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Journal of Virology, June 2005, p. 7162-7171, Vol. 79, No. 11
0022-538X/05/$08.00+0     doi:10.1128/JVI.79.11.7162-7171.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Inhibition of the Herpes Simplex Virus Type 1 DNA Polymerase Induces Hyperphosphorylation of Replication Protein A and Its Accumulation at S-Phase-Specific Sites of DNA Damage during Infection

Dianna E. Wilkinson and Sandra K. Weller*

Department of Molecular, Microbial and Structural Biology, University of Connecticut Health Center, Farmington, Connecticut 06030

Received 15 December 2004/ Accepted 1 February 2005


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ABSTRACT
 
The treatment of mammalian cells with genotoxic substances can trigger DNA damage responses that include the hyperphosphorylation of replication protein A (RPA), a protein that plays key roles in the recognition, signaling, and repair of damaged DNA. We have previously reported that in the presence of a viral polymerase inhibitor, herpes simplex virus type 1 (HSV-1) infection induces the hyperphosphorylation of RPA (D. E. Wilkinson and S. K. Weller, J. Virol. 78:4783-4796, 2004). We initiated the present study to further characterize this genotoxic response to HSV-1 infection. Here we report that infection in the presence of polymerase inhibitors triggers an S-phase-specific response to DNA damage, as demonstrated by induction of the hyperphosphorylation of RPA and its accumulation within viral foci specific to the S phase of the cell cycle. This DNA damage response occurred in the presence of viral polymerase inhibitors and required the HSV-1 polymerase holoenzyme as well as the viral single-stranded-DNA binding protein. Treatment with an inhibitor of the viral helicase-primase did not induce the hyperphosphorylation of RPA or its accumulation in infected cells. Taken together, these results suggest that the S-phase-specific DNA damage response to infection is dependent on the specific inhibition of the polymerase. Finally, RPA hyperphosphorylation was not induced during productive infection, indicating that active viral replication does not trigger this potentially detrimental stress response.


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INTRODUCTION
 
In response to agents that cause DNA damage or replication stress, mammalian cells activate signal transduction pathways that slow cell cycle progression and repair the damaged DNA. If the damage is irreparable, cells are eliminated through the induction of apoptosis. Defects in this stress response can compromise genomic stability, resulting in transformation and a predisposition to cancer (reviewed in references 28 and 42).

One of the early responders to DNA damage is replication protein A (RPA), a heterotrimeric single-stranded-DNA (ssDNA) binding protein consisting of 70-, 32-, and 14-kDa subunits (reviewed in reference 3). During an unperturbed cell cycle, RPA is associated with replication forks throughout S phase (12). Under DNA-damaging conditions, sites of DNA breaks or stalled replication forks generate stretches of ssDNA to which RPA binds. When bound to stretches of ssDNA, RPA undergoes a conformational change that results in hyperphosphorylation of the middle subunit (RPA32). The coating of hyperphosphorylated RPA at stretches of ssDNA exposed by stalled cellular forks or DNA damage is known to serve as a signal for DNA damage and to recruit proteins that participate in the repair of damaged DNA (reviewed in reference 3).

We have recently reported that in the presence of the viral polymerase inhibitor phosphonoacetic acid (PAA), herpes simplex virus type I (HSV-1) induces the hyperphosphorylation of RPA32. This DNA damage response appears to be specific to the inhibition of the viral polymerase since the hyperphosphorylation of RPA32 was not observed during productive infection or during infection with a polymerase null virus (50). We initiated the present study to further define this host stress response to HSV-1 infection.

HSV-1 encodes the following seven proteins that are essential for the replication of its genome: the origin-binding protein (UL9), the ssDNA-binding protein (UL29 or ICP8), the helicase-primase heterotrimer (UL5, UL8, and UL52), the viral polymerase (UL30), and its processivity subunit (UL42) (reviewed in references 48 and 51). Replication of the HSV-1 linear double-stranded DNA (dsDNA) genome occurs in the nucleus of the infected cell within globular domains called replication compartments (38). In addition to the seven essential viral replication proteins, cellular proteins that participate in DNA metabolism, including RPA, are also present in replication compartments (45, 46, 49, 50). We have shown that RPA and the recombination and repair proteins RAD51 and NBS1 are recruited to replication compartments and viral foci believed to be intermediates in the formation of replication compartments, consistent with the proposal that these proteins play a role in the viral life cycle (50).

If HSV-1 DNA replication is prevented by inhibiting the viral polymerase or infecting cells with a polymerase null virus, UL29 localizes to punctate foci called prereplicative sites (38). Two types of prereplicative sites have been described based on UL29 staining patterns (31, 46). Some infected cells contain few prereplicative sites (<20 UL29 foci per cell), while others contain numerous sites (50 to 200 UL29 foci per cell). The few prereplicative sites (called stage IIIa foci when formed in the absence of HSV-1 polymerase or stage IIIb foci when formed in the presence of an inhibited viral polymerase [5, 8]) form adjacent to nuclear structures called ND10 sites (31, 46) and are believed to be precursors to the formation of replication compartments (31). Numerous prereplicative sites, on the other hand, are not specifically associated with ND10 but instead colocalize with incorporated bromodeoxyuridine (BrdU), suggesting that they mark S-phase-dependent sites of ongoing DNA synthesis or repair (11, 31, 46). Since viral DNA synthesis is prevented under these conditions, we have proposed that these numerous prereplicative sites are not actual precursors of replication compartments, but instead represent areas of UL29 accumulation at ssDNA exposed at stalled or collapsed cellular replication forks (31). In this study, we show that infection in the presence of polymerase inhibitors triggers an S-phase-specific response to DNA damage, as demonstrated by the induction of the hyperphosphorylation of RPA and its accumulation at sites of DNA damage within numerous prereplicative sites. These results support our suggestion that numerous prereplicative sites generated under these conditions represent stalled or collapsed cellular replication forks. This DNA damage response, however, was not seen during infection with a polymerase null virus, indicating that viral polymerase is required for the PAA-induced DNA damage response to HSV-1 infection. Furthermore, although low levels of endogenous hyperphosphorylated RPA could be detected in cells infected in the absence of polymerase inhibitors, this isoform of RPA did not localize in either replication compartments or stage III prereplicative sites, which are the actual precursors of replication compartments. This endogenous, uninduced population of hyperphosphorylated RPA was, instead, localized away from sites relevant to viral DNA synthesis. These findings suggest that viral replication forks are not recognized by the cell as signals of DNA damage or replication stress. Thus, productive infection appears to avoid activating a host stress response that may otherwise be detrimental to viral genome replication.


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MATERIALS AND METHODS
 
Cell lines. Vero cells were obtained from the American Type Culture Collection (Manassas, Va.) and were grown in Dulbecco modified Eagle's medium (Invitrogen, Carlsbad, Calif.) supplemented with 5% fetal bovine serum (Gemini Bio-Products, Woodland, Calif.), penicillin, streptomycin, and amphotericin B (Invitrogen).

Reagents and antibodies. Camptothecin (CPT), glycerol gelatin, 1,4-diazobicyclo-[2.2.2]octane (DABCO), and the viral polymerase inhibitors PAA and acyclovir (ACV) were obtained from Sigma Chemical Co. (St. Louis, Mo.). The viral helicase-primase inhibitor BAY 57-1293 (16, 25, 26) was kindly provided by Gerald Kleymann (Bad Salzuflen, Germany).

Mouse monoclonal anti-UL29 (39S) and anti-ICP4 (58S) antibodies (44) were obtained from the American Type Culture Collection. A mouse monoclonal antibody against RPA32 (9H8) (15) was supplied by Marc Wold (University of Iowa). A phosphospecific rabbit polyclonal anti-phospho-RPA32 pSer4/pSer8 antibody (BL647) was purchased from Bethyl Laboratories, Inc. (Montgomery, Tex.). Secondary antibodies conjugated to Alexa Fluor 488 or Alexa Fluor 594 were purchased from Molecular Probes (Eugene, Oreg.).

Viruses and infections. Strain KOS was used as wild-type HSV-1. The UL29 deletion mutant virus HD2 (17) was kindly provided by David Knipe (Harvard Medical School). Mutants with ICP6::lacZ insertions in the helicase-primase genes for UL5 (hr99), UL8 (hr80), and UL52 (hr114) and the origin binding protein UL9 (hr94) were previously described (6, 19, 33, 53). A UL42 deletion mutant virus, Cgal{Delta}42 (24), was provided by Deborah Parris (Ohio State University). A polymerase null virus, HP66 (35), was obtained from Charles Hwang (Upstate Medical University). The polymerase mutant viruses PAAr5, {Delta}S1.1, 7E4A, and 6C4, which contain changes within the catalytic subunit of polymerase (10, 18, 35), were provided by Donald Coen (Harvard Medical School). The temperature-sensitive polymerase mutant viruses tsC4 and tsC7 (10) were provided by Priscilla Schaffer (Harvard Medical School). Two double mutants were also used in this study: they are 66/99, which is a chimera of HP66 and hr99R345K (a replication-defective UL5 helicase motif mutant virus), and RK/94, which is a chimera of hr99R345K and hr94 (31, 53). For infections, asynchronous, subconfluent cells were adsorbed for 1 h with 10 PFU per cell and incubated for 5.5 to 6.0 h postadsorption. Mock infections were carried out in parallel. Where indicated, 400 µg PAA per ml, 100 µM ACV, or 100 µM BAY 57-1293 was added at the time of adsorption and maintained throughout the course of infection.

Indirect immunofluorescence. An in situ extraction method that removes cytosolic and nucleosolic proteins was employed for the visualization of chromatin-bound or matrix-associated proteins. Cells were preextracted for 5 min on ice with cytoskeletal buffer as described previously (12) and then fixed in 4% paraformaldehyde for 10 min. Fixed cells were incubated with primary antibodies diluted in 3% normal goat serum (NGS) for at least 30 min. The 39S and BL647 antibodies were used at a concentration of 1:200. After extensive washing with phosphate-buffered saline (PBS), the cells were incubated for 30 min with secondary antibodies diluted in 3% NGS at a concentration of 1:200. After a final wash with PBS, coverslips were mounted in glycerol gelatin containing 2.5% DABCO to retard photobleaching.

Microscopy. Cells that were double-stained for immunofluorescence (IF) were examined under a Zeiss LSM 410 confocal microscope as described previously (50). Alexa Fluor 488 was excited at 488 nm, while Alexa Fluor 594 was excited at 568 nm. Appropriate emission filters were employed and channels were scanned individually using settings established with control slides. Channels were overlaid by a computer to create merged images. As controls, samples were stained with one primary antibody and the relevant secondary antibodies. No overlap between the optical channels was observed for any of the samples at the settings used. The collected images were arranged with Adobe Photoshop 7.0.

Western analysis. Protein expression was examined by Western blot analysis. Asynchronized cells were grown in 100-mm plates to 70% confluence. Cells were either mock infected or infected with wild-type or mutant HSV-1 at a multiplicity of infection of 10. When indicated, CPT, PAA, ACV, or BAY 57-1293 was added to the culture medium at the time of adsorption. At 7 h postinfection, whole-cell extracts were subjected to electrophoresis in sodium dodecyl sulfate-10% polyacrylamide gels and prepared for immunodetection as described previously (50). Primary antibodies in TBST (0.2% [vol/vol] Tween 20 in PBS) were used at a 1:3,000 dilution. After several washes in TBST, membranes were incubated with alkaline phosphatase-conjugated secondary antibodies at a dilution of 1:10,000 for 1 to 2 h and developed with alkaline phosphatase color detection according to the manufacturer's instructions (Promega, Madison, Wis.).


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RESULTS
 
HSV-1 infection in the presence of PAA induces an S-phase-specific response to DNA damage. RPA is known to associate with replication forks during an unperturbed S phase as well as to mark sites of DNA damage. Hyperphosphorylated RPA, however, is observed at sites of stalled replication forks or DNA damage and is not present at unperturbed replication forks (4, 13, 27, 39, 47). RPA colocalizes with UL29 in replication compartments, stage IIIb prereplicative sites, and numerous prereplicative sites during HSV-1 infection (46, 49, 50). Due to their inherent ability to bind ssDNA, the colocalization of RPA and UL29 may reflect the presence of ssDNA at these sites. Alternatively, the colocalization of RPA and UL29 may be based on interactions between these two proteins. Preliminary evidence for such an interaction was recently reported (45). The recruitment of RPA to sites of viral DNA synthesis during productive infection and of UL29 to sites of cellular DNA synthesis raises an interesting question as to how these ssDNA-binding proteins are recruited to viral and/or cellular replication forks.

The anti-phospho-RPA32 Ser4/Ser8 antibody (BL647) specifically recognizes hyperphosphorylated forms of RPA32 that are modified at serine 4 and serine 8 and has been used as a marker for genotoxic stress in human cells (39, 47). To determine whether BL647 could recognize similar signals in infected Vero cells, we compared BL647 with another antibody which detects all isoforms of RPA32 (9H8). Whole-cell lysates from Vero cells that were infected with wild-type virus or the polymerase null virus, HP66, in the absence or presence of PAA were examined by Western blot analysis (Fig. 1). The 9H8 antibody, which predominantly detected non- or hypophosphorylated isoforms of RPA32, also detected the hyperphosphorylated isoforms of RPA32 induced during infection in the presence of PAA (Fig. 1A, left panel). On the other hand, BL647 was specific in detecting only the hyperphosphorylated forms of RPA32 (Fig. 1A, right panel). Although one isoform of hyperphosphorylated RPA32 was detected at a basal level in all lysates probed with BL647, the amount of this isoform was greatly increased when the viral polymerase was inhibited with PAA (Fig. 1A, right panel, open arrow). Furthermore, a second hyperphosphorylated species detected by BL647 (Fig. 1A, right panel, closed arrow) was observed only in the presence of PAA. These results confirm our previous finding that, when treated with PAA, HSV-1 infection induces the hyperphosphorylation of RPA32 (Fig. 1A, KOS + PAA) (50).



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  		FIG. 1. (A) Rabbit anti-phospho RPA32 Ser4/Ser8 antibody BL647 recognizes only hyperphosphorylated forms of RPA32 in Vero cells. In the presence or absence of PAA, Vero cells were either mock infected or infected with either KOS or the polymerase null virus HP66. Lysates were prepared at 7 h postinfection. Aliquots of mock-, KOS-, and HP66-infected cell lysates were loaded onto an SDS-10% polyacrylamide gel in duplicate, separated by a molecular weight marker (M). After electrophoretic transfer of the samples, the membrane was cut down the middle of the marker lane. The left half of the membrane was probed with mouse anti-RPA32 (9H8), and the right half was probed with the rabbit anti-phospho RPA Ser4/Ser8 antibody (BL647). After immunodetection, the two membranes were aligned and the different migrating species of RPA32 were identified. Note that BL647 recognized only the hyperphosphorylated forms of RPA32 (open and closed arrows) and that wild-type infection in the presence of PAA induced their formation. (B) Merged images showing subcellular distribution of RPA and hyperphosphorylated RPA in Vero cells in the absence and presence of exogenously induced DNA damage. Cells were either left untreated or treated with camptothecin (CPT). As described in Materials and Methods, cells were extracted in situ and double-labeled with the general anti-RPA32 antibody 9H8 (green) and the phosphospecific anti-phospho RPA32 Ser4/Ser8 antibody BL647 (red). Yellow indicates colocalization of the two proteins. The endogenous forms of hyperphosphorylated RPA detected by BL647 in untreated cells likely represent the uninduced forms seen in the Western blot of mock-infected cells (panel A, open arrow).

The 9H8 and BL647 antibodies were also characterized by IF microscopy to determine whether they marked sites of damaged DNA in Vero cells treated with CPT, an inhibitor of cellular topoisomerase I that is known to cause DNA breaks (23) (Fig. 1B). Vero cells grown in the presence or absence of CPT were preextracted with Triton X-100 to remove soluble nuclear proteins, leaving behind chromatin-bound and matrix-associated proteins (12). The two untreated cells shown in Fig. 1B (left panel) displayed typical staining patterns of asynchronous cells that were doubly stained with the general 9H8 antibody (green) and the phosphospecific BL647 antibody (red). In untreated cells, BL647 detected endogenous hyperphosphorylated RPA that was distributed in a slightly rough granular pattern within the nucleus at sites distinct from presumed cellular replication forks marked by the isoforms of RPA detected with 9H8 (Fig. 1B, left panel). In cells that were apparently in S phase, hundreds of foci detected by 9H8 were observed which did not contain hyperphosphorylated RPA (Fig. 1B, left panel, top cell), consistent with previous reports that hyperphosphorylated forms of RPA do not localize to sites of ongoing DNA replication (47). In contrast, treatment of Vero cells with CPT induced increased staining of hyperphosphorylated forms of RPA and its accumulation with RPA into punctate foci which likely represent sites of DNA repair (Fig. 1B, right panel).

We next used the phosphospecific anti-RPA32 antibody to characterize the subcellular response to DNA damage during HSV-1 infection. Vero cells were infected with wild-type virus or the polymerase null virus, HP66, in the absence or presence of PAA. At 7 h postinfection, cells were prepared for IF microscopy and labeled with the phosphospecific BL647 antibody and a monoclonal anti-UL29 antibody (39S) (Fig. 2). Previous reports have described the efficient recruitment of RPA to replication compartments (46, 49, 50). In stark contrast, however, hyperphosphorylated RPA was not present in replication compartments, but instead was detected in smaller nuclear domains juxtaposed to the replication compartments (Fig. 2A to C). In mock-infected cells, hyperphosphorylated RPA was distributed in a granular pattern within the nucleus (Fig. 2D to F). The staining pattern of hyperphosphorylated RPA in mock- and KOS-infected cells probably represents the uninduced, endogenous isoform detected with the BL647 antibody via Western analysis (Fig. 1A, right panel, open arrow). The hyperphosphorylation of RPA32 was not induced during productive infection; furthermore, endogenous hyperphosphorylated RPA was found to be spatially distinct from the sites of ongoing viral DNA synthesis. Based on these two observations, we suggest that HSV-1 has evolved a mechanism to replicate its genome without activating a host stress response that may be deleterious to viral infection. The mechanism by which HSV-1 excludes hyperphosphorylation from sites of viral DNA synthesis remains to be determined. We also found that endogenous hyperphosphorylated RPA did not localize with UL29 in any of the infected cell populations displaying less numerous, stage IIIa (n = 299) (Fig. 2P to R and V to X) or stage IIIb prereplicative sites (n = 197) (Fig. 2J to L), which are considered to be precursors of replication compartments. Thus, neither replication compartments nor the less numerous prereplicative sites (stage IIIa and stage IIIb) represent sites of DNA damage as defined by the accumulation of hyperphosphorylated RPA.



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  		FIG. 2. Hyperphosphorylated RPA32 (P-RPA32) accumulates and colocalizes with UL29 only within numerous prereplicative sites generated under conditions in which the viral polymerase is inhibited. Vero cells were mock infected (D to F), infected with KOS either in the absence (A to C) or presence (G to L) of PAA, or infected with the polymerase null virus HP66 either in the absence (M to R) or presence (S to X) of PAA. For the detection of chromatin-bound or matrix-associated proteins, cells were extracted with Triton X-100 prior to fixation with paraformaldehyde as described in Materials and Methods. Cells were double-labeled with the mouse anti-UL29 antibody 39S (green) and the rabbit anti-phospho RPA32 Ser4/Ser8 antibody BL647 (red). Merged images are shown, with yellow indicating the colocalization of P-RPA with UL29. Viral foci examined included replication compartments (A to C), numerous prereplicative sites (G to I, M to O, and S to U), and stage III prereplicative sites (J to L, P to R, and V to X). Using BrdU labeling and IF microscopy, we confirmed that the numerous prereplicative sites presented in this report were S phase specific (11, 30-32, 46), while stage IIIa/IIIb prereplicative sites were S phase independent (31, 46) (data not shown).

We also asked whether hyperphosphorylated RPA was present in infected cells displaying S-phase-dependent numerous prereplicative sites. Approximately 77% of cells displaying numerous prereplicative sites formed in the presence of PAA exhibited an accumulation of hyperphosphorylated RPA which colocalized with UL29 (n = 263) (Fig. 2G to I). Notably, on the other hand, no increased staining of hyperphosphorylated RPA was ever observed within numerous prereplicative sites formed in cells infected with HP66 (i.e., in the absence of the viral polymerase) (n = 297) (Fig. 2M to O and S to U); furthermore, the hyperphosphorylated RPA that was observed in these cells did not colocalize with UL29. Thus, numerous prereplicative sites formed in the presence of an inhibited polymerase likely represent sites of DNA damage, while the numerous prereplicative sites formed in the absence of the viral polymerase do not. These results directly implicate the inhibited viral polymerase in the induction of an S-phase-specific response to DNA damage. It is known that cellular replication fork stalling and resulting replication stress can activate DNA damage responses, including the hyperphosphorylation of RPA and its redistribution to sites of DNA damage (39, 47). We propose that the DNA damage noted for this infected cell population (Fig. 2G to I) is due to a genotoxic effect of an inhibited polymerase on cellular forks.

The HSV-1 polymerase holoenzyme is required for hyperphosphorylation of RPA. To further define the requirements for the induction of RPA hyperphosphorylation during infection, we performed Western blot analysis on lysates from cells infected with different HSV-1 polymerase mutants, both in the presence and in the absence of PAA (Fig. 3). Three general types of polymerase mutant virus were examined: (i) nonviable polymerase mutants that make polymerase protein detectable by Western analysis (6C4 as well as tsC4 at 39.5°C and tsC7 at 39.5°C) (10, 35), (ii) nonviable polymerase mutants that do not make detectable protein (HP66, {Delta}S1.1, and 7E4a1) (35), and (iii) a viable polymerase mutant which is resistant to PAA (PAAr5) (10). Cell extracts were probed with the 9H8 antibody, which detects both unphosphorylated and phosphorylated forms of RPA32. To control for infection, we also probed the blots with a monoclonal antibody directed against ICP4 (58S) (Fig. 3). No induction of RPA hyperphosphorylation was noted in mock-infected cells or cells infected in the absence of PAA (Fig. 3). Furthermore, no induction of RPA hyperphosphorylation was observed in the presence of PAA for any of the nonviable polymerase mutants, regardless of whether the polymerase was present (Fig. 3, tsC4 at 39.5°C, tsC7 at 39.5°C, 6C4, {Delta}S1.1, 7E4a1, and HP66). The only conditions that were shown to induce the hyperphosphorylation of RPA32 were infections in which a functional polymerase was inhibited with PAA (Fig. 3, KOS + PAA at 34°C, 37°C, and 39.5°C as well as tsC4 + PAA at 34°C and tsC7 + PAA at 34°C). In order to rule out the possibility that the damage response was caused by a nonspecific effect of PAA on infected cells, we took advantage of a viral mutant whose polymerase has been shown to be resistant to PAA, namely, PAAr5 (10). Interestingly, PAAr5 infection did not induce the hyperphosphorylation of RPA32 in the presence of PAA (Fig. 3, PAAr5 + PAA), indicating that the effects of PAA during viral infection reflect the specific inhibition of the HSV polymerase and not toxic effects on other viral or cellular proteins. This result supports the hypothesis that this DNA damage response is not triggered when viral DNA synthesis is allowed to proceed. Taken together, these results indicate that during infection, an inhibited but otherwise functional viral polymerase is required for the induction of this DNA damage response.



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  		FIG. 3. RPA32 phosphorylation in cells infected with HSV-1 mutants defective in the polymerase or the polymerase accessory subunit. Infections were carried out either in the presence or in the absence of PAA. Lysates were harvested at 7 h postinfection and prepared for immunoblotting as described in Materials and Methods. Membranes were probed with the 9H8 antibody to detect all isoforms of RPA32. Black arrows indicate the hyperphosphorylated form of RPA32. As a control for the induction of RPA32 hyperphosphorylation, mock-infected cells were also treated with 1 µM camptothecin (CPT). To monitor infection, we also probed the membranes with the mouse anti-ICP4 antibody 58S. Results for lysates from mock-infected cells (mock), wild-type HSV-1 (KOS)-infected cells, and polymerase null mutant (HP66)-infected cells have been reported previously (50) and are included here as controls. The viable HSV-1 viruses examined were KOS, tsC4 at 34°C, tsC7 at 34°C, and the PAA-resistant virus PAAr5. Nonviable polymerase mutant viruses included HP66, tsC4 at 39.5°C, tsC7 at 39.5°C, 6C4, {Delta}S1.1, and 7E4a1. We also examined the polymerase accessory protein mutant virus Cgal{Delta}UL42.

We also determined whether the polymerase accessory subunit, UL42, played any role in inducing this stress response during infection. We found that the UL42-defective mutant Cgal{Delta}42 was unable to induce the hyperphosphorylation of RPA in the presence of PAA (Fig. 3), indicating that both the polymerase catalytic subunit, UL30, and its accessory protein, UL42, are necessary for this PAA-induced DNA damage response to infection.

Requirement of other HSV-1 replication proteins for induction of RPA32 hyperphosphorylation. We next examined HSV-1 mutants defective in each of the other five essential replication proteins for the ability to induce a DNA damage response. Figure 4 shows that RPA hyperphosphorylation was not induced in response to infection with any of the mutants in the absence of PAA. Interestingly, however, infections with HSV-1 mutants bearing a defect in the origin-binding protein (Fig. 4, hr94 + PAA) or in any of the members of the helicase-primase heterotrimer (Fig. 4, hr99 + PAA, hr80 + PAA, and hr114 + PAA) were still able to induce the hyperphosphorylation of RPA, provided that PAA was also present. These findings suggest that RPA hyperphosphorylation can be induced even though viral DNA synthesis is prevented, provided that an inhibited viral polymerase is still present. This finding was further supported by our observation that an HSV-1 replication-defective double mutant that lacks both viral polymerase and replicative helicase activity could not induce the hyperphosphorylation of RPA in the presence of PAA (Fig. 4, 66/99 + PAA), while another double mutant lacking functional helicase and origin-binding activity (yet encoding a functional polymerase) could induce this effect provided that PAA was also present (Fig. 4, RK/99 + PAA). Finally, we show that the UL29-defective mutant virus was unable to induce the hyperphosphorylation of RPA in the presence of the polymerase inhibitor (Fig. 4, HD2 + PAA). These results indicate that the ability to induce the damage response requires an inhibited viral polymerase holoenzyme as well as the viral ssDNA-binding protein. The origin-binding protein and the helicase-primase heterotrimer, on the other hand, are not required for the induction of this stress response.



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  		FIG. 4. RPA32 phosphorylation in cells infected with HSV-1 replication mutants. Infections were carried out either in the presence or in the absence of PAA and were prepared for Western blot analysis as described in the legend to Fig. 3. The black arrow indicates the hyperphosphorylated form of RPA32. The defect of each mutant is provided in parentheses.

PAA induces an S-phase-specific response to DNA damage in cells infected with a primase-defective virus. We were intrigued by the finding that infections with viruses deficient in origin-binding or helicase-primase activity were able to induce the hyperphosphorylation of RPA32 when PAA was present (Fig. 4). The ability to induce this DNA damage response correlates well with results of previous reports describing the ability of these mutants to form numerous prereplicative sites in the presence of a polymerase inhibitor (30, 32). This suggests that the inhibitor-induced numerous sites generated in these mutants also represent sites of stalled cellular forks or DNA damage. To further explore this correlation, we assessed the subcellular response to DNA damage during infection with a primase-defective virus, hr114 (Fig. 5). In the absence of PAA, all of the hr114-infected cells showed granular nuclear staining for UL29 which did not colocalize with endogenous hyperphosphorylated RPA (Fig. 5A), suggesting that these cells were not undergoing a DNA damage response. In the presence of the polymerase inhibitor, however, two populations of hr114-infected cells were observed: cells with granular UL29 staining which did not accumulate hyperphosphorylated RPA, as noted above (Fig. 5C), and the pattern of numerous prereplicative sites (Fig. 5B) which is specific to the S phase of the cell cycle (30, 32). The accumulation of hyperphosphorylated RPA and its localization with UL29 were observed in approximately 72% of hr114-infected cells displaying numerous prereplicative sites (n = 149) (Fig. 5B). These findings provide further evidence that cells infected under conditions that inhibit the viral polymerase undergo an S-phase-specific response to DNA damage and that the numerous prereplicative sites formed under these conditions mark DNA lesions present at sites of stalled or collapsed cellular replication forks.



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  		FIG. 5. Inhibition of the viral polymerase induces an S-phase-specific response to DNA damage in cells infected with the primase-defective mutant virus hr114. Vero cells were infected with hr114 either in the absence (A) or in the presence (B and C) of PAA. Cells were extracted with Triton X-100 prior to fixation with paraformaldehyde as described in Materials and Methods. Cells were double labeled with the mouse anti-UL29 antibody 39S (green) and the rabbit anti-phospho RPA32 Ser4/Ser8 antibody BL647 (P-RPA; red). Merged images are shown, with yellow indicating the accumulation of P-RPA with UL29, which was observed only in numerous prereplicative sites (B). We previously reported that hr114-infected cells exhibit a diffuse plus granular pattern of UL29 staining (32). The preextraction method used here (A and C) removes the diffusely staining UL29 population.

Analysis of host DNA damage response in the presence of other inhibitors of viral DNA synthesis. In our studies so far, the induction of RPA hyperphosphorylation and its accumulation in S-phase-specific sites of DNA damage had been observed only when the viral polymerase was inhibited with PAA. To determine whether this genotoxic response was dependent on the specific inhibition of the viral polymerase or was due to the pharmacological inhibition of viral replication proteins in general, we examined cells infected with wild-type virus in the presence of either ACV or BAY 57-1293. ACV is a nucleoside analog that inhibits the viral polymerase by a mechanism different from that of PAA, which is a nonnucleoside inhibitor of polymerase (9, 29, 34). BAY 57-1293, on the other hand, is a nonnucleoside compound that inhibits the ATPase and primase activities of the helicase-primase complex (26). A Western blot analysis of cells infected in the presence of ACV indicated that this polymerase inhibitor could also induce the hyperphosphorylation of RPA (Fig. 6). Furthermore, IF microscopic analysis of these cells showed that hyperphosphorylated RPA accumulated specifically within approximately 76% of the infected cell population displaying S-phase-dependent numerous prereplicative sites (n = 111) (Fig. 7A) and not within cells displaying less numerous prereplicative sites (n = 80) (Fig. 7B). In contrast, treatment of infection with BAY 57-1293 did not induce the hyperphosphorylation of RPA (Fig. 6), suggesting that the pharmacological inhibition of viral DNA replication via inactivation of the viral helicase-primase is not an inducer of the host response to DNA damage. IF microscopic analysis also indicated that cells infected in the presence of BAY 57-1293 failed to form replication compartments (Fig. 7C). This result is consistent with the inhibition of viral DNA replication in the presence of this compound (26). Finally, the endogenous population of hyperphosphorylated RPA was never found to localize with UL29 in infected cells that had been treated with BAY 57-1293 (n = 100) (Fig. 7C). We therefore concluded that the induction of the DNA damage response is not a general result of inhibiting HSV-1 DNA replication but instead requires specific inhibition of the viral polymerase.



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  		FIG. 6. RPA32 phosphorylation in cells infected with HSV-1 in the presence of pharmacological inhibitors of viral DNA replication. Vero cells were either mock infected (mock) or infected with wild-type HSV-1 (KOS) in the presence of the polymerase inhibitor ACV or the helicase-primase inhibitor BAY 57-1293. Western blot analysis was performed as described in the legend to Fig. 3. The black arrow indicates hyperphosphorylated RPA32.



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  		FIG. 7. ACV induces an S-phase-specific response to DNA damage during HSV-1 infection, while BAY 57-1293 does not. Vero cells were infected with wild-type HSV-1 (KOS) in the presence of the polymerase inhibitor ACV (A and B) or the helicase-primase inhibitor BAY 57-1293 (C). Cells were extracted with Triton X-100 prior to fixation with paraformaldehyde as described in Materials and Methods and were double-labeled with the mouse anti-UL29 antibody 39S (green) and the rabbit anti-phospho RPA32 Ser4/Ser8 antibody BL647 (P-RPA; red). Merged images are shown, with yellow indicating an accumulation of P-RPA with UL29, which was observed only in numerous prereplicative sites induced by ACV (A).

Taken together, these results suggest that numerous prereplicative sites generated under conditions which specifically inhibit the viral polymerase represent sites of DNA damage. We further propose that during infection, the inhibition of the viral polymerase is genotoxic to cellular DNA replication. This results in stress at cellular replication forks, leading to the induction of an S-phase-specific DNA damage response. Conversely, sites of active viral DNA replication (replication compartments) and their precursors (stage IIIa/b prereplicative sites) are not recognized by the cell as sites of genotoxic stress.


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DISCUSSION
 
The coating of hyperphosphorylated RPA on stretches of ssDNA exposed by stalled cellular forks or DNA damage is known to serve as a signal for cellular DNA damage and to mark nuclear foci where DNA repair is occurring (39, 47; reviewed in reference 3). We previously reported that RPA32 is hyperphosphorylated during wild-type HSV-1 infection in the presence of the viral polymerase inhibitor PAA (50). For the present study, IF microscopy and Western blot analyses were used to further characterize the induction of this cellular response to DNA damage during HSV-1 infection. Several observations were made, as follows: (i) inhibition of the HSV-1 polymerase during infection induced the hyperphosphorylation of RPA and its accumulation within numerous prereplicative sites, indicating an S-phase-specific host response to DNA damage; (ii) infections with mutant viruses demonstrated that in addition to an inhibited viral polymerase, the polymerase accessory subunit and viral ssDNA-binding protein were required for the induction of this DNA damage response but that neither the origin-binding protein nor the three-component helicase-primase was required; (iii) this genotoxic response was also observed with infections in the presence of another polymerase inhibitor, ACV, but not in the presence of BAY 57-1293, a specific inhibitor of the helicase-primase; (iv) endogenous hyperphosphorylated RPA did not colocalize with UL29 in either replication compartments or stage IIIa/IIIb prereplicative sites, which are the earliest precursors to replication compartments, suggesting that these viral assemblies are not recognized by the cell as sites of DNA damage or replication stress. Taken together, these findings directly implicate the inhibited viral polymerase as an agent that induces an S-phase-specific host response to DNA damage during HSV-1 infection. Furthermore, a productive infection appears not to trigger this stress response; instead, endogenous hyperphosphorylated RPA is located adjacent to replication compartments.

Inhibition of HSV-1 polymerase induces DNA damage at cellular replication forks. During HSV-1 infection, we observed the induction of RPA hyperphosphorylation only when the viral polymerase holoenzyme was inhibited. Furthermore, the accumulation of hyperphosphorylated RPA at repair centers was observed only within the infected cell population bearing numerous prereplicative sites. Since numerous prereplicative sites are specific to the S phase of the cell cycle (11, 30-32, 46; our unpublished results), we propose that these cells were undergoing an S-phase-specific DNA damage response to replication stress induced by an inhibited polymerase at cellular replication forks. The presence of an exogenous polymerase at cellular forks is surprising since the recruitment of proteins for replication is a very ordered process (14). In the context of viral DNA replication, a sequential recruitment of viral proteins has also been observed: UL29 and the helicase-primase heterotrimer are thought to form a scaffold at the viral fork and to recruit other viral proteins (5, 8, 30). The recruitment of HSV-1 polymerase to the viral scaffold appears to require an active primase, implying that the presence of a primer at the viral fork may be a prerequisite for polymerase recruitment (8). Since PAA- or ACV-induced numerous sites are able to form in cells infected with helicase-primase mutant viruses (30-32), the recruitment of viral polymerase to cellular forks is not dependent on the presence of the viral helicase-primase as it is at viral replication forks. It is possible that the laying down of RNA primers by the host polymerase {alpha}-primase at cellular forks (14) provides a signal for the recruitment of viral polymerase to cellular DNA. We further speculate that the presence of an inhibited viral polymerase at the cellular fork perturbs the cellular replication machinery and generates stretches of ssDNA at which RPA and UL29 accumulate. The binding of RPA to stretches of ssDNA which arise at stalled forks or DNA breaks is thought to cause a conformational change within RPA32, making it a better substrate for hyperphosphorylation (reviewed in reference 3). DNA damage-dependent signal transduction pathways are subsequently activated, and components of the repair machinery are recruited to the numerous prereplicative sites. The observation that an S-phase-specific damage response could also be induced by treating a wild-type infection with ACV, another inhibitor of the polymerase, but not with BAY 57-1293, which targets the helicase-primase (Fig. 6 and 7), provided further evidence that the S-phase-specific response to DNA damage is dependent on the inhibition of the viral polymerase.

This study demonstrated that neither the helicase-primase nor the origin-binding protein is necessary for the PAA-induced hyperphosphorylation of RPA32. The conditions under which we observed the induction of RPA32 hyperphosphorylation by these mutants (i.e., only when the viral polymerase was inhibited with PAA) correlate well with the occurrence of numerous prereplicative sites formed in cells infected with these mutants in the presence of viral polymerase inhibitors (30-32). To explore this correlation further, we used a primase-defective virus to demonstrate that hyperphosphorylated RPA accumulated specifically within the PAA-induced numerous prereplicative sites. This finding lends further support to the hypothesis that the presence of an inhibited viral polymerase at cellular forks may be genotoxic to the cell. Based on these findings, we propose that the PAA-induced numerous prereplicative sites seen in UL9, UL5, or UL8 mutant-infected cells (30-32) are also S-phase-specific repair foci resulting from an inhibited polymerase at cellular forks.

We have demonstrated that the pharmacological inhibition of HSV-1 polymerase during infection is genotoxic to the cell. We argue that caution should be used when interpreting experiments in which PAA or ACV is used to inhibit viral DNA replication. A method that does not rely on the pharmacological inhibition of the viral polymerase may serve as a more appropriate DNA-replication-negative control when assessing the host DNA damage response to HSV-1 infection.

Numerous prereplicative sites generated during infection in the absence of a polymerase holoenzyme are not sites of DNA damage. HP66, a polymerase null virus, failed to induce the hyperphosphorylation of RPA. Although the numerous prereplicative sites observed in HP66-infected cells resembled the numerous prereplicative sites induced by an inhibited viral polymerase, the sites formed in the absence of polymerase never accumulated hyperphosphorylated RPA, indicating that they were not sites of DNA damage. We propose that the presence of UL29 (or other viral proteins) at cellular forks does not cause genotoxic stress per se. The numerous foci formed in the absence of viral polymerase may simply represent UL29 localization at active cellular forks, as shown by colabeling with BrdU (32; our unpublished results) but not with hyperphosphorylated RPA (Fig. 1). Alternatively, cellular replication forks in cells infected in the absence of a functional viral polymerase may be stressed but the ssDNA generated at these sites may be below the threshold level required for the induction of a DNA damage response. Taking these results together, we propose that the numerous sites reported for infection with a polymerase accessory protein mutant virus or a temperature-sensitive polymerase mutant virus at the nonpermissive temperature (20, 30) are not sites of DNA damage but simply reflect the localization of UL29 to cellular replication centers.

Productive HSV-1 infection does not trigger replication stress but does induce other host responses to DNA damage. We have demonstrated that ongoing viral DNA synthesis with an active viral polymerase does not induce RPA32 hyperphosphorylation and that endogenous hyperphosphorylated forms of RPA are localized away from replication compartments. It will be interesting to determine whether the localization of potentially damaging signals away from replication compartments is part of a global mechanism by which HSV avoids triggering stress signals which could be deleterious to the replicating viral genome.

We have shown that NBS1, a component of the MRE11-RAD50-NBS1 (MRN) repair complex (7), is phosphorylated during productive HSV-1 infection, indicating that at least one component of the DNA damage response is activated during ongoing viral DNA synthesis (50). We envision that the active replication of viral DNA may avoid triggering one host response to DNA damage (i.e., the accumulation of hyperphosphorylated RPA) while co-opting another through the activation of the MRN repair complex. Replication intermediates that arise during viral DNA synthesis are complex, possibly branched, structures, indicating a role of recombination or repair during HSV-1 replication (2, 36, 40, 41, 43; reviewed in reference 51). We proposed that the MRN complex may play a role in generating or processing these intermediates (50). We also noted previously that NBS1 phosphorylation occurs during infection when the viral polymerase is inhibited (50). Thus, the MRN complex may serve two separate roles during HSV-1 infection, with one involving replication-dependent recombination of viral DNA intermediates and the other responding to cellular replication stress induced by an inhibited viral polymerase.

The roles of the MRN complex and RPA in HSV-1 infection will need to be studied in the larger context of the signaling pathways that are triggered by DNA breaks. Upstream regulators of DNA damage responses include the stress-related protein kinases ATM (ataxia telangiectasia mutated) and ATR (ATM and Rad3 related). These stress-related kinases serve as transducers of the DNA damage signal by phosphorylating and activating downstream molecules that either regulate or effect DNA repair (reviewed in references 28 and 42). ATM is predominately activated by dsDNA breaks, while ATR is also activated by RPA-bound ssDNA (1, 21, 22, 37, 52). Based on these different mechanisms of activation, we anticipate that ATM will have a major role during productive HSV-1 DNA replication, which has a high potential for generating dsDNA breaks, while ATR will likely regulate the host response to DNA damage arising from an inhibited viral polymerase at cellular forks. Unraveling the different cellular responses to DNA damage observed during HSV-1 infection will contribute to our understanding of both the cellular and viral processes of replication, recombination, and repair.


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ACKNOWLEDGMENTS
 
We thank Ellen Fanning and the members of our laboratory for helpful comments. We also thank Gerald Kleymann for supplying BAY 57-1293 and for helpful comments.

This research was supported by Public Health Service grant AI21747. D.E.W. was supported by NIH training grant F32AI054042.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Molecular, Microbial and Structural Biology, MC3205, University of Connecticut Health Center, 263 Farmington Ave., Farmington, CT 06030. Phone: (860) 679-2310. Fax: (860) 679-1239. E-mail: Weller{at}NSO2.uchc.edu. Back


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Journal of Virology, June 2005, p. 7162-7171, Vol. 79, No. 11
0022-538X/05/$08.00+0     doi:10.1128/JVI.79.11.7162-7171.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.



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