Rad51 replication fork recruitment is required for DNA damage tolerance
2013; Springer Nature; Volume: 32; Issue: 9 Linguagem: Inglês
10.1038/emboj.2013.73
ISSN1460-2075
AutoresRomán González‐Prieto, Ana M. Muñoz-Cabello, María J. Cabello-Lobato, Félix Prado,
Tópico(s)Plant Genetic and Mutation Studies
ResumoArticle5 April 2013free access Rad51 replication fork recruitment is required for DNA damage tolerance Román González-Prieto Román González-Prieto Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author Ana M Muñoz-Cabello Ana M Muñoz-Cabello Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author María J Cabello-Lobato María J Cabello-Lobato Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author Félix Prado Corresponding Author Félix Prado Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author Román González-Prieto Román González-Prieto Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author Ana M Muñoz-Cabello Ana M Muñoz-Cabello Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author María J Cabello-Lobato María J Cabello-Lobato Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author Félix Prado Corresponding Author Félix Prado Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain Search for more papers by this author Author Information Román González-Prieto1, Ana M Muñoz-Cabello1, María J Cabello-Lobato1 and Félix Prado 1 1Departamento de Biología Molecular, Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Consejo Superior de Investigaciones Científicas (CSIC), Seville, Spain *Corresponding author. Departamento de Biología Molecular, CABIMER-CSIC, Americo Vespucio s/n, Seville 41092, Spain. Tel.:+34 954468210; Fax:+34 954461664; E-mail: [email protected] The EMBO Journal (2013)32:1307-1321https://doi.org/10.1038/emboj.2013.73 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 Homologous recombination (HR) is essential for genome integrity. Recombination proteins participate in tolerating DNA lesions that interfere with DNA replication, but can also generate toxic recombination intermediates and genetic instability when they are not properly regulated. Here, we have studied the role of the recombination proteins Rad51 and Rad52 at replication forks and replicative DNA lesions. We show that Rad52 loads Rad51 onto unperturbed replication forks, where they facilitate replication of alkylated DNA by non-repair functions. The recruitment of Rad52 and Rad51 to chromatin during DNA replication is a prerequisite for the repair of the non-DSB DNA lesions, presumably single-stranded DNA gaps, which are generated during the replication of alkylated DNA. We also show that the repair of these lesions requires CDK1 and is not coupled to the fork but rather restricted to G2/M by the replicative checkpoint. We propose a new scenario for HR where Rad52 and Rad51 are recruited to the fork to promote DNA damage tolerance by distinct and cell cycle-regulated replicative and repair functions. Introduction In every cell cycle, DNA accumulates lesions that impair the advance of the replication forks. Eventually, this leads to an accumulation of single-stranded DNA (ssDNA), which activates a number of mechanisms aimed at ensuring that DNA replication passes through DNA lesions and repairing the gaps. Defects in this response cause replication fork stalling and genetic instability in yeast (Vázquez et al, 2008; Putnam et al, 2010) and are associated with cancer in humans (Moynahan and Jasin, 2010). This DNA damage tolerance (DDT) response relies in error-prone translesion synthesis (TLS) and error-free template switch (TS) mechanisms. TLS fills the gap by extending the 3′-end past the damaged template, using specialized DNA polymerases that are able to incorporate a nucleotide opposite the lesion, while TS uses the information of the sister chromatid to bypass damage (Friedberg, 2005). A crucial protein in this decision is the DNA polymerase processivity factor PCNA, which functions as a platform for factors involved in replication, repair, and chromatin assembly (Moldovan et al, 2007). In response to DNA damage, PCNA is ubiquitinated at lysine 164 by the Rad6/Rad18 ubiquitin ligase complex (Hoege et al, 2002), and this modification serves as a target for recruiting TLS polymerases (Lehmann et al, 2007). Alternatively, the ubiquitin residue at lysine 164 can be extended with a K63-linked polyubiquitin chain by the Ubc13/Mms2/Rad5 ubiquitin ligase complex to promote TS (Hoege et al, 2002). Thus, the choice between TLS and TS mechanisms determines whether the repair is mutagenic or not. Consequently, DDT is an essential process for cell-cycle progression, genome integrity, and cancer avoidance. Homologous recombination (HR) is also necessary for gap filling during DDT (Prakash, 1981; Friedberg, 2005; Heyer et al, 2010). The recombination proteins Rad52 and Rad51 are required for replication fork progression through alkylated DNA (Vázquez et al, 2008; Alabert et al, 2009), and in their absence cells accumulate ssDNA gaps (Lopes et al, 2006; Hashimoto et al, 2010). Since HR uses the genetic information of an intact molecule to repair a DNA break, it might provide the enzymatic activities required for TS. Accordingly, the RAD6 and RAD52 epistasis group of proteins cooperate through a mechanism that forms a sister-chromatid junction (SCJ), which is then further resolved by the helicase/topoisomerase Sgs1/Top3 complex (Liberi et al, 2005; Branzei et al, 2008; Minca and Kowalski, 2010). The requirement for recombination proteins during DDT is in apparent contradiction with the existence of anti-recombinogenic activities during DNA replication (Fabre et al, 2002). For instance, lysine K164 can also be SUMOylated, even in the absence of DNA damage. This modification is carried out by the Ubc9/Siz1 SUMO ligase complex and recruits the helicase Srs2 to prevent HR (Hoege et al, 2002; Papouli et al, 2005; Pfander et al, 2005). In fact, the activation of the replicative checkpoint by accumulation of ssDNA inhibits HR (Lisby et al, 2004; Meister et al, 2005; Alabert et al, 2009; Barlow and Rothstein, 2009). Thus, HR is often referred to as a double-edged sword; it is necessary for DNA damage repair and tolerance but can also generate genomic rearrangements when it is not properly regulated. However, the molecular scenarios that require or preclude recombination functions are still unknown. The HR mechanism has been extensively studied in response to DNA double-strand breaks (DSBs) (San Filippo et al, 2008). In contrast, much less is known about how HR is regulated during DDT, despite the fact that ssDNA gaps, and not DSBs, are the major lesions initiating spontaneous recombination (Fabre et al, 2002; Lettier et al, 2006). Indeed, its mode of action is still unclear. An important mechanistic question is whether gap repair by HR is coupled to the replication bypass across the lesion or whether it occurs post-replicatively. The efficiency of the response to replicative DNA damage is not affected when the expression of the group of RAD6 epistasis proteins is restricted to G2/M, suggesting that these mechanisms can operate uncoupled from the replication fork (Daigaku et al, 2010; Karras and Jentsch, 2010). The fact that not only rad18Δ, but also rad52Δ cells, accumulates ssDNA gaps behind the fork under replicative stress (Lopes et al, 2006; Hashimoto et al, 2010) supports the idea that HR can also work uncoupled from the fork. However, the recombination proteins Rad52 and Rad51 are required for replication fork progression through alkylated DNA (Vázquez et al, 2008; Alabert et al, 2009), suggesting that HR has additional, S phase-specific functions that remain to be determined. A major handicap for studying the role of HR during DDT is the difficulty of discriminating whether a recombinogenic lesion is associated with a ssDNA gap generated by replication fork impairment or with a DSB generated by processing a non-DSB DNA lesion. Furthermore, the few assays able to detect ssDNA gaps infer the role of HR from recombination mutants (Lopes et al, 2006; Gangavarapu et al, 2007), which can sometimes be misleading since the accuracy of HR relies on metastable and reversible intermediates (Heyer et al, 2010). To overcome these problems, we have used the Chromatin Endogenous Cleavage (ChEC) method (Schmid et al, 2004) to follow the binding of recombination proteins to replication forks and to DNA lesions other than DSBs during the cell cycle. We show that Rad52 and Rad51 are recruited to replication forks, where they facilitate DNA synthesis through alkylated DNA by a repair-independent process. Strikingly, the recruitment of Rad52 and Rad51 to chromatin during DNA replication is a prerequisite for the further repair of the lesion by HR, a process that is not coupled to the fork but rather restricted to G2/M by the replicative checkpoint. Results Physical evidence for the recruitment of Rad52 and Rad51 to replicative DNA damage other than DSBs To directly address whether recombination proteins are targeted to DNA lesions other than DSBs, we took advantage of the ChEC method developed by Laemmli and colleagues to map genomic interaction sites of chromatin proteins (Schmid et al, 2004). This method relies on the expression of proteins fused to the micrococcal nuclease (MN); the nuclease domain of these chimeras can be activated with Ca2+ ions and introduce a DNA DSB if they are bound to chromatin. For this, cells are permeabilized with digitonin, which does not affect protein–DNA interactions, and treated with Ca2+ for different times (Schmid et al, 2004). The rationale behind this approach is that a repair protein fused to MN will only generate a detectable cut in the DNA if it is targeted to a lesion that is not a DSB; by contrast, if the lesion is a DSB, the cleavage by the chimera will not enhance the appearance of DSBs (Figure 1A). Therefore, we can infer that recombination proteins bind to DNA if DNA is digested upon exposure to Ca2+. Figure 1.Physical evidence for Rad52 and Rad51 binding to non-DSB DNA lesions generated by replication through alkylated DNA. (A) Rationale of the approach. A DNA repair protein fused to MN will induce a detectable cut only if it is bound to a DNA lesion other than a DSB. (B, C) Rad51-MN and Rad52-MN are functional in MMS-induced DDT. (B) Response to chronic MMS treatment; cell growth analysis by 10-fold serial dilutions of the same number of mid-log phase cells. (C) Response to acute high-dose MMS treatment; wild-type, RAD52-MN, RAD51-MN, rad52Δ and rad51Δ cells were grown to mid-log phase, treated or not with 0.05% MMS for 2 h and plated onto YPD medium to determine their viability. The average and s.e.m. of three independent experiments are shown. (D) Rad52 binds to DNA in response to MMS. ChEC analysis of exponentially growing RAD52-MN cells incubated with 0.05% MMS for 2 h. The left and right panels show profiles for DNA content and DNA digestion. (E) Rad51 binds to DNA in response to MMS in a Rad52-dependent manner. ChEC analysis of RAD51-MN and RAD51-MN rad52Δ cells incubated with 0.05% MMS for 2 h. (F) DNA replication is required for Rad52 binding to MMS-induced DNA damage. RAD52-MN and Gp::CDC6 RAD52-MN cells were synchronized in G2/M in YP galactose with nocodazole, incubated for 1 h in YP glucose with nocodazole to remove Cdc6 (Tercero et al, 2003), and released into G1 in YP glucose with α factor. Finally, cells were released for 2 h in YP glucose with or without 0.05% MMS and analysed by ChEC. Download figure Download PowerPoint We fused MN to the C-terminal ends of Rad52 and Rad51 (Rad52-MN and Rad51-MN), and confirmed that these constructs were as proficient as the wild type in DNA damage repair and tolerance (Figures 1B and C; Supplementary Figure 1A). Rad52, which is essential for most HR events in yeast, binds to the 3′-ended ssDNA molecules generated by resection of a DSB and facilitates the formation of an ssDNA/Rad51 filament that mediates the search and exchange of homologous DNA sequences (San Filippo et al, 2008). To study the role of HR in repairing DNA lesions other than DSBs, cells were treated with methyl-methane sulfonate (MMS), a genotoxic agent that impairs replication fork progression by DNA alkylation (Tercero and Diffley, 2001) and that is highly toxic for cells defective in HR (Prakash and Prakash, 1977). After 2 h either with or without 0.05% MMS, cells were collected, permeabilized, and treated with Ca2+ for different times, and then total DNA was extracted and resolved in agarose gels. In the absence of Ca2+, a single, high molecular DNA band is detected on top of the gel (Figure 1D, time 0). In the absence of MMS, Rad52-MN digested DNA over a time course with Ca2+ (Figure 1D; note the gradual appearance of a smear below the top band in −MMS). Notably, the presence of MMS increased both the kinetics and extent (down to 1 kb) of DNA digestion by Rad52-MN (Figure 1D; compare+relative to −MMS at 30 and 45 min). The top band disappeared leading to a distribution of DNA fragments that peaked at ∼10 kb. This effect was not observed when we expressed only the MN (Supplementary Figure 1B). We also ruled out the possibility that the digestion was due to the fact that most MMS-treated cells remained at S phase (Figure 1D) by showing that DNA digestion by Rad52-MN in cells synchronized in G1, S, or G2/M and released in the absence of DNA damage did not change during the cell cycle (Supplementary Figure 1C). Importantly, and according to our hypothesis, introducing DNA DSBs with zeocin did not affect the kinetics of DNA cleavage by Rad52-MN (Supplementary Figure 1D). The expression of Rad51-MN also led to a Ca2+-dependent DNA digestion in the presence of MMS, even though less pronounced than that displayed by Rad52-MN. Importantly, this digestion was abolished in rad52Δ (Figure 1E), indicating that Rad52 mediates the binding of Rad51 to chromatin. Taken together, these results provide physical evidence for the recruitment of recombination proteins to non-DSBs DNA lesions. Next, we investigated whether Rad52 binding to MMS-induced DNA damage is dependent on replication. For this, we used cells expressing CDC6 under the control of the GAL1 promoter (Gp::CDC6). Expression of Cdc6 is essential for replication initiation but not for later cell-cycle events that depend on high levels of CDK activity (Piatti et al, 1995). Wild-type and Gp::CDC6 cells were maintained in glucose for 2 h during the synchronization steps to deplete Cdc6, after which G1 cells were released using medium with or without 0.05% MMS (Figure 1F). As reported, the absence of Cdc6 prevented DNA replication initiation and led to a 'reductional' cell division (Piatti et al, 1995). The absence of DNA replication also prevented Rad52-MN from binding to DNA in response to MMS, suggesting that Rad52 is recruited to non-DSBs replicative DNA lesions, presumably ssDNA gaps as suggested from Electron Microscopy studies with recombination mutants (Lopes et al, 2006; Hashimoto et al, 2010). Recruitment of recombination proteins to MMS-induced DNA damage is coupled to DNA replication In principle, Rad51 and Rad52 could be targeted to ssDNA gaps at or behind the fork, which are generated either by uncoupling the leading and lagging strands, or by repriming DNA synthesis downstream of damage, respectively. To determine this, cells were synchronized in G1 and released in the presence of 0.033% MMS for different times, and samples were analysed for Rad52-MN DNA cleavage (Figure 2A). Rad52-MN was bound to damaged DNA 1 h upon G1 release, when most cells were at S phase, as inferred from the DNA content profile. However, even at longer time points (3 and 4 h) at which bulk DNA was largely replicated, Rad52-MN was still bound to damaged DNA. This suggests that Rad52 can operate uncoupled from the replication fork at ssDNA gaps left behind the fork. Figure 2.The recruitment of Rad52 and Rad51 to MMS-induced DNA damage is coupled to DNA replication. (A) Rad52 can operate uncoupled from the replication fork in response to MMS. ChEC analysis of RAD52-MN cells synchronized in G1 and released in the presence of 0.033% MMS. (B) cRad52 is expressed specifically during G2/M. Western blot analysis of cRad52 in G2::cRAD52 cells synchronized in G1 and released into fresh medium. - indicates an asynchronous culture of a wild-type strain. Ponceau S staining and histone H3 levels were used as loading controls. (C) cRad52 is functional. G2::cRAD52, rad52Δ, and wild-type cells were synchronized in G2/M, incubated with or without 100 μg/ml zeocin for 30 min, and plated onto YPD medium to determine their viability. The average and s.e.m. of four independent experiments are shown. (D, E) Rad52 is required for S-phase progression in the presence of MMS, as determined by Flow cytometry analysis of cells synchronized in G1 and released in the presence of 0.033% MMS (D), and PFGE analysis of cells synchronized in G1, released in the presence of 0.033% MMS for 1 h (M), and then release into fresh media for the indicated times (min) (E). The quantification of lineal chromosomes is shown on the right. (F) Cells cannot tolerate MMS-induced DNA damage when HR is restricted to G2/M. Zeocin and MMS sensitivity of G2::cRAD52, rad52Δ, and wild-type cells. (G) Rad52 and Rad51 recruitment to MMS-induced DNA damage is coupled to DNA replication. ChEC analysis of RAD51-MN, G2::cRAD52 RAD51-MN, and rad52Δ RAD51-MN cells synchronized in G1 and released in the presence of 0.033% MMS for 270 min. The amount of cRad52 during the kinetics and of Rad51-MN 270 min after G1 release on MMS was determined by western blot. An asynchronous culture of rad51Δ was used as a negative control. Histone H3 and Pgk1 were used as loading controls. cRad52 was detected with an antibody against Clb2. Download figure Download PowerPoint If Rad52 can operate uncoupled from the fork, then one prediction is that HR should be able to repair DNA damage even if HR is restricted to G2/M. To address this point, we limited Rad52 expression to G2/M by inserting the promoter and degron sequences of CLB2 in front of the coding sequence of RAD52 (G2::cRAD52) (Figure 2B). The cRad52 chimera was functional, as determined by analysing the viability of cells arrested in mitosis and then treated for 30 min with zeocin (Figure 2C). As expected for cells lacking Rad52 during S phase, G2::cRAD52 cells were defective in completing DNA replication in the presence of 0.033% MMS as determined by DNA content (Figure 2D) and PFGE (Figure 2E), where only replicated chromosomes enter into the gel. The absence of Rad52 during S phase had little effect on the sensitivity to zeocin-induced DSBs (Figure 2F, top); however, in contrast to our prediction, G2::cRAD52 cells were highly sensitive to MMS (Figure 2F, bottom). This indicates that cells cannot tolerate replicative DNA damage if HR is restricted to G2/M. The fact that recombination proteins were bound to damaged sites after replication was largely completed but could not tolerate the damage when they were expressed only in G2/M suggests that their recruitment to chromatin is coupled to DNA replication. To assess this possibility, we followed Rad51-MN binding to chromatin in G2::cRAD52 cells by ChEC analysis. We first demonstrated that both cRad52 and Rad51-MN are functional when expressed together (Supplementary Figure 2). Cells were then synchronized in G1, released in the presence of MMS, and analysed after 270 min, at which point most G2::cRAD52 cells remained arrested in mitosis and expressed cRad52 (Figure 2G). As previously shown, Rad52 was required for Rad51-MN binding to chromatin in response to MMS. Importantly, Rad51-MN binding was also prevented when Rad52 expression was restricted to G2/M (Figure 2G). This result, which cannot be explained by reduced levels of Rad51-MN in the mutant relative to the wild type (Figure 2G), demonstrates that the binding of Rad52 and Rad51 to damaged chromatin has to occur during S phase. It is worth noting that this result discards the possibility that the ChEC signal measured the binding of HR factors to ssDNA during the Ca++ treatment, since G2::cRAD52 cells in G2/M have Rad52, Rad51-MN, and ssDNA. In conclusion, our results indicate that the recruitment of recombination proteins is coupled to DNA replication and this step is essential for DDT. The recruitment of Rad51 to replicating chromatin is required for MMS-induced HR repair According to our previous results, the recruitment of Rad51 to chromatin during DNA replication should be a prerequisite for the recombinational repair of replicative DNA lesions. DNA damage repair by HR is associated with repair centres that can be detected with the recombination proteins Rad52 and Rad51 fused to the green fluorescence protein (Lisby et al, 2001, 2004). While Rad52-YFP is biologically functional (Lisby et al, 2001), tagging of Rad51 with the fluorescence protein reduces its repair activity (Lisby et al, 2004). We thereby worked with yeast strains expressing both the chimera and the wild-type protein, which are fully functional in the repair of MMS and zeocine-induced DNA damage (Supplementary Figure 3A). Following the aforementioned strategy with Gp::CDC6 cells, we first confirmed that, as expected for replicative DNA damage, replication inhibition in the absence of Cdc6 prevented the assembly of MMS-induced HR foci (Figure 3A). Figure 3.Rad51 fork recruitment is a prerequisite for MMS-induced HR repair. (A) DNA replication is required for MMS-induced Rad52-YFP foci. Rad52 foci accumulation in wild-type and Gp::CDC6 cells transformed with pWJ1344 (RAD52-YFP), synchronized in G1 as shown in Figure 1F, and released in YP glucose with 0.01% MMS. (B) Sgs1 and Exo1 are required for zeocin-induced, but not for MMS-induced HR foci. Rad52 foci accumulation in wild-type, sgs1Δ, exo1Δ, and sgs1Δ exo1Δ cells transformed with pWJ1344 (RAD52-YFP), synchronized in G1, and released in the presence of 0.01% MMS (M) or 100 μg/ml zeocin (Z) for 90 and 120 min, respectively. (C) Rad52 expression during S phase is required for MMS-induced, but not for zeocin-induced HR foci. Rad51 foci accumulation in YFP-RAD51 and G2::cRAD52 YFP-RAD51 cells transformed with pWJ1278 (RAD51), synchronized in G1, and released in the presence of 0.01% MMS (left panel) or 100 μg/ml zeocin (right panel). The average and s.e.m. are shown. (D) Rad52 expression during S phase is required for MMS-induced SCJs. 2D gel analysis of X-shaped molecules in sgs1Δ and sgs1Δ G2::cRAD52 cells synchronized in G1 and released in the presence of 0.033% MMS. The amount of X-shaped molecules relative to the total amount of molecules, taken the highest value as 100, is shown. The experiment was repeated with similar results. (E) Rad52 expression during S phase is required for ssDNA gap repair. RPA foci accumulation and FACS analysis in RFA1-YFP1, RFA1-YFP1 G2::cRAD52 and RFA1-YFP1 rad52Δ cells synchronized in G1, released in 0.01% MMS for 1 h, treated with 2.5% sodium thiosulphate to inactivate the MMS, washed and incubated in fresh medium for different times. Bright field, DAPI fluorescence, and Rfa1-YFP foci of selected cells are shown. Download figure Download PowerPoint Since a single DSB is able to generate a Rad52 focus (Lisby et al, 2003), the foci induced by MMS might reflect the repair of residual DSBs caused by replication through alkylated DNA. To address this point, we analysed the role of Exo1 and Sgs1, required for DNA resection during DSB repair by HR (Gravel et al, 2008; Zhu et al, 2008; Mimitou and Symington, 2008), on MMS-induced HR foci. The double mutant exo1Δ sgs1Δ, and partially the single mutants sgs1Δ and exo1Δ, suppressed zeocin but not MMS-induced recombination foci (Figure 3B), suggesting that MMS-induced foci are indeed associated with non-DSBs DNA lesions. Consistently, exo1Δ sgs1Δ did not prevent the binding of Rad52 to MMS-induced DNA damage as inferred from ChEC analysis (Supplementary Figure 3B). Nevertheless, we do not rule out the possibility that ssDNA gaps may need to be enlarged for efficient DNA repair (e.g., for ssDNA/Rad51 filament formation). According with this idea, exo1Δ sgs1Δ accumulated MMS-induced recombination foci (Figure 3B) and were defective in SCJs formation (Vanoli et al, 2010) as compared to the wild type. In this frame, it is worth noting that this double mutant accumulated high levels of foci in the absence of DNA damage, supporting the idea that spontaneous recombinogenic damage is associated with replicative non-DSB DNA lesions (Fabre et al, 2002; Lettier et al, 2006). Once established that the repair by HR of MMS-induced non-DSBs replicative lesions is associated with foci, we examined the effect of restricting Rad52 expression to G2/M on MMS-induced YFP-Rad51 foci. G2::cRAD52 and wild-type cells expressing YFP-Rad51 were synchronized in G1 and released in the presence of 0.01% MMS. The lack of Rad52 during S phase in G2::cRAD52 suppressed the formation of Rad51 foci in response to MMS (Figure 3C, left panel). This effect was not due to reduced levels of YFP-Rad51 in the mutant relative to the wild type (Supplementary Figure 3C). Therefore, the recruitment of recombination proteins to chromatin during DNA replication is a prerequisite for the further repair of the lesion. This is an important difference to DSB repair by HR, which is independent of DNA replication (Alabert et al, 2009; Barlow and Rothstein, 2009) and can occur efficiently in G2 (Ira et al, 2004) even if Rad52 is not expressed during S phase (Figures 2C and F). Accordingly, G2::cRAD52 cells synchronized in G1 and released in the presence of zeocin formed YFP-Rad51 foci in G2/M (Figure 3C, right panel; Supplementary Figure 3D). In cells lacking Sgs1, the repair of MMS-induced DNA lesions leads to an accumulation of SCJs (X-shaped molecules), a process that requires Rad51 and Rad52 (Liberi et al, 2005). Thus, we followed the kinetics of X-shaped molecules in sgs1Δ and sgs1Δ G2::cRAD52 cells synchronized in G1 and released in the presence of 0.033% MMS. As shown in Figure 3D, the absence of Rad52 during S phase led to a dramatic drop in the amount of SCJs, suggesting that Rad52 cannot promote sister-chromatid recombination in response to non-DSB replicative lesions if it is not timely loaded during replication. An expectation of these results is that G2::cRAD52 cells accumulated unrepaired ssDNA gaps. ssDNA molecules are coated with replication protein A (RPA; formed by Rfa1-3 subunits), and can be detected by using the biologically functional chimera Rfa1-YFP (Lisby et al, 2004). Most wild-type cells released from G1 into medium containing 0.01% MMS for 1 h accumulated Rfa1-YFP foci (∼75%), and this value was even higher in G2::cRAD52 and rad52Δ cells (∼90%) (Figure 3E, top). Inactivation of the MMS and further incubation in fresh medium led to a gradual disappearance of the RPA foci and normal cell-cycle progression in the wild type, whereas both G2::cRAD52 and rad52Δ maintained the initial fraction of cells with foci 4 h after release from MMS and remained arrested in G2/M. Of note, the pattern of RPA foci during the kinetics was independent of Rad52; multiple faint foci after 1 h in the presence of MMS that gradually ended up into 1–2 bright foci after release in fresh medium (Figure 3E, bottom). Faint speckled foci and 1–2 bright foci have been associated with replication and HR, respectively (Lisby et al, 2004; Burgess et al, 2009). These results suggest that the presence of Rad52 during S phase is required for the repair of the replicative ssDNA gaps but not for the recruitment of the DNA lesions to the repair centres. Rad52 and Rad51 bind to replication forks regardless of the presence of DNA damage The fact that Rad52 and Rad51 have to be recruited onto chromatin during DNA replication in order to promote DDT suggests that their loading is coupled to the replication fork. To demonstrate at the molecular level that these proteins bind to forks, replication intermediates (RIs) from ChEC-treated cells were analysed by 2D-gel electrophoresis. Here, fork binding of a protein fused to MN and subsequent cleavage would generate a new population of RIs. To validate the assay, and once established that Ca++ treatment did not affect the pattern of RIs (Supplementary Figure 4A), we first analysed Rad27, a nuclease involved in the processing of Okazaki fragments (Zheng and Shen, 2011). We analysed RIs at a fragment that contains the early origin ARS305; RIs initiated f
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