©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Functional Compartmentation of dCTP Pools
PREFERENTIAL UTILIZATION OF SALVAGED DEOXYCYTIDINE FOR DNA REPAIR IN HUMAN LYMPHOBLASTS (*)

(Received for publication, September 13, 1994; and in revised form, October 28, 1994)

Yi-Zheng Xu Peng Huang William Plunkett (§)

From the Department of Clinical Investigation, The University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The utilization of dCTP derived from de novo synthesis through ribonucleotide reductase in exponentially growing CCRF-CEM cells was compared with the metabolic fate of dCTP produced by the salvage pathway. Exogenous dCyd was not effectively incorporated into replicating DNA; instead, dCTP derived from ribonucleotide reductase (labeled by [5-^3H]Cyd) was the main precursor for that purpose, apparently because of functional compartmentation of the dCTP pool in these cells. Studies of the metabolic route of incorporation of exogenous [5-^3H]dCyd into DNA of growing CCRF-CEM cells demonstrated that it was mainly incorporated through the DNA repair pathway. Incorporation of [5-^3H]dCyd into DNA of synchronized cell populations was maximal in G(1) cells, whereas [^3H]dThd incorporation occurred predominantly in S phase cells. When cellular DNA was density labeled by incubation with BrdUrd, repaired DNA, which was less dense than replicated DNA, was preferentially labeled by [5-^3H]dCyd. In contrast, replicated DNA was labeled by both [^3H]dThd and [5-^3H]Cyd. The DNA-damaging agents methylmethanesulfonate, ultraviolet irradiation, and -irradiation inhibited [^3H]dThd incorporation, whereas they stimulated the accumulation of [5-^3H]dCyd in DNA. Based on these results, we propose that the dCTP pool is functionally compartmentalized in growing CCRF-CEM cells. dCTP derived from the salvage pathway is utilized predominantly for DNA repair, whereas the de novo pathway supplies dCTP for DNA replication.


INTRODUCTION

Replicative DNA synthesis consumes the major portion of cellular deoxynucleoside triphosphates (dNTPs), whereas other processes such as repair of damaged DNA and deoxynucleotidyl intermediates of lipid metabolism probably place lesser demands on the cellular pools. Evidence from metabolic studies indicates that ribonucleotide reductase-mediated de novo synthesis of deoxynucleotides is tightly coupled to replicative DNA synthesis (Nicander and Reichard, 1983; Mathews and Ji, 1992; Reddy and Fager, 1993). Indeed, multiprotein complexes containing enzymes, including ribonucleotide reductase, that participate in both dNTP synthesis and DNA replication have been isolated and characterized (Noguchi et al., 1983; Harvey and Pearson, 1988; Hammond et al., 1989; Wu et al., 1994). This concept that the de novo pathway for dNTP synthesis is linked with DNA replication is further supported by indications that DNA is replicated at structurally distinct sites in the nucleus (Nakamura, et al., 1986; Mills et al., 1989; Cox and Laskey, 1991; Horzak et al., 1993; Coverley and Laskey, 1994). Kinetic studies of ribonucleoside uptake into the DNA of whole cells indicate that dNTP pools derived from de novo pathways are likely to be rather small and turn over rapidly (Nicander and Reichard, 1983). If the dNTP pool utilized for DNA replication derived from de novo synthesis is localized to the region surrounding a replication center, it is reasonable to ask whether dNTPs derived from different sources, such as the salvage pathways, could also be used for DNA replication.

It is clear from investigations in many cell types that [5-^3H]dCyd is a poor substrate for labeling replicating DNA (Plagemann et al., 1978; Cohen et al., 1983; Karle et al., 1983; Nicander and Reichard, 1983; Balzarini et al., 1984; Taljanidisz et al., 1986; Leeds and Mathews, 1987; Sasvari-Szekely et al., 1989; Xu and Plunkett, 1992; 1993). This stands in contrast to [5-^3H]Cyd which specifically labels dCMP in replicating DNA and to [^3H]dThd, which has been taken as the standard for measuring this process (Reichard, 1988). Incorporation of [5-^3H]dCyd and [^3H]dThd become comparable only in cells that are not active in DNA replication (Holmberg et al., 1988) or when de novo synthesis of dCDP is inhibited (Nicander and Reichard, 1983).

Relatively little is known of the metabolic fate of dNTPs derived from the salvage of deoxynucleosides. Evidence exists that salvaged dCyd is utilized in the formation of thymidylate used in replicative DNA synthesis in cell lines (Jackson, 1978; Xu and Plunkett 1992). Insight into the utilization of the dCTP pool derived from the salvage pathway for purposes other than replicative DNA synthesis was provided by Spyrou and Reichard(1987, 1989) who demonstrated that dCTP precursors for deoxyliponucleotide synthesis were derived from salvaged dCyd. Although some studies have indicated that labeled dCyd may be used to measure DNA repair (Snyder, 1984; Elliott and Downes, 1986; McKenna and McKelvey, 1986; Holmberg et al., 1988), it was only recently suggested that dCTP derived from the salvage pathway might be used as a specific substrate for DNA repair (Spasokukotskaja et al., 1992).

Following this lead, we sought to identify the source of dCTP utilized for DNA repair. To this end, we have compared the formation of dCTP from Cyd and dCyd and have investigated its utilization in human T lymphoblast CCRF-CEM cells. Our results support the hypothesis that there are two functionally separate dCTP pools in these cells; exogenous [5-^3H]dCyd labels one dCTP pool, and one is preferentially used for DNA repair.


EXPERIMENTAL PROCEDURES

Materials

[5-^3H]dCyd (26 Ci/mmol), [5-^3H]Cyd (26 Ci/mmol), [methyl-^3H]dThd (49 Ci/mmol), and [8-^3H]dATP (21.2 Ci/mmol) were obtained from ICN Biomedicals Inc. (Irvine, CA). Aphidicolin, calf thymus DNA, methanesulfonic acid methyl ester (MMS), (^1)BrdUrd, dCyd, dCTP, and dTTP were obtained from Sigma. DNA polymerase I was a product of Boehringer Mannheim Corp. Oligonucleotide templates for the assay of cellular deoxynucleoside triphosphate levels (Sherman and Fyfe, 1989) were synthesized by Genosys Biotechnologies, Inc. (The Woodlands, TX).

Cell Culture and Synchronization

Human lymphoblast CCRF-CEM cells (Foley et al., 1965) were maintained in RPMI 1640 medium (Life Technologies, Inc.) supplemented with 10% fetal bovine serum (Life Technologies, Inc.). During exponential growth, the population doubling time was 22 h. Cells were routinely checked for contamination with mycoplasma by the American Type Culture Collection and were consistently negative. Cell number and mean volume were determined by a Coulter counter (model ZM, Coulter Electronics, Hialeah, FL). To obtain synchronized cells, a double aphidicolin (2 µM) block method was applied as described by Matherly et al.(1989). The cell cycle phase distribution was determined by flow cytometry after the cells were treated with pepsin and RNase and stained with propidium iodide.

Determination of Specific Activities of [5-^3H]dCTP and [^3H]dTTP Pools

Cells in exponential growth were collected and resuspended in 3 ml of fresh culture medium (0.3-1.0 times 10^7/ml). Dialyzed fetal bovine serum was used for these experiments. Cells were labeled with trace amounts of [5-^3H]dCyd or [^3H]dThd (0.2 µCi/ml) and collected by centrifugation at 1500 revolutions/min for 5 min. Nucleotides were extracted in 0.4 N HClO(4) at 4 °C for 15 min. Following centrifugation the supernatant was neutralized with KOH, KClO(4) was removed by centrifugation, and the resulting supernatant was stored at -20 °C. High pressure liquid chromatographic separation and quantitation of nucleotides was carried out using a Partisil 10-SAX column (250 mm times 4 mm; Whatman Inc., Clifton, NJ) with a gradient generated by NH(4)H(2)PO(4) buffer as described previously (Xu and Plunkett, 1992). The radioactivity associated with [5-^3H]dCTP and [^3H]dTTP was quantitated with a radioactive flow detector (model A250, Packard Instrument Co., Meriden, CT). The amounts of dCTP and dTTP in extracts were quantitated using a DNA polymerase assay with synthetic oligonucleotide template/primers, as described by Sherman and Fyfe(1989); determinations were always made in duplicate or triplicate. The specific activity of each dNTP was expressed as disintegrations/minute/micromole.

Measurement of Incorporation of [5-^3H]dCyd, [^3H]dThd, and [5-^3H]Cyd into DNA

CCRF-CEM cells (3 times 10^6 in 10 ml) in exponential growth phase were incubated with either [^3H]dThd or [5-^3H]dCyd (0.2 µCi/ml) for 60 min. The radioactivity incorporated into HClO(4)-insoluble material was determined using 25-mm glass fiber discs as described previously (Huang et al., 1990). To quantitate DNA synthesis through the de novo pathway, 2 times 10^7 cells in 10 ml were incubated with 0.2 µCi/ml [5-^3H]Cyd for 30 min. Cells were then extracted with 0.4 N HClO(4). The acid-insoluble pellet was collected by centrifugation and washed once with 0.4 N HClO(4). RNA was removed by incubation with 0.5 N NaOH at 37 °C for 2 h in the presence of added calf thymus DNA (0.6 mg/ml). DNA was then precipitated by adding HClO(4) to a final concentration of 0.75 N. The pellet was collected using centrifugation and washed twice with 0.4 N HClO(4). This procedure eliminated greater than 99% of RNA and recovered more than 95% of the DNA in the pellet, as demonstrated by [^3H]Urd and [^3H]dThd labeling in control experiments. Radioactivity was quantitated after the pellet was solubilized with 2 N NaOH at 70 °C for 16 h. Use of [5-^3H]dCyd and [5-^3H]Cyd assured that the dUMP derived from these nucleosides lost its tritium at the thymidylate synthase reaction and did not label dTMP in DNA (Xu and Plunkett, 1993).

Separation of Repaired DNA from Replicated DNA

CCRF-CEM cells were labeled with [5-^3H]dCyd, [^3H]dThd, or [5-^3H]Cyd in the presence of 1 µM BrdUrd to density label DNA, which after alkaline CsCl centrifugation was separated into parental DNA (low density), newly replicated DNA (high density), and repaired DNA (intermediate density) according to Ball and Roberts(1971). DNA was isolated as described previously (Huang and Plunkett, 1992), sheared by eight passes through a 20-gauge needle, which produced DNA fragments with an average length of 15 kilobases. The DNA was banded by alkaline CsCl centrifugation as described by Smith et al.(1981) with minor modifications. Each centrifugation tube contained a total volume of 13 ml. Alkaline CsCl density centrifugation was carried out with a Beckman Ti75 rotor at 37,000 revolutions/min at 25 °C for 90 h. Fractions of 0.2 ml were aspirated from the top of the gradient with a Densiflow IIC apparatus (Buchler Instruments, Inc., Fort Lee, NJ) and diluted with H(2)O for measurements of radioactivity by liquid scintillation counting and of UV absorbance at 260 nm.

Effects of DNA-damaging Agents on [^3H]dThd and [5-^3H]dCyd Incorporation

The differential utilization of [^3H]dThd and [5-^3H]dCyd for DNA semiconservative replication and DNA repair was studied after cells were treated with the DNA-damaging agents MMS, UV, or irradiation. Cells were incubated with the indicated concentrations of MMS for 16 h; UV treatment at 254 nm or -irradiation with a Cs source (at 0 °C) was carried out immediately before [5-^3H]dCyd or [^3H]dThd was added to the cultures. Incorporation of [5-^3H]dCyd and [^3H]dThd into DNA was quantitated after cells were incubated with these nucleosides for 60 min as described above.


RESULTS

Incorporation of [5-^3H]dCyd and [^3H]dThd into DNA in CCRF-CEM Cells

To study functions of salvaged dCyd and dThd as precursors of DNA synthesis, [5-^3H]dCyd and [^3H]dThd were added to exponentially growing CCRF-CEM cells, and the relationship between the specific activities of the corresponding triphosphates and the rates of incorporation into DNA was determined. The size of each pool was undisturbed throughout the experiment; the dCTP pool was 24 ± 4 pmol/10^6 cells, and the dTTP pool was 81 ± 9 pmol/10^6 cells (mean ± S.D.). The specific activity of the [^3H]dTTP pool declined steadily after labeling with [^3H]dThd, whereas the specific activity of the [5-^3H]dCTP pool remained essentially constant for 2.5 h after labeling with [5-^3H]dCyd (Fig. 1A). The initial rate of accumulation of radioactivity in the acid-insoluble material from [^3H]dThd was more rapid than from [5-^3H]dCyd (Fig. 1B). Based on average pool-specific activities of 370 ± 22 dpm/pmol for [5-^3H]dCTP and 257 ± 13 dpm/pmol for [^3H]dTTP 30-50 min after addition of labeled nucleoside, the rate of incorporation was 7.8 and 0.36 pmol/10^6 cells/min for [^3H]dThd and [5-^3H]dCyd, respectively. In fact, the rate obtained from [^3H]dThd may underestimate the true rate of DNA synthesis in these cells, due to the decline in [^3H]dTTP pool specific activity of 8% during this period (Fig. 1A). Our results agree with those in studies of other cell lines, which demonstrated that the rate of dCyd incorporation into DNA is slower than that of dThd, indicating that salvaged dThd, but not dCyd, is readily accessible for use in DNA replication (Plagemann et al., 1978; Karle et al., 1983; Nicander and Reichard, 1983; Taljanidisz et al., 1986; Leeds and Mathews, 1987; Sasvari-Szekely et al., 1989).


Figure 1: Incorporation of [5-^3H]dCyd and [^3H]dThd in exponentially growing CCRF-CEM cells. CCRF-CEM cells were labeled with 0.2 µCi/mL [5-^3H]dCyd (bullet) or [^3H]dThd (circle) to quantitate the rate of incorporation of each into DNA. Aliquots of cells were withdrawn at indicated times and were subjected to HClO(4) extraction. The specific activity of [5-^3H]dCTP and [^3H]dTTP pools of HClO(4)-soluble extracts (A) and the radioactivity incorporated into HClO(4)-insoluble material (B) were determined as described in ``Experimental Procedures.'' The data represent the mean ± S.D. of three determinations.



Effect of Exogenous dCyd on [5-^3H]Cyd Incorporation

To investigate whether exogenous dCyd interfered with the incorporation of [5-^3H]Cyd into DNA through the de novo pathway regulated by ribonucleotide reductase, cells were preincubated with different concentrations of dCyd for 2 h before labeling with [5-^3H]Cyd for 30 min. The cellular dCTP pool was elevated by dCyd in a concentration-dependent manner (not shown), in agreement with our previous studies (Heinemann and Plunkett, 1989; Xu and Plunkett, 1993). This reduced the specific activity of the [^3H]dCTP pool to less than one-third of control cells (Fig. 2). However, addition of dCyd did not affect the incorporation of [5-^3H]Cyd into DNA. These results indicate that the expanded dCTP pool derived from salvaged dCyd did not mix with the dCTP pool used for DNA synthesis, which was derived from the de novo pathway via ribonucleotide reductase.


Figure 2: Effect of exogenous dCyd on [5-^3H]Cyd incorporation into DNA. Exponentially growing CCRF-CEM cells were preincubated with indicated concentrations of dCyd for 2 h before labeling with [5-^3H]Cyd (0.2 µCi/ml) for 30 min. Cells were then subjected to HClO(4) extraction. The specific activity of [5-^3H]dCTP pool (bullet) and the radioactivity incorporated into DNA (circle) were quantitated as described under ``Experimental Procedures.''



Incorporation of [5-^3H]dCyd and [^3H]dThd into Synchronized CCRF-CEM Cells

DNA repair occurs in all phases of the cell cycle (Russev and Boulikas, 1992; Kaufmann and Kaufman, 1993). A repair-specific substrate should demonstrate a different pattern of incorporation into DNA during the cell cycle from that of a precursor that is incorporated predominantly through S phase DNA replication. To test the hypothesis that dCyd may supply a dCTP pool that is specifically utilized for DNA repair, we compared the incorporation into DNA of [5-^3H]dCyd and [^3H]dThd in different phases of cell cycle. A double aphidicolin block method was used to synchronize CCRF-CEM cells in the late G(1) phase (Matherly et al., 1989). As shown in Fig. 3, [5-^3H]dCyd incorporation peaked 2 h after cells were released from the aphidicolin arrest, decreased as cells entered mid-S phase, and reached a second, lower peak between 12 and 18 h, as cells moved on to G(2) and M phases. In contrast, [^3H]dThd incorporation reached its maximum 4-6 h after cells were released from aphidicolin arrest, when most cells of the population were in S phase. These results are consistent with the hypothesis that [5-^3H]dCyd was not used as a substrate of DNA replication. Rather, we suspect that dCyd incorporation measures spontaneous DNA repair (Greer and Kaplan, 1986; Lindahl, 1993; Xiao and Samson, 1993) and possibly repair induced by the synchronizing treatment with aphidicolin.


Figure 3: Incorporation of [5-^3H]dCyd and [^3H]dThd in synchronized CCRF-CEM cells. Cells were synchronized by double aphidicolin block as described under ``Experimental Procdures.'' After cells were released from the second aphidicolin treatment, aliquots of cells were withdrawn at indicated times and were labeled with either [5-^3H]dCyd (bullet) or [^3H]dThd (circle) for 10 min. The radioactivity incorporated into HClO(4)-insoluble material was quantitated by liquid scintilation counting. Cell number was determined separately at the time of assay. The data were expressed as dpm/10^6 cells. Flow cytometry analysis demonstrated that about 70% of the cells entered mid-S phase between 4 and 6 h.



Incorporation of [5-^3H]dCyd, [5-^3H]Cyd, and [^3H]dThd into Replicated and Repaired DNA

To differentiate the functional utilization of exogenous dCyd from that of Cyd and dThd, the incorporation of these nucleosides into DNA was investigated under experimental conditions that would distinguish between replicative and repair synthesis of DNA. Cellular DNA was prelabeled with radioactive nucleosides, followed by density labeling with BrdUrd. After shearing and separation by alkaline density gradient centrifugation, DNA replicated during the BrdUrd incubation would band in alkaline CsCl at a higher density than parental DNA lacking BrdUrd. DNA containing the low level of BrdUrd incorporated during repair patch synthesis would be expected to band at an intermediate density.

As shown in Fig. 4A, when exponentially growing CCRF-CEM cells were labeled with [^3H]dThd for 39 h without BrdUrd and an additional 13 h in the presence of 1 µM BrdUrd, a bimodal distribution of [^3H]dThd-labeled DNA was observed. Radioactivity was associated with a UV-absorbing peak that banded at a high density (fractions 32-45) and was also incorporated into a peak of lesser density (fractions 5-15). These peaks represent newly replicated and unreplicated DNA after BrdUrd addition, respectively. The higher density peak shows less UV absorbance due to the effects of the BrdUrd treatment on DNA replication. When [^3H]dThd labeling and BrdUrd incorporation were carried out simultaneously for half of a cell cycle, only the high density peak was labeled (Fig. 4B). On the other hand, when cells were incubated with [5-^3H]dCyd, radioactivity was incorporated into a peak of intermediate density (Fig. 4C, fractions 16-25) in addition to incorporation into the high density peak. As was the case for dThd, when cells were incubated with [5-^3H]Cyd, only the high density peak was labeled (Fig. 4D), indicating that both [^3H]dThd and [5-^3H]Cyd served mainly as precursors for DNA replication.


Figure 4: Separation of DNA fragments labeled by [5-^3H]dCyd and [^3H]dThd by alkaline CsCl gradient centrifugation. Each experiment was carried out with 2 times 10^7 exponentially growing CCRF-CEM cells suspended in 60 ml of media. A, cells were labeled with 1 µM [^3H]dThd (0.2 µCi/ml) for 39 h. BrdUrd (1 µM) and fresh [^3H]dThd were added for the last 13 h of the incubation. B, cells were incubated with 1 µM [^3H]dThd (0.2 µCi/ml) for 13 h in the presence of 1 µM BrdUrd. C, cells were labeled with tracer amounts of [5-^3H]dCyd (0.2 µCi/ml) for 13 h in the presence of 1 µM BrdUrd. D, cells were labeled with tracer amounts of [5-^3H]Cyd (0.2 µCi/ml) for 13 h in the presence of 1 µM BrdUrd. Isolation of DNA and CsCl centrifugation were performed as described under ``Experimental Procdures.'' Fractions of 0.2 ml were collected and diluted for the measurement of radioactivity (circle) and UV absorption at 260 nm (times).



We hypothesize that the DNA labeled by dCyd that banded at the intermediate density was newly repaired DNA. Again, this is a relatively low level of incorporation and probably represents a background level of repair and possibly that stimulated by the actions of BrdUrd (Ashman et al., 1981; Shewach et al. 1992). Incorporation of dThd or Cyd into this peak was too minor to be detected. The overall incorporation of dCyd was still low compared with either Cyd or dThd in the high density peak. When both the intermediate density peak and the high density peak fractions were combined, the DNA specific activity (dpm/UV absorbance unit) from [^3H]dThd and [5-^3H]Cyd labeling was 23- and 13-fold higher, respectively, than that labeled by [5-^3H]dCyd. These results suggest the existence of functionally compartmentalized dCTP pools for DNA replication and DNA repair in CCRF-CEM cells. The portion of [5-^3H]dCyd that was incorporated into the high density peak may represent the upper limit of mixing of salvage pathway products with the dCTP pool used in replication. It is also possible that some DNA repair was taking place in the newly replicated DNA, probably because of the BrdUrd, which itself evokes DNA repair processes (Hopkins and Goodman, 1980; Shewach et al., 1992).

BrdUrd-enhanced [5-^3H]dCyd Incorporation into DNA

We expected the BrdUrd incorporated into newly replicated DNA in the experiments described above to induce DNA repair activity because BrdUrd is a mutagen. The DNA fragments that contain BrdUrd incorporated during both DNA replication and DNA repair are of high density and cannot be distinguished from DNA replication fragments using the density sedimentation technique. In agreement with this view, we found that incorporation of [5-^3H]dCyd in growing CCRF-CEM cells was indeed enhanced by BrdUrd in a concentration-dependent manner (Fig. 5). To exclude the possibility that the enhanced [5-^3H]dCyd incorporation was due to lowered cellular dCTP levels (Meuth and Green, 1974; Ashman et al., 1981; Shewach et al., 1992), we added dCyd to restore the dCTP pool before [5-^3H]dCyd labeling. This treatment, as shown in Fig. 5, did not change the stimulatory effect of BrdUrd on [5-^3H]dCyd incorporation. This result was consistent with the likelihood that at least part of the [5-^3H]dCyd incorporation into the high density peak (Fig. 4) was due to repair rather than replication.


Figure 5: Effect of BrdUrd on [5-^3H]dCyd incorporation into DNA. CCRF-CEM cells were incubated with BrdUrd at the indicated concentrations for 16 h. Cells were then incubated without (circle) or with (bullet) 1 µM non-radioactive dCyd for additional 2 h before they were labeled with [5-^3H]dCyd for 60 min. The radioactivity incorporated into DNA was quantitated as described under ``Experimental Procedures.''



Differential Actions of DNA-damaging Agents on [^3H]dThd and [5-^3H]dCyd Incorporation into DNA

To further investigate the possibility that dCyd may serve specifically as the precursor of DNA repair, we compared the incorporation of [5-^3H]dCyd and [^3H]dThd into DNA after cells were treated with MMS, UV, or -irradiation. As shown in Table 1, these agents had opposite effects on [5-^3H]dCyd and [^3H]dThd incorporation; they increased the incorporation of [5-^3H]dCyd and decreased the incorporation of [^3H]dThd. The lowered [^3H]dThd incorporation is consistent with the inhibition of semiconservative DNA replication by DNA damaging agents (Swenson and Setlow, 1966; Friedberg, 1985). The good correlation between treatment with DNA damaging agents and enhancement of [5-^3H]dCyd incorporation is consistent with the hypothesis that the dCTP pool labeled by salvaged [5-^3H]dCyd is used for DNA repair but, as demonstrated in earlier experiments, not for DNA replication.



Effect of DNA-damaging Agents on Labeling of Repaired DNA by [5-^3H]dCyd

Density labeling experiments were conducted to distinguish between utilization of salvaged dCyd for either replicative or repair DNA synthesis after DNA damage-inducing treatments. Cells were either incubated with MMS or treated with UV before their DNA was density labeled with BrdUrd during incubation with either [5-^3H]dCyd or [^3H]dThd. Fig. 6shows representative profiles of radioactivity associated with sheared DNA after separation in alkaline CsCl gradients. Labeling of the high density peak (fractions 35 45) by [^3H]dThd was decreased to only 14% of controls in the presence of 0.3 mM MMS (Fig. 6A). In contrast, MMS stimulated [5-^3H]dCyd labeling of the intermediate density peak (fractions 20 33) by 3-fold and the high density peak by 60% (Fig. 6B). Incubations with 0.01-1 mM MMS demonstrated that this effect was concentration dependent up to 0.3 mM MMS; 1 mM MMS appeared to be too toxic, producing visible cell lysis (data not shown). A similar but lesser effect was observed with UV illumination; the 6 J/m^2 dose increased dCyd incorporation in the intermediate density peak by 54%, whereas dThd incorporation was decreased to 87% of controls. The different magnitude of stimulation of dCyd into repaired DNA was due in part to the fact that MMS was present throughout the labeling, whereas UV was used only before [5-^3H]dCyd addition, perhaps giving UV-treated cells sufficient time to complete DNA repair early in the 13-h incubation period.


Figure 6: Effects of MMS and UV on [5-^3H]dCyd and [^3H]dThd incorporation into DNA fragments analyzed by CsCl gradient centrifugation. Exponentially growing CCRF-CEM cells were treated without (circle), or with 6 J/m^2 UV (times), or with 0.3 mM MMS (Delta). Cells were then labeled with [^3H]dThd (A) or with [5-^3H]dCyd (B) in the presence of 1 µM BrdUrd for 13 h as described in Fig. 4. For MMS-treated samples, labeling of [^3H]dThd or [5-^3H]dCyd was carried out in the presence of MMS. Isolation of DNA and CsCl centrifugation were performed as described under ``Experimental Procedures.''




DISCUSSION

The functional compartmentation of dCTP derived from the de novo and salvage pathways has important implications for cellular metabolism. Comparison of the rate of [5-^3H]Cyd incorporation into DNA with that of [5-^3H]dCyd indicates that Cyd is efficiently used for DNA replication in exponentially growing cells, whereas dCyd is a relatively poor precursor for this purpose. On the other hand, it is clear that cells utilize salvaged dCyd for the synthesis of dTTP through the dCMP deaminase pathway (Jackson, 1978; Xu and Plunkett, 1992) and for the synthesis of deoxyliponucleotides (Spyrou and Reichard, 1987, 1989). The origin of deoxynucleotides used for mitochondrial DNA synthesis may be derived from a distinct ribonucleotide reductase associated with this organelle (Young et al., 1994). The present study provides evidence that [5-^3H] dCyd was used selectively as a precursor of DNA repair in exponentially growing CCRF-CEM cells. Induction of DNA repair in growing cells with DNA-damaging agents enhanced the incorporation of [5-^3H]dCyd into DNA, whereas such treatments disrupted DNA replication (Fig. 6). Furthermore, density labeling experiments demonstrated that DNA undergoing repair was specifically labeled with [5-^3H]dCyd ( Fig. 4and Fig. 6).

Our approach involved investigating incorporation of dCyd and of Cyd into the dCMP of DNA and determining whether it occurred through DNA replication or DNA repair. The two pathways were distinguished by differential density labeling; BrdUrd-labeled replicating DNA banded in CsCl gradients at a high density, whereas the density of DNA in which BrdUrd was incorporated during repair was intermediate to replicating DNA and that of parental DNA without BrdUrd (Fig. 4). Greater than 60% of incorporated dCyd was found in the intermediate density peak, and so it appeared to be incorporated through DNA repair. Because some of dCyd identified in the high density peak was likely due to repair induced by BrdUrd, the actual proportion of dCyd used for DNA repair was probably even greater. In contrast, the majority of Cyd was incorporated into the high density peak of replicating DNA. Due to the much greater amount of replicating DNA relative to repaired DNA in these growing cells, estimation of Cyd incorporation into the intermediate density peak was uncertain, but appeared to be less than 5%. In control cells, the rate of [5-^3H]Cyd incorporation (59 pmol/10^6 cells/min, Fig. 2) was significantly greater than the rate of dThd incorporation (7.8 pmol/10^6 cells/min, Fig. 1). Assuming that dThd incorporation represents the true rate of DNA replication (Nicander and Reichard, 1983; Reichard, 1988), the apparent greater rate of [5-^3H]Cyd incorporation may be attributed to functional compartmentation of dCTP. These calculations were based on the average cellular pool-specific activities; because ribonucleotide reductase activity varies with the cell cycle (Eriksson et al., 1984) whereas the activity of dCyd kinase is relatively stable (Liliemark and Plunkett, 1986; Arner et al., 1988), it is possible that the specific activity of [^3H]dCTP depends on cell cycle stage and also on cell type. For example, Chinese hamster ovary cells exhibit a 10-fold difference in [^3H]dCTP-specific activity between G(1) and S phase populations (Leeds and Mathews, 1987); in contrast, the [^3H]dCTP-specific activity in exponentially growing CEM cells was only 60% greater than in S phase cells (Xu and Plunkett, 1993). Additionally, a compartmentalized dCTP pool used for DNA replication could have had a much higher specific activity. If so, the calculated rate of [5-^3H]Cyd incorporation into DNA would be decreased to a value comparable to that of [^3H]dThd incorporation. Although the average cellular [^3H]dCTP-specific activity decreased in cells that were preincubated with dCyd (Fig. 2), the rate of [5-^3H]Cyd incorporation into DNA was apparently unchanged. This suggests that the de novo metabolic route via ribonucleotide reductase is largely restricted from mixing with the dCTP pool generated by the salvage pathway. Furthermore, using synchronized cells, we found that the peak of [5-^3H]dCyd incorporation did not coincide with S phase DNA replication (Fig. 3), another indication that the dCTP pool labeled by [5-^3H]dCyd was excluded from DNA replication.

A compelling body of evidence supports the role of ribonucleotide reductase as the key enzyme in the functional compartmentation of dCTP (Moyer and Henderson, 1985; Nguyen and Sadee, 1986; Spyrou and Reichard, 1989; Mathews and Ji, 1992; Reddy and Fager, 1993). It has been calculated that CDP reduction is the most rapid among the four ribonucleoside diphosphate substrates (Jackson, 1992). Consistent with its central role in supplying dNTPs for DNA replication, the activity of this enzyme is known to be elevated during S phase. Because dCTP pools also increase in S phase cells (Liliemark and Plunkett, 1986; Arner et al., 1988) it is reasonable to conclude that ribonucleotide reductase is capable of producing an excess of dCTP beyond that consumed by DNA replication. In contrast, the activity of deoxycytidine kinase, the rate-limiting step in the salvage pathway, is fairly constant throughout the cell cycle (Liliemark and Plunkett, 1986; Arner et al., 1988). This is consistent with the notion that dCTP generated by the salvage pathway is not specifically required in S phase. Thus it is unlikely that dCTP derived from the salvage pathway would compete effectively with dCTP from the de novo pathway for incorporation into replicating DNA. If ribonucleotide reductase were localized near the DNA replication apparatus or were functionally part of that process, the preferential utilization of dCTP generated by the de novo pathway could be enhanced.

Due to technical limitations, the cellular location of ribonucleotide reductase has remained uncertain. Although some cellular fractionation studies (Leeds, et al., 1985) and investigations using immunocytochemistry (Engstrom and Rozell, 1988) have suggested that ribonucleotide reductase is a cytosolic enzyme, there is evidence that it may be associated with the nuclear membrane (Sikorska et al., 1990). Furthermore, several laboratories have characterized multienzyme complexes that contain ribonucleotide reductase and other enzymes involved in dNTP synthesis and DNA replication (Noguchi et al. 1983; Harvey and Pearson, 1988; Hammond et al., 1989, Reddy and Fager, 1993). A possible resolution to these apparently contradictory findings could be that ribonucleotide reductase exists both in a multiprotein DNA replication complex in the nucleus and as a free enzyme in the cytosol. It is possible that immunocytochemical methodologies may detect the soluble enzyme in the cytosol, but perhaps not a complexed form in the nucleus due to blockage of the epitope by other protein components.

Evidence from recent studies demonstrating that DNA replication forks are arranged in defined spatial patterns within the nucleus provides a structural context for DNA replication in which functionally compartmentalized dNTP pools may be a central component. It is now known that replication forks are tightly clustered in foci within the nucleus (Nakamura, et al., 1986; Mills et al., 1989; Cox and Laskey, 1991; Hozak et al., 1993; Coverley and Laskey, 1994). This arrangement is likely to facilitate rapid consumption of large amounts of dNTPs in each replication focus. It is doubtful that this rate of DNA synthesis can be supported by the relatively low concentration of dNTPs estimated assuming that the dNTPs are uniformly distributed in total cell water.

In contrast to the focal nature of DNA replication, it is likely that DNA repair in response to alkylating agents or radiation is dispersed throughout the genome, possibly with foci at transcriptionally active sites (Jackson et al., 1994). In this situation, DNA synthesis associated with nucleotide excision repair probably utilizes dCTP from a more general pool. We envision that this pool is composed of a mixture of dCTP from the salvage pathway and dCTP which has escaped from its source at ribonucleotide reductase in the replicating foci. In the process of diffusion, its concentration has been diluted relative to that within the replicating focus. When considering the utilization of exogenous dCyd, we propose that the salvage pathway contributes to this generalized pool of dCTP, which our experiments have demonstrated comprises the bulk of the nucleotide used for DNA repair, but makes only a small contribution to replicative DNA synthesis.

The relatively minor utilization of exogenous dCyd in replicating DNA should not be viewed as contradictory to the fact that a number of dCyd analogs, such as arabinosylcytosine, 2`,2`-difluoro-2`-deoxycytidine, 5-aza-2`-deoxycytidine, and 2`,3`-dideoxycytidine require the same salvage pathway for activation prior to incorporation into DNA (Major et al., 1981; Momparler, 1985; Huang et al., 1991, Starnes and Cheng, 1987). The extent to which these analogs are incorporated as a result of repair synthesis relative to DNA replication is unknown, although it has been assumed that the latter pathway is utilized in the absence of DNA damaging stimuli. Although the rate of exogenous dCyd incorporation into replicating DNA (0.36 pmol/10^6 cells/min) is about 5% of that of dThd incorporation (Fig. 1), the rate of 2`,2`-difluoro-2`-deoxycytidine incorporation was even less than that of dCyd (Huang et al., 1991). On the other hand, in comparison with dCyd, there are factors that favor the incorporation of analogs via the replication pathway. For example, cytidine nucleotide analogs accumulate to greater cellular concentrations than does dCTP (Plunkett and Gandhi, 1993), and the presence of the nucleotides of these drugs in the cell is prolonged because they are less likely to be eliminated by dCMP deaminase (Momparler, 1985; Heinemann et al., 1992). Thus, the incorporation of cytidine analogs may be taken as an indication of the upper limit of the mixing that occurs between the general dCTP pool generated by the salvage pathway with the dCTP pool produced by the de novo pathway for DNA replication.

In summary, it appears most of the dCTP generated by ribonucleotide reductase goes to a high throughput, low volume pool; some of this dCTP appears to become available to a more general pool in the nucleus. The salvage pathway also contributes dCTP and cytidine nucleotide analogs to this general pool. One of the functions of this pool in CCRF-CEM cells is to supply dCTP for repair of DNA. It is possible that this functional compartmentation could be used to advantage in chemotherapy by combining cytidine nucleotide analogs with agents and modalities that evoke a DNA repair response in tumor cells. Recent reports suggest the utility of this approach (Gregoire et al., 1994; Shewach et al., 1994).


FOOTNOTES

*
This work was supported in part by Grant DHP-1 from the American Cancer Society and Grant CA28596 from the National Cancer Institute, Department of Health and Human Services. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Clinical Investigation, Box 52, The University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030. Tel: 713-792-3335; Fax: 713-794-4316.

(^1)
The abbreviations used are: MMS, methanesulfonic acid methyl ester; dpm, disintegration/minute.


ACKNOWLEDGEMENTS

We gratefully acknowledge the expert editorial assistance of Walter Pagel in the preparation of this manuscript.


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