Non‐recombinogenic roles for Rad52 in translesion synthesis during DNA damage tolerance
2020; Springer Nature; Volume: 22; Issue: 1 Linguagem: Inglês
10.15252/embr.202050410
ISSN1469-3178
AutoresMaría I. Cano-Linares, Aurora Yáñez‐Vilches, Néstor García‐Rodríguez, Marta Barrientos‐Moreno, Román González‐Prieto, Pedro A. San-Segundo, Helle D. Ulrich, Félix Prado,
Tópico(s)Carcinogens and Genotoxicity Assessment
ResumoArticle2 December 2020free access Source DataTransparent process Non-recombinogenic roles for Rad52 in translesion synthesis during DNA damage tolerance María I Cano-Linares María I Cano-Linares Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, SpainThese authors contributed equally to this work Search for more papers by this author Aurora Yáñez-Vilches Aurora Yáñez-Vilches Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, SpainThese authors contributed equally to this work Search for more papers by this author Néstor García-Rodríguez Néstor García-Rodríguez Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Marta Barrientos-Moreno Marta Barrientos-Moreno Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain Search for more papers by this author Román González-Prieto Román González-Prieto orcid.org/0000-0001-8997-2321 Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain Search for more papers by this author Pedro San-Segundo Pedro San-Segundo Institute of Functional Biology and Genomics (IBFG), CSIC-University of Salamanca, Salamanca, Spain Search for more papers by this author Helle D Ulrich Helle D Ulrich orcid.org/0000-0003-0431-2223 Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Félix Prado Corresponding Author Félix Prado [email protected] orcid.org/0000-0001-9805-782X Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain Search for more papers by this author María I Cano-Linares María I Cano-Linares Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, SpainThese authors contributed equally to this work Search for more papers by this author Aurora Yáñez-Vilches Aurora Yáñez-Vilches Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, SpainThese authors contributed equally to this work Search for more papers by this author Néstor García-Rodríguez Néstor García-Rodríguez Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Marta Barrientos-Moreno Marta Barrientos-Moreno Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain Search for more papers by this author Román González-Prieto Román González-Prieto orcid.org/0000-0001-8997-2321 Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain Search for more papers by this author Pedro San-Segundo Pedro San-Segundo Institute of Functional Biology and Genomics (IBFG), CSIC-University of Salamanca, Salamanca, Spain Search for more papers by this author Helle D Ulrich Helle D Ulrich orcid.org/0000-0003-0431-2223 Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Félix Prado Corresponding Author Félix Prado [email protected] orcid.org/0000-0001-9805-782X Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain Search for more papers by this author Author Information María I Cano-Linares1, Aurora Yáñez-Vilches1, Néstor García-Rodríguez2,†, Marta Barrientos-Moreno1, Román González-Prieto1,†, Pedro San-Segundo3, Helle D Ulrich2 and Félix Prado *,1 1Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain 2Institute of Molecular Biology (IMB), Mainz, Germany 3Institute of Functional Biology and Genomics (IBFG), CSIC-University of Salamanca, Salamanca, Spain †Present address: Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine Center (CABIMER), CSIC-University of Seville-University Pablo de Olavide, Seville, Spain †Present address: Department of Cell and Chemical Biology, Leiden University Medical Center, Leiden, The Netherlands *Corresponding author. Tel: +34 954468210; E-mail: [email protected] EMBO Reports (2021)22:e50410https://doi.org/10.15252/embr.202050410 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 Abstract DNA damage tolerance relies on homologous recombination (HR) and translesion synthesis (TLS) mechanisms to fill in the ssDNA gaps generated during passing of the replication fork over DNA lesions in the template. Whereas TLS requires specialized polymerases able to incorporate a dNTP opposite the lesion and is error-prone, HR uses the sister chromatid and is mostly error-free. We report that the HR protein Rad52—but not Rad51 and Rad57—acts in concert with the TLS machinery (Rad6/Rad18-mediated PCNA ubiquitylation and polymerases Rev1/Pol ζ) to repair MMS and UV light-induced ssDNA gaps through a non-recombinogenic mechanism, as inferred from the different phenotypes displayed in the absence of Rad52 and Rad54 (essential for MMS- and UV-induced HR); accordingly, Rad52 is required for efficient DNA damage-induced mutagenesis. In addition, Rad52, Rad51, and Rad57, but not Rad54, facilitate Rad6/Rad18 binding to chromatin and subsequent DNA damage-induced PCNA ubiquitylation. Therefore, Rad52 facilitates the tolerance process not only by HR but also by TLS through Rad51/Rad57-dependent and -independent processes, providing a novel role for the recombination proteins in maintaining genome integrity. Synopsis The recombination protein Rad52 promotes translesion synthesis and mutagenesis by non-recombinogenic functions, which include the recruitment of the PCNA ubiquitylation complex Rad6/Rad18 to chromatin. The recombination protein Rad52 facilitates the repair of MMS-and UV-induced ssDNA lesions through Rad51/Rad57-dependent and independent non-recombinogenic functions Rad52, but not Rad51, Rad57and Rad54, is required for efficient DNA damage-induced mutagenesis Rad52, together with Rad51 and Rad57 but not Rad54, facilitates Rad6/Rad18 binding to chromatin and subsequent DNA damage-induced PCNA ubiquitylation Introduction Apart from a number of specialized DNA repair mechanisms aimed to deal with the different types of DNA damage, cells are endowed with mechanisms to tolerate DNA lesions that hinder the advance of replication forks, thus ensuring timely chromosome replication (Broomfield et al, 2001; Friedberg, 2005). The mechanisms of DNA damage tolerance (DDT) help replication forks to pass through the lesions and fill in the stretches of single-stranded DNA (ssDNA) generated during the blocking of the fork. The DDT response involves two different evolutionarily conserved strategies to fill in the ssDNA gaps: translesion synthesis (TLS) and homologous recombination (HR) (Sale, 2013; Prado, 2014, 2018; Branzei & Psakhye, 2016). TLS mechanisms use specialized polymerases that are able to incorporate a nucleotide opposite the lesion (Fig 1A), yet this process can be mutagenic as these polymerases display low fidelity. In Saccharomyces cerevisiae, there are three TLS polymerases: Rev1, Pol ζ (formed by the catalytic subunit Rev3 and the regulatory subunits Rev7, Pol31, and Pol32), and Pol η (encoded by RAD30) (Sale, 2013; Zhao & Washington, 2017). Their contribution to DDT is influenced by the dose and type of blocking lesions, and their activity (at least for Rev1/ Pol ζ) is most dominant in G2/M (Prakash, 1981; Paulovich et al, 1997; Baynton et al, 1998; Torres-Ramos et al, 2002; Lopes et al, 2006; Waters & Walker, 2006; Daigaku et al, 2010). Figure 1. Persistence of MMS-induced Rad52 foci in TS-deficient cells A. Scheme for the two major mechanisms of DDT: HR and TLS. An asterisk represents a blocking DNA lesion for the replicative polymerase. B. Representative image of wild-type cells with MMS-induced Rad52-YFP foci (bright field and fluorescence signal) C–G. ssDNA repair efficiency of the indicated TS mutants as determined from the maximal and final percentages of cells with Rad52 foci during the time course (right panels). Cells were synchronized in G1 and released into S phase in the presence of 0.033% MMS for 1 h (60+), treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. They were also released into medium without MMS for 1 h to control the formation of spontaneous Rad52-YFP foci (60-). The percentage of cells with foci at each point was normalized to the highest value of the wild type, taken as 100 (left panels). Cell cycle progression was determined by cell sorting. The mean and SEM of 3 (wild type, rad18∆, mms2∆, rad51∆, rad57∆, rad54∆), 4 (pol30-K164R), and 5 (rad5∆) independent experiments are shown. Statistically significant differences according to an unpaired two-tailed Student's t-test are shown, where three asterisks represent P-values <0.001. Source data are available online for this figure. Source Data for Figure 1 [embr202050410-sup-0004-SDataFig1.xlsx] Download figure Download PowerPoint In marked contrast to TLS, HR mechanisms use the intact sister chromatid to fill in the ssDNA gaps and are considered error-free (Fig 1A). These mechanisms have been extensively studied in yeast, where they require different components including Rad51, Rad52, Rad55/Rad57, and Rad54 (Prado, 2018). The recombinase Rad51 provides strand annealing and exchange activities. The mediator Rad52 competes with the ssDNA-binding replication protein A (RPA complex; formed by Rfa1, Rfa2, and Rfa3) to load Rad51 and form the ssDNA/Rad51 nucleofilament that initiates the search for homology. The helper complex Rad55/Rad57 is involved in the stabilization of the ssDNA/Rad51 nucleofilament. Finally, the translocase Rad54 is involved in the processing of early and late recombination intermediates. A key component in the control of DDT is the replication processivity factor PCNA (Hoege et al, 2002). In response to replicative DNA damage, but not double-strand breaks (DSBs), the Rad6/Rad18 complex monoubiquitylates PCNA at lysine 164 (Hoege et al, 2002; Davies et al, 2008), a modification that promotes the recruitment of the TLS polymerases (Bienko et al, 2005). This ubiquitin residue is further extended with a K63-linked polyubiquitin chain by the activity of the Mms2/Ubc13/Rad5 complex (Hoege et al, 2002; Lopes et al, 2006; Daigaku et al, 2010). The HR proteins act in concert with polyubiquitylated PCNA to form sister chromatid junctions (SCJs) as an intermediate to fill in the gaps of ssDNA (Liberi et al, 2005; Mankouri et al, 2007; Branzei et al, 2008; Minca & Kowalski, 2010; Vanoli et al, 2010; Karras et al, 2013; Giannattasio et al, 2014). HR can also deal with ssDNA gaps through an UbPCNA-independent mechanism. This UbPCNA-independent HR mechanism is also associated with SCJs (Branzei et al, 2008) and seems to deal with ssDNA fragments that remain unrepaired after S phase; accordingly, it is inhibited during S phase by Ubc9/Siz1-dependent PCNA sumoylation at lysines 164 and 127 and further recruitment of the antirecombinogenic helicase Srs2 (Schiestl et al, 1990; Pfander et al, 2005; Papouli et al, 2005). In contrast to UbPCNA/HR, the UbPCNA-independent HR pathway can lead to chromosomal rearrangements (Motegi et al, 2006). The UbPCNA-dependent and UbPCNA-independent HR pathways are also termed template switching (TS) and salvage pathway, respectively (Karras et al, 2013; Prado, 2014, 2018; Branzei & Psakhye, 2016). Epistatic analyses between HR and TLS mutants on DNA damage sensitivity have genetically separated the role of HR proteins from TLS (Rattray et al, 2002; Ball et al, 2009). In fact, the frequency of UV light-induced mutagenesis in cells defective in nucleotide-excision repair (NER)—essential for the removal of UV-induced photoproducts—is increased in a rad52∆ mutant. This has been interpreted as a consequence of channeling the filling of most ssDNA gaps to TLS (Paulovich et al, 1998). However, the interplay between HR proteins and TLS is more complex. Thus, the absence of Rad52 in NER proficient cells decreases the frequency and alters the type of UV- and ethyl methanesulfonate (EMS)-induced mutations (Prakash & Higgins, 1982; Kunz et al, 1992; Armstrong et al, 1994), which might reflect a role for Rad52 in TLS (Armstrong et al, 1994). However, these phenotypes might also be due to the loss of recombinational repair, a process that can increase the rate and type of mutations (Hicks et al, 2010). An additional level of complexity in the interplay between HR and TLS proteins comes from their role in spontaneous and DSB-induced events. However, a distinctive feature with the DDT mechanisms is that the repair of spontaneous and DSB lesions is not associated with PCNA ubiquitylation (Hoege et al, 2002; Stelter & Ulrich, 2003; Hirano & Sugimoto, 2006; Chen et al, 2006; Sharma et al, 2012). Here, we show that Rad52—but not Rad51 or Rad57—acts in concert with the TLS machinery to repair methyl methanesulfonate (MMS)- and UV-induced ssDNA gaps in a non-recombinogenic manner, as inferred from the different phenotypes displayed by rad52∆ and rad54∆ mutants. Accordingly, Rad52 is required for efficient DNA damage-induced mutagenesis. In addition, Rad52 cooperates with Rad51 and Rad57—but not Rad54—to promote efficient Rad6/Rad18 binding to chromatin and in response to replicative DNA lesions, PCNA ubiquitylation. Therefore, Rad52 can facilitate both HR and TLS at DNA repair centers, suggesting a regulatory function for this protein in the decision of tolerating DNA damage through mutagenic or recombinogenic mechanisms. Results Persistence of MMS-induced Rad52 foci in TS-deficient cells DNA damage induces the formation of DNA repair centers that can be visualized as nuclear foci using recombination proteins fused to a fluorescent protein (Fig 1B; Lisby et al, 2001). In the case of MMS, these foci mark post-replicative Rad52-mediated gap filling events (González-Prieto et al, 2013; Wong et al, 2020). We decided to follow the disappearance of Rad52 centers formed in response to MMS to explore the genetic contribution of the different DDT pathways to the post-replicative repair of ssDNA gaps. For this, cells expressing Rad52-YFP, which is functional in response to MMS (González-Prieto et al, 2013), were synchronized in G1 and released into fresh medium in the presence of 0.033% MMS for 1 h, treated with sodium thiosulfate to inactivate the MMS, washed, and incubated in fresh medium for different times. All time courses included a wild type to control the experimental variation in the G1 release and cell cycle progression between different sets of experiments. The maximal fraction of wild-type cells with Rad52 foci was ~40–60% and was reached 1–2 h after MMS inactivation, coinciding with completion of DNA replication. This percentage of wild-type cells with Rad52-YFP foci dropped to ~10% 5–6 h later (Fig 1, left panels; note that plotted values have been normalized to the highest value of the wild type, taken as 100). The efficiency of ssDNA repair, determined from the maximal and final percentages of cells with Rad52 foci during the time course, was ~80% (Fig 1, right panels; see Material and Methods for calculation). To address the relevance of DDT-induced PCNA modifications on Rad52-associated gap filling, we analyzed the effect of pol30-K164R, which expresses a PCNA complex mutated in its major site of ubiquitylation and sumoylation (Hoege et al, 2002). The efficiency of repair in pol30-K164R cells was reduced to ~20% (Fig 1C), suggesting that the modification of PCNA at lysine 164 is important for the resolution of Rad52 foci. Since PCNA polyubiquitylation at lysine 164 cooperates with recombination proteins in the process of TS (Prado, 2014, 2018; Branzei & Psakhye, 2016), we followed the accumulation of Rad52 foci in the absence of PCNA ubiquitylation (Rad18, Mms2, and Rad5) and HR (Rad51, Rad57, and Rad54) factors. The percentage of cells with Rad52 foci 1–2 h after MMS release was similar in these mutants to that displayed by the wild type, except for the rad57∆ and rad54∆ mutants that exhibited an unexpected lower percentage (Fig 1D–G). Importantly, in all cases the efficiency of ssDNA repair dropped to less than 20% (Fig 1D–G). These results suggest that the TS machinery is required for efficient post-replicative ssDNA gap repair at Rad52 foci. Persistence of MMS-induced Rad52 foci in TLS-deficient cells TS operates mostly in S phase (Karras & Jentsch, 2010; Daigaku et al, 2010; Ortiz-Bazán et al, 2014). Thus, PCNA polyubiquitylation should be dispensable for HR after completion of DNA replication. To address this point, the RAD18 promoter was replaced by the GAL1 promoter (Gp::RAD18), which is expressed in the presence of galactose and repressed in the presence of glucose. Accordingly, the kinetics of Rad52 foci accumulation in Gp::RAD18 cells grown in galactose was similar to that displayed by wild-type cells (Fig EV1A), indicating that Rad18 expression from the GAL1 promoter sustained DNA repair. Interestingly, if Rad18 expression was repressed after completion of DNA replication (1 h after MMS inactivation) by shifting the cells from galactose to glucose-containing medium, a significant percentage of Rad52 foci persisted at the end of the time course compared to the wild type (Fig 2A). This suggests that PCNA ubiquitylation is required for efficient ssDNA gap repair at Rad52 centers after completion of S phase. To confirm that this requirement was for PCNA monoubiquitylation and not polyubiquitylation, we repeated the analysis using a strain in which the MMS2 gene was under control of the GAL1 promoter (Gp::MMS2). In this case, cells expressing MMS2 from the GAL1 promoter in galactose repaired the MMS-induced ssDNA gaps with lower efficiency (~50%; Fig 2B, condition a) than wild-type cells (~80%; Fig EV1B). Repression of Mms2 expression for only 1 h before release into S phase was sufficient to cause a complete defect in Rad52-associated ssDNA repair (Fig 2B; compare conditions a and b). In contrast, repression of Mms2 expression after completion of DNA replication (2 h after MMS inactivation) did not compromise repair efficiency as compared with cells continuously expressing Mms2 (Fig 2B; compare conditions a and c), indicating that PCNA polyubiquitylation is unlikely to be important in G2/M. These results are consistent with a specific role for PCNA monoubiquitylation in the resolution of Rad52 foci. Click here to expand this figure. Figure EV1. Time course analyses of Rad52-YFP foci in DDT mutants A. ssDNA repair efficiency of cells expressing RAD18 from the GAL1 promoter as determined from the maximal and final percentages of cells with Rad52 foci during the time course (right panel). Cells were synchronized in G1 from galactose-containing medium and released into S phase in the presence of 0.033% MMS for 1 h, treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. Cells were grown in galactose-containing medium during the whole time course. The percentage of cells with foci at each point was normalized to the highest value of the wild type, taken as 100 (left panel). See Fig 2A for more details. The mean and SEM of four independent experiments are shown. B. ssDNA repair efficiency of mms2∆ (right) and wild-type (left) cells as determined from the maximal and final percentages of cells with Rad52 foci during the time course (right panels). Cells were synchronized in G1 from galactose-containing medium and shifted to glucose (a) or maintained in galactose (b, c) for 1 h in G1; then, cells were released into S phase in the presence of 0.033% MMS for 1 h, treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. Cells were maintained either in galactose (a), glucose (b), or shifted to glucose 2 h after MMS inactivation (c). The percentage of cells with foci at each point was normalized to the highest value in galactose, taken as 100 (left panels). See Fig 2B for more details. The mean and SEM of three independent experiments are shown. Data information: Statistically significant differences according to an unpaired two-tailed Student's t-test are shown, where three asterisks represents a P-value < 0.001. Download figure Download PowerPoint Figure 2. Persistence of MMS-induced Rad52 foci in TLS-deficient cells A. ssDNA repair efficiency of cells expressing RAD18 from the GAL1 promoter as determined from the maximal and final percentages of cells with Rad52 foci during the time course (right panel). Cells were synchronized in G1 from galactose-containing medium and released into S phase in the presence of 0.033% MMS for 1 h, treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. Cells were shifted to glucose-containing medium 1 h after MMS inactivation. The percentage of cells with foci at each point was normalized to the highest value of the wild type, taken as 100 (left panel). See Fig EV1A for control cells released in galactose-containing medium during the entire time course. B. ssDNA repair efficiency of cells expressing MMS2 from the GAL1 promoter as determined from the maximal and final percentages of cells with Rad52 foci during the time course (right panel). Cells were synchronized in G1 from galactose-containing medium and maintained in galactose (a, c) or shifted to glucose (b) for 1 h in G1; subsequently, cells were released into S phase in the presence of 0.033% MMS for 1 h, treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. Cells were maintained either in galactose (a), glucose (b) or shifted to glucose 2 h after MMS inactivation (c). The percentage of cells with foci at each point was normalized to the highest value in galactose, taken as 100 (left panel). Wild-type and mms2∆ controls were analyzed in parallel (Fig EV1B). C–E. ssDNA repair efficiency of the indicated TLS mutants as determined from the maximal and final percentages of cells with Rad52 foci during the time course (D and E, right panel). Cells were synchronized in G1 and released into S phase in the presence of 0.033% MMS for 1 h, treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. The percentage of cells with foci at each point was normalized to the highest value of the wild type, taken as 100 (C and E, left panel). Data information: The mean and SEM of 4 (A), 3 (B, E) and 3–12 (C, D; 12 (wild type), 6 (rev1∆, rev3∆, rad30∆, rev1∆ rad30∆, rev3∆ rad30∆), and 3 (rev1∆ rev3∆ and rev1∆ rev3∆ rad30∆) independent experiments are shown. Statistically significant differences according to an unpaired two-tailed Student's t-test are shown, where one, two, and three asterisks represent P-values < 0.05, < 0.01, and < 0.001, respectively. Source data are available online for this figure. Source Data for Figure 2 [embr202050410-sup-0005-SDataFig2.xlsx] Download figure Download PowerPoint In addition to priming PCNA polyubiquitylation for TS, Rad18-mediated PCNA monoubiquitylation is required for the recruitment of TLS polymerases to DNA lesions (Hoege et al, 2002; Stelter & Ulrich, 2003; Bienko et al, 2005). Moreover, Rev1-dependent TLS occurs preferentially in G2/M (Waters & Walker, 2006). Thus, we examined if blocking TLS by eliminating the TLS polymerases also impairs ssDNA processing at Rad52 repair centers. Remarkably, the absence of any of the highly mutagenic polymerases, Rev1 or Pol ζ, in rev1∆ or rev3∆ cells reduced the efficiency of ssDNA repair at Rad52 foci (~35–40%), whereas the absence of Pol η in rad30∆ had a much weaker effect (Fig 2C and D). Interestingly, the lack of a regulatory subunit of Pol ζ in pol32∆ cells caused a more severe defect (~20%; Fig 2E). This might be related to the role of Pol32 as part of the replicative polymerase Pol δ in the formation of SCJ during TS (Vanoli et al, 2010). The effects of rev1∆ and rev3∆ were not additive (Fig 2C and D), consistent with a cooperation between Rev1 and Pol ζ (Haracska et al, 2001). In fact, the double mutant rev1∆ rev3∆ increased ssDNA repair efficiency as compared to the single mutants, and this increase required the activity of the polymerase Pol η (Fig 2D; compare rev1∆ rev3∆ rad30∆ with rev1∆ rev3∆). These results indicate that Pol η can operate in the absence of the entire Rev1/Pol ζ complex but not of just either Rev1 or Pol ζ, suggesting that the presence of a non-functional Rev1/Pol ζ complex hampers the entry and/or activity of Pol η at the ssDNA gap. In sum, these results suggest that Rad6/Rad18-mediated PCNA monoubiquitylation, the activities of Rev1/Pol ζ, and to a lesser extent Pol η are required for efficient post-replicative processing of the ssDNA gaps that occurs at the Rad52 repair centers in response to MMS. Persistence of Rad52 foci in TLS mutants is not due to a demand for TLS polymerases during HR Since cells are released into S phase in the presence of MMS in our experimental setup, the blocking lesions accumulate not only ahead of the fork but also within the nascent strands. Thus, one possibility to explain the genetic connection between the processing of ssDNA gaps at Rad52 repair centers and the TLS machinery would be that DNA synthesis at the invaded sister chromatid during HR was blocked by additional lesions and therefore required TLS polymerases (Fig 3A; MMS in S/G2). To address this possibility, cells were synchronized in G1 and treated with MMS for 1 h; after that, the MMS was inactivated, and then, cells were released into S phase in the absence of DNA damage. Under these conditions, the MMS-induced blocking lesions are restricted to the parental molecule, leaving intact the nascent DNA molecules (Fig 3A; MMS in G1 only). In this case, Rad52 foci also persisted in the absence of Rev1 (Fig 3A). Therefore, the requirement of Rev1 for ssDNA processing at Rad52 repair centers is not due to a demand for TLS polymerases to bypass lesions in the newly synthesized DNA. Hence, TLS polymerases are unlikely to be required for the HR process. To confirm this, we analyzed HR using an unequal sister chromatid exchange (uSCE) recombination system (Fasullo & Davis, 1987) in cells growing on MMS-containing solid medium. Neither Rev1 nor Rev3 was required for recombination; actually, the frequency of MMS-induced HR increased in both rev1∆ and rev3∆ mutants (Fig 3B), consistent with the channeling of ssDNA gaps to HR in the absence of TLS. Figure 3. Persistence of Rad52 foci in TLS mutants is not due to a demand for TLS polymerases during HR A. ssDNA repair efficiency of the rev1∆ mutant treated with MMS only in G1 as determined from the maximal and final percentages of cells with Rad52 foci during the time course (right panel). Cells were synchronized in G1 and treated with 0.033% MMS for 1 h; after that the MMS was inactivated with 2.5% sodium thiosulfate and cells were released into fresh medium in the absence of DNA damage. The percentage of cells with foci at each point was normalized to the highest value of the wild type, taken as 100 (middle panel). A scheme on the left shows the experimental set up followed in this (MMS in G1 only) and the other experiments (MMS in S/G2). An asterisk represents a blocking DNA lesion for the replicative polymerase. B. Effect of the rev1∆ and rev3∆ mutations in spontaneous and MMS-induced uSCE. The frequency of spontaneous and MMS-induced recombinants was determined from colonies grown in the absence or presence of 0.01% MMS. The mean and SEM of four independent fluctuation tests are shown. C, D. ssDNA repair efficiency of the rad5∆ and rev1∆ mutants as determined from the maximal and final percentages of cells with Rad54 foci during the time course (right panels). Cells were synchronized in G1 and released into S phase in the presence of 0.033% MMS for 1 h, treated with 2.5% sodium thiosulfate to inactivate the MMS, washed, and released into fresh medium for different times. The percentage of cells with foci at each point was normalized to the highest value of the wild type, taken as 100 (left panels). Data information: The mean and SEM of 4 (A) and 3 (C, D) independent experiments are shown. Statistically significant differences according to an unpaired two-tailed Student's t-test are shown, where one, two, and three asterisks represent P-values < 0.05, < 0.01, and < 0.001, respectively. Source data are available online for this fi
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