The Products of the Herpes Simplex Virus Type 1 Im
http://www.100md.com
病菌学杂志 2006年第8期
Departments of Medicine and Microbiology and Molecular Genetics, Harvard Medical School at the Beth Israel Deaconess Medical Center, Boston, Massachusetts 02215
ABSTRACT
Herpes simplex virus type 1 ICP22–/US1.5– mutants initiate viral gene expression in all cells; however, in most cell types, the replication process stalls due to an inability to express 2 late proteins. Although the function of ICP22/US1.5 has not been established, it has been suggested that these proteins activate, induce, or repress the activity of cellular proteins during infection. In this study, we hypothesized that cell cycle-associated proteins are targets of ICP22/US1.5. For this purpose, we first isolated and characterized an ICP22–/US1.5– mutant virus, 22/n199. Like other ICP22–/US1.5– mutants, 22/n199 replicates in a cell-type-specific manner and fails to induce efficient 2 late gene expression in restrictive cells. Although synchronization of restrictive human embryonic lung cells in each phase of the cell cycle did not overcome the growth restrictions of 22/n199, synchronization of permissive Vero cells in S phase rendered them less able to support 22/n199 plaque formation and replication. Consistent with this finding, expression of cellular S-phase cyclins was altered in an ICP22/US1.5-dependent manner specifically when S-phase Vero cells were infected. Collectively, these observations support the notion that ICP22/US1.5 deregulates the cell cycle upon infection of S-phase permissive cells by altering expression of key cell cycle regulatory proteins either directly or indirectly.
INTRODUCTION
Many DNA-containing viruses alter cell cycle regulatory pathways to produce an environment that favors efficient viral replication. DNA viruses with small genomes do not carry the full complement of proteins required for viral DNA synthesis but utilize the cellular DNA synthesis machinery present during S phase to replicate their genomes. To access this machinery, parvoviruses depend on spontaneous progression of infected cells into S phase (43), whereas polyoma- and papillomaviruses contain proteins (large T antigen and E7, respectively) that drive and arrest cells in S phase (17, 20). Although DNA viruses with larger genomes, such as adeno- and herpesviruses, carry essential viral DNA replication proteins, these viruses also require selected S-phase activities for efficient replication (47, 48). During adenovirus infection, E1A drives cells and arrests them in S phase to free the cellular S-phase transcription factor, E2F, from the retinoblastoma protein (pRb) (46, 47). This activity is required for the transcription of adenovirus DNA replication proteins (7). Unlike the small DNA viruses and adenovirus, however, herpesviruses do not utilize machinery expressed exclusively during normal S phase for viral DNA synthesis, gene expression, or replication. Rather, they deregulate the cell cycle to access and utilize cellular activities normally expressed during G1 and S phases (21).
Herpes simplex virus type 1 (HSV-1) induces profound changes in cellular growth regulation pathways in infected cells. Studies of infected asynchronous cells and cells synchronized in each stage of the cell cycle have revealed that these alterations are dynamic and produce an environment that is unlike any phase of the normal cell cycle. At very early times postinfection (<2 h postinfection [hpi]) in cells in any phase of the cell cycle, an "S-phase-like" environment is created; however, cellular DNA synthesis does not occur in these cells as it would in normal S-phase cells. In these "S-phase-like" cells, the key S-phase transcription factor, E2F, is freed from hyperphosphorylated pRb (28), and expression of at least a subset of S-phase-specific genes is induced. At later times (>2 hpi) in cells infected during any stage of the cell cycle, cells appear to enter a "G1-like" phase. In contrast to "S-phase-like" cells in which pRb is hyperphosphorylated and E2F is freed, in "G1-like" cells, pRb is hypophosphorylated and E2F is found in complexes with pRb (21, 42). The ability of HSV-1 to activate S-phase genes very early in infection, while constraining expression of cell cycle-associated activities to a "G1-like" phase later in infection, independent of the phase of the cell cycle at the time of infection, suggests that HSV-1 infection alters the regulation of multiple cell cycle-associated activities.
Alteration of cell cycle machinery in HSV-1-infected cells is mediated in large part by the immediate-early (IE) regulatory proteins, infected cell protein 0 (ICP0), ICP27, ICP4, and ICP22. ICP0, when expressed alone, inhibits cellular DNA synthesis and arrests cells at the G1/S and G2/M interfaces of the cell cycle (29, 31). Additionally, replication of an ICP0 null mutant is complemented by unknown cellular factors expressed in transformed U2OS cells (49) but is severely impaired relative to the wild-type virus in confluent cells of most types (11), suggesting that ICP0 is required to induce cell cycle-associated activities while simultaneously arresting cells at the G1/S and G2/M interfaces. Although these observations suggest that ICP0 is responsible for cell cycle arrest and inhibition of host cell DNA synthesis, an ICP0 null mutant also inhibits cellular DNA synthesis and induces cell cycle arrest (31). Indeed, ICP27, like ICP0, also inhibits cellular DNA synthesis when expressed during HSV-1 infection (42). ICP27 is also required during HSV-1 infection to reduce levels of Cdk4 and cyclin D upon serum stimulation of quiescent cells (42). In cells infected with an ICP27 null mutant at later times (>2 hpi), pRb is hyperphosphorylated and E2F is released to induce S-phase-specific activities. Additionally, expression of both ICP27 and ICP4 is required to induce the expression of a subset of S-phase-specific proteins at early times postinfection of G1-arrested cells (28). Because the activities of the IE proteins have been examined primarily in cells arrested in G1, the role of IE proteins in deregulation of the cell cycle when S- and G2/M-phase cells are infected is poorly understood.
The reported properties of a fourth, less-well-characterized IE protein, ICP22, and its in-frame carboxyl-terminal variant, the US1.5 protein (13, 33) (Fig. 1A), suggest that these IE proteins also deregulate the cell cycle during HSV-1 productive infection. Studies of viral mutants have shown that ICP22 and/or US1.5 gene expression is required for efficient HSV-1 replication in most cell types in vitro and in vivo (restrictive cells) but is not required in certain immortalized cells (e.g., Vero, HeLa, and Hep-2 cells [permissive cells]) (34, 35, 41). In restrictive cells (but not in permissive cells), ICP22 and/or the US1.5 protein is required for efficient expression of 2 late (L) proteins (proteins whose expression is stringently dependent on viral DNA synthesis) but is not required for 1 L proteins (proteins synthesized at low levels in the absence of viral DNA synthesis) or viral DNA synthesis (35, 38, 41). As a result, ICP22–/US1.5– virus replication is severely reduced (100-fold) in restrictive cells compared to the wild-type virus. Although the products of the ICP22 and US1.5 genes are not required for efficient virus replication in permissive cells, replication is consistently 2- to 10-fold lower than replication of the wild-type virus. The basis for this modest reduction in virus replication has not been established. Like ICP0, ICP4, and ICP27, ICP22 and/or US1.5 expression in permissive cells has been reported to alter the expression and activities of cell cycle regulatory proteins, including cyclin A, cyclin B, wee-1, and myt-1, during HSV-1 infection (2-4, 32, 38). In addition to its ability to affect expression of cell cycle regulatory proteins, ICP22 expression is affected by the phase of the cell cycle at the time of infection. In infected permissive HeLa cells, ICP22 accumulation is greatest in S phase, suggesting a role for ICP22 during this phase of the cell cycle (10). Moreover, the electrophoretic mobility of ICP22 is altered when infected cells are treated with the Cdk inhibitor roscovitine (40). The ability of ICP22–/US1.5– viruses to replicate in a cell-type-dependent manner suggests that a primary role of ICP22 and/or the US1.5 protein may be to modify the expression or activities of cellular factors. It is possible that ICP22–/US1.5– viruses replicate in permissive but not restrictive cells because the cellular factors that ICP22 and/or the US1.5 protein may induce, activate, or repress are already induced, activated, or repressed in permissive cells but not in restrictive cells. The differential replication capabilities of ICP22–/US1.5– viruses in permissive and restrictive cells hint that these proteins either perform two independent functions in the two cell types or perform the same function that is manifested differently in the two cell types.
In this study, we attempted to establish a correlation between the reported functions of ICP22/US1.5 and HSV-1 replication efficiency. For this purpose, we first isolated and characterized an ICP22–/US1.5– mutant virus, 22/n199, for its ability to replicate in permissive and restrictive cells. 22/n199 replicated and synthesized viral DNA and L proteins in a manner similar to that reported for other ICP22–/US1.5– mutants in the two cell types (35, 41). Because cell cycle regulation is altered in all permissive cells, we hypothesized that deregulated expression of selected cell cycle-associated activities common only to permissive cells influences the replication efficiency of ICP22–/US1.5– mutants. To test this possibility, we examined the plating and replication efficiency of 22/n199 in restrictive and permissive cells synchronized in each phase of the cell cycle. Although infection of restrictive cells synchronized in G1, S, or G2/M did not overcome the growth restriction of 22/n199, infection of permissive cells synchronized in S phase, but not in G1 or G2/M phases, resulted in reduced replication efficiency of 22/n199. Because ICP22 and/or the US1.5 protein is required for the degradation of selected cyclins during HSV-1 infection (2), we analyzed the expression of selected cell cycle-associated proteins during infection of cells synchronized in each phase of the cell cycle. Although differences in cell cycle-associated protein expression were difficult to discern in restrictive cells, in S-phase permissive cells, in contrast to the wild-type virus, the ICP22–/US1.5– mutant virus failed to alter the expression of cyclins during S and G2/M phases. These findings indicate that one function of ICP22 and/or the US1.5 protein is to regulate S-phase-specific cell cycle-associated proteins necessary for efficient virus replication in permissive cells, and they suggest the possibility that other as yet unrecognized ICP22/US1.5-associated functions are required to facilitate HSV-1 replication in restrictive cells.
MATERIALS AND METHODS
Cells and viruses. Primary African green monkey kidney cells were obtained from Diagnostic Hybrids Inc. (Athens, OH). An immortalized African green monkey kidney cell line (Vero, ATCC CCL-81) and an immortalized rabbit skin cell line, (RAB-9, ATCC CRL-1414) were obtained from the American Type Culture Collection (Manassas, VA). A low-passage strain of human embryonic lung cells (HEL; strain 638) was derived at Baylor College of Medicine (39). ICP4-expressing Vero E5 cells were described previously (18). All cells were grown in Dulbecco's modified Eagle's medium, supplemented with 10% fetal bovine serum, 100 mM penicillin-streptomycin, and 2 mM glutamine, at 37°C in 5% CO2 unless otherwise noted. When grown in the presence of the proteasome inhibitor, MG132 (Calbiochem, San Diego, CA), the drug was first diluted in dimethyl sulfoxide and added to the medium at a final concentration of 2.5 μM as utilized by Everett et al. (22).
All viruses used in this study are derivatives of HSV-1 strain KOS. The ICP4 nonsense mutant, n12, was propagated in E5 cells (18). Wild-type as well as the US1/US1.5 mutant and rescuant viruses, 22/n199 and 22/n199R, respectively, generated in this study were propagated in Vero cells as previously described (18). Plaque assays of n12 were performed on E5 cells and plaque assays of all other viruses were performed on Vero cells as previously described (18).
Construction of the nonsense mutant plasmid, p22/n199. The strategy utilized by Post and Roizman (36) in construction of an ICP22–/US1.5– virus in strain F, R325-tk+, was modified for construction of 22/n199 in KOS. Like R325-tk+, 22/n199 is able to synthesize the same ICP22 truncation variant; however, unlike R325-tk+, 22/n199 contains the mutation only in US1/US1.5 and not in genes surrounding them. The 3.3-kb EcoRI-KpnI fragment of wild-type DNA (Fig. 1A) was cloned into pBR322 to produce pBR322-US1/US1.5. This fragment contains the entire ICP22 gene as well as 592 bp 5' of the transcription start site and 828 bp 3' of the poly(A) site. A single PvuII site lies within ICP22/US1.5 coding sequences in this fragment (Fig. 1B). pBR322-US1/US1.5 was digested with PvuII, and a 16-bp linker containing a HpaI site (5'-GGCTAGTTAACTAGCC-3') and encoding stop codons in all three reading frames was ligated into this site after codon 199 of US1. The resulting plasmid, pBR322-22/n199, was isolated after the products of the initial ligation were digested with HpaI and ligated under appropriate conditions to ensure that only one copy of the linker was present in the US1 open reading frame. Restriction digests and DNA sequencing confirmed the presence of a single HpaI linker in pBR322-22/n199.
Construction of the nonsense mutant virus, 22/n199, and rescuant virus, 22/n199R. To introduce the mutated US1 and US1.5 genes into the viral genome, the 21.5-kb EcoRI B fragment (24) of wild-type DNA, which contains all of IRL and most of IRS, including the entire ICP4 gene, was cloned into pBR325 to generate pBR325-EcoRIB (Fig. 1B). This plasmid was partially digested with EcoRI to linearize the plasmid at the EcoRI site within IRS. Concurrently, pBR322-22/n199 was digested with EcoRI and KpnI, and the resulting 3.3-kb fragment containing the mutant US1 and US1.5 genes was gel purified. This fragment was then ligated to partially digested pBR325-EcoRIB, and a construct containing the mutated US1 and US1.5 genes in the proper orientation, namely, pBR325-4/22/n199, was isolated. This plasmid was partially digested using KpnI and EcoRI, the 24.8-kb fragment of interest was gel purified, and the nonsense mutation in the US1 and US1.5 genes was introduced into the viral genome by homologous recombination as described previously (18). Specifically, Vero cells were cotransfected with infectious DNA from the ICP4 nonsense mutant, n12, and the purified 24.8-kb fragment using the CaPO4 method (12, 27). Marker rescue of the ICP4 nonsense mutation with wild-type ICP4 sequences in pBR325-4/22/n199 simultaneously introduced the nonsense mutation in the US1 and US1.5 genes into the viral genome. Three days after transfection, virus was harvested, Vero cell monolayers were infected with the virus after passage through a 0.2-μm filter to ensure that plaques were initiated by a single infectious virus particle, and cells were overlaid with 2% methylcellulose. Three days postinfection, individual plaques were collected, filtered, and used to infect new Vero cell monolayers. Plaques were picked three times and the virus produced on the final Vero cell monolayers was designated 22/n199. The plaque size of 22/n199 is smaller than wild-type plaques on Vero cell monolayers (Fig. 2C).
A control, rescuant virus of 22/n199, 22/n199R, was also isolated to ensure that the phenotypic properties of 22/n199 are due solely to the absence of functional ICP22 and/or US1.5 proteins. For this purpose, Vero cells were cotransfected with infectious 22/n199 viral DNA and the wild-type 3.3-kb EcoRI-KpnI fragment (Fig. 1A) of pBR322-US1/US1.5 (18). Following homologous recombination and three rounds of plaque purification, a stock of the rescuant virus was prepared. Potential rescuant viruses were identified by their plaque size, which is larger than 22/n199-derived plaques and approximately the same size as wild-type plaques (Fig. 2C).
Southern blot analysis. Two viral stocks each of the wild type, 22/n199, and 22/n199R were prepared on Vero cell monolayers as previously described (18). These stocks were diluted and used to infect Vero cell monolayers at a multiplicity of infection (MOI) of 5 PFU/cell. At 24 hpi, cells were harvested, cell suspensions extracted with phenol-chloroform, and total DNA precipitated. The amount of DNA in each sample was determined by UV spectroscopy and 5 μg of DNA of each sample digested with either EcoRI and PvuII or EcoRI and HpaI (Fig. 1C). Digested DNA was separated on a 0.7% agarose-Tris-borate-EDTA gel and transferred to a nylon membrane (Osmonics, Minnetonka, MN) by capillary transfer. An ICP22-specific probe was generated by random priming of ICP22-specific sequences in pBR322-US1/US1.5 in the presence of [-32P]ATP (Fig. 1B). The membrane was hybridized using this probe with ExpressHyb hybridization solution (BD Biosciences, Palo Alto, CA) per the manufacturer's instructions. The probed membranes were exposed to a PhosphorImager screen and analyzed using ImageQuant 3.3 software (Molecular Dynamics, Sunnyvale, CA).
Single-cycle replication assays. Vero, Rab-9, HEL, and primary AGMK cells were seeded at a density of 1 x 105 cells/ml in 35-mm dishes. Twenty-four hours later, monolayers were infected with 2.5 PFU/cell of wild-type virus, 22/n199, or 22/n199R in triplicate. Actual viral inocula were determined by standard plaque assay (18) and did not vary more than twofold. RAB-9 cells were not infected with the 22/n199R rescuant virus. At 1, 3, 6, 9, 12, 15, 18, 21, and 24 h postinfection, cells were scraped into medium, and suspensions were stored at –80°C. Samples were later thawed at room temperature and sonicated using a Misonix Sonicator 3000 (Misonix, Farmingdale, NY) at power level 8 for 1 min 40 s. Suspensions were clarified by low-speed centrifugation, and infectious virus in the supernatant fluids was measured by standard plaque assays on Vero cell monolayers (18).
Viral DNA slot blots. Vero, Rab-9, HEL, and primary AGMK cells were seeded at a density of 1 x 105 cells/ml in 35-mm dishes. Twenty-four hours later, replicate monolayers were infected at an MOI of 5 PFU/cell with wild-type or 22/n199 viruses. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. At 1, 3, 6, 9, 12, 15, 18, 21, and 24 h postinfection, cells were washed, treated with 0.25% trypsin, collected by low-speed centrifugation, and lysed by incubation for 4 h at 55°C in 0.5% sodium dodecyl sulfate (SDS) and 100 mg/ml proteinase K as described previously (8). Total cellular DNA was extracted with phenol-chloroform and ethanol precipitated, and 2.5 μg/sample of DNA was transferred to a nylon membrane using the Minifold II slot blot system (Schleicher and Schuell, Keene, NH) per the manufacturer's instructions. Probes used for DNA slot blots were generated by random priming of EcoRI fragments A, D, I, N, and O of HSV-1 DNA (24) in the presence of [-32P]dCTP. Hybridization of labeled probes to DNA slot blots was performed using ExpressHyb hybridization solution per the manufacturer's instructions. The probed membranes were exposed to a PhosphorImager screen and analyzed using ImageQuant 3.3 software (Molecular Dynamics).
Cell synchronization. (i) Isoleucine deprivation/refeeding. Actively dividing Vero cells were seeded in 60-mm dishes (5.5 x 105 cells per dish) and incubated at 37°C for 24 h. Cells were washed twice with phosphate-buffered saline and blocked in G1 by maintaining cells in isoleucine-free medium (Atlanta Biologicals, Atlanta, GA) containing 2% dialyzed serum (Atlanta Biologicals) for 42 h as described previously (11, 44). At this time, the medium was replaced with normal growth medium and cells were incubated at 37°C. Replicate cultures were harvested at 0, 3, 6, 9, 12, 15, 18, 21, and 24 h postaddition of normal growth medium (i.e., postrelease of the isoleucine deprivation block) for cell cycle analysis and determination of viral plating efficiency.
(ii) Double thymidine block. Vero cells were plated in 60-mm dishes (5.5 x 105 cells per dish) and incubated at 37°C for 24 h. Replicate cultures were blocked in early S phase by maintaining cells in normal medium for 24 h, adding 2 mM thymidine for 11 h, replacing with normal medium for 14 h, and adding 2 mM thymidine for an additional 11 h as previously described (9). HEL cells were blocked in early S phase by a similar protocol, with one exception: they were maintained in Dulbecco's modified Eagle's medium containing 2 mM thymidine and 0.1% fetal bovine serum during the first block. Following the second thymidine block, cultures were released into normal medium and processed at 0, 3, 6, 9, 12, 15, 18, 21, and 24 h using the procedures described below.
Cell cycle analysis and fluorescence-activated cell sorting (FACS). Cells were collected by treatment with 0.25% trypsin and low-speed centrifugation. Cells were resuspended in 70% ethanol and incubated at 4°C for 1 h. Fixed cells were pelleted by low-speed centrifugation and resuspended in 36 mM sodium citrate buffer containing 50 μg/ml propidium iodide and 400 μg/ml RNase A as previously described (25). The DNA content of individual cells was determined using a FACScan instrument (Becton Dickinson, Franklin Lakes, NJ) run on low speed, and quantified for single cells using CellQuest Pro 3.4 (Becton Dickinson) and Modfit 3.0 (Verity, Topsham, ME) software.
Plating efficiency. Stock preparations of the wild type and 22/n199 were diluted in normal medium to contain 100 PFU/400 μl of suspension as determined on Vero cell monolayers and 50 PFU/400 μl as determined on HEL cell monolayers, respectively. For each time postrelease of the cell cycle block, triplicate cultures were infected with 400 μl of virus suspension. After 1 h adsorption at 37°C, cells were overlaid with 0.5% methylcellulose. Four days later, methylcellulose was removed, cells were fixed and stained with 0.1% crystal violet in 20% ethanol, and plaques were counted.
Virus replication in synchronized cells. Vero and HEL cells were seeded as described for cell synchronization and infected either 24 h postplating (asynchronous) or, when synchronized to G1, S, or G2/M phases of the cell cycle, by the isoleucine deprivation or double thymidine block protocols described above. Vero cell cultures synchronized by isoleucine deprivation were infected at 6, 15, and 18.5 h postrelease and contained predominantly G1-, S-, and G2/M-phase cells, respectively. Vero cells synchronized by the double thymidine block protocol were infected at 3, 9, and 15 h postrelease and contained predominantly S, G2/M, and G1 cells, respectively. HEL cells synchronized by the double thymidine block protocol were infected at 6, 9, and 12 h postrelease and contained predominantly S, G2/M, and G1 cells, respectively. At these times, cells were infected with 2.5 PFU/cell of the wild-type virus or 22/n199 and harvested 24 h later. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. The amount of virus produced in synchronized, infected cultures was determined by standard plaque assays on 24-h-old Vero cell monolayers (18).
Western blot analysis. (i) Cell cycle-associated proteins. Cells were synchronized to G1, S, or G2/M phases of the cell cycle as described above and either collected, mock infected, or infected with 10 PFU/cell of the wild-type virus or 22/n199. Mock-infected and infected cells were collected at 12 hpi. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. At the time of harvest, cells were washed in phosphate-buffered saline, scraped, pelleted by low-speed centrifugation, and lysed in RIPA buffer (150 mM NaCl, 1% NP-40, 0.5% deoxycholate, 0.1% SDS, 50 mM Tris [pH 8.0]). After 30 min at 4°C, lysates were clarified by low-speed centrifugation, and an equal volume of 2x Laemmli sample buffer was added. All samples were volume standardized to contain 1 x 105 cell equivalents, separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) using 10% polyacrylamide minigels (Bio-Rad MiniProtean 3; Hercules, CA), and transferred to nitrocellulose membranes (Osmonics) per the manufacturer's instructions. Membranes were blocked in Tris-buffered saline (TBS) containing 2% nonfat dry milk and 0.05% Tween 20 (block solution) for 1 h at room temperature. Primary monoclonal antibodies were incubated at the following dilutions in block solution for 3 h: cyclin A (SC-239; Santa Cruz Biotechnology, Santa Cruz, CA), 1:500; cyclin B (610219; BD Biosciences), 1:500; and cyclin E (SC-247; Santa Cruz Biotechnology), 1:500. Membranes were washed six times in block solution over a 1-h period. Horseradish peroxidase-conjugated secondary antibodies (Jackson ImmunoResearch, West Grove, PA) were diluted 1:100,000 in block solution and incubated with the membranes for 2 h. Membranes were washed in block solution six times for a total time of 1 h and once in TBS with 0.05% Tween 20 for 10 min. Membranes were incubated with SuperSignal West Femto Maximum Sensitivity substrate (Pierce, Rockford, IL) for 5 min per the manufacturer's instructions. Membranes were washed once for 15 s in TBS and exposed on CL-X Posure film (Pierce).
(ii) Viral antigens. Vero and RAB-9 cells were seeded in 35-mm dishes (2 x 105 cells/dish). Twenty-four hours later, cells were infected with 2.5 PFU/cell of the wild-type virus or 22/n199. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. Every 3 h postinfection for 24 h, cells were collected and lysed as described above. Samples were volume standardized to contain 1 x 105 cell equivalents, proteins were separated on 10% SDS-PAGE gels and transferred to nitrocellulose membranes, and the membranes were blocked as described above. Primary polyclonal antibodies against gE (ab6510; Abcam, Cambridge, MA) and US11 (a kind gift from J. J. Diaz, Lyon, France) (19) were diluted 1:1,000, incubated with the membranes, and washed as described above. A polyclonal peptide antibody, Ab22, with reactivity against amino acids 14 to 29 of ICP22 as described by Ackermann et al. (1) was generated (Invitrogen). Ab22 was diluted 1:200, incubated with the membranes, and washed as described above. Secondary antibodies were diluted 1:100,000 in block solution and incubated with the membranes, and membranes were washed as above. Membranes were incubated with SuperSignal West Pico Chemiluminescent substrate (Pierce) for 5 min per the manufacturer's instructions and exposed on CL-X Posure film (Pierce).
RESULTS
Construction of an ICP22–/US1.5– virus, 22/n199, and a rescuant virus, 22/n199R. To date, the functional properties of the 420-amino-acid protein, ICP22, and the 249-amino-acid protein, US1.5, have been elucidated largely through studies of two mutants of HSV-1 strain F, R325-tk+ and del22Z. Both of these mutant viruses contain significant alterations in sequences surrounding the US1 gene, which encodes ICP22 (Fig. 1A). Both mutants not only contain large deletions in the US1/US1.5 genes, but also lack sequences which specify the L/ST transcripts (50). Characterization of these and other ICP22 mutants has led to the conclusion that the carboxyl-terminal half of ICP22 performs at least one essential function during HSV-1 lytic replication, namely, induction of 2 L protein synthesis (35, 38, 41).
In order to minimize alterations in US1/US1.5 and adjacent coding sequences, we constructed an ICP22–/US1.5– mutant plasmid, pBR322-22/n199, by introducing a 16-bp cDNA linker encoding stop codons in all three reading frames into a unique PvuII site in the coding sequences of the US1/US1.5 genes using standard cloning and mutagenesis techniques (Fig. 1A and B). Based on the genotype and phenotype of R325-tk+, the nonsense mutation in 22/n199 would eliminate the functions of both ICP22 and the US1.5 protein. The nonsense mutation was introduced into the viral genome by simultaneous marker rescue of the nonsense mutation in the essential ICP4 gene of mutant virus n12 and transfer into the genome by homologous recombination. As a control, a rescuant virus, designated 22/n199R, was also generated. Southern blot analysis confirmed the genotypes of 22/n199 and 22/n199R (Fig. 1B and C). Rescue of the n12 mutation in both copies of the ICP4 gene was confirmed by Southern blot analysis (data not shown). Western blot analysis of extracts of infected cells using Ab22 confirmed that wild-type- (Fig. 2D) and 22/n199R-infected cells (data not shown) expressed a 72-kDa ICP22-reactive species and 22/n199-infected cells (data not shown) expressed a 38-kDa truncated ICP22-reactive species.
Characterization of 22/n199. Previous studies have demonstrated that ICP22–/US1.5– mutants (R325-tk+ and del22lacZ) replicate efficiently (2- to 10-fold less than the wild type) in permissive Vero, HeLa, and Hep-2 cells, but inefficiently (100-fold less efficiently than the wild type) in all restrictive cell types tested (35, 38, 41). To determine whether 22/n199 also exhibits this host range phenotype, one-step growth curves of the wild type, 22/n199, and 22/n199R were performed in permissive Vero and restrictive RAB-9, HEL, and AGMK cells (Fig. 2A). In Vero cells, replication of 22/n199 was modestly impaired compared to the wild type and 22/n199R, exhibiting a fourfold reduction in viral yield at 24 hpi (KOS versus 22/n199, analysis of variance [ANOVA] P value = 0.039). In contrast to Vero cells, replication of 22/n199 was markedly reduced (50- to 100-fold reduction at 24 hpi) in RAB-9, HEL, and AGMK cells compared to wild-type or 22/n199R viruses (Fig. 2A). Consistent with its modestly impaired replication phenotype in Vero cells, 22/n199 plaques were smaller than wild-type or 22/n199R plaques on Vero cell monolayers (Fig. 2C). Further, 22/n199 plaques were barely visible on RAB-9 cell monolayers relative to wild-type and 22/n199R plaques (Fig. 2C). Although previous studies reported that ICP22–/US1.5– mutants synthesize wild type-like levels of viral DNA in both permissive and restrictive cells (38, 41), another study reported the opposite result (35). Consistent with the former reports, the kinetics and levels of viral DNA synthesis in 22/n199-infected cells were roughly equivalent to the kinetics and levels in wild-type-infected permissive and restrictive cells (Fig. 2B) (38, 41). Because other ICP22–/US1.5– viruses exhibit marked defects in 2 L protein synthesis in restrictive cells, we analyzed wild-type- and 22/n199-infected permissive Vero and restrictive RAB-9 cells for expression of representative 1 (gE) and 2 (US11) L proteins by Western blot analysis (Fig. 2D). Expression of the 1 L protein, gE, was delayed in 22/n199-infected Vero and RAB-9 cells compared to wild-type-infected cells. However, at 18 and 24 hpi, levels of gE were nearly equivalent in wild-type- and mutant-infected cells. In contrast, expression of the 2 L protein, US11, not only was delayed at early times in 22/n199-infected cells of both types, but did not reach wild-type levels even at 24 hpi, especially in RAB-9 cells. Collectively, these results demonstrate that the standard replication phenotypes of 22/n199 are similar to those of other ICP22–/US1.5– mutants (35, 41). Furthermore, since 22/n199R replicates in a wild-type-like manner, the replication defect of 22/n199 is due solely to the inability of this virus to synthesize ICP22 and/or US1.5 proteins.
Restrictive cells synchronized in each phase of the cell cycle remain restrictive for 22/n199 replication. (i) Plating efficiency. We hypothesized that cellular growth regulatory activities that are deregulated in permissive cells may be responsible for the ability of ICP22–/US1.5– viruses to replicate efficiently in these cells. To determine whether cellular factors expressed or activated during specific phases of the cell cycle influence the replication efficiency of 22/n199, we analyzed its ability to form plaques and replicate in cells synchronized in each phase of the cell cycle. To this end, restrictive HEL cells were synchronized in each phase of the cell cycle using a double thymidine block protocol. Replicate monolayers were arrested in S phase and released from growth arrest by incubation in normal growth medium. At the time of release and every 3 h thereafter for 24 h, cells were assayed for the level of synchrony in G1, S, or G2/M phases of the cell cycle by FACS analysis (Fig. 3A) or tested for their ability to support plaque formation by infecting with 50 PFU/plate of wild-type or 22n199 viruses (Fig. 3B). At the time of release, approximately 50% of the cells were synchronized in S phase, and the percentage of cells in S phase increased to 72% at 6 h postrelease. Because analysis of DNA content does not discriminate between cells in early S phase and cells in G1, it is likely that the highest percentage of cells in S phase actually occurred at the time of release. More than 50% of the cells were synchronized in G2/M at 9 h postrelease, and 68% were synchronized in G1 at 12 h postrelease.
At the time of release and every 3 h thereafter, wild-type virus produced approximately 50 plaques irrespective of the phase of the cell cycle at the time of infection (Fig. 3B), corroborating previous findings that wild-type HSV-1 produces plaques in cells independent of the phase of the cell cycle at the time of infection (11, 14). By contrast, 22/n199 failed to produce plaques at all times postrelease (Fig. 3B), indicating that activities expressed during each phase of the normal cell cycle are insufficient to support plaque formation by 22/n199. Although 22/n199 failed to produce plaques at any time post-release of the double thymidine block, 5 to 10 clusters of 4 to 20 rounded cells exhibiting cytopathic effects similar to those seen in Fig. 2C (bottom, middle panel) were observed when cells were infected during each phase of the cell cycle. This phenotype suggests the possibility that 22/n199 can initiate viral replication but cannot spread efficiently.
(ii) Replication efficiency. To determine the extent to which 22/n199 can replicate in synchronized HEL cells, cells were synchronized in S, G2/M, and G1 by releasing them from a double thymidine block for 6, 9, and 12 h, respectively. The percentages of cells at the time of infection in G1, S, and G2/M samples were 72.3, 68.5, and 58.9%, respectively. Synchronized cells were infected with 2.5 PFU/cell of wild-type or 22/n199 viruses and harvested 24 h later, and the amounts of infectious virus were determined by standard plaque assays on Vero cell monolayers (Fig. 3C). Although the wild-type virus replicated efficiently in asynchronous cells and in cells synchronized in G1, S, or G2/M, 22/n199 replicated 50-fold less efficiently than the wild-type virus in asynchronous cells and in cells synchronized in any phase of the cell cycle (Fig. 3C). In fact, the titer of 22/n199 in all these cells never exceeded the inoculum. Collectively, these results demonstrate that cell cycle-associated activities expressed specifically in G1, S, or G2/M are unable to complement the replication deficiencies of ICP22–/US1.5– mutants in restrictive cells.
ICP22 and/or the US1.5 protein is required for cell cycle-independent plaque formation and replication in permissive Vero cells. Although ICP22 and/or US1.5 is required for efficient plating and replication in restrictive cells, 22/n199 replicates to levels only 2- to 10-fold lower than wild-type virus in asynchronous Vero cells (Fig. 2A) (35, 38, 41). The reduction in viral replication efficiency may stem from reduced replication efficiency in all cells or the inability to replicate in cells in a specific stage(s) of the cell cycle. In this report we tested the latter hypothesis, and we are currently testing the former hypothesis by electron microscopy. To determine whether 22/n199 replicates in a cell cycle-dependent manner in permissive cells, Vero cell monolayers were arrested in G1 phase by isoleucine deprivation/refeeding (44) or in S phase by double thymidine block/release (9). At the time of release of the block and every 3 h thereafter for 24 h, cells were assayed for the extent of synchrony by FACS analysis (Fig. 3D and G). Vero cells were efficiently arrested in G1 when grown in isoleucine-free medium (78% of cells, Fig. 3D). At 15 h postaddition of isoleucine, most cells had recovered from the G1 block and progressed into S phase (75%). These cells then exited S phase and entered G2/M (35%) at 21 h postrelease. When Vero cells were arrested by the double thymidine block protocol, approximately 50% of the cells arrested in S phase (Fig. 3G). After release, the percentage of cells in S phase increased, with the highest percentage observed at 3 h postrelease (71%). At 9 h postrelease, the majority of cells (55%) exhibited 4n DNA content (G2/M). Most cells then cycled into G1 (81%).
Monolayers synchronized by either method were infected with 100 PFU/plate of wild-type or 22/n199 viruses (Fig. 3E and H). As published previously by our lab, Vero cell monolayers synchronized by isoleucine deprivation/refeeding and infected with 100 PFU/plate of wild-type virus produced approximately 100 plaques independent of the stage of the cell cycle at the time of infection (Fig. 3E) (11). Similar results were obtained for the wild-type virus in Vero cell monolayers synchronized by double thymidine block (Fig. 3H). In Vero cells synchronized in G1 by isoleucine deprivation/refeeding, 22/n199 produced 115 plaques when monolayers were infected with 100 PFU/plate of virus (Fig. 3E). As the percentage of S-phase cells increased over time, however, the number of 22/n199 plaques decreased, with the greatest reduction in plaque number (2- to 3-fold) occurring when the greatest proportion of cells was in S phase (Fig. 3E; wild type versus 22/n199, ANOVA P value = 2.7 x 10–3). Similarly, in Vero cells synchronized in S phase by the double thymidine block protocol, plaque formation in 22/n199-infected cells was reduced more than twofold compared to cells synchronized in G1 or G2/M (Fig. 3H; wild type versus 22/n199, ANOVA P value = 1.6 x 10–3). Furthermore, 22/n199 produced plaques most efficiently once cells entered G1. These experiments were repeated four times with similar results. Collectively, these observations indicate that ICP22/US1.5-associated functions are required for efficient HSV-1 plaque formation in S-phase Vero cells.
To determine whether the reduced plating efficiency of 22/n199 in Vero cells results from reduced replication in these cells, asynchronous cells and cells synchronized in G1, S, or G2/M were infected with wild-type, 22/n199, or 22/n199R viruses, and 24-h virus yields were determined. Cells were synchronized by both isoleucine deprivation and double thymidine block protocols in independent experiments, and asynchronous cells were tested in both experiments. For cells synchronized by isoleucine deprivation/refeeding, G1, S, and G2/M cells were obtained at 9, 15, and 18.5 h postrelease, yielding 82.5, 81.8, and 53.1% of cells in each phase, respectively. For cells synchronized by the double thymidine block protocol, G1, S, and G2/M cells were obtained at 15, 3, and 9 h postrelease, yielding 68.6, 54.7, and 90.1% of cells in each phase, respectively. Wild-type (Fig. 3F and I) and 22/n199R (data not shown) viruses replicated with equal efficiency in asynchronous and synchronized cells, producing 100 PFU/cell. Although 22/n199 replicated 2-fold less efficiently in asynchronous-, G1-, and G2/M-phase cells compared to the wild-type virus, it replicated the least efficiently (20% of the wild type) in S-phase cells synchronized by either method (Fig. 3F and I). Taken together, these results demonstrate that the functions of ICP22 and/or the US1.5 protein are required for cell cycle-independent growth in Vero cells and that these viral proteins are required to overcome growth restrictions specifically when Vero cells are infected during S phase. By extension, the inefficient replication of 22/n199 in S-phase cells may well be responsible for the modestly impaired replication phenotype of the virus in asynchronous permissive Vero cells (Fig. 2A).
ICP22 and/or US1.5 expression does not affect levels of Cdks but does affect levels of cyclins during wild-type infection of permissive Vero cells. Because the activities of ICP22 and/or the US1.5 protein are required for efficient replication in Vero S-phase cells, we hypothesized that these proteins alter, activate, or repress cellular proteins within S-phase-infected Vero cells but not in G1- or G2/M-infected Vero cells. To test this hypothesis, cells were synchronized in each phase of the cell cycle by isoleucine deprivation/refeeding and infected with wild-type or 22/n199 viruses, and cellular lysates were analyzed in duplicate by Western blot for levels of selected Cdks (data not shown) and cyclins (Fig. 4) at 12 hpi. The levels of expression of these specific Cdks and cyclins were examined because they regulate cell division during and immediately following S phase. Cells were also infected in the presence or absence of MG132, a proteasome inhibitor, to determine whether proteasome-mediated degradation plays a role in the levels of proteins observed.
(i) Cdks. During wild-type HSV-1 infection of asynchronous permissive and restrictive cells, levels of Cdks 1 and 4 have been reported to be reduced compared to mock-infected cells, whereas levels of Cdk2 have been reported to remain constant (2, 21, 42). Additionally, ICP22 and/or the US1.5 protein has been reported not to be required to reduce levels of Cdk1 and Cdk4 (2, 21, 42) during infection of asynchronous cells. In this analysis in synchronized Vero cells, Cdks 1, 2, and 4 were expressed in G1, S, and G2/M mock-infected cells. In wild-type-, 22/n199-, and 22/n199R-infected G1, S, and G2/M cells, levels of Cdk1 and 4 were equally reduced, and levels of Cdk2 remained constant compared to mock-infected cells (data not shown). Additionally, levels of Cdk2 remained constant independent of the stage of the cell cycle at the time of infection or of ICP22/US1.5 expression. Collectively, and as shown by others, these results demonstrate that expression of ICP22 and/or the US1.5 protein does not affect levels of Cdks 1, 2, and 4 during HSV-1 infection of synchronized Vero cells (2, 42).
(ii) Cyclins. Unlike Cdks, cyclins, as their name implies, "cycle" on and off during normal cell cycle progression. Since 22/n199 exhibited a reproducible growth impairment when S-phase cells were infected, we asked whether levels of S-phase cyclins (cyclins A and E) or a G2/M cyclin (cyclin B) are regulated differently following infection of synchronized Vero cell monolayers with the wild type or 22/n199 (Fig. 4). It has been reported that levels of cyclins A and B are reduced following HSV-1 infection of asynchronous permissive HeLa cells in an ICP22/US1.5-dependent manner (2).
In our tests in G1 cells, cyclin A was not expressed in mock-infected cells at the time of infection (0 h), at 12 h post-mock infection (at this time, 37% of the cells were in G1, 36% in S, and 27% in G2/M), or in either wild-type-infected or 22/n199-infected cells at 12 hpi (Fig. 4A). DNA content analysis of infected cells was not performed because newly synthesized viral DNA adds to the total DNA content of infected cells, making it impossible to determine the actual stage of the cell cycle using FACS analysis; however, expression of cyclins serves as a marker for cell cycle-associated activity. In the presence of MG132, however, cyclin A was expressed to a higher level in mock-infected G1 cells at 12 hpi. (Note: the 0 h samples in the left and right panels of Fig. 4 are duplicate samples. These were included to control for protein expression on different Western blots.) Additionally, cyclin A was expressed in wild-type- and 22/n199-infected MG132-treated cells, albeit at a lower level, indicating that cyclin A, an S-phase cyclin, is at least transiently expressed in both wild-type- and 22/n199-infected G1 cells (Fig. 4A).
In S-phase cells (mock, 0 h), cyclin A was expressed but was undetectable by 12 h post-mock infection (at this time, 59% of the cells were in G1, 40% in S, and 1% in G2/M; Fig. 4). Cyclin A was not expressed following wild-type infection at 12 hpi but was expressed in 22/n199-infected S-phase cells at this time, indicating that expression of ICP22 and/or the US1.5 protein is required to reduce cyclin A levels during HSV-1 infection of S-phase cells. In the presence of MG132, cyclin A was expressed in mock-, wild-type-, and 22/n199-infected cells at 12 hpi (Fig. 4B). Collectively, these observations indicate that cyclin A is degraded in a proteasome- and ICP22/US1.5-dependent manner specifically during infection of S-phase cells.
In cells infected in G2/M, cyclin A was expressed at the time of infection (mock, 0 h) but not at 12 hpi in mock-, wild-type-, or 22/n199-infected cells (Fig. 4C; in mock-infected cells at 12 hpi, 33% of the cells were in G1, 66% in S, and 0% in G2/M). Twelve hours after infection of cells in G2/M in the presence of MG132, cyclin A was present in mock-, wild-type-, and 22/n199-infected cells, indicating that cyclin A is degraded independent of ICP22 and/or the US1.5 protein when cells are infected in G2/M.
The kinetics of cyclin B expression in synchronized cells were similar to those of cyclin A, with two exceptions. First, cyclin B was not expressed in G1-infected cells in the presence of MG132 at 12 hpi (Fig. 4A), suggesting that wild-type- and 22/n199-infected cells do not induce cells infected during G1 to progress through S and into G2/M. Second, a low level of cyclin B was detected in cells synchronized in G2/M and infected with 22/n199 at 12 hpi (Fig. 4C). The presence of detectable cyclin B in these cells may imply that ICP22 and/or US1.5 protein expression is required to deregulate G2/M-specific expression during infection of G2/M cells. Similar results were obtained for cyclins A and B when cells were harvested at 6 hpi; however, the extent of degradation of cyclins A and B was less prominent (data not shown). Additionally, expression of cyclin A in 22/n199R-infected cells was similar to that observed in wild-type-infected cells (data not shown).
It has been reported that the intracellular localization and levels of cyclin E are unaffected in asynchronous HSV-1-infected cells (21). In these tests, two species of cyclin E were detected in mock-infected Vero cells in G1 (Fig. 4A). Because cyclin E is expressed only during late G1/early S phase in normal cells, it appears that cyclin E expression is markedly deregulated in Vero cells. In mock-infected G1 cells at 12 hpi, the faster-migrating cyclin E species (band c) was predominant. By contrast, wild-type- and 22/n199-infected G1 cells expressed both b and c forms of cyclin E seen in 0 h mock-infected cells, and form c was the predominant form. In the presence of MG132 in cells infected in G1 cells, band b was expressed under all conditions tested (Fig. 4A).
In S-phase cells at the time of infection, cyclin E was detected as two isoforms (Fig. 4B, bands b and c); however, in both mock- and wild-type-infected S-phase cells at 12 hpi, cyclin E was detected only as the fastest migrating form, band c. By contrast, in 22/n199-infected S-phase cells at 12 hpi, both bands b and c were expressed (Fig. 4B). In the presence of MG132 in S-phase-infected cells, two forms of cyclin E (bands b and c) were expressed in mock- and 22/n199-infected cells at 12 hpi (band c was more prominent in mock- than in 22/n199-infected cells), but the fast migrating form of the protein (band c) was expressed primarily in wild-type-infected S-phase cells.
Cells synchronized in G2/M expressed two forms of cyclin E at the time of infection (Fig. 4C, bands b and c). However, in mock-, wild-type-, and 22/n199-infected G2/M cells, the two slower-migrating forms of cyclin E (bands a and b) were prominent. In the presence of MG132, band b was the predominant form. Although previous reports have shown that ICP22 and/or US1.5 protein expression is not required to induce modifications of cyclins when G1 cells are infected, these results demonstrate that ICP22 and/or the US1.5 protein induces modifications of cyclins A, B, and E, directly or indirectly, specifically when S-phase Vero cells are infected.
DISCUSSION
Expression of proteins that regulate and/or are regulated by the cell cycle affect replication of DNA-containing viruses. While small DNA viruses and adenovirus utilize cell cycle-associated proteins expressed during normally regulated S phase to replicate, HSV-1 utilizes proteins expressed during multiple phases of the cell cycle. In a manner similar to IE and E regulatory proteins of small DNA viruses, HSV-1 IE proteins deregulate the expression of cell cycle-associated proteins (21, 42). In this study, we have shown that in addition to ICP0, ICP4, and ICP27, ICP22/US1.5 regulates the expression of cell cycle-associated proteins specifically when S-phase Vero cells are infected. The failure of the ICP22–/US1.5– mutant, 22/n199, to deregulate S- and G2/M-specific proteins (cyclins A, E, and B) may well be the basis for the inability of 22/n199 to replicate efficiently in S-phase-infected Vero cells. By extension, this property may also be the molecular basis for the modest replication impairment of ICP22–/US1.5– mutants in asynchronous permissive cells. Based on these findings we conclude that one function of ICP22 and/or the US1.5 protein is to deregulate the cell cycle, either directly or indirectly, when S-phase permissive cells are infected.
The replication defect(s) of 22/n199 is manifested differently in permissive and restrictive cells. ICP22–/US1.5– mutants initiate productive infection in restrictive cells (i.e., they express IE, E, and 1 L proteins and synthesize viral DNA) but complete the replication process and generate progeny virus only in the original infected cell and a few adjacent cells (35, 41). Virus spread is minimal. The most obvious defect in replication in restrictive cells is the inability to transcribe and synthesize selected 2 L proteins efficiently (38). It has therefore been proposed that the function of ICP22/US1.5 is to alter host cell gene expression to provide an environment conducive to 2 L gene expression (35, 41). A primary goal of this study was to determine whether the expression of cell cycle-associated factors during individual phases of the normal cell cycle controls ICP22–/US1.5– mutant permissivity. Similar approaches have identified S-phase cells as permissive hosts for an HSV-1 VP16 mutant (15), an adenovirus E1B 55-kDa mutant, (25) and wild-type bovine herpesvirus 4 (45). In contrast to these findings, however, we report here that cellular gene expression during any phase of the normal cell cycle was unable to complement 22/n199 in restrictive cells. This finding suggests that the dominant function associated with ICP22/US1.5 in restrictive cells is to alter expression of cellular proteins that are either not regulated by the cell cycle or expressed during multiple phases of the cell cycle.
Although ICP22–/US1.5– mutants replicate efficiently in permissive Vero, HeLa, and Hep-2 cells, the wild-type virus replicates 2 to 10 times more efficiently than ICP22–/US1.5– viruses (35, 41). Consequently, one may ask whether ICP22–/US1.5– viruses replicate less efficiently than the wild-type virus in all infected permissive cells, or only in a fraction of asynchronous cells. Here, we have established that the environment in S-phase Vero cells reduces 22/n199 plating and replication efficiency (Fig. 3), indicating that the phase of the cell cycle at the time of infection affects the efficiency of 22/n199 replication. In support of this observation, electron microscopic examination of asynchronous Vero cells infected with the wild type or 22/n199 revealed that a portion (20 to 30%) of 22/n199-infected but not wild-type-infected cells did not contain virions (data not shown). Although the stage of the cell cycle at the time of infection of these virion-free cells was not examined, the fraction of asynchronous cells in S phase is approximately 30%. Thus, a fraction of Vero cells (potentially those infected during S phase) apparently do not complete the replication process efficiently. Collectively, these results ascribe biological significance to the function of ICP22/US1.5 during wild-type virus replication in permissive cells.
ICP22 and/or the US1.5 protein is required to deregulate S and G2/M cyclins during infection of permissive S-phase cells: consequences for viral replication. The modest reduction in replication efficiency of 22/n199 in Vero cells infected during S phase suggests that ICP22 and/or the US1.5 protein functions during S phase. Although this is the first report demonstrating that ICP22 and/or the US1.5 protein has biological activity (i.e., its expression affects HSV-1 replication) specifically in S-phase permissive cells, previous studies have reported functions of ICP22 and the US1.5 protein consistent with this finding. ICP22 accumulates preferentially and is uniquely modified in HeLa cells infected during early S phase relative to G1 (10). The authors of that study also reported that ICP22 binds a 78-kDa cellular protein, p78, which accumulates specifically in physiologically normal S-phase cells and is constitutively expressed in many human tumor cell lines (37). This group also reported that ICP22 and/or the US1.5 protein reduces levels of cyclins A and B during HSV-1 infection of asynchronous HeLa cells (2). We show that cyclins A and B are degraded and cyclin E is modified either directly or indirectly in an ICP22/US1.5-dependent manner specifically when S-phase cells are infected (Fig. 4). It is possible that these properties of ICP22 and/or the US1.5 protein are at least partially responsible for the reduced replication efficiency of 22/n199 in permissive S-phase cells.
What is the function of ICP22/US1.5 during HSV-1 infection in S-phase permissive cells In a series of reports, Advani et al. suggested that an effect of ICP22/US1.5-dependent degradation of cyclins A and B is the activation of the cellular protein topoisomerase II (2-4). Since ICP22–/US1.5– mutants synthesize wild-type levels of viral DNA but do not synthesize 2 L proteins efficiently, these authors suggest that modified nonfunctional topoisomerase II reduces the number of newly replicated ICP22–/US1.5– genomes able to transcribe 2 L genes. Topoisomerase II is expressed in nontransformed cells during S phase of the normal cell cycle (26) and is also induced during HSV-1 infection (4). It is possible that in S-phase cells infected with ICP22–/US1.5– mutants, cyclin A expression precludes the activation of functional topoisomerase II and efficient expression of 2 L proteins. In the current study, Western blot analysis of the 2 L protein gC in wild-type- and 22/n199-infected S-phase Vero cells revealed that gC expression is reduced relative to wild-type-virus-infected cells or cells synchronized to G1 or G2/M and infected with 22/n199 (data not shown). By contrast, protein levels of gE, a 1 L protein, were equivalent in 22/n199-infected S-phase Vero cells relative to 22/n199-infected G1 and G2/M cells and wild-type-infected cells. Thus, like restrictive cells, the primary replication defect of 22/n199-infected S-phase Vero cells is evident as a defect in 2 L gene expression; however, whether topoisomerase II is altered differently in S-phase cells infected with 22/n199 relative to the wild type is unknown.
A potential role for cyclin E during HSV-1 replication. Identification of the role of cyclin E in HSV-1 replication is complicated by an apparent deregulation of cyclin E expression in Vero cells. Cyclin E is expressed during late G1 to early S phase in normal cells; Vero cells, however, express cyclin E during all phases of the cell cycle (Fig. 4). Indeed, this deregulation of cyclin E expression may be a factor in the immortalization of Vero cells. Since cyclin E expression or modification may also contribute to the permissivity of ICP22–/US1.5– mutants in Vero cells, the status of cyclin E in other permissive cell types is currently under investigation.
Cyclin E is expressed in different forms when S-phase Vero cells are infected with the wild-type virus or 22/n199. The faster migrating form is present in wild-type-infected S-phase cells, while a slower migrating form of the protein is also detected in 22/n199-infected S-phase cells. Since cyclin E levels are regulated by ubiquitination (30), and the slower-migrating form of cyclin E is 8 kDa (the size of ubiquitin) larger than the faster-migrating form, it is possible that the slower-migrating forms of cyclin E are ubiquitinated. In the presence of MG132, the slower-migrating species of cyclin E was the only species of cyclin E detected in all samples, suggesting that this species may be targeted for degradation via the proteasome. If this is the case, one role of ICP22 and/or the US1.5 protein may be to prevent the ubiquitination and potentially the proteolysis of cyclin E in S-phase infected cells. An active form of cyclin E may be required for its partners Cdks 1 and 2 to phosphorylate their normal targets (5). These targets include NPAT (a cellular transcription factor required for histone biosynthesis) (51), proliferating cell nuclear antigen (PCNA; required for efficient HSV-1 viral DNA replication) (6), and pRb (23). The targets of cyclin E function during cellular DNA synthesis, and it is possible that these proteins play accessory roles during HSV-1 DNA replication. Although DNA replication reaches wild-type levels in 22/n199-infected restrictive cells, it is possible that defects such as incomplete synthesis and concatemerization may contribute to the inability of the newly synthesized viral DNA to transcribe 2 L proteins. Alternatively, active Cdk2-cyclin E, Cdk1-cyclin E, Cdk1-cyclin A, Cdk2-cyclin A, or Cdk1-cyclin B may be required to phosphorylate viral targets. Indeed, Cdks modify the viral IE proteins ICP0 and ICP4 posttranslationally, and these modifications have been reported to confer specific activities on ICP0 during HSV-1 replication (16).
These studies add to the functions ascribed to ICP22 and the US1.5 protein and further link ICP22/US1.5 function to the cell cycle. Although factors expressed during S phase in permissive cells reduce 22/n199 replication, restrictive cells that are not in S phase do not complement virus replication. These properties distinguish two functions of ICP22 and the US1.5 protein or two consequences of the same function. Most studies of ICP22 and the US1.5 protein have been performed with permissive cells, and the reported functions of the proteins have not been associated with measurable effects on viral replication. The findings presented here provide biological relevance to several properties of ICP22 and the US1.5 protein, suggesting that expression of one or both of these proteins is required in S-phase permissive cells for efficient virus replication. The ability of ICP22/US1.5 to degrade or modify cyclins during infection of S-phase Vero cells may also be important for replication in restrictive cells; however, we have not observed similar degradation of cyclins A and B or modification of cyclin E by the wild-type virus in restrictive HEL cells (data not shown), indicating that an as yet unidentified role for ICP22/US1.5 exists in these cells.
REFERENCES
Ackermann, M., M. Sarmiento, and B. Roizman. 1985. Application of antibody to synthetic peptides for characterization of the intact and truncated 22 protein specified by herpes simplex virus 1 and the R325 22– deletion mutant. J. Virol. 56:207-215.
Advani, S. J., R. Brandimarti, R. R. Weichselbaum, and B. Roizman. 2000. The disappearance of cyclins A and B and the increase in activity of the G2/M-phase cellular kinase cdc2 in herpes simplex virus 1-infected cells require expression of the 22/US1.5 and UL13 viral genes. J. Virol. 74:8-15.
Advani, S. J., R. R. Weichselbaum, and B. Roizman. 2000. The role of cdc2 in the expression of herpes simplex virus genes. Proc. Natl. Acad. Sci. USA 97:10996-11001.
Advani, S. J., R. R. Weichselbaum, and B. Roizman. 2003. Herpes simplex virus 1 activates cdc2 to recruit topoisomerase II alpha for post-DNA synthesis expression of late genes. Proc. Natl. Acad. Sci. USA 100:4825-4830.
Aleem, E., H. Kiyokawa, and P. Kaldis. 2005. Cdc2-cyclin E complexes regulate the G1/S phase transition. Nat. Cell Biol. 7:831-836.
Arata, Y., M. Fujita, K. Ohtani, S. Kijima, and J. Y. Kato. 2000. Cdk2-dependent and -independent pathways in E2F-mediated S phase induction. J. Biol. Chem. 275:6337-6345.
Bagchi, S., P. Raychaudhuri, and J. R. Nevins. 1989. Phosphorylation-dependent activation of the adenovirus-inducible E2F transcription factor in a cell-free system. Proc. Natl. Acad. Sci. USA 86:4352-4356.
Balliet, J. W., J. C. Min, M. S. Cabatingan, and P. A. Schaffer. 2005. Site-directed mutagenesis of large DNA palindromes: construction and in vitro characterization of herpes simplex virus type 1 mutants containing point mutations that eliminate the oriL or oriS initiation function. J. Virol. 79:12783-12797.
Bostock, C. J., D. M. Prescott, and J. B. Kirkpatrick. 1971. An evaluation of the double thymidine block for synchronizing mammalian cells at the G1-S border. Exp. Cell Res. 68:163-168.
Bruni, R., and B. Roizman. 1998. Herpes simplex virus 1 regulatory protein ICP22 interacts with a new cell cycle-regulated factor and accumulates in a cell cycle-dependent fashion in infected cells. J. Virol. 72:8525-8531.
Cai, W., and P. A. Schaffer. 1991. A cellular function can enhance gene expression and plating efficiency of a mutant defective in the gene for ICP0, a transactivating protein of herpes simplex virus type 1. J. Virol. 65:4078-4090.
Cai, W. Z., and P. A. Schaffer. 1989. Herpes simplex virus type 1 ICP0 plays a critical role in the de novo synthesis of infectious virus following transfection of viral DNA. J. Virol. 63:4579-4589.
Carter, K. L., and B. Roizman. 1996. The promoter and transcriptional unit of a novel herpes simplex virus 1 gene are contained in, and encode a protein in frame with, the open reading frame of the 22 gene. J. Virol. 70:172-178.
Cohen, G. H., R. K. Vaughan, and W. C. Lawrence. 1971. Deoxyribonucleic acid synthesis in synchronized mammalian KB cells infected with herpes simplex virus. J. Virol. 7:783-791.
Daksis, J. I., and C. M. Preston. 1992. Herpes simplex virus immediate early gene expression in the absence of transinduction by Vmw65 varies during the cell cycle. Virology 189:196-202.
Davido, D. J., W. F. von Zagorski, W. S. Lane, and P. A. Schaffer. 2005. Phosphorylation site mutations affect herpes simplex virus type 1 ICP0 function. J. Virol. 79:1232-1243.
DeCaprio, J. A., J. W. Ludlow, J. Figge, J. Y. Shew, C. M. Huang, W. H. Lee, E. Marsilio, E. Paucha, and D. M. Livingston. 1988. SV40 large tumor antigen forms a specific complex with the product of the retinoblastoma susceptibility gene. Cell 54:275-283.
DeLuca, N. A., A. M. McCarthy, and P. A. Schaffer. 1985. Isolation and characterization of deletion mutants of herpes simplex virus type 1 in the gene encoding immediate-early regulatory protein ICP4. J. Virol. 56:558-570.
Diaz, J. J., D. Simonin, T. Masse, P. Deviller, K. Kindbeiter, L. Denoroy, and J. J. Madjar. 1993. The herpes simplex virus type 1 US11 gene product is a phosphorylated protein found to be non-specifically associated with both ribosomal subunits. J. Gen. Virol. 74:397-406.
Dyson, N., P. M. Howley, K. Munger, and E. Harlow. 1989. The human papilloma virus-16 E7 oncoprotein is able to bind to the retinoblastoma gene product. Science 243:934-937.
Ehmann, G. L., T. I. McLean, and S. L. Bachenheimer. 2000. Herpes simplex virus type 1 infection imposes a G(1)/S block in asynchronously growing cells and prevents G(1) entry in quiescent cells. Virology 267:335-349.
Everett, R. D., A. Orr, and C. M. Preston. 1998. A viral activator of gene expression functions via the ubiquitin-proteasome pathway. EMBO J. 17:7161-7169.
Geng, Y., E. N. Eaton, M. Picon, J. M. Roberts, A. S. Lundberg, A. Gifford, C. Sardet, and R. A. Weinberg. 1996. Regulation of cyclin E transcription by E2Fs and retinoblastoma protein. Oncogene 12:1173-1180.
Goldin, A. L., R. M. Sandri-Goldin, M. Levine, and J. C. Glorioso. 1981. Cloning of herpes simplex virus type 1 sequences representing the whole genome. J. Virol. 38:50-58.
Goodrum, F. D., and D. A. Ornelles. 1997. The early region 1B 55-kilodalton oncoprotein of adenovirus relieves growth restrictions imposed on viral replication by the cell cycle. J. Virol. 71:548-561.
Goswami, P. C., J. L. Roti Roti, and C. R. Hunt. 1996. The cell cycle-coupled expression of topoisomerase II during S phase is regulated by mRNA stability and is disrupted by heat shock or ionizing radiation. Mol. Cell. Biol. 16:1500-1508.
Graham, F. L., and A. J. van der Eb. 1973. A new technique for the assay of infectivity of human adenovirus 5 DNA. Virology 52:456-467.
Hilton, M. J., D. Mounghane, T. McLean, N. V. Contractor, J. O'Neil, K. Carpenter, and S. L. Bachenheimer. 1995. Induction by herpes simplex virus of free and heteromeric forms of E2F transcription factor. Virology 213:624-638.
Hobbs, W. E., II, and N. A. DeLuca. 1999. Perturbation of cell cycle progression and cellular gene expression as a function of herpes simplex virus ICP0. J. Virol. 73:8245-8255.
Koepp, D. M., L. K. Schaefer, X. Ye, K. Keyomarsi, C. Chu, J. W. Harper, and S. J. Elledge. 2001. Phosphorylation-dependent ubiquitination of cyclin E by the SCFFbw7 ubiquitin ligase. Science 294:173-177.
Lomonte, P., and R. D. Everett. 1999. Herpes simplex virus type 1 immediate-early protein Vmw110 inhibits progression of cells through mitosis and from G1 into S phase of the cell cycle. J. Virol. 73:9456-9467.
Long, M. C., V. Leong, P. A. Schaffer, C. A. Spencer, and S. A. Rice. 1999. ICP22 and the UL13 protein kinase are both required for herpes simplex virus-induced modification of the large subunit of RNA polymerase II. J. Virol. 73:5593-5604.
Ogle, W. O., and B. Roizman. 1999. Functional anatomy of herpes simplex virus 1 overlapping genes encoding infected-cell protein 22 and US1.5 protein. J. Virol. 73:4305-4315.
Poffenberger, K. L., A. D. Idowu, E. B. Fraser-Smith, P. E. Raichlen, and R. C. Herman. 1994. A herpes simplex virus type 1 ICP22 deletion mutant is altered for virulence and latency in vivo. Arch. Virol. 139:111-119.
Poffenberger, K. L., P. E. Raichlen, and R. C. Herman. 1993. In vitro characterization of a herpes simplex virus type 1 ICP22 deletion mutant. Virus Genes 7:171-186.
Post, L. E., and B. Roizman. 1981. A generalized technique for deletion of specific genes in large genomes: alpha gene 22 of herpes simplex virus 1 is not essential for growth. Cell 25:227-232.
Ren, Y., R. K. Busch, L. Perlaky, and H. Busch. 1998. The 58-kDa microspherule protein (MSP58), a nucleolar protein, interacts with nucleolar protein p120. Eur. J. Biochem. 253:734-742.
Rice, S. A., M. C. Long, V. Lam, P. A. Schaffer, and C. A. Spencer. 1995. Herpes simplex virus immediate-early protein ICP22 is required for viral modification of host RNA polymerase II and establishment of the normal viral transcription program. J. Virol. 69:5550-5559.
Schaffer, P. A., G. M. Aron, N. Biswal, and M. Benyesh-Melnick. 1973. Temperature-sensitive mutants of herpes simplex virus type 1: isolation, complementation and partial characterization. Virology 52:57-71.
Schang, L. M., A. Rosenberg, and P. A. Schaffer. 1999. Transcription of herpes simplex virus immediate-early and early genes is inhibited by roscovitine, an inhibitor specific for cellular cyclin-dependent kinases. J. Virol. 73:2161-2172.
Sears, A. E., I. W. Halliburton, B. Meignier, S. Silver, and B. Roizman. 1985. Herpes simplex virus 1 mutant deleted in the 22 gene: growth and gene expression in permissive and restrictive cells and establishment of latency in mice. J. Virol. 55:338-346.
Song, B., K. C. Yeh, J. Liu, and D. M. Knipe. 2001. Herpes simplex virus gene products required for viral inhibition of expression of G1-phase functions. Virology 290:320-328.
Tattersall, P. 1972. Replication of the parvovirus MVM. I. Dependence of virus multiplication and plaque formation on cell growth. J. Virol. 10:586-590.
Tobey, R. A., and K. D. Ley. 1970. Regulation of initiation of DNA synthesis in Chinese hamster cells. I. Production of stable, reversible G1-arrested populations in suspension culture. J. Cell Biol. 46:151-157.
Vanderplasschen, A., M. Goltz, J. Lyaku, C. Benarafa, H. J. Buhk, E. Thiry, and P. P. Pastoret. 1995. The replication in vitro of the gammaherpesvirus bovine herpesvirus 4 is restricted by its DNA synthesis dependence on the S phase of the cell cycle. Virology 213:328-340.
Whyte, P., K. J. Buchkovich, J. M. Horowitz, S. H. Friend, M. Raybuck, R. A. Weinberg, and E. Harlow. 1988. Association between an oncogene and an anti-oncogene: the adenovirus E1A proteins bind to the retinoblastoma gene product. Nature 334:124-129.
Whyte, P., N. M. Williamson, and E. Harlow. 1989. Cellular targets for transformation by the adenovirus E1A proteins. Cell 56:67-75.
Yanagi, K., A. Talavera, T. Nishimoto, and M. G. Rush. 1978. Inhibition of herpes simplex virus type 1 replication in temperature-sensitive cell cycle mutants. J. Virol. 25:42-50.
Yao, F., and P. A. Schaffer. 1995. An activity specified by the osteosarcoma line U2OS can substitute functionally for ICP0, a major regulatory protein of herpes simplex virus type 1. J. Virol. 69:6249-6258.
Yeh, L., and P. A. Schaffer. 1993. A novel class of transcripts expressed with late kinetics in the absence of ICP4 spans the junction between the long and short segments of the herpes simplex virus type 1 genome. J. Virol. 67:7373-7382.
Zhao, J., B. Dynlacht, T. Imai, T. Hori, and E. Harlow. 1998. Expression of NPAT, a novel substrate of cyclin E-CDK2, promotes S-phase entry. Genes Dev. 12:456-461.(Joseph S. Orlando, Todd L)
ABSTRACT
Herpes simplex virus type 1 ICP22–/US1.5– mutants initiate viral gene expression in all cells; however, in most cell types, the replication process stalls due to an inability to express 2 late proteins. Although the function of ICP22/US1.5 has not been established, it has been suggested that these proteins activate, induce, or repress the activity of cellular proteins during infection. In this study, we hypothesized that cell cycle-associated proteins are targets of ICP22/US1.5. For this purpose, we first isolated and characterized an ICP22–/US1.5– mutant virus, 22/n199. Like other ICP22–/US1.5– mutants, 22/n199 replicates in a cell-type-specific manner and fails to induce efficient 2 late gene expression in restrictive cells. Although synchronization of restrictive human embryonic lung cells in each phase of the cell cycle did not overcome the growth restrictions of 22/n199, synchronization of permissive Vero cells in S phase rendered them less able to support 22/n199 plaque formation and replication. Consistent with this finding, expression of cellular S-phase cyclins was altered in an ICP22/US1.5-dependent manner specifically when S-phase Vero cells were infected. Collectively, these observations support the notion that ICP22/US1.5 deregulates the cell cycle upon infection of S-phase permissive cells by altering expression of key cell cycle regulatory proteins either directly or indirectly.
INTRODUCTION
Many DNA-containing viruses alter cell cycle regulatory pathways to produce an environment that favors efficient viral replication. DNA viruses with small genomes do not carry the full complement of proteins required for viral DNA synthesis but utilize the cellular DNA synthesis machinery present during S phase to replicate their genomes. To access this machinery, parvoviruses depend on spontaneous progression of infected cells into S phase (43), whereas polyoma- and papillomaviruses contain proteins (large T antigen and E7, respectively) that drive and arrest cells in S phase (17, 20). Although DNA viruses with larger genomes, such as adeno- and herpesviruses, carry essential viral DNA replication proteins, these viruses also require selected S-phase activities for efficient replication (47, 48). During adenovirus infection, E1A drives cells and arrests them in S phase to free the cellular S-phase transcription factor, E2F, from the retinoblastoma protein (pRb) (46, 47). This activity is required for the transcription of adenovirus DNA replication proteins (7). Unlike the small DNA viruses and adenovirus, however, herpesviruses do not utilize machinery expressed exclusively during normal S phase for viral DNA synthesis, gene expression, or replication. Rather, they deregulate the cell cycle to access and utilize cellular activities normally expressed during G1 and S phases (21).
Herpes simplex virus type 1 (HSV-1) induces profound changes in cellular growth regulation pathways in infected cells. Studies of infected asynchronous cells and cells synchronized in each stage of the cell cycle have revealed that these alterations are dynamic and produce an environment that is unlike any phase of the normal cell cycle. At very early times postinfection (<2 h postinfection [hpi]) in cells in any phase of the cell cycle, an "S-phase-like" environment is created; however, cellular DNA synthesis does not occur in these cells as it would in normal S-phase cells. In these "S-phase-like" cells, the key S-phase transcription factor, E2F, is freed from hyperphosphorylated pRb (28), and expression of at least a subset of S-phase-specific genes is induced. At later times (>2 hpi) in cells infected during any stage of the cell cycle, cells appear to enter a "G1-like" phase. In contrast to "S-phase-like" cells in which pRb is hyperphosphorylated and E2F is freed, in "G1-like" cells, pRb is hypophosphorylated and E2F is found in complexes with pRb (21, 42). The ability of HSV-1 to activate S-phase genes very early in infection, while constraining expression of cell cycle-associated activities to a "G1-like" phase later in infection, independent of the phase of the cell cycle at the time of infection, suggests that HSV-1 infection alters the regulation of multiple cell cycle-associated activities.
Alteration of cell cycle machinery in HSV-1-infected cells is mediated in large part by the immediate-early (IE) regulatory proteins, infected cell protein 0 (ICP0), ICP27, ICP4, and ICP22. ICP0, when expressed alone, inhibits cellular DNA synthesis and arrests cells at the G1/S and G2/M interfaces of the cell cycle (29, 31). Additionally, replication of an ICP0 null mutant is complemented by unknown cellular factors expressed in transformed U2OS cells (49) but is severely impaired relative to the wild-type virus in confluent cells of most types (11), suggesting that ICP0 is required to induce cell cycle-associated activities while simultaneously arresting cells at the G1/S and G2/M interfaces. Although these observations suggest that ICP0 is responsible for cell cycle arrest and inhibition of host cell DNA synthesis, an ICP0 null mutant also inhibits cellular DNA synthesis and induces cell cycle arrest (31). Indeed, ICP27, like ICP0, also inhibits cellular DNA synthesis when expressed during HSV-1 infection (42). ICP27 is also required during HSV-1 infection to reduce levels of Cdk4 and cyclin D upon serum stimulation of quiescent cells (42). In cells infected with an ICP27 null mutant at later times (>2 hpi), pRb is hyperphosphorylated and E2F is released to induce S-phase-specific activities. Additionally, expression of both ICP27 and ICP4 is required to induce the expression of a subset of S-phase-specific proteins at early times postinfection of G1-arrested cells (28). Because the activities of the IE proteins have been examined primarily in cells arrested in G1, the role of IE proteins in deregulation of the cell cycle when S- and G2/M-phase cells are infected is poorly understood.
The reported properties of a fourth, less-well-characterized IE protein, ICP22, and its in-frame carboxyl-terminal variant, the US1.5 protein (13, 33) (Fig. 1A), suggest that these IE proteins also deregulate the cell cycle during HSV-1 productive infection. Studies of viral mutants have shown that ICP22 and/or US1.5 gene expression is required for efficient HSV-1 replication in most cell types in vitro and in vivo (restrictive cells) but is not required in certain immortalized cells (e.g., Vero, HeLa, and Hep-2 cells [permissive cells]) (34, 35, 41). In restrictive cells (but not in permissive cells), ICP22 and/or the US1.5 protein is required for efficient expression of 2 late (L) proteins (proteins whose expression is stringently dependent on viral DNA synthesis) but is not required for 1 L proteins (proteins synthesized at low levels in the absence of viral DNA synthesis) or viral DNA synthesis (35, 38, 41). As a result, ICP22–/US1.5– virus replication is severely reduced (100-fold) in restrictive cells compared to the wild-type virus. Although the products of the ICP22 and US1.5 genes are not required for efficient virus replication in permissive cells, replication is consistently 2- to 10-fold lower than replication of the wild-type virus. The basis for this modest reduction in virus replication has not been established. Like ICP0, ICP4, and ICP27, ICP22 and/or US1.5 expression in permissive cells has been reported to alter the expression and activities of cell cycle regulatory proteins, including cyclin A, cyclin B, wee-1, and myt-1, during HSV-1 infection (2-4, 32, 38). In addition to its ability to affect expression of cell cycle regulatory proteins, ICP22 expression is affected by the phase of the cell cycle at the time of infection. In infected permissive HeLa cells, ICP22 accumulation is greatest in S phase, suggesting a role for ICP22 during this phase of the cell cycle (10). Moreover, the electrophoretic mobility of ICP22 is altered when infected cells are treated with the Cdk inhibitor roscovitine (40). The ability of ICP22–/US1.5– viruses to replicate in a cell-type-dependent manner suggests that a primary role of ICP22 and/or the US1.5 protein may be to modify the expression or activities of cellular factors. It is possible that ICP22–/US1.5– viruses replicate in permissive but not restrictive cells because the cellular factors that ICP22 and/or the US1.5 protein may induce, activate, or repress are already induced, activated, or repressed in permissive cells but not in restrictive cells. The differential replication capabilities of ICP22–/US1.5– viruses in permissive and restrictive cells hint that these proteins either perform two independent functions in the two cell types or perform the same function that is manifested differently in the two cell types.
In this study, we attempted to establish a correlation between the reported functions of ICP22/US1.5 and HSV-1 replication efficiency. For this purpose, we first isolated and characterized an ICP22–/US1.5– mutant virus, 22/n199, for its ability to replicate in permissive and restrictive cells. 22/n199 replicated and synthesized viral DNA and L proteins in a manner similar to that reported for other ICP22–/US1.5– mutants in the two cell types (35, 41). Because cell cycle regulation is altered in all permissive cells, we hypothesized that deregulated expression of selected cell cycle-associated activities common only to permissive cells influences the replication efficiency of ICP22–/US1.5– mutants. To test this possibility, we examined the plating and replication efficiency of 22/n199 in restrictive and permissive cells synchronized in each phase of the cell cycle. Although infection of restrictive cells synchronized in G1, S, or G2/M did not overcome the growth restriction of 22/n199, infection of permissive cells synchronized in S phase, but not in G1 or G2/M phases, resulted in reduced replication efficiency of 22/n199. Because ICP22 and/or the US1.5 protein is required for the degradation of selected cyclins during HSV-1 infection (2), we analyzed the expression of selected cell cycle-associated proteins during infection of cells synchronized in each phase of the cell cycle. Although differences in cell cycle-associated protein expression were difficult to discern in restrictive cells, in S-phase permissive cells, in contrast to the wild-type virus, the ICP22–/US1.5– mutant virus failed to alter the expression of cyclins during S and G2/M phases. These findings indicate that one function of ICP22 and/or the US1.5 protein is to regulate S-phase-specific cell cycle-associated proteins necessary for efficient virus replication in permissive cells, and they suggest the possibility that other as yet unrecognized ICP22/US1.5-associated functions are required to facilitate HSV-1 replication in restrictive cells.
MATERIALS AND METHODS
Cells and viruses. Primary African green monkey kidney cells were obtained from Diagnostic Hybrids Inc. (Athens, OH). An immortalized African green monkey kidney cell line (Vero, ATCC CCL-81) and an immortalized rabbit skin cell line, (RAB-9, ATCC CRL-1414) were obtained from the American Type Culture Collection (Manassas, VA). A low-passage strain of human embryonic lung cells (HEL; strain 638) was derived at Baylor College of Medicine (39). ICP4-expressing Vero E5 cells were described previously (18). All cells were grown in Dulbecco's modified Eagle's medium, supplemented with 10% fetal bovine serum, 100 mM penicillin-streptomycin, and 2 mM glutamine, at 37°C in 5% CO2 unless otherwise noted. When grown in the presence of the proteasome inhibitor, MG132 (Calbiochem, San Diego, CA), the drug was first diluted in dimethyl sulfoxide and added to the medium at a final concentration of 2.5 μM as utilized by Everett et al. (22).
All viruses used in this study are derivatives of HSV-1 strain KOS. The ICP4 nonsense mutant, n12, was propagated in E5 cells (18). Wild-type as well as the US1/US1.5 mutant and rescuant viruses, 22/n199 and 22/n199R, respectively, generated in this study were propagated in Vero cells as previously described (18). Plaque assays of n12 were performed on E5 cells and plaque assays of all other viruses were performed on Vero cells as previously described (18).
Construction of the nonsense mutant plasmid, p22/n199. The strategy utilized by Post and Roizman (36) in construction of an ICP22–/US1.5– virus in strain F, R325-tk+, was modified for construction of 22/n199 in KOS. Like R325-tk+, 22/n199 is able to synthesize the same ICP22 truncation variant; however, unlike R325-tk+, 22/n199 contains the mutation only in US1/US1.5 and not in genes surrounding them. The 3.3-kb EcoRI-KpnI fragment of wild-type DNA (Fig. 1A) was cloned into pBR322 to produce pBR322-US1/US1.5. This fragment contains the entire ICP22 gene as well as 592 bp 5' of the transcription start site and 828 bp 3' of the poly(A) site. A single PvuII site lies within ICP22/US1.5 coding sequences in this fragment (Fig. 1B). pBR322-US1/US1.5 was digested with PvuII, and a 16-bp linker containing a HpaI site (5'-GGCTAGTTAACTAGCC-3') and encoding stop codons in all three reading frames was ligated into this site after codon 199 of US1. The resulting plasmid, pBR322-22/n199, was isolated after the products of the initial ligation were digested with HpaI and ligated under appropriate conditions to ensure that only one copy of the linker was present in the US1 open reading frame. Restriction digests and DNA sequencing confirmed the presence of a single HpaI linker in pBR322-22/n199.
Construction of the nonsense mutant virus, 22/n199, and rescuant virus, 22/n199R. To introduce the mutated US1 and US1.5 genes into the viral genome, the 21.5-kb EcoRI B fragment (24) of wild-type DNA, which contains all of IRL and most of IRS, including the entire ICP4 gene, was cloned into pBR325 to generate pBR325-EcoRIB (Fig. 1B). This plasmid was partially digested with EcoRI to linearize the plasmid at the EcoRI site within IRS. Concurrently, pBR322-22/n199 was digested with EcoRI and KpnI, and the resulting 3.3-kb fragment containing the mutant US1 and US1.5 genes was gel purified. This fragment was then ligated to partially digested pBR325-EcoRIB, and a construct containing the mutated US1 and US1.5 genes in the proper orientation, namely, pBR325-4/22/n199, was isolated. This plasmid was partially digested using KpnI and EcoRI, the 24.8-kb fragment of interest was gel purified, and the nonsense mutation in the US1 and US1.5 genes was introduced into the viral genome by homologous recombination as described previously (18). Specifically, Vero cells were cotransfected with infectious DNA from the ICP4 nonsense mutant, n12, and the purified 24.8-kb fragment using the CaPO4 method (12, 27). Marker rescue of the ICP4 nonsense mutation with wild-type ICP4 sequences in pBR325-4/22/n199 simultaneously introduced the nonsense mutation in the US1 and US1.5 genes into the viral genome. Three days after transfection, virus was harvested, Vero cell monolayers were infected with the virus after passage through a 0.2-μm filter to ensure that plaques were initiated by a single infectious virus particle, and cells were overlaid with 2% methylcellulose. Three days postinfection, individual plaques were collected, filtered, and used to infect new Vero cell monolayers. Plaques were picked three times and the virus produced on the final Vero cell monolayers was designated 22/n199. The plaque size of 22/n199 is smaller than wild-type plaques on Vero cell monolayers (Fig. 2C).
A control, rescuant virus of 22/n199, 22/n199R, was also isolated to ensure that the phenotypic properties of 22/n199 are due solely to the absence of functional ICP22 and/or US1.5 proteins. For this purpose, Vero cells were cotransfected with infectious 22/n199 viral DNA and the wild-type 3.3-kb EcoRI-KpnI fragment (Fig. 1A) of pBR322-US1/US1.5 (18). Following homologous recombination and three rounds of plaque purification, a stock of the rescuant virus was prepared. Potential rescuant viruses were identified by their plaque size, which is larger than 22/n199-derived plaques and approximately the same size as wild-type plaques (Fig. 2C).
Southern blot analysis. Two viral stocks each of the wild type, 22/n199, and 22/n199R were prepared on Vero cell monolayers as previously described (18). These stocks were diluted and used to infect Vero cell monolayers at a multiplicity of infection (MOI) of 5 PFU/cell. At 24 hpi, cells were harvested, cell suspensions extracted with phenol-chloroform, and total DNA precipitated. The amount of DNA in each sample was determined by UV spectroscopy and 5 μg of DNA of each sample digested with either EcoRI and PvuII or EcoRI and HpaI (Fig. 1C). Digested DNA was separated on a 0.7% agarose-Tris-borate-EDTA gel and transferred to a nylon membrane (Osmonics, Minnetonka, MN) by capillary transfer. An ICP22-specific probe was generated by random priming of ICP22-specific sequences in pBR322-US1/US1.5 in the presence of [-32P]ATP (Fig. 1B). The membrane was hybridized using this probe with ExpressHyb hybridization solution (BD Biosciences, Palo Alto, CA) per the manufacturer's instructions. The probed membranes were exposed to a PhosphorImager screen and analyzed using ImageQuant 3.3 software (Molecular Dynamics, Sunnyvale, CA).
Single-cycle replication assays. Vero, Rab-9, HEL, and primary AGMK cells were seeded at a density of 1 x 105 cells/ml in 35-mm dishes. Twenty-four hours later, monolayers were infected with 2.5 PFU/cell of wild-type virus, 22/n199, or 22/n199R in triplicate. Actual viral inocula were determined by standard plaque assay (18) and did not vary more than twofold. RAB-9 cells were not infected with the 22/n199R rescuant virus. At 1, 3, 6, 9, 12, 15, 18, 21, and 24 h postinfection, cells were scraped into medium, and suspensions were stored at –80°C. Samples were later thawed at room temperature and sonicated using a Misonix Sonicator 3000 (Misonix, Farmingdale, NY) at power level 8 for 1 min 40 s. Suspensions were clarified by low-speed centrifugation, and infectious virus in the supernatant fluids was measured by standard plaque assays on Vero cell monolayers (18).
Viral DNA slot blots. Vero, Rab-9, HEL, and primary AGMK cells were seeded at a density of 1 x 105 cells/ml in 35-mm dishes. Twenty-four hours later, replicate monolayers were infected at an MOI of 5 PFU/cell with wild-type or 22/n199 viruses. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. At 1, 3, 6, 9, 12, 15, 18, 21, and 24 h postinfection, cells were washed, treated with 0.25% trypsin, collected by low-speed centrifugation, and lysed by incubation for 4 h at 55°C in 0.5% sodium dodecyl sulfate (SDS) and 100 mg/ml proteinase K as described previously (8). Total cellular DNA was extracted with phenol-chloroform and ethanol precipitated, and 2.5 μg/sample of DNA was transferred to a nylon membrane using the Minifold II slot blot system (Schleicher and Schuell, Keene, NH) per the manufacturer's instructions. Probes used for DNA slot blots were generated by random priming of EcoRI fragments A, D, I, N, and O of HSV-1 DNA (24) in the presence of [-32P]dCTP. Hybridization of labeled probes to DNA slot blots was performed using ExpressHyb hybridization solution per the manufacturer's instructions. The probed membranes were exposed to a PhosphorImager screen and analyzed using ImageQuant 3.3 software (Molecular Dynamics).
Cell synchronization. (i) Isoleucine deprivation/refeeding. Actively dividing Vero cells were seeded in 60-mm dishes (5.5 x 105 cells per dish) and incubated at 37°C for 24 h. Cells were washed twice with phosphate-buffered saline and blocked in G1 by maintaining cells in isoleucine-free medium (Atlanta Biologicals, Atlanta, GA) containing 2% dialyzed serum (Atlanta Biologicals) for 42 h as described previously (11, 44). At this time, the medium was replaced with normal growth medium and cells were incubated at 37°C. Replicate cultures were harvested at 0, 3, 6, 9, 12, 15, 18, 21, and 24 h postaddition of normal growth medium (i.e., postrelease of the isoleucine deprivation block) for cell cycle analysis and determination of viral plating efficiency.
(ii) Double thymidine block. Vero cells were plated in 60-mm dishes (5.5 x 105 cells per dish) and incubated at 37°C for 24 h. Replicate cultures were blocked in early S phase by maintaining cells in normal medium for 24 h, adding 2 mM thymidine for 11 h, replacing with normal medium for 14 h, and adding 2 mM thymidine for an additional 11 h as previously described (9). HEL cells were blocked in early S phase by a similar protocol, with one exception: they were maintained in Dulbecco's modified Eagle's medium containing 2 mM thymidine and 0.1% fetal bovine serum during the first block. Following the second thymidine block, cultures were released into normal medium and processed at 0, 3, 6, 9, 12, 15, 18, 21, and 24 h using the procedures described below.
Cell cycle analysis and fluorescence-activated cell sorting (FACS). Cells were collected by treatment with 0.25% trypsin and low-speed centrifugation. Cells were resuspended in 70% ethanol and incubated at 4°C for 1 h. Fixed cells were pelleted by low-speed centrifugation and resuspended in 36 mM sodium citrate buffer containing 50 μg/ml propidium iodide and 400 μg/ml RNase A as previously described (25). The DNA content of individual cells was determined using a FACScan instrument (Becton Dickinson, Franklin Lakes, NJ) run on low speed, and quantified for single cells using CellQuest Pro 3.4 (Becton Dickinson) and Modfit 3.0 (Verity, Topsham, ME) software.
Plating efficiency. Stock preparations of the wild type and 22/n199 were diluted in normal medium to contain 100 PFU/400 μl of suspension as determined on Vero cell monolayers and 50 PFU/400 μl as determined on HEL cell monolayers, respectively. For each time postrelease of the cell cycle block, triplicate cultures were infected with 400 μl of virus suspension. After 1 h adsorption at 37°C, cells were overlaid with 0.5% methylcellulose. Four days later, methylcellulose was removed, cells were fixed and stained with 0.1% crystal violet in 20% ethanol, and plaques were counted.
Virus replication in synchronized cells. Vero and HEL cells were seeded as described for cell synchronization and infected either 24 h postplating (asynchronous) or, when synchronized to G1, S, or G2/M phases of the cell cycle, by the isoleucine deprivation or double thymidine block protocols described above. Vero cell cultures synchronized by isoleucine deprivation were infected at 6, 15, and 18.5 h postrelease and contained predominantly G1-, S-, and G2/M-phase cells, respectively. Vero cells synchronized by the double thymidine block protocol were infected at 3, 9, and 15 h postrelease and contained predominantly S, G2/M, and G1 cells, respectively. HEL cells synchronized by the double thymidine block protocol were infected at 6, 9, and 12 h postrelease and contained predominantly S, G2/M, and G1 cells, respectively. At these times, cells were infected with 2.5 PFU/cell of the wild-type virus or 22/n199 and harvested 24 h later. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. The amount of virus produced in synchronized, infected cultures was determined by standard plaque assays on 24-h-old Vero cell monolayers (18).
Western blot analysis. (i) Cell cycle-associated proteins. Cells were synchronized to G1, S, or G2/M phases of the cell cycle as described above and either collected, mock infected, or infected with 10 PFU/cell of the wild-type virus or 22/n199. Mock-infected and infected cells were collected at 12 hpi. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. At the time of harvest, cells were washed in phosphate-buffered saline, scraped, pelleted by low-speed centrifugation, and lysed in RIPA buffer (150 mM NaCl, 1% NP-40, 0.5% deoxycholate, 0.1% SDS, 50 mM Tris [pH 8.0]). After 30 min at 4°C, lysates were clarified by low-speed centrifugation, and an equal volume of 2x Laemmli sample buffer was added. All samples were volume standardized to contain 1 x 105 cell equivalents, separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) using 10% polyacrylamide minigels (Bio-Rad MiniProtean 3; Hercules, CA), and transferred to nitrocellulose membranes (Osmonics) per the manufacturer's instructions. Membranes were blocked in Tris-buffered saline (TBS) containing 2% nonfat dry milk and 0.05% Tween 20 (block solution) for 1 h at room temperature. Primary monoclonal antibodies were incubated at the following dilutions in block solution for 3 h: cyclin A (SC-239; Santa Cruz Biotechnology, Santa Cruz, CA), 1:500; cyclin B (610219; BD Biosciences), 1:500; and cyclin E (SC-247; Santa Cruz Biotechnology), 1:500. Membranes were washed six times in block solution over a 1-h period. Horseradish peroxidase-conjugated secondary antibodies (Jackson ImmunoResearch, West Grove, PA) were diluted 1:100,000 in block solution and incubated with the membranes for 2 h. Membranes were washed in block solution six times for a total time of 1 h and once in TBS with 0.05% Tween 20 for 10 min. Membranes were incubated with SuperSignal West Femto Maximum Sensitivity substrate (Pierce, Rockford, IL) for 5 min per the manufacturer's instructions. Membranes were washed once for 15 s in TBS and exposed on CL-X Posure film (Pierce).
(ii) Viral antigens. Vero and RAB-9 cells were seeded in 35-mm dishes (2 x 105 cells/dish). Twenty-four hours later, cells were infected with 2.5 PFU/cell of the wild-type virus or 22/n199. Actual viral inocula were determined by standard plaque assays (18) and did not vary more than twofold. Every 3 h postinfection for 24 h, cells were collected and lysed as described above. Samples were volume standardized to contain 1 x 105 cell equivalents, proteins were separated on 10% SDS-PAGE gels and transferred to nitrocellulose membranes, and the membranes were blocked as described above. Primary polyclonal antibodies against gE (ab6510; Abcam, Cambridge, MA) and US11 (a kind gift from J. J. Diaz, Lyon, France) (19) were diluted 1:1,000, incubated with the membranes, and washed as described above. A polyclonal peptide antibody, Ab22, with reactivity against amino acids 14 to 29 of ICP22 as described by Ackermann et al. (1) was generated (Invitrogen). Ab22 was diluted 1:200, incubated with the membranes, and washed as described above. Secondary antibodies were diluted 1:100,000 in block solution and incubated with the membranes, and membranes were washed as above. Membranes were incubated with SuperSignal West Pico Chemiluminescent substrate (Pierce) for 5 min per the manufacturer's instructions and exposed on CL-X Posure film (Pierce).
RESULTS
Construction of an ICP22–/US1.5– virus, 22/n199, and a rescuant virus, 22/n199R. To date, the functional properties of the 420-amino-acid protein, ICP22, and the 249-amino-acid protein, US1.5, have been elucidated largely through studies of two mutants of HSV-1 strain F, R325-tk+ and del22Z. Both of these mutant viruses contain significant alterations in sequences surrounding the US1 gene, which encodes ICP22 (Fig. 1A). Both mutants not only contain large deletions in the US1/US1.5 genes, but also lack sequences which specify the L/ST transcripts (50). Characterization of these and other ICP22 mutants has led to the conclusion that the carboxyl-terminal half of ICP22 performs at least one essential function during HSV-1 lytic replication, namely, induction of 2 L protein synthesis (35, 38, 41).
In order to minimize alterations in US1/US1.5 and adjacent coding sequences, we constructed an ICP22–/US1.5– mutant plasmid, pBR322-22/n199, by introducing a 16-bp cDNA linker encoding stop codons in all three reading frames into a unique PvuII site in the coding sequences of the US1/US1.5 genes using standard cloning and mutagenesis techniques (Fig. 1A and B). Based on the genotype and phenotype of R325-tk+, the nonsense mutation in 22/n199 would eliminate the functions of both ICP22 and the US1.5 protein. The nonsense mutation was introduced into the viral genome by simultaneous marker rescue of the nonsense mutation in the essential ICP4 gene of mutant virus n12 and transfer into the genome by homologous recombination. As a control, a rescuant virus, designated 22/n199R, was also generated. Southern blot analysis confirmed the genotypes of 22/n199 and 22/n199R (Fig. 1B and C). Rescue of the n12 mutation in both copies of the ICP4 gene was confirmed by Southern blot analysis (data not shown). Western blot analysis of extracts of infected cells using Ab22 confirmed that wild-type- (Fig. 2D) and 22/n199R-infected cells (data not shown) expressed a 72-kDa ICP22-reactive species and 22/n199-infected cells (data not shown) expressed a 38-kDa truncated ICP22-reactive species.
Characterization of 22/n199. Previous studies have demonstrated that ICP22–/US1.5– mutants (R325-tk+ and del22lacZ) replicate efficiently (2- to 10-fold less than the wild type) in permissive Vero, HeLa, and Hep-2 cells, but inefficiently (100-fold less efficiently than the wild type) in all restrictive cell types tested (35, 38, 41). To determine whether 22/n199 also exhibits this host range phenotype, one-step growth curves of the wild type, 22/n199, and 22/n199R were performed in permissive Vero and restrictive RAB-9, HEL, and AGMK cells (Fig. 2A). In Vero cells, replication of 22/n199 was modestly impaired compared to the wild type and 22/n199R, exhibiting a fourfold reduction in viral yield at 24 hpi (KOS versus 22/n199, analysis of variance [ANOVA] P value = 0.039). In contrast to Vero cells, replication of 22/n199 was markedly reduced (50- to 100-fold reduction at 24 hpi) in RAB-9, HEL, and AGMK cells compared to wild-type or 22/n199R viruses (Fig. 2A). Consistent with its modestly impaired replication phenotype in Vero cells, 22/n199 plaques were smaller than wild-type or 22/n199R plaques on Vero cell monolayers (Fig. 2C). Further, 22/n199 plaques were barely visible on RAB-9 cell monolayers relative to wild-type and 22/n199R plaques (Fig. 2C). Although previous studies reported that ICP22–/US1.5– mutants synthesize wild type-like levels of viral DNA in both permissive and restrictive cells (38, 41), another study reported the opposite result (35). Consistent with the former reports, the kinetics and levels of viral DNA synthesis in 22/n199-infected cells were roughly equivalent to the kinetics and levels in wild-type-infected permissive and restrictive cells (Fig. 2B) (38, 41). Because other ICP22–/US1.5– viruses exhibit marked defects in 2 L protein synthesis in restrictive cells, we analyzed wild-type- and 22/n199-infected permissive Vero and restrictive RAB-9 cells for expression of representative 1 (gE) and 2 (US11) L proteins by Western blot analysis (Fig. 2D). Expression of the 1 L protein, gE, was delayed in 22/n199-infected Vero and RAB-9 cells compared to wild-type-infected cells. However, at 18 and 24 hpi, levels of gE were nearly equivalent in wild-type- and mutant-infected cells. In contrast, expression of the 2 L protein, US11, not only was delayed at early times in 22/n199-infected cells of both types, but did not reach wild-type levels even at 24 hpi, especially in RAB-9 cells. Collectively, these results demonstrate that the standard replication phenotypes of 22/n199 are similar to those of other ICP22–/US1.5– mutants (35, 41). Furthermore, since 22/n199R replicates in a wild-type-like manner, the replication defect of 22/n199 is due solely to the inability of this virus to synthesize ICP22 and/or US1.5 proteins.
Restrictive cells synchronized in each phase of the cell cycle remain restrictive for 22/n199 replication. (i) Plating efficiency. We hypothesized that cellular growth regulatory activities that are deregulated in permissive cells may be responsible for the ability of ICP22–/US1.5– viruses to replicate efficiently in these cells. To determine whether cellular factors expressed or activated during specific phases of the cell cycle influence the replication efficiency of 22/n199, we analyzed its ability to form plaques and replicate in cells synchronized in each phase of the cell cycle. To this end, restrictive HEL cells were synchronized in each phase of the cell cycle using a double thymidine block protocol. Replicate monolayers were arrested in S phase and released from growth arrest by incubation in normal growth medium. At the time of release and every 3 h thereafter for 24 h, cells were assayed for the level of synchrony in G1, S, or G2/M phases of the cell cycle by FACS analysis (Fig. 3A) or tested for their ability to support plaque formation by infecting with 50 PFU/plate of wild-type or 22n199 viruses (Fig. 3B). At the time of release, approximately 50% of the cells were synchronized in S phase, and the percentage of cells in S phase increased to 72% at 6 h postrelease. Because analysis of DNA content does not discriminate between cells in early S phase and cells in G1, it is likely that the highest percentage of cells in S phase actually occurred at the time of release. More than 50% of the cells were synchronized in G2/M at 9 h postrelease, and 68% were synchronized in G1 at 12 h postrelease.
At the time of release and every 3 h thereafter, wild-type virus produced approximately 50 plaques irrespective of the phase of the cell cycle at the time of infection (Fig. 3B), corroborating previous findings that wild-type HSV-1 produces plaques in cells independent of the phase of the cell cycle at the time of infection (11, 14). By contrast, 22/n199 failed to produce plaques at all times postrelease (Fig. 3B), indicating that activities expressed during each phase of the normal cell cycle are insufficient to support plaque formation by 22/n199. Although 22/n199 failed to produce plaques at any time post-release of the double thymidine block, 5 to 10 clusters of 4 to 20 rounded cells exhibiting cytopathic effects similar to those seen in Fig. 2C (bottom, middle panel) were observed when cells were infected during each phase of the cell cycle. This phenotype suggests the possibility that 22/n199 can initiate viral replication but cannot spread efficiently.
(ii) Replication efficiency. To determine the extent to which 22/n199 can replicate in synchronized HEL cells, cells were synchronized in S, G2/M, and G1 by releasing them from a double thymidine block for 6, 9, and 12 h, respectively. The percentages of cells at the time of infection in G1, S, and G2/M samples were 72.3, 68.5, and 58.9%, respectively. Synchronized cells were infected with 2.5 PFU/cell of wild-type or 22/n199 viruses and harvested 24 h later, and the amounts of infectious virus were determined by standard plaque assays on Vero cell monolayers (Fig. 3C). Although the wild-type virus replicated efficiently in asynchronous cells and in cells synchronized in G1, S, or G2/M, 22/n199 replicated 50-fold less efficiently than the wild-type virus in asynchronous cells and in cells synchronized in any phase of the cell cycle (Fig. 3C). In fact, the titer of 22/n199 in all these cells never exceeded the inoculum. Collectively, these results demonstrate that cell cycle-associated activities expressed specifically in G1, S, or G2/M are unable to complement the replication deficiencies of ICP22–/US1.5– mutants in restrictive cells.
ICP22 and/or the US1.5 protein is required for cell cycle-independent plaque formation and replication in permissive Vero cells. Although ICP22 and/or US1.5 is required for efficient plating and replication in restrictive cells, 22/n199 replicates to levels only 2- to 10-fold lower than wild-type virus in asynchronous Vero cells (Fig. 2A) (35, 38, 41). The reduction in viral replication efficiency may stem from reduced replication efficiency in all cells or the inability to replicate in cells in a specific stage(s) of the cell cycle. In this report we tested the latter hypothesis, and we are currently testing the former hypothesis by electron microscopy. To determine whether 22/n199 replicates in a cell cycle-dependent manner in permissive cells, Vero cell monolayers were arrested in G1 phase by isoleucine deprivation/refeeding (44) or in S phase by double thymidine block/release (9). At the time of release of the block and every 3 h thereafter for 24 h, cells were assayed for the extent of synchrony by FACS analysis (Fig. 3D and G). Vero cells were efficiently arrested in G1 when grown in isoleucine-free medium (78% of cells, Fig. 3D). At 15 h postaddition of isoleucine, most cells had recovered from the G1 block and progressed into S phase (75%). These cells then exited S phase and entered G2/M (35%) at 21 h postrelease. When Vero cells were arrested by the double thymidine block protocol, approximately 50% of the cells arrested in S phase (Fig. 3G). After release, the percentage of cells in S phase increased, with the highest percentage observed at 3 h postrelease (71%). At 9 h postrelease, the majority of cells (55%) exhibited 4n DNA content (G2/M). Most cells then cycled into G1 (81%).
Monolayers synchronized by either method were infected with 100 PFU/plate of wild-type or 22/n199 viruses (Fig. 3E and H). As published previously by our lab, Vero cell monolayers synchronized by isoleucine deprivation/refeeding and infected with 100 PFU/plate of wild-type virus produced approximately 100 plaques independent of the stage of the cell cycle at the time of infection (Fig. 3E) (11). Similar results were obtained for the wild-type virus in Vero cell monolayers synchronized by double thymidine block (Fig. 3H). In Vero cells synchronized in G1 by isoleucine deprivation/refeeding, 22/n199 produced 115 plaques when monolayers were infected with 100 PFU/plate of virus (Fig. 3E). As the percentage of S-phase cells increased over time, however, the number of 22/n199 plaques decreased, with the greatest reduction in plaque number (2- to 3-fold) occurring when the greatest proportion of cells was in S phase (Fig. 3E; wild type versus 22/n199, ANOVA P value = 2.7 x 10–3). Similarly, in Vero cells synchronized in S phase by the double thymidine block protocol, plaque formation in 22/n199-infected cells was reduced more than twofold compared to cells synchronized in G1 or G2/M (Fig. 3H; wild type versus 22/n199, ANOVA P value = 1.6 x 10–3). Furthermore, 22/n199 produced plaques most efficiently once cells entered G1. These experiments were repeated four times with similar results. Collectively, these observations indicate that ICP22/US1.5-associated functions are required for efficient HSV-1 plaque formation in S-phase Vero cells.
To determine whether the reduced plating efficiency of 22/n199 in Vero cells results from reduced replication in these cells, asynchronous cells and cells synchronized in G1, S, or G2/M were infected with wild-type, 22/n199, or 22/n199R viruses, and 24-h virus yields were determined. Cells were synchronized by both isoleucine deprivation and double thymidine block protocols in independent experiments, and asynchronous cells were tested in both experiments. For cells synchronized by isoleucine deprivation/refeeding, G1, S, and G2/M cells were obtained at 9, 15, and 18.5 h postrelease, yielding 82.5, 81.8, and 53.1% of cells in each phase, respectively. For cells synchronized by the double thymidine block protocol, G1, S, and G2/M cells were obtained at 15, 3, and 9 h postrelease, yielding 68.6, 54.7, and 90.1% of cells in each phase, respectively. Wild-type (Fig. 3F and I) and 22/n199R (data not shown) viruses replicated with equal efficiency in asynchronous and synchronized cells, producing 100 PFU/cell. Although 22/n199 replicated 2-fold less efficiently in asynchronous-, G1-, and G2/M-phase cells compared to the wild-type virus, it replicated the least efficiently (20% of the wild type) in S-phase cells synchronized by either method (Fig. 3F and I). Taken together, these results demonstrate that the functions of ICP22 and/or the US1.5 protein are required for cell cycle-independent growth in Vero cells and that these viral proteins are required to overcome growth restrictions specifically when Vero cells are infected during S phase. By extension, the inefficient replication of 22/n199 in S-phase cells may well be responsible for the modestly impaired replication phenotype of the virus in asynchronous permissive Vero cells (Fig. 2A).
ICP22 and/or US1.5 expression does not affect levels of Cdks but does affect levels of cyclins during wild-type infection of permissive Vero cells. Because the activities of ICP22 and/or the US1.5 protein are required for efficient replication in Vero S-phase cells, we hypothesized that these proteins alter, activate, or repress cellular proteins within S-phase-infected Vero cells but not in G1- or G2/M-infected Vero cells. To test this hypothesis, cells were synchronized in each phase of the cell cycle by isoleucine deprivation/refeeding and infected with wild-type or 22/n199 viruses, and cellular lysates were analyzed in duplicate by Western blot for levels of selected Cdks (data not shown) and cyclins (Fig. 4) at 12 hpi. The levels of expression of these specific Cdks and cyclins were examined because they regulate cell division during and immediately following S phase. Cells were also infected in the presence or absence of MG132, a proteasome inhibitor, to determine whether proteasome-mediated degradation plays a role in the levels of proteins observed.
(i) Cdks. During wild-type HSV-1 infection of asynchronous permissive and restrictive cells, levels of Cdks 1 and 4 have been reported to be reduced compared to mock-infected cells, whereas levels of Cdk2 have been reported to remain constant (2, 21, 42). Additionally, ICP22 and/or the US1.5 protein has been reported not to be required to reduce levels of Cdk1 and Cdk4 (2, 21, 42) during infection of asynchronous cells. In this analysis in synchronized Vero cells, Cdks 1, 2, and 4 were expressed in G1, S, and G2/M mock-infected cells. In wild-type-, 22/n199-, and 22/n199R-infected G1, S, and G2/M cells, levels of Cdk1 and 4 were equally reduced, and levels of Cdk2 remained constant compared to mock-infected cells (data not shown). Additionally, levels of Cdk2 remained constant independent of the stage of the cell cycle at the time of infection or of ICP22/US1.5 expression. Collectively, and as shown by others, these results demonstrate that expression of ICP22 and/or the US1.5 protein does not affect levels of Cdks 1, 2, and 4 during HSV-1 infection of synchronized Vero cells (2, 42).
(ii) Cyclins. Unlike Cdks, cyclins, as their name implies, "cycle" on and off during normal cell cycle progression. Since 22/n199 exhibited a reproducible growth impairment when S-phase cells were infected, we asked whether levels of S-phase cyclins (cyclins A and E) or a G2/M cyclin (cyclin B) are regulated differently following infection of synchronized Vero cell monolayers with the wild type or 22/n199 (Fig. 4). It has been reported that levels of cyclins A and B are reduced following HSV-1 infection of asynchronous permissive HeLa cells in an ICP22/US1.5-dependent manner (2).
In our tests in G1 cells, cyclin A was not expressed in mock-infected cells at the time of infection (0 h), at 12 h post-mock infection (at this time, 37% of the cells were in G1, 36% in S, and 27% in G2/M), or in either wild-type-infected or 22/n199-infected cells at 12 hpi (Fig. 4A). DNA content analysis of infected cells was not performed because newly synthesized viral DNA adds to the total DNA content of infected cells, making it impossible to determine the actual stage of the cell cycle using FACS analysis; however, expression of cyclins serves as a marker for cell cycle-associated activity. In the presence of MG132, however, cyclin A was expressed to a higher level in mock-infected G1 cells at 12 hpi. (Note: the 0 h samples in the left and right panels of Fig. 4 are duplicate samples. These were included to control for protein expression on different Western blots.) Additionally, cyclin A was expressed in wild-type- and 22/n199-infected MG132-treated cells, albeit at a lower level, indicating that cyclin A, an S-phase cyclin, is at least transiently expressed in both wild-type- and 22/n199-infected G1 cells (Fig. 4A).
In S-phase cells (mock, 0 h), cyclin A was expressed but was undetectable by 12 h post-mock infection (at this time, 59% of the cells were in G1, 40% in S, and 1% in G2/M; Fig. 4). Cyclin A was not expressed following wild-type infection at 12 hpi but was expressed in 22/n199-infected S-phase cells at this time, indicating that expression of ICP22 and/or the US1.5 protein is required to reduce cyclin A levels during HSV-1 infection of S-phase cells. In the presence of MG132, cyclin A was expressed in mock-, wild-type-, and 22/n199-infected cells at 12 hpi (Fig. 4B). Collectively, these observations indicate that cyclin A is degraded in a proteasome- and ICP22/US1.5-dependent manner specifically during infection of S-phase cells.
In cells infected in G2/M, cyclin A was expressed at the time of infection (mock, 0 h) but not at 12 hpi in mock-, wild-type-, or 22/n199-infected cells (Fig. 4C; in mock-infected cells at 12 hpi, 33% of the cells were in G1, 66% in S, and 0% in G2/M). Twelve hours after infection of cells in G2/M in the presence of MG132, cyclin A was present in mock-, wild-type-, and 22/n199-infected cells, indicating that cyclin A is degraded independent of ICP22 and/or the US1.5 protein when cells are infected in G2/M.
The kinetics of cyclin B expression in synchronized cells were similar to those of cyclin A, with two exceptions. First, cyclin B was not expressed in G1-infected cells in the presence of MG132 at 12 hpi (Fig. 4A), suggesting that wild-type- and 22/n199-infected cells do not induce cells infected during G1 to progress through S and into G2/M. Second, a low level of cyclin B was detected in cells synchronized in G2/M and infected with 22/n199 at 12 hpi (Fig. 4C). The presence of detectable cyclin B in these cells may imply that ICP22 and/or US1.5 protein expression is required to deregulate G2/M-specific expression during infection of G2/M cells. Similar results were obtained for cyclins A and B when cells were harvested at 6 hpi; however, the extent of degradation of cyclins A and B was less prominent (data not shown). Additionally, expression of cyclin A in 22/n199R-infected cells was similar to that observed in wild-type-infected cells (data not shown).
It has been reported that the intracellular localization and levels of cyclin E are unaffected in asynchronous HSV-1-infected cells (21). In these tests, two species of cyclin E were detected in mock-infected Vero cells in G1 (Fig. 4A). Because cyclin E is expressed only during late G1/early S phase in normal cells, it appears that cyclin E expression is markedly deregulated in Vero cells. In mock-infected G1 cells at 12 hpi, the faster-migrating cyclin E species (band c) was predominant. By contrast, wild-type- and 22/n199-infected G1 cells expressed both b and c forms of cyclin E seen in 0 h mock-infected cells, and form c was the predominant form. In the presence of MG132 in cells infected in G1 cells, band b was expressed under all conditions tested (Fig. 4A).
In S-phase cells at the time of infection, cyclin E was detected as two isoforms (Fig. 4B, bands b and c); however, in both mock- and wild-type-infected S-phase cells at 12 hpi, cyclin E was detected only as the fastest migrating form, band c. By contrast, in 22/n199-infected S-phase cells at 12 hpi, both bands b and c were expressed (Fig. 4B). In the presence of MG132 in S-phase-infected cells, two forms of cyclin E (bands b and c) were expressed in mock- and 22/n199-infected cells at 12 hpi (band c was more prominent in mock- than in 22/n199-infected cells), but the fast migrating form of the protein (band c) was expressed primarily in wild-type-infected S-phase cells.
Cells synchronized in G2/M expressed two forms of cyclin E at the time of infection (Fig. 4C, bands b and c). However, in mock-, wild-type-, and 22/n199-infected G2/M cells, the two slower-migrating forms of cyclin E (bands a and b) were prominent. In the presence of MG132, band b was the predominant form. Although previous reports have shown that ICP22 and/or US1.5 protein expression is not required to induce modifications of cyclins when G1 cells are infected, these results demonstrate that ICP22 and/or the US1.5 protein induces modifications of cyclins A, B, and E, directly or indirectly, specifically when S-phase Vero cells are infected.
DISCUSSION
Expression of proteins that regulate and/or are regulated by the cell cycle affect replication of DNA-containing viruses. While small DNA viruses and adenovirus utilize cell cycle-associated proteins expressed during normally regulated S phase to replicate, HSV-1 utilizes proteins expressed during multiple phases of the cell cycle. In a manner similar to IE and E regulatory proteins of small DNA viruses, HSV-1 IE proteins deregulate the expression of cell cycle-associated proteins (21, 42). In this study, we have shown that in addition to ICP0, ICP4, and ICP27, ICP22/US1.5 regulates the expression of cell cycle-associated proteins specifically when S-phase Vero cells are infected. The failure of the ICP22–/US1.5– mutant, 22/n199, to deregulate S- and G2/M-specific proteins (cyclins A, E, and B) may well be the basis for the inability of 22/n199 to replicate efficiently in S-phase-infected Vero cells. By extension, this property may also be the molecular basis for the modest replication impairment of ICP22–/US1.5– mutants in asynchronous permissive cells. Based on these findings we conclude that one function of ICP22 and/or the US1.5 protein is to deregulate the cell cycle, either directly or indirectly, when S-phase permissive cells are infected.
The replication defect(s) of 22/n199 is manifested differently in permissive and restrictive cells. ICP22–/US1.5– mutants initiate productive infection in restrictive cells (i.e., they express IE, E, and 1 L proteins and synthesize viral DNA) but complete the replication process and generate progeny virus only in the original infected cell and a few adjacent cells (35, 41). Virus spread is minimal. The most obvious defect in replication in restrictive cells is the inability to transcribe and synthesize selected 2 L proteins efficiently (38). It has therefore been proposed that the function of ICP22/US1.5 is to alter host cell gene expression to provide an environment conducive to 2 L gene expression (35, 41). A primary goal of this study was to determine whether the expression of cell cycle-associated factors during individual phases of the normal cell cycle controls ICP22–/US1.5– mutant permissivity. Similar approaches have identified S-phase cells as permissive hosts for an HSV-1 VP16 mutant (15), an adenovirus E1B 55-kDa mutant, (25) and wild-type bovine herpesvirus 4 (45). In contrast to these findings, however, we report here that cellular gene expression during any phase of the normal cell cycle was unable to complement 22/n199 in restrictive cells. This finding suggests that the dominant function associated with ICP22/US1.5 in restrictive cells is to alter expression of cellular proteins that are either not regulated by the cell cycle or expressed during multiple phases of the cell cycle.
Although ICP22–/US1.5– mutants replicate efficiently in permissive Vero, HeLa, and Hep-2 cells, the wild-type virus replicates 2 to 10 times more efficiently than ICP22–/US1.5– viruses (35, 41). Consequently, one may ask whether ICP22–/US1.5– viruses replicate less efficiently than the wild-type virus in all infected permissive cells, or only in a fraction of asynchronous cells. Here, we have established that the environment in S-phase Vero cells reduces 22/n199 plating and replication efficiency (Fig. 3), indicating that the phase of the cell cycle at the time of infection affects the efficiency of 22/n199 replication. In support of this observation, electron microscopic examination of asynchronous Vero cells infected with the wild type or 22/n199 revealed that a portion (20 to 30%) of 22/n199-infected but not wild-type-infected cells did not contain virions (data not shown). Although the stage of the cell cycle at the time of infection of these virion-free cells was not examined, the fraction of asynchronous cells in S phase is approximately 30%. Thus, a fraction of Vero cells (potentially those infected during S phase) apparently do not complete the replication process efficiently. Collectively, these results ascribe biological significance to the function of ICP22/US1.5 during wild-type virus replication in permissive cells.
ICP22 and/or the US1.5 protein is required to deregulate S and G2/M cyclins during infection of permissive S-phase cells: consequences for viral replication. The modest reduction in replication efficiency of 22/n199 in Vero cells infected during S phase suggests that ICP22 and/or the US1.5 protein functions during S phase. Although this is the first report demonstrating that ICP22 and/or the US1.5 protein has biological activity (i.e., its expression affects HSV-1 replication) specifically in S-phase permissive cells, previous studies have reported functions of ICP22 and the US1.5 protein consistent with this finding. ICP22 accumulates preferentially and is uniquely modified in HeLa cells infected during early S phase relative to G1 (10). The authors of that study also reported that ICP22 binds a 78-kDa cellular protein, p78, which accumulates specifically in physiologically normal S-phase cells and is constitutively expressed in many human tumor cell lines (37). This group also reported that ICP22 and/or the US1.5 protein reduces levels of cyclins A and B during HSV-1 infection of asynchronous HeLa cells (2). We show that cyclins A and B are degraded and cyclin E is modified either directly or indirectly in an ICP22/US1.5-dependent manner specifically when S-phase cells are infected (Fig. 4). It is possible that these properties of ICP22 and/or the US1.5 protein are at least partially responsible for the reduced replication efficiency of 22/n199 in permissive S-phase cells.
What is the function of ICP22/US1.5 during HSV-1 infection in S-phase permissive cells In a series of reports, Advani et al. suggested that an effect of ICP22/US1.5-dependent degradation of cyclins A and B is the activation of the cellular protein topoisomerase II (2-4). Since ICP22–/US1.5– mutants synthesize wild-type levels of viral DNA but do not synthesize 2 L proteins efficiently, these authors suggest that modified nonfunctional topoisomerase II reduces the number of newly replicated ICP22–/US1.5– genomes able to transcribe 2 L genes. Topoisomerase II is expressed in nontransformed cells during S phase of the normal cell cycle (26) and is also induced during HSV-1 infection (4). It is possible that in S-phase cells infected with ICP22–/US1.5– mutants, cyclin A expression precludes the activation of functional topoisomerase II and efficient expression of 2 L proteins. In the current study, Western blot analysis of the 2 L protein gC in wild-type- and 22/n199-infected S-phase Vero cells revealed that gC expression is reduced relative to wild-type-virus-infected cells or cells synchronized to G1 or G2/M and infected with 22/n199 (data not shown). By contrast, protein levels of gE, a 1 L protein, were equivalent in 22/n199-infected S-phase Vero cells relative to 22/n199-infected G1 and G2/M cells and wild-type-infected cells. Thus, like restrictive cells, the primary replication defect of 22/n199-infected S-phase Vero cells is evident as a defect in 2 L gene expression; however, whether topoisomerase II is altered differently in S-phase cells infected with 22/n199 relative to the wild type is unknown.
A potential role for cyclin E during HSV-1 replication. Identification of the role of cyclin E in HSV-1 replication is complicated by an apparent deregulation of cyclin E expression in Vero cells. Cyclin E is expressed during late G1 to early S phase in normal cells; Vero cells, however, express cyclin E during all phases of the cell cycle (Fig. 4). Indeed, this deregulation of cyclin E expression may be a factor in the immortalization of Vero cells. Since cyclin E expression or modification may also contribute to the permissivity of ICP22–/US1.5– mutants in Vero cells, the status of cyclin E in other permissive cell types is currently under investigation.
Cyclin E is expressed in different forms when S-phase Vero cells are infected with the wild-type virus or 22/n199. The faster migrating form is present in wild-type-infected S-phase cells, while a slower migrating form of the protein is also detected in 22/n199-infected S-phase cells. Since cyclin E levels are regulated by ubiquitination (30), and the slower-migrating form of cyclin E is 8 kDa (the size of ubiquitin) larger than the faster-migrating form, it is possible that the slower-migrating forms of cyclin E are ubiquitinated. In the presence of MG132, the slower-migrating species of cyclin E was the only species of cyclin E detected in all samples, suggesting that this species may be targeted for degradation via the proteasome. If this is the case, one role of ICP22 and/or the US1.5 protein may be to prevent the ubiquitination and potentially the proteolysis of cyclin E in S-phase infected cells. An active form of cyclin E may be required for its partners Cdks 1 and 2 to phosphorylate their normal targets (5). These targets include NPAT (a cellular transcription factor required for histone biosynthesis) (51), proliferating cell nuclear antigen (PCNA; required for efficient HSV-1 viral DNA replication) (6), and pRb (23). The targets of cyclin E function during cellular DNA synthesis, and it is possible that these proteins play accessory roles during HSV-1 DNA replication. Although DNA replication reaches wild-type levels in 22/n199-infected restrictive cells, it is possible that defects such as incomplete synthesis and concatemerization may contribute to the inability of the newly synthesized viral DNA to transcribe 2 L proteins. Alternatively, active Cdk2-cyclin E, Cdk1-cyclin E, Cdk1-cyclin A, Cdk2-cyclin A, or Cdk1-cyclin B may be required to phosphorylate viral targets. Indeed, Cdks modify the viral IE proteins ICP0 and ICP4 posttranslationally, and these modifications have been reported to confer specific activities on ICP0 during HSV-1 replication (16).
These studies add to the functions ascribed to ICP22 and the US1.5 protein and further link ICP22/US1.5 function to the cell cycle. Although factors expressed during S phase in permissive cells reduce 22/n199 replication, restrictive cells that are not in S phase do not complement virus replication. These properties distinguish two functions of ICP22 and the US1.5 protein or two consequences of the same function. Most studies of ICP22 and the US1.5 protein have been performed with permissive cells, and the reported functions of the proteins have not been associated with measurable effects on viral replication. The findings presented here provide biological relevance to several properties of ICP22 and the US1.5 protein, suggesting that expression of one or both of these proteins is required in S-phase permissive cells for efficient virus replication. The ability of ICP22/US1.5 to degrade or modify cyclins during infection of S-phase Vero cells may also be important for replication in restrictive cells; however, we have not observed similar degradation of cyclins A and B or modification of cyclin E by the wild-type virus in restrictive HEL cells (data not shown), indicating that an as yet unidentified role for ICP22/US1.5 exists in these cells.
REFERENCES
Ackermann, M., M. Sarmiento, and B. Roizman. 1985. Application of antibody to synthetic peptides for characterization of the intact and truncated 22 protein specified by herpes simplex virus 1 and the R325 22– deletion mutant. J. Virol. 56:207-215.
Advani, S. J., R. Brandimarti, R. R. Weichselbaum, and B. Roizman. 2000. The disappearance of cyclins A and B and the increase in activity of the G2/M-phase cellular kinase cdc2 in herpes simplex virus 1-infected cells require expression of the 22/US1.5 and UL13 viral genes. J. Virol. 74:8-15.
Advani, S. J., R. R. Weichselbaum, and B. Roizman. 2000. The role of cdc2 in the expression of herpes simplex virus genes. Proc. Natl. Acad. Sci. USA 97:10996-11001.
Advani, S. J., R. R. Weichselbaum, and B. Roizman. 2003. Herpes simplex virus 1 activates cdc2 to recruit topoisomerase II alpha for post-DNA synthesis expression of late genes. Proc. Natl. Acad. Sci. USA 100:4825-4830.
Aleem, E., H. Kiyokawa, and P. Kaldis. 2005. Cdc2-cyclin E complexes regulate the G1/S phase transition. Nat. Cell Biol. 7:831-836.
Arata, Y., M. Fujita, K. Ohtani, S. Kijima, and J. Y. Kato. 2000. Cdk2-dependent and -independent pathways in E2F-mediated S phase induction. J. Biol. Chem. 275:6337-6345.
Bagchi, S., P. Raychaudhuri, and J. R. Nevins. 1989. Phosphorylation-dependent activation of the adenovirus-inducible E2F transcription factor in a cell-free system. Proc. Natl. Acad. Sci. USA 86:4352-4356.
Balliet, J. W., J. C. Min, M. S. Cabatingan, and P. A. Schaffer. 2005. Site-directed mutagenesis of large DNA palindromes: construction and in vitro characterization of herpes simplex virus type 1 mutants containing point mutations that eliminate the oriL or oriS initiation function. J. Virol. 79:12783-12797.
Bostock, C. J., D. M. Prescott, and J. B. Kirkpatrick. 1971. An evaluation of the double thymidine block for synchronizing mammalian cells at the G1-S border. Exp. Cell Res. 68:163-168.
Bruni, R., and B. Roizman. 1998. Herpes simplex virus 1 regulatory protein ICP22 interacts with a new cell cycle-regulated factor and accumulates in a cell cycle-dependent fashion in infected cells. J. Virol. 72:8525-8531.
Cai, W., and P. A. Schaffer. 1991. A cellular function can enhance gene expression and plating efficiency of a mutant defective in the gene for ICP0, a transactivating protein of herpes simplex virus type 1. J. Virol. 65:4078-4090.
Cai, W. Z., and P. A. Schaffer. 1989. Herpes simplex virus type 1 ICP0 plays a critical role in the de novo synthesis of infectious virus following transfection of viral DNA. J. Virol. 63:4579-4589.
Carter, K. L., and B. Roizman. 1996. The promoter and transcriptional unit of a novel herpes simplex virus 1 gene are contained in, and encode a protein in frame with, the open reading frame of the 22 gene. J. Virol. 70:172-178.
Cohen, G. H., R. K. Vaughan, and W. C. Lawrence. 1971. Deoxyribonucleic acid synthesis in synchronized mammalian KB cells infected with herpes simplex virus. J. Virol. 7:783-791.
Daksis, J. I., and C. M. Preston. 1992. Herpes simplex virus immediate early gene expression in the absence of transinduction by Vmw65 varies during the cell cycle. Virology 189:196-202.
Davido, D. J., W. F. von Zagorski, W. S. Lane, and P. A. Schaffer. 2005. Phosphorylation site mutations affect herpes simplex virus type 1 ICP0 function. J. Virol. 79:1232-1243.
DeCaprio, J. A., J. W. Ludlow, J. Figge, J. Y. Shew, C. M. Huang, W. H. Lee, E. Marsilio, E. Paucha, and D. M. Livingston. 1988. SV40 large tumor antigen forms a specific complex with the product of the retinoblastoma susceptibility gene. Cell 54:275-283.
DeLuca, N. A., A. M. McCarthy, and P. A. Schaffer. 1985. Isolation and characterization of deletion mutants of herpes simplex virus type 1 in the gene encoding immediate-early regulatory protein ICP4. J. Virol. 56:558-570.
Diaz, J. J., D. Simonin, T. Masse, P. Deviller, K. Kindbeiter, L. Denoroy, and J. J. Madjar. 1993. The herpes simplex virus type 1 US11 gene product is a phosphorylated protein found to be non-specifically associated with both ribosomal subunits. J. Gen. Virol. 74:397-406.
Dyson, N., P. M. Howley, K. Munger, and E. Harlow. 1989. The human papilloma virus-16 E7 oncoprotein is able to bind to the retinoblastoma gene product. Science 243:934-937.
Ehmann, G. L., T. I. McLean, and S. L. Bachenheimer. 2000. Herpes simplex virus type 1 infection imposes a G(1)/S block in asynchronously growing cells and prevents G(1) entry in quiescent cells. Virology 267:335-349.
Everett, R. D., A. Orr, and C. M. Preston. 1998. A viral activator of gene expression functions via the ubiquitin-proteasome pathway. EMBO J. 17:7161-7169.
Geng, Y., E. N. Eaton, M. Picon, J. M. Roberts, A. S. Lundberg, A. Gifford, C. Sardet, and R. A. Weinberg. 1996. Regulation of cyclin E transcription by E2Fs and retinoblastoma protein. Oncogene 12:1173-1180.
Goldin, A. L., R. M. Sandri-Goldin, M. Levine, and J. C. Glorioso. 1981. Cloning of herpes simplex virus type 1 sequences representing the whole genome. J. Virol. 38:50-58.
Goodrum, F. D., and D. A. Ornelles. 1997. The early region 1B 55-kilodalton oncoprotein of adenovirus relieves growth restrictions imposed on viral replication by the cell cycle. J. Virol. 71:548-561.
Goswami, P. C., J. L. Roti Roti, and C. R. Hunt. 1996. The cell cycle-coupled expression of topoisomerase II during S phase is regulated by mRNA stability and is disrupted by heat shock or ionizing radiation. Mol. Cell. Biol. 16:1500-1508.
Graham, F. L., and A. J. van der Eb. 1973. A new technique for the assay of infectivity of human adenovirus 5 DNA. Virology 52:456-467.
Hilton, M. J., D. Mounghane, T. McLean, N. V. Contractor, J. O'Neil, K. Carpenter, and S. L. Bachenheimer. 1995. Induction by herpes simplex virus of free and heteromeric forms of E2F transcription factor. Virology 213:624-638.
Hobbs, W. E., II, and N. A. DeLuca. 1999. Perturbation of cell cycle progression and cellular gene expression as a function of herpes simplex virus ICP0. J. Virol. 73:8245-8255.
Koepp, D. M., L. K. Schaefer, X. Ye, K. Keyomarsi, C. Chu, J. W. Harper, and S. J. Elledge. 2001. Phosphorylation-dependent ubiquitination of cyclin E by the SCFFbw7 ubiquitin ligase. Science 294:173-177.
Lomonte, P., and R. D. Everett. 1999. Herpes simplex virus type 1 immediate-early protein Vmw110 inhibits progression of cells through mitosis and from G1 into S phase of the cell cycle. J. Virol. 73:9456-9467.
Long, M. C., V. Leong, P. A. Schaffer, C. A. Spencer, and S. A. Rice. 1999. ICP22 and the UL13 protein kinase are both required for herpes simplex virus-induced modification of the large subunit of RNA polymerase II. J. Virol. 73:5593-5604.
Ogle, W. O., and B. Roizman. 1999. Functional anatomy of herpes simplex virus 1 overlapping genes encoding infected-cell protein 22 and US1.5 protein. J. Virol. 73:4305-4315.
Poffenberger, K. L., A. D. Idowu, E. B. Fraser-Smith, P. E. Raichlen, and R. C. Herman. 1994. A herpes simplex virus type 1 ICP22 deletion mutant is altered for virulence and latency in vivo. Arch. Virol. 139:111-119.
Poffenberger, K. L., P. E. Raichlen, and R. C. Herman. 1993. In vitro characterization of a herpes simplex virus type 1 ICP22 deletion mutant. Virus Genes 7:171-186.
Post, L. E., and B. Roizman. 1981. A generalized technique for deletion of specific genes in large genomes: alpha gene 22 of herpes simplex virus 1 is not essential for growth. Cell 25:227-232.
Ren, Y., R. K. Busch, L. Perlaky, and H. Busch. 1998. The 58-kDa microspherule protein (MSP58), a nucleolar protein, interacts with nucleolar protein p120. Eur. J. Biochem. 253:734-742.
Rice, S. A., M. C. Long, V. Lam, P. A. Schaffer, and C. A. Spencer. 1995. Herpes simplex virus immediate-early protein ICP22 is required for viral modification of host RNA polymerase II and establishment of the normal viral transcription program. J. Virol. 69:5550-5559.
Schaffer, P. A., G. M. Aron, N. Biswal, and M. Benyesh-Melnick. 1973. Temperature-sensitive mutants of herpes simplex virus type 1: isolation, complementation and partial characterization. Virology 52:57-71.
Schang, L. M., A. Rosenberg, and P. A. Schaffer. 1999. Transcription of herpes simplex virus immediate-early and early genes is inhibited by roscovitine, an inhibitor specific for cellular cyclin-dependent kinases. J. Virol. 73:2161-2172.
Sears, A. E., I. W. Halliburton, B. Meignier, S. Silver, and B. Roizman. 1985. Herpes simplex virus 1 mutant deleted in the 22 gene: growth and gene expression in permissive and restrictive cells and establishment of latency in mice. J. Virol. 55:338-346.
Song, B., K. C. Yeh, J. Liu, and D. M. Knipe. 2001. Herpes simplex virus gene products required for viral inhibition of expression of G1-phase functions. Virology 290:320-328.
Tattersall, P. 1972. Replication of the parvovirus MVM. I. Dependence of virus multiplication and plaque formation on cell growth. J. Virol. 10:586-590.
Tobey, R. A., and K. D. Ley. 1970. Regulation of initiation of DNA synthesis in Chinese hamster cells. I. Production of stable, reversible G1-arrested populations in suspension culture. J. Cell Biol. 46:151-157.
Vanderplasschen, A., M. Goltz, J. Lyaku, C. Benarafa, H. J. Buhk, E. Thiry, and P. P. Pastoret. 1995. The replication in vitro of the gammaherpesvirus bovine herpesvirus 4 is restricted by its DNA synthesis dependence on the S phase of the cell cycle. Virology 213:328-340.
Whyte, P., K. J. Buchkovich, J. M. Horowitz, S. H. Friend, M. Raybuck, R. A. Weinberg, and E. Harlow. 1988. Association between an oncogene and an anti-oncogene: the adenovirus E1A proteins bind to the retinoblastoma gene product. Nature 334:124-129.
Whyte, P., N. M. Williamson, and E. Harlow. 1989. Cellular targets for transformation by the adenovirus E1A proteins. Cell 56:67-75.
Yanagi, K., A. Talavera, T. Nishimoto, and M. G. Rush. 1978. Inhibition of herpes simplex virus type 1 replication in temperature-sensitive cell cycle mutants. J. Virol. 25:42-50.
Yao, F., and P. A. Schaffer. 1995. An activity specified by the osteosarcoma line U2OS can substitute functionally for ICP0, a major regulatory protein of herpes simplex virus type 1. J. Virol. 69:6249-6258.
Yeh, L., and P. A. Schaffer. 1993. A novel class of transcripts expressed with late kinetics in the absence of ICP4 spans the junction between the long and short segments of the herpes simplex virus type 1 genome. J. Virol. 67:7373-7382.
Zhao, J., B. Dynlacht, T. Imai, T. Hori, and E. Harlow. 1998. Expression of NPAT, a novel substrate of cyclin E-CDK2, promotes S-phase entry. Genes Dev. 12:456-461.(Joseph S. Orlando, Todd L)