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PARP-1 ensures regulation of replication fork progression by homologous recombination on damaged DNA
Correspondence to Kazuto Sugimura: sugimura{at}bio.mie-u.ac.jp; or Katsuzumi Okumura: katsu{at}bio.mie-u.ac.jp
Poly-ADP ribose polymerase 1 (PARP-1) is activated by DNA damage and has been implicated in the repair of single-strand breaks (SSBs). Involvement of PARP-1 in other DNA damage responses remains controversial. In this study, we show that PARP-1 is required for replication fork slowing on damaged DNA. Fork progression in PARP-1–/– DT40 cells is not slowed down even in the presence of DNA damage induced by the topoisomerase I inhibitor camptothecin (CPT). Mammalian cells treated with a PARP inhibitor or PARP-1–specific small interfering RNAs show similar results. The expression of human PARP-1 restores fork slowing in PARP-1–/– DT40 cells. PARP-1 affects SSB repair, homologous recombination (HR), and nonhomologous end joining; therefore, we analyzed the effect of CPT on DT40 clones deficient in these pathways. We find that fork slowing is correlated with the proficiency of HR-mediated repair. Our data support the presence of a novel checkpoint pathway in which the initiation of HR but not DNA damage delays the fork progression.
© 2008 Sugimura et al. This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.jcb.org/misc/terms.shtml). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).
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Poly-ADP ribosylation is a posttranslational modification catalyzed by poly-ADP ribose polymerase 1 (PARP-1) and PARP-2 and is one of the earliest cellular responses to DNA damage. PARP-1 and -2 bind to the damage sites and activate themselves by automodification. This process causes chromatin decondensation around damage sites, recruitment of repair machineries such as DNA ligase III–XRCC-1 base excision repair complexes, and accelerates DNA damage repair, especially in the case of SSBs (Caldecott et al., 1996; Masson et al., 1998; Leppard et al., 2003). PARP-1 also seems to affect DSB repair because PARP-1–deficient cells are hypersensitive to DSB-inducing agents, especially to CPT (Chatterjee et al., 1989; Bowman et al., 2001; Pommier et al., 2003). Biochemical studies showed that NHEJ proteins such as Ku and DNA-PK are poly-ADP ribosylated by PARP-1, and the affinity of Ku to DSBs was decreased (Ariumi et al., 1999; Galande and Kohwi-Shigematsu, 1999; Li et al., 2004). Moreover, a genetic study showed that PARP-1 could protect the HR pathway from toxic interference by Ku70 (in DT40 and mammalian cells) in response to CPT-induced DSBs (Hochegger et al., 2006). These studies suggest that PARP-1 can interfere with NHEJ, which might be unfavorable for DSB repair in S phase and provide access for the HR machineries. These findings have also suggested the possibility that PARP-1 is involved in the regulation of replication fork progression coupled with HR-dependent DSB repair via its poly-ADP ribosylation activity. However, the role of PARP-1 in HR-coupled replication fork modulation in vivo has not been fully investigated.
In this study, we investigated the contributions of PARP-1 to the progression of replication forks on CPT-damaged DNA using an in vivo replication labeling assay and a dynamic molecular combing technique that allowed us to analyze the progression of individual replication forks in vivo. Interestingly, the addition of CPT significantly slowed down fork progression when PARP-1 was active, whereas inhibition or deficiency of PARP-1 resulted in unchanged fork progression rates in the presence of CPT. We also found that this slowing down of the replication fork in wild-type cells depended on the HR pathway, which was diminished in PARP-1 knockout cells as a result of negative interference by Ku.
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-H2AX foci were detected only in the presence of CPT in S-phase nuclei (Fig. 2 A).
First, we performed Western blotting to detect poly-ADP ribosylated proteins using FITC-NAD+ as a substrate. Compared with untreated cells, the poly-ADP ribosylation level in CPT-treated cells was higher over a wide size range of proteins (Fig. 2 B). NU1025 treatment led to the disappearance of poly-ADP ribosylated proteins. Using fluorescence microscope analysis, we confirmed that poly-ADP ribosylation was inhibited by NU1025 in the presence of CPT and found that NU1025 sensitized cells to CPT (Fig. 2, C and D).
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To further evaluate the effect of PARP on replication forks, PARP-1 was reduced with a PARP-1–specific siRNA. PARP-1 was effectively decreased in HeLa cells at 48 and 72 h after transfection of two individual siRNAs (Fig. 3, A and B). Compared with cells transfected with the negative control siRNA, the poly-ADP ribosylation level was decreased to the background level (Fig. 3 C). The ratio of fork rates during IdU and CldU labeling was about twofold higher in cells transfected with PARP-1–specific siRNA compared with those with the negative control siRNA in the presence of CPT (Fig. 3 D). Consistent with this result, the entire distribution of fork rates during CldU labeling shifted to faster fork rates in the presence of CPT in those cells transfected with the PARP-1 siRNAs compared with cells transfected with negative control siRNA (Fig. 3 E). These results indicate that PARP-1 can affect replication fork progression on a damaged chromosome caused by CPT.
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To address a role of PARP-1 in SSB repair (SSBR), we analyzed the rate of fork progression in polymerase β–deficient DT40 cells, but CPT-induced fork slowing was not reversed in polymerase β–/– cells (Fig. S2, A and B, available at http://www.jcb.org/cgi/content/full/jcb.200806068/DC1). To confirm these results in mammalian cells, HeLa cells in which another SSBR factor, XRCC1, was knocked down by siRNA were used. The protein level of XRCC1 was significantly reduced at 48 or 72 h after siRNA transfection (Fig. S2 C). Similar to polymerase β–/– DT40 cells, CPT-induced fork slowing was not recovered in XRCC1-depleted HeLa cells (Fig. S2, D–F), suggesting that PARP-1 regulates replication fork progression in response to CPT-induced DNA damage independently of its SSBR-related activity.
We hypothesized that a PARP-1–related DSB repair pathway is responsible for CPT-induced fork slowing. To address this hypothesis, we tested two DT40 mutants in DNA DSB repair pathways, XRCC3–/– mutant cells that are defective in HR and Ku70–/– mutants that are defective in NHEJ. Interestingly, slowing of replication fork progression occurred at the same levels as wild type in the Ku70–/– single mutant but was clearly reversed in XRCC3–/– cells (Fig. 5 and Fig. S3, available at http://www.jcb.org/cgi/content/full/jcb.200806068/DC1). No effect of NU1025 was observed in XRCC3–/– cells. Similar results were observed in other HR-defective BRCA2tr/– DT40 cells (Fig. S3, C and D; Hatanaka et al., 2005). These results suggest that replication forks slow down in response to CPT as a result of the activation of the HR pathway.
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In this study, we found that PARP-1 can slow replication forks in response to DNA damage induced by CPT. We focused on the PARP-1–related repair pathways and found that the HR-dependent repair pathway may be involved in the modulation of fork progression on damaged templates in vertebrate cells (Fig. 5 C). Previous studies indicated that PARP-1 interacts with the Ku–DNA-PK complex that is required for the NHEJ pathway (Ariumi et al., 1999; Galande and Kohwi-Shigematsu, 1999; Li et al., 2004), and PARP-1 poly-ADP ribosylates Ku, decreasing the binding affinity of Ku to DSBs (Li et al., 2004). After these observations, Hochegger et al. (2006) suggested a biological role of the interaction of PARP-1 and NHEJ factors, proposing that PARP-1 minimized the suppressive effect of Ku and the NHEJ pathway on HR after replication fork collapse. Consequently, PARP-1 deficiency could result in reduced efficiency of HR because of the presence of Ku, resulting in enhanced progression of the replication fork. A previous study supports this idea, which found that replication fork arrest or slowing on damaged chromosomes was induced in an HR-dependent manner (Henry-Mowatt et al., 2003). Therefore, the reason why concomitant deletion of Ku70 in PARP-1–/– mutants resulted in fork slowing in response to CPT is likely to be reestablishment of HR-mediated DSB repair processes.
We think the reason why fork rates in PARP-1–/– cells were the same as those of wild-type cells in the presence of CPT because of the following points. A mutant of the checkpoint protein, Hus1, showed normal replication fork rates in the presence of CPT (Wang et al., 2004). This mutant is also deficient in the HR process (Wang et al., 2006). Intriguingly, PCNA dissociates from chromatin in response to CPT treatment in wild-type cells but not in the Hus1 mutant, indicating that restoration of chromatin binding of PCNA appears to be associated with a normal rate of fork progression in the presence of CPT. However, another study reported that Ku facilitated the association of PCNA to chromatin in response to DSBs (Park et al., 2004), so it is conceivable that restoration of chromatin binding of PCNA by Ku associates with normal fork rates on damaged DNA in PARP-1 or HR-defective cells. Indeed, we found that the chromatin-bound PCNA decreased in response to CPT but not in the presence of NU1025 (unpublished data). Heller and Marians (2006) showed that replication could be restarted and that leading-strand synthesis reinitiated downstream of an unrepaired block to leading-strand progression. If this occurs downstream of a DSB, de novo loading of PCNA by Ku might facilitate the restart. Further in-depth investigations are required to test this hypothesis.
We also suggest an alternative plausible model. Recently, the molecular details of a cytotoxic mechanism for CPT were suggested (Koster et al., 2007). They suggested that the accumulation of positive supercoils was induced by CPT ahead of the replication machinery and that DNA damages were induced within the supercoil. If PARP-1 or HR is deficient, Ku-mediated chromosomal translocation might occur within the supercoil. Presumably, the replication fork could then move on the translocated chromosome. However, we do not have any evidence to support this hypothesis.
In summary, we have shown that PARP-1 activity influences replication fork progression when replication forks collapse at CPT-induced single-strand gaps. The target of PARP-1 activity in this novel mechanism appears to be Ku, which interferes with HR-dependent DSB repair at the collapsed replication fork.
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Antibodies
Mouse anti-BrdU antibody was purchased from BD. Rat anti-BrdU antibodies were purchased from Oxford Biotechnology or Abcam. Mouse antiphosphohistone H2A.X (Ser139) antibody was purchased from Millipore. Mouse anti–PARP-1 and mouse anti–poly-ADP ribose antibodies were purchased from Trevigen. Goat anti–PARP-1 antibody was purchased from R&D Systems. Mouse anti–β-actin antibody was purchased from Sigma-Aldrich. Rhodamine-conjugated sheep antidigoxigenin antibody was purchased from Roche.
Colony formation assay
200 spontaneously growing HeLa cells were plated into each well of a 6-well dish. 24 h later, CPT (Topogen) was added at various concentrations to the medium in the presence or absence of 200 µM NU1025 (EMD). Cells were washed with PBS 22 h later and cultured in 10% FBS-containing medium for 6 or 7 d. Colonies consisting of >10 cells were counted.
Biochemical detection of poly-ADP ribose
Cells were grown on a coverslip (Matsunami Glass) and incubated for 1 h at 37°C in 56 mM Hepes, pH 7.5, 28 mM NaCl, and 0.01% digitonin in the presence or absence of 200 µM NU1025, and 25 µM FITC-NAD+ (Trevigen) was added with or without 10 µM CPT (Topogen). After 15 min, cells were fixed with ice-cold methanol/acetone (1:1) for 10 min. Fixed cells were counterstained with DAPI. For immunoblotting, 105 cells were lysed in 10 µl of 1x SDS gel loading buffer, and the lysate was loaded onto a 12% SDS-PAGE gel, transferred to nitrocellulose membrane (Biodyne), and detected with the mouse anti–FITC-HRP (1:2,500; Jackson ImmunoResearch Laboratories). Chemiluminescent signals were detected by the ECL system (GE Healthcare).
Plasmid constructs, transfection, and immunodetection of BrdU-labeled DNA
Expression constructs for PARP-1–EYFP and Topo I–DsRed were provided by M.S. Satoh (Laval University, Quebec City, Quebec, Canada), and expression construct PCNA-RFP was provided by M.C. Cardoso (Technische Universität Darmstadt, Darmstadt, Germany). Cells were plated in a 24-well plate containing 13-mm round coverslips (Thermo Fisher Scientific) or in a 6-well plate containing square coverslips (Matsunami Glass). After 24 h, 100 ng/ml expression constructs PARP-1–EYFP and 100 ng/ml Topo I–DsRed were transfected with Lipofectamine and Plus Reagent (Invitrogen) according to the manufacturer's manual. However, expression construct PCNA-RFP (20 ng/ml to COS-7 cells and 8 ng/ml to HeLa cells) was transfected instead of expression construct Topo I–DsRed. The cells were cultured for 24 h, pulse labeled with 25 µM BrdU (Sigma-Aldrich) for 5 min, and fixed in 4% PFA (Nacalai Tesque) in PBS at 25°C for 10 min. After washing in PBS, the cells were permeabilized in 0.5% Triton X-100 in PBS at 25°C for 10 min, washed three times in PBS, and incubated in blocking buffer (0.3% BSA and 0.02% Tween 20 in PBS) at 25°C for 10 min and in 50 µl of detection buffer (0.15% BSA, 0.01% Tween 20, 30 mM Tris-HCl, pH 8.0, 300 µM MgCl2, 1 mM 2-mercaptoethanol, and 0.5x PBS) containing the nucleases and the antibodies (20 µg/ml DNase I [Sigma-Aldrich], 700 U/ml Exonuclease III [Toyobo], and mouse monoclonal antibodies directed against BrdU; 1:100 [MBL International]) at 37°C for 30 min. After washing three times in PBST (0.1% Tween 20 in PBS) for 5 min, the cells were incubated in Alexa Fluor 647–conjugated goat anti–mouse IgG (1:250; Invitrogen) at 37°C for 30 min, washed three times in PBST, and counterstained with DAPI (20 ng/ml in PBS; Nacalai Tesque).
RNAi
10 nM siRNAs (Thermo Fisher Scientific) were transfected to 105 HeLa cells using Lipofectamine RNAiMAX (Invitrogen) according to the manufacturer's instructions. Cells were cultured for 48 or 72 h before preparation of total cellular proteins or replication labeling. The nucleotide sequences of the siRNA target sites in PARP-1 were as follows: siRNA against PARP-1 (a), 5'-AAGCCTCCGCTCCTGAACAAT-3' (nt 2,401–2,421); siRNA against PARP-1 (b), 5'-AAGATAGAGCGTGAAGGCGAA-3' (nt 2,671–2,691; Kameoka et al., 2004); and siRNA against XRCC1, 5'-AACUCGACUCACUGUGCAGAA-3' (nt 151–171; Luo et al., 2004). Cell lysates were subjected on an 8 or 12% SDS-PAGE gel, transferred to nitrocellulose membrane, and detected with mouse anti–PARP-1 antibody (Trevigen) or β-actin (Sigma-Aldrich) as a control. After incubation in HRP-conjugated goat anti–mouse secondary antibody (MBL International), signals were enhanced by chemiluminescence with the ECL system (GE Healthcare). Cells grown on coverslips were fixed for immunofluorescence using the mouse anti–PARP-1 antibody and Alexa Fluor 488–conjugated goat anti–mouse IgG (Invitrogen).
Dynamic molecular combing and immunofluorescent detection
Genomic DNA was prepared and combed onto the silanated coverslips (Matsunami Glass) as described previously (Michalet et al., 1997) with modification (see following paragraph). 4 x 106 DT40 cells were pulse labeled for 20 min with 25 µM IdU (Sigma-Aldrich) and sequentially pulse labeled for 20 min with 250 µM CldU (Sigma-Aldrich) with or without 1 µM CPT. 2 x 106 m5S and HeLa cells were pulse labeled for 20 min with 100 µM IdU, washed with PBS twice, and pulse labeled for 20 min with 100 µM CldU with or without 1 µM CPT. NU1025 was pretreated for 2 h before IdU labeling. For preparation of genomic DNA, to remove the mitochondrial genome, the nuclei were extracted with buffer A (250 mM sucrose, 20 mM Hepes, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 1 mM EDTA, pH 8.0, 1 mM EGTA, pH 6.8, 1 mM DTT, and 0.1 mM PMSF) before resuspension into low melting point agarose (Roche). Combed DNA molecules were heat denatured in 50% formamide (Roche) and 2x SSC at 72°C for 12 min. For immunodetection of IdU- and CldU-labeled DNA, denatured DNA molecules were incubated with mouse anti-BrdU monoclonal antibody (1:5; BD) and rat anti-BrdU monoclonal antibody (1:25; Oxford Biotechnology) for 1 h at 37°C. After washing with PBS and 0.05% Tween 20 for 5 min three times, they were incubated with Alexa Fluor 555–conjugated goat anti–mouse IgG (1:500; Invitrogen) and Alexa Fluor 488–conjugated rabbit anti–rat IgG (1:500; Invitrogen) for 30 min at 37°C. All antibodies were diluted in blocking solution (1% [wt/vol] blocking reagent in PBS and 0.05% Tween 20; Roche). After washing with PBS and 0.05% Tween 20 for 5 min three times, coverslips were mounted in VECTASHIELD (Vector Laboratories). To estimate the extension of DNA molecules, coverslips were prepared with 
-DNA, and the DNA molecules were stained with 6.7 mM YOYO-1 (Invitrogen) at 25°C for 1 h. YOYO-1–stained DNA molecules measured 21 ± 0.9 µm. As the virus genome is 48.5 kbp, the extension of DNA molecules is 2.32 ± 0.11 kbp/µm.
Microscopy
Nuclei were examined using a microscope (Axioplan 2 MOT; Carl Zeiss, Inc.) with a 63x Plan Apochromat NA 1.4 objective lens equipped with a charge-coupled device camera (MicroMAX; Princeton Instruments). For deconvolution analysis of the nucleus, z-stack images were captured at a step of 0.2 µm and processed using MetaMorph software (version 6.1; MDS Analytical Technologies).
Online supplemental material
Fig. S1 shows the effect of PARP inhibitor on replication fork progression in mouse m5S cells. Fig. S2 shows replication fork kinetics in HeLa cells treated with XRCC1 siRNA. Fig. S3 shows replication fork kinetics in BRCA2tr/– DT40 cells. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200806068/DC1.
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Acknowledgments |
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This work was supported partly by a Grant-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology, the Japan Society for the Promotion of Science (grant to K. Okumura and K. Sugimura), and the Sankyo Foundation of Life Science (grant to K. Okumura). S.-i. Takebayashi is supported by the Nakayama Foundation for Human Science.
Submitted: 11 June 2008
Accepted: 22 November 2008
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References |
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