Human Molecular Genetics, 2002, Vol. 11, No. 18 2189-2200
© 2002 Oxford University Press
Defects in homologous recombination repair in mismatch-repair-deficient tumour cell lines
1Institute for Cancer Studies, University of Sheffield School of Medicine, Beech Hill Road, Sheffield S10 2RX, UK, 2Department of Oncological Sciences and 3Department of Biochemistry, University of Utah School of Medicine, Salt Lake City, UT 84112-5330, USA
Received May 20, 2002; Accepted June 28, 2002
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Loss of mismatch repair (MMR) leads to a complex mutator phenotype that appears to drive the development of a subset of colon cancers. Here we show that MMR-deficient tumour cell lines are highly sensitive to the toxic effects of thymidine relative to MMR-proficient lines. This sensitivity was not a direct consequence of MMR deficiency or alterations of DNA precursor metabolism. Instead, MMR-defective tumour cell lines are also defective in homologous recombination repair (HRR) induced by DNA double-strand breaks. Furthermore, a frameshift mutation of the human RAD51 paralog XRCC2 found in the MMR-deficient uterine tumour cell line SKUT-1 can confer thymidine sensitivity when introduced into a MMR-proficient line. Like other cells with defective XRCC2, SKUT-1 is sensitive to mitomycin C, and MMR-proficient cells expressing the mutant XRCC2 allele become more sensitive to this agent. These data suggest that the thymidine sensitivity of MMR-deficient tumour cell lines may be a consequence of defects in the HRR pathway. The increased thymidine sensitivity and the loss of an important pathway for the repair of DNA double-strand breaks create new opportunities for therapies directed specifically against this subset of tumours.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Hereditary non-polyposis colon cancer (HNPCC) is an autosomal dominant inherited disorder characterized by an early onset of cancer of the colon and less frequently cancer of the stomach, small intestine, endometrium, ovary or upper urinary tract (1). HNPCC is primarily the result of mutations of two mismatch-repair (MMR) genes: hMSH2 and hMLH1 (2). The proteins encoded by MMR genes are essential for the correction of DNA replication errors (3–5), and recognize DNA modifications induced by some types of damaging agents (6,7). Inactivation of MMR results in an increase in mutation rate at microsatellite and coding sequences (8–11) and resistance to the cytotoxic effects of some types of DNA-damaging agents (6,12–14). In addition MMR appears to be necessary for induction of the G2 cell cycle checkpoint and apoptosis in response to DNA-alkylating agents or the purine analogue 6-thioguanine (15–17). MMR deficiency occurs as an early step in the development of tumours in HNPCC (9) and appears to drive the accumulation of mutations in genes required for the control of cellular growth [e.g. the TGFßRII receptor (18)] and apoptosis [e.g. BAX (19)].
MMR appears to interact with other DNA-repair pathways (5), although the importance of these interactions in the tumour-suppressive effects of MMR is unclear. MMR is known to have an antirecombinogenic role in bacteria and yeast, since this repair pathway corrects mismatches arising during strand exchange as well as replication (20). MMR proteins in Saccharomyces cerevisiae appear to interact with recombination proteins to regulate heteroduplex length (21) and to suppress homeologous recombination (22). Recent evidence suggests that MMR plays similar roles in mouse embryonic stem cells, since loss of MMR results in longer gene conversion tracts around double-strand breaks (DSBs) (23) and increases the frequency of recombination between diverged sequences (23,24). In addition, it has been shown that the S. cerevisiae MSH2/6 complex binds the Holliday junctions formed as intermediates in homologous recombination repair (HRR) and can facilitate their cleavage in in vitro assays (25).
Both MMR and HRR are affected by imbalances of the precursors of DNA synthesis, the deoxyribonucleoside triphosphates (dNTPs) (26,27). Imbalances in dCTP and dTTP pools, induced in cultured cells by adding thymidine to the medium, slow the DNA replication fork (28) and increase the rate of mutation caused by misinsertion of the dNTP in excess (26). dCTP and dTTP pool imbalances alter the sensitivity to DNA-alkylating agents (29), and depletion of the dTTP pool induces chromosome breakage that provides substrates for HRR (30). Cells have mechanisms that protect against the genetic effects of pool imbalances. Ribonucleoside triphosphate (rNTP) and dNTP pool imbalances trigger G1-and/or S-phase checkpoints (31,32) and apoptosis (33). Here we show that MMR-deficient tumour cell lines as a group are more sensitive to the toxic effects of thymidine. Our data also suggest that this hypersensitivity is the result of defects in the HRR pathway in MMR-deficient tumour cell lines.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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MMR-deficient tumour cell lines are hypersensitive to thymidine
To investigate potential synergistic effects between dNTP pool imbalances and MMR, we first determined the sensitivity of the repair-deficient tumour cell lines to the toxic effects of thymidine relative to MMR-proficient tumour cell lines (see Table 1 for a list of cell lines). Dose–response studies revealed that the MMR-deficient tumour cell lines were 2.9–15-fold more sensitive to the toxic effects of thymidine (Fig. 1). Cell lines showing hypersensitivity originated from a variety of tumours (colon, endometrial and ovarian). They had varying p53 status, although all were deficient in MMR (Table 1).
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Thymidine sensitivity is not a direct consequence of MMR deficiency
We next determined whether correction of the repair deficiency made cells more resistant to the cytotoxic effects of thymidine. HCT116 +3 is a strain of the hMLH1-deficient colon cancer cell line HCT116 in which the MMR deficiency (and the mutator phenotype) was corrected by the introduction of human chromosome 3, which contains the wild-type repair gene (34). The response of HCT116 +3 was compared with that of a derivative (called M2) that specifically lost the complementing hMLH1 gene but retained most of chromosome 3. M2 is MMR-deficient and regains the mutator phenotype characteristic of MMR-deficient cells. M2 was used in this experiment because the introduction of other genes on chromosome 3 in HCT116 +3 could have affected its response. Furthermore, HCT116 +3 is a clonal derivative of HCT116, and, considering the genetic instability of this line, a clonal derivative may have properties different from the uncloned population. However HCT116+3 and M2 responded similarly to thymidine (Fig. 2). In addition, a strain of the hMSH6-deficient line of DLD-1 made MMR-proficient by the transfer of human chromosome 2 (35) remained as sensitive to thymidine as the parental DLD-1 (data not presented).
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dNTP pool levels in MMR-proficient and -deficient tumour cell lines are not altered
Since the differences in thymidine sensitivity were not directly attributable to MMR status, we measured dNTP levels in the tumour cell lines to determine whether the sensitivity was the result of imbalances of intracellular dNTP pools. Excess thymidine inhibits cell growth by depleting cells of dCTP through the allosteric regulation of ribonucleotide reductase (28). Regulatory mutations of enzymes involved in dNTP synthesis can confer resistance to thymidine, and such mutations typically lead to alterations in the balance of the dCTP pools (26). Measurements of dNTP pools in tumour cell lines revealed some variations of dNTP content (Table 2). Given the diversity of tumour cell lines assayed, this was not surprising. However, there were no consistent differences in the level or balance of the dNTPs in MMR-proficient or -deficient cell lines that could account for the altered thymidine sensitivity. For example, the repair-proficient colorectal carcinoma cell line SW480 had the lowest level of dCTP despite being relatively resistant to the cytotoxic effects of thymidine. Furthermore, dNTP pools in both repair-proficient (SW480) and -deficient (HCT116) cell lines responded similarly to treatment with thymidine (Table 2). dTTP levels increased to virtually the same value in both cell lines after a 24-hour treatment with 0.5 or 1.5 mM thymidine, and dCTP levels declined. Thus differences in dNTP pools do not appear to account for the altered sensitivity to thymidine.
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MMR-deficient cells arrest in S phase at a lower level of thymidine
Since the depletion of dCTP by thymidine treatment slows DNA replication fork progression, thymidine-treated cells accumulate in S phase of the cell cycle. To determine whether the hypersensitive HCT116 cells arrested in S phase following treatment with low levels of thymidine, we measured the cell cycle distribution in HCT116 and MMR-proficient SW480 cells treated with equimolar and equitoxic doses of thymidine for 24 hours. HCT116 cells accumulated in S phase after treatment with 0.25 mM thymidine (Fig. 3). In contrast, this concentration of thymidine had little or no effect on SW480 cells. If SW480 cells were treated with an equitoxic dose of thymidine (1.5 mM for SW480), the fraction of cells in S phase increased to a level similar to that found in HCT116 cells treated with 0.25 mM. HCT116 cells treated with 1.5 mM thymidine did not arrest in mid-S phase, although there was a pronounced broadening of the G1 peak at the G1–S border. This indicated that many of the cells arrested at the G1–S border or very early in S phase. The reduced level of G2 cells could be explained by the cycling of the G2 cells initially present through to G1 and the paucity of cells entering G2 because of the stringent block of DNA synthesis. Again a higher concentration of thymidine (5 mM) was necessary to obtain the same response in SW480. Thus both MMR-proficient and -deficient lines arrest in S phase after treatment with cytotoxic levels of thymidine. However, a 3–6-fold lower level of thymidine was sufficient to produce arrest in HCT116.
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MMR-deficient tumour cell lines are defective in homology-based recombination induced by DNA DSBs
Given our recent finding that a hamster cell line deficient in HRR is sensitive to thymidine (36), we determined whether defects in HRR could underlie the thymidine sensitivity found in MMR-deficient tumour cell lines. To test this possibility, we used the assay developed by Jasin and co-workers (37), which measures the homology-based recombination of two defective G418 genes following the introduction of a DNA DSB by a transiently expressed endonuclease, I-SceI (Fig. 4A). The recombination reporter SCneo was electroporated into the MMR-proficient tumour cell lines SW480 and SW620 and the MMR-defective lines DLD-1, HEC1A and SKUT-1 (Fig. 4B). Following transfection of the I-SceI expression vector pCMV3nls–I-SceI, strains of SW480 and SW620 containing intact single copies of SCneo produced G418+ colonies at a frequency of 0.8–3.5x10-3 (Fig. 4C). The frequencies in SW480 and SW620 cells that were not transfected with the I-SceI expression vector were 370–1000-fold lower. These frequencies are very similar to those reported by Jasin and co-workers (37) for hamster V79 cells. In contrast, the frequencies of G418+ colonies were not increased in SCneo-containing strains of DLD-1, HEC1A or SKUT-1 following transfection of the I-SceI vector (Fig. 4C). As a result, the frequencies in the MMR-deficient strains transfected with the I-SceI vector were 200–15 000-fold lower than those in the corresponding SW480 or SW620 cells.
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We next considered the possibility that the difference in recombination frequency could be the result of an inability of the MMR-deficient cell lines to take up the plasmid expressing the I-SceI endonuclease. To test this, we measured the frequency of cells taking up a construct expressing the green fluorescent protein (GFP) from the same cytomegalovirus promoter in the expression vector pcDNA3. However, there was no significant difference in the frequency of GFP+ cells when this construct was transfected into MMR-proficient or -deficient cell lines (data not presented). In addition, we tested the possibility that the low level of recombinants in the MMR-deficient cell lines was the result of an induction of cell death following expression of the I-SceI endonuclease specifically in the MMR-deficient cells. To resolve this question, the GFP and I-SceI expression constructs were cotransfected into MMR proficient and deficient cells. The fraction of GFP+ cells undergoing apoptosis was then determined 24 hours later by Hoechst staining, which detects condensed apoptotic nucleii. This experiment did not reveal any difference in the fraction of apoptotic cells in the two cell types. Thus the depressed frequency of G418+ cells in the MMR-deficient cell lines DLD-1, HEC1A and SKUT-1 appears to be a result of a deficiency in homology-based recombination.
The MMR-deficient tumour cell line SKUT-1 has a mutation of the Rad51 paralog XRCC2
To determine whether the deficiency in the formation of G418+ cells in the MMR-deficient tumour cell lines was the result of mutations of genes known to be required for HRR, we amplified cDNA sequences encoding the human RAD51 paralogues XRCC2 and XRCC3. We initially chose these genes because cells deficient in them have been shown to be defective in the homology based recombination assay used above (37,38). Sequence analysis of cDNAs amplified from the cell lines revealed a frameshift in XRCC2 in one of the MMR-deficient tumour cell lines, SKUT-1 (Fig. 5A). This mutation was a single nucleotide deletion in a run of eight thymine residues at nucleotides 342–350 of the XRCC2 coding sequence. The mutation appeared to be heterozygous, since both the wild-type and -1 frameshifted mutant sequences can be seen in the XRCC2 sequencing image for SKUT-1. To verify the presence of the mutant allele, we subcloned the amplified fragment covering this region into pcDNA3.1. Sequence analysis revealed a roughly equal proportion of plasmids containing the wild-type XRCC2 fragment and those carrying the -1 frameshift. In addition, we considered the possibility that the SKUT-1 cell population was heterozygous in that it contained cells carrying either the mutant or wild-type allele. To test this possibility, DNAs purified from single-cell clones of SKUT-1 were used to amplify and sequence the XRCC2 fragment containing the frameshift. Of six single-cell clones tested in this way, all were heterozygous for the mutation. This frameshift results in the loss of the wild-type amino acid sequence after amino acid 116 (of the 280 encoded by the gene) and produces a stop codon 48 nucleotides downstream from the frameshift (Fig. 5B). No mutations were found in XRCC3 in any of the cell lines.
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To determine whether this frameshift mutation affected the level of the XRCC2 protein in SKUT-1, extracts from MMR-proficient and -deficient cell lines were analysed by western blotting. These blots revealed that the level of XRCC2 was indeed depressed in SKUT-1, but not in any other cell lines (Fig. 5C).
MMR-proficient cells expressing the mutant XRCC2 allele become sensitive to thymidine
Given that the mutation in SKUT-1 was heterozygous, we next determined whether it was responsible for the thymidine sensitivity found in this line. The mutant allele was cloned into the expression vector pcDNA3.1 and transfected into the MMR-proficient tumour cell line SW480. Expression of the mutant construct was confirmed by detection of the frameshift mutation in XRCC2 cDNAs amplified from the transfected SW480 clones (Fig. 6A). Those expressing the mutant allele were then assayed for sensitivity to thymidine. As shown in Figure 6B, SW480 clones expressing the mutant allele became more sensitive to thymidine. The degree of sensitivity in the transfectants is not as great as that found in SKUT-1; however, the level of expression of the mutant allele appears to be lower in the transfectants.
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Cells expressing the mutant XRCC2 allele are mitomycin C-sensitive
A distinctive property of cells defective in XRCC2 is their sensitivity to mitomycin C (39–41). To determine whether SKUT-1 or the transfectants of SW480 containing the mutant allele were also sensitive to mitomycin C, we measured the viability of these cells in the presence of varying concentrations of this agent. SKUT-1 was sensitive to mitomycin C relative to the MMR-proficient line SW480 (Fig. 7). Similarly, transfectants expressing the mutant XRCC2 allele showed increased sensitivity to this agent.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Cells that develop MMR deficiency acquire a mutator phenotype that appears to drive genetic changes required for tumorigenesis. Here we show that MMR-deficient tumour cell lines become hypersensitive to the cytotoxic effects of thymidine. Our data further indicate that this thymidine sensitivity is the result of defects in the HRR pathway. Thymidine sensitivity is not a direct result of loss of MMR. Previous work with MMR-deficient mouse embryonic stem cells demonstrates that the knockout of MSH2 does not result in HRR deficiency (23). Thus the thymidine sensitivity and the loss of HRR appear to be the result of events downstream from the loss of MMR. In support of this hypothesis, we found a frameshift mutation in the HRR gene XRCC2 typical of mutations found in MMR-deficient tumour cells.
Since MMR-deficient tumour cells are known to acquire a wide range of mutations, other mechanisms that could lead to thymidine sensitivity were investigated. MMR deficiency itself is not directly responsible for this effect. Two different MMR-deficient tumour cell lines in which the repair defect was corrected by chromosome transfer remained as sensitive to thymidine as the uncorrected parental cells. Alterations in dNTP pools also do not account for the thymidine sensitivity. Earlier work has shown that mutations affecting the allosteric regulation of ribonucleotide reductase or CTP synthetase lead to thymidine resistance through an elevation of the dCTP pool (26). Other regulatory alterations of these enzymes (leading to increased dTTP, for example) could conceivably induce sensitivity. However, there were no consistent alterations of dNTP content in repair-proficient and -deficient tumour cell lines, and pools responded similarly to exogenous thymidine. Despite these observations, thymidine induces S-phase arrest at a 3–6-fold lower concentration in MMR-deficient cells relative to proficient ones, indicating that the repair-deficient cells display an acute sensitivity to changes in dNTP balance.
Our recent observation that an HRR-deficient hamster cell line lacking functional XRCC3 is thymidine-sensitive (36) provided a strong rationale for measuring the integrity of this repair pathway in MMR-deficient tumour cell lines. Further support for this hypothesis came from the increased sensitivity of these tumour cell lines to the DSB-inducing agents camptothecin and etoposide (42). Consistent with these reports, the MMR-defective tumour cell lines tested here were defective in the production of G418+ colonies by homology-based recombination following induction of a site-specific DSB.
In addition, we found that the MMR-deficient tumour cell line SKUT-1 acquired a frameshift mutation in XRCC2. The mutation identified in SKUT-1 was heterozygous. However transfection of the mutant allele into a MMR-proficient tumour cell line conferred thymidine and mitomycin C sensitivity, indicating that this allele functions in a dominant-negative manner. Transfectants expressing the mutant allele were not as sensitive to these agents as SKUT-1. This may be due to a lower level of expression of the mutant allele relative to the wild-type genes. Alternatively, there may be other alterations in SKUT-1 cells that further contribute to the sensitivity. XRCC2 is a member of the recA/RAD51 family of HRR proteins (39–41) and is required for the homology-based recombination repair of DNA DSBs (37). The truncated protein resulting from the frameshift retains the Walker Box A that interacts with the phosphates of ATP, but it loses the Walker Box B (Fig. 5B). It is not clear how the truncation would affect interactions with other human Rad51 paralogs (43,44), and this is the subject of further investigation. The XRCC2 mutation was only found in one of the MMR-deficient cells analysed here. However, it is interesting to note that frameshift mutations in MRE11 have been found in a high proportion of MSI+ tumours and in one of the thymidine-sensitive MMR deficient cell lines (HCT116) used in this work (45). MRE11 has been implicated in sensing of DNA damage, and plays a role in checkpoint responses as well as both non-homologous end joining and HRR at DSBs (46,47). We are currently testing the effect of this mutant allele on thymidine sensitivity and HRR to determine the role that it plays in this phenotype.
There is ample evidence from studies with bacteria and yeast to suggest that loss of HRR could account for the increased sensitivity to thymidine. In these organisms, HRR is required to reactivate stalled replication forks (48,49). Since thymidine slows replication forks in mammalian cells by limiting dCTP supply, our findings are consistent with the idea that HRR is necessary to resolve aberrant replication structures that may be induced by thymidine-induced dNTP pool imbalances in human cells. In the absence of HRR, such structures may trigger S-phase arrest and cell death.
An intriguing issue raised by these findings is the potential effect of the HRR deficiency on the development of this subset of tumours. As discussed above, the HRR dysfunction does not appear to be a direct result of MMR deficiency. Frameshifts in mononucleotide runs of coding sequences are a signature of mutations resulting from loss of MMR (10,18,19). Thus the frameshift found in XRCC2 provides evidence supporting our argument that loss of HRR is a downstream event, following loss of MMR, not unlike the mutations in the TGFßRII or BAX genes (18,19). It is possible that the disregulation of recombination resulting from loss of MMR may pose a growth disadvantage for cells. Long gene conversion tracts or unregulated homeologous exchanges may retard S-phase progression or trigger checkpoints. Therefore loss of HRR may provide a selective growth advantage through the suppression of such genetic exchanges. Alternatively, given the role of HRR in maintaining the integrity of DNA replication forks, it is possible that loss of HRR may compromise S-phase checkpoints in the tumour cells. Consistent with this possibility, Takeda and co-workers (50) have presented evidence that supports the hypothesis that ATM and HRR are on the same damage response pathway. The importance of the loss of this pathway in the development of tumours has been well documented (51).
The hypersensitivity of MMR-deficient tumour cells to thymidine strikingly contrasts the resistance of these lines to DNA-alkylating agents, 6-thioguanine and cisplatin (6,12–14). The MMR-mediated ‘correction’ of lesions induced by these agents appears to be lethal, since deficient cells are strongly resistant to their cytotoxic effects. This resistance to important classes of chemotherapeutic agents has created a major difficulty in devising therapies directed against tumours showing microsatellite instability (52). The increased sensitivity of MMR-deficient cell lines to thymidine and the loss of HRR in such cells raise the possibility that agents inducing dNTP pool imbalances may improve the effectiveness of treatment of this distinctive subset of tumours.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Cell lines and culture
MMR-proficient and -deficient tumour cell lines used in these experiments are presented in Table 1. Most were obtained from American Type Culture Collection (Manassas, VA). The immortalized human diploid fibroblast cell line LM217E was from John Murname (University of California at San Francisco). The +3 and M2 derivatives of HCT116 (34) were provided by Dr C. Richard Boland (University of California at San Diego). +3 and M2 were grown in alpha minimum essential medium with 10% fetal bovine serum (Gibco BRL, Rockville, MD), and the remaining cell lines were maintained in Dulbecco's modified eagle's medium with 10% FBS. The MMR defects present in the MMR-defective cell lines used in these experiments were confirmed by western blotting (data not presented).
Cytotoxicity assays
Cytotoxic response to thymidine and mitomycin C was measured in medium supplemented with dialysed serum (to remove exogenous sources of deoxynucleosides). 100–500 cells were plated in each well of six-well dishes and treated with varying concentrations of thymidine in triplicate (Sigma). Cells were allowed to grow for 6–15 days before staining with 0.4% methylene blue/50% methanol (Fisher Scientific). Colonies of >50 cells were scored. The surviving fraction was determined by dividing the average number of colonies for each treatment by the average number of colonies in the control wells.
Cell cycle analysis
Approximately 1x106 cells were plated into appropriate medium with 10% dialysed serum 48 h prior to treatment with 0.25–5 mM thymidine. At appropriate time intervals, 1.5x106 cells were isolated, washed with PBS, fixed with methanol (Fisher), and stored at -20°C. The samples were resuspended in 0.01% propidium iodide (ICN Biomedicals, Inc., Lisle, IL) and 2 mg/ml RNase A (Calbiochem, San Diego, CA) and incubated at room temperature for 30 min in the dark. DNA content was determined by fluorescence-activated cell sorting using a FACScan machine with Cell Quest software (Becton Dickinson). The data were analysed with the Modfit program (Verity House Software, Topsham, MA).
dNTP measurements
Exponentially growing cells maintained in dialyzed serum were washed with cold TBS (Tris-buffered saline) and then incubated in cold 60% methanol/1% toluene. The methanol/toluene solution was dried, resuspended in 5% TCA and neutralized with 1.5 volumes of Freon–amine (0.5 M Freon and tri-n-octylamine, Sigma).
Contaminating rNTPs were removed by passing cell extracts over a boronate column (Affigel 601, Bio-Rad Laboratories, Hercules, CA) and by treatment with 0.1 M sodium periodate. The pH of the samples were then adjusted to the HPLC mobile phase ( pH 3.7) with phosphoric acid.
dNTP concentrations in the extracts were measured by anion-exchange chromatography using a Beckman HPLC (high-performance liquid chromatography) system. The dNTPs were fractionated using a Partisil 10 SAX column (10 µmx25 cm, Whatman, Clifton, NJ) with a monobasic ammonium phosphate gradient consisting of 0.075 M ammonium phosphate pH 3.7 (buffer A) and 1.0 M ammonium phosphate pH 3.7 (buffer B) at a flow rate of 1.5 ml/min. Individual dNTPs were identified by comparison with retention times of standards (Boehringer Manheim, Indianapolis, IN) that were monitored at 254 and 260 nm. Peak areas were measured using Beckman Gold software, and concentrations were determined by comparison with a calibration curve for each dNTP.
Recombination assay
The SCneo recombination substrate and the pCMV3nls–I-SceI expression vector were kind gifts from Dr Maria Jasin, Memorial Sloan Kettering Cancer Center. To obtain strains of the MMR-proficient and -deficient lines carrying single copies of the SCneo recombination reporter, 1.5x107 cells of each cell line were electroporated at 0.4 kV/50 µF with the uncut SCneo substrate and plated in a non-selective medium. Hygromycin (final concentration 0.1 mM) was added to the medium after 48 h, and hygR colonies were isolated and expanded from each cell line. To identify strains containing SCneo, Southern blotting was performed on genomic DNA (10 µg) isolated from each clone. The 1.1 kb XhoI–HindIII fragment radio labelled with [
-32P]dCTP was used as probe, and detection was carried out by autoradiography. Isolates carrying SCneo (SW480 clones SN3, 4 and 5, SW620 clone SN2, DLD-1 clones SN2 and SN6, Hec-1A clones SN2 and 3, and the SKUT-1 clone SN3) were transfected with the pCMV3nls–I-SceI expression vector (10 ng) using Lipofectamine (5 h treatment, with a final concentration 10 µg/ml; Gibco BRL). Recombination frequencies were determined by selection in media containing 1 mg/ml G418 (at a density of 13–25 cells/mm2). In addition, two dishes were plated with 500 cells each to measure cloning efficiency. Cells were allowed to grow for 10–14 days before staining with 0.4% methylene blue/50% methanol. Colonies of >50 cells were subsequently scored. All experiments were repeated independently three to six times.
cDNA sequencing
mRNA was extracted from each individual cell line using the RNeasy protect mini kit (Qiagen). The coding regions of the XRCC2 and XRCC3 genes were amplified using the Reverse iTONE-STEP RT–PCR Kit (Abgene) according to the manufacturer's protocol. The primer pair XRCC2f (5'-AGTTG GTGAA TGGCG TTGGT-3') and XRCC2r (5'-CGTAG TACCC TGCAA AAGAC-3') was used at 0.4 µM to amplify XRCC2 cDNA. The XRCC3 cDNA was amplified in two fragments. Primers for fragment 1 were XRCC31f (5'-TGAAT TGAAG GCGAG TGCCT-3') and XRCC31r (5'-AGCAG TACGG GGACC TTCTT-3'), and those for fragment 2 were XRCC32f (5'-CCAGG AGAGC TGCTT CAGAA-3') and XRCC32r (5'-GCAGA CGCGT TTTAA AGGCC-3'). The RT–PCR cycle was 47°C for 30 min; 94°C for 2 min; (94°C for 40 s; 52–56°C for 45 s; 72°C for 1 min)40; 72°C for 10 min. cDNA products were purified using the QIAquick PCR Purification Kit (Qiagen) and sequenced using IRD-labelled primers with the SequiTerm EXCEL II kit (Epicentre Technologies) according to the manufacturer's protocol. Samples were run on PAGE and analysed using the LONG READIR 4200 DNA sequencer (LI-COR).
Plasmid construction and transfection
The XRCC2 cDNA from SKUT-1 was cloned into the pcDNA3.1 Topo expression vector (Invitrogen) according to the manufacturer's protocol. The inserted XRCC2 cDNA was sequenced and a vector carrying the mutated allele of XRCC2 was enriched. The pcDNAXRCC2m was transfected into the SW480 cell line using Lipofectamine (Gibco–BRL) according to the manufacturer's protocol. Three colonies were isolated and expanded in G418-selective medium (1 mg/ml). To verify expression of the mutated allele, mRNA was extracted from each clone using the RNeasy Protect Mini Kit (Qiagen), and the XRCC2 cDNA was sequenced as described above.
Western blot analysis
Cells were trypsinized and counted before lysis in RIPA buffer in the presence of the proteinase inhibitor PMSF (100 µg/ml). Extracts were prepared from 5x106–1x107 cells. 100 µg/µl of each protein sample was separated on 8–16% Tris–glycine precast gels (BioWhittaker Molecular Applications) and transferred to nitrocellulose membrane (Immun-Blot PVDF, BioRad) using a transblot semidry transfer unit (BioRad). This membrane was blocked in 5% milk for 1 h, and the XRCC2 protein was detected using a goat polyclonal antibody (Santa Cruz Biotechnologies) at a 1 : 200 dilution in 3% milk overnight. Anti-goat peroxidase conjugates (Santa Cruz Biotechnologies) were used as a secondary antibody at a dilution of 1 : 50 000. Immunoreactive protein was visualized using ECL reagents (Amersham Pharmacia) following the manufacturer's instructions.
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ACKNOWLEDGEMENTS |
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We are particularly grateful to Herbert Ley and Michael Mathews for their help with the HPLC analysis. This work was supported by a programme grant from Yorkshire Cancer Research to M.M. and by Grant CA62244 from the US National Cancer Institute to M.M. and B.D.P.
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FOOTNOTES |
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* To whom correspondence should be addressed. Tel: +44 1142713288; Fax: +44 1142713515; Email: m.meuth{at}sheffield.ac.uk
The author wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors.
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