Intrinsic and acquired resistance to cisplatin remains a primary hurdle to treatment of high-grade serous ovarian cancer (HGSOC). Cisplatin selectively kills tumor cells by inducing DNA crosslinks that block replicative DNA polymerases. Single-stranded DNA (ssDNA) generated at resulting stalled replication forks (RF) is bound and protected by heterotrimeric replication protein A (RPA), which then serves as a platform for recruitment and activation of replication stress response factors. Cells deficient in this response are characterized by extensive ssDNA formation and excessive RPA recruitment that exhausts the available pool of RPA, which (i) inhibits RPA-dependent processes such as nucleotide excision repair (NER) and (ii) causes catastrophic failure of blocked RF. Here, we investigated the influence of RPA availability on chemosensitivity using a panel of human HGSOC cell lines. Our data revealed a striking correlation among these cell lines between cisplatin sensitivity and the inability to efficiently repair DNA via NER, specifically during S phase. Such defects in NER were attributable to RPA exhaustion arising from aberrant activation of DNA replication origins during replication stress. Reduced RPA availability promoted Mre11-dependent degradation of nascent DNA at stalled RF in cell lines exhibiting elevated sensitivity to cisplatin. Strikingly, defective S-phase NER, RF instability, and cisplatin sensitivity could all be rescued by ectopic overexpression of RPA. Taken together, our findings indicate that RPA exhaustion represents a major determinant of cisplatin sensitivity in HGSOC cell lines.

Significance: The influence of replication protein A exhaustion on cisplatin sensitivity harbors important implications toward improving therapy of various cancers that initially respond to platinum-based agents but later relapse due to intrinsic or acquired drug resistance. Cancer Res; 78(19); 5561–73. ©2018 AACR.

High-grade serous ovarian cancer (HGSOC) is the most lethal gynecologic malignancy, with a 5-year survival rate of approximately 50% (1). Frontline treatment consists of debulking surgery followed by combination chemotherapy with a platinum-based drug such as cisplatin (CDDP). This regimen yields a high initial response rate, often resulting in clinical remission. However, most patients eventually relapse, at which point their cancer manifests strong resistance to CDDP (2). The therapeutic efficacy of CDDP is attributable to the capacity of its platinum atom to form covalent bonds involving the N7 position of purines in DNA generating 1,2- and 1,3-intrastrand crosslinks (∼98%) in addition to a much lower yield (∼2%) of interstrand crosslinks (3). These adducts strongly block the progression of DNA polymerases, inducing a state of “replication stress” that selectively promotes the elimination of rapidly proliferating cancer cells (4).

The mechanism of clinical resistance to CDDP is complex and remains incompletely understood. The cytotoxicity of CDDP can be reduced by processes that counter its ability to react with DNA, including increased efflux/reduced uptake and inactivation by sulfur-containing molecules (5). In addition, CDDP-induced intrastrand crosslinks are removed by nucleotide excision repair (NER; ref. 6), and cells deficient in this pathway are sensitive to the drug (7, 8). Indeed, based on various lines of evidence, increased NER capacity has been proposed as an important mechanism of CDDP resistance in ovarian cancer (5).

In addition to mechanisms outlined above that remove DNA lesions or inhibit their formation, modulation of the cellular response to replication stress, i.e., pathways that contribute to the stabilization/resolution of stalled replication forks (RF), is emerging as a key determinant of CDDP resistance in cancer (9). For example, germline mutations in BRCA1 or BRCA2 greatly increase susceptibility to ovarian and breast tumors (10). Although BRCA1/2 were initially characterized for their key roles in DNA double-strand break (DSB) repair by homologous recombination (HR), recent data reveal that these proteins also stabilize stalled RF by inhibiting Mre11-dependent nucleolytic degradation of nascent DNA (11, 12). Moreover, sensitivity to replication-blocking drugs such as CDDP was recently shown to correlate strongly with the inability of BRCA1/2-defective cells to protect nascent DNA from nuclease activity (12).

Following exposure to replication stress–inducing drugs, uncoupling of DNA polymerases from the MCM helicase complex at blocked RF generates regions of single-stranded DNA (ssDNA), which are rapidly coated by heterotrimeric replication protein A (RPA; ref. 13). RPA-ssDNA acts as a recruitment/activation platform for DNA damage response (DDR) factors that mitigate replication stress, notably the apical kinase ataxia telangiectasia and Rad3-related (ATR; ref. 14). Activated ATR phosphorylates multiple substrates, which cooperate to (i) prevent further origin activation, (ii) stabilize stalled RF, and (iii) block cell-cycle progression. Abrogation of ATR activity during replication stress leads to unrestricted firing of DNA replication origins, excessive accumulation of ssDNA, and greatly increased levels of chromatin-bound RPA (15). This eventually results in global exhaustion of cellular RPA and extensive DSB formation at persistently stalled RF, a process recently termed “replication catastrophe” (16). However, the precise mechanisms leading to RF instability in response to RPA exhaustion, and their potential influence on cancer chemoresistance, remain incompletely characterized.

Under conditions of limited RPA availability, important RPA-dependent processes aside from DNA replication can be compromised, which might contribute to increased RF stalling/failure. Indeed, we and others have shown, in various mutant yeast strains and mammalian cell lines exhibiting replication stress response defects, that sequestration of RPA at stalled RF exhausts the cellular pools of this complex, thus interfering with its ability to perform essential functions in NER during S phase (17, 18). Moreover, we previously demonstrated that a majority of human melanoma cell lines exhibit striking defects in NER exclusively during S, although the molecular basis was not thoroughly investigated (19). Here, we probed the impact of RPA exhaustion on DNA repair and RF stability, thereby elucidating the basis of chemoresistance, in ovarian cancer cells.

Cell culture

Ovarian cancer cell lines were established from patient tumors (TOV) or ascites (OV) as described (20–23), and cultured in OSE medium (Wisent) supplemented with 10% FBS, l-glutamine, and antibiotics (ThermoFisher). U2OS osteosarcoma cells were purchased from the ATCC and cultured in DMEM (Corning) supplemented with 10% FBS, l-glutamine, and antibiotics. U2OS and all ovarian cancer cell lines were authenticated in 2018 using short tandem repeat (STR) profiling by the McGill University Genome Center (Montreal, Canada). The model ovarian cancer cell line A2780 and its isogenic CDDP-resistant counterpart A2780Cis, both of which were authenticated by the distributor (Sigma-Aldrich) by STR profiling and used within 1 month following purchase, were grown in RPMI-1640 (Sigma-Aldrich) supplemented with 10% FBS, l-glutamine, and antibiotics. All cell lines were used within 20 passages after thawing and routinely tested for mycoplasma contamination by DAPI staining/fluorescence microscopy.

Ectopic RPA expression and siRNA treatment

The RPA expression plasmid (pAC-GFP-RPA; ref. 16), a generous gift of Dr. J. Lukas (University of Copenhagen, Copenhagen, Denmark), was transfected using Lipofectamine 2000 (ThermoFisher). Stable clones were sorted by FACS in 96-well plates and selected with 500 μg/mL Geneticin (ThermoFisher). siRNA smartpools for knockdown of RPA1 and BRCA1 were purchased from Dharmacon and transfected using RNAiMax (ThermoFisher). Pools of nontargeting (NT) duplexes were used as controls.

Cell irradiation and drug treatment

Cell monolayers were irradiated with 254-nm UV (hereafter UV) at a fluence rate of 0.4 J/m2/s, or with ionizing radiation (dose rate of 5 × 10−2 Gy/s) as previously described (24). The ATR inhibitor VE-821 (Cedarlane), hydroxyurea (HU; Bioshop Canada), cyclin-dependent kinase inhibitor roscovitine (Abcam), or MRE11 inhibitor mirin (Sigma-Aldrich) were added to cell monolayers in fresh media.

NER assay

Removal of 6-4 pyrimidine-pyrimidone photoproducts (6-4PP) as a function of cell cycle was quantified as described (19). For repair determinations in the presence of roscovitine, cells were treated with 20 μmol/L of the drug (or mock treated) for 2 or 72 hours before the assay and during post-UV incubations.

Clonogenic survival

Cells were seeded on 100 mm dishes and treated with CDDP (Cedarlane) or mitomycin C (MMC; Abcam) for 2 hours in serum-free medium. Camptothecin (CPT; Bioshop Canada) treatment was carried out for 2 hours in complete medium. Hydroxyurea was added for the duration of the experiment. Clonogenic survival was evaluated as previously described (19).

Immunoblotting

Western blotting was performed with whole-cell extracts using standard protocols and the following antibodies: mouse anti-RPA2 (Calbiochem; NA18), rat anti-tubulin (Abcam; ab6161), rabbit anti-RPA1 (Abcam; ab79398), mouse anti-RPA3 (Abcam; ab6432), rabbit anti-RPA2 phospho S33 (Abcam; ab211877), rabbit anti-Chk1 phospho S345 (Cell Signaling Technology; #2348), mouse anti-Chk1 (Cell Signaling Technology; #2360), mouse anti-BRCA1 (Santa Cruz Biotechnology; sc-6954), mouse anti-GAPDH (Santa Cruz Biotechnology; sc-365062), and rabbit anti-YY1 (Santa Cruz Biotechnology; sc-1703).

Quantification of CDDP-induced DNA adducts

Immediately after treatment with 20 μmol/L CDDP (2 hours in serum-free medium), genomic DNA was extracted in SNET buffer (20 mmol/L Tris-HCl, pH 8, 400 mmol/L NaCl, 1% SDS, 5 mmol/L EDTA, and 0.4 mg/mL proteinase K), followed by phenol/chloroform extraction and RNase A (Qiagen) treatment. DNA was quantified with PicoGreen (ThermoFisher) using a TBS-380 fluorimeter (Turner Biosystems). Note that 100 ng of DNA was analyzed by slot blotting with rat anti–cisplatin-modified DNA antibody (Abcam, ab103261) as described (18).

Immunofluorescence microscopy

Cells were seeded on glass slides in 6-well dishes and irradiated with UV or mock-treated. Posttreatment incubations were carried out in the presence of 30 μmol/L BrdUrd (Sigma-Aldrich). RPA and BrdUrd immunofluorescence were evaluated as described (18) using mouse anti-RPA2 (Calbiochem; NA18) and rat anti-BrdUrd (Abcam; ab6326). Secondary antibodies were goat anti–mouse-Alexa488 (ThermoFisher, A10667) and chicken anti–rat-Alexa594 (ThermoFisher; A21471). Images were acquired using a DeltaVision Elite system (GE Healthcare). Intensity of nuclear RPA2 signals was evaluated using custom software, as described (18). At least 200 cells combined from 2 independent experiments were quantified for each condition.

Quantification of EdU foci by fluorescence microscopy

Cells were seeded on glass slides in 6-well dishes and irradiated with 20 J/m2 of UV or mock-irradiated, followed by 1-hour incubation in complete media and then 30-minute incubation in the presence of 50 μmol/L EdU. Cells were then washed with PBS and fixed with 3% formaldehyde in PBS on ice for 20 minutes. Samples were washed with cold PBS, permeabilized on ice for 10 minutes with 0.5% triton X-100 in PBS, and washed again with PBS. Labeling was performed using click chemistry in PBS + 2 mmol/L CuSO4, 1 μmol/L Alexa Fluor 647 Azide, and 10 mmol/L l-ascorbate for 30 minutes at room temperature. Samples were washed with PBS and then labeled with 10 μg/mL DAPI in PBS. Images were acquired using a DeltaVision Elite system (GE Healthcare) with a 100x objective and deconvoluted using softWoRx 7.0.0. The intensity of nuclear EdU signals and number of foci were quantified using custom software as described previously (25). At least 200 cells combined from 2 independent experiments were quantified for each condition. Statistical analysis (Student t test) was used to compare the fold change in number of foci after UV relative to mock between SPR+ and SPR cell lines. Because different cell lines vary in the average size of their nucleus, the number of foci per cell was normalized to an equal arbitrary surface for easier visualization on a box plot. (This does not affect the statistical analysis above.)

Dual detection of γH2AX and RPA following CDDP treatment by flow cytometry

Exponentially growing cultures were treated or mock-treated with 5 μmol/L CDDP for 2 hours in serum-free medium, washed with PBS, and incubated for 24 hours. Following immunolabeling using anti-RPA1 (Abcam; ab79398) and anti–γH2AX Ser139 (Millipore), flow cytometry was performed as described (16).

DNA fiber analysis

DNA fiber assays were performed as described (12). Briefly, for DNA RF progression, cells were labeled for 15 minutes with 5 μmol/L IdU, washed, UV-irradiated, and then labeled for 90 minutes with 25 μmol/L CldU. For RF progression in the presence of HU, following labeling for 15 minutes with 5 μmol/L IdU, cells were washed whereupon 0.2 mmol/L HU was added together with 25 μmol/L CldU for a further 60-minute incubation. For RF progression experiments, the CldU/IdU ratio of each fiber in treated samples was normalized by dividing by the mean of CldU/IdU ratios observed after mock treatment. For RF protection assays, cells were labeled with 5 μmol/L IdU for 20 minutes followed by 25 μmol/L CldU for 20 minutes. After washing, cells were incubated with 30 mmol/L HU ± 100 μmol/L mirin for 3 hours. The CldU/IdU ratio of each fiber was normalized by dividing by the mean of CldU/IdU ratios observed immediately after CldU labeling (time 0). Imaging was performed using a DeltaVision Elite system (GE Healthcare) in conjunction with FIJI software (NIH). Experiments were performed at least twice independently, and a minimum of 125 fibers were counted for each experiment (250 fibers total per condition).

Pulsed-field gel electrophoresis

Cells were treated with 10 μmol/L CDDP in complete medium for 24 hours, trypsinized, washed, and resuspended in PBS. Agarose plugs composed of low melting point agarose (1% w/v) in PBS and containing 500,000 cells were prepared using a CHEF disposable plug mold (Bio-Rad). The plugs were digested with a solution of 1 mg/mL proteinase K, 100 mmol/L EDTA, 0.2% (w/v) sodium deoxycholate, and 1% (w/v) N-lauroyl sarcosinate at 37°C with low agitation for 2 days. Plugs were then washed 3 times for 1 hour with 20 mmol/L Tris HCl (pH 8.0), 50 mmol/L EDTA, and once with 0.5X TBE for 20 minutes. Pulsed-field gel electrophoresis was performed in a 0.9% (w/v) agarose gel in 0.5X TBE at 14°C under the following conditions. Block I: 12 hours at 5.5 V/cm, 0.1 to 30 seconds at 120°C. Block II: 12 hours at 3.6 V/cm, 0.1 to 5 seconds at 120°C. The gel was partially dried at room temperature for 30 minutes and then at 55°C for 30 minutes before staining overnight with SYBR Green (ThermoFisher) diluted in 0.5X TBE. Image acquisition was performed using a ChemiDoc imaging system (Bio-Rad).

Cisplatin resistance correlates with NER efficiency during S phase in ovarian cancer cell lines

Our overarching goal was to investigate the potential influence of replication stress–induced RPA exhaustion on CDDP resistance in HGSOC cells. We employed a panel of cell lines including eight HGSOC (OV866(2), TOV2223G, OV1369(R2), OV90, OV3331, OV1946, TOV2835EP, and TOV3041G) and two “non HGSOC” cell lines, i.e., one clear cell adenocarcinoma (TOV21G), and one endometrioid adenocarcinoma (TOV112D); see Supplementary Table S1 for cell line characterization). Clonogenic survival assays revealed a broad spectrum of CDDP sensitivities among cell lines (Fig. 1A), with LD50 differing by as much as 180-fold, ranging from 0.049 μmol/L for TOV21G to 9 μmol/L for TOV112D (Fig. 1A; Supplementary Table S2). As noted earlier, well-studied mechanisms contributing to CDDP resistance act by preventing the generation of DNA damage. We therefore quantified the initial induction of CDDP-induced intrastrand crosslinks in our HGSOC cell lines by slot blot (Fig. 1B, left) and found no correlation with drug sensitivity (Fig. 1B, right). This suggests that the ability to respond to DNA damage, as opposed to mechanisms that prevent its induction, might play a primary role in chemoresistance among cell lines of our panel.

Figure 1.

Cisplatin resistance correlates with SPR efficiency in ovarian cancer lines. A, Sensitivity to CDDP was measured by clonogenic survival. Values represent the mean ± SEM of three independent experiments. B, Cisplatin LD50s do not correlate with initial DNA damage induction as analyzed by slot blot (see text for details). Band intensities from three independent experiments were quantified by densitometry. C, Flow cytometry–based assay for measurement of 6-4PP excision as a function of cell cycle. Representative bivariate dot plot of 6-4PP versus DNA content (propidium iodide) for SPR-proficient OV866(2) and SPR-deficient TOV3041G. D, 6-4PP repair as a function of cell cycle at 6 hours post-UV in 12 ovarian cancer cell lines. Cells are considered SPR-deficient when SPR is significantly slower relative to other phases (*, P < 0.01; two-tailed paired t test). E, Cisplatin LD50s plotted against the percentage of unrepaired 6-4PP at 6 hours post-UV. F, UV LD50s were plotted against percentage of unrepaired 6-4PP. P value, F test of the linear regression.

Figure 1.

Cisplatin resistance correlates with SPR efficiency in ovarian cancer lines. A, Sensitivity to CDDP was measured by clonogenic survival. Values represent the mean ± SEM of three independent experiments. B, Cisplatin LD50s do not correlate with initial DNA damage induction as analyzed by slot blot (see text for details). Band intensities from three independent experiments were quantified by densitometry. C, Flow cytometry–based assay for measurement of 6-4PP excision as a function of cell cycle. Representative bivariate dot plot of 6-4PP versus DNA content (propidium iodide) for SPR-proficient OV866(2) and SPR-deficient TOV3041G. D, 6-4PP repair as a function of cell cycle at 6 hours post-UV in 12 ovarian cancer cell lines. Cells are considered SPR-deficient when SPR is significantly slower relative to other phases (*, P < 0.01; two-tailed paired t test). E, Cisplatin LD50s plotted against the percentage of unrepaired 6-4PP at 6 hours post-UV. F, UV LD50s were plotted against percentage of unrepaired 6-4PP. P value, F test of the linear regression.

Close modal

As mentioned earlier, defective responses to DNA replication stress cause excessive RPA sequestration at stalled RF, which reduces the availability of this complex for NER specifically during S (hereafter, S phase-specific nucleotide excision repair is referred to as SPR, and SPR-deficient and -proficient cell lines are denoted SPR and SPR+, respectively). Because NER is an important determinant of CDDP sensitivity, we employed a flow cytometry–based immunoassay (26) to evaluate the efficiency of this pathway as a function of cell cycle in our cell lines. This assay has been fully optimized to quantify removal of UV-induced 6-4PPs; however, we emphasize that results on 6-4PPs are extrapolatable to CDDP-induced intrastrand crosslinks because removal of either adduct is dependent on NER. We found that five HGSOC cell lines were SPR, i.e., exhibiting profound defects in NER during S versus G0–G1 or G2–M (Fig. 1C and D), whereas three repaired efficiently in all phases. We also characterized the two “non-HGSOC” cell lines, one of which was SPR+ and the other SPR (Fig. 1D). Remarkably, all SPR cell lines were significantly more sensitive to CDDP versus SPR+ counterparts (Fig. 1E; Supplementary Table S2). We also measured NER efficiency in A2780 and A2780Cis, an isogenic pair of ovarian cancer lines differing in CDDP sensitivity (27), and found that CDDP-resistant A2780Cis exhibited significantly more efficient SPR versus the parental CDDP-sensitive counterpart A2780 (Fig. 1D). Because we evaluated NER capacity by quantifying repair of UV-induced DNA adducts, we also determined clonogenic survival in response to this model mutagen and found a significant correlation between UV LD50 and SPR capacity (Fig. 1F; Supplementary Fig. S1A and Supplementary Table S2). The above data indicate that SPR defects are frequent in HGSOC cell lines and correlate strongly with CDDP and UV sensitivity.

We also evaluated sensitivity of our HGSOC cell lines to replication stress–inducing drugs for which, compared with the situations for UV and CDDP, NER plays only a limited role in removing the primary genotoxic adduct. The bifunctional alkylating agent MMC induces mainly DNA interstrand crosslinks resolved by the Fanconi anemia pathway in collaboration with HR (28), and the topoisomerase I inhibitor CPT generates replication-associated DSB repaired by HR and nonhomologous end-joining (29). Sensitivity to either MMC or CPT correlated significantly with SPR deficiency (Fig. 2A and B; Supplementary Fig. S1B–S1C; Supplementary Table S2); however, no correlation was noted for ionizing radiation, which induces DSB in all cell-cycle phases and does not kill cells primarily by causing replication stress (Fig. 2C; Supplementary Fig. S1D; Supplementary Table S2). Moreover, SPR deficiency correlated with sensitivity to HU, which causes replication stress without directly inducing DNA damage (Fig. 2D; Supplementary Fig. S1E; Supplementary Table S2). The above results suggest that defective SPR, while presumably contributing to enhanced CDDP/UV sensitivity in ovarian cancer cells, might represent only one consequence of abnormal DNA replication stress responses among others that confer such sensitivity.

Figure 2.

SPR efficiency correlates with sensitivity to MMC, CPT, and HU, but not ionizing radiation, in ovarian cancer cell lines. The percentage of 6-4PP remaining at 6 hours post-UV was plotted against the LD50 of MMC (A), CPT (B), ionizing radiation (C), and HU (D). P value, F test of the linear regression.

Figure 2.

SPR efficiency correlates with sensitivity to MMC, CPT, and HU, but not ionizing radiation, in ovarian cancer cell lines. The percentage of 6-4PP remaining at 6 hours post-UV was plotted against the LD50 of MMC (A), CPT (B), ionizing radiation (C), and HU (D). P value, F test of the linear regression.

Close modal

SPR-deficient CDDP-sensitive HGSOC cell lines exhibit abnormal DNA replication dynamics

Defective responses to replication stress often lead to excessive recruitment of RPA at stalled RF (17, 18). We measured chromatin-bound RPA at 3 hours post-UV in 3 SPR and 3 SPR+ HGSOC lines using immunofluorescence microscopy. We observed significantly increased chromatin-bound RPA during S (BrdUrd+ cells) in SPR cell lines compared with SPR+ counterparts (Fig. 3A); moreover, this could not be attributed to variations in RPA expression (Fig. 3B). We next evaluated RF progression post-UV using DNA fiber analysis and observed significantly reduced incorporation of labeled nucleoside analogs (CldU) post-UV in SPR versus SPR+ lines (Fig. 3C). These results suggest that elevated recruitment of RPA to chromatin, reflecting abnormal DNA replication dynamics during genotoxic stress, might underlie defective SPR in CDDP-sensitive HGSOC cell lines.

Figure 3.

SPR-defective HGSOC cell lines exhibit abnormal DNA replication dynamics. A, RPA2 recruitment to chromatin post-UV was measured by immunofluorescence microscopy. BrdUrd labeling distinguishes S-phase cells. Left, representative images for TOV2835EP. Right, fluorescence intensity of the nuclear RPA signal in HGSOC lines. Values for the mock treatment (black) are compared with values post-UV (blue). Red lines, median values. B, Immunoblot analysis of RPA subunit levels in SPR and SPR+ ovarian cancer lines. C, Effect of UV (20 J/m2) on RF progression was measured using DNA fiber analysis. Red lines, median values. D, Effect of UV on the number of active RF following UV irradiation (see text for details). Left, fluorescence microscopy of EdU foci in TOV2835EP. Right, the number of EdU foci per nucleus in each line was quantified using custom software. E, Left, mean intensity of nuclear EdU signal post-UV, relative to mock, from D. Right, ratio of EdU incorporation post-UV (relative to mock-treated cells) divided by the normalized ratio of RF progression as measured in C to estimate the number of active origins post-UV. F, Effect of roscovitine on SPR following 2- or 72-hour treatment. P values, Student t test comparing mean between SPR+ and SPR lines. G, Induction of phospho-Chk1 (S345), phospho-RPA2 (S33), and γH2AX 4 hours after CDDP treatment.

Figure 3.

SPR-defective HGSOC cell lines exhibit abnormal DNA replication dynamics. A, RPA2 recruitment to chromatin post-UV was measured by immunofluorescence microscopy. BrdUrd labeling distinguishes S-phase cells. Left, representative images for TOV2835EP. Right, fluorescence intensity of the nuclear RPA signal in HGSOC lines. Values for the mock treatment (black) are compared with values post-UV (blue). Red lines, median values. B, Immunoblot analysis of RPA subunit levels in SPR and SPR+ ovarian cancer lines. C, Effect of UV (20 J/m2) on RF progression was measured using DNA fiber analysis. Red lines, median values. D, Effect of UV on the number of active RF following UV irradiation (see text for details). Left, fluorescence microscopy of EdU foci in TOV2835EP. Right, the number of EdU foci per nucleus in each line was quantified using custom software. E, Left, mean intensity of nuclear EdU signal post-UV, relative to mock, from D. Right, ratio of EdU incorporation post-UV (relative to mock-treated cells) divided by the normalized ratio of RF progression as measured in C to estimate the number of active origins post-UV. F, Effect of roscovitine on SPR following 2- or 72-hour treatment. P values, Student t test comparing mean between SPR+ and SPR lines. G, Induction of phospho-Chk1 (S345), phospho-RPA2 (S33), and γH2AX 4 hours after CDDP treatment.

Close modal

Increased frequency of stalled RF due to aberrant activation of replication origins, e.g., upon abrogation of ATR activity, can cause RPA exhaustion (16). Moreover, other carcinogenic mechanisms including oncogene activation that increase the number of active RF (30, 31) might also diminish RPA availability. No correlation was observed between SPR status and rate of proliferation, percentage of cells in S, or S-phase duration (Table 1; Supplementary Fig. S2A–S2C), indicating that SPR cells do not exhibit significantly higher rates of replication versus SPR+ counterparts in the absence of replication stress. We next evaluated the number of active origins during replication stress in SPR+ versus SPR HGSOC lines by quantifying the number of sites of DNA synthesis per nucleus, i.e., EdU foci representing replication origins, using fluorescence microscopy. This analysis revealed that in SPR+ cell lines, the number of EdU foci at 1 hour post-UV is comparable with that in the absence of UV treatment (Fig. 3D). However, SPR cell lines exhibit a significant increase in EdU focus formation post-UV, suggesting dysregulated origin firing during replication stress. To further validate these results, we adapted a method reported elsewhere (32) to estimate the number of active RF in our cell lines upon replication stress. From the experiment immediately above, we evaluated total EdU signal intensity after UV relative to mock treatment and found that it was similar in SPR+ versus SPR cells (Fig. 3E, left). Because the progression of individual RF is reduced in SPR versus SPR+ cell lines post-UV (Fig. 3C), the above data suggest that SPR cells present elevated sites of EdU incorporation. To quantify this, nuclear EdU signal intensity was divided by the CldU/IdU ratios presented in Fig. 3C. For SPR+ cell lines, this value was below one, consistent with robust inhibition of origin firing after UV. In contrast, SPR cell lines displayed significantly higher ratios, indicating increased origin activation relative to SPR+ counterparts (Fig. 3E, right plot).

Table 1.

Rates of proliferation and S-phase duration of cell lines used in this study

SPRDoubling time (days)% cells in SS-phase duration (hours)
OV866(2) 1.9 ± 0.1 33 ± 4 9.8 ± 0.8 
OV1369(R2) 2.2 ± 0.1 33 ± 2 11 ± 1 
TOV2223G 2.1 ± 0.1 28 ± 3 7.8 ± 0.7 
TOV2835EP – 2.8 ± 0.4 38 ± 2 10.2 ± 0.6 
OV1946 – 1.8 ± 0.1 43 ± 2 8.4 ± 0.5 
TOV3041G – 2.8 ± 0.1 37 ± 3 10.6 ± 0.9 
U2OS RPA-GFP 1.2 ± 0.1 50.0 ± 0.7 9.3 ± 0.2 
U2OS GFP – 0.95 ± 0.05 49.7 ± 0.9 9.0 ± 0.3 
SPRDoubling time (days)% cells in SS-phase duration (hours)
OV866(2) 1.9 ± 0.1 33 ± 4 9.8 ± 0.8 
OV1369(R2) 2.2 ± 0.1 33 ± 2 11 ± 1 
TOV2223G 2.1 ± 0.1 28 ± 3 7.8 ± 0.7 
TOV2835EP – 2.8 ± 0.4 38 ± 2 10.2 ± 0.6 
OV1946 – 1.8 ± 0.1 43 ± 2 8.4 ± 0.5 
TOV3041G – 2.8 ± 0.1 37 ± 3 10.6 ± 0.9 
U2OS RPA-GFP 1.2 ± 0.1 50.0 ± 0.7 9.3 ± 0.2 
U2OS GFP – 0.95 ± 0.05 49.7 ± 0.9 9.0 ± 0.3 

NOTE: Values are the mean ± SEM of three independent experiments. Quantification of S-phase duration is described in Supplementary Fig. S2A–S2C.

We next evaluated the impact of aberrant origin activity on the phenotypes of CDDP-sensitive (SPR) HGSOC cell lines. CDK activity permits activation of origins during S (33), and inhibition of these kinases by roscovitine diminishes origin firing and RPA recruitment to chromatin upon replication stress (16). Strikingly, we found that roscovitine treatment significantly rescued SPR defects (Fig. 3F), consistent with a key role of aberrant origin activity in abrogating SPR in ovarian cancer cell lines. We note that phosphorylation of Chk1 (S345), RPA2 (S33), and γH2AX are all comparable in SPR+ and SPR cell lines following CDDP treatment (Fig. 3G). Thus, although the precise mechanism underlying abnormal activation of replication origins in SPR cell lines remains unknown, it is unlikely to be attributable to defective ATR/Chk1 activation upon replication stress.

Exhaustion of RPA due to elevated origin activity can eventually engender widespread induction of DSB at stalled RF (16). Using a flow cytometry assay (16), we found that 24 hours after CDDP treatment, the proportion of SPR cell lines displaying elevated signals for both chromatin-bound RPA and γH2AX (a marker of DSB) was significantly higher in all cases compared with SPR+ counterparts (Fig. 4A). We also used pulsed-field gel electrophoresis to directly evaluate generation of DSB 24 hours after CDDP treatment (Fig. 4B). In agreement with the aforementioned result, SPR cell lines manifested significantly higher DSB induction compared with SPR+ counterparts. Taken together, our results indicate that CDDP sensitivity in HGSOC cell lines strongly correlates with abnormal activation of replication origins, SPR defects, and RF instability upon replication stress.

Figure 4.

SPR cell lines exhibit increased γH2AX and DNA DSB after CDDP treatment. A, RPA recruitment to chromatin and DSB formation after CDDP treatment. Left, representative bivariate plots of γH2AX and RPA for TOV3041G. ATR inhibition (VE-821) was used as positive control. Right, percentage of cells with high γH2AX and RPA chromatin levels for HGSOC lines. B, PFGE analysis of CDDP-induced DSB. Left, representative gel. The bands corresponding to DSB were quantified by densitometry. Right, fold induction of DSB signal after CDDP, relative to mock. Values are mean ± SEM of three independent experiments. P values, Student t test comparing mean between SPR+ and SPR lines.

Figure 4.

SPR cell lines exhibit increased γH2AX and DNA DSB after CDDP treatment. A, RPA recruitment to chromatin and DSB formation after CDDP treatment. Left, representative bivariate plots of γH2AX and RPA for TOV3041G. ATR inhibition (VE-821) was used as positive control. Right, percentage of cells with high γH2AX and RPA chromatin levels for HGSOC lines. B, PFGE analysis of CDDP-induced DSB. Left, representative gel. The bands corresponding to DSB were quantified by densitometry. Right, fold induction of DSB signal after CDDP, relative to mock. Values are mean ± SEM of three independent experiments. P values, Student t test comparing mean between SPR+ and SPR lines.

Close modal

Nascent DNA at stalled RF is unstable in SPR versus SPR+ HGSOC cell lines

We used DNA fiber analysis to further evaluate RF dynamics in our panel of cell lines upon exposure to low concentrations of HU, a ribonucleotide reductase inhibitor that induces RF stalling without directly generating DNA lesions (34, 35). Consistent with our observation that SPR HGSOC cell lines are sensitive to HU (Fig. 2D), we observed that all three SPR HGSOC lines exhibit reduced replication progression rate in response to this drug compared with SPR+ counterparts (Fig. 5A). This further supports the notion that the observed impairment in RF progression upon genotoxin exposure in SPR cell lines is unlikely to result solely from defective SPR.

Figure 5.

Nascent DNA at stalled RF is unstable in SPR-deficient HGSOC cell lines. A, Effect of HU on RF progression by DNA fiber analysis. B, RF protection upon HU treatment (± mirin) by DNA fiber analysis. C, Top plot, depletion of BRCA1 in SPR+ lines using siRNA versus NT controls. Bottom, RF protection ± mirin in BRCA1 knockdown SPR+ lines. D, 6-4PP removal after BRCA1 knockdown. Cells treated with ATR inhibitor were used as positive control for defective SPR. E, 6-4PP removal ± mirin. For bar graphs, values represent the mean ± SEM of three independent experiments. P values, Student t test (A, B, D, and E) or Mann–Whitney U test (C). For DNA fiber experiments, red lines are median values.

Figure 5.

Nascent DNA at stalled RF is unstable in SPR-deficient HGSOC cell lines. A, Effect of HU on RF progression by DNA fiber analysis. B, RF protection upon HU treatment (± mirin) by DNA fiber analysis. C, Top plot, depletion of BRCA1 in SPR+ lines using siRNA versus NT controls. Bottom, RF protection ± mirin in BRCA1 knockdown SPR+ lines. D, 6-4PP removal after BRCA1 knockdown. Cells treated with ATR inhibitor were used as positive control for defective SPR. E, 6-4PP removal ± mirin. For bar graphs, values represent the mean ± SEM of three independent experiments. P values, Student t test (A, B, D, and E) or Mann–Whitney U test (C). For DNA fiber experiments, red lines are median values.

Close modal

As mentioned previously, the HR factors BRCA1/2 and Rad51 ensure protection of nascent DNA at stalled RF from Mre11-dependent degradation (hereafter “RF protection”; refs. 12, 36), which in turn promotes overall RF progression upon exposure to replication-blocking genotoxins (37). Moreover, RF protection significantly mitigates the cytotoxicity of chemotherapeutics, including CDDP (12). Given the strong association between SPR efficiency and chemosensitivity, we explored possible correlations between SPR status and RF protection in HGSOC cell lines. Quantification of RF protection was performed by successive incubations with IdU and CldU, followed by prolonged incubation with a concentration of HU that completely inhibits RF progression. Under these conditions, nascent DNA degradation upon RF stalling leads to CldU/IdU ratios <1. Interestingly, SPR cell lines all manifest strong RF protection defects in response to HU, which were rescued by concomitant treatment with the Mre11 inhibitor mirin (Fig. 5B). Importantly, only one of the interrogated cell lines was found to have undetectable BRCA1 expression, which probably explains its RF protection defect, whereas all six lines had normal BRCA2 and Rad51 levels (Supplementary Fig. S3). Moreover, DNA sequencing did not reveal mutations in BRCA1/2 in these cell lines (Supplementary Table S1); therefore, the molecular basis of RF protection defects in SPR cell lines OV1946 and TOV2835EP currently remains unknown. Nevertheless, our data indicate that RF protection and SPR defects may often coexist in chemosensitive ovarian cancer cell lines, suggesting possible mechanistic overlap between these phenomena.

RPA availability modulates chemosensitivity and nascent DNA stability in ovarian cancer cell lines

Because nascent DNA degradation is expected to generate ssDNA and consequent RPA recruitment to stalled RF, we reasoned that RF protection defects might contribute to SPR inhibition by diminishing RPA availability. Contrary to this notion, although siRNA-mediated knockdown of BRCA1 in 3 SPR+ cell lines caused strong Mre11-dependent RF protection defects (Fig. 5C), it did not compromise SPR (Fig. 5D). Importantly, exposure to VE-821, a potent ATR inhibitor, generated marked SPR defects in the 3 SPR+ cell lines as expected, ruling out the possibility that these might somehow be immune to the impact of replication stress–induced RPA sequestration. Conversely, we found that mirin treatment, which resolves Mre11-dependent RF protection defects (Fig. 5B), did not restore SPR in deficient cell lines (Fig. 5E). Overall, the above data indicate that Mre11-dependent degradation of nascent DNA at stalled RF does not engender SPR defects in HGSOC cell lines.

We considered the alternative possibility that RPA exhaustion leading to compromised SPR might also contribute to RF protection defects in HGSOC lines. For technical reasons, i.e., high cloning efficiency, we used U2OS osteosarcoma cells as model for our initial experiments. We previously showed that U2OS exhibits profound SPR defects that can be rescued by ectopic overexpression of RPA (18). We generated a U2OS clone stably expressing the three RPA subunits in a stoichiometric manner, as well as a control GFP-expressing clone. In agreement with our previous finding using transient overexpression (18), stable RPA overexpression rescued SPR defects in the U2OS clone (Fig. 6A), confirming that limited availability of RPA is a major cause of SPR deficiency in this system. Moreover, consistent with the notion that RPA sequestration influences nascent DNA stability, we found (i) that SPR-defective U2OS cells present Mre11-dependent RF protection defects upon HU treatment and (ii) that both RF protection and progression defects in this cell line are rescued upon RPA overexpression (Fig. 6B). We emphasize that the effects of RPA overexpression on NER and replication stress described immediately above are not likely caused by differences in the rate of DNA synthesis, as no differences were noted in the rate of S-phase progression/duration, or the fraction of cells in S phase, between the RPA-overexpressing and control U2OS clones (Table 1).

Figure 6.

RPA availability modulates chemosensitivity and nascent DNA stability. A, A clone of U2OS was selected for stable stoichiometric overexpression of RPA-GFP (GFP only as control). Left, immunoblot of RPA subunits. Recombinant RPA1 and RPA2 retain a P2A cleavage sequence and have a higher MW. RPA3 is fused to GFP. Right, removal of 6-4PP. B, RF protection and progression assays as in Fig. 5. C, RPA binding to chromatin after cisplatin treatment in U2OS following siRNA against RPA1 and/or BRCA1 was measured by flow cytometry. D, RF protection in U2OS during HU treatment following siRNA against BRCA1 and/or RPA1. E, Clones of SPR-deficient OV1946 were selected that overexpress RPA-GFP. F, 6-4PP removal post-UV. G, Fork progression post-UV by DNA fiber assay. H, RF protection during HU treatment ± mirin. I, Clonogenic survival following cisplatin treatment. P values, Student t test (A and FI) or Mann–Whitney U test (B and D). For DNA fiber experiments, red lines are median values.

Figure 6.

RPA availability modulates chemosensitivity and nascent DNA stability. A, A clone of U2OS was selected for stable stoichiometric overexpression of RPA-GFP (GFP only as control). Left, immunoblot of RPA subunits. Recombinant RPA1 and RPA2 retain a P2A cleavage sequence and have a higher MW. RPA3 is fused to GFP. Right, removal of 6-4PP. B, RF protection and progression assays as in Fig. 5. C, RPA binding to chromatin after cisplatin treatment in U2OS following siRNA against RPA1 and/or BRCA1 was measured by flow cytometry. D, RF protection in U2OS during HU treatment following siRNA against BRCA1 and/or RPA1. E, Clones of SPR-deficient OV1946 were selected that overexpress RPA-GFP. F, 6-4PP removal post-UV. G, Fork progression post-UV by DNA fiber assay. H, RF protection during HU treatment ± mirin. I, Clonogenic survival following cisplatin treatment. P values, Student t test (A and FI) or Mann–Whitney U test (B and D). For DNA fiber experiments, red lines are median values.

Close modal

To further characterize the link between RPA availability and RF protection, we used siRNA to knockdown BRCA1 and RPA1, either alone or in combination, in U2OS (Fig. 6C, left). BRCA1 depletion, while not causing a significant increase in RPA recruitment to chromatin following CDDP treatment as assessed by flow cytometry (Fig. 6C, right), strongly exacerbated the existing RF protection defect in U2OS (Fig. 6D). We note that the lack of effect of BRCA1 depletion on chromatin-bound RPA is consistent with results indicating that RF protection defects associated with BRCA1 depletion do not compromise SPR in HGSOC cells (Fig. 5C and D). Strikingly, siRNA-mediated knockdown of RPA1 impaired RF protection to a similar extent as for BRCA1-depleted cells, which could be reversed by Mre11 inhibition (Fig. 6D). Moreover, codepletion of RPA1 and BRCA1 caused an even stronger Mre11-dependent RF protection defect (Fig. 6D). These data imply that reduced RPA availability (i) promotes Mre11-dependent degradation of nascent DNA at stalled RF and (ii) can synergize with other genetic anomalies, e.g., BRCA1 deficiency, to cause RF protection defects.

We sought to perform similar experiments in SPR HGSOC cell lines, but for unknown reasons could only generate RPA-overexpressing clones in OV1946. We selected three independent OV1946 clones overexpressing recombinant RPA (Fig. 6E). Although GFP-expressing controls exhibited profoundly reduced SPR, RPA overexpression completely restored NER proficiency (Fig. 6F). In addition, RPA overexpression rescued RF protection and progression defects in response to HU (Fig. 6G and H), as well as markedly increased CDDP resistance (Fig. 6I). These data indicate that reduced RPA availability contributes to RF instability and chemosensitivity in HGSOC cells.

Here, we sought to elucidate the role of DNA replication stress–induced RPA exhaustion in promoting CDDP sensitivity using a panel of patient-derived HGSOC cell lines. We initially turned our attention to NER, which, as discussed earlier, (i) is believed to represent an important determinant of CDDP resistance in ovarian cancer and (ii) is sensitive to replication stress–induced fluctuations in RPA availability during S phase. Specifically, we exploited a cell-cycle–specific repair assay to demonstrate that a majority of HGSOC cell lines exhibit strong defects in NER during S phase relative to G0–G1 and G2–M. Moreover, remarkably, these SPR cell lines are all significantly (as much as 180-fold) more sensitive to CDDP relative to SPR+ counterparts. Together with our previous observations in melanoma and model cancer cell lines (19, 26), this suggests (i) that reduced SPR capacity represents a common feature of human cancers and (ii) that SPR tumors in vivo, although challenging at present to identify as such, may respond much better to replication-blocking chemotherapeutic drugs including CDDP.

Our data indicating that SPR defects in HGSOC lines can be attributed to sequestration of RPA at aberrantly activated replication origins are consistent with recently published results showing that defective ATR signaling causes RPA exhaustion at least in part due to misregulated origin firing (16). Although our results do not support a causative role for ATR/Chk1 activation defects in compromising SPR in HGSOC cells, we cannot exclude that downstream components of S-phase checkpoint-dependent inhibition of origin firing might be involved. In addition, activation of various oncogenes, e.g., Ras, Myc, and cyclin E1, dysregulates replication origin initiation programs in human cells (31, 38), although their impact on RPA availability and SPR remains unknown. Collectively, our data suggest that yet-to-be-identified cancer-associated genetic alterations resulting in failure to prevent unscheduled origin activity might enhance chemosensitivity in HGSOC cells by causing RPA exhaustion and SPR defects. We note that although RPA is well characterized for its essential roles in DNA replication and repair, this multifunctional complex has also recently been implicated in transcription (39–41). Thus, it is interesting to speculate that generation of ssDNA at sites of RNA polymerase stalling at CDDP-damaged sites in DNA might (i) sequester RPA and (ii) promote replication stress via collisions between the blocked transcriptional machineries and DNA polymerases (42).

Defective SPR is not the sole consequence of RPA exhaustion, as this complex is involved in a number of other important cellular processes. Indeed, we found that exposure to CDDP causes increased RPA recruitment to chromatin and DSB formation in SPR versus SPR+ HGSOC lines, which is reminiscent of the situation for RPA exhaustion in cells treated with ATR inhibitors (16). Although RPA is required to promote recruitment/activity of a multitude of DDR factors at stalled RF (14), we note that the precise mechanisms through which reduced RPA availability compromises RF stability remain unclear. Notably, recent proteomic surveys of RF-associated proteins in situations where RPA pools are exhausted did not reveal striking changes in protein recruitment to stalled RF (43). Alternatively, it is possible that unprotected ssDNA at stalled RF might be more susceptible to breakage arising from enzymatic activities present in HGSOC cells. For example, recently published data demonstrate that ssDNA at stalled RF is frequently targeted by the APOBEC family of cytidine deaminases, eventually leading to DSB induction via base excision repair–mediated incisions (44). Elucidation of the complete spectrum of molecular mechanisms leading to DSB induction at RF upon RPA exhaustion in HGSOC cells will require further investigation.

We found that, relative to SPR+ counterparts, SPR HGSOC cell lines exhibit extensive Mre11-dependent degradation of nascent DNA at HU-stalled RF, raising the possibility that regions of ssDNA thus generated might cause RPA sequestration and associated phenotypes. However, we provide several lines of evidence arguing against this possibility: (i) Abrogation of BRCA1, which causes strong RF protection defects, is not sufficient to trigger either RPA exhaustion or inhibition of SPR, and (ii) preventing nascent DNA degradation at stalled RF via pharmacologic inhibition of Mre11 did not rescue SPR. Although RF protection defects do not cause RPA exhaustion, the converse appears to be true since we found that siRNA-mediated knockdown of RPA1 results in RF protection defects comparable with those observed in BRCA1-depleted cells. These findings are generally in accord with recently published data indicating that disruption of RPA1 interaction with ssDNA due to PTEN downregulation diminishes RF progression rates upon HU exposure (45). The mechanisms responsible for the impact of RPA exhaustion on Mre11-dependent degradation of nascent DNA at RF are unknown. Phosphorylated RPA32 interacts with PALB2, which itself promotes BRCA2 recruitment (46, 47); however, as mentioned above, a recent study revealed that RPA availability does not significantly alter protein recruitment at stalled RF (43). We note that histone posttranslational modifications strongly influence nascent DNA stability (12). It is therefore possible that diminished RPA availability might influence chromatin structure at RF, in turn causing increased susceptibility to nuclease-mediated nascent DNA degradation.

Importantly, we show that ectopic RPA overexpression protects the SPR HGSOC cell line OV1946 from CDDP-induced cell death while rescuing SPR proficiency and RF stability. The notion that RPA availability influences CDDP resistance is in agreement with published data showing that cell killing by this drug is potentiated by mutations that cripple RPA binding to DNA (48). Moreover, our data indicate that RPA sequestration can cause RF protection defects, which are also expected to strongly contribute to chemosensitivity (12). In this regard, it is interesting to note that high RPA expression, which based on our results might potentially overcome RF instability upon replication stress, has been reported to be predictive of adverse outcome in ovarian cancer treatment (48, 49). This raises the interesting possibility that elevated RPA availability might be selected for during the course of chemotherapy treatment, leading to reversal of chemosensitivity. Overall, our results lead us to propose that modulation of RPA protein levels/dynamics during S is a critical determinant of CDDP resistance in ovarian tumors via multiple mechanisms. Cellular pathways that influence RPA availability may therefore be expected to harbor useful biomarkers for predicting treatment response in HGSOC.

No potential conflicts of interest were disclosed.

Conception and design: F. Bélanger, E. Fortier, M. Dubé, J.-Y. Masson, A.-M. Mes-Masson, H. Wurtele, E. Drobetsky

Development of methodology: F. Bélanger, E. Fortier, A. Elsherbiny, H. Wurtele, E. Drobetsky

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): F. Bélanger, E. Fortier, M. Dubé, J.-F. Lemay, R. Buisson, A. Elsherbiny, E. Drobetsky

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): F. Bélanger, E. Fortier, M. Dubé, J.-F. Lemay, R. Buisson, J.-Y. Masson, A. Elsherbiny, S. Costantino, H. Wurtele, E. Drobetsky

Writing, review, and/or revision of the manuscript: F. Bélanger, E. Fortier, M. Dubé, R. Buisson, S. Costantino, E. Carmona, A.-M. Mes-Masson, H. Wurtele, E. Drobetsky

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): M. Dubé, E. Drobetsky

Study supervision: H. Wurtele, E. Drobetsky

This study was funded by operating grant #364096 from the Canadian Institutes of Health Research (CIHR) awarded to E. Drobetsky, H. Wurtele, and A.-M. Mes-Masson. J.-Y. Masson is recipient of a CIHR Foundation Award. H. Wurtele, E. Fortier, and J.-F. Lemay are supported by salary awards from the Fonds de Recherche du Québec-Santé (FRQS). The Centre de Recherche de l'Hôpital Maisonneuve-Rosemont and Centre Hospitalier de l'Université de Montréal (CRCHUM)/Institut du Cancer de Montréal receive institutional funding from the FRQS. For ovarian tumor banking, A.-M. Mes-Masson is supported by the Banque de Tissus et de Données of the Réseau de Recherche sur le Cancer and FRQS, affiliated with the Canadian Tumor Repository Network (CTRNet).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

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