DNA Mismatch Repair Proteins Are Required for Efficient Herpes Simplex Virus 1 Replication

Abstract

Herpes simplex virus 1 (HSV-1) is a double-stranded DNA virus that replicates in the nucleus of its human host cell and is known to interact with many cellular DNA repair proteins. In this study, we examined the role of cellular mismatch repair (MMR) proteins in the virus life cycle. Both MSH2 and MLH1 are required for efficient replication of HSV-1 in normal human cells and are localized to viral replication compartments. In addition, a previously reported interaction between MSH6 and ICP8 was confirmed by coimmunoprecipitation and extended to show that UL12 is also present in this complex. We also report for the first time that MLH1 associates with ND10 nuclear bodies and that like other ND10 proteins, MLH1 is recruited to the incoming genome. Knockdown of MLH1 inhibits immediate-early viral gene expression. MSH2, on the other hand, which is generally thought to play a role in mismatch repair at a step prior to that of MLH1, is not recruited to incoming genomes and appears to act at a later step in the viral life cycle. Silencing of MSH2 appears to inhibit early gene expression. Thus, both MLH1 and MSH2 are required but appear to participate in distinct events in the virus life cycle. The observation that MLH1 plays an earlier role in HSV-1 infection than does MSH2 is surprising and may indicate a novel function for MLH1 distinct from its known MSH2-dependent role in mismatch repair.

INTRODUCTION

Herpes simplex virus 1 (HSV-1) is a large double-stranded DNA virus that replicates in the nucleus of the host cell. Although cis- and trans-acting functions involved in HSV-1 replication in infected cells have been identified (69), the mechanism of HSV-1 DNA replication remains poorly understood. The appearance of viral genomes in the nucleus might be expected to induce a cellular DNA damage response, and it is now clear that HSV-1 has evolved a complex relationship with the host DNA damage response pathways (33, 48, 60, 66, 70, 71). During infection, cellular factors that are beneficial to the virus are hijacked and recruited, while other factors and pathways are degraded or inactivated. Previous work from our group and others has shown that the ATR (ATM- and Rad3-related) and DNA-PK (DNA-dependent protein kinase) DNA damage-sensing kinases are attenuated in HSV-1-infected cells, while the ATM (ataxia-telangiectasia-mutated) kinase is activated (31, 34, 48, 50). In addition, other components of the DNA damage repair and response pathways also appear to be beneficial to the virus and are utilized during viral replication (35). Many host cellular DNA damage proteins are known to localize to viral replication compartments (33, 48, 60, 66, 71). Furthermore, in a recent proteomic analysis of replication compartments, several DNA repair proteins were identified as potential interacting partners of the viral single-stranded DNA (ssDNA) binding protein ICP8 (66). One such potential interacting partner, the DNA mismatch repair (MMR) protein MSH2, was shown to localize to the replication compartments in infected cells (66).

The DNA MMR system is highly conserved from prokaryotes to humans and executes several important cellular functions in order to maintain the integrity of the genome (20, 32). Higher eukaryotes possess three MutS homologs that function in MMR (MSH2, MSH3, and MSH6). MSH2 interacts with either MSH6 or MSH3 to form two distinct but partially redundant heterodimers. The MSH2-MSH6 heterodimer recognizes both single-base-pair mismatches and 1- or 2-base-pair insertion/deletion loops (IDLs), while the MSH2-MSH3 heterodimer recognizes larger IDLs. Four MutL homologs (MLH1, MLH3, PMS1, and PMS2) have been identified, which form three distinct heterodimers, with MLH1 acting as the common subunit. Together, the MSH2-MSH6, MSH2-MSH3, and MLH1-PMS2 heterodimers are required for the repair of DNA replication errors and the maintenance of genomic stability (20, 23). This is highlighted by the autosomal dominant disorder Lynch syndrome caused by inherited mutations of the DNA MMR genes (41), in which patients are at an increased risk of developing early onset colorectal, endometrial, and other cancers. Additionally, the MMR proteins play key roles in nonrepair mechanisms as well (23). In response to certain forms of DNA damage, the MMR proteins play an as-yet-undefined role in signaling cell cycle checkpoints and apoptosis. These proteins have also been reported to prevent recombination between heterologous sequences (11).

In this paper we address the question of whether HSV-1 utilizes MMR pathway components to promote its own replication or inactivates it. We report here for the first time that cellular MMR proteins are required for efficient HSV-1 replication and are recruited to viral replication compartments following infection. Furthermore, we demonstrate that MLH1 but not MSH2, MSH6, or MSH3 localizes to ND10 nuclear bodies in uninfected human fibroblasts. ND10 or promyelocitic leukemia (PML) nuclear bodies are punctate subnuclear structures that play important roles in intrinsic antiviral defense, transcriptional regulation, apoptosis, DNA damage response, and chromatin remodeling (13). Interestingly, although many other ND10 components studied to date have antiviral effects (15, 39), we show that MLH1 is required for efficient viral infection. Depletion of MLH1 exerts a profound effect on immediate-early (IE) viral gene expression, while depletion of MSH2 exerts a later effect. These results suggest that we have identified a novel function for MLH1 distinct from its usual role downstream of MSH2.

MATERIALS AND METHODS

Cells and reagents.

HeLa, HFF-1, U2OS, Vero, and pEAK cells were obtained from the American Type Culture Collection (ATCC). All cells except Vero cells were maintained in Dulbecco's modified Eagle medium (DMEM) with 10% fetal bovine serum; Vero cells were maintained in 5% fetal bovine serum. Vero-derived E11 cells were obtained from Neal DeLuca (University of Pittsburgh School of Medicine, Pittsburgh, PA). These cells stably express ICP4 and ICP27 and were maintained in 400 μg/ml G418 (Gibco) (57). Hec59 and HCT-116 cells were grown in DMEM–nutrient mixture F-12 (DMEM–F-12). Hec59+chromosome 2 and HCT-116+chromosome 3 cells were kindly provided by Thomas Kunkel and Alan Clark and were maintained in DMEM–F-12 supplemented with 400 μg/ml G418 (Gibco). Phosphonoacetic acid (PAA) was purchased from Sigma Chemical Co. (St. Louis, MO) and used at a concentration of 400 μg/ml. N-Methyl-N′-nitro-N-nitrosoguanidine (MNNG) (Chemical Abstracts Service [CAS] no. 70-25-7; obtained from the National Cancer Institute Chemical Carcinogen Reference Standard Repository) was dissolved in dimethyl sulfoxide (DMSO) to a concentration of 10 mM and stored at −20°C until use. O⁶-Benzylguanine (O⁶-BG; CAS no. 19916-73-5) was purchased from Sigma, dissolved in DMSO to a concentration of 25 mM, and stored at −80°C until use.

Viruses.

The KOS strain was used as wild-type HSV-1. The ICP0-null virus 0β and the ICP0-expressing virus d106 are derived from KOS and were provided by Neal DeLuca (University of Pittsburgh School of Medicine, Pittsburgh, PA). Virus 0β contains LacZ insertions in both copies of the ICP0 gene (58), and d106 is deleted for all immediate-early genes except ICP0 and contains the green fluorescent protein (GFP) gene under the control of the human cytomegalovirus (HCMV) promoter inserted in the ICP27 gene (56). The ICP0-null virus dl1403 is derived from strain 17+ and was provided by Roger Everett (MRC Virology Unit, Glasgow, Scotland) (63). Virus in1863 is derived from strain 17+ and was obtained from Chris Preston (MRC Virology Unit, Glasgow, Scotland). Virus in1863 contains the lacZ gene under the control of the HCMV promoter/enhancer inserted into the tk gene. Viruses vECFP-ICP4 and vEGdl110 were also derived from strain 17+ and were provided by Roger Everett (MRC Virology Unit, Glasgow, Scotland). Virus vECFP-ICP4 contains a functional ECFP-ICP4 fusion protein expressed from both copies of the ICP4 gene (18). Virus vEGdl110 is ICP0 null, is derived from dl1403, and contains enhanced green fluorescent protein (EGFP) insertions in both copies of the ICP0 gene (38). All viruses were grown and titrated on Vero cells except the ICP0-null viruses 0β, dl1403, and vEGdl110, which were grown and titrated on U2OS cells. Virus d106 was grown and titrated on E11 cells.

Lentivirus.

The pLKO.1 system was used to package lentiviruses and deliver short hairpin RNA (shRNA) as previously described (48). The following shRNA target sequences were cloned into pLKO.1-TRC (Addgene plasmid 10878), according to the manufacturer's suggestions: shRNA targeting MSH2 (shMSH2), 5′ CCCAUGGGCUAUCAACUUAAU; shRNA targeting MSH6 (shMSH6), 5′ AUCGCCAUUGUUCGAGAUUUA; shRNA targeting MLH1 (shMLH1), 5′ GUGGCUCAUGUUACUAUUACA; shRNA targeting MSH3 (shMSH3), 5′ UCGAGUCGAAAGGAUGGAUAA; and shRNA targeting PML (shPML), 5′ AGAUGCAGCUGUAUCCAAG (16). The sequence for shRNA targeting GFP (shGFP), 5′ GCAAGCUGACCCUGAAGUUCA, was previously described (48). Lentiviral particles were produced by transient transfection of pEAK cells with pLKO.1, psPAX2 (Addgene plasmid 12260), and pMD2.G (Addgene plasmid 12259) in a ratio of 4:3:1.

HSV-1 growth curves.

All growth curves were done with in1863 at a multiplicity of infection (MOI) of 0.1 PFU/cell. Virus yield was collected at 0, 6, 12, and 24 h postinfection and titrated on Vero cells by staining for β-galactosidase-positive plaques as previously described (48). HeLa and HFF-1 cells were infected with lentivirus, selected with 2 μg/ml puromycin, and infected with HSV-1 at 72 h post-lentiviral infection.

Immunofluorescence analysis.

Immunofluorescence (IF) analysis was performed as described with the following modifications (48). Briefly, cells adhered to glass coverslips were washed with phosphate-buffered saline (PBS), fixed with 4% paraformaldehyde, and permeabilized with either 1% Triton X-100 in PBS (Fix/Perm type 1) or with ice-cold acetone for 2 min at room temperature (Fix/Perm type 2). Cells were blocked with 3% normal goat serum and reacted with antibodies as indicated. Where indicated, cells were preextracted with cytoskeletal extraction buffer (CSK) on ice for 2 min prior to fixation to remove soluble proteins as previously described (12, 37). Primary antibodies included monoclonal mouse anti-ICP0 5H7 (1:200; East Coast Bio), monoclonal mouse anti-ICP4 (1:200; US Biological), monoclonal mouse anti-ICP8 11E2 (1:200; Abcam), polyclonal rabbit anti-ICP8 367 (1:400; provided by William Ruyechan, State University of New York, Buffalo, NY) (59), monoclonal mouse anti-MSH2 Ab-2 (1:100; Calbiochem), polyclonal rabbit anti-MSH6 (1:100; Santa Cruz), polyclonal rabbit anti-MSH3 H-300 (1:100; Santa Cruz), monoclonal mouse anti-MLH1 (1:200; BD Biosciences), and polyclonal rabbit anti-PML H-238 (1:200; Santa Cruz). TO-PRO-3 (1:1,000; Molecular Probes) was used as a nuclear counterstain. Alexa Fluor secondary antibodies (1:200; Molecular Probes) with fluorophores excitable at wavelengths of 488, 594, or 647 were used. For experiments shown in the figures (see Fig. 5C and 6B), all three antibodies were directly conjugated to fluorophores to allow the simultaneous use of mouse anti-ICP0 and anti-MLH1 (see Fig. 5C) or mouse anti-ICP4 and anti-MLH1 (see Fig. 6B). Conjugation was performed using the Zenon mouse IgG labeling kit (Molecular Probes) for ICP0, ICP4, and MLH1 and the Zenon rabbit IgG labeling kit (Molecular Probes) for PML, according to the manufacturer's suggested protocol. ICP0 and ICP4 were conjugated to Alexa Fluor 488, MLH1 was conjugated to Alexa Fluor 546, and PML was conjugated to Alexa Fluor 647. Images were captured using a Zeiss LSM 510 confocal nonlinear optics (NLO) microscope or a Zeiss LSM 780 confocal microscope equipped with argon and HeNe lasers and a Zeiss 63× objective lens (numerical aperture, 1.4). Images were processed and arranged using Adobe Photoshop CS3 and Illustrator CS3.

Fig. 5.

Punctate MLH1 localization requires intact ND10. HFF-1 cells were infected with lentiviruses expressing the indicated shRNA and selected with puromycin. (A, B) Localization of MLH1 and PML was determined by immunofluorescence analysis (A), and knockdown was determined by Western blotting (B). (C) HFF-1 cells were either mock infected or infected with d106, which is deleted for all IE genes except ICP0, at an MOI of 10 PFU/cell, fixed at 6 h postinfection, and stained for ICP0, MLH1, and PML. (D) HFF-1 cells were either mock infected or infected with HSV-1 (KOS), the ICP0-null virus 0β, or d106 at an MOI of 10 PFU/cell. Cells were harvested at 6 h postinfection for Western blot analysis as indicated. For the triple-label experiments shown in panel C, all antibodies were directly conjugated to fluorophores, as described in Materials and Methods, to allow for the simultaneous use of the monoclonal mouse antibodies for MLH1 and ICP0. All immunofluorescence samples were prepared using Fix/Perm type 1, and TO-PRO-3 was used as a nuclear counterstain in panel A.

Fig. 6.

MLH1 is recruited to the incoming viral genome. (A) HFF-1 cells were infected with vECFP-ICP4, which expresses a functional cyan fluorescent protein (CFP)-ICP4 fusion protein at an MOI of 0.01 PFU/cell, or the ICP0-null virus vEGdl110, which expresses GFP from the ICP0 promoter at an MOI of 1 PFU/cell. (B) HFF-1 cells were infected with lentiviruses expressing shRNA to either GFP or PML and selected with puromycin. Following selection, cells were infected with the ICP0-null virus dl1403 at an MOI of 0.5, fixed at 24 h postinfection, and stained for ICP4, MLH1, and PML. All cells were processed for immunofluorescence at 24 h postinfection using Fix/Perm type 1, as described in Materials and Methods, and stained for the indicated proteins. For the triple-label experiments shown in panel B, all antibodies were directly conjugated to fluorophores, as described in Materials and Methods, to allow for the simultaneous use of the monoclonal mouse antibodies for MLH1 and ICP4.

Western blot analysis.

Cells in 35-mm dishes were lysed in 2× SDS sample buffer (4% SDS, 20% glycerol, 100 mM Tris at pH 6.8, 100 mM dithiothreitol [DTT], 10% β-mercaptoethanol, 1× protease inhibitor cocktail [Roche], and 0.1% bromophenol blue). Alternatively, cells were collected by centrifugation and washed with PBS. Cells were then resuspended in standard RIPA buffer supplemented with protease inhibitors. Proteins were resolved by SDS-PAGE. Primary antibodies used included monoclonal mouse anti-ICP0 (1:1,000; East Coast Bio), monoclonal mouse anti-ICP4 (1:1,000; US Biological), monoclonal mouse anti-ICP8 11E2 (1:1,000; Abcam), polyclonal rabbit anti-UL12 BWp12 (1:10,000; provided by Joel Bronstein and Peter Weber [4, 44]), polyclonal goat anti-PML N19 (1:1,000; Santa Cruz), monoclonal mouse anti-β-actin (1:15,000; Sigma), polyclonal rabbit anti-β-actin (1:15,000; Sigma), monoclonal mouse anti-MSH2 (1:1,000; BD Biosciences), monoclonal mouse anti-MLH1 (1:1,000; BD Biosciences), polyclonal rabbit anti-MSH3 H-300 (1:1,000; Santa Cruz), polyclonal rabbit anti-MSH6 (1:1,000; Bethyl laboratories), monoclonal mouse anti-Ku70 (1:5,000; NeoMarkers), and monoclonal mouse anti-GFP (1:5,000; Zymed).

Immunoprecipitation analysis.

Cells were collected by centrifugation, washed with PBS, and then resuspended in standard RIPA buffer supplemented with protease inhibitors. Immunoprecipitation was performed overnight at 4°C using monoclonal mouse anti-MSH2 Ab-2 (5 μg; Calbiochem) or monoclonal mouse anti-MSH6 (5 μg; BD Biosciences).

Far-Western blot analysis.

Recombinant UL12, MSH2-MSH3, and MSH2-MSH6 were expressed and purified from insect cells as previously described (9, 53). Proteins were denatured and separated by SDS-PAGE and transferred to polyvinylidene difluoride (PVDF) membranes. After transfer, proteins were visualized by Coomassie blue staining. Then membranes were incubated with purified UL12 protein overnight at 4°C, washed extensively, and then blotted for UL12. Purified UL12 was included in all gels as a positive control for the Western blots. As a negative control for nonspecific binding of UL12 to any of the other proteins, membranes were probed directly with UL12 antibody prior to incubation with purified UL12.

Mismatch repair assay.

The heteroduplex MMR substrate was prepared as previously described (45). To assess MMR activity during HSV-1 infection, HeLa cells were infected with HSV-1 at an MOI of 2 PFU/cell and incubated for 3 h. Cells were then transfected with 1 μg of heteroduplex plasmid and 1 μg of red fluorescent protein (RFP)-expressing plasmid pDsRed2-N1 (Clontech) using Lipofectamine 2000. Following incubation for an additional 16 h, cells were harvested and analyzed for fluorescence intensity with a FACS LSRII-B flow cytometer (BD Biosciences) using BD FACSDiva software. GFP fluorescence was normalized to RFP expression to correct for differences in transfection efficiency.

RESULTS

Mismatch repair proteins are required for efficient HSV-1 replication.

To test whether MMR is required for HSV-1 replication, we utilized a lentivirus system to provide efficient delivery of shRNA targeting MSH2 and MLH1. We also included a control targeting a nonspecific protein, GFP. HeLa cells were infected with lentiviruses expressing shRNA and selected with puromycin, and knockdown was determined by Western blot analysis (Fig. 1A). Consistent with previous reports (43), shRNA-mediated knockdown of MSH2 resulted in depletion not only of MSH2 but also of the MSH2-interacting proteins MSH6 and MSH3 (Fig. 2A and data not shown). Knockdown of MLH1 resulted in a significant decrease in MLH1 levels but did not significantly affect levels of MSH2, MSH6, or MSH3 (Fig. 1A and data not shown). As expected, the shGFP control did not alter the levels of any of the proteins examined (Fig. 1A).

Fig. 1.

MSH2 is required for efficient HSV-1 replication in cancer cell lines. (A) HeLa cells were infected with lentiviruses expressing the indicated shRNA and selected with puromycin. After selection, knockdown was verified by Western blot analysis, and cells were infected with HSV-1 (in1863) at an MOI of 0.1 PFU/cell. (B, C) Isogenically matched cell lines Hec59 and Hec59+chromosome 2 (Hec59 +2) (B) or HCT-116 and HCT-116+chromosome 3 (HCT-116 +3) (C) were infected with HSV-1 at an MOI of 0.1 PFU/cell. For all growth curves, progeny virus was collected at the indicated times postinfection, and titers were determined on Vero cells. Values represent the averages of three independent experiments, and error bars represent the standard errors of the means.

Fig. 2.

MSH2 and MLH1 are required for efficient HSV-1 replication in normal human cells. HFF-1 cells were infected with lentiviruses expressing the indicated shRNA and selected with puromycin. (A to C) After selection, knockdown was verified by Western blot analysis (A), cells were infected with HSV-1 (in1863) at an MOI of 0.1 PFU/cell, progeny virus was collected at the indicated times, and titers were determined on Vero cells (B and C). The growth curves presented in panels B and C were done at the same time, and the shGFP values are the same in both graphs. Values represent the averages of three independent experiments, and error bars represent the standard errors of the means.

In order to determine whether MMR proteins are required for efficient HSV-1 growth, HeLa cells were infected with lentiviruses expressing shRNA to MSH2 and MLH1, selected with puromycin, and infected with HSV-1 at an MOI of 0.1 PFU/cell (Fig. 1A). Depletion of MSH2 with shRNA resulted in a 4- to 5-fold decrease in viral yield at both 12 and 24 h postinfection. Depletion of MLH1 with shRNA had no effect on viral yield at 12 and 24 h postinfection. Thus, in HSV-1-infected HeLa cells, knockdown of MSH2 has a more deleterious effect on HSV-1 growth than does knockdown of MLH1.

In order to further test the requirements of MSH2 and MLH1 in HSV-1 replication, we used two pairs of isogenic cell lines defective in MMR. Hec59 cells contain biallelic mutations in MSH2 and fail to make MSH2 protein; thus, they are deficient in MMR. The complemented cell line Hec59+chromosome 2 restores MSH2 expression and MMR (68). Efficient virus growth on Hec59 cells was delayed compared to the complemented cell line (Fig. 1B). After 24 h, however, virus growth on Hec59 cells recovered, resulting in final yields similar to those seen on Hec59+chromosome 2 cells, which expresses wild-type MSH2. To test the role of MLH1, we compared HSV-1 growth in HCT-116 cells that do not express MLH1 and are deficient in MMR to growth in the complemented cell line HCT-116+chromosome 3 in which MLH1 expression has been restored (27). Virus growth on HCT-116 cells closely paralleled the growth kinetics seen on HeLa cells (Fig. 1C). There was very little difference between the growth of HSV-1 on the parental and the complemented cell, with a slightly higher yield of virus on the complemented cell at the 24 h time point. Together, these data indicate a beneficial role for MSH2 in HSV-1 replication and a minimal role for MLH1 in HeLa or HCT-116 cells.

Because transformed cells contain mutations that have the potential to alter various repair pathways, we next asked whether MMR proteins play a positive or negative role during HSV-1 infection of normal diploid limited-passage human foreskin fibroblasts (HFF-1). Lentivirus was used to deliver shRNA targeting MSH2, MLH1, and GFP as described above. We also included controls specific for MSH6 and MSH3 to allow us to distinguish between the different MSH2 heterodimers. Transduced cells were selected with puromycin, and knockdown of targeted protein was verified by Western blot analysis (Fig. 2A). All knockdown cells generated in these experiments behaved similarly to the control cells in terms of growth and doubling times during the course of the experiments. To avoid problems associated with long-term knockdown of DNA repair proteins, we generated a new batch of lentiviral knockdown cells for each repeat of every experiment.

Consistent with the results obtained in HeLa cells treated with shRNA to MSH2 and the Hec59 cells, HFF-1 cells depleted of MSH2 resulted in a 10-fold decrease in virus yield (Fig. 2B). Surprisingly, HFF-1 cells depleted of MLH1 resulted in a similar decrease in virus yield, approximately 20-fold. This is in stark contrast to the apparently dispensable role of MLH1 in both HeLa cells and HCT-116 cells.

MSH2 is known to participate in two heterodimers, consisting of MSH2-MSH6 and MSH2-MSH3. The observation that MSH2 is required for efficient virus production prompted us to ask whether one or both of these binding partners were also important for HSV-1 growth. The silencing of MSH6 in HFF-1 cells resulted in a 20-fold decrease in viral yield, while virus growth in cells depleted of MSH3 resembled that in the control knockdown cells (Fig. 2C). A similar dependency on MSH2 and MSH6 was observed in HeLa cells (data not shown). Knockdown of MSH3 in HeLa cells resulted in a less severe viral growth defect than knockdown of either MSH2 or MSH6. Taken together, these data suggest that in both HFF-1 and HeLa cells, MSH2 and MSH6 are required for efficient HSV-1 replication, while MSH3 is not. MLH1 is necessary for efficient HSV-1 replication in HFF-1 but appears to be dispensable in certain transformed cells.

DNA mismatch repair protein interactions during HSV-1 infection.

Since HSV-1 infection results in the degradation of some components of the DNA repair machinery, we next asked whether MMR proteins are stable in HSV-1-infected cells. HeLa and HFF-1 cells were infected with HSV-1 at an MOI of 5 PFU/cell, and whole-cell extracts were prepared at the indicated times. MSH2, MSH6, MSH3, and MLH1 were stable in HSV-1-infected HeLa and HFF-1 cells, and the known ICP0 target, PML, was degraded efficiently during the same time points (Fig. 3A and B). The stability of MSH6 and MSH3 suggests proper formation of a heterodimer with MSH2, as these proteins depend upon MSH2 binding for their stabilization (43, 45). To confirm this, we examined the heterodimeric interactions between MSH2-MSH6 and MSH2-MSH3 during infection. Figure 3C shows that both MSH6 and MSH3 coimmunoprecipitated with MSH2 in mock- and HSV-1-infected cells.

Fig. 3.

Mismatch repair proteins are stable and interact during HSV-1 infection. HeLa (A) or HFF-1 (B) cells were infected with HSV-1 (KOS) at an MOI of 5 PFU/cell, and whole-cell extracts were collected at the indicated times postinfection. Western blotting was performed for the indicated MMR proteins, and actin was used as a loading control. (C and D) HeLa cells were infected with HSV-1 at an MOI of 5 PFU/cell and harvested at 6 h postinfection. Immunoprecipitation (IP) was performed from cell lysates using antibody against MSH2 (C) or MSH6 (D). Products were separated by SDS-PAGE and blotted with antibodies against MSH2, MSH6, MSH3, ICP8, and UL12 as indicated. (E) For far-Western blot analysis, purified UL12, MSH2-MSH3, MSH2-MSH6, and bovine serum albumin (BSA) were denatured, separated by SDS-PAGE, transferred to a PVDF membrane, and stained with Coomassie blue (left). The membrane was then destained and either probed directly with the anti-UL12 antibody (middle) or incubated with purified UL12 protein, washed, and then probed with the anti-UL12 antibody (right). Arrows indicate the molecular weights of the purified proteins.

We next asked whether viral proteins could be detected in complexes with MMR proteins. Figure 3D indicates that the viral ssDNA binding protein ICP8 can interact with MSH6, consistent with previous reports of an ICP8-MSH6 interaction (66); however, it is not clear whether this interaction is direct or indirect. Since ICP8 is also known to interact with the viral alkaline nuclease UL12 (66, 67), we tested whether UL12 was present in this complex. Figure 3D indicates that UL12 was precipitated by anti-MSH6 antisera. We have previously shown that UL12 interacts directly with the cellular MRN complex (2), and we wanted to see if UL12 could interact directly with MSH6. To test this, purified MSH2-MSH6 and MSH2-MSH3 complexes were analyzed for interaction with purified UL12 by far-Western blot analysis. UL12 was able to specifically interact with MSH6 and MSH3, though no direct interaction with MSH2 was detected (Fig. 3E). Thus, the interaction between MSH2-MSH6 with UL12 and ICP8 may be mediated by a direct interaction between MSH6 and UL12, though we cannot rule out direct interactions between ICP8 and the MMR proteins.

Mismatch repair proteins are recruited to HSV-1 replication compartments.

Since MMR proteins are important for efficient HSV-1 infection, we wanted to determine if they are recruited to replication compartments during infection. HFF-1 cells were infected with HSV-1 at an MOI of 2 PFU/cell and analyzed by immunofluorescence (IF) analysis at 6 h postinfection. MSH2 and MSH6 exhibited a nuclear diffuse staining pattern in uninfected cells and were partially reorganized in infected cells to colocalize with ICP8 in viral replication compartments (Fig. 4A). Consistent with these results in HFF-1 cells, we also observed that MSH6 exhibited a nuclear diffuse staining pattern in uninfected HeLa cells and was recruited to HSV-1 replication compartments (data not shown). These results are consistent with the previous finding that MSH2 could be detected in replication compartments (66). MSH3 exhibited a nuclear diffuse staining pattern in uninfected cells and remained in a nuclear diffuse pattern following infection. The lack of MSH3 recruitment to replication compartments is consistent with its dispensable role in the viral life cycle (Fig. 2C). We verified the specificity of the MSH2, MSH6, and MSH3 antibodies by demonstrating a loss of fluorescence intensity following infection with lentiviruses expressing shRNAs that specifically target these proteins (data not shown).

Fig. 4.

Localization of mismatch repair proteins during HSV-1 infection. HFF-1 cells were infected with HSV-1 (KOS) at an MOI of 2 PFU/cell, processed for immunofluorescence at 6 h postinfection using Fix/Perm type 2, as described in Materials and Methods, and stained for MSH2, MSH6, MSH3, and ICP8 (A) or MLH1, PML, and ICP8 (B). The ICP0-null virus used was 0β.

MLH1 is an ND10 component in HFF-1 cells.

We were surprised to observe that in uninfected HFF-1 cells, MLH1 was present in a punctate nuclear staining pattern reminiscent of ND10. This pattern is different than the pattern observed in HeLa cells in which MLH1 exhibits a nuclear diffuse staining pattern with a few nuclear foci (46). Several cellular proteins, including PML, sp100, ATRX, and hDaxx, are known to localize in ND10, and many of these have been shown to have antiviral properties (15, 16, 39). To test whether MLH1 colocalized with ND10, HFF-1 cells were costained with antisera to MLH1 and PML. The complete colocalization of MLH1 and PML suggests that MLH1 is in fact an ND10 component in HFF-1 cells (Fig. 4B). Many ND10 proteins are recruited to incoming viral genomes, forming ND10-like foci which are subsequently disrupted by the degradation of PML and other targets of the viral E3 ubiquitin ligase ICP0 (22, 47); however, we showed that MLH1 was not degraded following infection (Fig. 3A and B). Thus, the fate of MLH1 after infection was of interest. Figure 4B shows that MLH1 colocalizes with ICP8 in replication compartments, consistent with its positive role in viral infection. MLH1 was also recruited to replication compartments in infected HeLa cells (data not shown). In the absence of ICP0, however, PML is not degraded, ND10 proteins are not disrupted, and the viral genome remains associated with ND10 bodies. In cells infected with the ICP0-null virus, MLH1 remained in ND10 proteins, some of which are associated with the developing replication compartments (Fig. 4B).

To further test whether MLH1 is indeed an ND10 component, we chose to silence PML and monitor the localization of MLH1. PML is the central organizer of ND10, and many ND10 components become nuclear diffuse or have altered localization in the absence of PML (16). Figure 5 demonstrates that PML was successfully silenced in shPML cells, as no PML protein was detectable by IF (Fig. 5A, middle) or Western blotting (Fig. 5B). TO-PRO-3 dye was used to detect cellular DNA and mark the boundary of the nucleus. In control shGFP cells, MLH1 and PML colocalized; however, in cells expressing shPML, MLH1 was observed in a nuclear diffuse staining pattern (Fig. 5A, middle). Although MLH1 staining in shPML cells appeared less intense than that in uninfected cells (Fig. 5A, middle), this was likely due to the diffuse localization and not a result of degradation, as no decrease was observed in the total amount of MLH1 in shPML cells by Western blotting (Fig. 5B). Figure 5B indicates that in shMLH1 cells, MLH1 has been silenced efficiently. PML levels were also decreased in these cells. This may suggest that MLH1 can stabilize PML; however, additional experiments will be required to test this hypothesis. In shMLH1 cells, PML foci remained intact (Fig. 5A). However, as the knockdown of MLH1 was not complete, we cannot rule out that the remaining MLH1 is sufficient to stabilize these ND10 bodies.

As stated previously, ICP0 is sufficient to degrade PML and disrupt ND10. We wanted to determine if ICP0 was sufficient to reorganize MLH1. To test this, HFF-1 cells were either mock infected or infected with the mutant virus d106. d106 is deleted for all IE genes except ICP0 and therefore is a useful tool for overexpression of ICP0. In mock-infected cells, PML and MLH1 colocalized in ND10, and no ICP0 was detectable (Fig. 5C). Following infection with d106, large ICP0 nuclear inclusions were formed, PML was degraded, and MLH1 was redistributed to a nuclear diffuse staining pattern with a few small foci. We also observed that ICP0 was sufficient to alter the cellular localization of MLH1 in a transient-transfection assay with ICP0 and that this activity was dependent on the E3-ubiquitin ligase activity of ICP0 (data not shown). To confirm that MLH1 levels remained constant, we infected HFF-1 cells with wild-type virus, the ICP0-null virus, or d106. PML was efficiently degraded during infection with wild-type virus and d106 but was not degraded during infection with the ICP0-null virus (Fig. 5D). Furthermore, the levels of MLH1 and MSH2 do not change during infection with any of the three viruses. Thus, ICP0 expression alone is sufficient to redistribute MLH1 and does not alter the levels of MLH1. Together, these data suggest that MLH1 is a bona fide ND10 component and that disruption of ND10 is required for efficient MLH1 reorganization to replication compartments. Thus, disruption of ND10 not only may play a role in relieving the silencing properties of PML and sp100 but also may function to release positive factors such as MLH1 for recruitment to replication compartments.

MLH1 is recruited to the incoming viral genome prior to MSH2.

As described above, during infection with the wild-type virus, ICP0 degrades PML and other ND10 components such as sp100; however, in the absence of ICP0, PML and sp100 are not degraded and remain associated with the incoming genome. This can be most easily visualized at the edge of a developing plaque, where asymmetric ICP4 foci in the nucleus represent the incoming viral genome in newly infected cells (14, 17). Using this assay, we asked whether MLH1 is recruited to the incoming viral genome, as would be expected for other ND10 proteins. HFF-1 cells were infected with HSV-1 vECFP-ICP4 (labeled HSV-1) and ICP0-null HSV-1 vEGdl110 (labeled ΔICP0) at MOIs of 0.01 and 1 PFU/cell, respectively. These dilutions were chosen to optimize the edge of the plaque assay. The vECFP-ICP4 virus makes a functional enhanced cyan fluorescent protein (ECFP)-ICP4 fusion protein from both copies of the ICP4 gene (18). In uninfected cells, MLH1 and PML colocalized in ND10 (Fig. 6A, top panel). Following infection with vECFP-ICP4 (labeled HSV-1), PML was completely degraded, and MLH1 colocalized to some extent with ECFP-ICP4, presumably on the viral genome (Fig. 6A, second panel). Not all of the MLH1 was recruited to the incoming genome, as some remained nuclear diffuse. We also analyzed the location of MLH1 at the edge of a developing plaque in cells infected with the ICP0-null virus vEGdl110, which expresses GFP in place of ICP0 under the control of the ICP0 promoter (38). Under these conditions, PML was observed in asymmetric foci at the edge of the developing plaque that represent the incoming genome (Fig. 6A, third panel). MLH1 completely colocalized with PML in these cells. These data demonstrate that MLH1 behaves as a ND10 component and is reorganized to the incoming genome into ND10-like foci. Of interest is the observation that MLH1 remained associated with the incoming genome even when other ND10 components, such as PML, were degraded. MSH2, however, was not recruited to the incoming genome at the edge of a developing plaque during infection with either the wild-type or ICP0-null virus (Fig. 6A, fourth and bottom panels). This is surprising, as recruitment of MLH1 to mismatched or damaged DNA is generally thought to depend on prior MSH2 recruitment (1, 46). The observation that MLH1 was recruited to the viral genome while MSH2 was not may indicate a novel function for MLH1 that is independent of MSH2.

Since we have identified MLH1 as a novel ND10 component and MLH1 appears to be recruited to incoming genomes more efficiently during infection with the ICP0-null virus, we wanted to determine if PML itself was contributing to the recruitment of MLH1. To test this, HFF-1 cells were transduced with lentiviruses expressing shGFP or shPML and subsequently infected with the ICP0-null virus at an MOI of 0.5 PFU/cell. Cells were fixed at 24 h postinfection and stained for ICP4, MLH1, and PML. In shGFP cells, both MLH1 and PML were efficiently recruited to the incoming genome (Fig. 6B, top). In shPML cells, we observed a significant reduction in the amount of PML, yet MLH1 was still efficiently recruited to the incoming genome (Fig. 6B, bottom). Therefore, PML is not required for MLH1 recruitment to the incoming genome.

Several prereplicative structures leading to the formation of replication compartments have been described by our group and others (5, 36, 37, 40). They are characterized by an ordered addition of viral and cellular proteins (48, 52, 71). The earliest detectable prereplicative sites have been called stage II microfoci, which contain ICP8, the origin binding protein UL9, and the three-subunit helicase-primase complex, UL5/UL8/UL52, as well as the cellular proteins RPA and ATRIP (37, 48). Stage IIIa prereplicative sites have a composition similar to that of stage II microfoci but are slightly larger. Stage IIIb prereplicative sites are characterized by the presence of the polymerase and the polymerase accessory protein and form under conditions in which the viral polymerase is inhibited with PAA. HFF-1 cells were infected with HSV-1 at an MOI of 2 PFU/cell in the presence and absence of PAA and examined for the localization of MSH2 (Fig. 7A) and MLH1 (Fig. 7B). In the absence of PAA, large replication compartments formed, and MSH2 and MLH1 colocalized with ICP8, consistent with the data shown in Fig. 4. In contrast, in the presence of PAA, small punctate ICP8 foci, representing stage IIIb prereplicative sites, that did not colocalize with MSH2 were visible, which remained nuclear diffuse. In addition, MSH2 did not colocalize with stage II or stage IIIa prereplicative sites (data not shown). Interestingly, during infection in the presence of PAA, MLH1 was detectable in sites adjacent to and partially overlapping with ICP8 (Fig. 7B). Thus, it appears that MSH2 and MLH1 are recruited to replication compartments via two different mechanisms. MLH1 recruitment occurs prior to viral DNA replication and may occur via direct interactions with the viral genome, while MSH2 recruitment requires viral DNA synthesis.

Fig. 7.

MSH2 is not recruited to HSV-1 prereplicative sites. HFF-1 cells were infected with HSV-1 (KOS) at an MOI of 2 PFU/cell in the presence or absence of PAA. Cells were processed for immunofluorescence analysis using Fix/Perm type 2, as described in Materials and Methods, at 6 h postinfection and stained with ICP8 and MSH2 (A) or MLH1 (B) antibodies.

MSH2 and MLH1 have separate roles in the HSV-1 life cycle.

Lentiviral depletion of MSH2 and MLH1 from HFF-1 cells resulted in similar reductions in viral yields (10- and 20-fold, respectively). However, MLH1 was recruited to incoming viral genomes, while recruitment of MSH2 to the replication compartments required viral DNA synthesis. To more specifically define the defects in HSV-1 replication following knockdown of these two proteins (Fig. 8A, top), we infected HFF-1 cells expressing shMSH2 or shMLH1 with HSV-1 at an MOI of 2 PFU/cell and monitored expression of the IE and E gene products ICP4 and ICP8, respectively. Figure 8A shows that shMSH2 cells exhibited a slight reduction in ICP4 expression combined with a delay in ICP8 expression. A similar defect was observed in HSV-1 gene expression in HeLa cells transfected with siRNA targeting MSH2 (data not shown). Surprisingly, shMLH1 cells showed a more dramatic reduction in ICP4 expression and undetectable levels of ICP8 by Western blotting at 6 h postinfection. In order to further define the difference between shMSH2 and shMLH1 cells, we monitored the intranuclear localization of ICP4 and ICP8 in infected cells at 6 h postinfection. Figure 8B shows that control shGFP cells exhibited large replication compartments that costained for ICP4 and ICP8 (80% of cells). shMSH2 cells showed markedly smaller replication compartments, as indicated by ICP8 staining, that represent bona fide replication compartments, as ICP4 was recruited to them (64% of cells). Consistent with the HSV-1 gene expression profiles, little to no ICP8 was detectable in foci in the shMLH1 cells, and only very small ICP4 foci were detectable (65% of cells) (Fig. 8B). Figure 8C shows the quantification of these data. Taken together with the gene expression data, it appears that MSH2 and MLH1 participate in distinct stages of the viral life cycle, with MLH1 depletion acting at an earlier stage than MSH2 depletion.

Fig. 8.

MSH2 and MLH1 participate in separate events in the HSV-1 life cycle. HFF-1 cells were infected with lentiviruses expressing the indicated shRNA and selected with puromycin. (A) Knockdown was verified by Western blotting, and cells were infected with HSV-1 (KOS) at an MOI of 2 PFU/cell, harvested at the indicated times postinfection, and analyzed by Western blotting for ICP4 and ICP8. (B and C) Cells were infected with HSV-1 at an MOI of 2 PFU/cell and fixed at 6 h postinfection using Fix/Perm type 1, as described in Materials and Methods. (B) Cells were analyzed by immunofluorescence with ICP4 and ICP8 antibodies. (C) Cells were scored for progression of infection by ICP4 staining patterns. Representative images of each condition are shown in panel B; shGFP shows large replication compartments (RC), shMSH2 shows small RC, and shMLH1 shows ICP4 foci. The values represent greater than 250 cells per condition counted between 2 independent experiments.

DNA repair and damage response functions of the MSH2-MSH6 heterodimer are not disrupted during HSV-1 infection.

We next wished to test whether the cellular functions of the DNA MMR pathway remain intact during HSV-1 infection. An intact DNA MMR pathway is required for activation and downstream response to the DNA-alkylating agent N-methyl-N′-nitro-N-nitrosoguanidine (MNNG) (62). We have previously demonstrated that DNA alkylation damage results in the localization of MMR proteins to the chromatin (46). To examine if MMR proteins were able to respond to DNA damage during HSV-1 infection, HFF-1 cells were infected with HSV-1 at an MOI of 5 PFU/cell and, at 4 h postinfection, were treated with 10 μM MNNG for 2 h. Insoluble protein populations were examined by treatment of cells with detergent prior to fixation. MSH2 colocalized with ICP8 in viral replication compartments in untreated cells; however, this population of MSH2 remained soluble, as it was completely removed by detergent extraction (Fig. 9A). In contrast, treatment with MNNG resulted in the accumulation of insoluble MSH2, which remained in viral replication compartments following extraction (Fig. 9A). These data suggest that MMR proteins were recruited to damaged viral DNA and that the first step in the response to DNA alkylation damage remained intact.

Fig. 9.

MSH2-MSH6 functions, including the response to alkylation damage and DNA repair, are intact during HSV-1 infection. (A) HFF-1 cells were infected with HSV-1 (KOS) at an MOI of 5 PFU/cell. At 2 h postinfection, cells were pretreated with O⁶-benzylguanine, and at 4 h postinfection, 10 μM MNNG was added. Cells were processed for immunofluorescence analysis at 6 h postinfection using Fix/Perm type 2, as described in Materials and Methods, and stained for MSH2 and ICP8. Where indicated, cells were pretreated with detergent prior to fixation to extract soluble proteins. (B and C) HeLa cells were infected with HSV-1 at an MOI of 2 PFU/cell. At 3 h postinfection, cells were transfected with a heteroduplex plasmid encoding EGFP with a premature stop codon, which can be corrected by cellular MMR proteins, and RFP as a transfection control. At 24 h postinfection, cells were harvested and analyzed for EGFP expression by flow cytometry (B) or Western blot analysis (C). EGFP expression was normalized to RFP expression. The values represents the averages of two independent experiments done in triplicate, and the error bars represent the standard errors of the means.

In addition to activating a DNA checkpoint response, the DNA MMR pathway repairs a variety of DNA lesions, including single-base-pair mismatches. To examine repair in infected cells, we performed an in vivo repair assay that utilizes an EGFP reporter with a premature stop codon conferred by a single nucleotide mismatch. Efficient repair of the mismatch results in EGFP expression, which can be monitored by flow cytometry (73). We infected HeLa cells with HSV-1 at an MOI of 2 PFU/cell and, at 3 h postinfection, transfected cells with the EGFP reporter. Both mock- and HSV-1-infected cells demonstrated equal and robust repair, suggesting that this pathway remained functional during HSV-1 infection and that infection does not enhance repair activity (Fig. 9B). Infection was confirmed by Western blot analysis of ICP8 expression (Fig. 9C).

DISCUSSION

Mismatch repair proteins are required for efficient growth on normal human cells.

In this study, we investigated the interaction of herpes simplex virus 1 (HSV-1) with the cellular mismatch repair (MMR) pathway. MSH2, MSH6, and MLH1 were shown to localize to replication compartments (Fig. 4). The localization of MSH2 in replication compartments was consistent with a previous report from Taylor and Knipe (66) and raised the possibility that components of the MMR pathway are required for efficient replication of HSV-1. We have now shown that MSH2 is required for efficient viral growth in both human foreskin fibroblasts (HFF-1) and HeLa cells. Interestingly, in HFF-1 cells, depletion of MLH1 had a result similar to that of depletion of MSH2; however, virus growth was not significantly compromised in HeLa cells depleted of MLH1 or in HCT-116 cells lacking MLH1. The requirement for MLH1 in HFF-1 cells but not immortalized cells may suggest that MLH1 plays a previously unrecognized role distinct from its known role downstream of MSH2-MSH6 recognition of mismatched bases. It is possible that the cancer cells tested provide an alternate factor that can substitute for MLH1 or that they provide a cellular environment in which MLH1 is no longer required for efficient HSV-1 growth.

MSH2 is known to participate in the repair of single nucleotide mismatches and insertion/deletion loops (IDLs) via two distinct heterodimers, MSH2-MSH6 or MSH2-MSH3, respectively. The data shown in Fig. 2 indicated that functions of the MSH2-MSH6 heterodimer are important for productive HSV-1 infection. We also showed that MSH2 can bind damaged DNA in viral replication compartments and that single-base-pair mismatches were efficiently repaired in HSV-1-infected cells. Together these experiments indicated that MMR is functional in HSV-1-infected cells. It will be of interest to determine whether MMR proteins contribute to the fidelity of HSV-1 DNA replication. It is known that while the HSV-1 DNA polymerase possesses 3′-to-5′ exonuclease activity (26, 42), it exhibits a higher misincorporation rate than many other viral and cellular replicative polymerases (6, 21). Alternatively, it is possible that the MSH2-MSH6 heterodimer may play a role in regulation of recombination, since the MSH2-MSH6 heterodimer has been shown to have antirecombinogenic functions (11). Although we and others have proposed that homologous recombination is important during HSV-1 DNA replication (72), many questions remain about how recombination is regulated. Additional experiments will be required to determine whether the role of MSH2-MSH6 in HSV-1 infection is related to prevention of mutations or regulation of recombination at dissimilar sequences.

We have also confirmed the observation that ICP8 interacts directly or indirectly with the MMR protein MSH6 by coimmunoprecipitation and have extended this observation by showing that UL12 is also present in this complex and that UL12 is able to specifically interact with MSH6. We have previously proposed that ICP8 and UL12 function as a viral recombinase important for the homologous recombination observed in infected cells (54). Furthermore, UL12 is able to directly interact with the host MRN (Mre11/Rad50/Nbs1) complex (2), which is required for cellular homologous recombination (65). It will be of interest to determine if the MSH2-MSH6 interaction with UL12 functions to regulate homologous recombination during HSV-1 infection.

MMR proteins have been implicated in the life cycles of other DNA viruses. MSH2, MSH6, and MLH1 were detected in replication compartments induced by Epstein-Barr virus (10). MSH2, MSH6, and MSH3 were also detected in adeno-associated virus (AAV) replication compartments when HSV-1 was used as the helper virus but not when adenovirus was used as a helper (49). UL12 and ICP8 are also recruited to the AAV replication compartments, and it is possible that they are responsible for the recruitment of MMR proteins (49). Thus, it appears that several DNA viruses interact with components of the MMR pathway, and one report suggests that these proteins may be playing a proofreading role following viral DNA replication (64). Our results are consistent with these observations but highlight an even earlier role in the viral life cycle for MLH1, as MLH1 is recruited to the viral genome prior to initiation of replication and is required for efficient IE gene expression.

MLH1 is a novel ND10 protein.

We observed MLH1 staining in bright nuclear foci in HFF-1 cells, consistent with a previous report that MLH1 localizes to punctate foci in human embryonic lung fibroblasts (51). We demonstrated here that these foci also costained for PML, indicating that MLH1 is a previously unrecognized component of ND10 structures. When the central organizing protein of ND10, PML, is depleted in primary human fibroblasts, the location of other ND10 proteins such as sp100 and hDaxx becomes diffuse (16). In this report we show that MLH1 behaved in a similar fashion. We also show that ICP0 expression is sufficient to alter the cellular localization of MLH1 from ND10 to nuclear diffuse, presumably due to the degradation of PML. Also consistent with our identification of MLH1 as a bona fide ND10 component, MLH1 has been shown to interact with another ND10 protein, the Bloom Syndrome helicase BLM (30, 51). Furthermore, MLH1 has a predicted SUMO interaction motif, another hallmark of ND10 proteins (3). It has been suggested that ND10 structures may serve as reservoirs for stress response proteins that reorganize in response to a variety of cellular insults (29). Consistent with this idea, ND10 structures have been shown to disperse following alkylation damage (8), which likely allows for MLH1 recruitment to sites of DNA damage (46).

The ND10 proteins PML, sp100, hDaxx, and ATRX have been characterized extensively during HSV-1 infection, and they all appear to contribute to repression of the incoming viral genome (15, 16, 39). Interestingly, although MLH1 is recruited to the incoming viral genome like other ND10 proteins and this recruitment is enhanced in the absence of ICP0, a subset of MLH1 remains associated with the viral genome even after ND10 proteins are disrupted during wild-type infection. We also show that MLH1 recruitment to the incoming genome is independent of PML, as it was recruited efficiently in cells depleted of PML. Unlike other ND10 proteins that play antiviral roles, MLH1 is required for viral gene expression, and viral growth is inhibited when MLH1 is depleted. MLH1 may bind the incoming viral genome directly, as MLH1 has some affinity for both double- and single-stranded DNA (19). Alternatively, it may associate with viral genomes indirectly. Taken together, our findings indicate that ND10 may not function solely to inhibit viral infection. ND10 may also serve positive functions for viral replication by acting as a reservoir for beneficial proteins, which promote gene expression and DNA replication. This situation may be reminiscent of another ND10-associated protein, the histone acetyltransferase CLOCK, that has been reported to play a positive role in viral gene expression (24, 25).

MSH2 and MLH1 play separable roles in HSV-1 replication.

Depletion of MSH2 and MLH1 from HFF-1 cells results in similar decreases in virus yield (10- and 20-fold, respectively); however, analysis of the block in the virus life cycle revealed that MLH1 acts at a step prior to that of MSH2. The most striking defect in MLH1-depleted cells is reduced IE gene expression and a failure to make E gene products. Furthermore, MLH1 is recruited to the incoming genome, while MSH2 is not. Lastly, we observed that MLH1 is required in normal human cells but not in any of the human cancer cell lines tested, whereas MSH2 is required in both the normal and transformed lines. Together, these data suggest that MLH1 plays an important function in the viral genome required for IE gene expression in normal cells and that this role is independent of MSH2. This result is surprising in light of the reported functions for MLH1 downstream of MSH2. As the incoming viral genome is a linear DNA molecule which contains nicks and gaps (55), it is possible that DNA repair is required prior to viral gene expression and that this repair is mediated by an MLH1-dependent mechanism. One intriguing hypothesis is the recent identification of MLH1 as an interacting partner of FAN1 (Fanconi-associated nuclease 1), which participates in interstrand cross-link repair (ICL) (28, 61). ICL proteins have been studied mostly in response to chemical cross-linkers such as mitomycin C (7) and have not been studied in the context of HSV-1 infection. It will be of significant interest to both the HSV-1 and cellular DNA repair fields to exploit the possible separation of function of MLH1 from MSH2 and its impact on DNA repair and gene expression.