Ctf4 Prevents Genome Rearrangements by Suppressing DNA Double-Strand Break Formation and Its End Resection at Arrested Replication Forks

Article

Ctf4 Prevents Genome Rearrangements by Suppressing DNA Double-Strand Break Formation and Its End Resection at Arrested Replication Forks Graphical Abstract

Authors Mariko Sasaki, Takehiko Kobayashi

3′ 5′

replication fork arrest

Correspondence [email protected]

replisome Fob1

In Brief Ctf4 +Ctf4

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MRX-dependent HR-independent repair

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Exo1 Sgs1-Dna2 HR-mediated repair

DSB repair without rDNA amplification

DSB repair with rDNA amplification

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The Mre11-Rad50-Xrs2 complex is important for DSB repair at arrested forks

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Homologous recombination is dispensable for DSB repair at arrested forks

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Ctf4 is a key regulator in suppressing DSB end resection

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Suppression of end resection is important to protect DSBs from genome rearrangements

Sasaki & Kobayashi, 2017, Molecular Cell 66, 533–545 May 18, 2017 ª 2017 Elsevier Inc. http://dx.doi.org/10.1016/j.molcel.2017.04.020

Sasaki and Kobayashi demonstrate that DNA double-strand breaks (DSBs) at arrested forks are normally repaired by homologous recombination-independent pathways. They reveal Ctf4 as a key player in preventing chromosome rearrangements by protecting arrested forks from breakage and end resection. DSB repair at arrested forks is regulated differently from replicationindependent DSBs.

Molecular Cell

Article Ctf4 Prevents Genome Rearrangements by Suppressing DNA Double-Strand Break Formation and Its End Resection at Arrested Replication Forks Mariko Sasaki1 and Takehiko Kobayashi1,2,* 1Laboratory of Genome Regeneration, Institute of Molecular and Cellular Biosciences, University of Tokyo, 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-0032, Japan 2Lead Contact *Correspondence: [email protected] http://dx.doi.org/10.1016/j.molcel.2017.04.020

SUMMARY

Arrested replication forks lead to DNA double-strand breaks (DSBs), which are a major source of genome rearrangements. Yet DSB repair in the context of broken forks remains poorly understood. Here we demonstrate that DSBs that are formed at arrested forks in the budding yeast ribosomal RNA gene (rDNA) locus are normally repaired by pathways dependent on the Mre11-Rad50-Xrs2 complex but independent of HR. HR is also dispensable for DSB repair at stalled forks at tRNA genes. In contrast, in cells lacking the core replisome component Ctf4, DSBs are formed more frequently, and these DSBs undergo end resection and HR-mediated repair that is prone to rDNA hyper-amplification; this highlights Ctf4 as a key regulator of DSB end resection at arrested forks. End resection also occurs during physiological rDNA amplification even in the presence of Ctf4. Suppression of end resection is thus important for protecting DSBs at arrested forks from chromosome rearrangements.

INTRODUCTION Accurate and complete replication of the genome is essential for the maintenance of genome stability. DNA replication, however, is constantly challenged by various types of replication stress, which stall or arrest replication forks (Branzei and Foiani, 2010; Mirkin and Mirkin, 2007). Replication stress can be induced by exogenous sources such as DNA damaging or crosslinking agents as well as by hydroxyurea, which reduces the nucleotide pool. These clastogens have been used for conventional chemotherapy, which takes advantage of the detrimental consequence of excessive replication stress induced by these drugs. Replication forks also encounter endogenous fork-pausing elements, such as non-histone proteins tightly associated with the template DNA, DNA secondary structures, and transcription machineries, which impact genome stability even in unchallenged conditions (Branzei and Foiani, 2010; Mirkin and Mirkin, 2007).

The replisome is a multi-protein complex that carries out DNA synthesis at the replication fork (O’Donnell and Li, 2016). The Cdc45-MCM-GINS (CMG) complex is thought to be the replicative helicase in eukaryotes. The leading strand is synthesized by DNA polymerase ε, whereas synthesis of the lagging strand is initiated by the DNA-polymerase-a-primase complex and completed by DNA polymerase d. Other factors in the replisome ensure efficient DNA replication. Ctf4 in Saccharomyces cerevisiae (Mcl1 in Schizosaccharomyces pombe [Tsutsui et al., 2005; Williams and McIntosh, 2002] and AND-1 in mammals [Im et al., 2009; Zhu et al., 2007]) binds as a homotrimer to a subunit of GINS and to DNA polymerase a and has been suggested to facilitate coupling of the CMG helicase to lagging strand synthesis (Gambus et al., 2009; Miles and Formosa, 1992; Simon et al., 2014; Tanaka et al., 2009). Ctf4 is important for errorfree DNA damage tolerance (Fumasoni et al., 2015). Moreover, along with the Chl1 helicase, Ctf4 promotes sister chromatid cohesion accompanied by DNA replication (Borges et al., 2013; Hanna et al., 2001; Kouprina et al., 1992; Samora et al., 2016). Through its ability to interact with numerous proteins, Ctf4 has been suggested to function as a key hub that recruits proteins with diverse functions to the replisome in order to coordinate DNA synthesis with chromosome metabolism (Samora et al., 2016; Simon et al., 2014; Villa et al., 2016). Cells have evolved multiple layers of mechanisms in order to respond properly to replication fork stalling (Branzei and Foiani, 2010). For example, checkpoint factors maintain replisome stability and coordinate the restart of replication or appropriate DNA repair with cell cycle progression. Failure to resume DNA synthesis results in the generation of DNA double-strand breaks (DSBs)—a major source of the genome rearrangements that are a hallmark of cancer cells, cause human genomic disorders, and influence genome diversity (Branzei and Foiani, 2010; Liu et al., 2012; Sasaki et al., 2010). Homologous recombination (HR) is important for repairing replication-independent DSBs such as meiotic DSBs and DSBs generated in the G2/M phase of the mitotic cell cycle (Krogh and Symington, 2004). Therefore, it has long been assumed that HR is required for the repair of DSBs at arrested forks. However, there is little direct evidence to address whether cells use HR as the main pathway to repair broken forks. This is largely due to a lack of experimental systems that can detect physiologically relevant DSBs forming at arrested forks at a defined site at high frequency. Molecular Cell 66, 533–545, May 18, 2017 ª 2017 Elsevier Inc. 533

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Figure 1. Ctf4 Prevents rDNA Hyper-amplification upon Replication Fork Arrest (A) Fob1-dependent fork arrest and DSB formation at the RFB. Top: organization of an rDNA repeat unit; bottom, DNA replication in rDNA. 35S, 35S rRNA; 5S, 5S rRNA; ARS, autonomously replicating sequence; RFB, replication fork barrier; Bg, BglII recognition site; pink bar, the position of the probe used for Southern blotting. (B) PFGE analysis of the size of chr XII in independent haploid clones isolated from the ctf4D fob1 heterozygous diploid strain. DNA was stained with ethidium bromide. Sizes of DNA markers and approximate numbers of rDNA repeats are indicated. (C) Quantitative PCR-based rDNA copy number assays. The amount of rDNA relative to a single-copy MCM2 gene was determined in the indicated strains (bars show mean ± SEM). One-way ANOVA was used for multiple comparisons (*p < 0.05 between WT and mutants and between ctf4D and ctf4D fob1/tof1D/rrm3D mutants). (D) Cellular growth of ctf4D and ctf4D fob1 mutants isolated by tetrad dissection of the ctf4D fob1 heterozygous diploid strain. The ctf4D and ctf4D fob1 colonies are indicated. See also Figure S1.

The budding yeast ribosomal DNA (rDNA) locus represents a unique region among eukaryotic genomes in which DSBs arising from replication fork arrest can be detected by Southern blotting (Burkhalter and Sogo, 2004; Kobayashi et al., 2004; Weitao et al., 2003). The rDNA region contains a tandem array of
150 copies of 9.1-kb rDNA sequence and is located at a single locus on chr XII. Each repeat contains 35S and 5S rRNA transcription units as well as an origin of DNA replication and a replication fork barrier (RFB) sequence (Figure 1A). After DNA replication is initiated, progression of the fork motion against the 35S rDNA is blocked by Fob1 protein bound to the RFB site (Brewer et al., 1992; Kobayashi, 2003; Kobayashi et al., 1992; Kobayashi and Horiuchi, 1996), which leads to generation of a one-ended DSB (Burkhalter and Sogo, 2004; Kobayashi et al., 2004; Weitao et al., 2003; Figure 1A). DSB repair can be accompanied by a gain or a loss of rDNA repeats and by the production of extrachromosomal rDNA circles (ERCs) in a manner that is dependent mainly on

534 Molecular Cell 66, 533–545, May 18, 2017

the HR protein Rad52 (Ivessa et al., 2000; Kobayashi et al., 2004). How these DSBs are repaired, however, remains unknown (Fritsch et al., 2010). We previously conducted a genome-wide screen to search for mutants that carry abnormal rDNA copy numbers; this was based on the hypothesis that mutants defective in the proper response to replication fork arrest or to DSBs undergo aberrant changes in rDNA repeat number (Kobayashi and Sasaki, 2017; Saka et al., 2016). We discovered that the mutant lacking the CTF4 gene showed hyper-amplified rDNA arrays (Saka et al., 2016). In the present study, we show that Ctf4 plays a pivotal role in preventing rDNA hyper-amplification in response to replication fork arrest. By conducting time course experiments and highly quantitative physical analysis of DSBs and their repair intermediates, we provide evidence that Ctf4 is a key factor in restricting DSB formation and its end resection—and, thus, in preventing DSBs from being repaired by HR. Even in the presence of

Ctf4, DSB end resection occurs in rDNA low-copy strains, which need to undergo rDNA amplification to restore normal rDNA copy numbers. These findings indicate that suppression of end resection is a critical regulatory step that protects DSBs at arrested forks from being repaired by HR pathways that are prone to genome rearrangements. HR is also dispensable for repair of DSBs formed at stalled forks at tRNA genes. Our findings reveal that repair of DSBs at arrested forks is regulated differently from replication-independent DSBs. RESULTS Ctf4 Prevents rDNA Hyper-amplification in Response to Replication Fork Arrest We first examined whether rDNA hyper-amplification in ctf4D occurs in response to Fob1-mediated replication fork arrest at the RFB. To this end, we isolated haploid clones from a diploid strain heterozygous for ctf4D and fob1 by tetrad dissection and compared their rDNA repeat numbers by pulsed-field gel electrophoresis (PFGE) and qPCR-based copy number assay. The size of chr XII was larger in ctf4D clones than in wild-type (WT) clones (Figure 1B). The copy number assay showed that ctf4D clones had, on average, twice as many rDNA repeats as did WT clones (Figure 1C), demonstrating that the absence of Ctf4 results in rDNA hyper-amplification; this is consistent with our previous finding (Saka et al., 2016). The rDNA expansion in ctf4D was suppressed when fork stalling was inhibited by a fob1 mutation (Kobayashi and Horiuchi, 1996) or by deletion of TOF1 (Mohanty et al., 2006; Tourrie`re et al., 2005; Figures 1B, 1C, and S1A). Based on the comparison of the colony sizes between ctf4D and ctf4D fob1, growth defects of ctf4D appeared partially alleviated by a fob1 mutation (Figure 1D), although we did not find a statistically significant difference in the doubling times of single and double mutants of ctf4D with fob1 (data not shown). Conversely, rDNA amplification was exacerbated when fork arrest was increased by inactivation of the 50 -30 helicase Rrm3, which facilitates progression of stalled forks across protein-DNA complexes (Ivessa et al., 2000, 2003; Mohanty et al., 2006; Figures 1C, S1B, and S1C). These findings show that Ctf4 plays an important role in preventing rDNA hyper-amplification specifically in the context of replication fork arrest. Arrested Forks at the RFB Are Broken More Frequently in the Absence of Ctf4 An arrested fork can collapse and be converted into a DSB, a major initiator of HR-mediated genome rearrangements. Aberrant rDNA amplification might thus be the consequence of an increased number of recombination-inducing DSBs and/or their rearrangement-prone repair. To test these possibilities, we determined the level and kinetics of DSB formation and repair in the presence or absence of Ctf4. We conducted time course experiments during synchronous S phase progression in cells carrying the FOB1 gene under the GAL1 promoter followed by direct detection of arrested fork intermediates and DSBs by Southern blotting (Figures 1A and S2A–S2C). In these experiments, GAL-FOB1 ctf4D cells had a rDNA copy number similar to that of GAL-FOB1 cells (Figure S2D), because

we kept FOB1 expression turned off prior to time courses (Figure S2A). Both GAL-FOB1 CTF4 and ctf4D cells exhibited Fob1-dependent, S-phase-specific replication-fork blockage and DSB formation at the RFB (Figures 2A, 2B, S2E, and S2F). In the CTF4 background, arrested forks and DSBs accounted for
10% and 0.6% of the total rDNA signals at the peak time, respectively (Figure 2C). Considering that arrested forks and DSBs had a different number of probe-annealing sites in our Southern blot assay (Figure 1A), we estimated that approximately one in ten arrested forks had been converted into oneended DSBs (Figure 2D; STAR Methods). By taking into account the proportion of arrested forks and DSBs in all detectable rDNA molecules as well as the average number of rDNA repeats carried, we estimated that each cell had, on average, eight arrested forks and one DSB at the peak time (STAR Methods). It should be noted that, in addition to quantifying the arrested-fork and DSB signals as a proportion of the total radioactive signal, we quantified them relative to the 3.0-kb restriction fragment containing the telomere-proximal rDNA and its adjacent non-rDNA fragment, referred to as the terminal fragment, with the same conclusion regardless of the quantification method used (Figures 2C and S2G; data not shown). Our estimation of the numbers of arrested forks and DSBs is smaller than a previous estimation made from two-dimensional (2D) gel electrophoresis analysis of genomic DNA isolated from asynchronously growing cultures (Fritsch et al., 2010). Nonetheless, because a single, unrepaired DSB is sufficient to induce cell death (Weiffenbach and Haber, 1981), DSB(s) at the RFB place a substantial burden on cell viability in unchallenged conditions. In ctf4D cells, arrested forks were detected at a timing and level similar to that found in WT cells (Figure 2C). However, DSBs were seen at a
1.6-fold higher steady-state level in ctf4D cells than in CTF4 cells (Figure 2C). The kinetics of the disappearance of DSBs was similar between CTF4 and ctf4D (Figure 2E), indicating that the increase in DSBs did not result from a transient accumulation of unrepaired DSBs caused by a delay in the initiation of DSB repair. Instead, DSBs were most likely formed at a higher frequency at the RFB in the absence of Ctf4. Ctf4 Restricts End Resection of DSBs at Arrested Forks In our Southern blot assay, we noticed that cells lacking Ctf4 displayed a 13-fold increase in steady-state levels of previously unidentified DNA species migrating faster than DSBs, which were barely seen in the presence of Ctf4 (Figure 2F and lower panels in Figures 2A and 2B). To reveal their identities, we performed 2D neutral/neutral gel electrophoresis, where the first dimension separates DNA molecules according to their mass and the second dimension resolves molecules according to their mass and shape. In contrast to DSBs migrating along the arc of linear molecules, the faster-migrating DNA species were seen below this arc (Figure S3A), which was indicative of the presence of a single-stranded DNA region. We then determined the strand compositions of these DNA molecules by 2D neutral/denaturing gel analysis, in which the component strands were separated in the second dimension; this was followed by Southern blotting with strand-specific

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Figure 2. Absence of Ctf4 Causes DSBs and Their End Resection to Occur More Frequently

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(A and B) Time-course experiments and physical assays to monitor arrested forks and DSBs in GALFOB1 (A) and GAL-FOB1 ctf4D (B). Time-course experiments were performed as described in Figure S2A. Genomic DNA was prepared in agarose plugs, digested with BglII, separated by conventional agarose gel electrophoresis, and analyzed by Southern blotting with the rDNA probe indicated in Figure 1A. Open circle indicates the terminal fragment. Asterisks indicate cross-hybridizing bands. Bands corresponding to arrested forks, linear fragments, DSBs, and resected DSBs are indicated. Lower panels show a more exposed contrast image of the phosphorimager signal around DSBs marked by dashed lines. (C) Levels of arrested forks and DSBs quantified as percent of total hybridization signal in the lane (bars show mean ± SEM). (D) Proportion of DSBs relative to arrested forks. The value at t = 15 was not included due to the difficulty of accurately measuring low levels of arrested forks and DSBs at this time point. Bars show mean ± SEM. (E) DSBs expressed as percent of maximum values. Bars show mean ± SEM. (F) Level of resected DSBs normalized to the terminal fragment. Bars show mean ± SEM. (G) DSBs and resected DSBs expressed as percent of maximum values in one representative culture of GAL-FOB1 ctf4D. See also Figures S2 and S3.

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end resection occurs in S phase (Doksani et al., 2009; Zierhut and Diffley, 2008). In contrast, we found that DSBs at the RFB are rarely resected in WT cells (Figures 2A, 2F, S3A, and S3B). Thus, the significance of end resection for DSB repair remains unclear. To address this question, we investigated whether canonical end-resection factors are required for DSB repair in the CTF4 background. The Mre11-Rad50-Xrs2 (MRX) complex (Xrs2 is a homolog of Nbs1 in mammals) is central to the DNA damage response (Stracker and Petrini, 2011). In the absence of any component of the MRX complex, a majority of DSBs at the RFB remained unrepaired at the time when most cells had completed bulk DNA synthesis and arrested forks had almost disappeared (Figures 3A, 3C, and S4A–S4D), demonstrating that the MRX complex is required for DSB repair at the RFB. The MRX complex, along with Sae2, functions to initiate 50 -30 DSB end resection through the nuclease activity of Mre11 (Mimitou and Symington, 2009; Stracker and Petrini, 2011). It also plays a structural role in DSB repair by keeping DSB ends or sister chromatids close together (Mimitou and Symington, 2009; Stracker and Petrini, 2011). To examine which of these functions is important for DSB repair, we performed DSB assays in the nuclease-deficient mre11 mutant (Moreau et al., 1999) and in sae2D; however, both mutants were proficient in DSB repair (Figures 3D, 3F, S4E, S4F, and S4H), indicating that the catalytic role of the MRX complex is dispensable for DSB repair. These results support the previous finding that disrupting MRX complex formation, but not the nuclease activity of Mre11, renders cells sensitive to Fob1 overproduction (Bentsen et al., 2013). Exo1 and Sgs1-Dna2 act redundantly to carry out extensive resection (Mimitou and Symington, 2009). Absence of both Exo1 and Sgs1 leads to severe defects in DSB end resection and recombination-mediated repair of replication-independent DSBs (Mimitou and Symington, 2008). DSB repair at the RFB, however, was found to be proficient in cells lacking both Exo1 and Sgs1 (Figures S4I and S4L). Collectively, these results suggest that DSB end resection is dispensable for DSB repair at the RFB in CTF4 cells. Cells Lacking Ctf4 Depend on DSB End Resection for Repair The MRX complex was also important for DSB repair in cells lacking Ctf4 (Figures 3B and 3C), although the nuclease activity of Mre11 was not required for DSB repair (Figures S4E–S4G). It should be noted that DSB levels were higher in ctf4D mre11D than in CTF4 mre11D (Figure 3C), supporting our findings that DSBs are formed more frequently in the absence of Ctf4 (Figure 2C). In marked contrast to DSB repair in the CTF4 sae2D mutant, DSB repair was severely delayed in the ctf4D sae2D double mutant (Figures 3D–3F and S4H). Resected DSBs were almost undetectable in ctf4D mre11D and ctf4D sae2D mutants (Figures 3B, 3E, and 3G). These results demonstrate that both the MRX complex and Sae2 are necessary for DSB end resection and for DSB repair in the ctf4D background. As compared with the ctf4D mutant, the proportion of resected DSBs relative to intact DSBs was reduced by
4 and 2-fold in ctf4D cells lacking Exo1 and Sgs1, respectively (Figures 3G, S4J, S4K, and S4M). The kinetics of DSB repair could not be examined in ctf4D exo1D sgs1D cells due to their severe growth

defects. Therefore, in the absence of Ctf4, DSB ends undergo end resection in a manner that is dependent on canonical end resection factors. Our results revealed that the presence or absence of Ctf4 determines the fate of DSBs—in other words, whether they undergo end resection for repair or not. Cells Lacking Ctf4 Depend on Rad52 for DSB Repair and Cellular Viability We next sought to determine whether DSB repair is dependent on homology-directed repair. To this end, we performed DSB assays in mutants lacking two key HR factors, Rad51 and Rad52. Although these cells displayed severe defects in the repair of replication-independent DSBs (Krogh and Symington, 2004), they were both proficient in DSB repair at the RFB (Figures 4A, S5A, S5C and S5D). These results support previous findings that overproduction of Fob1 does not compromise growth of cells lacking Rad52 (Bentsen et al., 2013; Calzada et al., 2005). Thus, Rad52 and Rad51 are not required for DSB repair at the RFB in the presence of Ctf4. In contrast, cells lacking both Ctf4 and Rad52 displayed DSB repair defects. In the ctf4D rad52D mutant, DSBs disappeared with a similar timing to those in the rad52D mutant (Figures 4B and S5A). The amount of resected DSBs was reduced in the double mutants as compared with the single mutants (Figure S5B). However, we noticed the appearance above the linear fragments of two additional signals that were rarely seen in the rad52D mutant (Figures 4A and 4B). On the basis of their size and the timing of their appearance (Figures 4B and 4C), we concluded that they corresponded to DSBs that have undergone extensive resection past the adjacent BglII recognition site, and thus have become resistant to restriction digestion due to the exposure of long ssDNA regions (Figure 4B, right). The level of resected DSBs combined was
22-fold higher in ctf4D rad52D than in rad52D (Figure 4D). Hyper-resected DSBs were also seen in the ctf4D rad51D mutant at a level
9-fold higher than in rad51D (Figures S5C and S5E). Cells lacking Ctf4 and Rad51 were able to enter into the next cell cycle at later time points (Figures S5C and S5D), whereas the ctf4D rad52D mutant displayed further delays in entry into the next cycle (Figure S5A). These results imply that Rad52 is more important than Rad51 for engagement of resected DSBs for repair at the RFB in ctf4D cells. In addition to its function as a Rad51 mediator, Rad52 has strand-annealing activity in vitro and carries out single-strand annealing (SSA)-mediated DSB repair independently of Rad51 (Krogh and Symington, 2004). To test whether Rad52-dependent SSA pathways are involved in DSB repair, we performed DSB assays in ctf4D mutants lacking SSA factors such as Saw1, Rad1, and Rad10; these mutants were proficient in DSB repair, did not accumulate extensively resected DSBs, and were able to enter into the next cell cycle (Figures S5F–S5H; data not shown). Thus, SSA is unlikely to be the major pathway of DSB repair at the RFB. Cells lacking Ctf4 display synthetic growth defects with rad52D, but not with rad51D (Fumasoni et al., 2015; Kouprina et al., 1992). Those findings, along with our results, raise the possibility that growth defects in the ctf4D rad52D mutant may arise from their defects in DSB repair at the RFB. To test this possibility, we examined the genetic interaction of ctf4D with rad52D in

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Figure 3. Requirements of the MRX Complex and Sae2 for DSB Repair in CTF4 and ctf4D Cells (A and B) Time-course experiments and physical assays in GAL-FOB1 mre11D (A) and ctf4D mre11D (B). (C) Level of DSBs in GAL-FOB1 mre11D and ctf4D mre11D quantified as percent of total hybridization signal in the lane. (D and E) Time-course experiments and physical assays in GAL-FOB1 sae2D (D) and ctf4D sae2D (E). (F) Level of DSBs in GAL-FOB1 sae2D and ctf4D sae2D quantified as percent of total hybridization signal in the lane. (G) Proportion of maximum values of resected DSBs relative to maximum DSB levels in the indicated strains. Each point represents the value of independent cultures. Bars are mean ± SD. See also Figure S4. Open circles, asterisks, and lower panels in (A), (B), (D), and (E) are as described in Figure 2A. Arrested forks, DSBs, and resected DSBs are indicated as in Figure 2A. CTF4 and ctf4D indicate genomic DNA samples collected from GAL-FOB1 and GAL-FOB1 ctf4D time courses, respectively. Black and red horizontal lines in (C) and (F) indicate the maximum DSB level seen during the time courses in GAL-FOB1 and GAL-FOB1 ctf4D cells, respectively; bars in (C) and (F) show mean ± SEM.

both FOB1 and fob1 backgrounds by comparing the colony sizes of haploid clones isolated from a diploid strain heterozygous for ctf4D, rad52D, and fob1 (Figure 4E). The ctf4D rad52D double mutants exhibited synthetic growth defects because they formed colonies of a size substantially smaller than would be expected if these genes did not show genetic in-

538 Molecular Cell 66, 533–545, May 18, 2017

teractions (70-fold [observed] versus
9-fold [expected]) (Figure 4E). In contrast, ctf4D and rad52D exhibited only additive effects on growth in the fob1 background (Figure 4E). The ctf4D rad51D double mutants did not show synthetic growth defects in either the FOB1 or the fob1 background (Figure S5I). From these results, we infer that cellular growth of ctf4D cells is

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Figure 4. Cells Lacking Ctf4 Depend on Rad52 for Engagement of Resected DSBs for Repair (A and B) Time-course experiments and physical assays in GAL-FOB1 rad52D (A) and ctf4D rad52D (B). Open circles, asterisks, and lower panels are as described in Figure 2A. Arrested forks, DSBs, and resected DSBs are indicated as in Figure 2A. Extensively resected DSBs are indicated by arrows with a triangle or square head. The right panel shows the probe position and strand compositions of DSBs and DSBs that are resected at varying degrees. (C) DSBs and DSBs resected at varying degrees expressed as percent of maximum values in one representative culture of GAL-FOB1 ctf4D rad52D. (D) Summed level of DSBs that are resected at varying degrees in the indicated strains (bars show mean ± SEM). (E) Tetrad dissection of the diploid strain heterozygous for fob1, ctf4D, and rad52D. Each point represents the size of independent colonies of the indicated genotypes. Bars represent mean ± SD. One-way ANOVA was used for multiple comparisons between WT and other mutants. p values and fold difference relative to WT colonies are indicated. See also Figure S5.

dependent on RAD52, which is needed mainly to repair DSBs formed at the RFB. DSB End Resection Occurs During Physiological rDNA Amplification in rDNA Low-Copy Strains DSB end resection at the RFB may represent a pathological situation that is evident only when Ctf4 is absent. Alternatively, it may occur even in the presence of Ctf4 when rDNA copy number changes are required. Previous studies demonstrated that absence of Rpa135, an essential subunit of RNA polymerase I responsible for 35S rDNA transcription, causes a reduction

of rDNA repeats to almost half the normal number (Kobayashi et al., 1998). When RPA135 is complemented back into cells, cells gradually undergo rDNA amplification and eventually reach the normal rDNA repeat number in a manner dependent on Fob1 and Rad52 (Kobayashi et al., 1998). Based on the requirement of Rad52 for this amplification, we hypothesized that, in rDNA low-copy strains, DSBs undergo end resection to induce HR. We performed DSB assays in WT cells carrying different rDNA copy numbers (Figure 5A), ranging from
20 to 110 (Ide et al., 2010). The proportion of DSBs arising from arrested forks was highest in the 20-copy strain and decreased as the rDNA copy

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Figure 5. DSB End Resection Occurs in rDNA Low-Copy Strains (A) Time course experiments and physical assays in GAL-FOB1 cells carrying the indicated number of rDNA repeats. Open circles, asterisks, and lower panels are as described in Figure 2A. Arrested forks, DSBs, and resected DSBs are indicated as in Figure 2A. (B) Proportion of DSBs relative to arrested forks (bars show mean ± SEM). (C) Proportion of maximum values of resected DSBs relative to maximum DSB levels in the indicated strains. Each point represents the value of independent cultures. Bars show mean ± SD.

number increased (Figure 5B), implying that arrested forks are more fragile in the rDNA low-copy strain. Importantly, we observed that the amount of resected DSBs generated from DSBs was highest in the 20-copy strain and was inversely correlated with rDNA copy number (Figure 5C). Thus, even in the presence of Ctf4, DSB end resection was induced in rDNA low-copy strains that were to undergo recombination-dependent rDNA amplification. DSB end resection occurs less frequently as the necessity to undergo rDNA amplification is lessened after copy number is restored, pointing to the existence of an active process that suppresses end resection in WT cells that carry the normal rDNA copy number. These findings indicate that end resection is a key regulatory step to control rDNA copy numbers. DSBs at Stalled Forks at tRNA Genes Are Repaired by HR-Independent Pathways The budding yeast genome contains an estimated 1,400 natural replication fork pause sites (Ivessa et al., 2003). These sites include tRNA genes where nonhistone protein-DNA complexes can form, and fork progression past them is promoted by the Rrm3 helicase similarly to the rDNA RFB. We sought to examine whether HR is required for DSB repair at these natural pause sites. The levels of fork stalling at these sites are extremely low in the WT background owing to the action of Rrm3 (Ivessa et al., 2003); therefore, we used the rrm3D background to enhance fork stalling and DSB formation (if formed at all). We performed time courses and physical assays in three genomic regions on chr VI (Figures 6A–6C), matching the regions that were examined in the previous study using 2D gel electrophoresis (Ivessa et al., 2003).

540 Molecular Cell 66, 533–545, May 18, 2017

First, we detected S-phase-specific fork stalling, consistent with the previous findings (Ivessa et al., 2003; Figure 6). Second, we detected DSBs by probe 1; their sizes were suggestive of DSB formation at the tRNA genes (Figures 6A–6C, 6D, 6G, and 6I). DSBs at tP(UGG)F were detected at a level above the limit of detection by Southern blotting (Figure 6E), whereas DSBs at the other two sites were too low to quantify accurately. These DSBs were detected only by probe 1 and not by probe 2 (Figures 6D and 6F–6J), suggesting that they represented one-ended DSBs formed at stalled forks. Lastly, DSBs in these regions disappeared at later time points, regardless of the presence or absence of Rad52 (Figure 6). Collectively, these results suggest that DSBs formed at stalled forks at non-rDNA sites are also repaired by Rad52-independent pathways. DISCUSSION We have revealed that DSBs that form as a consequence of replication fork arrest at the RFB rarely undergo end resection. We identified two situations where DSB end resection is induced. First, DSB ends are resected in the absence of Ctf4, and subsequent DSB repair results in aberrant rDNA amplification. Second, end resection is enhanced in rDNA low-copy strains, where rDNA amplification is required to restore normal rDNA repeat numbers. These results have enabled us to propose a model of DSB repair at an arrested fork (Figure 7). Movement of the CMG helicase is blocked by Fob1 protein bound to the RFB, leading to stalling of the replisome (Calzada et al., 2005). We demonstrated that Ctf4 affects the fate of replication fork arrest by preventing DSB formation (Figure 2C) and

Figure 6. Rad52 Is Dispensable for DSB Repair at Stalled Forks at tRNA Genes

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by suppressing DSB end resection (Figure 2F). Ctf4 connects the DNA polymerase a–primase complex to the CMG helicase, which is linked to DNA polymerase ε (Gambus et al., 2009; Samora et al., 2016; Simon et al., 2014; Tanaka et al., 2009; Villa et al., 2016). Previous studies demonstrated that replisome components associated with hydroxyurea-induced stalled forks are reduced in ctf4D as compared with WT (Tanaka et al., 2009; Tittel-Elmer et al., 2009). Ctf4 may maintain the association of replisome factors with the arrested fork. The large, multi-protein complexes bound to the arrested fork could suppress DSB formation and block access to DSB ends by end resection factors (Figure 7, left). In a previous study, Fritsch et al. (2010) investigated DSB levels at the RFB in DSB-repair-deficient mutants such as rad52 and dnl4 using 2D gel electrophoresis analysis and demonstrated that none of these mutants altered DSB levels (Fritsch et al., 2010). They raised the possibility that most DSBs detected in WT represent DSBs that are formed in vitro and generated when forks are arrested at the RFB, around which there exist nicks on the parental, unwound DNA strands; the melting of these strands during DNA preparation might generate

(A–C) Organization of the regions around (A) tP(UGG)F (the complementary strand), (B) tA(AGC) F, and (C) tY(GUA)F1 (the complementary strand) on chr VI. tRNA genes, restriction sites, probes used for Southern blotting, and expected sizes of linear and DSB fragments are indicated. (D and F–J) Time-course experiments and physical assays in the indicated strains around tP(UGG)F (the complementary strand) in (D) and (F), tA(AGC)F in (G) and (H), and tY(GUA)F1 (the complementary strand) in (I) and (J) on chr VI. Southern blotting was performed with probe 1 in (D), (G), and (I) and with probe 2 in (F), (H), and (J). Arrested forks and DSBs are indicated as in Figure 2A. Triangles indicate DNA fragments partially digested by the restriction enzyme. Asterisks indicate cross-hybridizing bands. (E) Levels of arrested forks and DSBs at tP(UGG)F quantified as percent of total hybridization signal in the lane (bars show range).

one-ended DSBs (Fritsch et al., 2010). The reason why DSBs were resistant to end resection in WT in our study can be explained by the possibility that these DSBs are artifactual DSBs and thus blind to end resection. Alternatively, end resection is actively suppressed in WT. To distinguish between these possibilities, it is important to identify mutations that cause DSB repair defects. We showed that DSB repair was severely delayed in mre11D mutants (Figure 3), although Bentsen et al. (2013) did not observe an accumulation of DSBs in mre11D mutants. The differences between the studies remain unclear. In our study, however, the mutants lacking any component of the MRX complex showed defects in repair of most DSBs (Figure S4). Thus, the MRX complex is most likely important for DSB repair. While we cannot definitively exclude the possibility that DSBs seen in WT are a mixture of DSBs formed in vitro and in vivo, the identification of mrx mutants deficient in DSB repair provides strong evidence that most, if not all, DSBs are formed in vivo. Thus, we favor the idea that DSB end resection is actively suppressed in WT. The catalytic role of the MRX complex in DSB end resection is dispensable for DSB repair at the RFB, which is in contrast to its function in canonical DSB repair pathways (Figure S4). A previous study suggests that the MRX complex is important for stabilizing replisome components at the hydroxyurea-induced stalled forks through tethering of sister chromatids (Tittel-Elmer et al., 2009). Although it remains to be determined how the MRX complex acts during DSB repair at arrested forks, this structural role might be important for keeping broken forks close together for DSB repair. To our surprise, DSBs at the RFB were found to be repaired by HR-independent pathways. In a previous study, the bacterial

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Figure 7. Model of the Repair Pathways of DSBs at Arrested Forks at the RFB Ctf4 may keep replisome factors bound to the arrested forks through its ability to bind GINS and DNA polymerase a. The presence of large protein complexes prevents access to DNA ends by factors responsible for DSB formation and end resection. Replisomes at the broken forks may be competent to repair DSBs independently of HR pathways, possibly by template switching and subsequent DNA synthesis. The MRX complex facilitates DSB repair, possibly by allowing the broken forks to come into close proximity. In the absence of Ctf4, replisome factors fall off from the broken forks. DSB repair now becomes dependent on DSB end resection and HR. DSB end resection also occurs during physiological rDNA amplification in Ctf4-expressing low-rDNA-copy strains. Ctf4 may have an additional role during DSB repair by suppressing the usage of intrachromosomal rDNA repeats or free ERCs as a template, which can lead to rolling-circle DNA replication.

Tus/Ter system was introduced into the budding yeast genome to induce site-specific fork stalling (Larsen et al., 2014). Neither the extent or timing of fork stalling nor the cell cycle progression

542 Molecular Cell 66, 533–545, May 18, 2017

was compromised in rad50, rad51, or rad52 strains, indicating that HR factors are not required to overcome fork stalling at Tus/Ter sites. This system has also been introduced into mouse

embryonic stem cells to induce fork stalling within the recombination reporter (Willis et al., 2014). Fork stalling resulted in the generation of HR-mediated repair products, pointing to the role of HR in response to fork stalling, which might seem contradictory to our findings and the study of Larsen et al. (2014). However, it remains to be determined whether HR is induced by DSBs formed at stalled forks or during the restart of stalled forks independent of DSBs. Furthermore, because this system is designed to detect the outcomes of HR and not those of other repair pathways, findings by Willis et al. (2014) suggest that HR can occur in response to fork stalling, but do not necessarily exclude the possibility that HR is not the main pathway for DSB repair at stalled forks. The rDNA RFB sequence was integrated near early-firing origins on chr III (Calzada et al., 2005). Induction of fork stalling at this non-rDNA locus by FOB1 expression did not compromise the viability of rad52D cells. We showed that Rad52 was not required for repair of DSBs formed at tRNA genes (Figure 6). It is important to determine in future studies whether these DSBs also undergo end resection in the absence of Ctf4 and, if so, whether their repair depends on Rad52. Therefore, HR-independent DSB repair is unlikely to be unique to the rDNA region. Our findings indicate that repair pathways exist that are specific to DSBs formed at arrested forks. The mechanisms of DSB repair remain to be identified in future work; however, any of the replisome components associated with DSBs via Ctf4 and the MRX complex might be competent to restart DNA synthesis beyond the break, possibly by HR-independent template switching (Figure 7; Houseley and Tollervey, 2011). When cells lack Ctf4, DSB repair becomes dependent on end resection and HR. In the absence of Ctf4, replisome components may be destabilized and fall off from broken ends (Figure 7). DSBs that are no longer bound by replisome factors might become accessible to end processing factors, and their repair might rely on Rad52-mediated recombination reactions (Figure 4). Due to the repetitive nature of the rDNA region, DSB ends have several template options to invade or to anneal to: for example, the repeat on the replicated arms of sister chromatids that lies at an equal position, the repeat at an unequal position relative to DSBs, the repeat on the same chromosome, and also free ERCs (Figure 7). The latter three options can lead to changes in rDNA copy number. By restricting DSB end resection, Ctf4 may limit HR-dependent DSB repair, which is prone to rDNA rearrangements. Because the rate of rDNA amplification in ctf4D cells (Figures 1B and 1C) is much greater than that found during rDNA amplification from low-copy strains (Kobayashi et al., 1998), Ctf4 may have an additional role in restricting the use of repair templates that are prone to processive rDNA amplification (Figure 7). DSB end resection also occurs in rDNA low-copy strains, which undergo rDNA amplification to restore the normal repeat number (Kobayashi et al., 1998), even in the presence of Ctf4 (Figure 5). The mechanism that enhances DSB end resection in these strains might also be the disassembly of replisome factors from broken forks, much like the proposed mechanism in cells lacking Ctf4 (Figure 7). Alternatively, the chromatin environment might influence DSB end resection. In strains with normal rDNA copy numbers, the histone deacetylase Sir2 acts to repress the bidirectional, non-coding RNA promoter E-pro (Kobayashi and

Ganley, 2005). In rDNA low-copy strains, in contrast, Sir2-mediated silencing is weakened, and transcript levels of non-coding RNAs are elevated (Kobayashi and Ganley, 2005). It is possible that the chromatin environment around DSBs might be more open in rDNA low-copy strains than in normal-copy strains, which may favor DSB end resection. Our findings suggest that regulation of end resection of DSBs at the arrested fork is physiologically important in maintaining the proper rDNA copy number. HR is often considered to be an error-free DSB repair pathway. However, DSBs formed in or near repetitive elements can undergo non-allelic HR, resulting in genome rearrangements (Liu et al., 2012; Sasaki et al., 2010). Repetitive elements such as retrotransposable elements and tRNAs are known to pause fork progression and hotspots of chromosome rearrangements (Ivessa et al., 2003; Lemoine et al., 2005). Therefore, suppression of end resection might be an important mechanism to minimize the risk of HR-mediated genome rearrangements in response to DSBs at arrested forks, especially when these are generated in or near repetitive regions. DSBs at the arrested forks are distinct from two-ended DSBs generated independently of replication in terms of physical structure, local chromatin context, and the presence or absence of replisome factors bound to DNA. These characteristics might render HR-mediated repair of these DSBs highly prone to complex chromosome rearrangements (Liu et al., 2012). Furthermore, it may explain why DSBs at arrested forks need to be regulated differently from replication-independent DSBs. Our results provide a framework for understanding the repair pathways of replication-associated DSBs formed upon replication fork arrest at the protein-DNA complex. STAR+METHODS Detailed methods are provided in the online version of this paper and include the following: d d d d

d d

KEY RESOURCES TABLE CONTACT FOR REAGENT AND RESOURCE SHARING EXPERIMENTAL MODEL AND SUBJECT DETAILS METHOD DETAILS B Yeast Strains and Culture Methods B Fluorescence-Activated Cell Sorter Analysis B Genomic DNA Preparation B PFGE Analysis B rDNA Copy Number Assays B Southern Blotting B Analysis of Colony Sizes QUANTIFICATION AND STATISTICAL ANALYSIS DATA AND SOFTWARE AVAILABILITY

SUPPLEMENTAL INFORMATION Supplemental Information includes five figures and one table and can be found with this article online at http://dx.doi.org/10.1016/j.molcel.2017.04.020. AUTHOR CONTRIBUTIONS M.S. and T.K. designed the experiments, analyzed the data, and wrote the paper. M.S. performed the experiments.

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ACKNOWLEDGMENTS We thank H. Araki and S. Keeney for discussions and comments on the manuscript. We are grateful to L. Symington and T. Usui for yeast strains. We thank members of the Kobayashi laboratory, especially K. Saka, Y. Akamatsu, and T. Iida, for discussions. This work was supported by a Grant-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT), Japan (23114002, to T.K.), a Grant-in-Aid for JSPS Fellows (24-3076, to M.S.), and a Grant-in-Aid for Young Scientists (B) (15K18581, to M.S.). Received: November 24, 2016 Revised: March 20, 2017 Accepted: April 26, 2017 Published: May 18, 2017

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STAR+METHODS KEY RESOURCES TABLE

REAGENT or RESOURCE

SOURCE

IDENTIFIER

Chemicals, Peptides, and Recombinant Proteins Alpha factor mating pheromone

Zymo Research

Y1001

Pronase E

Sigma

P6911

RNase A

Macherey-Nagel

740505

Proteinase K

Merck Millipore

1.24568

Propidium iodide

Sigma

P4864

Zymolyase 100T

Nacalai

07665-55

SeaPlaque GTG Agarose

Lonza

50111

Seakem LE Agarose

Lonza

50004

Pulsed-Field Certified Agarose

Bio-Rad

1620138

NEBuffer 3.1

New England Biolabs

B7203S

BglII

New England Biolabs

R0144M

CutSmart buffer

New England Biolabs

B7204S

Eco RV-HF

New England Biolabs

R3195M

Bovine Serum Albumin

Nacalai

01859-76

Hybond-XL

GE Healthcare

RPN303S

High Prime DNA Labeling Kit

Roche

11585592001

Qubit dsDNA HS Assay Kits

Thermo Fisher Scientific

Q32854

SYBR Premix Ex TaqII (Tli RNaseH Plus)

TaKaRa

RR820A

This paper

http://dx.doi.org/10.17632/ b66s2sg297.1

Critical Commercial Assays

Deposited Data Mendeley Data dataset Experimental Models: Organisms/Strains S. cerevisiae: Strain background: W303

This paper (see Table S1)

Oligonucleotides Primer: NatNT2-GALL-FOB1 Forward: GGAGAACAATTTAACGATTGTG TGAGTGTGAATTTGTGCTGAGGATAACACGTACGCTGCAGGTCGAC

This paper

N/A

Primer: NatNT2-GALL-FOB1 Reverse: ACCGAGTCATCATCATCAAACA ACACGTCATTGTAACGCGGTTTCGTCATTCCGGGGGGGATCCACTAG

This paper

N/A

Janke et al., 2004

EUROSCARF #P30296

Recombinant DNA pYM-N28 Software and Algorithms ImageJ

https://imagej.nih.gov/ij/

Prism 6 (version 6.0d or 6.0h)

http://www.graphpad.com/ scientificsoftware/prism/

Multi Gauge

FUJIFILM

CONTACT FOR REAGENT AND RESOURCE SHARING Further information and requests for reagents should be directed to and will be fulfilled by the Lead Contact, Takehiko Kobayashi ([email protected]). EXPERIMENTAL MODEL AND SUBJECT DETAILS Yeast strains were derivatives of the laboratory strain W303 and are listed in Table S1.

e1 Molecular Cell 66, 533–545.e1–e5, May 18, 2017

METHOD DETAILS Yeast Strains and Culture Methods Yeast strains used in this study are listed in Table S1. Gene replacement was performed by a one-step gene replacement procedure using standard methods. Strains carrying a truncated form of the GAL1 promoter (GALL) upstream of the FOB1 ORF were constructed by a one-step gene replacement using NatNT2-GALL-FOB1 cassette amplified using primers listed in KEY RESOURCE TABLE and pYM-N28 as a template, as described previously (Janke et al., 2004). Heterozygous gene mutations were introduced into the diploid strain by sequentially replacing one allele of the gene of interest with appropriate selection markers. Correct replacement was confirmed by PCR-based genotyping. The nuclease-deficient mre11-H125N allele (Moreau et al., 1999) was introduced into the GAL1-FOB1 background by crossing LSY716A (a kind gift of L. Symington) to the MATp GAL1-FOB1 strain to generate MSY724. Diploid cells were sporulated by incubating cells in 1% potassium acetate at 30 C for 1–3 days and tetrads were dissected to isolate haploid clones for further analysis. For PFGE and rDNA copy number assays, haploid strains were grown overnight in YP medium (1% yeast extract and 2% peptone) containing 2% glucose at 30 C. To prepare synchronous cultures for time course experiments, cells were grown in 5–10 mL of YP medium containing 2% glucose at 30 C overnight. Cells were inoculated into 100–400 mL of YP medium containing 2% raffinose and one drop of antifoam 204 (Sigma) at an appropriate density to reach
1 3 107 cells/ml the following morning, and grown overnight at 23 C. To arrest cells in the G1 phase of the cell cycle, a-factor (ZYMO RESEARCH) was added to the cultures at a final concentration of 20 nM. Cells were incubated at 23 C for 3–4 hr until > 85% of cells were arrested in G1. Some mutants, especially those deficient in HR pathways, showed inefficient G1 arrest, and only cultures with > 70% of G1-arrested cells were used for the time course experiments. For MSY360 (GAL-FOB1) and MSY375 (GAL-FOB1 ctf4D), the culture was spilt into two flasks. To one culture, 20% galactose was added at a final concentration of 2% to induce FOB1 expression (FOB1 ON). To the other culture, the same volume of 20% raffinose was added (FOB1 OFF). The cultures were further incubated for 30 min at 23 C. Cells were collected by centrifugation, washed twice, and resuspended at
1 3 107 cells/ml in fresh medium (YP + 2% raffinose + 2% galactose (FOB1 ON) or YP + 4% raffinose (FOB1 OFF)) containing 0.1 mg/ml of pronase E (Sigma) and one drop of antifoam 204 (Sigma). Cells were incubated at 23 C, collected at various time points, immediately treated with 0.1% sodium azide, and washed twice with 50 mM EDTA (pH 7.5). For other strains, time course experiments were performed only under the conditions where FOB1 expression was induced. The number of time course experiments performed per GAL-FOB1 strain was as follows: WT (n = 5); ctf4D (n = 5); rrm3D (n = 3); mre11D (n = 3); ctf4D mre11D (n = 3); mre11-nd (n = 3); ctf4D mre11-nd (n = 3); rad50D (n = 4); xrs2D (n = 3); sae2D (n = 3); ctf4D sae2D (n = 3); exo1D sgs1D (n = 2); ctf4D exo1D (n = 3); ctf4D sgs1D (n = 3); rad52D (n = 3); ctf4D rad52D (n = 4); rad51D (n = 3); ctf4D rad51D (n = 3); saw1D (n = 2); ctf4D saw1D (n = 2); WT carrying
20 rDNA copies (n = 3); WT carrying
40 rDNA copies (n = 3); WT carrying
80 rDNA copies (n = 3); and WT carrying
110 rDNA copies (n = 3). To determine the requirement of HR for DSB repair at stalled forks outside the rDNA in Figure 6, we performed time courses and physical assays using rrm3D and rrm3D rad52D in the GAL-FOB1 background for the following reasons. First, because fork stalling at natural pause sites is suppressed by Rrm3 (Ivessa et al., 2003), the level of DSBs arising from such stalled forks is expected to be extremely low in the WT background, especially at the single-copy sequence. Thus, the rrm3D background was used to enhance fork stalling and subsequent DSB formation. Second, the rrm3D rad52D cells showed severe growth defects in the FOB1 background (data not shown); therefore, we used the GAL-FOB1 background to keep FOB1 expression turned off prior to the time courses, which alleviated the severe growth defects of rrm3D rad52D double mutants. Two independent time course experiments were performed per strain. Fluorescence-Activated Cell Sorter Analysis 1 3 107 cells were collected at various time points in the time course experiments described above. Cell cycle analysis was performed as previously described with slight modifications (Tanaka and Diffley, 2002). Briefly, cells were fixed with 70% ethanol at –20 C overnight. Cells were resuspended into 200 mL of 50 mM sodium citrate at pH 7.4 containing 0.25 mg/ml RNaseA (Macherey-Nagel) and incubated for 1 hr at 50 C. After 100 mL of 50 mM sodium citrate at pH 7.4 containing 1 mg/ml Proteinase K (Merck Millipore) were added, cells were incubated for 1 hr at 50 C. After 300 mL of 50 mM sodium citrate at pH 7.4 containing 4 mg/mml propidium iodide (Sigma) were added, cells were sonicated and further diluted with 50 mM sodium citrate at pH 7.4 containing 4 mg/mml propidium iodide (Sigma) when necessary and were analyzed using a BD Accuri C6 Flow Cytometer (BD Bioscience). Genomic DNA Preparation For PFGE, physical assays and 2D gel analyses, genomic DNA was prepared in low melting temperature agarose plugs as described previously (Murakami et al., 2009) with slight modification. For PFGE, cells were collected and washed once with 50 mM EDTA (pH 7.5) (5 3 107 cells per plug). For physical assays and 2D gel analyses, cells were collected, immediately treated with 0.1% sodium azide, and washed twice with 50 mM EDTA (pH 7.5) (5 3 107 cells per plug, except for 8–10 3 107 cells per plug for time courses with GAL-FOB1 rrm3D and GAL-FOB1 rrm3D rad52D to detect DSBs at tRNA genes in Figure 6). Cells were resuspended in 50 mM EDTA (pH 7.5) (33 mL per plug) and incubated at 42 C. Cell suspension was mixed with the solution (66 mL per plug) containing 0.83% low-melting-point agarose SeaPlaque GTG (Lonza), 170 mM sorbitol, 17 mM sodium citrate, 10 mM EDTA (pH 7.5), 0.85% b-mercaptoethanol, 0.17 mg/ml Zymolyase 100 T (Nacalai) and poured into plug mold (Bio-RAD). Agarose was allowed to solidify Molecular Cell 66, 533–545.e1–e5, May 18, 2017 e2

at 4 C for > 30 min. Plugs were incubated for 1–1.25 hr at 37 C in the solution containing 450 mM EDTA (pH 7.5), 10 mM Tris-HCl (pH 7.5), 7.5% b-mercaptoethanol and 10 mg/ml RNaseA (Macherey-Nagel). Plugs were then incubated overnight at 50 C in the solution containing 250 mM EDTA (pH 7.5), 10 mM Tris-HCl (pH 7.5), 1% sodium dodecyl sulfate and 1 mg/ml Proteinase K (Merck Millipore). Plugs were washed four times with 50 mM EDTA (pH 7.5) and stored at 4 C in 50 mM EDTA (pH 7.5). For rDNA copy number assays, genomic DNA was prepared by the conventional method as previously described (Murakami et al., 2009). Cells were washed once with 50 mM EDTA (pH 8.0) and resuspended in 200 mL of 0.5 M EDTA (pH 8.0). Cell suspension was mixed with 800 mL of spheroplast buffer (1 M sorbitol, 0.1 M EDTA (pH 8.0), 1% b-mercaptoethanol, 1.25 mg/ml Zymolyase 100T) and incubated for 45 min at 37 C. The spheroplasts were collected by centrifugation for 5 min at 4,500g, lysed in 500 mL of lysis buffer (50 mM EDTA (pH 8.0), 50 mM Tris-HCl (pH 8.0), 0.5% sodium dodecyl sulfate, 200 mg/ml Proteinase K) and incubated for 1 hr at 65 C. Proteins were precipitated by adding 200 mL of 5 M potassium acetate and incubating on ice, followed by centrifugation at 15,000 rpm for 15 min at 4 C. DNA in the supernatant was isopropanol precipitated and suspended in 100 mL of TE (10 mM Tris-HCl [pH 7.5] and 1 mM EDTA [pH 8.0]) containing 0.1 mg/ml RNaseA (Macherey-Nagel). PFGE Analysis In Figures 1B, S1A and S1B, one-third of a plug and Hansenula wingei chromosomal DNA markers (Bio-Rad) was cut and placed on a tooth of the comb. After extra buffer was removed using lab wipers, the comb was set in the gel tray and 1.0% agarose solution (Pulsed Field Certified Agarose, Bio-Rad) in 0.5x TBE (44.5 mM Tris base, 44.5 mM boric acid and 1 mM EDTA, pH 8.0) was poured. PFGE was performed on a Bio-Rad CHEF DR-III system in 2.2 L of 0.5x TBE at 14 C under the following conditions: 68 hr at 3.0 V/cm, 120 included angle, initial switch time of 300 s, and final switch time of 900 s. DNA was stained with 0.5 mg/mL of ethidium bromide (EtBr) and photographed. In Figure S1C, one-third of a plug was loaded on 1.0% agarose gels in 1x TAE (40 mM Tris base, 20 mM acetic acid and 1 mM EDTA, pH 8.0), along with Hansenula wingei and Schizosaccharomyces pombe chromosomal DNA markers (Bio-Rad), as described above. PFGE was performed on a Bio-Rad CHEF DR-III system in 2.2 L of 1x TAE at 14 C under the following conditions: 73 hr 10 min at 2.0 V/cm, 106 included angle, constant switch time of 32 min 26 s, during which 1x TAE buffer was changed with fresh buffer every 24 hr. DNA was stained with 0.5 mg/mL of EtBr and photographed. In Figures S1B and S1C, DNA was vacuum-transferred to Hybond-XL (GE Healthcare) as described previously with slight modification (Murakami et al., 2009), using VacuGene XL (GE Healthcare). Briefly, gels were incubated for 20 min in freshly prepared 0.25N HCl. DNA was then vacuum-transferred to the membrane at 55 cm Hg for 20 min with denaturing buffer (1.5 M NaCl, 0.5N NaOH) and 2 hr with transfer buffer (1.5 M NaCl, 0.25 N NaOH). DNA was fix to the membrane by soaking it in freshly prepared 0.4 N NaOH for 10 min, followed by washing it with 2X SSC (0.3 M NaCl, 0.03 M citrate) for 10 min. rDNA Copy Number Assays Tetrads were dissected from MSY281, 285 and 437, and genomic DNA was prepared as described above from at least four independent colonies per genotype. The concentration of genomic DNA was determined by a Qubit dsHS assay on a Qubit 2.0 Fluorometer (Life technologies). 100 ng of genomic DNA was digested with the restriction enzyme BglII (TaKaRa) and diluted to 10 pg/ml. Reaction mixtures were prepared using SYBR Premix Ex TaqII (Tli RNaseH Plus) (TaKaRa) in a total volume of 20 ml, where 50 pg of BglII-digested DNA was mixed with each primer at a final concentration of 0.2 mM in 1x Master mix. Quantitative real-time PCR was performed in triplicate, using a Thermal Cycler Dice Real Time System II (TaKaRa) according to the manufacturer’s instructions. The primer sequences used are available upon request. Relative quantification of the amount of rDNA was performed by the DCt method, where the average cycle threshold (Ct) value for the MCM2 gene was subtracted from the average Ct value for rDNA in each sample. The amount of rDNA in one of the WT clones was set at 1, and the relative fold enrichment of rDNA in other clones was determined by the following formula: F(x) = 2-(X-Y), where X is the DCt of interest and Y is the averaged DCt of one of the WT clones. In Figure 1C, the average of WT clones was set at one. One-way ANOVA was used for multiple comparisons using GraphPad Prism software (version 6.0d). P values of less than 0.05 were considered as statistically significant. Southern Blotting Single-dimension agarose gel electrophoresis for physical assays Approximately one-third of an agarose plug was cut, transferred to 2.0 mL flat bottom tubes, and equilibrated four times in 1 mL of TE for 15 min at room temperature. In all the physical assays except for the one in Figures 6I and 6J, agarose plugs were then equilibrated twice for 30 min at room temperature in 1 mL of 1x NEBuffer 3.1 (New England Biolabs). After the buffer was discarded completely, each plug was incubated in 160–200 mL of 1x NEBuffer 3.1 containing 1 unit/ml of BglII (New England Biolabs) overnight at 37 C. In Figures 6I and 6J, agarose plugs were equilibrated with 1x CutSmart buffer (New England Biolabs) for subsequent digestion with EcoRV-HF (New England Biolabs), respectively. Plugs were placed on a tooth of the comb. After extra buffer was removed using lab wipers, the comb was set in the gel tray (15 3 25 cm) and 0.7% agarose gels in 1x TBE was poured. DNA was separated in 1.45L of 1x TBE at 2.0 V/cm for 20–22 hr with buffer circulation in Sub-cell GT electrophoresis system (Bio-Rad). DNA was vacuum-transferred to Hybond-XL (GE Healthcare) as described previously (Murakami et al., 2009) with slight modification (Murakami et al., 2009), using VacuGene XL (GE Healthcare). DNA was vacuum-transferred to the membrane at 55 cm Hg for 20 min with 0.25 N HCl, 20 min with denaturing buffer (1.5 M NaCl, 0.5N NaOH) and 2 hr with transfer buffer (1.5 M NaCl, 0.25 N NaOH). DNA

e3 Molecular Cell 66, 533–545.e1–e5, May 18, 2017

was fix to the membrane by soaking it in freshly prepared 0.4 N NaOH for 10 min, followed by washing it with 2 X SSC (0.3 M NaCl, 0.03 M citrate) for 10 min Two-dimensional agarose gel electrophoresis To identify replication intermediates detected by single-dimension agarose gel electrophoresis (Figure S2C), two sets of BglIIdigested DNA (one separated by single-dimension gel electrophoresis and the other by 2D gel electrophoresis) were separated on 0.7% agarose (SeaKem Agarose LE, Lonza) in 1x TBE at 2.0 V/cm for 22 hr with buffer circulation, as described above. Gel slices containing DNA of interest were excised, rotated 90 , and cast in a 1.2% agarose gel (SeaKem Agarose LE, Lonza) containing 0.3 mg/ml of EtBr in 1x TBE. The second-dimension gel electrophoresis was performed in 1x TBE containing 0.3 mg/ml of EtBr at 6.0 V/cm for 5.5 hr with buffer circulation. DNA was transferred to Hybond-XL (GE Healthcare), using VacuGene XL (GE Healthcare), as described above. For the 2D neutral/neutral gel analysis in Figure S3A, genomic DNA samples isolated from GAL-FOB1 and GAL-FOB1 ctf4D samples at t = 70 (min) were analyzed. Approximately one-half of an agarose plug was digested with BglII as described above. DNA was separated on 0.9% agarose (SeaKem Agarose LE, Lonza) in 1x TBE at 2.0 V/cm for 21.5 hr with buffer circulation, as described above. DNA was stained with 0.3 mg/mL of EtBr and photographed. Gel slices containing DNA of interest were excised, rotated 90 , and cast in a 1.2% agarose gel (SeaKem Agarose LE, Lonza) in 1x TBE containing 0.3 mg/mL of EtBr. The second-dimension gel electrophoresis was performed at 6.0 V/cm for 4–5 hr in 1x TBE containing 0.3 mg/ml of EtBr with buffer circulation. DNA was vacuum-transferred to Hybond-XL (GE Healthcare), using VacuGene XL (GE Healthcare), as described above. In Figure S3B, genomic DNA samples isolated from GAL-FOB1 and GAL-FOB1 ctf4D samples at t = 70 (min) were analyzed by 2D neutral/denaturing gel analysis as described previously (Oh et al., 2009) with modifications. Approximately one-half of an agarose plug was digested with BglII (New England Biolabs) as described above. DNA was separated on 0.9% agarose (SeaKem Agarose LE, Lonza) in 1x TBE at 2.0 V/cm for
18–21 hr with buffer circulation, as described above. DNA was stained with 0.5 mg/mL of EtBr and photographed. The gel was washed twice for 30 min in 10 mM EDTA (pH 8.0). Gel slices containing DNA of interest were excised, rotated 90 , and cast in a 1.2% agarose gel (SeaKem Agarose LE, Lonza) in dH2O. The gel was equilibrated in 5x alkaline running buffer (250 mM NaOH, 5 mM EDTA) and then in 1x alkaline running buffer. The second-dimension gel electrophoresis was performed at 1.0 V/cm for
24–48 hr at 4 C in 1x alkaline running buffer with buffer circulation. DNA was vacuum-transferred to Hybond-XL (GE Healthcare) at 55 cm Hg for 2.5 hr in denaturation buffer (1.5 M NaCl, 0.5 N NaOH), using VacuGene XL (GE Healthcare). Probe preparation and hybridization Primer sequences used to prepare probes were as follows: rDNA probe 8 in Figure 1A (50 -ACGAACGACAAGCCTACTCG and 50 -AAAAGGTGCGGAAATGGCTG); probe 1 in Figures 6A and 6D (50 -AATGGCATCATTTTGTGATGTG and 50 -AAGTAATATGCAA ATTTGTTATGACTTC); probe 2 in Figures 6A and 6F (50 -CTCTTCGTTTCGTCTGACAG and 50 -TACTACGTTTCTATGTTATTTTGTGC); probe 1 in Figures 6B and 6G (50 -TGAATAGCAGCACAAGATCAC and 50 -TCTGGAGCTTCAAGTTATTATGC); probe 2 in Figures 6B and 6H (50 -AAACCTTTTGGTACACGAACAG and 50 -TTACATTGTCGACAGATTCCC); probe 1 in Figures 6C and 6I (50 -GAATCTCAA GCCATTCACAAG and 50 -AGTCCATTCATTAGTATAAACTCACG); probe 2 in Figures 6C and 6J (50 -GACTCTGTTTGTAAAGAGATT TGG and 50 -TCACGTTCAGCAAATTGTGG); rDNA probe 1 in Figures S3B and S3C (50 -CATTTCCTATAGTTAACAGGACATGCC and 50 -AATTCGCACTATCCAGCTGCACTC). To prepare double-stranded rDNA probes, PCR products were generated by two rounds of PCR. In brief, the first round of PCR was performed with primers using genomic DNA as a template. The PCR products were gel-purified and used to seed a second round of PCR with the same primers, after which the PCR products were gel-purified. Radioactive probes were generated using High Prime (Roche), according to the manufacturer’s instructions. Briefly, 100 ng of DNA that was heat-denatured was incubated for 30 min at 37 C in a total volume of 20 mL containing 4 mL of High Prime, 5 mL of [a-32P]-dCTP (3,000 Ci/mmol, 10 mCi/ml, Perkin Elmer), followed by removal of unincorporated nucleotides using ProbeQuant G-50 Micro Columns (GE Healthcare). The strand-specific probes used for Figures S3B and S3C were prepared by linear PCR. The reactions were set up in a final volume of 20 mL containing 5 ng of gel-purified PCR products as a template, 0.2 mM dATP, 0.2 mM dTTP, 0.2 mM dGTP, 5 mL of [a-32P]-dCTP (3,000 Ci/mmol, 10 mCi/ml, Perkin Elmer), 1 mM primer, 1.25 units of ExTaq (TaKaRa) and 1x ExTaq buffer. PCR was initiated by a denaturation step at 94 C for 3 min, followed by 35 cycles of amplification (96 C for 20 s, 51 C for 30 s, and 72 C for 1 min), and a final step at 72 C for 2 min. Hybridization was performed as described previously (Murakami et al., 2009). Briefly, the membrane was pre-wetted with 0.5 M phosphate buffer (pH 7.2) and was pre-hybridized for > 1 hr at 65 C with 25 mL of hybridization buffer (1% bovine serum albumin (Nacalai), 0.5 M phosphate buffer (pH 7.2), 7% sodium dodecyl sulfate, 1 mM EDTA (pH 8.0)). The membrane was incubated overnight at 65 C with 25 mL of fresh hybridization buffer and heat-denatured probe. The membrane was washed four times for 15 min at 65 C with wash buffer (40 mM phosphate buffer [pH 7.2], 1% SDS, 1 mM EDTA [pH 8.0]) pre-equilibrated at 65 C. The membrane was exposed to phosphor screen. Image analysis The radioactive signal was detected using Typhoon FLA9000 or FLA7000 (GE Healthcare), and images were analyzed by FUJIFILM Multi Gauge version 2.0 software (Fujifilm). To quantify DSBs and replication intermediates in physical assays, the membranes were exposed to phosphor screens for an appropriate exposure time without any saturation of signals. Scanned images were quantified by creating profile regions for all of the lanes and selecting the bands of interest at the same time as setting polygonal background lines

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for each band. We quantified the signals of all bands detected in the lanes. Next, the levels of DSBs and replication intermediates were calculated as percent of signal intensity of each band relative to the total lane signal, followed by subtracting the value at t = 0 min from those at other time points. To determine the ratio of DSBs to arrested forks in Figure 2D, the number of probe annealing sites in each type of molecule was taken into consideration (Fritsch et al., 2010): DSB fragments contain one annealing site, whereas arrested forks contain two sites (Figure 1A). Thus, the proportion of DSBs relative to arrested forks was determined by dividing twice the signal intensity of DSBs by the signal intensity of arrested forks. The average number of arrested forks in GAL-FOB1 cells at the peak in Figure 2C was estimated as follows. Among DNA molecules detected in physical assays, arrested fork and converged fork intermediates contain two sites where the probe can anneal. Taking this into consideration, we halved their signal intensities and calculated the proportion of corrected arrested fork signal among the total DNA molecules detected. In GAL-FOB1 cells, the level of arrested forks ranged from 3.9% to 8.4% across five time courses, with a mean of 5.7%. As shown in Figure S2D, GAL-FOB1 cells carried roughly 140 rDNA copies, leading us to estimate that the average number of arrested forks in each cell at peak times ranged from 5.4 to 11.8, with a mean of 8.1. The average number of DSBs at peak time in Figure 2C was determined as described above. The DSB levels ranged from 0.45% to 0.86%, with a mean of 0.60%, leading to an estimate of 0.8 DSBs on average. Resected DSBs were often below the limit of detection by Southern blotting (< 0.1% of DNA) when quantified by the methods above. To better compare their levels, we over-exposed the membrane (so that the signals in the linear fragments, but not other bands, usually reached saturation) and quantified the signals in DSBs, resected DSBs, and the terminal fragment containing the telomere-proximal rDNA repeat and its adjacent non-rDNA fragment indicated by the open circle in Figure 2A. The signal intensity of DSBs and resected DSBs was calculated relative to the terminal fragment, and the value at t = 0 min was then subtracted from those at other time points. The level of resected DSBs in various mutants in Figures 3G, 5C, and S3G was compared by determining the proportion of resected DSBs relative to DSBs, instead of comparing the steady-state level of resected DSBs. To this end, DSBs and resected DSBs were quantified relative to the terminal fragment as described above. Next, we divided the maximum level of resected DSBs in each time course by the maximum level of DSBs in the same experiment. The hyper-resected DSBs seen in GAL-FOB1 rad52D, ctf4D rad52D, rad51D and ctf4D rad51D were quantified relative to the terminal fragment, as described above. Analysis of Colony Sizes Tetrads were dissected from MSY281, 287 and 389, and plates were incubated at 30 C for 2 days. Plates were scanned, images were imported, and the areas occupied by different colonies were analyzed by ImageJ (NIH). Plates were replica-plated onto appropriate media to genotype colonies. Colony sizes of different genotypes were compared by one-way ANOVA, using GraphPad Prism software (version 6.0d). QUANTIFICATION AND STATISTICAL ANALYSIS Statistical analysis was performed using GraphPad Prism software (version 6.0d or 6.0h). Statistical parameters are described in the Figures, Figure legends and METHOD DETAILS. DATA AND SOFTWARE AVAILABILITY Data have been deposited to Mendeley Data and are available at http://dx.doi.org/10.17632/b66s2sg297.1.

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