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Failure to reactivate stalled or collapsed DNA replication forks is a potential source of genomic instability. Homologous recombination (HR) is a major mechanism for repairing the DNA damage resulting from replication arrest. The single-strand DNA (ssDNA)-binding protein, replication protein A (RPA), plays a major role in multiple processes of DNA metabolism. However, the role of RPA2 hyperphosphorylation, which occurs in response to DNA damage, had been unclear. Here, we show that hyperphosphorylated RPA2 associates with ssDNA and recombinase protein Rad51 in response to replication arrest by hydroxyurea (HU) treatment. In addition, RPA2 hyperphosphorylation is critical for Rad51 recruitment and HR-mediated repair following HU. However, RPA2 hyperphosphorylation is not essential for both ionizing radiation (IR)-induced Rad51 foci formation and I-Sce-I endonuclease-stimulated HR. Moreover, we show that expression of a phosphorylation-deficient mutant of RPA2 leads to increased chromosomal aberrations following HU treatment but not after exposure to IR. Finally, we demonstrate that loss of RPA2 hyperphosphorylation results in a loss of viability when cells are confronted with replication stress whereas cells expressing hyperphosphorylation-defective RPA2 or wild-type RPA2 have a similar sensitivity to IR. Thus, our data suggest that RPA2 hyperphosphorylation plays a critical role in maintenance of genomic stability and cell survival after a DNA replication block via promotion of HR.
DNA double-strand breaks (DSBs) may arise spontaneously, e.g. during DNA replication or following exposure to DNA-damaging agents (1), such as ionizing radiation (IR) or chemotherapeutic drugs. Homologous recombination (HR) is a major pathway in the repair of DSBs, especially those arising from stalled/collapsed replication forks (2–4). Defective HR may lead to genetic exchanges that result in genomic instability. In addition, cells deficient in HR are sensitive to IR and some chemotherapeutic drugs, especially S phase cells because HR preferentially repairs DSBs in this phase of the cell cycle.
The molecular mechanism of HR can be subdivided further, depending on the nature of the DNA structure. Two-ended DSBs can be caused directly by IR; however, only one-ended DSBs or no DSBs are produced as a result of damage or discontinuities during interrupted replication (5,6). The HR mechanism required for repairing two-ended DSBs has been extensively studied in the past. In general, it is believed that HR is triggered when a two-ended DSB is processed to a 3′ single-strand DNA (ssDNA) tail via resection. Once the ssDNA is generated, it is rapidly bound by the ssDNA-binding protein replication protein A (RPA) that in turn is displaced by Rad51. The resultant Rad51 filament facilitates DNA strand invasion and exchanges steps. Although the mechanisms required for HR after DNA replication stalling have not yet been defined in mammalian cells, it appears that eukaryotic cells have evolved a mechanism similar to that described in bacteria for re-establishment of replication forks after their progression has been impeded by lesions in the template. It has been suggested that similar to RuvABC complex in Escherchia coli, the endonuclease Mus81 in mammalian cells contributes to the generation of one-ended DSBs by cleavage of stalled DNA replication forks, which subsequently promotes HR (7–9). In some instances, replication may be restarted downstream of the unrepaired parental strand lesion, leaving a ‘daughter strand gap’ (DSG)—a ssDNA lesion in which no DSB-end is involved. It has been suggested that DSG repair in E.coli requires the RecFOR complex, which initiates with RecA, an equivalent of human Rad51, loading at ssDNA region (10). Studies in mammalian cells have shown the existence of DSGs during stalled DNA replication (11–14), but it is not clear how this process is regulated. Therefore, the detailed molecular mechanisms of the HR that occur in response to replication arrest in mammalian cells is much less understood compared with the HR process stimulated by two-ended DSBs.
RPA is a heterotrimer composed of three tightly associated subunits, RPA1, RPA2 and RPA3. The major ssDNA-binding subunit is RPA1. However, the other two subunits also participate in the interactions with ssDNA (15,16). RPA is required for almost all aspects of cellular DNA metabolism, including DNA replication, DNA damage checkpoint activation, DNA repair and recombination. During DNA replication, RPA binds to ssDNA and is required for the initiation as well as elongation processes (17,18). In addition to its role in DNA replication, RPA is also critical for cell cycle checkpoint activation: RPA-coated ssDNA recruits the ataxia-telangiectasia and Rad3-related (ATR)–ATR interacting protein complex, which facilitates S phase checkpoint signaling under the condition of replication arrest (19,20). RPA is also essential for multiple DNA repair pathways, including HR-mediated repair. RPA is thought to facilitate two-ended DSBs stimulated HR by removing secondary structures in ssDNA that may prevent the formation of the Rad51 filament (21,22) or by actively regulating protein–protein interactions (23). However, the role of RPA in HR after replication block remains unknown.
RPA becomes phosphorylated in cells when bound to ssDNA. Although no precise phosphorylation site has been identified in RPA1, nine potential phosphorylation sites have been suggested within the unstructured N-terminal domain of RPA2 (24–29). A novel DNA damage-induced phosphorylation site at Thr-98 in RPA2 was also recently identified (26). RPA2 is phosphorylated in both cell cycle and DNA damage-dependent manner. S phase-dependent phosphorylation of RPA2 is mediated by the cyclin-dependent kinase2 kinase. Cyclin-dependent kinase modifies RPA2 at serine residues 23 and 29 in mitotic cells. DNA damage-induced RPA2 phosphorylation, so-called RPA2 hyperphosphorylation includes modification of threonine 21, serines 4, 8 and 33 as well as at least one phosphoserine in residues 11–13. These events are carried out by the phosphatidylinositol 3-OH-kinase-related kinase family, which includes ataxia-telangiectasia mutated, ATR and DNA-dependent protein kinase (17). A recent study suggests that cyclin-dependent kinase is also required for DNA damage-induced RPA2 hyperphosphorylation (27). However, the question remains as to whether and how RPA2 hyperphosphorylation regulates the function of RPA in DNA replication, cell cycle checkpoint and DNA repair.
Earlier studies suggested that hyperphosphorylation of RPA2 may not be essential for RPA function in vitro or in vivo (18) since deletion of the N-terminus of RPA32 does not affect ssDNA-binding activity, heterotrimer complex formation or the ability to support DNA replication in vitro. However, recent evidence suggests a role for RPA2 hyperphosphorylation in DNA metabolism. It has been suggested that RPA2 phosphorylation facilitates mitotic exit in response to mitotic DNA damage (30). Several studies indicated that RPA2 hyperphosphorylation negatively regulates the role of RPA in DNA replication both in vitro and in vivo (17) although the mechanism underlying the inhibition of replication by RPA2 hyperphosphorylation is controversial. Despite the negative regulation of DNA replication, RPA2 hyperphosphorylation is likely to be involved in DNA repair, particularly HR-mediated repair due to the following reasons. First, RPA2 phosphorylation stimulates repair of chromosomal DNA damage caused either by camptothecin or bleomycin treatment (27). Second, cells expressing a RPA2 mimetic that has persistent phosphorylation exhibit normal checkpoint activation and re-enter the cell cycle normally after recovery but display a pronounced defect in the repair of DNA breaks (31). Since HR-mediated repair is a major mechanism to repair DNA damage caused by stalled/collapsed DNA replication forks, these results suggest a role for RPA2 phosphorylation in HR. Third, hyperphosphorylated RPA2 was found to directly interact with Rad51 in vitro (32) and phosphorylation of RPA2 increased the affinity between RPA and Rad51. Fourth, the observation that hyperphosphorylated RPA2 occurs predominantly in late S and G2 phases (27), where HR is a major repair mechanism, suggests a role for RPA hyperphosphorylation in HR. Therefore, we tested the biological function of RPA2 phosphorylation in HR that occurs in response to DNA damage, particularly after replication arrest.
Here, we demonstrate that hyperphosphorylated RPA2 colocalizes with ssDNA and Rad51 in response to hydroxyurea (HU)-induced DNA replication fork stalling. Cells expressing phosphorylation-defective RPA2 exhibit a defect in Rad51 foci formation and a decreased frequency of HR in response to replication arrest but have no obvious defect on IR-induced Rad51 recruitment and I-Sce-I-induced HR. In addition, an increased frequency of chromosomal abnormalities after treatment with HU, but not upon exposure to IR, was observed in cells expressing phosphorylation-defective RPA2. Therefore, we propose that RPA2 hyperphosphorylation is required for HR, which plays a critical role in the maintenance of genomic stability and cell survival in response to replication arrest.
MCF-7 cells were obtained from the American Type Culture Collection (Manassas, VA). Retroviral vectors carrying a HA-tagged wild-type RPA2 or phosphorylation mutant RPA2, RPA2-(7xA)-(S4A/S8A/S11A/S12A/S13A/T21A/S33A), have been described previously (33). RPA2 small hairpin RNA (shRNA) was generated based on lentiviral pLKO.1-puro vector (Sigma, St Louis, MO). Lentivirus transfer vector in which RNA polymerase III U6 promoter drives the expression of small hairpin RNAs complementary to RPA2 3′-untranslated region of messenger RNA target sequence was produced based on previously published sequence (34). In brief, the oligonucleotides encoding RPA2 was cloned into the AgeI and EcoRI sites of the pLKO.1-puro vector. All the DNA plasmid transfections were performed using Lipofectamine according to the manufacturer's recommendations (Invitrogen, Carlsbad, CA).
HA-tagged wild-type RPA2 (RPA2-WT) or RPA2-(7xA)-mutant form (RPA2-A) were introduced into MCF7 cells by three rounds of retroviral infection. Silencing of endogenous RPA2 in these cells was accomplished by two rounds of lentiviral infection using pLKO.1 vector expressing the sequence targeting RPA2 (RPA2 shRNA). The cells were irradiated with X-rays or treated with HU for Rad51 foci analysis or chromosome aberration and survival assay.
The cells were treated with 0.5% Triton 100 cytoskeletal buffer and fixed with 4% formaldehyde. The fixed cells were permeablized using 0.1% Triton X-100 phosphate-buffered saline for 15 min, followed by blocking with 10% fetal bovine serum and then incubated with primary antibodies. The bound antibodies were revealed by goat anti-mouse IgG Alexa fluor 594 and chicken anti-rabbit IgG Alexa Fluor 488. The slides were viewed at ×1000 magnification on an Olympus fluorescence microscope (BX40 with Magna-Fire CCD camera). The protocol for ssDNA detection was conducted according to a protocol described previously (35).
Proteins from whole cell lysates were resolved by sodium dodecyl sulphate–polyacrylamide gel electrophoresis. The primary antibodies used for this analysis are anti-HA tag antibody (Covance, Princeton, NJ; 1:1000 dilution), rabbit polyclonal antibody phosphor RPA32 Ser4/Ser8 (Bethyl, Montgomery, TX; BL647, 1: 1000 dilution), monoclonal antibody RPA2 (EMD Bioscience, Brookfield, WI; Ab-2, 1:100 dilution) and polyclonal antibodies RPA2 (Abcam, Cambridge, MA; 1:1000 dilution). Secondary antibodies used were goat anti-mouse IgG–horseradish peroxidase conjugated and goat anti-rabbit IgG horseradish peroxidase conjugated (Pierce, Rockford, IL), both at 1:5000 dilutions.
The Neutral Comet Assay was performed using the Comet Assay kit from Trevigen (Gaithersburg, MD) following manufacturer's instructions. The lyses occurred at 4°C for 30 min. Comets were analyzed using CometScore software (TriTek, Sumerduck, VA).
The HR analysis induced by I-Sce-I expression has been described previously (36). In brief, the HR was measured in MCF7-pDR-green fluorescent protein (GFP) cells with RPA2-WT or RPA2-A expression where the endogenous RPA2 was depleted by shRPA2. The HR analysis upon replication arrest was conducted according to the protocol described previously (36). Briefly, the cells were treated for 24 h with 2 mM thymidine (Thy). The cells were subsequently washed in phosphate-buffered saline and incubated in fresh medium for 3 h and then the cells were treated with 2 mM HU for 24 h. All cells were left for 24 h to recover before being subjected to flow cytometric analysis for GFP-positive cells. For each analysis, at least 1000000 cells were processed.
Florescence in situ hybridization was performed using pan-telomeric and pan-centromeric peptide nucleic acid probes. Telomere (C3TA2)3-specific and centromere (16-mer repeat DNA)-specific probes directly labeled with Cy3 and fluorescein isothiocyanate fluorescent dyes, respectively, were obtained from Applied Biosystems (Foster City, CA).The cells were radiated (2 Gy) or treated with HU (1 mM, 6 h) and processed with florescence in situ hybridization analysis 24 h later. The cells were incubated with colcemid (0.1 μg/ml, Sigma) for 2.5 h before fixation onto slides. The slides were treated with 1 mg/ml pepsin/0.01 N HCl for 10 min at 37°C, after fixed in 4% formaldehyde, dehydrated in ethanol series (70, 90 and 100%), stained with a telomeric and centromeric probe mixture, denatured 3 min at 80°C and hybridized in a moist chamber for 2 h at reverse transcription, then dehydrated again and sealed in anti-fading 4′,6-diamidino-2-phenylindole mounting (Vector laboratories Burlingame, CA).
Cells expressing RPA2-WT or RPA2-A where endogenous RPA2 has been deleted were used in a colony formation assay to measure cell survival after HU or IR treatment. The cells were harvested, filtered with cell strainers (BD Falcon™) to achieve single-cell suspension and plated in triplicates into P60 dishes at various cell densities, with a target number of surviving colonies at 50–100 per dish. Treatment with HU or IR was carried out 18 h after cell plating. After a 24 h exposure to HU at 0.1, 0.5, 1, 2 and 10 mM of concentrations or IR at 2, 4 and 6 Gy of doses, cells were rinsed twice with phosphate-buffered saline and allowed to grow in fresh medium for 14–21 days. Visible colonies were identified with methanol fixation and 0.35% methylene blue staining. The number of colonies (>50 cells) per dish was counted, and the surviving fractions were calculated as the ratio of the plating efficiencies of treated cells to untreated cells.
To test whether hyperphosphorylation of RPA2 is critical for the DNA damage response, we first analyzed RPA2 hyperphosphorylation profiles in response to various DNA-damaging agents, including the replication inhibitors HU and Thy and the DSB inducer IR. HU inhibits ribonucleotide reductase and leads to DNA synthesis arrest. Thy disrupts DNA synthesis by altering the balance between thymidine triphosphate and deoxycytidine triphosphate through the allosteric inhibition of ribonucleotide reductase. Both, HU and Thy, inhibit the DNA elongation step of replication. While Thy does not generate detectable DSBs (37), HU causes DSBs that are obvious after ~12 h of continuous treatment as assessed by pulsed-field gel electrophoresis (36). In contrast to the replication inhibitors HU and Thy, IR can directly cause DNA DSBs. In agreement with previous studies (33), exposure to HU and Thy significantly increased RPA2 hyperphosphorylation (Figure 1A, first panel), as shown by using an antibody specific for phosphorylated RPA2. However, this hyperphosphorylation can only be detected after exposure to a high dose of IR (Figure 1B, first panel) (39,40). We also detected the amount of DSBs in the cells treated with replication inhibitors as well as IR using the comet assay under neutral conditions, which allows detection of DSBs but not ssDNA breaks. No obvious DSBs were detected after 6 h or 24 h of continuous treatment with HU or Thy. In contrast, IR causes enormous DSBs at the doses utilized (Figure 1C). Therefore, our observation is consistent with the previous report that RPA2 hyperphosphorylation is more prevalent when replication forks are stalled rather than when DSBs are generated (33). Since it has been shown that at high doses of IR DNA replication chain elongation is blocked (41), the observation that only higher doses of IR cause RPA2 hyperphosphorylation may be caused by the blockage of DNA replication forks. Alternatively, since it has been suggested that ssDNA resection is restricted to S and G2/M phase cells and RPA2 hyperphosphorylation induced by stalled/collapsed DNA replication is ATR dependent (33), a second explanation is that higher doses of IR can cause more ssDNA, which can activate ATR and subsequently RPA2 hyperphosphorylation. Together, the results shown in Figure 1 and previous reports (33) suggest that RPA2 hyperphosphorylation is more prevalent when DNA replication forks are stalled.
Here, we need to note that our observation and others (33) are not inconsistent with a recent publication showing that rapid hyperphosphorylation of RPA2 are observed in response to IR which is ataxia-telangiectasia mutated and DNA-dependent protein kinase dependent. This form of IR-induced RPA2 hyperphosphorylation was only observed in mitotic cells but not in interphase cells (42). Thus, hyperphosphorylation of RPA2 can take place in specific mitotic cells when DSBs are generated, a process that is dependent on ataxia-telangiectasia mutated and DNA-dependent protein kinase (42) but primarily upon stalling of replication forks, an ATR-dependent process (33).
Arrest of replication fork progression should lead to the accumulation of ssDNA. We tested if hyperphosphorylated RPA2 is localized at ssDNA regions during replication block. We first examined whether HU induces ssDNA by immunofluorescence staining. ssDNA was detected by using an assay that permits identification of sites of ssDNA >1 kb in situ. This assay is based on the observation that the nucleotide base analogue bromodeoxyuridine (BrdU) is recognized by an anti-BrdU antibody when incorporated into ssDNA but not dsDNA (35). The approach has been successfully utilized in different laboratories (35,43,44). We found that HU treatment significantly increased the percentage of cells with ssDNA in vivo, indicating that ssDNA accumulated in response to replication arrest (Figure 2A). To determine if hyperphosphorylated RPA colocalized with ssDNA, cells treated with HU were costained with BrdU and phospho-specific RPA2 antibody. An extensive colocalization of hyperphosphorylated RPA2 with ssDNA foci was observed (Figure 2B) indicating that hyperphosphorylated RPA2 associates with ssDNA regions after a replication block. Given the requirement of DNA binding for RPA2 hyperphosphorylation, we reasoned that phosphorylated RPA2 foci occur later than RPA2 recruitment. Therefore, we examined the recruitment of RPA2 and hyperphosphorylated RPA2 at various time points after HU treatment by immunofluorescence staining using anti-RPA2 and specific anti-phosphorylated RPA2 antibodies. We found that RPA2 recruitment starts to increase as early as 1 h following HU treatment. In contrast, HU-induced phosphorylated RPA2 foci were not observed until 2 h after HU treatment (Figure 2C). Representative RPA2 and hyperphosphorylated RPA2 foci in response to HU are shown in Figure 2C. Moreover, RPA2 foci and phosphorylated RPA2 foci show nearly complete colocalization (Figure 2C). This result indicates that RPA2 is recruited to ssDNA and then is further hyperphosphorylated, which is consistent with previous observations that RPA2 can only be hyperphosphorylated after binding to DNA or chromatin (33,45,46).
The accumulated ssDNA during stalled/collapsed DNA replication forks could lead to DSBs. The increased DSBs induced by HU treatment were detected by pulsed-field gel electrophoresis (36). Consistent with this report, an increase in cells with γ-H2AX foci after HU treatment has also been observed (47). DSBs and other lesions associated with DNA replication are repaired by HR in mammalian cells (37,48). A key protein in HR-mediated repair is RAD51, and cells deficient in RAD51 accumulate DSBs after replication or at stalled replication forks (49). Since it has been demonstrated that hyperphosphorylated RPA2 directly interacts with Rad51 (32), we next determined if hyperphosphorylated RPA2 colocalized with Rad51 in response to replication arrest. Hyperphosphorylated RPA2 was observed to extensively colocalize with Rad51 in response to HU treatment (Figure 3A), suggesting the association of hyperphosphorylated RPA2 with Rad51 in vivo in stalled/collapsed DNA replication response pathways. To further study the role of phosphorylated RPA2 in Rad51 recruitment, we used an RPA2 mutant construct in which seven DNA damage-associated phosphorylation sites (33) were substituted by alanine (RPA2-A). Retroviral vectors carrying a HA-tagged wild-type RPA2 or phosphorylation mutant RPA2, RPA2-(7xA)-(S4A/S8A/S11A/S12A/S13A/T21A/S33A) have been described previously (33) (Figure 3B). Like wild-type RPA2 (RPA2-WT), the RPA2-(7xA)-mutant (RPA2-A) co-immunoprecipitated with RPA1 (33) (data not shown), indicating that substitution of the serine residues in the N-terminus of the protein did not affect the interaction of RPA2 with RPA1. Also, disruption of the phosphorylation sites of RPA2 does not affect RPA heterotrimer formation, the normal S phase functions of RPA2 or cell growth (33,34). We tested Rad51 recruitment in cells expressing RPA2-WT or RPA2-A where endogenous RPA2 was depleted by shRNA targeting the 3′-untranslated region of the RPA2 messenger RNA (34). As shown in Figure 3C, endogenous RPA2 expression was reduced to undetectable levels via transduction with RPA2 shRNA. Exogenous RPA2-WT and RPA2-A proteins were resistant to shRNA RPA2 and expressed the proteins at similar levels. Strikingly, exogenous RPA2-A expression showed a major reduction in percentage of cells with positive Rad51 foci after HU treatment compared with cells expressing RPA2-WT (Figure 3D) (T-test, P <0.01, P=0.002), indicating that RPA2 hyperphosphorylation is required for Rad51 assembly. The specificity of Rad51 antibody was verified by the observation that depletion of Rad51 using Rad51 small interfering RNA abolished HU-induced Rad51 foci formation in cells expressing RPA2-WT (data not shown). Representative Rad51 foci are shown in Figure 3E (upper panel). The effect of RPA2-A on Rad51 recruitment was not caused by a deficiency in RPA recruitment to chromatin since normal RPA foci formation was observed in cells expressing RPA2-WT or RPA2-A in response to HU (See supplementary Figure S1, available at Carcinogenesis Online). This is consistent with previous reports showing that defective RPA2 phosphorylation doesnot affect recruitment of RPA to chromatin or the sites of damaged DNA (33,34). In order to test if RPA2 hyperphosphorylation plays a role in Rad51 recruitment in response to two-ended DSBs, the formation of Rad51 foci was compared in cells expressing RPA2-WT or RPA2-A after exposure to IR. We found that Rad51 foci formation was not affected by RPA2 hyperphosphorylation since a similar proportion of cells with IR-induced Rad51 foci was observed in cells expressing RPA2-WT or RPA2-A (Figure 3D), indicating the RPA2 hyperphosphorylation is not essential for the Rad51 recruitment in response to the classical two-ended DSBs. Representative Rad51 foci are shown in Figure 3E (bottom panel).
Notably, RPA2 hyperphosphorylation has no obvious effect on the fraction of cells with Rad51-positive foci in untreated cells (Figure 3D, T-test, P>0.05). This finding is consistent with our observation that no obvious RPA2 hyperphosphorylation was observed in cells without any treatment (Figure 1A and B). Our finding further supports the previous study suggesting that the repair mechanism of spontaneous HR is different to that induced by replication inhibitors (50). Collectively, the data in Figure 3 demonstrate that hyperphosphorylated RPA2 accumulates in ssDNA regions and is important for the formation of Rad51 foci after replication block.
The observation that RPA2 hyperphosphorylation affects Rad51 recruitment in response to replication arrest led us to speculate that RPA2 hyperphosphorylation plays a role in HR after replication arrest. One of the main systems established for the study of HR processes is the measurement of the occurrence of HR in artificial recombination substrates. This assay is broadly used to study two-ended DSBs stimulated HR by overexpressing the I-Sce-I endonuclease protein (51) or to study the HR in response to replication block by HU and Thy or topoisomerase I inhibitor camptothecin (36,50,52–54). Therefore, we next examined the biological consequence of expressing RPA2-A on a previously established HR assay system (51). The established MCF7 cell line containing an integrated copy of the pDR-GFP reporter (MCF7 DR-GFP) was described previously (55). It is based on reconstruction of the wild-type GFP from two non-functional fragments of GFP complementary DNA (51). The protocol for HR induced by stalled/collapsed DNA replication forks was conducted according to a previous publication (36). To improve the frequency of HR, the cells were first synchronizated by double thy block that results in 70–90% of cells in the S phase (data not shown). The resulting S phase cells were treated with HU. We found that cells expressing RPA2-A displayed a decreased frequency of HR triggered by HU treatment in comparison to cells expressing RPA2-WT (Figure 4A) (T-test, P<0.05, P=0.006), suggesting that RPA2 hyperphosphorylation is critical for HR-mediated repair after replication arrest.
We next examined the biological consequence of expressing RPA2-A on HR induced by rare-cutting I-Sce-I endonuclease. We found no changes in the frequency of HR in cells expressing RPA2-A compared with cells expressing RPA2-WT (Figure 4B) (T-test, P >0.05, P=0.18), indicating that RPA2 hyperphosphorylation has no obvious role on HR induced by I-Sce-I expression. This data suggests that RPA2 hyperphosphorylation is specifically required for HR after replication block but not in response to two-ended DSBs. Our findings could be explained by the fact that no detectable RPA2 hyperphosphorylation or RPA2 phosphorylation foci were observed by I-Sce-I overexpression, whereas they were easily detected in cells after HU treatment (Figures 1A and 4C and D). We need to note that the frequency of the HR induced by HU is lower than that stimulated by I-Sce-I expression due to the genome-wide effect of replication inhibition compared with the targeted DSB induced with I-Sce-I (50). Together, these findings strongly support a key role for RPA2 hyperphosphorylation in HR triggered by DNA replication arrest.
Failure to restart stalled/collapsed replication forks is a potential source of genomic instability. In cells without DNA damage, DNA replication duplicates each chromosome during S phase to create a sister chromatid. If the chromosomal DNA replication is blocked, the ssDNA generated due to replication arrest could be processed into DSBs (36). In normal cells, the DSBs are repaired by HR since only one-end DSBs are created, which is not an appropriate substrate for non-homologous end joining. However, if DSBs are not repaired by HR, these persistent DSBs will result in chromatid breaks or chromosome breaks. The deficient HR and the lack of HU-induced Rad51 focus formation in cells expressing RPA2-A indicates a defective response to stalled/collapsed DNA replication forks, which could lead to genomic instability. Therefore, we examined whether a deficiency in RPA2 hyperphosphorylation results in the accumulation of chromosomal abnormalities in response to HU using florescence in situ hybridization. We observed that cells expressing RPA2-A displayed higher frequencies of chromatid and chromosome breaks after exposure to HU (Figure 5A and B) compared with cells expressing RPA2-WT. The representative metaphases spread after HU treatment in cells expressing RPA2-A is shown in Figure 5B.
Most importantly, cells expressing RPA2-A exhibited a higher rate of radial structures (Figure 5C), such as tri-radial and quadri-radial aberrations (Figure 5D), generated by the increased frequency of chromatid breaks and subsequent interchromatid fusions. The presence of radial chromosomes in cells expressing RPA2-A further reveals a link between RPA2 hyperphosphorylation and HR since cells deficient in HR often show similar chromosome aberrations, such as the cells deficient in the FA/BRCA2 pathway (56,57). These findings highlight an important role of RPA2 hyperphosphorylation in maintenance of genomic stability during DNA replication. Next, we tested the role of hyperphosphorylated RPA2 in maintenance of genomic stability after IR. We found that cells expressing RPA2-A display similar frequencies of chromatid and chromosome breaks after exposure to IR (Figure 5E). Thus, Figure 5 indicates that RPA2 hyperphosphorylation is specially required for maintenance of genomic stability in response to replication arrest, which perfectly fits its role in Rad51 foci formation and HR induced by HU (Figures 2 and and33).
The HR-mediated repair is required to restart stalled or collapsed replication forks under replication stress conditions. The role of hyperphosphorylated RPA2 in Rad51 recruitment and HR in response to HU indicates that RPA2 hyperphosphorylation plays a role in cell survival after replication arrest. To test this hypothesis, we determined the effect of RPA2 hyperphosphorylation on survival following HU treatment or IR exposure. As shown in Figure 6A, cells expressing RPA2-A produced a phenotype of greater sensitivity to HU by clonogenic cell survival (Figure 6A). In contrast, the cells expressing hyperphosphorylation mutant RPA2 showed a similar sensitivity to IR (Figure 6B). Thus, Figure 6 suggests that RPA2 hyperphosphorylation is required for cell survival under conditions of replication arrest, which is correlated with its role in Rad51 recruitment and HR stimulated by HU treatment (Figure 3). It should be noted that although hyperphosphorylation of RPA2 plays a role in inhibition of DNA replication after activation of S phase checkpoint during replication stress, the defects of HR in cells expressing phosphorylation defective RPA2 may explain the sensitivity to HU (Figure 6A) since it has been indicated that S phase checkpoint is not required for cell viability under replication stress conditions (58).
The activation of oncogenes and inactivation of tumor suppressor genes frequently results from cancer-promoting mutations. These genetic alterations are often mediated by impaired DSB repair. RPA is an essential factor for DSB repair and cell cycle checkpoint activation. In the present report, we demonstrate that hyperphosphorylated RPA2 colocalizes with ssDNA and Rad51 and is essential for Rad51 recruitment and HR in response to replication arrest. In addition, we also demonstrate that RPA2 hyperphosphorylation is critical for maintenance of genomic stability and cell viability after replication stalling. These findings provide direct evidence that RPA2 hyperphosphorylation facilitates HR and is important for maintenance of genomic stability and cell survival under conditions of replication stress in addition to its role on mitotic exit in response to mitotic DNA damage (30).
Consistent with previous reports, we found that RPA2 hyperphosphorylation profoundly occurs following replication arrest (Figure 1). This result can be explained by studies showing that it is not only the ssDNA but also the nucleic acid structure at stalled replication forks that regulates function of ATR (59–61). The single-strand/double-strand junctions, which reflect the structure at the replication forks, are able to stimulate RPA2 hyperphosphorylation by ATR more effectively than ssDNA fragment (61). In our study, we observed that hyperphosphorylation of RPA2 is required for HR in response to replication arrest but not two-ended DSBs-induced HR. Our work further supports the notion that the HR mechanisms required for repairing classical two-end DSBs and restart for stalled/collapsed DNA replication are different (36,50,53). Several models have been proposed in response to the stalled or collapsed DNA replication in E.coli, including one-ended DSBs-stimulated recombination-dependent replication and direct-restart mechanism of reactivating forks without any DSBs creation (4). A potential mechanism required for repair of ssDNA gaps, DSG, has also been proposed (4,62). Whether RPA2 hyperphosphorylation regulates any or all these subpathways of HR in mammalian cells is a very interesting topic to be addressed in the future.
A question raised by our studies is why RPA2 hyperphosphorylation is specifically required for HR after replication arrest. It has been shown that hyperphosphorylation induced a conformational change in RPA (63). Because RPA hyperphosphorylation does not significantly alter the high affinity of RPA for ssDNA (28,63), a likely scenario is that hyperphosphorylated RPA located at DNA damage sites after replication arrest facilities the recruitment of factors for DNA repair, such as Rad51, a key player for HR-mediated repair. The components that associate with RPA during replication arrest most likely differ from those associated with RPA in response to two-ended DSB. Rad51 may not be efficiently recruited to the sites of stalled/collapsed DNA replication forks. The conformational change caused by RPA2 hyperphosphorylation is necessary for Rad51 recruitment to the damaged DNA sites under the conditions of replication arrest. This speculation is supported by a recent study showing that the affinity of RPA to Rad51 is increased when RPA is hyperphosphorylated. In addition, we observed that deficient RPA2 phosphorylation failed to recruit Rad51 (Figure 3) although RPA is recruited to the damaged DNA sites normally (supplementary Figure S1 is available at Carcinogenesis Online) (33,34).
Given the complexity of DNA replication, S phase is a vulnerable period for the genome in the cell-division cycle. During replication arrest, long regions of RPA-coated ssDNA at replication forks are necessary for ATR checkpoint signaling. Interestingly, RPA2 is also a direct substrate of ATR (33). The evidence suggests a regulated process in which ssDNA-bound RPA activates the ATR kinase, which in turn induces RPA hyperphosphorylation. In addition, it has been documented that hyperphosphorylation of RPA2 prevents RPA from participating in DNA replication in response to DNA damage. In the present study, we show that RPA2 hyperphosphorylation is essential for HR after stalled or collapsed DNA replication forks (Figure 4). Therefore, hyperphosphorylated RPA2 appears to serve as a central regulator in response to replication arrest, which is evidenced by the increased chromosome instability and cell toxicity observed in cells lacking RPA2 hyperphosphorylation (Figures 5 and and6).6). Therefore, a possible scenario under conditions of replication stress is that ssDNA bound by RPA actives ATR and subsequently RPA2 hyperphosphorylation induced by ATR promotes HR via facilitation of Rad51 recruitment (supplementary Figure S2 is available at Carcinogenesis Online).
Interestingly, a recent study suggests that dephosphorylation of RPA2 is also important to DNA repair process after replication arrest, and persistent RPA2 phosphorylation leads to the increased DSBs (31). The complexity of DNA DSB repair processing requires that the individual reaction steps are ordered. The HR-mediated repair process in a highly organized process. RPA2 hyperphosphorylation and dephosphorylation could be an important mechanism to regulate the assembly and/or disassembly of protein complexes at the site of stalled/collapsed DNA lesions in a sequential order. Therefore, HR-mediated repair in response to stalled/collapsed DNA replication forks is a strictly regulated process in which both phosphorylation and dephosphorylation of RPA is required. Defects in either part of the regulation process results in genomic instability.
We have shown that defective RPA2 hyperphosphorylation results in the enhanced sensitivity to HU, which is an approved therapeutic agent for chronic myelogenous leukemia. HU is commonly used for other myeloproliferative disorders and also used occasionally for metastatic melanoma and other cancers. Given that hyperphosphorylated RPA2 can also be induced by topoisomerase I inhibitor camptothecin, cross-linking agents cisplatin and other chemotherapeutic drugs (50,64), our studies should have significant clinical implications. A recent study has suggested that RPA2 hyperphosphorylation confers resistance to cisplatin and other chemotherapy drugs (65). Therefore, the evaluation of RPA2 hyperphosphorylation can be potentially used as an indicator for the sensitivity/resistance to these chemotherapeutic agents. The understanding of the molecular mechanism by which RPA2 phosphorylation promotes HR should eventually lead to the development of a novel class of anticancer drugs targeting RPA2 hyperphosphorylation.
A seed grant and startup fund from the Department of Radiation Oncology, Washington University School of Medicine by the Dr J.R.R.; Stephanie Pagano Fund for Mesothelioma Research by a grant from the American Cancer Society (IRG-58-010-51); Wendy Will Case Cancer Fund to J.Z.
We apologize for those whose work was not cited due to the limited space. We thank M.Jasin for their generous contribution of materials.
Conflict of Interest Statement: None declared.