cellular distribution of the small 3.4S DNA polymerase remains unchanged. ..... Fransler,B., and L. A. Loeb. 1972. Sea urchin nuclear. DNA polymerase. IV.
JOURNAL OF VIROLOGY, Nov. 1975, p. 1095-1100 Copyright ( 1975 American Society for Microbiology
Vol. 16, No. 5 Printed in U.S.A.
DNA Polymerases in Polyoma Virus-Infected Mouse Kidney Cells ULRIKE WINTERSBERGER* AND ERHARD WINTERSBERGER Institut fur Krebsforschung der Universitat Wien, A-1090 Vienna, Austria*; Physiologisch-Chemisches Institut der Universitat Wurzburg, D-87 Wurzburg, Federal Republic of Germany; and Departement de Biologie Moleculaire, Universite de Geneve, Switzerland Received for publication 7 July 1975
Infection of arrested mouse kidney cells by polyoma virus results in the induction of the cellular 6-8S DNA polymerase activity. Levels of this enzyme increase two- to threefold in the cytoplasm but seven- to tenfold in nuclei and nuclear extracts, suggesting an accumulation of the enzyme in the nucleus. Experiments using the inhibitor of DNA synthesis, fluordeoxyuridine, indicate that this accumulation is linked to active DNA synthesis. The activity and cellular distribution of the small 3.4S DNA polymerase remains unchanged.
Polyoma virus infection of nondividing mouse cells leads to an induction of host DNA synthesis concomitant with the onset of viral DNA replication (7, 24). At the same time activity of cellular enzymes involved in the synthesis of DNA increases. Earlier studies had shown that, in addition to enzymes catalyzing various steps in the production of deoxynucleoside triphosphates, DNA polymerase, which was then measured in total cell homogenates, increases after infection (10, 13, 14, 23). In the meantime it was demonstrated by several laboratories that cells from higher organisms generally contain two types of DNAdependent DNA polymerases (1, 3, 4, 20, 25), a large so-called 6-8S enzyme (DNA polymerase a; see reference 22 for nomenclature) which is predominantly found in the cytoplasm, and a small 3.4S enzyme (DNA polymerase ,B) present in the nucleus and in the cytoplasm. A third distinct enzyme is localized in mitochondria. In various cases of induced DNA synthesis, such as in cells entering the S phase of the cell cycle or during liver regeneration, the large enzyme was found to increase in activity in the cytoplasm whereas little change in the concentration of the small DNA polymerase was observed (2, 5, 21). This led to the tentative assumption that DNA polymerase a is involved in DNA replication. We have studied the process of DNA polymerase induction in polyoma virus-infected mouse kidney cells and have observed that also in this system only DNA polymerase a increases in activity. In addition evidence was obtained for an accumulation of this enzyme in nuclei synthesizing DNA.
MATERIALS AND METHODS Cell culture. Kidney cells from 10-day-old inbred swiss mice were grown to confluence in plastic petri dishes (100-mm diameter) as described by Winocour (27). Each plate was infected with 0.4 ml of a polyoma virus suspension containing about 10' PFU/ml. Two hours after addition of the virus, cells were covered with 10 ml of Eagle medium per dish without serum and incubated in a CO2 incubator at 37 C. Twentyfour hours after infection, the medium was removed and cells were washed twice with 5 ml of ice-cold phosphate-buffered saline. Mock-infected cells were treated similarly except that 0.4 ml of medium was used in place of the virus suspenision. If infection was carried out in the presence of 5-fluordeoxyuridine, the medium contained 15 ug of the drug per ml (18). To release cells from the fluordeoxyuridine block, the medium was removed 24 h after infection, and cells were washed twice with warm medium and covered with 10 ml of medium containing 5 jg of thymidine per ml. After 10 min of incubation cells were extracted as described below. Simian virus 40-transformed mouse kidney cells were grown by serial passage in Eagle medium containing 5% fetal calf serum. They were used when the cell density was about 107 cells/dish. Under these conditions cells are nearly confluent, but they are not inhibited in DNA synthesis and growth as the density of these cells goes up to about 5 x 107/dish. Cell fractionation. Cells from 5 to 10 plates (containing about 107 cells each) were employed for every experiment. They were removed from the plates and lysed with buffer A (50 mM Tris-hydrochloride [pH 7.5], 25 mM KCl, 5 mM MgCl,) containing 0.25 M sucrose and 1% Triton X-100 (0.2 ml of buffer was used per dish). The suspension was layered on top of 2 volumes of buffer A containing 1 M sucrose, and nuclei were centrifuged through this cushion (10 min,
WINTERSBERGER AND WINTERSBERGER
1,500 x g, Sorval rotor HB 4, 0 C). The cytoplasmic fraction was removed from the top and saved, and the nuclei were washed twice with buffer A containing 0.25 M sucrose. By inspection under a phase contrast microscope these nuclei appeared intact and free from, contaminating whole cells or cytoplasm. To test DNA polymerase activity in nuclei, these were suspended in 0.1 M potassium-phosphate buffer (pH 7.5) and 10 Ml of the suspension was added to the assay mixture described below. Nuclear extracts were prepared by suspending nuclei in 0.2 M potassium-phosphate buffer (pH 7.5) containing 1 M NaCl (0.1 ml of buffer was used for nuclei obtained from one plate of cells). DNA was sheared and enzyme extraction was achieved by 20 times passage of the suspension through a 20-gauge needle with a plastic syringe. A clear extract containing over 90% of the nuclear DNA polymerase activity was obtained by centrifugation for 30 min at 40,000 x g. Nuclear extracts were dialyzed against buffer lacking NaCl to reduce the ionic strength of the solution for the enzyme test. DNA polymerase assay. DNA polymerase activity was determined by measuring the incorporation of ['HITMP into acid-insoluble material essentially as described earlier (28). The incorporation medium (0.125 ml) contained: 0.05 M Tris-hydrochloride buffer (pH 7.4 or 8.8); 10 mM MgCl,; 1 mM dithiothreitol; 50 MM each of dATP, dGTP, and dCTP; 50 gM [3H NITP (64 counts/min per pmol); 5 mM phosphoenolpyruvate; 10 Mg of pyruvate kinase; 12.5 Mg of activated salmon sperm DNA; and 10 to 50 Ag of protein from the enzyme solutions. After 30 min of incubation at 35 C, reactions were stopped by rapid cooling in ice and addition of 2 ml of cold 5% trichloroacetic acid containing 0.02 M sodium-pyrophosphate. Precipitates were filtered onto Whatman GF/C filters, washed with trichloroacetic acid and ethanol-ether (1:1), dried, and counted. Assays of all cell and enzyme fractions were dependent upon externally added DNA and the four deoxyribonucleoside triphosphates. Tests were linear with time for at least 60 min and with enzyme protein up to 50 Mg/assay. Protein was determined by the method of Lowry et al.
Aliquots of 10 Ml were used to assay for DNA polymerase activity at pH 7.4 and 8.8 as described above, except that after incubation 100-Ml aliquots of the reaction mixture were pipetted onto Whatman GF/C filters (25-mm diameter) which were processed further as described earlier (28). The activity of the marker enzymes was determined in parallel.
RESULTS Infection of stationary mouse kidney cells with polyoma virus leads to about a threefold increase of total DNA polymerase activity in the cell homogenate, which reaches its maximum around 24 h after infection (23). We separately analyzed the cytoplasm, nuclei, and nuclear extracts for DNA polymerase activity. To distinguish between enzyme a and ,B we made use of the different activities at pH 7.4 and 8.8 and of the distinct sedimentation properties in sucrose gradients of the large and the small DNA polymerase. Enzyme levels were determined 24 h after infection and compared to those of uninfected or mock-infected controls. As shown in Table 1, infection resulted in a twofold increase of DNA polymerase activity in the cytoplasm. In nuclei and nuclear extracts, however, a much more pronounced enhancement could be measured. Here the polymerase activity exceeded the control levels seven- to tenfold (five independent experiments). The first indication as to which enzyme, a or A, was responsible for the increased activity came from the analysis of enzyme activities at the two pH values, 7.4 and 8.8. Whereas DNA polymerase a is more active at pH 7.4 than at pH 8.8, the small 3.4S enzyme shows higher activity at pH 8.8 (5) or equal activities at both pH values (this study). Considering these properties the results of Table 1 can be interpreted to mean the following: the increase of DNA (15). Specific activities of DNA polymerase are ex- polymerase activity in the cytoplasm as well as pressed as nanomoles of TMP incorporated per milli- in nuclei is due to the large polymerase. Moregram of protein per 30 min. over at the onset of DNA synthesis this enzyme Sucrose gradients. Sedimentation analyses were is found associated with the nucleus. This carried out in 12-ml linear gradients of 5 to 20% sucrose in 0.2 M potassium-phosphate (pH 7.5)-i mM conclusion is further substantiated by sedimendithiothreitol. Cytoplasm and nuclear extracts were tation analyses in sucrose gradients shown in dialyzed against 0.2 M potassium-phosphate buffer Fig. 1. Both large and small enzyme are present (pH 7.5) prior to sedimentation analyses, and 0.5 to 1 in the cytoplasm of uninfected cells but only the ml were layered onto the gradients and centrifuged for faster sedimenting DNA polymerase rises upon 24 h at 35,000 rpm and 0 C in the SW40 rotor of infection (Fig. 2a and b). Nuclear extracts of the Spinco centrifuge. Protein (3 mg) of the cytoplas- uninfected or mock-infected cells, in contrast, mic fraction or 1.5 mg of protein of the nuclear extract contain only small amounts of the large enzyme plus 50 Mg of alcoholdehydrogenase (sedimentation besides the 3.4S polymerase and infection recoefficient 7.5S) and 150 Mg of malate dehydrogenase sults in a six- to eightfold increase of enzyme a (sedimentation coefficient 4.4S) were applied onto each gradient, the latter enzymes serving as marker in accord with the data of Table 1. Under our proteins. After centrifugation, gradients were frac- conditions of cell fractionation 5% of the total tionated from the bottom into 25 to 28 equal fractions. cellular DNA polymerase activity is found in
DNA POLYMERASES IN POLYOMA-INFECTED CELLS
VOL. 16, 1975
TABLE 1. DNA polymerase activity in cell fractions of polyoma virus-infected kidney cells Sp acta in: Determinants
Uninfected or mockinfected cells Infected cells SV40-transformed cells' Cells infected in presence of 5-fluordeoxyuridine 10 min after release from fluordeoxyuridine block
0.63 1.36 0.66
0.43 0.85 0.45
0.80 1.54 0.53
0.53 0.95 0.38
1.41 1.30 0.88
0.89 0.81 0.55
Nanomoles of TMP incorporated per milligram of protein per 30 min. 'SV40, Simian virus 40.
nuclei of uninfected cells but at least 25% is present in nuclei of infected cells. Analyses of a the separated polymerases at the two pH values 500 1000 also support the conclusion drawn from the results reported in Table 1: the activity of DNA 250 500 polymerase a is about twofold higher at pH 7.4 than at pH 8.8, whereas DNA polymerase B is equally active at both pH values. Chang et al. 1500 (5) observed the enzyme being more active at b 8.8 than at pH 7.4. The difference in our pH 500 1000 results may be due to the use of other buffer substances. a 250 L 500 The sedimentation analyses were performed ~~~~~~ under conditions of high ionic strength (0.2 M C~~~~~~~~ Ex phosphate buffer) which favor the dissociation x of high molecular aggregates of DNA polymer750 .n 1500 c ~~~~~~~~cc ase a to the 6-8S form. Our analysis under these a 0 conditions resulted in an S value of 7.5 to 8.0 for 500 io1000 this enzyme. There is no difference in the 500 250 sedimentation behavior of DNA polymerases from uninfected and infected mouse cells. Any other differences that could possibly exist be1500 tween the 6-8S polymerases of uninfected and d infected mouse cells, however, would not be 500 1000 revealed by our experiments. High levels of the large DNA polymerase were 500 250 also found in nuclei of simian virus 40-transformed mouse kidney cells (Fig. lc). In fact, 0 5 10 15 20 25 0 5 10 15 20 25 specific activities of DNA polymerase in nuclei Fraction number of these transformed cells were similar to those FIG. 1. Sedimentation of DNA polymerases from of nuclei from lytically infected cells from the cytoplasm and nuclear extracts in sucrose gradients. same origin. Enzyme levels in the cytoplasm Cytoplasm and nuclear extracts of polyoma-infected were nearly twofold higher in transformed commouse kidney cells were prepared and analyzed by pared to lytically infected mouse cells (Table 1 sedimentation in sucrose gradients. Gradients were and Fig. lc). aligned with respect to the position of the markers. (a) Figure 2 depicts the time course of the DNA Mock-infected cells; (b) infected cells; (c) simian polymerase induction. In agreement with the cells virus 40-transformed mouse kidney cells; (d) infected in the presence of 5-fluordeoxyuridine. Sym- observations of Weil and Kara (23), enzyme bols: *, activity at pH 7.4; 0, activity at pH 8.8. Note activity starts to increase between 12 and 16 h the different scale on the left and the right ordinate. postinfection. The appearance of DNA polym1500
Cytoplasm 75S 44S
Nucleus 75S 44S
WINTERSBERGER AND WINTERSBERGER
ingly, however, the accumulation of the enzyme in the nucleus is significantly reduced under the 70 influence of fluordeoxyuridine. This finding has substantiated been by sedimentation analyses of nuclear extracts (Fig. ld). A release of cells 60 from the fluordeoxyuridine block by addition of thymidine, which results in a rapid and synchronous resumption of DNA synthesis (18), is 50 1/ - X ,' accompanied by an immediate increase of the >1 amount of DNA polymerase a in nuclei (Table > ,' > 1). Already 10 min after addition of thymidine, 40 / fK A DNA polymerase activity in nuclei reaches a / / > C% /, level identical with that measured in cells u -30 -/ 5f infected in absence of the drug and this level - _____remains / -a---__________ constant for several hours. -
DISCUSSION 10 , 0
Hours after infection FIG. 2. Time course of the induction of DNA polymerases in polyoma virus-infected mouse kidney cells. Confluent mouse kidney cells were infected with polyoma virus. Five plates were used for each time point to prepare cytoplasm and nuclei. Specific activities of DNA polymerase in the cytoplasm and in nuclear suspensions were determined. Solid lines, pH 7.4; broken lines, pH 8.8; 0 and 0, cytoplasm; A and A, nuclei.
Much fundamental information on the mechanism of polyoma virus DNA replication has been obtained recently (6, 9, 11, 12, 16, 17, 19, 26). Our experiments are a first attempt to learn about the enzymes involved in this process. We have shown in this report that infection of nondividing mouse kidney cells with polyoma virus results in the induction of the large cellular 6-8S DNA polymerase (DNA polymerase a) which sets in between 12 and 16 h after infection. This suggests that DNA polymerase a .
may function iin the replication of cellularDNA which in this system iS turned on simultaneously with the synthesis of viral DNA. As an example of rapidly dividing cells of the same origin we have chosen simian virus 40-transerase a in the nucleus occurs together or slightly formed mouse kidney cells a great proportion of earlier than the activity enhancement in the which is in the S phase and therefore synthesizcytoplasm. Comparison of DNA polymerase ing DNA. As expected the level of DNA polymactivities at the two pH values gave results in erase a in the cytoplasm as well as in the accordance with those reported in Table 1 and nucleus of these cells is high and comparable to Fig. 1: although the DNA polymerase activity of that of lytically infected mouse kidney cells. nuclei is identical at pH 7.4 and 8.8 before the These observations are in accord with reports of induction (mainly due to the 3.4S enzyme), other authors on the behavior of DNA polymerduring induction activity rises more drastically ases in the cell cycle of tissue culture cells (5, at the lower pH value than at the higher one 21) or in regenerating liver and hepatomas (2). (due to the increase of 6-8S DNA polymerase in In each of these cases the cytoplasmic 6-8S the nucleus). DNA polymerase increases in activity at the If infection of confluent mouse kidney cells is time of DNA synthesis, whereas the nuclear carried out in the presence of 5-fluordeoxyuri- 3.4S enzyme, as in our experiments, remains at dine, viral and cellular DNA replication are a fairly constant level. Since the induction of strongly inhibited and only early viral functions the cellular polymerase activity is a prerequisite are expressed. The induction of cellular en- for the replication of viral DNA our results zymes involved in DNA replication is unim- indicate, though they do not prove, an involvepaired and takes place just as in cells infected in ment of DNA polymerase a in the synthesis of the absence of the nucleoside derivative (18). As polyoma virus DNA. expected, we found that the activity of DNA In addition to the increase of enzyme activity polymerase a reaches the same level in the in the cytoplasm, our data show an accumulacytoplasm of cells infected in the presence as in tion of DNA polymerase a in the nucleus. We the absence of the drug (see Table 1). Interest- found that whereas the increase of DNA polym-
DNA POLYMERASES IN POLYOMA-INFECTED CELLS
VOL. 16, 1975
erase activity in the cytoplasm is two- to threefold, in nuclei it is seven- to tenfold. This finding excludes the possibility that our results are merely caused by a contamination of our nuclear preparations by cytoplasm. It this were the case one would expect an about equal rise of enzyme activity in both cytoplasm and nuclei. Elevated nuclear levels of DNA polymerase a were found by Baril et al. (2) in regenerating liver cells and in hepatomas. A slight enhancement is also apparent in the S phase of L cells and HeLa cells (5, 21), and Fansler and Loeb (8) reported a reversible association of DNA polymerase with nuclei during the cell cycle of sea urchin embryos. The presence of high concentrations of DNA polymerase a in the nucleus of virus-infected and virus-transformed cells appears to be coupled to active DNA synthesis. Significantly lower enzyme levels were measured in nuclei of infected cells in which DNA synthesis is inhibited by fluordeoxyuridine, whereas the increase of DNA polymerase activity in the cytoplasm is unimpaired. The block exerted by fluordeoxyuridine may not be absolute; in particular, initiation of DNA replication might still occur although further synthesis soon comes to a halt. This initiation could suffice, however, to cause some accumulation of DNA polymerase a in the nucleus as indicated by our experiments. The fact that a release of cells from the fluordeoxyuridine block results in a rapid increase of the level of nuclear DNA polymerase a to that measured in cells infected in the absence of the drug supports the assumption of a link between DNA synthesis and the appearance of DNA polymerase a in the nucleus. DNA polymerase a might therefore in fact be a nuclear enzyme which is bound and retained more strongly in nuclei synthesizing DNA. Its almost exclusive occurrence in the cytoplasm of resting cells could be due to a very rapid and efficient leakage of the enzyme during the preparation of nuclei in aqueous media. ACKNOWLEDGMENTS This work was carried out during a visit in the laboratory of R. Weil at the Department of Molecular Biology, University of Geneva. We are most grateful to Roger Weil for his kind hospitality and his support (Swiss National Foundation grant no. 3097.73). We thank him and Hans Turler for many stimulating discussions and N. Bensemmane for help with the cell culture. We are also grateful to P. Reichard for comments on the manuscript. U. W. thanks the Austrian Ministery of Science and Research for a grant, from the Swiss-Austrian exchange program. LITERATURE CITED 1. Baril, E. F., 0. E. Brown, M. D. Jenkins, and J. Laszlo. 1971. DNA-polymerase with rat liver ribosomes and
6. 7. 8.
smooth membranes. Purification and properties of the enzyme. Biochemistry 10:1981-1992. Baril, E. F., M. D. Jenkins, 0. E. Brown, J. Laszlo, and H. P. Morris. 1973. DNA-polymerases I and II in regenerating rat liver and Morris hepatomas. Cancer Res. 33:1187-1193. Chang, L. M. S., and F. J. Bollum. 1971. Low molecular weight DNA polymerase in mammalian cells. J. Biol. Chem. 246:5835-5837. Chang, L. M. S., and F. J. Bollum. 1972. Low molecular weight DNA polymerase from rabbit bone marrow. Biochemistry 11:1264-1272. Chang, L. M. S., McK. Brown, and F. J. Bollum. 1973. Induction of DNA polymerase in mouse L cells. J. Mol. Biol. 74:1-8. Crawford, L. V., C. Syrett, and A. Wilde. 1973. The replication of polyoma DNA. J. Gen. Virol. 21:515-521. Dulbecco, R., L. H. Hartwell, and M. Vogt. 1965. Induction of cellular DNA synthesis by polyoma virus. Proc. Natl. Acad. Sci. U.S.A. 53:403-410. Fransler, B., and L. A. Loeb. 1972. Sea urchin nuclear DNA polymerase. IV. Reversible association of DNA polymerase with nuclei during the cell cycle. Exp. Cell Res. 75:433-441. Francke, B., and T. Hunter. 1974. In vitro polyoma DNA synthesis: discontinuous chain growth. J. Mol. Biol. 83:99-121. Hartwell, L. H., M. Vogt, and R. Dulbecco. 1965. Induction of cellular DNA synthesis by polyoma. II. Increase in the rate of enzyme synthesis after infection with polyoma virus in mouise embryo cells. Virology
27:262-272. 11. Hunter, T., and B. Francke. 1974. In vitro polyoma DNA synthesis: characterization of a system from infected 3T3 cells. J. Virol. 13:125-139. 12. Hunter, T., and B. Francke. 1974. In vitro polyoma DNA synthesis: involvement of RNA in discontinuous chain growth. J. Mol. Biol. 83:123-130. 13. Kara, J., and R. Weil. 1967. Specific activation of the DNA synthesizing apparatus in contact inhibited mouse kidnev cells by polvoma virus. Proc. Natl. Acad. Sci. U.S.A. 57:63-70. 14. Kit, S., D. R. Dubbs, and P. M. Frearson. 1966. Enzymes of nucleic acid metabolism in cells infected with polyoma virus. Cancer Res. 26:638-646. 15. Lowry, 0. H., N. J. Rosebrough, A. L. Farr. and R. .J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. 16. Magnusson, G., V. Pigiet, E. L. Winnacker, R. Abrams, and P. Reichard. 1973. RNA-linked short DNA fragments during polvoma replication. Proc. Natl. Acad. Sci. U.S.A. 70:412-415. 17. Magnusson, G., E. L. Winnacker, R. Eliasson, and P. Reichard. 1972. Replication of polvoma DNA in isolated nuclei. II. Evidence for semi-conservative replication. J. Mol. Biol. 72:539-552. 18. Petursson, G., and R. Weil. 1968. A study on the mechanism of polvoma-iniduced activation of the cellular DNA-synthesizing apparatus: synchronization bv FdU of virus-induced DNA-svnthesis. Arch. Gesamte Virusforsch. 24:1-29. 19. Pigiet, V., R. Eliasson, and P. Reichard. 1974. Replication of polvoma DNA in isolated nuclei. III. The nucleotide sequence at the RNA-DNA junction of nascent strands. J. Mol. Biol. 84:197-216. 20. Sedwick, W. D.. T. Shu-Fong Wang, and D. Korn. 1972. Purification and properties of nuclear and cytoplasmic DNA polymerases from huiman KB cells. J. Biol. Chem. 247:5026-5033. 21. Spadari, S., and A. Weissbach. 1974. Interrelation between DNA synthesis and various DNA polymerase
23. 24. 25.
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activities in synchronized HeLa cells. J. Mol. Biol. 86:11-20. Spadari, S., and A. Weissbach. 1975. RNA-primed DNA synthesis: specific catalysis by HeLa cell DNA polymerase a. Proc. Natl. Acad. Sci. U.S.A. 72:503-507. Weil, R., and J. Kara. 1970. Polyoma "tumor antigen": an activator of chromosome replication? Proc. Natl. Acad. Sci. U.S.A. 67:1011-1017. Weil, R., M. Michel, and G. Ruschmann. 1965. Induction of cellular DNA synthesis by polyoma virus. Proc. Natl. Acad. Sci. U.S.A. 53:1468-1475. Weissbach, A., A. Schlabach, B. Fridlender, and A.
Bolden. 1971. DNA polymerases from human cells. Nature (London) 231:167-170. 26. Winnacker, E. L., G. Magnusson, and P. Reichard. 1972. Replication of polyoma DNA in isolated nuclei. I. Characterisation of the system from mouse fibroblast 3T6 cells. J. Mol. Biol. 72:523-537. 27. Winocour, E. 1963. Purification of polvoma virus. Virology 19:158-168. 28. Wintersberger, U., and E. Wintersberger. 1970. Studies on DNA polymerases from yeast. 1. Partial purification and properties of two DNA polymerases from mitochondria-free cell extracts. Eur. J. Biochem. 13:11-19.