamin A/C in the nuclear fraction (Fig. 6). At 4 hpi, equal amounts of activated IRF-3 (pIRF-3 S396) were seen in the nuclei of cells infected with equal-genome amounts of d109 (MOI = 10) and d106 (MOI = 50) (Fig. 6, lanes 14 and 16). Cells infected with an equal MOI of d106 (MOI = 10) showed delayed IRF-3 accumulation, most likely because of decreased viral DNA presence during infection (Fig. 6, lane 15). Interestingly, by 6 hpi pIRF-3 S396 had continued to accumulate in the nuclei of cells infected with d109 but was at very reduced levels in cells expressing ICP0 (d106). Additional experiments in RAW264.7 cells revealed a similar difference in d109- and d106-infected cells at 2 and 4 hpi (Fig. S3). Because differences in ISG54 expression were observed by 4 hpi (Fig. 2B), the results taken together suggest that ICP0 is capable of blocking IRF-3 activity at early times in the nucleus as well as upstream of IRF-3 activation at later stages of infection.

Fig. 6.

ICP0 inhibits nuclear accumulation of activated IRF-3. HFF were mock-infected or infected with the d109 or d106 viruses. Nuclear and cytoplasmic fractions were prepared at 4, 6, and 8 hpi. Fractions were probed using antibodies specific for ICP0, IRF-3, and phospho-IRF-3 (Ser396). Fractionation efficiency was determined by localization of GAPDH (cytoplasm) and lamin A/C (nucleus).

Previously, we had shown that ICP0 expression causes the relocalization of SeV-activated IRF-3 in the nuclei of infected cells (26). We therefore hypothesized that ICP0 sequesters IRF-3 from its normal nuclear activity. To determine whether this sequestration model applied to the situation at 4 hpi in infected HFF, we examined IRF-3 localization by indirect immunofluorescence. At 6 hpi, we fixed d109- and d106-infected HFF and stained them with antibodies specific for IRF-3 and ICP0. The IRF-3 antibody used in this experiment (SL-12) preferentially binds to activated IRF-3 (22); therefore, we detected diffusely nuclear IRF-3 in d109-infected cells (Fig. 7E). In contrast, cells infected with d106 showed a relocalization of IRF-3 to ICP0 foci (Fig. 7 I and L). These results argued that nuclear ICP0 inhibits IFN-β expression early during infection by sequestering IRF-3 in the nucleus of infected HFF.

Fig. 7.

ICP0 sequesters nuclear IRF-3. HFF were infected with d109 (MOI 10) or d106 viruses (MOI 10 or 50) for 6 hpi. Samples were fixed and stained using antibodies specific for ICP0 (shown in green) and IRF-3 (shown in red). (Scale bar, 5 μm.)

Effects of ICP0 Expression on IFI16.

In WT HSV-1 infection, ICP0 is localized to the nucleus initially but accumulates in the cytoplasm at late time points postinfection (39). This change in localization requires viral DNA replication and late gene expression (40). Our nuclear fractionation and immunofluorescence studies suggested that ICP0 expressed by d106 is mainly, if not entirely, nuclear (Figs. 6 and 7), consistent with the replication-incompetent nature of the d106 virus. Given the nuclear localization of ICP0 and its apparent effect upstream of IRF-3 activation, we tested whether ICP0 expression affects nuclear IFI16. We infected HFF with d106 or d109, prepared protein lysates at 6 hpi, and performed Western blot analysis. Notably, cells infected with d106 at both an MOI of 10 and 50 (lanes 3 and 4) showed a marked reduction in the steady-state levels of IFI16 compared with d109 virus- (lane 2) or mock-infected (lane 1) cells (Fig. 8A). Interestingly, d106 infection in RAW264.7 cells also promoted the loss of p204, the murine IFI16 ortholog (Fig. S3, lanes 15 and 16). In addition, we examined IFI16 localization in d106- and d109-infected cells by immunofluorescence at 2, 4, and 6 hpi. At 2 hpi, IFI16 partially localized with nuclear ICP0 foci and subsequently was lost from the cell by 4 and 6 hpi (Fig. 8B). Together these results suggested that ICP0 expression during HSV-1 infection promotes the relocalization and subsequent degradation of IFI16.

Fig. 8.

ICP0 expression promotes IFI16 relocalization and degradation. HFF cells were infected with d109, d106, or WT KOS strain virus. (A) Whole-cell lysates were harvested at 6 hpi and probed for ICP0, IFI16, IRF-3, phospho-IRF-3 (Ser396), STING, and GAPDH using specific antibodies. (B) Infected cells were fixed and stained at 2, 4, and 6 hpi for IFI16 (shown in green) and ICP0 (shown in red). MOIs for d106 are stated in parentheses.

HSV-1 Promotes Degradation of IFI16 via an ICP0 RING Finger-Dependent Mechanism.

Previous studies have shown that ICP0 promotes the degradation of cellular proteins in a proteasome-dependent manner through the E3 ligase activity of its RING finger domain (41). This domain coordinates the ubiquitination of specific target proteins and defines ICP0 as an E3 ubiquitin ligase. To test the involvement of the ubiquitin–proteasome pathway in the loss of IFI16, we examined IFI16 protein levels in the presence of the proteasome inhibitor MG132. We treated HFF with MG132 or DMSO for 30 min before infection with WT HSV-1 KOS strain, and drug treatment was continued throughout infection. At 8 hpi we harvested cell lysates and probed for IFI16 expression. Treatment with MG132 inhibited the loss of IFI16 observed during infection (Fig. 9A). To test whether loss of IFI16 in our system was dependent on RING finger domain activity, we constructed an ICP0 RING finger mutant virus by mutational alteration of cysteine residues 116 and 156 to alanine and glycine, respectively. These mutations abolish ICP0 ligase activity by disrupting zinc coordination (42, 43). The RING finger mutant virus, KOS.RFm, was incapable of degrading USP7 (44) compared with the rescued virus, KOS.RFr, but KOS.RFm ICP0 localized normally in the nucleus of infected cells. We harvested cell lysates from KOS.RFm- or KOS.RFr-infected cells at 8 hpi and probed for ICP0 expression and IFI16 levels. We found that although KOS.RFm and KOS.RFr expressed similar levels of ICP0, KOS.RFm was unable to promote the degradation of IFI16 (Fig. 9B). Based on these results, we concluded that ICP0 promotes the degradation of IFI16 in a proteasome- and RING finger domain-dependent manner.

Fig. 9.

ICP0 promotes the degradation of IFI16 in a proteasome- and RING finger-dependent manner. (A) HFF were pretreated with DMSO (control) or MG132 for 30 min before and throughout infection with WT HSV (KOS) virus at an MOI of 10. Whole-cell lysates were harvested at 8 hpi. Western blot analysis of ICP0, IFI16, and tubulin is shown for each of these samples. (B) Cells were infected with the ICP0 RING-finger mutant virus (KOS.RFm) and its rescue (KOS.RFr). Whole-cell lysates were harvested and analyzed as in A.

Discussion

Sensing of microbial macromolecules by innate immune mechanisms has been demonstrated to take place at the cell plasma membrane, in internal vesicles, and in the cytoplasm, but there has been little clear evidence that innate sensing takes place in the cell nucleus. Sensors of viral or bacterial DNA have been considered to be cytosolic in origin (45). In this study, we show that HSV-1 DNA must be delivered to the cell nucleus for sensing by the nuclear IFI16 sensor and that, although IFI16 appears to remain in the nucleus after viral DNA recognition, signaling takes place by exportation of an unidentified molecule to activate IRF-3 through STING in the cytoplasm. The viral ICP0 protein in its nuclear form can relocalize IFI16 within the nucleus and promote its degradation, thereby blocking further signaling. The observations that viral DNA is delivered to the nucleus, that the sensor is nuclear, and that an inhibitor is nuclear strongly support the hypothesis that sensing of HSV-1 DNA occurs in the nucleus in HFF.

Nuclear Sensing of HSV-1 DNA.

IFI16 has been defined as being involved in IFN-β and CXCL10 induction in HSV-infected THP-1 cells (17) and epithelial cells (29), respectively. Originally described as a cytosolic DNA sensor (17), IFI16 recently was implicated in sensing KSHV in the nucleus of infected cells (36). The latter study found that IFI16 colocalized with nuclear KSHV genomes and subsequently was translocated to the cytoplasm where it induced inflammasome and NF-κB signaling. We found that IFI16, which is nuclear in HFF, is involved in the induction of an IFN response to HSV-1 in this cell line, because knockdown of IFI16 decreased IFN-β expression during infection with the d109 virus. Endosomally localized TLR9 was the first sensor identified as recognizing HSV-1 DNA (46); however, its expression is restricted to plasmacytoid dendritic cells and B cells in humans, and thus it is unlikely to be a sensor of HSV-1 in HFF. Indeed, Rasmussen et al. (47) previously reported that conventional dendritic cells, macrophages, and mouse embryonic fibroblasts infected with HSV-1 could secrete IFN-β in a TLR9-independent but viral DNA-dependent manner. Additional cytosolic sensors have been identified that recognize HSV-1 DNA; however, it has remained unclear how this viral DNA gains access to the cytosolic compartment during infection, given that the capsid likely protects viral DNA from cytosolic sensors during transport through the cytoplasm. Some have proposed that viral DNA may be made available to the cytosolic compartment by the degradation of viral capsid (48). Our experiments revealed that chemical inhibition of the proteasome in HFF had no effect on ISG54 expression in response to HSV-1. Instead, blocking the release of viral DNA into the nucleus of infected cells by TPCK treatment greatly diminished the cellular response to HSV-1 infection (30). These results are consistent with a need to deliver the viral DNA to the nucleus for IFN-β induction in HFF. Other cell types, however, may have means for freeing viral DNA in the cytoplasm or endosomes so that other receptors can initiate innate responses.

The sensing of viral DNA in the nucleus raises the question of how IFI16 distinguishes between nuclear viral and cellular DNA. The issue of specificity was one of the reasons that the compartmentalized cytosolic and endosomal sensing of DNA was attractive. IFI16 is known to bind to both ss- and dsDNA in vitro (17); thus, it should be able to bind to cellular or viral DNA. IFI16 may bind preferentially to the under-chromatinized HSV DNA, DNA ends, or nicks and gaps in HSV DNA. HSV DNA is known to be chromatinized rapidly upon entry into the cell nucleus (49, 50), although the histone association with HSV DNA is less densely packed (49) and looser (51) than cellular chromatin. The altered chromatin structure may allow IFI16 binding and activation of the signaling pathway. IFI16 has been implicated in the DNA damage response by its interactions with BRCA1 and p53 (52, 53). HSV infection is known to activate DNA damage response pathways (54–56); therefore, the free ends of HSV DNA or nicks and gaps could provide DNA-binding sites for IFI16 to activate the IRF-3 signaling pathway and potentially other signaling cascades also. Currently there are conflicting reports as to whether DNA damage itself induces IFN-β expression. A recent study in primary human monocytes suggested that IFN-α and -γ, but not -β, were induced in response to the DNA damaging agent etoposide (57). However, additional reports have shown that treatment of young human diploid fibroblasts with bleomycin increased IFI16 expression (58), and irradiation of bone marrow-derived mouse macrophages increased ISG expression in an IFN-β–dependent manner (59). Interestingly, no detectable double-stranded break response has been detected in UV-inactivated HSV-1-infected cells (55), which presumably would mimic d109 infection in our system. However, current methods of detecting double-stranded breaks may be insufficient to detect an initial DNA damage response from nonreplicating viral DNA. Therefore, further studies are needed to understand the relationship between IFI16 sensing of viral DNA and the DNA damage response.

As we were submitting this manuscript, Li et al. (60) reported results showing that IFI16 must be nuclear to sense HSV-1 infection. In this study, the authors showed that a HEK293 cell line expressing IFI16 that lacked a functional nuclear localization signal was unable to induce IFN-β expression in response to HSV-1 compared with WT IFI16-expressing cells. Our results are consistent with and complementary to their results and expand on the mechanisms of activation and inhibition of IFI16 sensing of HSV-1 infection.

Role for Nuclear IFI16 in HSV-1–Induced IFN Expression.

It is currently unclear how nuclear IFI16 induces the cytoplasmic IRF-3 signaling cascade in HFF. Previously, cytosolic IFI16 was shown to associate with STING upon activation with immunostimulatory DNA (17). In our system, STING knockdown reduced the expression of IFN in response to HSV-1 infection, suggesting the involvement of this protein in the activation of IRF-3 signaling in HFF. However, we do not observe a measurable relocalization of IFI16 from the nucleus to the cytoplasm during infection. Additional experiments using the CRM1 inhibitor leptomycin B revealed that a nuclear export event is required for the autophosphorylation of the TBK1 kinase, suggesting that additional factors may translocate to the cytoplasm to initiate signaling upon IFI16 activation. The factor(s) that link nuclear IFI16 to cytosolic STING are unknown currently and represent a high priority for future studies.

Inhibition of IRF-3 Signaling by Nuclear ICP0.

In this study we found that HSV ICP0 can inhibit the IRF-3 pathway through degradation of IFI16. HSV-1 infection has long been known to antagonize the IRF-3 signaling pathway, and the expression of ICP0 plays a major role in inhibiting this cellular response (22, 25, 26). Our previous studies have shown that ICP0 can block RIG-I–induced IRF-3 signaling in SeV-infected cells by sequestering nuclear IRF-3 and reducing its levels (22, 26). In this study we have examined the mechanisms of ICP0 inhibition of IRF-3 activation in HSV-infected cells. In HFF, we observed that ICP0 inhibits the IRF-3 signaling pathway at two distinct steps. Nuclear/cytoplasmic fractionation and analysis of IFN-β and ISG expression revealed that ICP0 initially inhibits type I IFN expression at a stage after nuclear accumulation of phosphorylated IRF-3. This early inhibition is associated with a relocalization of IRF-3 to ICP0 nuclear foci and is consistent with the sequestration of IRF-3 in the SeV coinfection system. At later times after infection ICP0 inhibits the accumulation of IRF-3 in the nucleus coincident with the loss of IFI16.

ICP0 is an E3 ubiquitin ligase and promotes the degradation of cellular proteins to enhance virus replication and inhibit host innate responses (41, 61). Others have observed that the ability of ICP0 to inhibit IRF-3 signaling is dependent, at least in part, on functional proteasomes (25, 62); however, no cellular target for degradation has been identified. In addition to sequestering IRF-3 in the nucleus, we observed that ICP0 promotes the degradation of the IFI16 DNA sensor in a proteasome- and RING finger-dependent manner. ICP0 is associated with the degradation of a number of cellular proteins, but only a limited number of these have been shown to interact directly with ICP0 and/or be directly ubiquitinated in a reaction involving ICP0 (reviewed in ref. 63). Therefore, it will be important to determine the mechanism by which ICP0 promotes degradation of IFI16. Most importantly, this provides a mechanism by which a virus can inhibit nuclear IFI16 from activating IRF-3 signaling.

ICP0 may also inhibit IRF-3 signaling in the cytoplasm. At early times postinfection ICP0 is localized to the nucleus, whereas upon viral DNA replication the protein accumulates in the cytoplasm. A recent study showed that an ICP0 mutant protein without a nuclear localization signal could inhibit signaling in human embryonic lung fibroblasts (62). It is conceivable that ICP0 can affect IRF-3 activation in the cytoplasm when it accumulates there. Our results do not rule out additional effects of ICP0 in the cytoplasm of infected cells; however, our results argue strongly for the ability of nuclearly localized ICP0 to inhibit IRF-3 signaling early during infection.

Based on the studies described here, we propose the following model for the activation and ICP0-mediated inhibition of type I IFN expression during HSV-1 infection (Fig. 10). In HFF, HSV-1 infection is sensed initially by IFI16 upon the release of viral DNA into the nucleus. A nuclear-to-cytoplasmic signaling cascade is initiated that activates IRF-3 and induces its accumulation in the nucleus. ICP0 expressed at early times during infection sequesters this nuclear IRF-3 from cellular promoters and blocks type I IFN expression. In addition, ICP0 targets IFI16 for degradation, inhibiting additional signaling and activation of IRF-3. Determination of the mechanism of specific detection of viral DNA in the nucleus within the same cellular compartment as cellular DNA likely will shed light on basic cellular mechanisms for detecting “foreign” or altered DNA within the cell. HSV DNA activates DNA damage response pathways, and IFI16 has been implicated in DNA damage response; thus, these results raise the possibility that the IFN response to viral DNA and the DNA damage response pathways share sensing or signaling components.

Fig. 10.

Model of nuclear HSV-1 DNA sensing and inhibition by ICP0. HSV-1 fusion at the plasma membrane or via endosomal compartments deposits viral capsids in the cytoplasm. Capsids traffic to nuclear pores where viral DNA is released into the nucleus. Nuclear IFI16 senses accumulating viral DNA, inducing a nuclear-to-cytoplasmic signaling cascade activating IRF-3, which dimerizes and translocates to the nucleus. Immediate-early expression of ICP0 sequesters IRF-3 from cellular promoters and promotes degradation of IFI16 to inhibit IFN-β expression.

Materials and Methods

Cell Culture and Viruses.

HFF were grown in DMEM supplemented with 15% (vol/vol) fetal bovine serum (FBS). RAW246.7 macrophages, U2OS, Vero, and FO6 cells were maintained in DMEM supplemented with 5% (vol/vol) FBS, 5% (vol/vol) bovine calf serum (BCS), and glutamine with appropriate selection media as needed.

The WT HSV-1 KOS strain virus was propagated and titered by plaque assay on Vero cells (64). The d106 and d109 viruses were propagated on E11 and FO6 cells, respectively, and were titered in parallel on FO6 cells (27). The ICP0-RING finger mutant virus (KOS.RFm) and its corresponding rescued virus (KOS.RFr) were constructed by homologous recombination into the ICP0-null 7134 virus (65). Infectious viral DNA was cotransfected with linearized plasmid containing ICP0-flanking regions and mutations within the ICP0 protein (C116G/C156A). Cells were harvested at 3 d posttransfection and were freeze-thawed, and dilutions were plated on U2OS cells to isolate plaques. An agarose overlay containing X-Gal was used to distinguish recombinant plaques that did not express β-galactosidase. White plaques underwent three rounds of purification before viral DNA was harvested. The ICP0 RING finger domain was amplified, and mutations were confirmed by sequencing and restriction endonuclease digestion. KOS.RFm infectious DNA was used to construct the KOS.RFr virus in a similar manner. Virus stocks were grown and titered on U2OS cells.

Infections.

Virus was diluted in PBS containing 0.1% (wt/vol) glucose and 1% (vol/vol) heat-inactivated BCS. Cells were infected at the stated MOI for 1 h at 37 °C, washed twice with PBS, and overlaid with DMEM containing 1% (vol/vol) heat-inactivated BCS. Infected cells were incubated at 37 °C for the indicated length of time.

Drugs.

Cells were treated with DMEM containing 5 ng/mL leptomycin B (Sigma-Aldrich) for 30 min before infection. For proteasome-inhibition studies, cells were treated with 1 μM MG132 (Sigma) or 0.01% DMSO. TPCK was used at a concentration of 5 μg/mL. Drugs were included throughout the adsorption period as well as in overlay medium.

Viral DNA Analysis by Quantitative PCR.

At 2 hpi total cellular DNA was harvested using a Qiagen Generation Capture Column Kit. DNA levels of specific sequences were determined by quantitative real-time PCR using the Power SYBR Green PCR master mix and a Prism 7300 sequence detection system (Applied Biosystems). PCR reactions were carried out in duplicate, and relative copy numbers were determined by comparison with standard curves. Viral DNA was normalized to cellular γ-actin (human) or GAPDH (mouse) levels. A list of primer sequences used can be found in SI Materials and Methods.

Cellular RNA Analysis by Quantitative PCR.

Total RNA was extracted using the Qiagen RNeasy Kit and was DNase treated using the DNA-free kit (Ambion). DNase-treated RNA then was reverse-transcribed and quantified by real-time PCR as above. Mock reverse-transcribed samples were included as negative controls. Experiments were conducted three times, and the values were averaged. Samples were normalized to either γ-actin mRNA or 18S rRNA. The Student t test was used to determine the statistical significance of differences between samples. A list of primer sequences used can be found in SI Materials and Methods.

Nuclear Cytoplasmic Fractionation.

Nuclear and cytoplasmic extracts were prepared with the NE-PER Nuclear and Cytoplasmic Extraction Kit (Thermo Scientific). The purity of nuclear and cytoplasmic extracts was assessed by immunoblotting with anti-lamin A/C (cell signaling) and either anti-tubulin or anti-GAPDH antibodies (Abcam), respectively.

Western Blots.

Cells were lysed in NuPAGE LDS Sample Buffer, and proteins were resolved on NuPAGE 4–12% Bis Tris Gels (Invitrogen). Proteins were transferred to PVDF membranes, and Western blots were developed using Luminate Forte Western HRP substrate (Millipore). A list of antibodies used and their dilutions can be found in SI Materials and Methods.

Indirect Immunofluorescence.

HSV-1–infected HFF grown on coverslips were fixed with 2% (vol/vol) formaldehyde, permeabilized with 0.5% Nonidet P-40, and blocked in 5% (vol/vol) normal goat serum. Fixed cells were incubated with antibodies for 30 min at 37 °C and washed two times with PBS containing 0.05% Tween 20 followed by one washing with PBS. Alexa Fluor 488- and 594-conjugated secondary antibodies were incubated with cells for 2 h at 25 °C. The coverslips were washed as above and mounted in ProLong Gold antifade reagent (Invitrogen). Images were acquired using an Axioplan 2 microscope (Zeiss) with a 63× objective and Hamamatsu CCD camera (model C4742-95). Images were arranged in figures using Adobe Photoshop CS4 (Adobe Systems). A list of antibodies used and their dilutions can be found in SI Materials and Methods.

siRNA Transfections.

Double-stranded IFI16-specific, STING-specific, and nontarget control siRNAs were purchased from Dharmacon. The pooled siRNA were transfected into HFF using the DarmaFECT 2 transfection reagent (Dharmacon) at a final siRNA concentration of 5 nM according to the manufacturer’s instructions. At 3 d posttransfection cells were assayed for IFI16 or STING levels by immunoblotting and/or were infected with HSV-1.

Flow Cytometry.

Infected HFF were trypsinized and washed once with ice-cold PBS containing 1 mM sodium orthovanadate. Cells were fixed for 20 min with 1.5% (vol/vol) formaldehyde and incubated in permeabilization buffer (0.2% saponin, 0.5% BSA, 1 mM sodium orthovanadate, 1 mM β-glycerophosphate, 50 mM sodium fluoride) for 15 min. Cells were incubated in permeabilization buffer with primary antibody for 1 h at 25 °C and then were washed and stained with secondary goat anti-rabbit FITC-conjugated antibody (Jackson ImmunoResearch Laboratories) for 1 h in permeabilization buffer and analyzed by flow cytometry. A list of antibodies used and their dilutions can be found in SI Materials and Methods.