Article | Published:

Transcriptionally active chromatin recruits homologous recombination at DNA double-strand breaks

Nature Structural & Molecular Biology volume 21, pages 366374 (2014) | Download Citation


Although both homologous recombination (HR) and nonhomologous end joining can repair DNA double-strand breaks (DSBs), the mechanisms by which one of these pathways is chosen over the other remain unclear. Here we show that transcriptionally active chromatin is preferentially repaired by HR. Using chromatin immunoprecipitation–sequencing (ChIP-seq) to analyze repair of multiple DSBs induced throughout the human genome, we identify an HR-prone subset of DSBs that recruit the HR protein RAD51, undergo resection and rely on RAD51 for efficient repair. These DSBs are located in actively transcribed genes and are targeted to HR repair via the transcription elongation–associated mark trimethylated histone H3 K36. Concordantly, depletion of SETD2, the main H3 K36 trimethyltransferase, severely impedes HR at such DSBs. Our study thereby demonstrates a primary role in DSB repair of the chromatin context in which a break occurs.


Cells have developed HR and nonhomologous end joining (NHEJ) to repair highly toxic DNA DSBs1. In contrast to HR, which involves extensive DNA end resection and uses an intact copy of the damaged locus, NHEJ ligates broken DSB ends with no or limited processing. Although mostly faithful, both mechanisms can be sources of genomic instability. For example, HR can trigger amplification and depletion of repetitive sequences and can yield loss of heterozygosity when the homologous chromosome is used as the template rather than the sister chromatid. In contrast, NHEJ can induce point mutations and small deletions, especially when repair is performed by the error-prone backup NHEJ pathway (alt-NHEJ). Considering the wide range of DNA products that can arise from these different repair pathways, the choice between NHEJ and HR represents a central aspect of DSB repair.

Although cell-cycle phase contributes to this choice, these pathways coexist in S- and G2-phase cells2,3,4, thus implying that other factors participate in this decision. Several repair proteins can orient repair toward one pathway or the other. For example, critical roles have been assigned to the Ku heterodimer, whose binding inhibits resection and channels repair toward NHEJ5, to 53BP1–RIF1 (which promotes NHEJ at least in part via counteracting BRCA1 functions6,7,8,9,10,11,12) and to CtIP, MRE11 and Exo1 (which promote resection and thus HR13,14). However, how these factors are targeted to DSBs to achieve their functions is largely unknown.

An appealing hypothesis, supported by recent studies, is that the chromatin context in which a DSB occurs could contribute to this decision. For example, mono- and dimethylation of histone H4 K20 (refs. 15,16,17,18,19), as well as acetylation of H4 K16 by Tip60 (ref. 20), are involved in regulating the recruitment and anchoring of 53BP1, which is proposed to counteract HR21. Because these histone modifications are not evenly distributed throughout the genome, this raises the hypothesis that chromatin may have a role in the repair-pathway choice. In addition, recent studies have provided interesting insights regarding the differences in DSB repair between heterochromatin and euchromatin2,3,22.

Previous studies have benefited from the development of new techniques, going beyond classical damage-induction methods, including focused lasers and sequence-specific DSB-inducible cell systems (which greatly increased the resolution of molecular studies on DSBs by permitting the use of ChIP23,24). However, although these techniques helped to identify new chromatin players targeted to DSBs25,26 as well as their kinetics of recruitment, they hindered any investigation of the influence of preexisting chromatin structure on repair. Indeed, on the one hand, lasers induce DNA damage along tracks in the nucleus in a random fashion with regard to genomic position. On the other hand, the available sequence-specific DSB-inducible systems trigger either one single break (the HO, I-SceI and FokI systems24,27,28) or several DSBs that are mainly located in ribosomal DNA (the I-PpoI system23), thus precluding the comparison of repair events occurring at different genomic locations.

Recently, we developed stable human cell lines expressing a restriction enzyme (AsiSI) targeting an 8-bp recognition sequence fused to a modified estrogen receptor (ER) ligand-binding domain (henceforth called DIvA for DSB inducible via AsiSI)29,30,31. Treating such cells with 4-hydroxytamoxifen (4OHT) triggers nuclear localization of the AsiSI enzyme and the ensuing rapid induction of approximately 150 sequence-specific DSBs dispersed across the genome31. This system provides a unique opportunity to simultaneously study, at a molecular level, the repair events that transpire at many different DSBs located within various chromatin states.

Using ChIP-seq approaches, we found here that distinct DSBs induced across the genome are not necessarily repaired by the same pathway and that transcriptionally active trimethylated histone H3 K36 (H3K36me3)-enriched chromatin is preferentially repaired by homologous recombination, thereby pointing out a critical role of preexisting chromatin in addressing the DSB repair pathway at sites of breaks.


Differential binding of RAD51 and XRCC4 at distinct DSBs

To investigate how chromatin context might influence choice of DSB repair pathways, we performed chromatin immunoprecipitation followed by massively parallel sequencing in 4OHT-treated DIvA cells with antibodies to RAD51 and XRCC4, core components of the HR and NHEJ machineries, respectively. We also performed a control S139-phosphorylated histone H2AX (γH2AX) ChIP-seq to evaluate the efficiency of DSB induction at each AsiSI site, because γH2AX levels reflect the extent of DSB induction30. As expected, γH2AX showed a typical pattern, with wide spreading away from the DSBs (~1–2 Mb) and a clear drop at the break sites themselves, presumably reflecting reduced nucleosome occupancy (~1 kb) (Supplementary Fig. 1a,b). ChIP-seq analyses indicated clear recruitment of XRCC4 and RAD51 to AsiSI sites (Fig. 1). Notably, although XRCC4 localization was highly focused around the break point, RAD51 spread farther away (~2–4 kb), consistently with its role in binding single-stranded DNA (ssDNA) generated by DNA end resection (Fig. 1a).

Figure 1: A subset of AsiSI-induced DSBs recruits RAD51.
Figure 1

(a) ChIP-seq analyses in DIvA cells after 4OHT treatment (4 h), using anti-XRCC4 or anti-RAD51 antibodies. Averaged XRCC4 and RAD51 signals, over a 20-kb region flanking annotated AsiSI sites, are shown. (b) Profiles of XRCC4 and RAD51 around four selected AsiSI sites (indicated by arrows). Chr, chromosome. (c) Averaged XRCC4 and RAD51 signals over 20-kb windows and centered at the AsiSI site, shown for each category (RAD51-bound or RAD51-unbound subsets). (d) ChIP against XRCC4 and RAD51 in 4OHT-treated DIvA cells, analyzed by qPCR. The ratios between the signals observed for XRCC4 and RAD51 are shown for eight AsiSI-induced DSBs, either bound or not by RAD51 in ChIP-seq experiments (RAD51-bound DSBs are labeled with roman numerals, and RAD51-unbound DSBs are labeled with arabic numerals). Mean and s.e.m. (n = 4 technical replicates) from a representative experiment (out of 5 experiments) are shown.

A closer inspection of various AsiSI-induced DSBs revealed that some displayed readily detectable binding of both RAD51 and XRCC4 (Fig. 1b). By contrast, other sites displayed little or no discernible RAD51 accrual despite exhibiting robust γH2AX induction and XRCC4 binding (Fig. 1b and Supplementary Fig. 1c). We confirmed these findings by a second biological-replicate experiment (Supplementary Fig. 2a,b). We defined and further characterized two subsets (containing 20 DSBs each) of RAD51-bound DSBs and RAD51-unbound DSBs on the basis of respective enrichment of XRCC4 and RAD51 (RAD51/XRCC4 ChIP-seq signal ratio, Online Methods). The averaged profiles of XRCC4 and RAD51 on these DSB subsets validated our categorization process (Fig. 1c). To further confirm these data, we determined the relative enrichments of XRCC4 and RAD51 by ChIP followed by real-time quantitative PCR (qPCR) for certain DSBs in each subset. Because our ChIP-seq and ChIP-qPCR analyses revealed that XRCC4 binding was restricted to the DSB, whereas RAD51 was mainly found further away from the break (Fig. 1a,b and Supplementary Fig. 2c), we always analyzed subsequent XRCC4 and RAD51 ChIP by qPCR at 100 and 800 bp from the DSB, respectively. This analysis by ChIP-qPCR clearly differentiated the two categories, with RAD51-bound DSBs I, II, III, IV and V exhibiting high RAD51/XRCC4 ratios, which contrasted with the substantially lower RAD51/XRCC4 ratios displayed by DSBs 1, 2 and 3, which we categorized as RAD51 unbound (Fig. 1d).

To investigate the influence of the cell-cycle phase in RAD51 and XRCC4 recruitment at DSBs, we performed ChIP experiments in G1- or G2-synchronized DIvA cells after a double thymidine block (Fig. 2a). We found that RAD51 accumulated at DSBs much more strongly in G2 than in G1, whereas XRCC4 recruitment was similar in both cell-cycle phases, results consistent with the cell-cycle characteristics of HR (Fig. 2b,c). To exclude a potential bias due to unequal DSB induction at investigated AsiSI annotated sites across the cell cycle, we controlled AsiSI efficiency in synchronized cells, using our previously reported protocol30,32 based on the ligation of biotinylated double-stranded oligonucleotides, cohesive with AsiSI ends and subsequent streptavidin purification (cleavage assay; Online Methods). Importantly, AsiSI-mediated DSB induction was constant throughout the cell cycle (Supplementary Fig. 3a,c).

Figure 2: RAD51 recruitment at RAD51-bound DSBs occurs mainly in G2.
Figure 2

(a) Cell-cycle distributions as measured by fluorescence-activated cell sorting (FACS) in synchronized DIvA cells. (b) ChIP using XRCC4 (left) or RAD51 (right) antibodies in DIvA cells synchronized in G1 and G2. Enrichments in XRCC4 and RAD51 measured by qPCR at 80 bp and 800 bp from the DSB-I, respectively (from the RAD51-bound subset) are shown. Mean and s.e.m. (n = 4 technical replicates) of a representative experiment (out of three experiments) is shown. (c) RAD51/XRCC4 ratio obtained by ChIP-qPCR, calculated for various DSBs from the RAD51-unbound subset (labeled with arabic numerals, in blue) and from the RAD51-bound subset (labeled with roman numerals, in red) in both G1- and G2-synchronized cells. Mean and s.e.m. (n = 4 technical replicates) of a representative experiment (out of three experiments) are shown.

Therefore, our data together indicate that in G1 phase, XRCC4 recruitment dominated at all DSBs investigated and that in G2 RAD51 is recruited, in addition to XRCC4, but only at a subset of DSBs. This suggested that in G2 distinct DSBs induced throughout the genome are not necessarily repaired by the same pathway.

RAD51-bound DSBs exhibit HR features

To determine whether RAD51 binding correlated with 5′-to-3′ DSB resection, which is characteristic of HR-engaged DSBs, we probed for the presence of ssDNA at RAD51-bound or RAD51-unbound DSBs by using biotinylated double-stranded oligonucleotides with a specific 15-base 3′-end extension that allowed hybridization to the exposed ssDNA (Online Methods). The five RAD51-bound sites investigated indeed bore this hallmark of resection, in contrast with the three sites investigated that we had defined as RAD51 unbound (Fig. 3a). These differences were not due to variations in DSB induction efficiencies (Supplementary Fig. 3d).

Figure 3: RAD51-bound AsiSI-DSBs are resected and repaired by a RAD51-dependent pathway.
Figure 3

(a) ssDNA arising through resection, assessed at each of the eight previously analyzed DSBs. Pulldown efficiency measured by qPCR is shown as ratios between treated and untreated cells. Mean and s.e.m. (n = 3 technical replicates) of a representative experiment (out of three experiments) are shown. (b) Western blot analysis of AID-DIvA stably transfected cells induced by 4OHT for 4 h and subsequently treated (or not) with auxin for the indicated time. Uncropped images are shown in Supplementary Figure 8. (c) The presence of γH2AX foci, monitored by immunofluorescence in untreated cells, in 4OHT-induced cells (4 h) and in 4OHT-induced cells further incubated with auxin (2 h). (d) Cleavage assay in AID-DIvA cells, treated as indicated, followed by qPCR close to a cleaved AsiSI site. Normalized pulldown efficiencies from a representative experiment (out of six experiments) are shown (error bars, s.e.m.; n = 4 technical replicates). (e) Western blot analyses of AID-DIvA cells transfected with a control siRNA or siRNA against RAD51 (top) or against XRCC4 (bottom). Uncropped images are shown in Supplementary Figure 8. (f) Cleavage assay in AID-DIvA cells transfected with control siRNA or siRNA against RAD51 or against XRCC4. Analysis of immunoprecipitated DNA close to six DSBs is shown for RAD51-bound (top) and RAD51-unbound (bottom) subsets. The percentage of sites that remained broken for each DSB after the indicated times of auxin treatment is plotted. A representative experiment (out of three experiments) is shown.

To investigate whether the RAD51 binding that we observed was biologically relevant and reflected HR repair, we next asked whether DSB repair kinetics at such sites was dependent on RAD51 and/or XRCC4. The DIvA system does not permit ready study of the kinetics of DSB repair, because the enzyme regenerates DSBs at AsiSI sites even after they have been accurately religated. We therefore further refined our system by adding an auxin-inducible degron (AID) to the AsiSI-ER fusion enzyme, thus allowing us to remove it and stop AsiSI activity upon auxin addition33. The AID-DIvA cells behaved similarly to the DIvA cells with regard to kinetics of DSB induction (refs. 30,31 and Supplementary Fig. 4a,b). Our analyses revealed that auxin induced the efficient degradation of AID-AsiSI-ER, whose kinetics of loss correlated well with decreases in γH2AX levels (Fig. 3b). Auxin also triggered the rapid disappearance of γH2AX foci, which reflect the presence of DSBs (Fig. 3c and Supplementary Fig. 4c,d). We next analyzed the repair kinetics of an AsiSI-induced DSB by using the cleavage assay described above that uses adaptor ligation and qPCR to measure the presence of DSB ends (refs. 30,32, Supplementary Fig. 3 and Online Methods). This revealed that repair was nearly completed 1 h after auxin addition (Fig. 3d). We then compared repair kinetics for DSBs that we had designated as RAD51 bound or RAD51 unbound, upon short interfering RNA (siRNA)-mediated depletion of either XRCC4 or RAD51 (Fig. 3e). Strikingly, RAD51 depletion substantially slowed the repair kinetics of RAD51-bound DSBs (Fig. 3f and Supplementary Fig. 4e), whereas it had little or no effect on the repair of RAD51-unbound DSBs (Fig. 3f and Supplementary Fig. 4e). By contrast, XRCC4 depletion influenced the repair of RAD51-unbound DSBs but had little or no effect on the repair of RAD51-bound DSBs (Fig. 3f and Supplementary Fig. 4e).

Collectively, these data substantiated our ChIP data indicating that all DNA DSBs induced across the genome are not repaired equivalently and revealed that individual DSBs are indeed repaired by the pathway whose factor exhibited preferential binding at each site. DSBs defined as RAD51 bound behave as HR-prone DSBs because they undergo resection and rely on RAD51 for their effective repair. In contrast, other DSBs do not undergo detectable resection and rely on XRCC4 for repair.

RAD51-bound DSBs lie in transcriptionally active chromatin

The above findings strongly suggested that genomic or epigenomic features influence choice of DSB repair pathways in human cells. To identify such features, we compared our ChIP-seq data with publicly available epigenomic data sets (ENCODE project34; Online Methods), particularly focusing on histone marks generally associated with transcriptionally active or inactive regions. Interestingly, in comparison to our experimentally characterized set of RAD51-unbound DSB sites, the RAD51-bound DSBs that we identified were reproducibly enriched in chromatin marks associated with active transcription, such as H3K36me3 or H3 acetylated on K9 (Supplementary Fig. 5a). By contrast, repressive chromatin marks (such as H3 trimethylated on K9 or K27) were higher on RAD51-unbound sites than on RAD51-enriched sites (Supplementary Fig. 5a). These findings therefore highlighted a potential connection between a locus being transcribed and its propensity to engage in HR. To investigate any association between active transcription and HR repair, we profiled, by ChIP-seq in undamaged DIvA cells, the S2-phosphorylated C-terminal-domain form of RNA polymerase II (PolII-S2P) that is associated with transcriptional elongation. As expected, PolII-S2P spread along the transcribed units, thus reflecting active transcription (Supplementary Fig. 5b). Whereas almost all the DSBs from our two subsets were located either within or proximal to (<1,000 bp) a gene (Supplementary Table 1), the genes close to RAD51-bound DSBs exhibited considerably higher PolII-S2P levels than genes located near RAD51-unbound DSBs, either when taken individually or collectively (Fig. 4a,b). We also observed this tendency of RAD51-bound DSBs to be located in actively transcribed regions when taking all DSBs into account (Supplementary Fig. 5c). Furthermore, the two RAD51-bound AsiSI sites that turned out not to be in proximity to an annotated gene (Supplementary Table 1a) were actually located in PolII-S2P enriched regions (Supplementary Fig. 5d), thus implying that these DSBs in fact lie within transcribed loci. Collectively, the above data strongly suggested that active transcription is a feature that helps target a particular locus for HR repair.

Figure 4: RAD51-bound DSBs lie within transcribed units.
Figure 4

(a) PolII-S2P ChIP-seq in untreated DIvA cells. PolII-S2P enrichment is shown around the four AsiSI-induced DSBs presented in Figure 1b. Positions are in bp. (b) Average PolII-S2P enrichments around the transcription start site, calculated for genes associated with each DSB from the RAD51-bound and RAD51-unbound subsets. (c) XRCC4 and RAD51 ChIP in 4OHT-treated DIvA incubated with DRB or not. The RAD51/XRCC4 ratios for three RAD51-bound DSBs (DSB-I, DSB-II and DSB-III), one RAD51-unbound DSB (DSB-1) and two DSBs located far from any gene (DSB-5 and DSB-6) are shown. Mean and s.e.m. (n = 4 technical replicates) of a representative experiment (out of three experiments) are shown. (d) As in c, except that cells were treated with actinomycin D or not. (e) Cell-cycle distribution measured by staining with 5-ethynyl-2′-deoxyuridine and propidium iodide and FACS analysis, in untreated or DRB- or actinomycin D (actD)-treated DIvA cells, as indicated. Mean and s.e.m. of percentage of cells in each phase are shown (n = 3 independent experiments).

Transcription-dependent RAD51 recruitment

To test whether transcription was involved in RAD51 recruitment, we first inhibited transcription by using 5,6-dichloro-1-β-D-ribofuranosylbenzimidazole (DRB) or actinomycin D. A ChIP experiment performed against PolII-S2P confirmed that transcription was indeed impaired under these conditions (Supplementary Fig. 6a,b). We studied XRCC4 and RAD51 recruitment at selected DSBs by ChIP followed by qPCR. We included in our analysis two DSBs arbitrarily chosen from among the cleaved AsiSI sites (DSB-5 and DSB-6), located far from any genes. As expected, these DSBs behaved as RAD51-unbound DSBs, showing a low RAD51/XRCC4 ratio (Fig. 4c,d) with a low RAD51 recruitment (Supplementary Fig. 6c). Strikingly, transcription inhibition led to a strong decrease in the RAD51/XRCC4 ratio at RAD51-bound sites, whereas it induced far less pronounced changes at other AsiSI-induced DSBs studied (Fig. 4c,d). These changes in RAD51/XRCC4 ratio were due to a sharp decrease in RAD51 binding upon transcription inhibition (Supplementary Fig. 6c) while XRCC4 recruitment remained unchanged. Importantly, the effects of transcriptional inhibition on RAD51-bound sites did not reflect appreciable changes in cell-cycle distributions (Fig. 4e) and AsiSI cutting efficiencies in asynchronous cells (Supplementary Fig. 6d) and also across the cell cycle (Supplementary Fig. 6e), thus excluding a potential bias due to the influence of cell cycle on this observed impairment in RAD51 binding. Furthermore, transcription inhibition did not substantially alter the abundance of XRCC4 and RAD51 (data not shown).

To test the converse—that transcription activation could enhance RAD51 targeting—we used a transcription activator–like effector nuclease (TALEN)35 to generate a DSB within an inactive gene that can be easily turned on. We chose to use a GBP1-TALEN to induce a DSB in the interferon-γ (IFN-γ)-responsive gene GBP1, which under basal conditions showed very little enrichment of PolII-S2P in DIvA cells (Fig. 5a). Upon GBP1-TALEN transfection, a DSB was indeed induced within GBP1, as demonstrated by γH2AX induction detected by ChIP specifically at the vicinity of the DSB (Fig. 5b). Importantly, IFN-γ treatment induced GBP1 activation, as detected by both increased GBP1 mRNA levels (Fig. 5c) and increased H3K36me3 levels on the GBP1 gene body (Fig. 5d). We next analyzed RAD51 and XRCC4 recruitment in TALEN-transfected cells, either pretreated with IFN-γ (GBP1 thus being transcriptionally active) or not (GBP1 thus being transcriptionally silent). IFN-γ treatment substantially increased RAD51 binding to the TALEN-induced DSB without modifying XRCC4 levels (Fig. 5e). Collectively, these data therefore supported a model in which transcriptionally active loci display preferential RAD51 recruitment and thus HR repair.

Figure 5: Transcriptional activation of an unexpressed gene leads to repair by HR.
Figure 5

(a) PolII-S2P profile obtained in DIvA cells, shown on the GBP gene cluster, located on chromosome 1. A TALEN pair was ordered to specifically induce a DSB in the GBP1 gene, part of the GBP cluster located on chromosome 1. (b) γH2AX ChIP in DIvA cells transfected either with pCDNA3 or with the GBP1-TALEN plasmids. Enrichment measured by qPCR at 80 bp from the expected DSB or on a control genomic region is shown (error bars s.e.m.; n = 4 technical replicates of a representative experiment (out of two experiments). (c) Levels of GBP1 mRNA measured by RT-qPCR in DIvA cells treated (or not) with IFN-γ as indicated. cDNA levels were normalized to ribosomal protein P0 cDNA levels. (d) H3K36me3 ChIP in control or TALEN-transfected cells subjected (or not) to an IFN-γ treatment. H3K36me3 levels analyzed by qPCR on the GBP1 gene are shown as mean and s.e.m. (n = 4 technical replicates) of a representative experiment (out of two experiments). (e) XRCC4 and RAD51 ChIP in GBP1-TALEN–transfected cells, treated or not with IFN-γ. Enrichment of XRCC4 and RAD51 measured by qPCR at 80 bp and 800 bp from the TALEN-induced DSB, respectively, is shown as mean and s.e.m. (n = 4 technical replicates) of a representative experiment (out of three experiments).

RAD51 recruitment depends on an H3K36me3-LEDGF (p75) axis

A recent study identified LEDGF (p75) as a protein that facilitates binding of the HR- and resection-promoting factor CtIP on damaged chromatin, probably via a direct interaction between LEDGF and H3K36me3 (ref. 36). Because this histone mark is also associated with transcriptional elongation and is highly enriched on active genes37,38, we investigated its potential role in directing RAD51 to transcribed loci. First we performed a genome-wide mapping of H3K36me3 distribution in untreated DIvA cells (Fig. 6). As expected, H3K36me3 was strongly enriched on gene bodies, thus validating our ChIP-seq data (Fig. 6a). Notably, on average, the genes close to RAD51-bound DSBs exhibited much higher H3K36me3 levels than those located near RAD51-unbound DSBs (Fig. 6b). In addition, the regions surrounding RAD51-bound DSBs were significantly enriched in H3K36me3 as compared to loci that encompassed RAD51-unbound DSBs (Fig. 6c and examples in Fig. 6d). Thus, our H3K36me3 profiling in DIvA cells revealed that RAD51-bound DSBs are located in chromatin regions enriched for H3K36me3, results consistent with our previous finding indicating a preferential recruitment of RAD51 to transcriptionally active loci.

Figure 6: H3K36me3 enrichment on chromatin correlates with the use of HR.
Figure 6

(a) H3K36me3 ChIP-seq in untreated DIvA cells. Average H3K36me3 signals calculated around transcriptional start site (TSS) of all annotated genes (RefSeq, hg18) are shown. (b) Average H3K36me3 enrichments around the TSS, calculated for genes associated with each DSB from the RAD51-bound and RAD51-unbound subsets. (c) H3K36me3 average signal calculated over a 4-kb window centered on the DSB, for each category. Error bars, s.e.m. *P < 0.05 (Mann–Whitney test; Online Methods). (d) Profiles of H3K36me3 in undamaged DIvA cells at two RAD51-bound DSBs (DSB-I and DSB-II), one DSB identified in the RAD51-unbound subset (DSB-1) and a DSB located far from any gene (DSB-5).

In light of these observations, we investigated whether LEDGF and H3K36me3 participate in targeting RAD51 and HR repair to actively transcribed loci. Depletion of LEDGF by siRNA (Supplementary Fig. 7a) did not affect H3K36me3 levels (as detected by ChIP, Supplementary Fig. 7b) but led to a clear decrease in the RAD51/XRCC4 ratio at all RAD51-bound DSBs studied (Fig. 7a) due to a strong impairment of RAD51 recruitment (Supplementary Fig. 7c). Notably, LEDGF depletion had no or only minor effects on RAD51-unbound DSBs (Fig. 7a and Supplementary Fig. 7c). Such a dramatic effect of LEDGF depletion on RAD51 binding was not due to detectable changes in cell-cycle distributions (Fig. 7c) or HR protein levels (data not shown).

Figure 7: The H3K36me3-LEDGF axis targets RAD51 at DSBs induced in active genes.
Figure 7

(a) XRCC4 and RAD51 ChIP in 4OHT-treated DIvA cells transfected with control or LEDGF siRNAs. RAD51/XRCC4 ratios analyzed by RT-qPCR around six RAD51-bound DSBs (DSB-I to DSB-VI), four RAD51-unbound DSBs (DSB-1 to DSB-4) and two DSBs far from any gene (DSB-5 and DSB-6) are shown. *P < 0.05; **P < 0.01; ***P < 0.005; NS, nonsignificant (two-sided unpaired Student's t test). Error bars, s.e.m. (n = 4 technical replicates). A representative experiment (out of three experiments) is shown. (b) As in a, except that a siRNA against SETD2 was used. (c) Cell-cycle distribution measured by staining with 5-ethynyl-2′-deoxyuridine and propidium iodide and FACS analysis in DIvA cells transfected with siRNAs, as indicated. Mean and s.e.m. of percentage of cells in each phase are shown (n = 3 independent experiments). (d) U2OS cells, transfected with siRNAs as indicated, laser-microirradiated and stained for cyclin A, γH2AX (red) and RAD51 (green). Percentages of cyclin A–positive (CYCA+) cells that exhibit RAD51 recruitment at the laser line are shown. (e) Reporter assay on RG37-GFP-HR cells transfected with siRNAs as indicated and with a vector expressing I-SceI or not. GFP-positive cells were assayed by flow cytometry and signal expressed relative to the amount of GFP-positive cells in control siRNA–transfected cells. Mean and s.e.m. of independent experiments are shown (n = 4 independent experiments). *P < 0.05; **P < 0.01; ***P < 0.005; *****P < 0.001 (two-sided unpaired Student's t test). (f) H3K36me3 ChIP-seq in untreated and 4OHT-treated DIvA cells. Profiles obtained around the RAD51-bound DSB-I (indicated by an arrow) are shown. Positions are in bp. (g) H3K36me3 signals obtained by ChIP-seq, averaged around all cleaved AsiSI sites.

Similarly, we used siRNA to deplete SETD2, the main H3K36 trimethyltransferase38, to investigate the consequence of the loss of H3K36me3 on RAD51 binding. SETD2 siRNA led to a strong decrease of SETD2 mRNA levels (Supplementary Fig. 7e) and to an almost complete disappearance of H3K36me3 on chromatin (Supplementary Fig. 7f). Notably, SETD2 depletion triggered a strong reduction of RAD51/XRCC4 ratios at RAD51-bound DSBs (Fig. 7b) due to a dramatic decrease in RAD51 recruitment (Supplementary Fig. 7g). As for LEDGF siRNA, SETD2 depletion did not notably modify cell-cycle distribution (Fig. 7c) and HR-factor level (data not shown). Of note, we observed similar results when using a second siRNA for SETD2 and LEDGF (data not shown). Importantly, both LEDGF and SETD2 depletions led to decreased RAD51 binding without detectable reduction of the level of actively transcribing RNA polymerase II on nearby genes (Supplementary Fig. 7d,h). Taken together, these data therefore indicated a function for this histone mark in RAD51 binding independent from transcription per se.

To further confirm the role of H3K36me3 in RAD51 recruitment at DSBs, we analyzed the effect of SETD2 depletion on RAD51 binding at sites of damage generated by laser microirradiation. At those laser-induced DSBs, SETD2 siRNA also led to a clear decrease in RAD51 recruitment (Fig. 7d).

To investigate whether the loss of RAD51 binding observed upon H3K36me3 removal was associated with a HR repair defect, we next used the well-characterized DR-GFP reporter assay to analyze the consequences of SETD2 depletion on HR repair39. As expected, depletion of RAD51 by siRNA completely abolished the use of HR to repair the I-SceI–induced DSB, whereas 53BP1 siRNA led to an increased usage of HR. Strikingly, SETD2 depletion almost completely abolished HR repair, to an extent similar to that caused by RAD51 depletion (Fig. 7e).

Thus, altogether, our data indicate that SETD2 depletion not only impaired RAD51 binding at AsiSI DSBs and sites of laser-induced damage but also severely decreased HR repair at an I-SceI–induced DSB.

To assess the potential role for de novo H3K36me3 deposition occurring during repair, which could have explained the effect of SETD2 depletion on HR, we analyzed H3K36me3 distribution by ChIP-seq after 4OHT-dependent DSB induction. We did not detect marked induction of H3K36 trimethylation at any DSB sites tested upon 4OHT treatment (either taken individually (Fig. 7f) or collectively (Fig. 7g)). Therefore, we argue for a role of preexisting H3K36me3 in the targeting of RAD51 to DSBs.


In this study, by using a human cell line (DIvA) expressing a restriction enzyme fused to the ligand-binding domain of the estrogen receptor (AsiSI-ER) together with a ChIP-seq approach, we have found that distinct DSBs induced across the genome are not necessarily repaired by the same pathway. We identified an HR-prone subset of DSBs that recruit the HR protein RAD51, undergo resection and rely on RAD51 for efficient repair. Taken together, our data clearly indicate that the choice of repair pathway at AsiSI-induced DSBs depends, at least in part, on the chromatin context in which the break occurs. More specifically, it is influenced by the transcriptional status of the closest nearby gene, through at least one histone mark, H3K36me3, associated with transcriptional elongation. This chromatin mark fosters HR-mediated repair in transcribed genes, probably through being recognized by LEDGF, which then can promote CtIP recruitment to initiate resection at nearby DSB sites, thereby leading to RAD51 loading and repair by HR (Fig. 8).

Figure 8: Targeting of HR at transcriptionally active loci.
Figure 8

The H3K36me3 mark associated with transcriptional elongation would be recognized by LEDGF, which itself interacts with CtIP, thus bringing a resection-promoting factor to the site of the break. This would subsequently trigger RAD51 binding and HR repair on actively transcribed genes. DSBs occurring within inactive genes or intergenic regions would be unable to recruit a resection factor, thus leading to repair by NHEJ.

Repair of AsiSI-induced DSBs across the cell cycle

We have also addressed the influence of the cell cycle on AsiSI-induced DSB repair. In G1, as expected, we could not detect RAD51 binding while XRCC4 recruitment predominated at all investigated DSBs, which were located in either transcriptionally active or inactive chromatin. At this cell-cycle phase, although DSBs induced on silent chromatin are very likely to be repaired by NHEJ, the repair status of the DSBs induced on active chromatin still remains to be investigated. Indeed, even if both H3K36me3 and LEDGF are present in G1 (data not shown), the resection-promoting activity of CtIP is tightly controlled by phosphorylation events at the S-G2 transition40,41. Therefore, although it is still present in G1 phase, the H3K36me3-LEDGF pathway may not be able to trigger CtIP-dependent resection. DSBs induced in loci either rich or poor in H3K36me3 would thus be repaired by NHEJ.

However, our data also indicate that even though XRCC4 is recruited at DSBs associated with active loci, it may be inefficient in G1, because depletion of XRCC4 did not impair repair of these DSBs. One appealing hypothesis is that DSBs induced in active loci may be repaired by HR in the subsequent S phase. Of note, a recent study showed that some DSBs induced in late G1 are repaired by HR, as cells progress to S phase42. Such HR repair at transcribed loci may be initiated in G1; indeed, in G0-G1–synchronized cells I-PpoI–dependent DSBs induced in transcriptionally active genes, and not in intergenic regions, are found to undergo 'chromosome kissing' (a process believed to be linked to homology search)43. More investigation is clearly required to address the fate of these DSBs in G1, i.e., to state whether they are left unrepaired until S phase, are repaired by NHEJ or are repaired by an alternative pathway.

In G2, we could identify DSBs both able or unable to recruit RAD51, in agreement with previous reports, amounting to only about 15% of the DSBs repaired by HR in G2 (refs. 2,42). We show here that the chromatin structure that preexists at the site of the break is one of the major factors that influence the use of HR or NHEJ in G2 (described in next section).

Repair in heterochromatin versus in euchromatin

We found that the DSBs located in silent loci and intergenic regions were still unable to recruit RAD51 in G2, whereas DSBs induced in active chromatin, rich in H3K36me3, were channeled to HR repair.

This conclusion is seemingly at odds with studies reporting that heterochromatin (generally considered to be devoid of active genes) is repaired by a mechanism involving DNA synthesis by an Artemis-dependent HR pathway2,3,4,44, whereas euchromatin is mainly repaired by a DNA synthesis–independent pathway, probably NHEJ. Our work in DIvA cells did not allow us to investigate repair in heterochromatin, because AsiSI does not induce DSBs in these regions (probably because of its dense structure and/or its highly methylated status30). However, we note that, beyond its enrichment on active genes, H3K36me3 is also enriched in heterochromatin45 and that, although it is still poorly characterized, RNA production has been shown to be a feature of constitutive heterochromatin46,47,48. Consequently, as for DSBs induced in active genes, DNA breaks occurring within certain heterochromatic regions could also be targeted to HR repair through H3K36me3-dependent mechanisms. Concerning repair in euchromatin, we found that actively transcribed and H3K36me3-enriched loci are repaired by HR, whereas H3K36me3-depleted regions are repaired by NHEJ. This is in agreement with the studies from the Jeggo and Lobrich groups, because H3K36me3-enriched loci represent only a minor fraction (about 5%) of the genome49, the vast majority of euchromatin being composed of intergenic regions and silent genes.

Preexisting chromatin contributes to repair-pathway choice

Our study demonstrates a critical role of the preexisting chromatin context on the decision to use HR or NHEJ to repair a DSB induced in the human genome. Notably, changing the transcriptional status of a locus prior to DSB induction modified the pathway used for repair: a DSB induced in an inactive gene switched repair from a RAD51-independent to a RAD51-dependent pathway upon transcriptional activation (Fig. 5).

We identified a histone mark, H3K36me3, to be essential in the recruitment of RAD51 through LEDGF, a protein that possesses a PWWP domain. This specific module is involved in H3K36me3 recognition36,50,51 and was found to interact with CtIP36. Additional H3K36me3-binding factors are probably at work, because LEDGF depletion led to a less dramatic effect than did SETD2 depletion on RAD51 recruitment at DSBs. Importantly, LEDGF-CtIP anchoring occurs on H3K36me3 that preexists on chromatin. Indeed, our ChIP-seq data obtained in 4OHT-treated DIvA cells indicated that H3K36me3 is not increased after DSB induction, thus suggesting a role of preexisting H3K36me3 in RAD51 binding (Fig. 7f,g). Of note, a study recently conducted in the Humphrey laboratory also identified SETD2 as required for HR repair in mammalian cells (T.C. Humphrey, personal communication).

Interestingly, other studies have already suggested a role of preexisting chromatin in the choice of NHEJ versus HR. Indeed, mono- or dimethylated histone H4 K20 serves as an anchoring module for 53BP1, known to promote NHEJ. Although it is still controversial whether these marks are induced at DSBs, their involvement in 53BP1 stabilization at the site of damage is well documented15,16,17,18,19. In addition, a recent study revealed a role of acetylated K16 on histone H4 (H4K16ac), one of the histone marks associated with transcriptional activity, in counteracting 53BP1 binding to dimethylated H4 K20. Similarly to our findings, transcriptional activation of a locus before DSB induction led to an increased recruitment of BRCA1 and to a reduction of 53BP1 binding attributed to the higher H4K16ac levels triggered by transcription20. How exactly the H3K36me3-LEDGF-CtIP axis cooperates with H4K16ac-Tip60-BRCA1 to tightly regulate HR at DSBs will require further investigation, although our experiments using HR-DRGFP reporter assays suggest that both pathways act independently to promote resection (data not shown).

In conclusion, our study has revealed that DSBs do not all behave in the same manner across the genome and that these differences depend on whether the breaks occur in a transcribed locus. These data underline the extraordinary potential of chromatin in regulating genome stability: beyond the choice between classical NHEJ and HR, explored in this study, chromatin status could also conceivably control the use of alt-NHEJ, unequal HR, single-strand annealing and other less characterized repair pathways, in order to adapt the repair mechanism to the type and function of the locus to repair. This chromatin-dependent control could help to minimize the risks associated with repair events and could have a critical role in maintaining genome integrity.


AID-DIvA cell generation.

The AsiSI-ER fusion was cloned into the pAID1.1-N vector (BioROIS13). Cloning was performed with a modified Escherichia coli strain (AsiSI-met) provided by New England Biolabs. The AID-AsiSI-ER plasmid was transfected into U20S cells with the Cell Line Nucleofactor kit V (Amaxa), and selection was performed with 800 μg/mL G418.

Cell culture.

DIvA (AsiSI-ER-U20S), AID-DIvA (AID-AsiSI-ER-U20S) and RG37 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with antibiotics, 10% FCS (Invitrogen) and either 1 μg/mL puromycin (DIvA cells) or 800 μg/mL G418 (AID-DIvA cells) at 37 °C under a humidified atmosphere with 5% CO2. For AsiSI-dependent DSB induction, cells were treated with 300 nM 4OHT (Sigma, H7904) for 4 h. When indicated, 4OHT-treated cells were washed three times in prewarmed PBS and further incubated with 500 μg/mL auxin (Sigma, I5148). Pol II was inhibited with 5 μM actinomycin D (Sigma, A9415) for 10 h or 100 μM of DRB (Sigma, D1916) for 7 h. siRNA transfections were performed with the Cell Line Nucleofactor kit V (Amaxa) according to the manufacturer's instructions. Sequences for siRNAs are shown in Supplementary Table 2. For cell synchronization, cells were incubated with 2 mM thymidine for 18 h, released for 11 h and subjected to the second thymidine treatment for 18 h. G1 cells were collected after 15 h of release, and G2 cells were collected after 7 h.

TALEN-dependent DSB induction.

A TALEN pair targeting the sequence TCCTCACCTGATGAGAATGA gaatgaggttgaGGATTCAGCTGACTTTGTGA in the GBP1 gene was ordered at (H114918). 5.106 U20S cells were transiently transfected with 12.5 μg of each of the two TALENs by the calcium phosphate coprecipitation method for 24 h, and IFN-γ treatment (InVivogen, rhifn-g) was performed for 8 h at 50 ng/mL where indicated.

I-SceI–induced DSB HR assay.

105 RG37 cells, stably expressing an I-SceI–based HR-GFP substrate (kind gift of B. Lopez, Institut Gustave Roussy), were transfected with siRNA (10 nM final) (INTERFERin, PolyPlusTransfection, 409-10) and then transfected 24 h later with 1 μg of I-SceI vector (jetPEI, PolyPlusTransfection, 101-10). After 72 h, cells were collected, and GFP-positive cells were analyzed by flow cytometry.

Western blot.

Western blot analysis was performed with NuPAGE Tris-acetate 3–8% gels and reagents (Invitrogen) according to the manufacturer's indications. Briefly, cells were rinsed twice with ice-cold PBS and then lysed in the appropriate lysis buffer with sample reducing agent (Invitrogen). Liquid transfer of resolved proteins onto PVDF membranes (Invitrogen) was performed. After a 1-h block in 5% nonfat dry milk with 0.5% PBS-Tween, membranes were incubated overnight with the following primary antibodies: anti-HA (HA-11, Babco, 1:2,000), anti-γH2AX (Cell Signaling, 2577s, 1:500), anti-RAD51 (Santa Cruz, SC-8349, 1:200), anti-XRCC4 (Abcam, ab145, 1:1,000), anti-lamin A–C (Santa Cruz, H110, 1:500) and anti–α-tubulin (Sigma, DM1A, 1:100,000). Validations for these antibodies are available on the manufacturers' websites. Horseradish peroxidase–coupled secondary antibodies were from Sigma (anti-mouse, A2554, 1:10,000; anti-rabbit, A0545, 1:10,000), and the chemiluminescence Lumilight reagent was from Roche Diagnostic. Original images of blots used in this study can be found in Supplementary Figure 8.

Chromatin immunoprecipitation.

ChIP assays were carried out according to the protocol described in ref. 30 with the following modifications: 200 μg of chromatin was immunoprecipitated with 2 μg of anti-γH2AX (Epitomics, 2212-S), anti-XRCC4 (Abcam, ab145), anti-RAD51 (Santa Cruz, SC-8349) or no antibody (mock). These antibodies were validated in for ChIP in previous studies30,31. 50 μg of chromatin was used for the PolII-S2P immunoprecipitation (Abcam, ab5095, 2 μg), and 10 μg of chromatin was used for H3K36me3 ChIP (Abcam, ab9050, 2 μg). Both antibodies were validated in ChIP by the providers. Immunoprecipitated DNA and input DNA were analyzed in triplicate by RT-qPCR (primer sequences are provided in Supplementary Table 3). Because RAD51 recruitment often showed a significant decrease in the immediate vicinity of the DSB (Fig. 1a,b and Supplementary Fig. 2c), primer pairs used to measure XRCC4 enrichment were located at less than 100 bp from the DSB, whereas those used to assess RAD51 recruitment were located at 800 bp. IP efficiency was calculated as percentage of input DNA immunoprecipitated. Data are shown either as a ratio of RAD51/XRCC4 signals or as ChIP efficiency, as indicated. For ChIP-seq, sequencing libraries were prepared with 10 ng of purified DNA (average size 250–300 bp) and subjected to high-throughput sequencing (single read) by the Genomic Core facility (EMBL, Heidelberg, for XRCC4 and RAD51) or BGI (for γH2AX, PolII-S2P and H3K36me3), with a HiSeq 2000 sequencing system, or by the Genomics core facility at the Cancer Research Institute (CRI), Cambridge, UK, with an Illumina Genome Analyzer 2. After quality-filtering, the number of uniquely mapped sequencing reads (aligned to hg18) was counted over 200-bp windows.

Cleavage assay/resection assay.

The full procedure for the cleavage assay has been previously described30,32. Briefly, a biotinylated double-stranded oligonucleotide, ligatable with AsiSI sites, was ligated in vitro to genomic DNA after break induction. T4 ligase was heat-inactivated at 65 °C for 10 min, DNA was fragmented by EcoRI digestion at 37 °C for 2 h, and heat inactivation was then performed at 70 °C for 20 min. After a preclearing step, DNA was pulled down with streptavidin beads (Sigma) at 4 °C overnight and then washed five times in RIPA buffer and twice in TE. Beads were resuspended in 100 μL of water and digested with HindIII at 37 °C for 4 h. After phenol/chloroform purification and precipitation, DNA was resuspended in 100 μL water.

The resection assay is based on the cleavage efficiency assay, but a longer cohesive extremity (15 bp) allows specific pulldown of resected ends. Specific biotinylated double-stranded oligonucleotides were thus designed for each analyzed DSB (Supplementary Table 4). For both assays, precipitated DNA was quantified for each site by RT-qPCR with primers described in Supplementary Table 3.

Immunofluorescence and quantification.

Detailed methods for immunofluorescence have already been described in ref. 30. γH2AX foci were quantified with a Thermo Scientific Cellomics ArrayScan VTI HCS Reader. Laser microirradiation was essentially carried out as previously described13,52. Cells were plated on glass-bottomed dishes (Willco-Wells) and presensitized for 24 h with 10 μM BrdU before being exposed to a UV-A laser beam with a confocal inverted microscope (Olympus FluoView 1000), a 405-nm laser diode (6 mW, SIM scanner) and a 60× UPlanSApo/1.35 oil objective. Laser settings of 0.2 mW output (50 scans) restricted the generated DNA damage to the laser path in a presensitization-dependent manner without noticeable cytotoxicity. After a recovery time of 30–50 min, cells were stained with antibodies against RAD51 (Santa Cruz sc-8349), γH2AX (Millipore 05-636) and cyclin A (BD Biosciences 611268).

RAD51-bound and RAD51-unbound category design.

We previously reported that only a subset of annotated AsiSI sites over the genome are efficiently cleaved by the AsiSI-ER fusion protein in cells, mostly owing to their methylation status, and that efficient cleavage was always associated with both a wide γH2AX domain and a clear drop in close vicinity to the break30. A subset of the 100 most cleaved AsiSI sites in the genome was therefore determined on the basis of γH2AX enrichment over a 20-kb window and γH2AX depletion on 1,000 bp around the DSB. This subset was further sorted on the basis of the RAD51/XRCC4 ratio (a 1,000-bp window surrounding the DSB was taken into account for XRCC4, and a 4-kb window was used for RAD51) to identify two categories. The 20 best sites either in favor of XRCC4 or RAD51 were selected (Supplementary Table 1) and used to generate results shown in Figures 4 and 6 and Supplementary Figure 5a.

Average profiles around DSBs and TSSs.

To plot data with respect to DSBs, AsiSI site positions were retrieved from the human genome (hg18). ChIP-seq counts were retrieved for 20 kb around each of these DSBs and averaged with a 200-bp window. In order to plot data with respect to transcription start sites (TSS), for each gene associated with the studied DSBs (i.e., located not farther than 1,000 bp from the AsiSI site), transcript positions and orientations were obtained from the refFlat table from UCSC (hg18) at Unique genes were taken into account, and ChIP-seq counts were averaged with a 1,000-bp window.

Correlation with epigenomic features.

For each of the 20 DSBs of the two subsets, the averaged signal of various histone marks and proteins obtained by ChIP-seq and available from the ENCODE project ( was calculated over a 4,000-bp window centered on the DSBs. Data sets used for Supplementary Figure 5a were generated by the Broad Institute (Bernstein) or by the USC (Snyder) as indicated. Because some of these data are available only aligned against hg19, AsiSI positions from the two HR-prone and non–HR-prone subsets were converted to hg19 coordinates with the liftOver tool (UCSC).

Accession codes.

High-throughput sequencing data have been deposited with Array Express under accession number E-MTAB-1241.


Primary accessions



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We thank New England Biolabs for providing AsiSI genomic DNA. We thank B. Lopez (Institut de Cancérologie Gustave-Roussy) for RG37-HR I-SceI GFP cells. We thank V. Benes and the Solexa team at the European Molecular Biology Laboratory Genomic Core Facility, the Beijing Genomic Institute (BGI) and the Genomics core facility at the Cancer Research Institute (CRI) in Cambridge for high-throughput sequencing. We thank the Flow Cytometry platform from the Fédération de Biologie de Toulouse at the Laboratoire de Biologie Cellulaire et Moléculaire du Contrôle de la Prolifération (LBCMCP-FRBT). F.A. is supported by a grant from the Ligue Nationale Contre le Cancer (LNCC); P.C. is supported by a grant from the Association Contre le Cancer (ARC); and S.B. and E.G. are supported by grants from the Fondation pour la Recherche Médicale (FRM). Research in S.P.J.'s laboratory is supported by grants from Cancer Research UK (C6/A11226), the European Research Council and the European Community's Seventh Framework Program (DDResponse) and by core infrastructure funding from Cancer Research UK and the Wellcome Trust. K.M.M. was funded by a Wellcome Trust Project grant and C.K.S. by a Return-to-Europe Federation of European Biochemical Societies fellowship. S.P.J.'s salary is provided by the University of Cambridge and supplemented by Cancer Research UK. Funding to G.L. was provided by grants from the ARC, Agence Nationale pour la Recherche (ANR-09-JCJC-0138), Canceropole Grand Sud Ouest (GSO) and Research Innovation Therapeutic Cancerologie (RITC).

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Author notes

    • Kyle M Miller

    Present address: Department of Molecular Biosciences, Institute for Cellular and Molecular Biology, University of Texas at Austin, Austin, Texas, USA.


  1. Laboratoire de Biologie Cellulaire et Moléculaire du Contrôle de la Prolifération, Université de Toulouse, Université Paul Sabatier, Toulouse, France.

    • François Aymard
    • , Beatrix Bugler
    • , Emmanuelle Guillou
    • , Pierre Caron
    • , Sébastien Briois
    • , Virginie Daburon
    •  & Gaëlle Legube
  2. CNRS, Laboratoire de Biologie Cellulaire et Moléculaire du Contrôle de la Prolifération, Toulouse, France.

    • François Aymard
    • , Beatrix Bugler
    • , Emmanuelle Guillou
    • , Pierre Caron
    • , Sébastien Briois
    • , Virginie Daburon
    •  & Gaëlle Legube
  3. Gurdon Institute, University of Cambridge, Cambridge, UK.

    • Christine K Schmidt
    • , Kyle M Miller
    •  & Stephen P Jackson
  4. Department of Biochemistry, University of Cambridge, Cambridge, UK.

    • Christine K Schmidt
    •  & Stephen P Jackson
  5. Wellcome Trust Sanger Institute, Hinxton, Cambridge, UK.

    • Christine K Schmidt
    •  & Stephen P Jackson
  6. Bioinformatic Plateau I2MC, INSERM, University of Toulouse, Toulouse, France.

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F.A., P.C., E.G. and V.D. performed experiments. B.B. developed the AID-DIvA cell line. C.K.S. suggested and performed immunofluorescence studies on laser-induced damage. J.S.I. and S.B. performed bioinformatic analyses of the ChIP-seq data. K.M.M. performed XRCC4 ChIP library preparation and sequencing. G.L. conceived and analyzed experiments. F.A., J.S.I., K.M.M., S.P.J. and G.L. wrote the manuscript.

Competing interests

The authors declare no competing financial interests.

Corresponding author

Correspondence to Gaëlle Legube.

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