were labeled for two generations during expo- ... exponential culture was grown for two generations at 30°C in the presence of .... Lysozyme + 20-s sonic. 1.18.
JOURNAL OF BACTERIOLOGY, Apr. 1979, p. 245-248 0021-9193/79/04-0245/04$02.00/0
Vol. 138, No. 1
Relationship between Deoxyribonucleoside Triphosphate Pools and Deoxyribonucleic Acid Synthesis in an nrdA Mutant of Escherichia coli JOHN D. MANWARING AND JAMES A. FUCHS* Department of Biochemistry, College of Biological Sciences, University of Minnesota, St. Paul, Minnesota 55108
Received for publication 17 October 1979
A shift to 420C in an nrdA mutant causes a decrease in deoxyribonucleic acid synthesis without a concomitant decrease in deoxynucleotide triphosphate pools.
Ribonucleoside-diphosphate reductase (RDP reductase) (EC 1.17.4.1) in Escherichia coli catalyzes the reduction of the four RDPs to the corresponding deoxyribonucleoside diphosphates and thus generates all precursors for DNA synthesis (13, 18). Wechsler and Gross (22) partially characterized and mapped a group of mutants unable to replicate DNA at 420C. Among this group of mutants was E101, a mutant that was characterized as having an immediate reduction in the rate of DNA synthesis, and the mutation was designated dnaF. Fuchs et al. (11) found that E101 contained a thermolabile Bl subunit of RDP reductase and changed the designation of this mutation to nrdA. They found that incubation of either crude extract or a 90% pure preparation of this mutant B1 subunit for 2 min at 45°C essentially destroyed all activity, whereas an equivalently purified parental enzyme lost no activity. Thus, it was assumed that a shift of a culture of E101 to 420C inactivated the B1 subunit of RDP reductase which depleted deoxyribonucleotide triphosphate pools and thus arrested DNA synthesis. However, in additional studies, it was observed that the deoxyribonucleotide triphosphate pools of E101 grown at 42°C were not depleted, and we began studies to explain this discrepancy. In an attempt to correlate the abrupt change in rate of DNA synthesis with changes in the deoxyribonucleoside triphosphate pools, the nrdA mutant E101 and its parent, CR34 (22), were labeled for two generations during exponential growth at 300C with 32P, and the nucleotide pools were analyzed at 300C and for 2.5 h after a shift to 420C. All four deoxyribonucleotide pools of E101 were lower than the level of the pools of its parent during growth at 300C (Fig. 1). The nrdA mutation was transferred to different genetic backgrounds via P1 transduction, and these strains were analyzed in a like manner and were found to behave similarly
(data not shown). This finding was not unexpected because it had previously been found that in derepressed cultures of E101 and CR34 grown at 300C the RDP reductase activity of E101 was only 5% of that found in CR34 (11). Lower deoxyribonucleotide pools in E101 would be expected to decrease the rate of DNA elongation which would be compensated for by an increased number of replication forks which would increase the level of RDP reductase synthesized (8), thus allowing the cell to compensate for the defect in RDP reductase (the data in Fig. 3 suggest that there are more replication forks). However, it was surprising to find that the deoxyribonucleotide pools did not rapidly decrease after the shift to 420C. In fact, the pattern of pool changes seen in E101 is quite similar to that in CR34 for the entire 2.5-h period (Fig. 1). During this period the relative differences in pool levels observed at 300C is maintained. Similar results were found for thymine prototrophic strains harboring the same mutation (data not shown). These data are difficult to reconcile with the observation of Wechsler and Gross that E101 exhibits an immediate decrease in DNA synthesis at 420C. Extensive mapping of nrdA (10) as well as examination of revertants (4 and unpublished data) has shown that the defects of DNA synthesis and RDP reductase are caused by a single mutation. Although Dermody et al. also reported that E101 was defective in DNA synthesis, they reported a delay before DNA synthesis decreased after a shift to 420C (4). CR34 and E101 were grown exponentially under the same conditions used for nucleotide pool analysis except that no 32P was used. Samples were removed from each culture and pulsed for 3 min with [3H]thymidine. After the 3-min pulse, the samples were processed and counted (2). In CR34 the rate of DNA synthesis immediately increased after the shift to 420C, whereas in 245
246
NOTES
J. BACTERIOL.
0 HOURS AT 420C FIG. 1. Nucleoside triphosphate pools of CR34 (A) and E101 (B) after transfer from 30 to 42°C. An exponential culture was grown for two generations at 30°C in the presence of [32P]orthophosphate. The labeled culture was then shifted to a 42°C water bath (zero time). Incorporation of label was measured as
described previously (14). 0 E101 there was an immediate and significant 10 decrease in DNA synthesis (Fig. 2). The rate of DNA synthesis continued to increase at a new 0 r exponential rate at 10 min after the shift in 4 CR34. In E101, the rate of DNA synthesis decreased exponentially after a large initial drop in N ° 2 rate after the shift. Similar results were found in x other strains harboring the nrdA mutation. These results basically agree with those of CLI Wechsler and Gross (22), although the resolu0 tion in our experiments was greater because we used pulse-labeling of DNA rather than DNA 0.4 accumulation. In studying these strains harboring the nrdA -20 0 20 40 60 mutation, we have found them to exhibit a sharp 80 decrease in their rate of DNA synthesis upon MINUTES AT 42°C shifting to 420C while at the same time displayFIG. 2. In vivo DNA synthesis (pulse-labeling) in ing little or no decrease in the deoxyribonucleo- CR34 (0) and EIOI (0) before and after transfer from side triphosphate pools. Two possibilities that 30 to 42°C. Cultures were exponentially growing in may explain this apparent dichotomy are, first, media containing 10 pg of thymidine per ml and 100 RDP reductase may have a more direct effect pg of deoxyadenosine per ml at 30°C and were transon DNA synthesis than just production of de- ferred to 42°C at zero time. At indicated times 35 ,ul of culture was placed in an equal volume, at the same oxyribonucleotides. We wanted to test the pos- temperature of the above media containing [3HJthysibility that ribonucleotide reductase was part of midine (150 ,uCi/ml). After 3 min 50-pl samples were the DNA replication complex (1, 19) and that treated and counted as previously described (2). changes in the mutant enzyme at 420 would inactivate the replication complex. For T4-in- deoxyribonucleoside triphosphates and that mufected E. coli cells, it has been reported that the tations in genes coding for these enzymes could T4 DNA replication complex contains at least directly affect DNA synthesis (3, 5, 15, 24). A some of the enzymes needed for synthesis of second possibility is that there may be a localQ
VOL. 138, 1979
NOTES
ized pool of DNA precursors immnediately available to the replication fork that could conceivably be seriously lowered without immediately affecting the total cellular concentration of the deoxynucleotides. Good evidence for this type of compartmentalization of the thymidine triphosphate pool in eucaryotic cells has been recently reported (9). This localized pool would be dependent upon RDP reductase being in close proximity to the replication site. To test the first possibility, we made CR34 and E101 permeable to deoxyribonucleoside triphosphates by ether treatment, and we assayed DNA synthesis (20). In this system, DNA synthesis is dependent on added deoxyribonucleotides and not the catalytic activity of RDP reductase. P1 grown on a nalA nrdA strain KK395 was used to transduce a polA strain, P3478, to nalA nrdA (KK1083) and nalA nrdA' (KK1082). Cultures of KK1082 and KK1083 were grown exponentially at 30°C and then shifted to 42°C. Samples were harvested at 30°C and at various times after the shift, and ether treated (20). These ether-treated cells were assayed for in vitro DNA synthesis in the presence of the four deoxyribonucleoside triphosphates (20) at both 30 and 42°C. The in vitro rate of DNA synthesis is two- to threefold greater at 42°C than at 30°C for both strains when grown at 30°C or even after 2 h after the shift to 42°C (Fig. 3). Thus, the effect of the nrdA mutation on DNA synthesis is not present when deoxyribonucleoside triphosphates are supplied in vitro. Figure 3 also indicates that the mutant has a 2.5-fold greater specific activity of DNA synthesis than its parent when grown at 30°C. This would be expected if the decreased deoxyribonucleotide pools found for the mutant decreased the rate of DNA elongation in vivo and caused increased initiation of DNA replication as has been reported for suboptimal concentrations of TTP (16). The parent increases the in vitro specific activity of DNA synthesis after a shift to 42°C and thus faster
247
growth conditions, whereas the mutant decreases its specific activity of in vitro DNA synthesis suggesting that either DNA replication forks in the mutant are being degraded or there is a failure to initiate new replication forks while protein synthesis continues to increase (Fig. 3). This experiment failed to indicate any direct effect of the nrdA mutation on DNA synthesis. However, caution must be taken in this interpretation because the in vivo complex may not be preserved during treatment of these cells. To test the possibility of the RDP reductase enzyme being situated near the replication complex, thus allowing for a localized pool of deoxyribonucleotides, we sought evidence for a physical association of the enzymes and cellular structural components. Such an association has been previously suggested (6, 21). Folded genomes (12) were assayed and found to have
z
a-
E
'E
._
0
0
E
_ _
0
_
_
. _ _
120 30 60 90 CULTURE TIME AT 420C (min)
FIG. 3. Rates of in vitro DNA synthesis at 30 and 42°C of ether-treated cells of KK1082 (nrdA+) and KK1083 (nrdA) that had been exponentially grown at 30°C and transferred to 42°C. Cell samples that had been ether treated at 0, 30, 60, 90, and 120 min after the temperature shift were assayed for DNA synthesis at 30 and 42°C (20). Symbols: KK1082 assayed at 30°C (0), and at 42°C (0); K1083 assayed at 30°C (A) and at 42°C (A).
TABLE 1. RDP reductase activity in fractions of lysed cells Activity in supernatant Activity in pellet (43,500 Strain Strain
T~3reatment Treatment
in Sp act crude extract
(43,500 x g) % of crude exSp act tract
KK1081" KK1081
Lysozyme Lysozyme + 20-s sonic
0.90 1.18
55 42
x g)
% of crude ex-
Sp act
tract
1.28 1.02
42 11
2.70 0.35
disruption 12 0.35 53 0.77 0.79 80-s sonic disruption 23 99.2 66 28.9 44.4 Lysozyme KK516b aAn E. coli F- polA deoB nalA strain. bA strain of E. coli that harbors a temperature-inducible A phage carrying the genes coding for RDP reductase (7). The strain was grown and induced as described (7).
KK1081a
248 NOTES higher RDP reductase specific activity than the soluble fraction (data not shown). Cells were lysed with lysozyme (23), centrifuged at 1,200 x g for 10 min to remove whole cells, and centrifuged at 43,500 x g for 60 min. The resulting pellet contained large amounts of RDP reductase activity (Table 1), suggesting that RDP reductase may be associated with the rapidly sedimenting fraction. Sonic disruption of whole cells or lysozyme extracts greatly decreased the specific activity of the 43,500 x g pellet. In a strain that greatly overproduces RDP reductase, a significant amount of the activity was also found in the rapidly sedimenting fraction. However, we were unable to find conditions in which we could further characterize this association. An association of the enzyme with a structural component could position the enzyme in the close proximity of the replication site and thus allow a localized pool of deoxynucleotides that is immediately utilized in replication as has been suggested by Reddy et al. (17) and Fridland (9). This work was supported by Public Health Service grant GM 20884 from the National Institute of General Medical Sciences to J.A.F. J.D.M. was supported by Public Health Service training grant GM 00345 from the National Institute of General Medical Sciences.
LITERATURE CITED 1. Alberts, B., and R. Sternglanz. 1977. Recent excitement in the DNA replication problem. Nature (London) 269: 655-661. 2. Bollum, F. J. 1966. Filter paper disk techniques for assaying radioactive macromolecules, p. 296-300. In G. L. Cantoni and D. R. Davies (ed.), Procedures in nucleic acid research. Harper & Row, Publishers, New York. 3. Collinsworth, W. L., and C. K. Mathews. 1974. Biochemistry of DNA-defective amber mutants of bacteriophage T4. IV. DNA synthesis in plasmolyzed cells. J. Virol. 13:908-915. 4. Dermody, J. J., G. J. Bourguignon, P. D. Fogelsong, and R. Sternglanz. 1974. Nalidixic acid-sensitive and resistant modes of DNA replication in Escherichia coli. Biochem. Biophys. Res. Commun. 61:1340-1347. 5. Dicou, L., and N. R. Cozzarelli. 1973. Bacteriophage T4-directed DNA synthesis in toluene-treated cells. J. Virol. 12:1293-1302. 6. Eriksson, S. 1975. Ribonucleotide reductase from Escherichia coli. Demonstration of a highly active form of the enzyme. Eur. J. Biochem. 56:289-294. 7. Eriksson, S., B. Sjoberg, S. Hahne, and 0. Karlstrom. 1977. Ribonucleoside diphosphate reductase from Escherichia coli. An immunological assay and a novel purification from an over-producing strain lysogenic for
J. BACTERIOL. phage A dnrd. J. Biol. Chem. 252:6132-6138. 8. Filpula, D., and J. A. Fuchs. 1978. Regulation of the synthesis of ribonucleoside diphosphate reductase in Escherichia coli: specific activity of the enzyme in relationship to perturbations of DNA replication. J. Bacteriol. 135:429-435. 9. Fridland, A. 1973. DNA precursors in eukaryotic cells. Nature (London) New Biol. 243:105-107. 10. Fuchs, J. A., and H. 0. Karlstrom. 1976. Mapping of nrdA and nrdB in Escherichia coli K-12. J. Bacteriol. 128:810-814. 11. Fuchs, J. A., H. 0. Karlstrom, H. R. Warner, and P. Reichard. 1972. Defective gene product in dnaF mutant of Escherichia coli. Nature (London) New Biol. 238:69-71. 12. Kornberg, T., A. Lockwood, and A. Worcel. 1974. Replication of the Escherichia coli chromosome with a soluble enzyme system. Proc. Natl. Acad. Sci. U.S.A. 71:3189-3193. 13. Larsson, A., and P. Reichard. 1966. Allosteric effects and substrate specificity of the ribonucleoside diphosphate reductase system from Escherichia coli B. Biochim. Biophys. Acta 113:407-408. 14. Manwaring, J. D., and J. A. Fuchs. 1977. Nucleoside triphosphate pools in minicells of Escherichia coli. J. Bacteriol. 130:960-962. 15. North, T. W., M. E. Stafford, and C. K. Mathews. 1976. Biochemistry of DNA-defective mutants of bacteriophage T4. VI. Biological functions of gene 42. J. Virol. 17:973-982. 16. Pritchard, R. H., and K. G. Lark. 1964. Induction of replication by thymine starvation at the chromosome origin in Escherichia coli. J. Mol. Biol. 9:288-307. 17. Reddy, G. P. V., A. Singh, M. E. Stafford, and C. K. Mathews. 1977. Enzyme associations in T4 phage DNA precursor synthesis. Proc. Natl. Acad. Sci. U.S.A. 74: 3152-3156. 18. Reichard, P. 1967. The biosynthesis of deoxyribose. Ciba lectures in biochemistry. John Wiley & Sons, Inc., New York. 19. Schekman, R., A. Weiner, and A. Kornberg. 1974. Multienzyme systems of DNA replication. Science 186: 987-993. 20. Vosberg, H., and H. Hoffman-Berling. 1971. DNA synthesis in nucleotide-permeable Escherichia coli cells. I. Preparation and properties of ether-treated cells. J. Mol. Biol. 58:739-753. 21. Warner, H. R. 1973. Properties of ribonucleoside diphosphate reductase in nucleotide-permeable cells. J. Bacteriol. 115:18-22. 22. Wechsler, J. A., and J. D. Gross. 1971. Escherichia coli mutants temperature-sensitive for DNA synthesis. Mol. Gen. Genet. 113:273-284. 23. Wickner, W., R. Schekman, K. Geider, and A. Kornberg. 1973. A new form of DNA polymerase III and a copolymerase replicate a long, single-stranded primertemplate. Proc. Natl. Acad. Sci. U.S.A. 70:1764-1767. 24. Wovcha, M. G., P. K. Tomich, C. Chiu, and G. R. Greenberg. 1973. Direct participation of dCMP hydroxymethylase in synthesis of bacteriophage T4 DNA. Proc. Natl. Acad. Sci. U.S.A. 70:2196-2200.
