Peter Glazer, and Leena Kalghatgi for help. We also thank Harvey. Friedman and ... U.-K. Freese, B. Fleckenstein, and H. zur Hausen, J. Virol. 61:119-124, 1987; ...
JOURNAL OF VIROLOGY, Apr. 1987, p. 1291-1295 0022-538X/87/041291-05$02.00/0 Copyright © 1987, American Society for Microbiology
Vol. 61, No. 4
Isolation and Characterization of Phosphonoacetic Acid-Resistant Mutants of Human Cytomegalovirus RICHARD T. D'AQUILAl* AND WILLIAM C. SUMMERS2
Departments of Therapeutic Radiology,'2 Molecular Biophysics and Biochemistry,2 Human Genetics,2 and Medicine,1 Yale University School of Medicine, New Haven, Connecticut 06510-8039 Received 8 September 1986/Accepted 12 December 1986
Mutants of the human cytomegalovirus (HCMV) that were 6- to 13-fold more resistant to phosphonoacetic acid than the wild-type HCMV (Towne) were isolated. Extracts from mycoplasma-free, mutant-infected cells had phosphonoacetate-resistant DNA polymerase activity in vitro. This strongly suggests that the selected mutations are in the HCMV DNA polymerase genes of these viruses.
Although a few genes have been mapped by biochemical the human cytomegalovirus (HCMV) genome (11, 32, 34, 36, 37, 45, 46), no locus has yet been genetically defined, and only a few mutant strains of HCMV have been isolated and characterized (2, 25, 30, 49, 50). None of the mutants of HCMV so far isolated has drug-resistant HCMV DNA polymerase activity, although temperature-sensitive salt stimulation of viral DNA polymerase activity in vitro has been demonstrated in one mutant HCMV strain (25). In contrast, the DNA polymerase gene of herpes simplex virus type 1 (HSV-1) was mapped by conventional genetics and by marker rescue and gene transfer studies with wellcharacterized temperature-sensitive and drug-resistant mutants with viral DNA polymerases that were biochemically distinguishable from the wild-type viral polymerase (1, 3, 4, 6-10, 12, 14, 15, 17, 18, 21, 22, 26-28, 38-40). We report here the isolation and characterization of phosphonoacetic acid (PAA)-resistant mutants of HCMV (Towne). These mutants have PAA-resistant DNA polymerase activity in vitro under conditions that strongly favor the viral polymerase. Because PAA, an inorganic PP1 analog, binds directly to herpesvirus polymerases (12, 24, 29, 31), the data presented here strongly suggest that the mutations underlying PAA resistance in these viruses are in the HCMV DNA polymerase gene. Although the degree of PAA resistance of these viruses has not been quantitated well in these studies, the data suggest that the degree of PAA resistance of these HCMV mutants will be adequate for drug resistance marker transfer studies to genetically map the HCMV DNA polymerase gene and its PAA-binding site. Isolation of phosphonacetate-resistant mutants. The wildtype parental virus HCMV (Towne) was provided by H. Friedman and S. Plotkin (University of Pennsylvania, Philadelphia) and was plaque purified by limiting dilution three times to ensure a homogeneous stock. Primary human fibroblasts used for virus propagation and assay were prepared from neonatal foreskins and grown in minimum essential medium (GIBCO Laboratories, Grand Island, N.Y.) supplemented with 10% fetal calf serum (GIBCO), penicillin G (100 U/ml), and streptomycin (100 ,ug/ml). To select for spontaneous viral mutants that were resistant to PAA, wild-type HCMV (Towne) was serially passaged at a multiplicity of infection (MOI) of 0.1 in human foreskin
fibroblasts in increasing concentrations of PAA (Abbott Laboratories, North Chicago, Ill.). This process may involve the sequential selection of mutants that are progressively better adapted to growth in the presence of the drug, possibly due to more than a single mutation (22). After nine passages over 6 months, the progeny of infection in the presence of 100 ,ug of PAA per ml were plaque-purified twice by limiting dilution in the presence of 100 ,ug of PAA per ml. All subsequent studies reported here were done on the first three doubly plaque-purified mutants isolated (PAAr-8B, PAAr-8C, PAAr-8D). A fourth PAA-resistant mutant (PAAr13A) was isolated but has not yet been characterized in detail. High-titered, frozen stocks were made of all these independently derived mutant virus isolates on the third passage after the second plaque purification. Virions of the wild-type and first three PAA-resistant HCMV (Towne) isolates were each prepared from the supernatant fluids of cultures infected at an MOI of 0.1 3 days after a complete cytopathic effect was evident by ultracentrifugation of the clarified culture supernatant through a cushion of 20% sucrose in 10 mM Tris hydrochloride (pH 8)-0.15 M NaCl. The virion pellet was suspended in 10 mM Tris hydrochloride (pH 8)-i mM EDTA (TE). After digestion in 1% sodium dodecyl sarcosinate-0.5% sodium dodecyl sulfate-300 ,ug of proteinase K (20 U/mg; BoehringerMannheim Biochemicals, Indianapolis, Ind.) per ml at 37°C for 4 h, virion DNA was extracted with phenol and chloroform-isoamyl alcohol, dialyzed extensively against TE, and then precipitated with ethanol. Restriction endonuclease digestion of virion DNA with three different enzymes confirmed that these mutants were highly related, if not identical, to wild-type HCMV (Towne) (data not shown). Characterization of PAA resistance in tissue culture. The PAA sensitivity of these virus isolates was determined by a modification of the procedure of Dreesman and BeyneshMelnick (13) described by Coen et al. (7). Figure 1 presents the means of three experiments in which the effects of a range of PAA concentrations (0 to 200 xug/ml) on plaque formation by the parental HCMV (Towne) and the mutants PAAr-8C and PAAr-8D are compared. The result of one experiment of the plaquing efficiency of the mutant PAAr-8B from 75 to 200 ,ug/ml is also shown in Fig. 1. The 50% effective dose of PAA for the mutants was 6- to 13-fold higher than that of the parental virus. The efficiency of plaquing of PAAr-8C and PAAr-8D was statistically significantly different from that of the parental wild-type HCMV
means on
*
Corresponding author. 1291
J. VIROL.
NOTES
1292
100' cm
80'
Ea)
60
cn
40.-
0~=3
Cr2 0-
20
0
50
150 100 [PAA] (mcg./ml.)
200
FIG. 1. Efficiency of plaquing of wild-type and mutant (PAAr8B, PAAr-8C, and PAAr-8D) HCMV (Towne) viruses in the presence of PAA. The 50% effective dose for each virus is as follows: wild type, 7 ,ug/ml; PAAr-8C, 40 jig/ml; PAAr-8D, 60 ,ug/ml; PAAr8B, 90 jig/ml. Symbols: A, wild type; A, PAAr-8B; x, PAA'-8C; *, PAAr-8D.
(Towne) (P < 0.05; two-tailed Student t test) at 75 and 100 Vig of PAA per ml. In preliminary experiments performed before the isolation of these PAAr mutants, the effect of PAA on wild-type HCMV (Towne) was confirmed to be at the level of viral DNA synthesis. DNA was isolated from wild-type HCMV (Towne) that was grown in [3H]thymidine (New England Nuclear Corp., Boston, Mass.) and either the presence or absence of 50 ,ug of PAA per ml. Analysis of cesium chloride equilibrium density gradients revealed no incorporation of label into DNA at the viral density in the presence of PAA (data not shown). Based on the data presented in Fig. 1, these PAAr mutants of HCMV (Towne) do not appear to be as highly resistant as some HSV-1 PAAr mutants (6, 7); however, the degree of resistance is of practical importance to determine if the PAAr phenotype of these HCMV mutants can be selected in a mixed infection with wild-type virus to allow genetic mapping of the site of the mutation(s) conferring PAA resistance. By mixing various amounts of wild-type and mutant virus stocks and duplicate plating in the presence and absence of PAA, it was shown that as little as 1 PFU of each of the mutants tested could be detected in the presence of 1.5 x 10' PFUs of wild-type HCMV (Towne) in 100 ,ug of PAA per ml (data not shown). While their exact degree of PAA resistance remains uncertain, these mutants are distinguishable enough from the wild type to be useful for drug resistance marker transfer studies. Characterization of PAA resistance in cellular extracts in vitro. To demonstrate that the PAA resistance of the mutant viruses correlated with alterations in HCMV DNA polymerase activity, DNA polymerase assays were done in vitro by using extracts of virus-infected and uninfected cells. Extracts were prepared by the method described by Nishiyama et al. (35) from either HCMV-infected cells (3 days after infection at an MOI of 2) or uninfected cells (at 70%
confluence). Assay conditions (primer template, salt concentration, and buffer) were chosen that would maximize the HCMV and minimize the host cell polymerase activity of these unfractionated extracts. In a direction comparison, the uninfected cell extract yielded greater incorporation into a gapped, duplex DNA primer template (activated calf thymus
DNA [51]) than into an oligodeoxynucleotide-primed, singlestranded DNA (phage M13 [44]) primer template, but the wild-type HCMV-infected cell extract used the M13 primer template more efficiently (data not shown). This is in keeping with earlier findings of others that the partially purified HCMV polymerase, in contrast to host cell polymerases, used an oligodeoxynucleotide-primed, single-stranded DNA template more efficiently than a gapped, duplex DNA primer template (Table 1 in reference 23). In fact, the preference for the M13 primer template over the activated calf thymus DNA primer template observed here was five-fold greater at salt concentrations that stimulate HCMV polymerase activity and that inhibit the major host cell polymerases (see below). Therefore, in all these experiments the M13 primer template was used. An additional degree of selectivity may have been provided by the phosphate buffer system (35), which inhibits the salt-stimulated mammalian DNA polymerase beta (48). Primed M13 single-stranded DNA was prepared by annealing 17-mer M13 sequencing primer [5'(GTAAAACGAC GGCCAGT)3'; New England BioLabs, Inc., Beverly, Mass.] to single-stranded M13 DNA (44) in 10 mM Tris hydrochloride (pH 7.5)-10 mM MgCl2 at a ratio of 1:100 (wt/wt) by slowly cooling from 80°C to room temperature. The in vitro polymerase assays were performed by the method described by Nishiyama et al. (35) and incorporation of [3H]dTTP ([methyl-3H]dTTP; 79.6 Ci/mmol; New England Nuclear) into 5% trichloroacetic acid-insoluble material was measured. Each assay was done in duplicate, and background incorporation (0 min of incubation) was subtracted from each assay result. Prior studies of the partially purified HCMV DNA polymerase (19, 20, 23, 35) and unfractionated extracts of HCMV-infected cells (33) demonstrated marked inhibition of host cell DNA polymerase activity and stimulation of the HCMV DNA polymerase activity at high salt concentrations in vitro. Wild-type-, PAAr-8C-, and PAAr-8D-infected cell extracts showed maximal polymerase activity on the M13 primer template at 25 to 75 mM ammonium sulfate (Fig. 2).
200-
4-
co 0
100
0
50 100 [Ammonium sulfate] (mM)
FIG. 2. Effect of ammonium sulfate concentration (in millimolar) on the in vitro DNA polymerase activity of cellular extracts on
primed single-stranded (M13) DNA template. Of note is the relative increase in activity in all infected cell extracts, except for that of PAAr-8B, from their individual base lines in the absence of ammonium sulfate at 25 to 50 mM (or 25 to 75 mM) ammonium sulfate. The essentially background level activity of the uninfected, logarithmically growing cell extract at and above 75 mM ammonium sulfate is also of note. The error bars represent the standard error of the mean. Symbols: l, uninfected; A, wild type; A, PAAr-8B, X, PAAr-8C; *, PAAr-8D.
NOTES
VOL. 61, 1987
It is of interest that the PAAr-8B-infected cell extract lacked such salt stimulation (Fig. 2). In the absence of ammonium sulfate, the uninfected, logarithmically growing cell extract had only 1.7 to 8% of the polymerase activity seen in any of the HCMV-infected cell extracts (data not shown), and only the HCMV-infected cell extracts consistently showed polymerase activity above background levels at and above 75 mM ammonium sulfate (Fig. 2). These assay conditions maximize differences between HCMV and host cell polymerases. The use of unfractionated extracts, however, does not allow a clear distinction of viral polymerase activity from the host cell polymerase activity that is induced by HCMV infection (23, 50). DNA polymerase assays were done over a range of PAA concentrations with the M13 primer template at a high salt concentration (80 mM ammonium sulfate). Higher levels of polymerase activity, relative to the base line in the absence of PAA, were seen at every PAA concentration tested in each of the mutant-infected cell extracts compared with wild-type-infected and uninfected cell extracts (Fig. 3). These PAA-resistant mutant HCMV strains had PAAresistant DNA polymerase activity in cellular extracts in vitro under conditions that strongly favor the viral polymerase over the host cell polymerases. The relatively small differences in in vitro DNA polymerase activity between wild-type-infected and mutant-infected cell extracts may suggest that the viral polymerases of these mutants are not highly resistant to PAA. However, because these unfractionated extracts included HCMV-induced host cell polymerases as well as the viral polymerase and because in vivo viral density DNA could not be detected in wild-type HCMV in the presence of PAA (see above), much of the in vitro polymerase activity at high PAA concentrations may be due to PAA-resistant cellular polymerase(s), despite the attempts to utilize selective assay conditions. An accurate assessment of the relative resistances of these viruses therefore cannot be drawn from these in vitro data. Because mycoplasma contamination has also been reported to induce PAA-resistant, salt-stimulated polymerase activity in cellular extracts (33), all cells and virus stocks were tested for evidence of mycoplasma with the two most sensitive methods available (42). A simple cytochemical staining technique, using the specific DNA-binding benzamidine fluorochrome Hoechst compound no. 33258 (Polysciences, Inc.), was used as described by others (41) with unstained, fixed positive and negative control slides (Flow Laboratories, Inc., McLean, Va.) stained in parallel with experimental slides. Uninfected HCMV-permissive cells lacked any cytoplasmic fluorescence which is suggestive of mycoplasma contamination (data not shown). When virus stocks were plated on either permissive or nonpermissive cells (BHK cells) at an MOI of 2, however, cytopathic effect occurred; and coarse fluorescence, possibly due to cell debris, viral DNA, or both (41), made interpretation difficult (data not shown). Therefore a uridine/uracil incorporation ratio (43), using [5,6-3H]uridine (40 Ci/mmol) and [5,63H]uracil (44 Ci/mmol; New England Nuclear) labeling of parallel cultures in vivo, was determined for the primary human foreskin fibroblasts (both uninfected and infected at an MOI of 2 from the same virus stocks used in making the infected cell extracts). Previous work has documented that mammalian cells, unless contaminated by mycoplasma, incorporate greater than 200-fold more uridine than uracil into RNA (42, 43). Others have also shown that HCMV infection does not alter the uridine/uracil incorporation ratio (33). All of the primary human fibroblasts and virus stocks used here
1293
A. 100' 80 .5
60
.XaI-R
40
20 0o 0 1
50
100
[PM] (mcg./ml.)
B.
100 80-
.5
60-
t5 co
40200-
[PM] (mcg./ml.)
Co iooj 80
.Z
60 X
40
20
20
0I 0
50
100
[PM] (mcg./ml.) FIG. 3. Effect of PAA concentration on the in vitro DNA polyactivity of HCMV (Towne)-infected cell extracts on a primed single-stranded (M13) DNA template. The data are expressed relative to the activity in the absence of PAA, and each mutant-infected cell extract is compared with the same wild-typeinfected cell extract and uninfected, logarithmically growing cell extract. The data for wild-type- and PAAr-8C-infected cell extracts are duplicate assays from individual experiments. The data for the uninfected cell extract and the PAAr-8B- and PAAr-8D-infected cell extracts represent two sets of duplicate assays from two separate experiments each. The absolute levels of polymerase activity of the uninfected cell extract were only minimally above that of the background (data not shown). The error bars represent the standard error of the mean. Symbols: al uninfected; A, wild type; A, merase
PAAr-8B;
X,
PAAr-8C;
*,
PAAr-8D.
1294
J. VIROL.
NOTES
had a ratio of specific activity of [3H]uridine/[3H]uracil of greater than 400 (data not shown), which effectively excludes mycoplasma contamination (43) of cells or virus stocks as a confounding variable in these experiments. The biochemical characterization of the mutant polymerase activity in unfractionated cellular extracts described here excludes the possibility that the PAA resistance of these mutant viruses resulted from changes in the cellular entry or disposition of PAA, because such factors would not be operative in cell lysates in vitro. The extensive evidence that PAA (and its analog phosphonoformate) binds directly to the polymerases of HSV-1 (12, 31), the herpesvirus of turkeys and Marek's disease herpesvirus (29), the salmon herpesvirus (47), and HCMV (Towne) (20, 24, 35) and competes with PPi for binding (12, 29) suggests that the drug resistance mutations in these viruses are in the HCMV DNA polymerase gene. Genetic confirmation of the genomic site of the mutation(s) will be necessary, because induction of a PAAresistant host cell polymerase activity by these mutants or a mutation in an HCMV-encoded enzyme other than the polymerase might also underly the PAA resistance based on the data presented here. In vivo differences in viral DNA synthesis have not been determined here because this would not conclusively establish that the HCMV DNA polymerase gene, and not another HCMV gene, is the site of the mutation responsible for PAA resistance. Rather, we have attempted to map the site of the PAA resistance mutation by DNA-mediated drug resistance marker transfer. The genomic location of the HCMV polymerase gene has been tentatively identified in the Towne strain based on nucleic acid hybridization studies (R. D'Aquila and W. C. Summers, 10th International Herpesvirus Workshop, 1985). Results of the mixing experiments (described above) demonstrate that the PAA resistance of these mutants is a useful selectable marker for genetic studies. Therefore, the focus of our efforts has been to simultaneously confirm that the PAA resistance in these mutants is due to a HCMV DNA polymerase gene mutation and that the HCMV DNA polymerase gene maps to the genomic region with biochemical homology to polymerases of other herpesviruses by a marker transfer experiment. Cotransfections of an HCMV (Towne) DNA fragment (limited to the region of biochemical homology that is the likely physical site of the HCMV polymerase gene) cloned from one of these HCMV (Towne) PAA-resistant mutants with intact, infectious HCMV (Towne) wild-type DNA to transfer the PAA resistance phenotypic marker to recombinant progeny are in progress in our laboratory. In addition to genetic confirmation of the location of the HCMV DNA polymerase locus, future work may define the PPi (and PAA) binding site of the HCMV polymerase by comparison of the nucleotide sequence and crystal structure of the polymerases of these mutants to that of the wild-type HCMV (Towne) polymerase. Because most HSV-1 mutants with an altered sensitivity to PPi analogs also have altered sensitivities to nucleoside analogs and aphidicolin (6-9, 12, 15, 21), the HCMV polymerase deoxynucleotide triphosphate binding site may also be characterized by comparative sequence and crystallographic study of the polymerases of these mutants and wild-type HCMV (Towne). Mapping of the mutation(s) in the polymerase of PAAr-8B that is responsible for the lack of stimulation of viral polymerase activity in vitro by high salt concentrations (Fig. 2) might allow identification of the unique structural features of the polymerases of herpesviruses associated with salt stimulation (20, 23, 35, 48). Other HCMV genes may also be mapped by using these mutants, including the 3' to 5' exonuclease
activity that has been copurified with the HCMV DNA polymerase (35) and the HCMV major DNA-binding protein gene (5, 16). We thank Wilma P. Summers, Saumyen Sarkar, Marcia Lewis, Peter Glazer, and Leena Kalghatgi for help. We also thank Harvey Friedman and Stanley Plotkin for providing the virus and Abbott Laboratories for PAA. This work was supported by Public Health Service grants CA-16038, CA-09259, and AM-01423 from the National Institutes of Health. ADDENDUM
The putative HCMV (AD169) DNA polymerase gene has recently been localized by nucleotide sequence and transcriptional analysis (R. Heilbronn, G. Jahn, A. Burkle, U.-K. Freese, B. Fleckenstein, and H. zur Hausen, J. Virol. 61:119-124, 1987; T. Kouzarides, A. T. Bankier, S. C. Satchwell, K. Weston, P. Tomlinson, and B. G. Barrell, J. Virol. 61:125-133, 1987). The genomic region of HCMV (Towne) identified in our studies as the site of homology to the polymerase genes of other herpesviruses (cited above) appears to map to the same location as this putative HCMV (AD169) DNA polymerase gene. LITERATURE CITED 1. Aron, G. M., D. J. M. Purifoy, and P. A. Schaffer. 1975. DNA synthesis and DNA polymerase activity of herpes simplex type 1 temperature-sensitive mutants. J. Virol. 16:498-507. 2. Biron, K. K., J. A. Fyfe, S. C. Stanat, L. K. Leslie, J. B. Sorrell, C. U. Lambe, and D. M. Coen. 1986. A human cytomegalovirus mutant resistant to the nucleoside analog 9-{[2 hydroxy-1(hydroxymethyl)ethoxy]methyl}guanine (BW B759U) induces reduced levels of BW B759U triphosphate. Proc. Natl. Acad. Sci. USA 83:8769-8773. 3. Chartrand, P., C. S. Crumpacker, P. A. Schaffer, and N. M. Wilkie. 1980. Physical and genetic analysis of the herpes simplex virus DNA polymerase locus. Virology 103:311-326. 4. Chartrand, P., N. D. Stow, M. C. Timbury, and N. M. Wilkie. 1979. Physical mapping of paar mutations of herpes simplex virus type 1 and type 2 by intertypic marker rescue. J. Virol. 31:265-276. 5. Chiou, H. C., S. K. Weller, and D. M. Coen. 1985. Mutations in the herpes simplex virus major DNA binding protein gene leading to altered sensitivity to DNA polymerase inhibitors. Virology 145:213-226. 6. Coen, D. M., D. P. Aschman, P. T. Gelep, M. J. Retondo, S. K. Weller, and P. A. Schaffer. 1984. Fine mapping and molecular cloning of mutations in the herpes simplex virus DNA polymerase locus. J. Virol. 49:236-247. 7. Coen, D. M., H. E. Fleming, L. K. Leslie, and M. J. Retondo. 1985. Sensitivity of arabinosyladenine-resistant mutants of herpes simplex virus to other antiviral drugs and mapping of drug hypersensitivity mutations to the DNA polymerase locus. J. Virol. 53:477-488. 8. Coen, D. M., P. A. Furman, D. P. Aschman, and P. A. Schaffer. 1983. Mutations in the herpes simplex virus DNA polymerase gene conferring hypersensitivity to aphidicolin. Nucleic Acids Res. 11:5287-5297. 9. Coen, D. M., P. A. Furman, P. T. Gelep, and P. A. Schaffer. 1982. Mutations in the herpes simplex virus DNA polymerase gene can confer resistance to 9-p-D-arabinofuranosyladenine. J. Virol. 41:909-918. 10. Crumpacker, C. S., L. E. Schnipper, P. N. Kowalsky, and D. M. Sherman. 1982. Resistance of herpes simplex virus to adenine arabinoside and E-5-(2-bromovinyl)-2'-deoxyuridine: a physical analysis. J. Infect. Dis. 146:167-172. 11. DeMarchi, J. M. 1981. Human cytomegalovirus DNA: restriction enzyme cleavage maps and map locations for immediateearly, early and late RNAs. Virology 114:23-38.
VOL. 61, 1987
12. Derse, D., K. F. Bartow, and Y.-C. Cheng. 1982. Characterization of the DNA polymerases induced by a group of herpes simplex virus type 1 mutants selected for growth in the presence of phosphonoformic acid. J. Biol. Chem. 257:10251-10260. 13. Dreesman, G. R., and M. Beynesh-Melnick. 1967. Spectrum of human cytomegalovirus complement-fixing antigens. J. Immunol. 99:1106-1114. 14. Furman, P. A., D. M. Coen, M. H. St. Clair, and P. A. Schaffer. 1981. Acyclovir-resistant mutants of herpes simplex virus type 1 express altered DNA polymerase or reduced acyclovir phosphorylating activities. J. Virol. 40:936-941. 15. Gibbs, J. S., H. C. Chiou, J. D. Hall, D. W. Mount, M. J. Retondo, S. K. Weller, and D. M. Coen. 1985. Sequence and mapping analyses of the herpes simplex virus DNA polymerase gene predict a C-terminal substrate binding domain. Proc. Natl. Acad. Sci. USA 82:7969-7973. 16. Gibson, W., T. L. Murphy, and C. Roby. 1981. Cytomegalovirus infected cells contain a DNA binding protein. Virology. 111:251-262. 17. Hay, J., H. Moss, A. T. Jamieson, and M. C. Timbury. 1976. Herpesvirus proteins: DNA polymerase and pyrimidine deoxynucleoside kinase activities in temperature-sensitive mutants of herpes simplex virus type 2. J. Gen. Virol. 31:65-73. 18. Hay, J., and J. Subak-Sharpe. 1976. Mutants of herpes simplex type 1 and 2 that are resistant to phosphonoacetic acid and induce altered DNA polymerase activities in infected cells. J. Gen. Virol. 31:145-148. 19. Hirai, K., T. Furakawa, and S. A. Plotkin. 1976. Induction of DNA polymerase in WI-38 and guinea pig cells infected with human cytomegalovirus. Virology 70:251-255. 20. Hirai, K., and Y. Watanabe. 1976. Induction of alpha-type DNA polymerases in human cytomegalovirus infected WI-38 cells. Biochim. Biophys. Acta 447:328-329. 21. Honess, R. W., D. J. M. Purifoy, D. Young, R. Gopal, N. Cammack, and P. O'Hare. 1984. Single mutations at many sites within the DNA polymerase locus of herpes simplex viruses can confer hypersensitivity to aphidicolin and resistance to phosphonoacetic acid. J. Gen. Virol. 65:1-17. 22. Honess, R. W., and D. H. Watson. 1977. Herpes simplex virus resistance and sensitivity to phosphonoacetic acid. J. Virol. 21:584-600. 23. Huang, E.-S. 1975. Human cytomegalovirus. III. Virus-induced DNA polymerase. J. Virol. 16:298-310. 24. Huang, E.-S. 1975. Human cytomegalovirus. IV. Specific inhibition of virus-induced DNA polymerase activity and viral DNA replication by phosphonoacetic acid. J. Virol. 16:1560-1565. 25. Ihara, S., K. Hirai, and Y. Watanabe. 1978. Temperaturesensitive mutants of human cytomegalovirus: isolation and partial characterization of DNA-minus mutants. Virology 84:218-221. 26. Jofre, J. T., P. A. Schaffer, and D. S. Parris. 1977. Genetics of resistance to phosphonoacetic acid in strain KOS of herpes simplex virus type 1. J. Virol. 23:833-836. 27. Knipe, D. M., W. T. Ruyechan, and B. Roizman. 1979. Molecular genetics of herpes simplex virus. III. Fine mapping of a genetic locus determining resistance to phosphonoacetate by two methods of marker transfer. J. Virol. 29:698-704. 28. Knopf, K. W., E. R. Kaufman, and C. S. Crumpacker. 1981. Physical mapping of drug-resistance mutations define an active center of the herpes simplex virus DNA polymerase enzyme. J. Virol. 39:746-757. 29. Leinbach, S. S., J. M. Reno, L. F. Lee, A. F. Isbell, and J. A. Boezi. 1976. Mechanism of phosphonoacetate inhibition of herpesvirus-induced DNA polymerase. Biochemistry 15:426-430. 30. Maeda, F., S. Ihara, and Y. Watanabe. 1979. Morphogenesis of nuclear inclusions and virus capsids in HEL cells infected with temperature-sensitive mutants of human cytomegalovirus. J. Gen. Virol. 44:419-432. 31. Mao, J. C.-H., and E. E. Robishaw. 1975. Mode of inhibition of
NOTES
32. 33. 34.
35.
36.
37.
38.
39. 40. 41.
42. 43.
44.
45. 46. 47.
48. 49. 50.
51.
1295
herpes simplex virus DNA polymerase by phosphonoacetate. Biochemistry 14:5475-5479. McDonough, S. H., and D. H. Spector. 1983. Transcription in human fibroblasts permissively infected by human cytomegalovirus. Virology 125:31-46. Miller, R. L., and F. Rapp. 1976. Distinguishing cytomegalovirus, mycoplasma, and cellular DNA polymerases. J. Virol. 20:564-569. Mocarski, E. S., L. Pereira, and N. Michael. 1985. Precise localization of genes on large animal virus genomes: use of lambda gtll and monoclonal antibodies to map the gene for a cytomegalovirus protein family. Proc. Natl. Acad. Sci. 82:1266-1270. Nishiyama, Y., K. Maeno, and S. Yoshida. 1983. Characterization of human cytomegalovirus induced DNA polymerase and the associated 3'-to 5', exonuclease. Virology 124:221-231. Nowak, B., A. Gmeiner, P. Sarnow, A. J. Levine, and B. Fleckenstein. 1984. Physical mapping of human cytomegalovirus genes: identification of DNA sequences coding for a virion phosphoprotein of 71Kda and a viral 65Kda polypeptide. Virology 134:91-102. Pande, H., S. W. Baak, A. D. Riggs, B. R. Clark, J. E. Shively, and J. A. Zaia. 1984. Cloning and physical mapping of a gene fragment coding for a 64-kilodalton major late antigen of human cytomegalovirus. Proc. Natl. Acad. Sci. USA 81:4965-4969. Purifoy, D. J. M., and M. Beynesh-Melnick. 1975. DNA polymerase induction by DNA-negative temperature-sensitive mutants of herpes simplex virus type 2. Virology 68:374-386. Purifoy, D. J. M., and K. L. Powell. 1977. Identification of the herpes simplex virus DNA polymerase gene. Nature (London) 269:621-623. Purifoy, D. J. M., and K. L. Powell. 1977. Herpes simplex virus DNA polymerase as the site of phosphonoacetate sensitivity: temperature-sensitive mutants. J. Virol. 24:470-477. Russell, W. C., C. Newman, and D. H. Williamson. 1975. A simple cytochemical technique for demonstration of DNA in cells infected with mycoplasma and viruses. Nature (London) 253:461-462. Schneider, E. L., and E. J. Stanbridge. 1975. Comparison of the methods for the detection of mycoplasma contamination of cell cultures: a review. In Vitro 11:20-34. Schneider, E. L., E. J. Stanbridge, and C. J. Epstein. 1974. Incorporation of 3H-uridine and 3H-uracil into RNA-a simple technique for the detection of mycoplasma contamination of cultured cells. Exp. Cell Res. 84:311-318. Schrier, P. H., and R. Cortese. 1979. A fast and simple method for sequencing DNA cloned in the single stranded bacteriophage M13. J. Mol. Biol. 129:169-172. Stenberg, R. M., D. R. Thomsen, and M. F. Stinski. 1984. Structural analysis of the major immediate-early gene of human cytomegalovirus. J. Virol. 49:190-199. Stinski, M. F., D. R. Thomsen, R. M. Stenberg, and L. C. Goldstein. 1983. Organization and expression of the immediateearly genes of human cytomegalovirus. J. Virol. 46:1-14. Suzuki, S., T. Kimura, and M. Saneyoshi. 1986. Characterization of DNA polymerase induced by salmon herpesvirus, Oncorhynchus masou virus. J. Gen. Virol. 67:405-408. Weissbach, A. 1981. Cellular and viral induced eukaryotic polymerases, p. 67-86. In The enzymes, vol. XIV. Academic Press, Inc., New York. Yamanishi, K., and F. Rapp. 1977. Temperature-sensitive mutants of human cytomegalovirus. J. Virol. 24:416-418. Yamanishi, K., and F. Rapp. 1979. Induction of host DNA synthesis and DNA polymerase by DNA-negative temperaturesensitive mutants of human cytomegalovirus. Virology 94:237-241. Yoshida, S., T. Kondo, and T. Ando. 1974. Multiple molecular species of cytoplasmic DNA polymerase from calf thymus. Biochim. Biophys. Acta 353:463-474.
