Factors determining DNA double-strand break repair pathway choice in G2 phase
2011; Springer Nature; Volume: 30; Issue: 6 Linguagem: Inglês
10.1038/emboj.2011.27
ISSN1460-2075
AutoresAtsushi Shibata, Sandro Conrad, Julie Birraux, Verena Geuting, Olivia Barton, M. Ismail, Andreas Kakarougkas, Katheryn Meek, G. Taucher-Scholz, Markus Löbrich, Penny A. Jeggo,
Tópico(s)Carcinogens and Genotoxicity Assessment
ResumoArticle11 February 2011free access Factors determining DNA double-strand break repair pathway choice in G2 phase Atsushi Shibata Atsushi Shibata Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Sandro Conrad Sandro Conrad Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Julie Birraux Julie Birraux Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Verena Geuting Verena Geuting Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Olivia Barton Olivia Barton Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Amani Ismail Amani Ismail Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Andreas Kakarougkas Andreas Kakarougkas Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Katheryn Meek Katheryn Meek Department of Pathobiology and Diagnostic Investigation, College of Veterinary Medicine, Michigan State University, East Lansing, MI, USA Search for more papers by this author Gisela Taucher-Scholz Gisela Taucher-Scholz Biophysics Department, GSI Helmholtzzentrum Schwerionenforschung GmbH, Darmstadt, Germany Search for more papers by this author Markus Löbrich Corresponding Author Markus Löbrich Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Penny A Jeggo Corresponding Author Penny A Jeggo Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Atsushi Shibata Atsushi Shibata Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Sandro Conrad Sandro Conrad Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Julie Birraux Julie Birraux Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Verena Geuting Verena Geuting Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Olivia Barton Olivia Barton Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Amani Ismail Amani Ismail Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Andreas Kakarougkas Andreas Kakarougkas Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Katheryn Meek Katheryn Meek Department of Pathobiology and Diagnostic Investigation, College of Veterinary Medicine, Michigan State University, East Lansing, MI, USA Search for more papers by this author Gisela Taucher-Scholz Gisela Taucher-Scholz Biophysics Department, GSI Helmholtzzentrum Schwerionenforschung GmbH, Darmstadt, Germany Search for more papers by this author Markus Löbrich Corresponding Author Markus Löbrich Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany Search for more papers by this author Penny A Jeggo Corresponding Author Penny A Jeggo Genome Damage and Stability Centre, University of Sussex, East Sussex, UK Search for more papers by this author Author Information Atsushi Shibata1, Sandro Conrad2, Julie Birraux1, Verena Geuting2, Olivia Barton2, Amani Ismail1, Andreas Kakarougkas1, Katheryn Meek3, Gisela Taucher-Scholz4, Markus Löbrich 2 and Penny A Jeggo 1 1Genome Damage and Stability Centre, University of Sussex, East Sussex, UK 2Radiation Biology and DNA Repair, Darmstadt University of Technology, Darmstadt, Germany 3Department of Pathobiology and Diagnostic Investigation, College of Veterinary Medicine, Michigan State University, East Lansing, MI, USA 4Biophysics Department, GSI Helmholtzzentrum Schwerionenforschung GmbH, Darmstadt, Germany *Corresponding authors: Darmstadt University of Technology, Radiation Biology and DNA Repair, Darmstadt 64287, Germany. Tel.: +49 6151 167460; Fax: +49 6151 167462; E-mail: [email protected] Damage and Stability Centre, University of Sussex, East Sussex BN1 9RQ, UK. Tel.: +44 127 367 8482; Fax: +44 127 367 8121; E-mail: [email protected] The EMBO Journal (2011)30:1079-1092https://doi.org/10.1038/emboj.2011.27 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info DNA non-homologous end joining (NHEJ) and homologous recombination (HR) function to repair DNA double-strand breaks (DSBs) in G2 phase with HR preferentially repairing heterochromatin-associated DSBs (HC-DSBs). Here, we examine the regulation of repair pathway usage at two-ended DSBs in G2. We identify the speed of DSB repair as a major component influencing repair pathway usage showing that DNA damage and chromatin complexity are factors influencing DSB repair rate and pathway choice. Loss of NHEJ proteins also slows DSB repair allowing increased resection. However, expression of an autophosphorylation-defective DNA-PKcs mutant, which binds DSBs but precludes the completion of NHEJ, dramatically reduces DSB end resection at all DSBs. In contrast, loss of HR does not impair repair by NHEJ although CtIP-dependent end resection precludes NHEJ usage. We propose that NHEJ initially attempts to repair DSBs and, if rapid rejoining does not ensue, then resection occurs promoting repair by HR. Finally, we identify novel roles for ATM in regulating DSB end resection; an indirect role in promoting KAP-1-dependent chromatin relaxation and a direct role in phosphorylating and activating CtIP. Introduction DNA double-strand breaks (DSBs) are significant biological lesions that cause cell death if unrepaired but promote translocations and genomic instability if misrepaired. Regulating how DSBs are processed to limit genomic instability is critical for cancer avoidance. DSBs can arise from exogenous DNA-damaging agents and via endogenously generated reactive oxygen species, or following metabolic processes such as replication, V(D)J recombination and meiosis. These processes can generate DSBs directly, for example following exposure to ionising radiation (IR) or indirectly for example following replication fork collapse. DSBs can be structurally distinct depending on their origin; for example, one-ended DSBs arise at replication forks, IR generates two-ended DSBs and the DSBs generated during V(D)J recombination are four ended and held in a synaptonemal complex. DSBs are repaired by two major pathways; DNA non-homologous end joining (NHEJ) or homologous recombination (HR). NHEJ repairs DSBs in all cell-cycle phases and represents the major pathway in G1, while HR functions in S/G2. HR also has a significant role at the replication fork in promoting replication fork recovery in a manner that does not necessitate DSB formation (Helleday et al, 2007). NHEJ involves the binding of the heterodimeric Ku protein to double-stranded DNA ends, recruitment of the DNA-dependent protein kinase catalytic subunit (DNA-PKcs) to generate the DNA-PK holoenzyme and DNA-PK kinase activation (Lieber, 2010). The assembled DNA-PK complex on the DNA end then helps to recruit a complex involving DNA ligase IV, XRCC4 and the less tightly associated XLF/Cernunnos protein, which effects the rejoining step (Lieber, 2010). HR is a more complex process involving 5′–3′ end resection to generate a 3′ single-stranded (ss)DNA overhang (Wyman and Kanaar, 2006). The ssDNA is rapidly bound by RPA, which is subsequently displaced by Rad51 in a process that involves BRCA2. Invasion of a homologous sequence to generate a Holliday junction and heteroduplex DNA then follows. Subsequent steps involve branch migration, fill-in of the ssDNA regions and Holliday junction resolution. In addition to these core DSB repair processes, recent studies have shown that ∼15–20% of IR-induced DSBs require ATM, the nuclease Artemis, the MRN complex, γH2AX, MDC1, RNF8, RNF168 and 53BP1 for repair (Riballo et al, 2004; Noon et al, 2010). The DSBs whose repair requires these additional proteins are those located at or in close proximity to heterochromatin (HC) and the process requires ATM-dependent phosphorylation of Kruppel-associated box-associated protein-1 (KAP-1), an HC-building protein (Goodarzi et al, 2008). Since HC-DSBs appear to be repaired with slow kinetics in control cells, the findings suggest that HC is a barrier to DSB repair that ATM relieves via KAP-1 phosphorylation. Recent studies have shown that in G2 phase, the slowly repaired DSBs are ATM-Artemis-Rad51/BRCA2 dependent, showing that the process represents HR (Beucher et al, 2009). A current critical question is what regulates the choice of DSB repair pathway usage. Studies have shown that the level of several critical HR proteins increases from S to G2 phase (e.g. Rad51, Rad52 and BRCA1) and that steps of HR are activated by CDKs (Shrivastav et al, 2008). Most significantly, CDK phosphorylation upregulates CtIP-dependent resection (Huertas et al, 2008). In yeast and chicken DT40 cells, the cell-cycle-dependent activation of CDK rigorously promotes the switch from NHEJ to HR during S/G2 phase (Aylon et al, 2004; Sonoda et al, 2006). However, in mammalian cells, recent studies have shown that, even though HR has the capacity to function in G2 phase, NHEJ represents the major DSB repair pathway in G2 as in G1 with ∼75–85% of IR-induced DSBs undergoing repair by NHEJ (Beucher et al, 2009). Thus, even though HR is activated and available in G2 phase, additional factors must influence which pathway is utilised. In this study, we examine factors that regulate the choice between usage of HR and NHEJ. Since it is likely that the pathway chosen will differ depending upon whether the DSBs are one, two or four ended (i.e. their origin), we have focused our study on pathway usage at IR-induced two-ended DSBs in G2 phase. Moreover, chromatin superstructure may be distinct in S phase. Guided by the finding that in G2 phase, NHEJ rejoins the majority of DSBs, while HR predominantly rejoins the slowly repaired HC-DSBs, we examined whether the speed of DSB repair influences pathway choice. We show that both DNA damage complexity and chromatin complexity influence the magnitude of DSB end resection. Our findings suggest a model whereby NHEJ makes the first attempt to repair DSBs but allows access to the resection machinery when rejoining does not rapidly ensue. Results DNA end complexity determines the speed of DSB repair and the level of DSB end resection in G2 phase Our previous findings showed that HR preferentially repairs HC-DSBs in G2 phase (Beucher et al, 2009). Since HC influences the speed of DSB repair, we examined the impact of DSB repair kinetics on repair pathway usage in G2 phase (Goodarzi et al, 2008). Previous findings have suggested that the complexity of DNA damage is another parameter that influences DSB repair kinetics (Ritter and Durante, 2010). We compared the kinetics of DSB repair and the magnitude of end resection in G2 phase following exposure to high linear energy transfer (LET) carbon ions, X-rays and etoposide (Etp), a topoisomerase II inhibitor. Carbon ions induce highly complex DSBs compared with X-rays (Hada and Georgakilas, 2008; Tobias et al, 2010), while we have previously observed that Etp-induced DSBs are repaired with fast kinetics (Riballo et al, 2004). DSB repair was examined by γH2AX foci enumeration in unsynchronised cycling cells since cell-cycle synchronisation methods tested led to genomic damage. G2 phase cells were identified using CENP-F. Aphidicolin (APH) was added to prevent irradiated S phase cells progressing into G2 phase during analysis. S phase cells, identified by marked pan-nuclear γH2AX staining, were excluded from analysis. G1 phase cells represented those remaining (see Supplementary Figure S1 for details and Beucher et al (2009) for additional control experiments). DSBs induced by carbon ions were repaired with substantially slower kinetics compared with X-ray-induced DSBs in G1 consistent with the notion that damage complexity influences DSB repair kinetics (Figure 1A). We also observed slower repair of carbon-ion-induced DSBs in G2 compared with X-ray-induced DSBs (Figure 1B). In contrast, Etp-induced DSBs are rapidly repaired in G1 and G2 cells (Figure 1A and B). Thus, the magnitude of damage complexity correlates with DSB repair kinetics for these three agents in G1 and G2 phase. We next examined HR-processing steps following exposure to these agents by monitoring the percentage of induced DSBs that undergo resection or load Rad51 in G2 phase (Figure 1C). We assess resection or Rad51 loading by monitoring RPA or Rad51 foci numbers from 2 h after treatment and divide this by number of induced DSBs assessed from the γH2AX foci numbers at 15 min after treatment. We have previously observed that by 2 h post-IR, RPA/Rad51 foci predominantly (>90%) overlap with γH2AX and 53BP1 foci (see also Supplementary Figure S2) (Beucher et al, 2009). Surprisingly, RPA foci form at nearly all carbon-ion-induced DSBs in contrast to X-ray-induced DSBs, when only 20–30% undergo resection consistent with previous findings (Beucher et al, 2009). In contrast, only a low percentage of Etp-induced DSBs undergo resection. To exclude the possibility that end resection might be underestimated by ongoing HR repair, we enumerated RPA foci in BRCA2-defective cells, which form RPA foci but fail to load Rad51. Consistent with our data after X-rays (Beucher et al, 2009), while BRCA2-defective cells show sustained RPA foci after carbon ions, the number of foci present at 2 h, which we consider represents the maximum level, was similar in control and BRCA2-defective cells (Figure 1D; see Supplementary Figure S3A for representative images). Thus, the data for control cells in Figure 1C is a good representation of the maximum level of resection. To consolidate that RPA foci formation represent DSB repair by HR, we enumerated the rate of γH2AX foci loss in BRCA2-defective cells. Approximately 90% of γH2AX persist in BRCA2 cells 24 h after carbon-ion irradiation (Figure 1E). Although the high percentage of DSBs remaining unrepaired in control cells by 24 h precludes a full assessment of the contribution of HR, the finding nonetheless demonstrates a significant role for HR. DSB repair kinetics in BRCA2-deficient cells after X-rays is similar to our previous findings, suggesting that HR represents the slow DSB repair component repairing 20–30% of induced DSBs. In stark contrast, DSB repair kinetics in control and BRCA2-defective cells were similar following exposure to Etp. We further confirmed this result in mouse embryonic fibroblasts (MEFs), which show slightly faster repair kinetics than human cells. Rad54-deficient MEFs exhibit a substantial DSB repair defect and substantial persistent RPA foci (Figure 1F; DSB repair kinetics by γH2AX foci is shown in Supplementary Figure S3B). Taken together, these results provide evidence that the speed of DSB repair correlates with the level of DSB end resection. Figure 1.DSB end complexity determines the speed of DSB repair and the level of DSB end resection in G2 phase. (A, B) Positive correlation between DNA damage complexity and repair kinetics. The kinetics of DSB repair following exposure to 2 Gy carbon ions, 2 Gy X-rays and 20 (A) or 5 (B) μM Etp for 15 min, was monitored by enumerating γH2AX foci in HSF1 primary cells in G1 (A) or G2 (B) cells. The percent of γH2AX foci is normalised to γH2AX foci numbers at 15 min after damage (representing the number of DSBs induced). The dose of Etp chosen produced a similar number of DSBs to that induced by 2 Gy X-rays. Irradiated G1 and G2 cells were identified as CENP-F negative and positive, respectively. S phase cells identified by pan-nuclear γH2AX staining were excluded from analysis (Supplementary Figure S1). (C) The magnitude of resection correlates with the speed of DSB repair. RPA or Rad51 foci numbers were enumerated following exposure to 2 Gy carbon ions, 2 Gy X-rays and 5 μM Etp in HSF1 primary cells. The number of RPA or Rad51 foci at the times indicated was normalised to γH2AX foci numbers at 15 min after damage. RPA and Rad51 foci numbers were similar (data not shown). The actual numbers of γH2AX and RPA–Rad51 foci scored per cell in G2 phase are given in Supplementary Figure S3C and D. We have previously undertaken kinetic analyses of RPA foci formation and shown maximal numbers 2 h post-IR (Beucher et al, 2009). Here, we observe a similar finding for Rad51 foci formation (Supplementary Figure S2C). Further discussion of this analysis is given in Supplementary Figure S2 legend. (D) Total DSB end resection events in HSF1 (WT) and HSC62 (BRCA2) cells. Following exposure to 1 or 2 Gy carbon ions, the number of RPA foci was scored in G2 cells. (E) The contribution of BRCA2-dependent HR to G2 DSB repair following exposure to 2 Gy carbon ions, 2 Gy X-rays and 5 μM Etp. Approximately 90% of DSBs remained in HSC62 (BRCA2) cells at 24 h after carbon ions compared with ∼30% of X-ray-induced DSBs. There is no detectable DSB repair defect in BRCA2-deficient cells following exposure to Etp. (F) More than ∼90% of RPA foci persist in Rad54−/− MEFs up to 24 h after carbon-ion irradiation. The number of RPA foci was enumerated after 2 Gy carbon ions. G2 cells were identified as speckled p-histoneH3 Ser10 staining. Error bars represent the s.e.m. of three experiments. Download figure Download PowerPoint Slowly repaired Etp-induced DSBs that localise to HC undergo resection We reproducibly observed a significant number of Etp-induced Rad51 foci (∼10% of γH2AX foci at 15 min) in G2 cells (Figure 1C). To consolidate the notion that Etp induces a small percentage of Rad51 foci formation, we exposed cells to increasing Etp concentrations and enumerated Rad51 foci at varying times after exposure (Figure 2A). These results confirmed that Rad51 foci form in a dose-dependent manner after Etp treatment at a low percentage of DSBs. Since topoisomerase II introduces a uniform type of DSB end with a 4-bp 5′-overhang (Spitzner et al, 1995), we speculated that there must be a unique aspect that promotes DSB end resection at a subset of these DSBs. Given our finding that after X- or γ-rays, the slowly repaired HC-DSBs (∼15–25% of induced DSB) are preferentially repaired by HR in G2 phase, we examined whether the subset of Etp-induced DSBs that undergo resection might localise to HC regions. We used a high concentration of Etp to examine the kinetics of DSB repair in G1 cells and, importantly, observed biphasic kinetics (Figure 2B). (N.B. Etp treatment causes >10-fold more γH2AX foci in G2 than in G1, necessitating the use of higher concentrations to investigate G1 cells; G0/G1 cells were used to allow analysis up to 24 h after treatment). The fraction of slowly repaired DSBs was substantially lower than that induced by X-rays (∼90% of DSBs were repaired with the fast kinetics (t1/2=2.2 h); ∼10% were repaired more slowly (t1/2=12.7 h)) (Figure 2B; Supplementary Table S1). Moreover, there was a correlation between the fraction of slowly repaired DSBs and those harbouring RPA/Rad51 foci consistent with our proposed causal relationship (Figures 1C and 2B). To examine whether these might indeed represent HC-DSBs, we examined their co-localisation with pS824-KAP-1, a heterochromatic marker (Noon et al, 2010). Approximately 8–24 h following IR exposure, pS824-KAP-1 foci form uniquely at HC-DSBs in an ATM-dependent manner (Noon et al, 2010). We also confirmed that pKAP-1 foci do not form at all persistent DSBs by examining their formation in DSB repair defective, XLF cells (Supplementary Figure S4A). To facilitate quantification, we treated cells with a high concentration of Etp (150 μM) and observed ∼8 γH2AX foci and 6.9 pKAP-1 foci with strong co-localisation at 8 h, when the fast DSB repair process is completed (Figure 2C). The high number of DSBs present at 30 min after 150 μM Etp precluded an accurate estimation of DSB induction. However, exposure to 50 μM Etp gave ∼48 DSBs at 30 min, from which we estimated an induction level of 150 DSBs following 150 μM Etp assuming a dose-linear induction rate. Thus, a concentration of Etp that induced 150 DSBs yielded ∼7 DSBs, associating with pKAP-1 at 8 h (HC-DSBs); in contrast, 2 Gy IR induces ∼50 DSBs and ∼5 pKAP-1 foci at 8 h (Supplementary Table S1). Thus, the slow DSB repair component after Etp treatment is approximately three-fold lower compared with X-rays, correlating with the lower frequency of resection after Etp (Figures 1C and 2C). We further consolidated the notion that HC-DSBs form at a lower frequency after Etp compared with γ-rays using NIH3T3 cells, which have readily visualised dense DAPI regions, which represent HC regions (Supplementary Figure S4B). To substantiate that Etp generates HC-DSBs, we examined the requirement for ATM in DSB repair after Etp with or without KAP-1 siRNA, a treatment that has previously been shown to relieve the requirement for ATM for DSB repair (Goodarzi et al, 2008). We observed a small fraction of unrepaired DSBs following ATMi addition at 24 h after high-dose Etp treatment; further, the requirement for ATM could be relieved by KAP-1 siRNA (Figure 2D). Finally, we observed that RPA, Rad51 and γH2AX foci co-localise with pKAP-1 foci in G2 cells at 8 h after Etp treatment (Figure 2E). Collectively, these findings strongly suggest that Etp-induced DSBs that undergo end resection localise to HC regions as observed for IR-induced DSBs. Taken together, we suggest that two factors can influence the speed of DSB repair, DNA damage complexity and chromatin complexity, and that slowly repaired DSBs as a consequence of either damage or chromatin complexity preferentially undergo DSB end resection. Figure 2.Slowly repaired etoposide-induced DSBs localise to heterochromatin and undergo resection. (A) Rad51 foci arise following exposure to Etp for 30 min in G2 cells in a dose-dependent manner. Cells were exposed to increasing doses of Etp and Rad51 foci enumerated in HSF1 primary cells. The number of Rad51 foci in untreated cells ( 60–70% of induced DSBs to undergo resection, demonstrating that HR can occur at euchromatic (EC)-DSBs. Further, the results clearly show that loss of DNA-PKcs (which does not impact upon Ku80 protein stability or DNA-binding capacity) results in substantially increased DSB end resection (Drouet et al, 2005). We consolidated this finding using a FACS-based approach involving the detection of chromatin-bound RPA in G2 cells using α-RPA following detergent extraction. Consistent with our foci analysis, we observed increased RPA retention in G2 phase cells following Ku80+DNA-PKcs siRNA after IR (Figure 3C). Additionally, we observed increased IR-induced SCEs arising from G2 phase cells in Ku80+DNA-PKcs double-knockdown cells (Figure 3D). Thus, loss of either Ku or DNA-PKcs can enhance resection, demonstrating that the DNA-PK holoenzyme (Ku+DNA-PKcs) functions as a complex to ensure the appropriate regulation of resection at DNA ends. Figure 3.DNA-PK competes with DSB end resection. (A) Ku80 siRNA significantly increases RPA foci numbers post-IR. A549 cells were exposed to 1 Gy X-rays and RPA foci were enumerated as indicated either with or without BRCA2 siRNA. G2 cells were identified with CENP-F. Knockdown efficiency and typical images are shown in Supplementary Figure S5A. (B) Ku80 siRNA, DNA-PKcs siRNA or combined siRNA causes increased DSB end resection in A549 G2 cells. RPA and 53BP1 foci were enumerated after 1 Gy X-rays. Solid bars represent the number of RPA foci, and solid+hashed bars represent the total number of 53BP1 foci. The knockdown efficiencies are shown in the right panel. (C) Loss of Ku80 and DNA-PKcs increased IR-induced RPA retention in G2. RPA retention in G2 cells was analysed using α-RPA antibody after detergent extraction. Chromatin-associated RPA was detected as an Alex488 signal following FACS. APH alone does not induce detectable RPA signal by FACS (data not shown), although some level of RPA signal was detected by IF (Supplementary Figure S2). Percent of RPA positive was shown in the right panel. Error bars represent two independent experiments. (D) Increased IR-induced SCEs in Ku80 and DNA-PKcs double-knockdown G2 cells. IR-induced SCEs in G2 phase were analysed following 2 Gy X-rays. (E) A significant reduction of IR-induced BrdU foci formation in CHO cells expressing DNA-PKcs ABCDE S>A. In all, 20 μM BrdU was added 24 h before IR. Cells were extracted with 0.2% Triton for 1 min at 2 h after 2 Gy and stained with α-BrdU antibody without denaturation. (F, G) CHO cells expressing DNA-PKcs ABCDE S>A fail to form Rad51 foci at 1 h after 2 Gy X-rays in contrast to control cells and cells expressing the phosphorylation mimic, DNA-PKcs ABCDE S>D. G2 cells were identified by DAPI intensity using ImageJ. Similar results were obtained examining RPA foci (data not shown). DNA-PK expression levels in the V3 strains are shown in Supplementary Figure S5B. Further characterisation of the strains has been described previously (Chan and Lees-Miller, 1996; Cui et al, 2005; Meek et al, 2007). Download figure Download PowerPoint The finding that there is increased DSB end resection in the presence of Ku but absence of DNA-PKcs implies that Ku can be removed from or vacates the DSB end to allow resection when NHEJ does not progress. To gain insight into this, we examined HR processing in a DNA-PKcs-defective cell line, V3, and derivatives expressing wild type (WT) DNA-PKcs, and DNA-PKcs mutants with alanine or aspartic acid substitutions at all six phosphorylation s
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