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. Author manuscript; available in PMC: 2013 May 8.
Published in final edited form as: Mol Cell. 2012 Mar 7;46(1):79–90. doi: 10.1016/j.molcel.2012.02.004

Viral E3 ubiquitin ligase-mediated degradation of a cellular E3: viral mimicry of a cellular phosphorylation mark targets the RNF8 FHA domain

Mira S Chaurushiya 1,3, Caroline E Lilley 1, Aaron Aslanian 2,4, Jill Meisenhelder 2, Daniel C Scott 5, Sebastien Landry 1, Simina Ticau 1, Chris Boutell 6, John R Yates III 4, Brenda A Schulman 5, Tony Hunter 2, Matthew D Weitzman 1,7,*
PMCID: PMC3648639  NIHMSID: NIHMS366769  PMID: 22405594

Abstract

Viral hijacking of cellular processes relies on the ability to mimic the structure or function of cellular proteins. Many viruses encode ubiquitin ligases to facilitate infection, although the mechanisms by which they select their substrates are often unknown. The Herpes Simplex Virus type-1 encoded E3 ubiquitin ligase, ICP0, promotes infection through degradation of cellular proteins, including the DNA damage response E3 ligases RNF8 and RNF168. Here we describe a mechanism by which this viral E3 hijacks a cellular phosphorylation-based targeting strategy to degrade RNF8. By mimicking a cellular phosphosite, ICP0 binds RNF8 via the RNF8 forkhead associated (FHA) domain. Phosphorylation of ICP0 T67 by CK1 recruits RNF8 for degradation and thereby promotes viral transcription, replication, and progeny production. We demonstrate that this mechanism may constitute a broader viral strategy to target other cellular factors, highlighting the importance of this region of the ICP0 protein in countering intrinsic antiviral defenses.

Introduction

Cellular networks are coordinated by protein-protein interactions that employ a variety of structural domains to recognize specific short linear motifs, such as those created by phosphorylation events (Pawson and Scott, 1997; Ren et al., 2008). These interactions can be inducible and reversible, and play important roles in cellular signaling pathways. The DNA damage response (DDR) is a complex network of signals coordinated through post-translational modifications that facilitate interactions between structured domains and short linear motifs (Polo and Jackson, 2011). Induction of a double-stranded DNA break is recognized by the cellular Mre11-Rad50-Nbs1 complex (Carson et al., 2003; Falck et al., 2005; Lee and Paull, 2005; Uziel et al., 2003), which activates the ataxia-telangiectasia mutated (ATM) kinase to phosphorylate histone H2AX on the chromatin surrounding the break (Burma et al., 2001; Downs et al., 2000; Rogakou et al., 1998). The resulting phosphosite on γH2AX is bound by the BRCA C-terminal (BRCT) domains of the checkpoint mediator protein Mdc1 (Hammet et al., 2007; Lee et al., 2005; Stucki et al., 2005), which then serves as a primary platform for recruitment of downstream repair factors. Two E3 ubiquitin ligases, RNF8 and RNF168, function in the DDR to bridge these early phosphorylation signals with subsequent ubiquitin-based signaling events. The RNF8 protein contains a fork-head associated (FHA) domain that binds ATM-catalyzed phospho-TQXF motifs on Mdc1 (Huen et al., 2007; Kolas et al., 2007; Mailand et al., 2007). RNF8 then ubiquitinates proteins at the site of the break, including histones H2A and H2AX, and ubiquitin chains in turn serve as a recruiting platform for the ubiquitin-binding motifs of RNF168 (Doil et al., 2009; Stewart et al., 2009). These events facilitate downstream accumulation of mediator proteins such as 53BP1, and effector functions including DNA processing and repair, transcriptional silencing, and cell cycle modulation.

The DDR is intimately involved in the cellular response to virus infection, and is manipulated by many virally-encoded proteins (Weitzman et al., 2010). In the case of Herpes Simplex Virus type-1 (HSV-1), the virus-encoded E3 ubiquitin ligase, ICP0, targets RNF8 and RNF168 for degradation (Lilley et al., 2010). This degradation is important to counteract RNF8- and RNF168-mediated recognition and transcriptional silencing of the viral genome (Lilley et al., 2011). HSV-1 undergoes lytic and latent stages of infection, and ICP0 promotes both of these in a manner predominantly dependent on its catalytic RING domain (Cai et al., 1993; Everett, 2000; Halford et al., 2001; Halford and Schaffer, 2001; Leib et al., 1989). In addition to RNF8 and RNF168, ICP0 has been shown to mediate degradation of DNA-PKcs (Parkinson et al., 1999), ND10 components (Everett et al., 1998), and centromeric proteins (Everett et al., 1999; Lomonte and Morency, 2007; Lomonte et al., 2001). While it is clear that ICP0 renders the host cell environment amenable to viral infection through degradation of specific cellular factors, the mechanisms by which it targets these diverse proteins remain unclear.

Here we identify a short linear motif within ICP0 that is phosphorylated by the cellular CK1 kinase, mimicking the phosphosites induced on Mdc1 during propagation of DDR signals. This phosphosite, on ICP0 T67, directly binds the RNF8 FHA domain and is required for ICP0-mediated ubiquitination and degradation of RNF8. We demonstrate that ICP0 T67 is required for targeting RNF8 in the context of HSV-1 infection, and is important for countering cellular attempts to repress transcription from the viral genome. Finally, we show that ICP0 pT67 can bind other cellular proteins, suggesting that ICP0 pT67 binds additional cellular proteins to intersect with diverse cellular pathways.

Results

The FHA domain of RNF8 is required for targeting by ICP0

ICP0 localizes to discrete foci in the nucleus corresponding to ND10 domains (Everett and Maul, 1994; Maul and Everett, 1994). RNF8 is diffusely nuclear, but re-localizes to ICP0 foci upon ICP0 expression (Lilley et al., 2010). In order to determine the regions of RNF8 required for targeting by ICP0, we constructed Flag-tagged fragments of RNF8 comprising different functional domains (Figure 1A) and co-expressed them with eGFP-ICP0. We observed that only RNF8 fragments containing the FHA domain were recruited to ICP0 foci, indicating that the interaction is coordinated through the RNF8 FHA domain. Supporting these observations, introduction of an inactivating R42A point mutation into the FHA domain was sufficient to abolish RNF8 recruitment to ICP0 foci (Figures 1B, 1C, and S1A).

Figure 1. The RNF8 FHA domain is required for targeting by ICP0.

Figure 1

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(A) Functional regions of RNF8 and fragments generated. (B) Flag-RNF8 fragments and mutants were co-transfected into HeLa cells with eGFP-ICP0ΔRING and processed for immunofluorescence 24 h post-transfection. (C) Cells expressing ICP0 and Flag-RNF8 (50 cells for each condition imaged in panel B) were scored for co-localization. (D) Flag-RNF8 or the R42A mutant were transfected into HeLa cells and infected with WT HSV-1 (17syn+) 24 h post-infection. Cells were harvested 8 h post-infection (see also Supplemental Figure S1).

We next sought to determine whether the FHA domain of RNF8 was required for RNF8 degradation during HSV-1 infection. We found that WT Flag-RNF8 was degraded 8 h post-infection (hpi), but that the R42A mutant was resistant to degradation (Figure 1D). Furthermore, all Flag-RNF8 fragments lacking or mutated in the FHA domain were resistant to degradation during infection (Figures S1B and S1C). Supporting these observations, recombinant 6xHis-ICP0 protein ubiquitinated recombinant 6xHis-Flag-RNF8 R42A protein in vitro much less efficiently than WT His-Flag-RNF8 (Figure S1D). We thus concluded that ICP0 targets RNF8 for ubiquitination and degradation via interaction with the FHA domain of RNF8.

ICP0 phosphorylation is required for interaction with the RNF8 FHA domain

FHA domains are well-described phospho-Thr binding motifs (Durocher and Jackson, 2002), and ICP0 is heavily phosphorylated (Davido et al., 2005). To address whether the RNF8-ICP0 interaction was phosphorylation-dependent, recombinant GST-RNF8-FHA protein and the R42A mutant were purified from E. coli (Figure S2A) and used to precipitate ICP0 from lysates of transfected 293T cells. We observed that GST-RNF8-FHA but not the R42A mutant bound ICP0 (Figure 2A). To determine whether the interaction was phosphorylation-dependent, we treated lysates with alkaline phosphatase (CIP) prior to GST pulldown. GST-RNF8-FHA was unable to retrieve ICP0 from lysates after CIP treatment, and this was rescued when the phosphatase inhibitor sodium orthovanadate (Na3VO4) was included (Figure 2A). We also observed faster migration of ICP0 after phosphatase treatment, consistent with dephosphorylation of the protein. These data demonstrate that the FHA domain of RNF8 is sufficient to interact with ICP0 in a phosphorylation-dependent manner.

Figure 2. ICP0 requirements for RNF8 interaction.

Figure 2

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(A) ICP0 was transfected into 293T cells and harvested 24h post-transfection. Cells were lysed in GST lysis buffer and, where indicated, lysates were treated with CIP or CIP plus sodium Na3VO4, and then incubated with GST-RNF8-FHA or GST-RNF8-FHA-R42A and precipitated using glutathione-Sepharose beads. (B) Top panel, functional domains of ICP0 and location of the RNF8 consensus motif. Bottom panel, eGFP-ICP0 or the T67A mutant were transfected into 293T cells, harvested 24h post-transfection, and lysates were used in GST pulldowns with GST-RNF8-FHA or GST-RNF8-FHA-R42A. (C) eGFP-ICP0 WT or T67A were co-transfected with Flag-RNF8 into HeLa cells, fixed 24 h post-transfection, and localization assessed by immunofluorescence. (D) eGFP-ICP0 or the T67A mutant was co-transfected into HeLa cells with eGFP-ICP0 in 3× excess over Flag-RNF8. Cells were harvested 24 h post-transfection. (E) eGFP-ICP0 or the T67A mutant were co-transfected into HeLa cells with eGFP-ICP0 in 3× excess over HA-RNF168 and analyzed by immunoblotting. (F) HeLa cells were co-transfected with RNF8 and/or RNF168 in excess of ICP0, subject to 10 Gy IR 24 h post-transfection, and fixed and processed for immunofluorescence 1 h post-IR. (G) Cells expressing ICP0 (30 cells for each condition imaged in (F) were scored for 53BP1 co-localization with γH2AX. Cells in which 53BP1 was localized to non-IRIF foci were excluded from this analysis. In a parallel analysis, it was confirmed that cells expressing ICP0 or mutants thereof had also been transfected with RNF8 and/or RNF168 (see also Supplemental Figures S2, S3, and S4).

ICP0 T67 is required for interaction with the RNF8 FHA domain

The FHA domain of RNF8 has a preference for phospho-Thr with Phe or Tyr in the pT+3 position (Huen et al., 2007). We scanned the ICP0 amino acid sequence for Thr residues conforming to the binding consensus motif for the RNF8 FHA domain and detected only one such consensus motif, TELF, at amino acids 67-70 (Figure 2B, upper panel). This sequence is conserved in other strains of HSV-1 and in ICP0 of HSV-2 (Figure S2B), which shares 61% identity with the HSV-1 ICP0 sequence. We generated a T67A mutant of eGFP-ICP0 and tested its ability to interact with RNF8 using GST pulldown assays. GST-RNF8-FHA retrieved WT but not T67A eGFP-ICP0 from cell lysates (Figure 2B, lower panel). Mutations in other functional domains of ICP0 and at previously described ICP0 phosphorylation sites did not affect ICP0 interaction with RNF8 (Figure S2C).

We next examined the sub-cellular localization of eGFP-ICP0 T67A by immunofluorescence. In the presence of eGFP-ICP0, Flag-RNF8 was re-localized to ICP0 foci, but in the presence of the T67A mutant Flag-RNF8 remained diffusely nuclear (Figure 2C). Interestingly, we noted that the F70A mutant of ICP0 could still co-localize with RNF8 and that this was through the RNF8 FHA domain (Figure S2D). Together, these experiments demonstrate that ICP0 T67 is required for interaction with the RNF8 FHA domain, although in the context of this transfection experiment ICP0 F70 is not strictly required.

To determine whether the ICP0 T67A mutation affected the ability of ICP0 to degrade RNF8 or RNF168, we co-transfected eGFP-ICP0 or the T67A mutant with Flag-RNF8 or HA-RNF168. We observed that WT eGFP-ICP0 but not T67A expression resulted in a decrease in Flag-RNF8 levels (Figure 2D). In contrast, both WT eGFP-ICP0 and the T67A mutant were able to degrade HA-RNF168 (Figure 2E). These data demonstrate that RNF8 is targeted for degradation by the ICP0 T67 residue, whereas RNF168 is targeted by ICP0 via a different mechanism.

We previously showed that ICP0 expression blocks 53BP1 recruitment to ionizing radiation-induced foci (IRIF) due to ICP0-mediated degradation of RNF8 and RNF168 (Lilley et al., 2010). We thus analyzed the ability of ICP0 T67A to block 53BP1 recruitment to IRIF. We observed that 53BP1 recruitment to sites of DNA damage was blocked by both WT eGFP-ICP0 and eGFP-ICP0-T67A (Figure 2F, left two panels). This block is likely due to the T67A mutant retaining the ability to degrade RNF168, which is also required for 53BP1 recruitment. Supporting this hypothesis, we observed that overexpression of RNF168 alone was sufficient to rescue 53BP1 recruitment to IRIF in the presence of eGFP-ICP0 T67A, while recruitment of 53BP1 in the presence of eGFP-ICP0 WT required overexpression of both RNF8 and RNF168 (Figures 2F, 2G, and S3A) (Lilley et al., 2010). Finally, in the presence of eGFP-ICP0-T67A, the RNF8 that did not localize to ICP0 foci was able to accumulate at IRIF (Figure S3B). In addition, we observed that ICP0 expression has no effect on ATM signaling after IR or on the levels or subcellular localization of Ubc13 and Uve1, E2 enzymes proposed to be involved in this process (Figures S4A, S4B, S4C, and data not shown). These data indicate that targeting of RNF8 by the T67 residue on ICP0 is functionally important for ICP0 to prevent recruitment of 53BP1 to IRIF.

Phosphorylation of ICP0 T67 is necessary and sufficient for binding the RNF8 FHA domain

To investigate whether phosphorylation of ICP0 T67 directly mediates the RNF8-ICP0 interaction, we purified the untagged RNF8 FHA domain from E. coli (Figure S5A) and used isothermal titration calorimetry (ITC) to measure the affinity of synthetic ICP0 peptides to the FHA domain (Figure 3). A peptide containing phosphorylated T67 bound the RNF8 FHA domain (Figure 3A), whereas a nonphosphorylated peptide of the same sequence did not (Figure 3B). The interaction was not observed when the FHA domain contained the R42A mutation (Figure 3C). We measured the dissociation constant to be 0.833 μM, which is comparable to or better than those observed for the TQXF motifs in Mdc1 (3.1 μM to 11.7 μM) (Huen et al., 2007). Additionally, we tested the F70A mutant pT67 peptide and found that the affinity of this peptide was 10× to 20× lower than for the WT peptide (Figures 3A, S5B, and S5C). These data demonstrate that ICP0 pT67 is both necessary and sufficient to bind directly to the RNF8 FHA domain, and that the F70 residue does increase the affinity for this interaction in vitro, consistent with previous observations (Huen et al., 2007).

Figure 3. Phosphorylation of ICP0 at T67.

Figure 3

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(A) The RNF8 FHA domain or R42A mutant were purified from E. coli and peptides comprising the region around ICP0 T67 were synthesized. Binding of the peptides to the FHA domains was measured using isothermal calorimetry. 1.2mM peptides were titrated into solutions containing 100mM RNF8 FHA domain. pT67 peptide bound the RNF8 domain. (B) Nonphosphorylated peptide did not bind the RNF8 FHA domain. (C) R42A mutation in the FHA domain prevented binding to pT67 peptide. (D) Antibodies were raised in rabbits against a synthetic peptide containing ICP0 pT67 and affinity purified. Peptide (1 μg) was spotted onto nitrocellulose and analyzed by immunoblotting using sera purified over nonphosphorylated peptide (α-T67 total) or double purified over nonphosphorylated and pT67 peptide (α-pT67). (E) eGFP-ICP0 WT and T67A were transfected into HeLa cells and processed for immunofluorescence using α-pT67 antibodies. (F) eGFP-ICP0 WT or T67A were transfected into HeLa cells and lysates were prepared. Before analysis, one lysate expressing WT eGFP-ICP0 was subjected to dephosphorylation by l phosphatase for 30 min at 37°C (see also Supplemental Figure S5).

ICP0 T67 is phosphorylated in cells

To demonstrate that ICP0 T67 is indeed phosphorylated in cells, we generated antibodies specific for the phosphorylated T67 residue of ICP0 (Figure 3D). We observed that the α-pT67 antibodies recognized and co-stained with eGFP-ICP0 WT by immunofluorescence, but failed to recognize the eGFP-ICP0 T67A mutant, demonstrating specificity for the ICP0 T67 residue (Figure 3E). Additionally, the α-pT67 antibodies recognized WT eGFP-ICP0 protein by immunoblotting, but this signal was lost when proteins in the lysates were dephosphorylated using λ phosphatase, or if the lysate contained the ICP0 T67A mutant (Figure 3F). Combined, these data demonstrate that the ICP0 protein is phosphorylated on T67 in cells and validate the specificity of the α-pT67 antibodies.

CK1 phosphorylates ICP0 T67 to facilitate interaction with RNF8

Bioinformatic analysis of the ICP0 T67 region indicated the presence of high probability consensus sites for the cellular kinases CK2 and CK1, which phosphorylate Ser and Thr residues in acidic regions. In vitro phosphorylation reactions using purified GST-ICP0-241 protein, containing the N-terminal 241 amino acids of ICP0 (Figure S6A), revealed that CK1, but not CK2, could phosphorylate T67 as measured by reactivity to the α-pT67 antibodies (Figure 4A). The T67A mutant of GST-ICP0-241 was not phosphorylated by either CK1 or CK2, providing further evidence that the observed CK1-mediated phosphorylation was specifically on T67 (Figure 4A).

Figure 4. Mechanism of ICP0 T67 phosphorylation.

Figure 4

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(A) Recombinant GST-ICP0-241 or the T67A mutant were purified from E. coli, phosphorylated in vitro by recombinant CK1δ̣ or CK2α̃β, and analyzed by SDS-PAGE and immunoblotting. (B) GST-ICP0-241 or the indicated mutants were phosphorylated in vitro by recombinant rat CK1d and used to precipitate Flag-RNF8 from lysates via glutathione-Sepharose. (C) The region of ICP0 containing T67 (red, bold) and the upstream serines (red) is indicated. Recombinant GST-241 or the indicated mutants were phosphorylated in vitro by CK1. Reactions were quenched at 5 min intervals between 0 and 20 min, and phosphorylation of T67 was measured by reactivity to α-pT67 antibodies. (D) Plasmids expressing the ICP0-nls241, the S64A, or T67A mutants were transfected into HeLa cells and analyzed by SDS-PAGE and immunoblotting with α-pT67 antibodies. (E) Plasmids expressing the indicated ICP0 sequences were transfected into 293T cells, lysed in GST lysis buffer and used in GST pulldown assays with recombinant GST-RNF8-FHA or the R42A mutant. (F) Plasmids expressing the indicated ICP0 proteins were co-transfected into HeLa cells with Flag-RNF8 and localization was assessed by immunofluorescence. (G) Cells expressing ICP0 and Flag-RNF8 proteins (50 cells for each condition represented in F) were counted and scored for the ability of RNF8 to localize with ICP0. Cells in which any ICP0 foci co-localized with RNF8 were scored as positive for RNF8/ICP0 co-localization (see also Supplemental Figure S6).

We next tested whether CK1-mediated phosphorylation of GST-ICP0-241 could facilitate interaction with RNF8. Lysates from 293T cells transfected with Flag-RNF8 were incubated with GST-ICP0-241 WT, T67A, or F70A proteins and glutathione-Sepharose beads. The nonphosphorylated GST-ICP0-241 protein was unable to retrieve Flag-RNF8 from lysates, but did interact with RNF8 after phosphorylation by CK1 (Figure 4B), and this was dependent on the RNF8 R42 residue (Figure S6B). Furthermore, we observed that GST-ICP0-241 T67A and F70A mutants could not retrieve WT Flag-RNF8 from lysates even after phosphorylation by CK1 (Figure 4B). In the case of the T67A mutant this is due to the inability to generate the pT67 phosphosite as demonstrated by lack of reactivity to α-pT67 antibodies. In the case of F70A, the T67 phosphosite was generated after phosphorylation by CK1 (Figure 4B), but RNF8 was unable to bind. We thus concluded that CK1-mediated phosphorylation of ICP0 facilitates interaction with RNF8 via the FHA domain.

CK1 phosphorylates Ser and Thr residues containing phosphoamino acids in the -3 position, or to a lesser extent, Ser or Thr residues in acidic regions (Flotow et al., 1990; Flotow and Roach, 1991). ICP0 contains three Ser residues upstream of T67: S58, S60, and S64, which are interspersed with acidic Asp and Glu residues (Figure S2B). To determine whether these residues are required for T67 phosphorylation by CK1, in vitro phosphorylation reactions were performed using GST-ICP0-241 WT, T67A, or S58/60/64A (S3A) recombinant proteins (Figure S6A) as substrates. We observed CK1-mediated T67 phosphorylation on WT GST-ICP0-241 over a phosphorylation time course, as measured by reactivity to the α-pT67 antibodies. In contrast, T67 was not phosphorylated on the T67A and S3A mutant proteins (Figure 4C). Lack of α-pT67 reactivity after CK1 phosphorylation of the S3A mutant was not due to epitope loss resulting from the Ala mutations, since these antibodies were generated and purified using peptides lacking the upstream Ser residues (Figure 3D). Furthermore, the α-pT67 antibodies robustly reacted with a synthetic S3A peptide containing phosphorylated T67 (Figure S6C). GST-ICP0-241 S3A protein could not retrieve Flag RNF8 from cell lysates even after phosphorylation by CK1 (Figure S6B), providing additional evidence that T67 was not phosphorylated in the S3A mutant. These observations indicate that Ser 58, 60, and/or 64 contribute to CK1-mediated phosphorylation of ICP0 T67. In these experiments it is likely that CK1 is catalyzing its own S64 priming phosphorylation, facilitated by acidic residues in this region. However, bioinformatic analysis indicates that Ser 58, 60, and 64 are much higher probability CK2 consensus sites, and it is possible that in the cellular context, CK2 phosphorylation of S64 primes for CK1-mediated phosphorylation of T67. Consistent with these observations, both CK1 and CK2 appear to be able to catalyze phosphorylation at S58, S60, and/or S64 in vitro (Figures S6D and S6E).

To test the individual contribution of ICP0 S64, which is the pT-3 residue that most likely contributes to CK1-mediated phosphorylation of T67, we used mammalian expression plasmids encoding the first 241 amino acids of ICP0 fused to a nuclear localization signal (ICP0-nls241). Plasmids encoding WT, T67A, or S64A proteins were transfected into 293T cells. We observed phosphorylation at T67 on WT ICP0-nls241 but not the T67A or S64A mutants (Figure 4D), indicating that S64 is required for T67 phosphorylation on ICP0 in cells. We next used GST-pulldown assays with S58A, S60A, and S64A mutants of ICP0-nls241 in order to dissect the individual contributions of these Ser residues to the RNF8-ICP0 interaction. WT or mutants of ICP0-nls241 were expressed in 293T cells, and lysates were incubated with purified recombinant GST-RNF8-FHA protein or the R42A mutant. We observed that GST-RNF8-FHA was able to retrieve ICP0-nls241 WT, S58A, and S60A from lysates, whereas T67A and S64A mutants did not interact (Figure 4E). Supporting these observations, full length ICP0 protein containing the S64A mutation was unable to co-localize with RNF8 in cells, whereas the S58A mutant was more similar to WT ICP0, in that it was able to partially co-localize with RNF8 (Figures 4F and 4G). The inability of S64A to bind the RNF8 FHA domain most likely reflects its in vivo contribution to T67 phosphorylation, since pS64 does not significantly contribute to direct binding in vitro (Figure S6F). We thus conclude that S64 is required as a priming phosphorylation site for CK1-mediated phosphorylation of T67 and subsequent RNF8 binding.

Relevance of the ICP0 T67A mutation during HSV-1 infection

We next constructed HSV-1 containing the T67A mutation within the ICP0 gene to determine the effects of the mutation during HSV-1 infection. HeLa cells were infected with WT, ΔICP0, or T67A HSV-1 at MOI=3 and lysates were collected at 2 h intervals from 0 to 8 hpi. We observed decreased levels of ICP0 targets over the course of WT HSV-1 infection (Figure 5A). During infection with T67A virus, RNF8 levels were comparable to those observed during infection with ΔICP0 virus, indicating that T67 promotes RNF8 degradation during viral infection. We sometimes observed slight RNF8 loss in an ICP0-independent manner at late time points during infection. This may be due to HSV-1 effects on cell cycle, (de Bruyn Kops and Knipe, 1988) since it has been reported that RNF8 levels are cell cycle regulated (Plans et al., 2008). In contrast to the stabilized RNF8 levels during T67A virus infection, we observed that DNA-PKcs, USP7, and RNF168 were still degraded by the T67A virus (Figure 5A). Combined, these observations indicate that T67 is required to promote ICP0-mediated RNF8 degradation during HSV-1 infection, while other substrates tested in this experiment are targeted via different mechanisms.

Figure 5. Effect of the T67A mutation on virus infection.

Figure 5

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(A) WT, ΔICP0, or T67A mutants of HSV-1 (strain KOS) were used to infect HeLa cells at MOI=3 and harvested at the indicated time points. (B) The indicated viruses were infected at MOI=0.01 in HepaRG cells expressing a scrambled shRNA (shNeg) or shRNA targeting RNF8 (shRNF8) and harvested at the indicated time points. Transcripts were quantified via qPCR with primers targeted to the immediate-early ICP27 gene and are represented as the relative transcript level at each time point. Error bars represent standard deviation of triplicate samples. (C) 25% of the cell pellets from the samples in (B) was used to isolate genomic DNA. ICP27 gene copies, indicative of the relative number of viral genomes, were quantified and analyzed by qPCR as described in (B). Error bars represent standard deviation of triplicate samples (see also Supplemental Figure S7).

We have previously observed that early transcription events during HSV-1 infection are repressed by RNF8 (Lilley et al., 2011). We therefore examined whether the ICP0 T67A virus, which is impaired for RNF8 degradation during infection, was transcriptionally repressed as compared to WT HSV-1 during early stages of infection. HepaRG cells expressing a control shRNA (shNeg) or shRNA targeting RNF8 (shRNF8) (Lilley et al., 2011) were infected with WT, T67A, or ΔICP0 HSV-1 at MOI=0.01 and harvested at 2, 6, and 12 hpi. RNA was isolated, reverse transcribed, and transcript levels were measured using qPCR with primers targeted to the viral ICP27 transcript. We observed that the T67A virus was impaired in its ability to generate ICP27 transcripts as compared to WT virus at all time points, and that the ΔICP0 virus was significantly more impaired than the WT or T67A viruses (Figure 5B), likely due to loss of its ability to target any ICP0 substrates. These ICP0- and T67A-dependent defects were also observed during infection of HFFF cells and mouse embryonic fibroblasts (MEFs) (data not shown). Furthermore, we observed that RNF8 knockdown by shRNF8 expression improved the ability of all viruses, including WT HSV-1, to generate ICP27 transcripts as compared to shNeg cells. In the case of the T67A virus, the increased transcript levels in shRNF8 cells did not reach WT HSV-1 levels in either shNeg or shRNF8 cells, indicating that there may be other cellular proteins targeted through ICP0 pT67 that additionally contribute to transcriptional repression of the viral genome. The ICP0-, T67-, and RNF8-dependent effects on transcription are not attributable to lower viral genome entry into cells, as input genomes were well matched between all three viruses in shNeg and shRNF8 cells (Figure 5C, 2 h time point). Interestingly, when assessing replication of these input viral genomes, the T67A virus was impaired approximately 6-fold in the amount of replicated DNA at 12 hpi (Figure 5C). This defect, however, does not appear to be RNF8-dependent as similar T67-dependent replication defects were observed in shNeg as compared to shRNF8 cells. Finally, we observed that the T67A virus was deficient for progeny production, as progeny production at 16 hpi yielded approximately 10-fold less progeny than WT virus (Figure S7).

ICP0 T67 can bind other cellular proteins

Since the T67A virus is impaired in viral transcript generation in both RNF8-dependent and -independent manners, and replication appears to be impaired in an RNF8-independent manner, we next sought to identify other proteins that could be targeted through the ICP0 pT67 region. We used biotinylated nonphosphorylated peptide or peptides phosphorylated at T67 (pT67), S64 (pS64), or S64 and T67 (pS64/pT67) (Figure 6A) to isolate cellular proteins from 293T lysates. The pT67 and pS64/pT64 peptide pulldowns contained unique bands at approximately 60 and 102 kDa (Figure 6B). We analyzed proteins co-purifying with nonphosphorylated and pT67/S64 peptides via mass spectrometry and identified several hundred proteins that co-purified uniquely with pS64/pT67 (Table S1). Chk2 was one of the most abundant proteins in this pulldown (Figure 6C), and additional mass spectrometric analysis of the 60 kDa bands in the pT67 and pS64/pT67 purifications confirmed that they predominantly contained Chk2 peptides. Interestingly, this protein and two others identified in the pS64/T67 pulldowns, Nbs1 and FOXK2, also contain FHA domains. To validate the mass spectrometry results and dissect the individual contributions of pS64 and pT67 to the interactions, we repeated the peptide pulldowns and analyzed precipitated proteins by SDS-PAGE and immunoblotting. In all cases, the nonphosphorylated and pS64 peptides did not pull down RNF8, Chk2, Nbs1, or FOXK2, whereas pT67 and pS64/p67 peptides did (Figure 6D). These observations demonstrate a dependence on pT67.

Figure 6. ICP0 pT67 can bind other cellular proteins.

Figure 6

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(A) Biotinylated peptides were synthesized that were nonphosphorylated or phosphorylated at T67, S64, or S64/T67. (B) Peptides were used to precipitate cellular proteins from 293T lysates using Streptavidin beads and analyzed by silver staining after SDS-PAGE. (C) Co-purifying FHA domain proteins were identified by mass spectrometry. FHA domain proteins purifying uniquely in the presence of pS64/pT67 peptide are listed, with the corresponding number of peptides identified. (D) Peptide pulldowns were analyzed by immunoblotting using antibodies to proteins identified by mass spectrometry in (C). (E) TAP-ICP0 or GFP-ICP0 was co-transfected with HA-Chk2 into 293T cells and TAP-ICP0 was precipitated using streptavidin-Sepharose. (F) 293T cells were co-transfected with HA-Chk2 and eGFP-ICP0 WT or the T67A mutant, and eGFP-ICP0 proteins were immunoprecipitated from the resulting lysates using α-GFP antibodies and protein A/G-agarose beads. (G) ICP0 pT67 peptide (0.35 mM) was titrated into a solution containing purified Chk2 FHA (27 μM) and binding affinity measured by ITC. The peptide bound with comparable affinity to RNF8 FHA domain. (H) The unphosphorylated peptide did not bind. (I) The F70A mutation did not significantly affect the binding to Chk2 FHA (see also Supplemental Table S1).

We next analyzed the ability of Chk2 to interact with full length ICP0 protein in cells. ICP0 protein tagged with a tandem affinity purification (TAP) tag, comprising streptavidin-binding and calmodulin-binding peptides, was co-transfected into 293T cells together with HA-Chk2, and TAP-ICP0 was affinity purified from lysates using streptavidin Sepharose. As a negative control, eGFP-ICP0 was co-transfected with HA-Chk2. We observed HA-Chk2 to co-purify with TAP-ICP0 but not eGFP-ICP0 on streptavidin-Sepharose, providing evidence that Chk2 and ICP0 interact in cells (Figure 6E). We then assessed the contribution of T67 to this interaction. 293T cells were co- transfected with HA-Chk2 in the presence of WT or T67A eGFP-ICP0 and immunoprecipitated using α-GFP antibodies. We observed that Chk2 co-immunoprecipitated with WT but not T67A eGFP-ICP0, demonstrating that the Chk2-ICP0 interaction is T67 dependent (Figure 6F). Interestingly, we found that the FHA domain of Chk2 was sufficient to bind ICP0 pT67 by ITC, with a similar dissociation constant as observed for the RNF8 FHA domain (Figure 6G). This interaction was dependent on pT67 (Figure 6H), but unlike RNF8 FHA, the interaction was not significantly affected by the F70A mutation (Figure 6I). It is interesting to note it has previously been reported that Chk2 promotes ICP0-mediated cell cycle arrest and enhancement of viral growth (Li et al., 2008), providing functional relevance for the interaction we describe. Combined, these experiments demonstrate that other cellular proteins can be targeted via the ICP0 pT67 residue, validating this approach to identify relevant targets. Thus the pT67 region of ICP0 may be utilized in a generalized strategy to bind and redirect other cellular proteins, including other FHA domain proteins, functioning in pathways relevant to the viral lifecycle.

Discussion

The studies described here demonstrate that the HSV-1 encoded E3 ubiquitin ligase ICP0 has evolved a short linear motif, STD(p)TELF, to mimic a cellular phosphosite normally catalyzed on Mdc1 by ATM after DNA damage. The significance of this targeting is highlighted by reduced viral transcription and replication from HSV-1 containing the T67A mutation in ICP0. These observations indicate that ICP0 pT67 functions to help counteract the repressive effects of RNF8 on viral transcription, thereby promoting the viral lifecycle. Our results are in keeping with the recently described effects of RNF8 on transcription following DNA damage (Shanbhag et al., 2010). We observed decreased accumulation of ICP27 transcripts for the T67A virus, and it will be interesting to assess the impact on other viral genes. Furthermore, it appears that ICP0 can bind other cellular proteins via pT67, implicating this targeting mechanism as a broader viral strategy to modulate cellular pathways at key regulatory nodes (summarized in Figure 7).

Figure 7. ICP0 mimicry of cellular phosphorylation marks targets cellular FHA domain proteins, promoting viral infection.

Figure 7

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Parallels between RNF8-Mdc1 interaction during the DNA damage response and RNF8-ICP0 interaction during HSV-1 infection are shown. The motifs on ICP0 and Mdc1 use distinct mechanisms to catalyze phosphorylation, but both employ the induced phosphosite to bind RNF8. Also depicted are consequences for the virus and the possibility that ICP0 (or other viral proteins) can use this mimicking approach to target other cellular proteins.

Chk2 is not degraded during infection with HSV-1 (Lilley et al., 2005) and it has been reported that Chk2 promotes growth of WT virus in an ICP0-dependent manner (Li et al., 2008). Combined with our finding that pT67 is involved in binding Chk2, these data implicate this region in targeting additional cellular factors in order to harness signaling pathways that enhance virus growth. They also suggest that the Chk2 FHA domain may bind sequences that diverge from the optimal consensus (Li et al., 2002) in vitro and in the cellular context, although binding is less dependent on the +3 Phe residue than for the RNF8 FHA domain.

While ICP0 mimics Mdc1-like short linear motifs in conforming to the binding consensus sequence for the RNF8 FHA domain, it is interesting to note that differences between the Mdc1 and ICP0 sequences lead to distinct phosphorylation mechanisms. The presence of Glu instead of Gln at pT+1 and the presence of Ser at pT-3 result in loss of the ATM phosphorylation site (TQ) and generation of a CK1 site (pSXXT) for ICP0 T67. A key difference between ATM and CK1 is that while ATM normally exists as an inactive dimer in cells that is activated in response to DNA damage (Bakkenist and Kastan, 2003), several of the six human CK1 isoforms are constitutively active kinases, and all display similar preferences for the target sequences that they phosphorylate, regulated by hierarchical priming mechanisms (Cheong and Virshup, 2011). This means that CK1 is available to be recruited by ICP0 once its synthesis is completed, without the requirement for upstream signaling events. While it has been demonstrated that ATM is activated upon lytic HSV-1 infection, this activation requires the formation of pre-replicative sites or the onset of viral replication (Lilley et al., 2005; Wilkinson and Weller, 2004). ICP0 is most important during the earliest stages of infection, before the onset of viral replication, at a time when ATM may not be fully activated.

Although the studies presented here do not address the mechanism of RNF168 targeting by ICP0, our data indicate that it is clearly not targeted through pT67. RNF168 loss in the presence of ICP0 does not appear to be an indirect consequence of the RNF8 degradation, since RNF168 levels do not change in RNF8 knockdown cells, and RNF168 is still degraded when RNF8 degradation is impaired in the presence of ICP0 T67A. Since ICP0 is heavily autoubiquitinated in cells, it is possible that interaction with the cellular ligase may be mediated through RNF168 MIU/UMI recognition of ubiquitin chains on ICP0. Conversely, ICP0 has recently been described to bind SUMO via several SUMO-interacting-motif-like sequences (Boutell, et al., 2011), and it may therefore be possible that SUMOylation of RNF168 plays a role in targeting by ICP0. The E3-E3 interactions between RNF8 and ICP0, and RNF168 and ICP0, are part of an evergrowing list of E3-E3 interactions (Weissman et al., 2011). Further analysis of these interactions and the mechanisms regulating E3-E3 targeting may provide insights into more general mechanisms regulating E3 turnover, in particular highlighting the regulatory role of post-translational modifications.

Experimental Procedures

Details on cell lines, antibodies, viruses, plasmids, qPCR, protein purification, and mass spectrometry can be found in the Supplemental Information.

GST pulldowns

ICP0 or RNF8 and mutants thereof were expressed in 293T cells and harvested 24 hpi. Cells were lysed in GST lysis buffer (20 mM Tris-HCl 8, 200 mM NaCl, protease inhibitors (Roche), 2 mM PMSF, and 0.5% NP-40). Lysate (500 μg) was incubated with 10 μg GST-RNF8-FHA or GST-ICP0-241 and mutants thereof, and 20 μl glutathione-Sepharose in 500 μl total volume for 2 h at 4°C. Where indicated, 100 U CIP and/or 200 μM sodium orthovanadate was added to the lysate and incubated for 30 min at 30°C prior to pulldown. For in vitro phosphorylation of GST-ICP0-241 before pulldown, 10 mg protein was incubated with 1000 U rat CK1δ (NEB) for 30 min at 30°C in a 20 μl reaction containing 50 mM Tris pH 7.5, 10 mM MgCl2 and 5 mM DTT, with or without ATP (10 mM). The entire phosphorylation reaction was added to the pulldown.

Isothermal Titration Calorimetry

RNF8-FHA WT and R42A proteins were buffer matched over a NAP-5 column (GE Healthcare) into ITC buffer (25 mM Tris-HCl pH 8.0, 200 mM NaCl). The indicated peptides were synthesized at the Hartwell Center at St. Jude Children's Research Hospital. Peptide solutions were injected into a sample cell containing RNF8-FHA or Chk2 FHA domain as detailed in Supplemental Methods. Measurements were performed on a Microcal ITC200 and binding isotherms were plotted and analyzed using Origin (v7.0).

qPCR

1 ×106 HepaRG cells expressing shNeg or shRNF8 were infected with HSV-1 and mutants thereof at MOI=0.01. Cells were harvested at the indicated time points. RNA was isolated (RNeasy, Qiagen) and 500 ng RNA was reverse transcribed using SuperScript III and oligo dT (Invitrogen) in a 20 μl reaction. qPCR reactions were prepared in at least triplicate in 25 μl volumes using 3 μl cDNA or 100 ng genomic DNA, and SYBR Green qPCR master (ABI). The viral ICP27 transcript was detected using primers GCATCCTTCGTGTTTGTCATT (F) and GCATCTTCTCTCCGACCCCG (R) and normalized to endogenous RPLPO transcript detected using primers CTGGAAGTCCAACTACTTCC (F) and TGCTGCATCTGCTTGGAGCC (R).

Peptide pulldowns

Biotinylated peptides were synthesized and conjugated to biotin at the Salk Institute Peptide Synthesis Core. Peptide (10 μg) was incubated with 60 μl streptavidin beads and 5 mg cell lysate (20 mM Tris-HCl (pH 8.0), 200 mM NaCl, protease inhibitors (Roche), 2 mM PMSF, and 0.5% NP-40), and rotated for 2 h at 4°C. Beads and precipitated proteins were washed 3× in lysis buffer, boiled in 2× SDS loading dye, separated by SDS-PAGE, and visualized by silver staining. For mass spectrometry analysis, bead-bound proteins were directly denatured, reduced, and digested with trypsin.

Co-immunoprecipitation and co-affinity purification

293T cells were co-transfected with indicated plasmids and harvested 24 h post-transfection. Cells were lysed in lysis buffer (20 mM Tris-HCl (pH 8.0), 200 mM NaCl, protease inhibitors (Roche), 2 mM PMSF, and 0.5% NP-40) and 500 μg cleared lysate was incubated with 40 μl streptavidin-Sepharose (GE) or 4 μl α-GFP antibodies (Clontech) plus 40 μl protein A/G-agarose (Santa Cruz) for 2 h at 4°C in 500 μl total volume. Precipitated proteins were washed in lysis buffer, resuspended and boiled in SDS loading dye, and analyzed by immunoblotting.

Supplementary Material

Figure S1
Table S1

Highlights.

  • HSV-1 ICP0 encodes a mimicking phosphosite to bind RNF8 via the RNF8 FHA domain

  • CK1 catalyzes phosphorylation at ICP0 T67 to enable RNF8 binding

  • ICP0 T67 is important to promote RNF8 degradation and the viral life cycle

  • ICP0 can bind other cellular proteins through pT67

Acknowledgments

We thank members of the Weitzman Lab for discussions and Jennifer Olszewski for help with protein purifications. We are grateful to M. Yaffe, D. Durocher, D. Davido, R. Everett, D. Stern, and J. Lukas for gifts of reagents. Work on viruses and DNA repair in the Weitzman Lab has been supported by grants from NIH (AI067952, CA097093, and AI051686) and a Pioneer Developmental Chair from the Salk Institute. M.S.C. is supported by a predoctoral Ruth L. Kirschstein National Research Service Award (NIH/NCI T32 CA009523) and by a gift from the H.A. & Mary K. Chapman Charitable Trust. S.L. is supported by the Natural Sciences & Engineering Research Council of Canada. A.A. is supported by an American Cancer Society Postdoctoral Fellowship. The Peptide Synthesis Facility at Salk is supported by a Cancer Center Support grant from NCI (CA14195). Work in the Schulman Lab is supported by ALSAC, the St. Jude Cancer Center grant (NIH5P30CA021765), NIH (R01GM069530), and the Howard Hughes Medical Institute. B.A.S. is an Investigator of the Howard Hughes Medical Institute. T.H. is a Frank and Else Schilling American Cancer Society Professor, and the work in his group was supported by a grant from the NCI (CA80100). Work in the Yates Lab is supported through an NIH grant (P41 RR011823).

Footnotes

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Supplementary Materials

Figure S1
NIHMS366769-supplement-Figure_S1.doc (31.7MB, doc)
Table S1
NIHMS366769-supplement-Table_S1.xls (58.5KB, xls)

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