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Carcinogenesis, Vol. 22, No. 1, 179-186, January 2001
© 2001 Oxford University Press

CANCER BIOLOGY

Excessive base excision repair of 5-hydroxymethyluracil from DNA induces apoptosis in Chinese hamster V79 cells containing mutant p53

Li-Jun Mi, Wenren Chaung, Robert Horowitz1, George W. Teebor and Robert J. Boorstein

2Department of Pathology, New York University School of Medicine, Sackler Institute of Graduate Biomedical Sciences and The Rita and Stanley Kaplan Cancer Center, New York, NY 10016 and
1 Department of Oncology, Montefiore Hospital, Albert Einstein School of Medicine, Bronx, NY 10467, USA


   Abstract
Top
 Abstract
Introduction
 Materials and methods
 Results
 Discussion
 References
 
We have demonstrated previously that the toxicity of 5-hydroxymethyl-2'-deoxyuridine (hmdUrd) to Chinese hamster fibroblasts (V79 cells) results from enzymatic removal of large numbers of hydroxymethyluracil residues from the DNA backbone [Boorstein,R. et al. (1992) Mol. Cell. Biol., 12, 5536–5540]. Here we report that a significant portion of the hmdUrd-induced cell death that is dependent on DNA base excision repair in V79 cells is apoptosis. Incubation of V79 cells with pharmacologically relevant concentrations of hmdUrd resulted in the characteristic changes of apoptosis as measured by gel electrophoresis, flow cytometry and phase contrast microscopy. However, hmdUrd did not induce apoptosis in V79mut1 cells, which are deficient in DNA base excision repair of 5-hydroxymethyluracil (hmUra). Apoptosis was not prevented by addition of 3-aminobenzamide, which inhibits synthesis of poly(ADP–ribose) from NAD, indicating that apoptosis was not the direct consequence of NAD depletion. Pulsed field gel electrophoresis indicated that hmdUrd treatment resulted in high molecular weight (2.2–4.5 Mb) DNA double-strand breaks prior to formation of internucleosomal ladders in V79 cells. Simultaneous measurement of DNA strand breaks with bromodeoxyuridine/terminal deoxynucleotidyl transferase–fluorescein isothiocyanate labeling and of cell cycle distribution indicated that cells with DNA strand breaks accumulated in late S/G2 and that hmdUrd-treated cells underwent apotosis after arrest in late S/G2 phase. Our results indicate that excessive DNA base excision repair results in the generation of high molecular weight DNA double-strand breaks and eventually leads to apoptosis in V79 cells. Thus, delayed apoptosis following DNA damage can be a consequence of excessive DNA repair activity. Immunochemical analysis showed that both V79 and V79mut1 cells contained mutant p53, indicating that apoptosis induced by DNA base excision repair can be independent of p53.

Abbreviations: AP site, apyrimidinic site; BrdUrd, bromodeoxyuridine; FITC, fluorescein isothiocyanate; hmdUrd, 5-hydroxymethyl-2'-deoxyuridine; hmUDG, hydroxymethyluracil–DNA glycosylase; hmUra, 5-hydroxymethyluracil; PARP, poly(ADP–ribose) polymerase; PS, phosphotidylserine; TdT, terminal deoxynucleotidyl transferase.


   Introduction
Top
 Abstract
 Introduction
Materials and methods
 Results
 Discussion
 References
 
We have previously demonstrated that the thymidine analog 5-hydroxymethyl-2'-deoxyuridine (hmdUrd) is toxic to mammalian cells following incorporation into cellular DNA. However, it is only toxic to cells that contain the repair enzyme hydroxymethyluracil–DNA glycosylase (hmUDG), demonstrating that the toxicity of hmdUrd to these cells is a consequence of an intact DNA base excision repair system (1). hmUDG initiates the base excision repair process by excising 5-hydroxymethyluracil (hmUra) from DNA, generating an apyrimidinic (AP) site (2). Such AP sites are acted on by an AP endonuclease (3,4) and then a 2'-deoxyribophosphoesterase (5), resulting in transient single strand breaks in DNA (reviewed in ref. 6). The gap can then be filled by DNA polymerase and the nick is finally resealed by DNA ligase.

We now ask whether the hmdUrd-induced cell death seen in repair-competent mammalian cells is best classified as apoptosis. Apoptosis, or programmed cell death, can result from exposure of cells to a wide variety of DNA-damaging agents including ionizing radiation, ultraviolet radiation, oxidizing agents and drugs used in cancer chemotherapy (reviewed in refs 7 and 8). Apoptosis is usually characterized by nuclear fragmentation, by the generation of high molecular weight DNA fragments and by internucleosomal DNA fragmentation (`ladder' formation), production of apoptotic bodies (sub G0/G1 population in flow cytometer) (9,10), along with characteristic alterations in cytoplasmic morphology. Normal p53, the product of a tumor suppressor gene, is required for or associated with most apoptosis induced by DNA damage (11–13).

While many previous studies have documented that DNA-damaging agents can produce apoptosis, interpretation of the mechanisms by which damage results in apoptosis has been complicated by the fact that most such agents produce heterogeneous damage to DNA while interfering with other cellular functions. Furthermore, since DNA damage can be repaired, it is difficult to ascribe apoptotic consequences to unrepaired DNA damage in DNA, or to alterations in DNA following initial modification by DNA repair enzymes. In our system, cells are incubated in the presence of hmdUrd, resulting in the introduction of large quantities of a single type of lesion into cellular DNA. Our isolation of an hmUDG-deficient cell line makes it possible to study the consequences of DNA damage in the presence or absence of the initiation of DNA repair. Therefore, we can ask whether the base excision repair process itself results in apoptosis, how the apoptosis is mediated, which phase of the cell cycle the cells have gone through before apoptosis and whether such apoptosis is dependent on the presence of the wild-type p53 gene.


   Materials and methods
Top
 Abstract
 Introduction
 Materials and methods
Results
 Discussion
 References
 
Cells
Chinese hamster lung fibroblasts (V79 cells) containing hmUDG and V79mut1 cells were maintained as previously described (1). hmUDG-deficient and hmdUrd-resistant V79mut1 cells were derived from V79 cells (1) and have the same phenotype as the parental cells upon exposure to DNA-damaging agents (14). p53–/– human promyelocytic HL-60 cells (15) were grown in RPMI 1640 medium containing 10% heated fetal bovine serum (FBS) and 1% penicillin and streptomycin. Mouse embryo fibroblasts known to have wild-type p53 (16) were generously provided by Dr Robert Carroll (NYUMC). Primary Chinese hamster embryo fibroblasts were generously provided by John Lehman (Albany Medical College).

DNA gel electrophoresis
Internucleosomal fragmentation (ladder formation) was analyzed as described by Wyllie (17) with minor modification. Cells were trypsinized, washed with phosphate-buffered saline (PBS) and lysed with neutral lysis buffer (18). Genomic DNA was extracted by the phenol–chloroform method and analyzed by gel electrophoresis at 46 V for 2 h on a 1.0% agarose gel containing 0.4 µg/ml ethidium bromide. Internucleosomal fragmentation was determined by inspection under UV light.

DNA staining and flow cytometry
Apoptotic cells (defined as those with less than the G0/G1 DNA content) were measured according to the protocol of Nicoletti et al. (9). Briefly, cells were trypsinized and then fixed with 70% ethanol at 4°C overnight. Cells were centrifuged at 400 x g for 10 min in a GKPR centrifuge (Beckman) and the cell pellets were then resuspended in 1% FBS/PBS. Cells were treated with RNase and stained with 50 µg/ml propidium iodide (Sigma). Samples were then analyzed on a FACScan flow cytometer (Becton Dickinson, San Jose, CA) with a LYSYS II program.

Pulsed field gel electrophoresis
High molecular weight DNA double-strand breaks were analyzed by pulsed field gel electrophoresis as described (19). Cells were labeled with [3H]thymidine for 24 h; 3H was then removed and 4 µM hmdUrd was added. At the end of the indicated treatment, cells were collected and suspended in 0.6% agarose (pulsed field gel electrophoresis grade; Sigma), in a solution consisting of ~15 µl cell pellet, 45 µl L buffer (50 mM EDTA, 50 mM Tris, 20 mM NaCl) and 60 µl 2x TAE, and placed into plug molds (Bio-Rad). After being cooled, samples were digested in digestion buffer containing 0.5 mg proteinase K per sample and washed with TE buffer (20 mM Tris, 50 mM EDTA, pH 8.0). Samples were then analyzed by pulsed field gel electrophoresis for 3 days in 0.6% chromosomal-grade agarose (Bio-Rad) running at 2 V/cm with a 30 min switch interval for 72 h at 14°C in 1x TAE. Each lane of the gel was sliced at 0.5 cm intervals and [3H]thymidine content was determined using a scintillation counter. Cells were irradiated with 0.5–5.0 Gy of {gamma}-irradiation (137Cs) as a positive control for formation of double-strand breaks.

Analysis of DNA strand breaks as a function of cell cycle stage by flow cytometry
To identify the cell cycle stage where DNA strand breaks accumulated and where apoptotic bodies were formed, cells exposed to hmdUrd were collected and labeled with bromodeoxyuridine (BrdUrd; Sigma) by terminal deoxynucleotidyl transferase–fluorescein isothiocyanate (TdT–FITC) labeling (TdT kit; Beohringer-Mannheim, Indianapolis, IN; FITC-conjugated anti-BrdUrd was from Becton Dickinson, San Jose, CA) and stained with propidium iodide as described elsewhere (20–23) with minor modifications. Briefly, cells were exposed to 4 µM hmdUrd for 0 or 24 h, and then harvested immediately or 24 h after the termination of the 24 h exposure. Cells were then fixed in 70% ethanol and washed with PBS and 0.4–0.6 x 106 cells were then suspended in a total volume of 50 µl containing 0.25 nmol of BrdUrd, TdT reaction buffer (TdT kit), 25 µM cobalt chloride and 12.5 units of TdT. The reaction mixture was maintained at 37°C for 1 h. After centrifugation, the pellet was washed with PBS and resuspended in TBF staining buffer (0.5% Tween20 and 0.5% FBS in PBS) and incubated with 0.625 µg of FITC-conjugated anti-BrdUrd antibody at 37°C for 0.5 h. Samples were washed with staining buffer and suspended in 500 µl of PBS, treated with RNase and stained with 2.5 µg propidium iodide. Cells exposed to camptothecin, which is known to generate DNA strand breaks) were collected and incubated in the absence or presence of TdT and used as negative and positive controls, respectively. Samples were analyzed by a FASCan flow cytometer. The fluorescent signals from FITC and propidium iodide were quantified using LYSYS II software.

Detection of membrane phosphotidylserine with FITC–annexin V by flow cytometry
The externalization of phosphotidylserine (PS), another reliable early apoptosis marker, was detected in V79 cells using an ApAlert annexin V kit (ClonTech). Briefly, V79 cells were treated with 0 or 4 µM hmdUrd for 24 h and were then further incubated for 0, 8 or 24 h in fresh medium. Cells were then collected and suspended in binding buffer (provided in the ClonTech kit) and incubated with FITC–annexin V (ClonTech) for 10 min. Samples were immediately analyzed by FACscan flow cytometry (Becton-Dickinson).

Immunocytochemical analysis of p53 protein
The presence or absence of functional p53 in V79 and V79mut1 cells was analyzed by immunocytochemistry (24) using the monoclonal antibody p53 DO-1 (Santa Cruz Biotechnology), which recognizes both wild-type and mutant p53 protein, and with p53 pAb240 (Oncogene Science), which recognizes only mutant p53 protein. Cells were collected and cytocentrifuged on to silane-coated slides. The slides were then air dried, fixed in 10% neutral-buffered formalin and washed with PBS. One hundred microliters of a 1:80 diluted primary antibody was applied to the slides and incubated at room temperature for 1 h. Slides were washed with PBS three times; biotinylated horse anti-mouse IgG (1:200) was added to the slides which were then incubated at room temperature for 30 min. Elite ABC (Vector Laboratories, Burlingame, CA) (1:100) was added to the slide and incubated at room temperature for another 30 min. DAB staining was performed after washing with PBS. Hemotoxylin was used as a counterstain. The slides were then examined by conventional microscopy.


   Results
Top
 Abstract
 Introduction
 Materials and methods
 Results
Discussion
 References
 
Induction of apoptosis by hmdUrd in V79 cells is delayed and dose dependent
We have shown previously that removal of large amounts of the pyrimidine base hmUra from DNA via DNA base excision repair was lethal to V79 cells (1). To determine whether this cell death was apoptosis, V79 cells were exposed to hmdUrd for 24 h and cellular DNA was analyzed by gel electrophoresis to observe the characteristic DNA internucleosomal fragmentation (`ladder' formation). As shown in Figure 1Go, cells analyzed immediately after the initial 24 h exposure to hmdUrd showed no clear formation of ladders. However, when cells were analyzed after 24 h of additional incubation following removal of hmdUrd from the growth medium, ladder formation was readily observed (Figure 1Go, lanes 5 and 7). The delayed apoptosis suggests that hmdUrd-induced apoptosis did not appear to be a direct toxic effect. Rather, it is supportive of the hypothesis that hmdUrd-induced apoptosis is associated with cellular repair activity.



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  		Fig. 1. DNA gel electrophoresis of hmdUrd-treated V79 cells. V79 cells were exposed to 0 (lane 1), 1 µM (lanes 2 and 3), 4 µM (lanes 4 and 5) or 10 µM hmdUrd (lanes 6 and 7) for 24 h, the hmdUrd was removed and cells were analyzed immediately (lanes 2, 4 and 6) or incubated for an additional 24 h (lanes 3, 5 and 7). Apoptosis (ladder formation) can be readily seen with the highest doses of hmdUrd 24 h after its removal from the medium (lanes 5 and 7). As a positive control, HL-60 cells were exposed to 10 µM camptothecin (lane 8). {lambda}/HindIII and {phi}X174/HaeII molecular weight markers are displayed in the rightmost lane.

 
The apoptosis observed was also dose dependent. With 10 µM hmdUrd treatment, marked fragmentation was clearly visible (Figure 1Go, lane 7) and with 4 µM hmdUrd, a faint ladder was seen (Figure 1Go, lane 5), whereas with 1 µM hmdUrd, no ladders could be observed. These data are consistent with our previous results that removal of large amounts of hmUra from DNA via hmUDG was toxic to V79 cells and suggest that the excessive DNA base excision repair of hmUra induces apoptosis in V79 cells.

Induction of apoptosis by hmdUrd in V79 cells but not in V79mut1 cells
To confirm that hmdUrd induces apoptosis as a consequence of DNA repair, both repair-competent V79 cells and hmUDG-deficient V79mut1 cells were exposed to hmdUrd and examined by phase contrast microscopy. With 24 h exposure to hmdUrd, followed by additional incubation in the absence of hmdUrd for an additional 24 h, characteristic cytoplasmic blebbing and apoptotic bodies were clearly seen in V79 cells (Figure 2AGo, panel b). No such changes were seen in V79mut1 cells exposed to hmdUrd (Figure 2AGo, panel d). These observations further support the hypothesis that hmdUrd-induced apoptosis is related to intact DNA base excision repair.





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  		Fig. 2. (A) Morphology of V79 and V79mut1 cells treated with hmdUrd. V79 cells (a, b) or V79mut1 cells (c, d) were exposed to 0 (a, c) or 10 µM hmdUrd (b, d) for 24 h, the hmdUrd was removed and the cells were incubated for an additional 24 h. Apoptosis was seen only in V79 cells. (Phase contrast microscopic magnification is 20x). (B) Flow cytometric analysis of hmdUrd-induced apoptosis in V79 and V79mut1 cells. V79 cells (left) and V79mut1 cells (right) were treated with 0 (top row) or 4 µM hmdUrd for 24 h (middle and bottom rows). Cells were analyzed immediately after the removal of hmdUrd (middle) or incubated for an additional 24 h (bottom row). Cells were analyzed by FASCan flow cytometry using LYSYS II software. Histograms of DNA content are displayed on a log scale. The apoptotic cells are shown with sub-G1 DNA content. (C) Flow cytometric analysis of camptothecin-induced apoptosis in V79 and V79mut1 cells. V79 cells (left) and V79mut1 cells (right) were treated with 0 (top) or 20 µM camptothecin (bottom) for 24 h. Cells were analyzed as described in Materials and methods. Apoptotic cells are indicated with arrows.

 
The association of hmdUrd-induced apoptosis with the repair activity of hmUDG was further analyzed quantitatively in both V79 cells and V79mut1 cells by measuring the fraction of cells containing less than the G0/G1 DNA content as measured by flow cytometry (9). As shown in Figure 2BGo, apoptotic cells were readily found in the repair-competent V79 cells (left) but not in the repair-deficient V79mut1 cells (right), providing strong evidence that hmdUrd-induced apoptosis is associated with repair.

To prove that V79mut1 cells are not deficient in apoptotic machinery, they were exposed to the topoisomerase I inhibitor, camptothecin, a known inducer of apoptosis (reviewed in ref. 25). Both V79 cells and V79mut1 cells showed the same pattern and similar extent of apoptosis when stained with propidium iodide and analyzed by flow cytometry (Figure 2CGo).

The quantitative analysis in V79 cells during the different time intervals shown in Figure 2BGo is consistent with the gel electrophoresis results showing <10% apoptotic cells at the end of 24 h exposure to 4 µM hmdUrd, but <=25% apoptotic cells 24 h after termination of the 24 h treatment. In summary, the finding that hmdUrd induces apoptosis in repair-competent cells, but not in repair-deficient cells demonstrates that one of the consequences of excessive DNA base excision repair can be apoptosis.

Double-strand break formation after exposure to hmdUrd
Next we explored possible mechanistic links between DNA base excision repair and the hmdUrd-induced apoptosis in V79 cells. While DNA base excision repair is known to produce transient single-strand DNA breaks, we previously hypothesized that double-strand breaks might be generated during extensive repair activity (26). To test this hypothesis, we measured the formation of double-strand breaks by pulsed field gel electrophoresis, a sensitive way of detecting high molecular weight double-strand DNA breaks. hmdUrd induced a high level of cleavage of DNA to high molecular weight (<=4.6 Mb) fragments by 24 h after 4 µM hmdUrd exposure in V79 cells (Figure 3Go). These high molecular weight fragments have a similar molecular weight to the >1 Mb fragments demonstrated in thymocytes after dexamethasone exposure (27). The percentage of DNA breaks was equivalent to or higher than that seen in cells irradiatiated with 2.5 Gy of {gamma}-irradiation (data not shown). There was no detectable double-strand breakage in cells treated with hmdUrd for only 8h, while increased breakage was seen 12 h after a 24 h hmdUrd treatment. These results confirm that the double-strand breaks result from the repair of hmdUrd. It remains to be clarified whether the double-strand breaks that are generated in the repair competent cells are (i) direct signals for apoptosis, (ii) an early consequence of apoptosis or (iii) activators of other pathways in the apoptosis machinery.



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  		Fig. 3. Detection of DNA double-strand breaks after hmdUrd exposure in V79 cells by pulsed gel electrophoresis. V79 cells were prelabeled with [3H]thymidine for 24 h. They were then grown in the absence or presence of 4 µM hmdUrd for 12 or 24 h. Cell were then harvested immediately or were washed and incubated for an additional 24 h before harvesting; they were then analyzed by pulsed field gel electrophoresis as described in Materials and methods. Molecular weight standards of 5.7, 4.6, 3.5 and 2.2 Mb migrated 1, 2, 3.5 and 5.5 cm, respectively.

 
Preferential labeling with BrdU/TdT–FITC of cells arrested in G2 by hmdUrd exposure
We have shown here and elsewhere that exposure of V79 cells to pharmacologically relevant concentrations of hmdUrd induces cell cycle arrest at late S/G2 phase (Figure 2BGo and ref. 26). To understand the relationship between generation of DNA strand breaks and induction of apoptosis in terms of the cell cycle, V79 cells were exposed to hmdUrd for different lengths of time and analyzed using BrdUrd/TdT–FITC labeling combined with propidium iodide staining. As seen in Figure 4Go, at 0 h, <5% of cells were labeled with BrdUrd/TdT–FITC and these cells showed a similar cell cycle distribution to unlabeled cells. At 24 h (Figure 4bGo), as previously reported (26), the overall population of cells was clustered at late S/G2, but these cells preferentially become labeled with BrdUrd/TdT–FITC. G1 cells showed essentially no strand breaks as judged by this assay. Of the cells collected 24 h after the end of the 24 h hmdUrd treatment, >60% were labeled with BrdUrd/TdT–FITC; these cells were also mainly accumulated at late S. Alternatively, these cells could have been arrested in G2, and a reduction in DNA content made them appear to be in S phase. A population with less than the G0/G1 DNA content, representing apoptotic bodies, was also present (indicated by arrow Fig. 4CGo), which was consistent with Figure 2BGo. These cells also were positive for BrdUrd/TdT–FITC.



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  		Fig. 4. Simultaneous measurement of DNA strand breaks with BrdUrd/TdT–FITC and of cell cycle compartment by flow cytometry. V79 cells (a–c) and V79mut1 cells (d and e) were exposed to 4 µM hmdUrd for 0 (a, d) or 24 h (b, e). Cells were harvested immediately after the treatment (a, b, d and e) or after another 24 h of incubation in the absence of hmdUrd after the end of the 24 h treatment with hmdUrd (c). Cell samples were labeled with BrdUrd/TdT–FITC followed by propidium iodide staining and analyzed as described in Materials and methods. Left-hand panels show dot plots of cells, with the y axis showing the labeling with BrdUrd/TdT–FITC (R2 region), and the x axis showing the DNA content (propidium iodide staining). The percentage of labeled cells is indicated. The right panels are histograms showing the cell cycle distribution pattern of individual regions (R2 on the upper and R1 on the lower). The amount of cells is plotted on the y axis and the DNA content on the x axis. G1, S and G2 phase are indicated in the 0 h treatment (a, d). An arrow indicates the apoptotic bodies (c). The figure shows representative data from one of two experiments.

 
In contrast, when V79mut1 cells were treated with hmdUrd for 24 h (Figure 4eGo), no increase in strand breakage was seen, indicating that DNA strand breakage after hmdUrd exposure is entirely dependent on hmUDG activity. Our results suggest a mechanism in which excessive removal of hmUra produces many strand breaks in DNA, thus inducing late S/G2 arrest. Apoptosis appears to proceed directly from S/G2 or G2/M phase rather than from G1/S as for other DNA damage-induced apoptosis (reviewed in ref. 1).

hmdUrd-induced apoptosis in V79 cells causes membrane externalization of phosphotidylserine
Since the processes of apoptosis DNA base excision both generate 3'-OH ends in DNA, it is hard to differentiate the two cell populations using the TdT assay alone. Although co-staining with propidium iodide staining would be helpful in this matter, ideal evidence of apoptosis in this system would be the production of a signal that is independent of DNA strand breaks. One such signal is membrane externalization of PS (28–30). Figure 5Go shows that there was four times more externalized PS in hmdUrd-treated cells than in untreated cells. In addition, PS seemed to be a marker that can be detected earlier (8 h after the termination of 24 h hmdUrd treatment) than internucleosomal DNA fragmentation. This result further confirms that excessive hmUDG repair activity induces apoptosis characterized both by the DNA fragmentation and by cell membrane changes.



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  		Fig. 5. HmdUrd-induced apoptosis in V79 cells causes membrane externalization of PS. V79 cells were exposed to 0 (top panel) or 10 µM hmdUrd for 24 h and then were grown for another 8 h in the absense of hmdUrd. Cells were collected and stained with FITC–annexinV as described in Materials and methods. Samples were then analyzed by FACscan flow cytometry. The percentage of positively stained cells (apoptotic cells) is indicated in the R2 region of the left panel presented as a dot scatter, and in the right panel presented as a histogram. The figure shows representative data from one of three experiments.

 
Effects of 3-aminobenzamide on apoptosis induced by DNA base excision repair
Recent studies indicate not only that the enzyme poly(ADP–ribose) polymerase (PARP) is activated after damage and repair (26,31,32) but also that cleavage of this enzyme is integral to apoptosis (33,34). The reduction in cellular NAD+ concentration as a consequence of poly(ADP–ribose) synthesis has also been proposed to play a role in apoptosis (35). To study a potential role for this enzyme in apoptosis, cells were exposed to hmdUrd followed by the PARP inhibitor, 3-aminobenzamide. Non-toxic concentrations of 3-aminobenzamide cause marked toxicity to hmdUrd-treated cells, which die predominantly in G2. 3-Aminobenzamide also prevents the reduction in cellular NAD+ after incorporation and repair of hmdUrd by blocking poly(ADP–ribose) synthesis (26,31). Addition of 3-aminobenzamide did not prevent cells from entering apoptosis after exposure to hmdUrd (Figure 6Go).



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  		Fig. 6. Effect of 3-aminobenzamide, a PARP inhibitor, on hmdUrd-induced apoptosis. Cells were exposed to 4 µM hmdUrd for 24 h and, after removal of hmdUrd, incubated in the presence or absence of 4 µM 3-aminobenzamide. The population with sub-G0/G1 DNA content was quantified by flow cytometry. Data represent the average of four experiments.

 
Detection of p53 in V79 and V79mut1 cells
To begin to analyze the molecular basis of hmdUrd-induced apoptosis, it was important to determine if the p53 gene product is involved in this process. Are the effects of the repair of hmdUrd that result in apoptosis mediated by p53? p53 expression increases in response to other types of DNA damage (36,37), and p53 is involved in binding to and reannealing strand breaks (38) and, integrally, in apoptosis (11). We have previously shown that the p53 gene of V79 cells contains point mutations that affect the ability to induce mdm2 (39). In the current study, we examined whether further alterations in p53 might explain the lack of apoptosis in V79mut1 cells. Therefore, we evaluated the status of p53 in V79 and V79mut1 cells by immunocytochemistry. Both cell lines stained strongly with the anti-p53 monoclonal antibody PAb-240, which recognizes only mutant p53 (Figure 7Go). PAb-240 did not stain HL-60 cells (which are known to lack p53) or mouse embryo fibroblasts (which have only wild-type p53). Staining with antibody p53 DO-1 was also strongly positive. Although p53 DO-1 recognizes both wild-type and mutant p53, strong staining is consistent with the p53 protein being mutated in both lines, because wild-type p53 has a very short half-life. When the p53 cDNA for the V79mut1 cell line was sequenced, it showed mutations at the same sequence as in V79 cells (39), excluding the possibility that the p53 gene had undergone mutation during the establishment of the V79mut1 cells from V79 cells. The differences in hmdUrd-induced apoptosis between V79 and V79mut1 cells are therefore not the consequence of differences in p53. Our immunocytochemical analysis of p53 protein in the V79 cell line is also consistent with the findings of another recent study (40). Taken together, our results indicate that the V79 and V79mut1 cell lines both lack normal p53, so we can conclude that the lack of excision repair-mediated apoptosis in V79mut1 cells was not a consequence of an alteration in p53.



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  		Fig. 7. Detection of p53 protein in V79 cells by immunohistochemistry. Immunohistochemical staining with p53 antibody DO-1 (a and b) and Pab-240 (c–f) as primary antibody on slides of V79 cells (a, c), V79mut1 cells (b, d), mouse embryo fibroblasts (e) and human promyelocytic HL-60 cells (f). DAB was used as an indicator system and shows brown staining in positively stained cells. Hemotoxylin was used as a counterstain (60x).

 
As an additional control for the p53 analysis, primary Chinese hamster embryo cells were also analyzed by immunocytochemistry. These cells did not stain with PAb-240 but showed positive nuclear staining with DO-1 (39; J.Lehman, unpublished results). When these cells were exposed to hmdUrd, they also underwent apoptosis as measured by morphology and flow cytometry, but to a lesser extent than V79 cells (data not shown). The qualitative differences in apoptosis between the primary Chinese hamster embryo cells are best explained by the fact that the primary cells have less hmdUrd incorporation and greater cell cycle transit time than V79 cells. These results demonstrate that hmdUrd-induced apoptosis as a consequence of DNA base excision repair can occur in cells with normal p53 as well.


   Discussion
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
References
 
In previous studies, we have demonstrated that when V79 cells are exposed to hmdUrd for 24 h, the hmdUrd is incorporated into cellular DNA. When the hmdUrd is removed from the growth medium, the cells continue to remove the remaining hmUra from the cellular DNA, thereby remaining sensitive to the inhibitor of poly(ADP–ribosylation), 3-aminobenzamide (26,34). The dose-dependent apoptosis described here supports our earlier hypothesis that the excessive base excision repair of hmUra is lethal to cells and is consistent with other recent studies on imbalanced DNA repair (41). The time dependence found here indicates that hmdUrd-induced apoptosis is a slow, repair-dependent phenomenon and differs from the apoptosis that occurs after exposure to other DNA-damaging agents, such as UV radiation, free radicals and oxidants, which induce apoptosis very soon after DNA damage (reviewed in ref. 8).

Our results show that cells that incorporate large amounts of hmdUrd during S phase then arrest in late S/G2, allowing repair. Cells that are able to complete repair can subsequently divide, but the S/G2 cells unable to complete repair accumulate double-strand breaks and proceed directly to apoptosis (pre-G1 cells). The mechanism of double-strand break formation is uncertain. The single-strand breaks resulting from repair of hmUra may be on opposite strands from those generated by repair of endogenous damage such as spontaneous deamination, depurination or methylation. Thus the closely juxtaposed single-strand breaks generated by hmUDG repair activity might be converted to double-strand breaks (42). Second, if the repair-generated single-strand breaks are not resealed in time, other enzymes, particularly nucleases, or possibly a trypsin-activated endo-exonuclease (43), might recognize the break and cleave the opposite strand. Other more general mechanisms of double-strand break formation might be occurring. Regardless of the mechanism, we see here a process dependent on base excision repair which proceeds from late S/G2, not G1.

Activated PARP is known to bind to the ends of broken DNA strands and to poly(ADP–ribosylate) certain nuclear proteins including itself and topoisomerase I (44). We have previously demonstrated that 3-aminobenzamide does not block initiation of DNA base excision repair, but does block repair-dependent NAD depletion, which results from synthesis of poly(ADP–ribose) (31). Our current results with inhibition of PARP thus indicate that repair-induced apoptosis can occur under conditions in which poly(ADP–ribose) synthesis is inhibited and in which cellular NAD+ is not being depleted. Thus our data would indicate that, in this system, apoptosis is not the direct consequence of NAD+ depletion. Whether the slight enhancement of hmdUrd induced apoptosis by 3-aminobenzamide results from increased production of strand breaks (45) remains to be further studied.

Our present results thus demonstrate that repair of non-lethal base modifications by an intact DNA base excision repair system can produce apoptosis. This process involves the generation of DNA strand breaks to produce high molecular weight DNA fragments and late S/G2 arrest.

Although p53 is known to play a role in cellular responses to DNA damage, the DNA base excision repair of hmUra, the resultant alterations in cell cycle progression to G2 arrest, the generation of high molecular weight DNA double-strand breaks and, ultimately, apoptosis are independent of completely normal p53 function in this system. These results are consistent with recent findings that p53 is not required for the delayed G2 arrest following ionizing radiation and for radiation-induced apoptosis. Colon cell lines lacking p53 fail to arrest in G1 in response to {gamma}-irradiation, but undergo apoptosis with substantially more slowly than p53 wild-type cells or tissues (46). Furthermore, HL-60 cells, which lack p53, undergo radiation-induced apoptosis following progression to a G2 checkpoint. Maximal apoptosis was seen 72 h after radiation treatment (47). Our results raise the possibility that the delayed p53-independent apoptosis following treatment with ionizing radiation may result from the gradual repair of base damage produced by ionizing radiation, in contrast to the immediate p53-dependent apoptotic response to the damage produced directly by the radiation.


   Notes
 
2To whom correspondence should be addressed Email: robert.boorstein{at}med.nyu.edu


   Acknowledgments
 
We would like to thank Dr John Hirst for flow cytometric analysis, Dr Herman Yee for immunohistochemical staining and Dr Ross Basch for helpful discussions. This work was supported by an American Cancer Society Junior Faculty Award, by NIH grants CA-51060 and AM-07421, by the Rita and Stanley Kaplan Cancer Center, the Irma T.Hirschl Trust and a gift from Charles and Helen Lazars.


   References
Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received November 19, 1999; revised August 22, 2000; accepted August 30, 2000.


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S. Ulbert, M. Cross, R. J. Boorstein, G. W. Teebor, and P. Borst
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