Stalled replication forks generate a distinct mutational signature in yeast Nicolai B. Larsena,b,1,2, Sascha E. Libertia,b,1, Ivan Vogela, Signe W. Jørgensena,b, Ian D. Hicksona,b,3, and Hocine W. Mankouria,b,3 a Center for Chromosome Stability, Department of Cellular and Molecular Medicine, University of Copenhagen, 2200 Copenhagen, Denmark; and bCenter for Healthy Aging, Department of Cellular and Molecular Medicine, University of Copenhagen, 2200 Copenhagen, Denmark

Proliferating cells acquire genome alterations during the act of DNA replication. This leads to mutation accumulation and somatic cell mosaicism in multicellular organisms, and is also implicated as an underlying cause of aging and tumorigenesis. The molecular mechanisms of DNA replication-associated genome rearrangements are poorly understood, largely due to methodological difficulties in analyzing specific replication forks in vivo. To provide an insight into this process, we analyzed the mutagenic consequences of replication fork stalling at a single, site-specific replication barrier (the Escherichia coli Tus/Ter complex) engineered into the yeast genome. We demonstrate that transient stalling at this barrier induces a distinct pattern of genome rearrangements in the newly replicated region behind the stalled fork, which primarily consist of localized losses and duplications of DNA sequences. These genetic alterations arise through the aberrant repair of a single-stranded DNA gap, in a process that is dependent on Exo1- and Shu1-dependent homologous recombination repair (HRR). Furthermore, aberrant processing of HRR intermediates, and elevated HRR-associated mutagenesis, is detectable in a yeast model of the human cancer predisposition disorder, Bloom’s syndrome. Our data reveal a mechanism by which cellular responses to stalled replication forks can actively generate genomic alterations and genetic diversity in normal proliferating cells.

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RecQ helicase DNA replication stress recombination

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losses or duplications of DNA sequences are both prevalent and variable between different tissues within individuals (10). These CNVs have been proposed to arise due to DNA replication-based mechanisms and can cause chromosomal structural changes that drive somatic cell evolution (11). However, the molecular mechanisms that promote mutagenesis at stalled replication forks remain poorly characterized, especially in normal proliferating cells. A number of studies have used programmed DNA replication fork barriers as a tool to perturb DNA replication at a specific locus (12). These DNA replication barriers can be engineered into precise regions of the genome to monitor cellular responses to a stalled replication fork in the absence of any genome-wide stress, or global checkpoint activation. In this context, our previous work revealed that the Escherichia coli Tus/Ter complex could function as a polar DNA replication barrier that can transiently arrest replication forks when engineered into the Saccharomyces cerevisiae genome (13, 14). To better understand mutational processes occurring at a single perturbed replication fork, we examined mutagenic signatures arising at a Tus/Ter barrier in wild-type cells and in the sgs1 mutant, which is the yeast model of the human cancer predisposition disorder, Bloom’s syndrome. Using this approach, we have identified a distinctive mutagenic signature, consisting predominantly of localized deletions and duplications of DNA sequences, in the newly replicated region behind the stalled replication fork. Furthermore, we demonstrate that these mutations arise through aberrant Shu1-dependent HR repair of an

M

utations accumulate during successive rounds of somatic cell division, and these can contribute to cancer development and aging throughout the course of our life span. Indeed, stochastic mutations arising during DNA replication have been proposed as the major contributory factor that determines cancer incidence (1). Examples of DNA replication errors include the misincorporation of bases during the elongation step of DNA replication, or mutagenic processes that are deployed when the DNA replication machinery encounters various types of impediments such as DNA secondary structures, DNA-bound proteins, or DNA adducts (2). The precise stress response pathway used in response to DNA replication perturbation depends on a number of factors, including the nature of the impediment, the genomic locus, and the precise stage of S-phase. However, some DNA replication stress response processes are inherently error-prone, such as translesion DNA synthesis (3) or replication fork restart via homologous recombination-mediated mechanisms (4–6). In these scenarios, cells adopt error-prone repair processes that prioritize cellular survival over the fidelity of DNA replication. However, a corollary to this is that DNA replication stress can serve as a driver of mutagenesis in cancer cells, which often have a dysregulated DNA replication program due to oncogene activation (7). Mutation signatures have been characterized extensively in cancer cells. Interestingly, and despite the methodological difficulties involved in their detection, normal cells can also exhibit mutational signatures and types of genome rearrangements that are commonly observed in cancer cells of the same cell type (8, 9). Furthermore, copy-number variations (CNVs) caused by

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Significance The molecular mechanisms that generate genome alterations and genetic heterogeneity in proliferating cells are technically challenging to delineate. To analyze mutagenic outcomes at a perturbed replication fork, we engineered an inducible replication fork barrier, coupled with a genetic reporter, into the yeast genome. We demonstrate that replication fork stalling triggers a cellular response mechanism that can generate localized losses and duplications of DNA sequences as an associated cost. Because the key proteins involved in this process are evolutionarily conserved in eukaryotes, we propose these findings may reveal a ubiquitous cellular response to DNA replication stress, as well as a conserved mechanism of DNA replication-associated mutagenesis. Author contributions: N.B.L., S.E.L., I.D.H., and H.W.M. designed research; N.B.L., S.E.L., S.W.J., and H.W.M. performed research; N.B.L., S.E.L., S.W.J., and H.W.M. contributed new reagents/analytic tools; N.B.L., S.E.L., I.V., I.D.H., and H.W.M. analyzed data; and I.D.H. and H.W.M. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. 1

N.B.L. and S.E.L. contributed equally to this work.

2

Present address: The Novo Nordisk Foundation Center for Protein Research, Faculty of Health and Medical Sciences, University of Copenhagen, 2200 Copenhagen, Denmark.

3

To whom correspondence may be addressed. Email: [email protected] or hocine@sund. ku.dk.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10. 1073/pnas.1706640114/-/DCSupplemental.

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Edited by Philip C. Hanawalt, Stanford University, Stanford, CA, and approved July 28, 2017 (received for review April 22, 2017)

Exo1-generated ssDNA gap. Our data reveal a mechanism by which nucleolytically generated ssDNA gaps at perturbed replication forks can potentially fuel genome rearrangements, including CNVs, in the human genome. Results Site-Specific Stalling at a Tus/Ter Barrier Triggers Mutagenesis Behind the Replication Fork. To analyze the phenotypic consequences of

replication fork stalling at a nonprogrammed DNA replication barrier, we engineered yeast strains harboring 14xTer sites in place of the HIS2 locus on ChrVI. This location is ∼4 Kb away from the efficient, early-firing DNA replication origin, ARS607. Therefore, the vast majority of replication forks encountering the Tus/Ter barrier emanate from this origin. A URA3 reporter gene, which permits the positive selection for ura3 mutations when cells are exposed to 5-fluoroorotic acid (5-FOA), was coengineered in either of two locations relative to the Ter sites: origin-proximal (ARS607→URA3→Tus/Ter; Fig. 1A) or origin-distal (ARS607→Tus/ Ter→URA3; Fig. 1B). The origin-proximal configuration reveals mutagenic events occurring in the newly replicated DNA behind the Tus/Ter-arrested replication fork, whereas the origin-distal configuration reveals mutagenic events associated with replication fork resumption, DNA replication termination, or replication fork restart. In both configurations of the URA3 reporter, we observed that the Tus/Ter barrier caused site-specific replication fork stalling, as detected by 2D gel electrophoresis (2DGE; Fig. 1 A and B and Fig. S1A). As demonstrated previously, replication fork stalling at Tus/Ter was also associated with increased levels of unprocessed, X-shaped DNA (X-DNA) in strains lacking the RecQ helicase, Sgs1 (13). We assessed whether the URA3 reporter constructs positioned on either side of the Tus/Ter barrier were prone to increased mutagenesis after replication fork stalling. In wild-type cells,

induction of the Tus/Ter barrier did not influence the URA3 mutation rate when the reporter was placed in either the originproximal or origin-distal configuration (Fig. 1C). Similarly, there was no pronounced increase in URA3 mutagenesis in the origindistal reporter in sgs1 mutants. Interestingly, however, we observed an approximately ninefold increase in the mutation rate in the origin-proximal URA3 reporter in sgs1 mutants when the Tus/Ter barrier was induced (Fig. 1C). This increased mutagenesis in sgs1 mutants was not detectable when the polar Tus/Ter barrier (13) was orientated in the nonblocking (permissive) orientation. Furthermore, increased mutagenesis in sgs1 mutants was independent of the direction of URA3 transcription (Fig. S1 A and B). Spontaneous CAN1 (on ChrV) mutation rates were similar in all the wild-type or sgs1 strains harboring Tus/Ter barriers (Fig. 1D), indicating global mutation rates were generally unaffected by Tus expression or by deletion of SGS1. To compare for locus-specific effects, we also engineered the origin-proximal URA3 construct into another location adjacent to ARS305 on ChrIII. Consistent with the ChrVI data (Fig. 1C), we observed that the URA3 mutation rate was not significantly altered when the Tus/Ter barrier was induced in wild-type cells, but was elevated approximately ninefold in the sgs1 mutant (Fig. S1C). This demonstrates that the Tus/Ter-induced mutagenesis observed in the origin-proximal URA3 reporter in sgs1 mutants is a general consequence of replication fork stalling, and not specific for the ChrVI his2 locus. Taken together, our data reveal that transient replication fork stalling in the sgs1 mutant leads to unprocessed X-DNA, and an elevated incidence of mutations in the newly replicated region behind the Tus/Ter barrier. Replication Fork Stalling at Tus/Ter Barriers Causes Heterogeneous Types of Mutations. To define the types of mutations arising in

the origin-proximal URA3 reporter, we analyzed the sequences

Fig. 1. Site-specific fork stalling at a Tus/Ter barrier triggers X-DNA and localized mutagenesis. (A) Wildtype and sgs1 mutants harboring an origin-proximal his::URA3-14xTer cassette were released from G1arrest after the induction of Tus. Genomic DNA was extracted after 35 min, and an MfeI-MfeI restriction fragment was analyzed by 2DGE, using a ChrVI-specific probe. The blue arrow on the 2DGE images indicates stalled forks detectable at the Tus/ Ter barrier, whereas the green arrow indicates unprocessed X-DNA in the sgs1 mutant. Cell-cycle profiles are shown below the respective 2DGE image. (B) Wild-type and sgs1 strains harboring an origindistal his2::14xTer-URA3 cassette were analyzed as above, with the exception that a ClaI-NruI restriction fragment was analyzed by 2DGE. (C) URA3 mutation rates and (D) CAN1 mutation rates were measured simultaneously in strains harboring the indicated Ter modules and plasmids. Error bars indicate 95% confidence limits, and numerical values above columns indicate the fold-difference in mutation rate between isogenic strains. Statistical analysis of differences in mutation rates was performed using a one-sided Mann–Whitney U test, and statistical significance in our assays was indicated when P < 0.01 (**P < 0.01; ****P < 0.0001).

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of the mutated ura3 locus in wild-type and sgs1 5-FOAR colonies harboring either an empty vector control or a Tus-expression plasmid. Mutations were assigned to the following categories: base errors (base substitutions, or −1/+1 frame-shifts), deletions, duplications, “complex” (>1 base error within a 12-bp run), or no detectable mutation in the URA3 ORF [i.e., due to extragenic mutations that cause 5-FOA resistance (15)]. In both wild-type and sgs1 mutants harboring an empty (no Tus) vector, ∼70% of detectable ura3 mutations were single-base substitutions (Fig. 2), indicative of spontaneous mutations arising within URA3 (15). Interestingly, despite the lack of change in the URA3 mutagenesis rate in wild-type cells (Fig. 1C), Tus/Ter-mediated fork stalling caused a notable change in the mutation spectrum (Fig. 2). Moreover, in sgs1 cells, the increased Tus/Ter-induced mutation rate (Fig. 1C) was associated with an altered mutation spectrum (Fig. 2). Similar results were observed at both the ChrVI and ChrIII Tus/Ter barriers (Datasets S1 and S2), indicating that the observed mutations were independent of the chromosomal location. The most prominent type of Tus/Ter-induced mutation in both wild-type and sgs1 cells was a deletion of variable size (Fig. 2 and Fig. S2). Duplications of localized sequences were also detectable, but were less frequent (Fig. 2). The majority of deletions and duplications exhibited detectable homology or homeology at their breakpoints

Shu1-Dependent HRR Generates Mutations at a Tus/Ter-Stalled Replication Fork. Because the Tus/Ter barrier generates losses

and duplications of sequences flanked by homology/homeology (Datasets S1 and S2), we investigated the consequences of fork stalling when homologous recombination repair (HRR) was disabled. In agreement with our previous results (13), we observed that deletion of RAD51 (the central HRR factor required for strand invasion) prevented X-DNA accumulation in sgs1 mutants after transient fork stalling at the Tus/Ter barrier (Fig. 3A). To investigate the type of HRR occurring at the Tus/Ter barrier, we analyzed the effects of deleting SHU1. Shu1 is a member of the heterotetrameric Shu protein complex that is implicated in the repair of DNA replication-associated damage (16, 17). Whereas RAD51 deletion

Fig. 3. Shu1-dependent homologous recombination occurs at a Tus/Ter barrier. (A) The indicated yeast strains harboring his2::URA3-14xTer were analyzed by 2DGE at 35 min after release from G1. The green arrow indicates unprocessed X-DNA in the sgs1 mutant. Cell-cycle profiles are shown below the respective 2DGE image. (B) URA3 mutation rates were determined in the indicated strains harboring his2::URA3-14xTer. Statistical analysis of differences in mutation rates was performed using a one-sided Mann–Whitney U test, and statistical significance is indicated when P < 0.01 (****P < 0.0001). (C) Mutations in the ChrVI origin-proximal URA3 reporter were identified in individual 5-FOAR colonies by DNA sequencing. Pie charts indicate the relative proportion of mutation types identified in the shu1 and shu1 sgs1 mutants expressing Tus. For cross-reference, the wild-type and sgs1 (+Tus) data from Fig. 2 are repeated here. A list of mutations arising in shu1 and shu1 sgs1 strains expressing Tus can be found in Dataset S3.

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Fig. 2. Replication fork stalling at the Tus/Ter barrier triggers heterogeneous types of mutations. Mutations in the ChrVI origin-proximal URA3 reporter were identified in individual 5-FOAR colonies by DNA sequencing. Pie charts indicate the relative proportion of mutation types identified in each strain, and the mutations are listed in Dataset S1.

(Datasets S1 and S2), suggesting these genome rearrangements might have arisen via a common mechanism. A comparison of deletion sizes showed a tendency for the deletions to be larger in sgs1 cells than in wild-type cells (Fig. S2). For example, 70% of the deletions in wild-type cells were 100 bp in sgs1 mutants. While the deletions in wild-type cells were highly heterogeneous (32 of 34 different deletions detected), certain deletion types recurred (namely, 114-, 147-, 358-, and 423-bp deletions) in sgs1 cells. However, it should be noted that each of these recurrent deletions was also detected at least once in wildtype cells (Datasets S1 and S2), suggesting they arise via similar mechanisms in wild-type and sgs1 mutants, but that Sgs1 minimizes their accumulation. Another Tus/Ter-induced mutation type we observed was defined as “complex” and consisted of multiple (>1) base errors within a 12-bp run. These complex mutations were detectable at a very low frequency in wild-type cells after induction of Tus/Ter, but were prevalent in sgs1 mutants (Fig. 2 and Datasets S1 and S2). Taken together, we conclude that fork stalling at a Tus/ Ter barrier can generate localized deletions, duplications, and complex mutations behind the replication fork. Furthermore, large deletions and complex mutations are overrepresented in sgs1 mutants, suggesting Sgs1 counteracts the mutagenic processes that generate these.

Fig. 4. Exo1 generates ssDNA at Tus/Ter-stalled forks. (A) URA3 mutation rates were determined in the indicated strains harboring his2::URA3-14xTer. Statistical analysis of differences in mutation rates was performed using a one-sided Mann–Whitney U test, and statistical significance is indicated when P < 0.01 (****P < 0.0001). (B) Mutations in the ChrVI origin-proximal URA3 reporter were identified in individual 5-FOAR colonies. Pie charts indicate the relative proportion of mutation types identified in the exo1 and exo1 sgs1 mutants expressing Tus. For cross-reference, the wild-type and sgs1 (+Tus) data from Fig. 2 are repeated here. A list of mutations arising in exo1 and exo1 sgs1 strains expressing Tus can be found in Dataset S4. (C) Diagrammatic representation of the amplicons that were quantified by qPCR. The numbers in the diagram indicate the distance (bp) from the first Ter site. (D) Yeast strains harboring his2::URA3-14xTer were released from G1-arrest after the induction of Tus. After 35 min, DNA-associated proteins were crosslinked and extracted, and RPA-associated chromatin was enriched using an anti-RPA antibody. RPA enrichment at specific loci was detected by qPCR analysis of various (colorindicated) amplicons located anterior to the 14xTer array. The data shows the average RPA enrichment observed in each amplicon over three independent experiments, normalized to an averaged signal for replicating chromatin on ChrVI (see Methods). Error bars indicate the SD. (E) RPA enrichment profiles of the indicated strains harboring his2::URA3-14xTer, and expressing Tus, are shown.

completely eliminates X-DNA accumulation in methyl methanesulfonate (MMS)-treated sgs1 mutants (18), deletion of SHU1 only partially reduces the accumulation of X-DNA (17). This suggests that Shu1 either affects the overall kinetics or efficiency of HRR, or that different types of DNA lesions generated by MMS exposure can lead to genetically heterogeneous types of X-DNA in S-phase. Interestingly, and in support of the latter hypothesis, 2DGE revealed that deletion of SHU1 abolished X-DNA accumulation in the sgs1 mutant after stalling at the Tus/Ter barrier (Fig. 3A). Therefore, Shu1-dependent HRR is likely the principal mechanism that generates X-DNA after replication fork arrest at a Tus/Ter barrier. Next, we examined mutation rates in strains lacking Rad51 or Shu1. Previous findings have demonstrated that spontaneous mutagenesis rates are elevated ∼10-fold in rad51 strains, or approximately sixfold in shu1 strains (19). We observed that the Tus/Ter-induced URA3 mutagenesis rate was not markedly enhanced (above that observed in the corresponding empty vector control strain) in shu1, shu1 sgs1, rad51, or rad51 sgs1 mutants (Fig. 3B). To examine whether Tus/Terinduced deletions and duplications were Shu1-dependent, we performed DNA sequence analysis of ura3 mutations in shu1 and shu1 sgs1 strains expressing Tus. We observed that deletions and duplications were largely eliminated when SHU1 was deleted (50% for wild-type or sgs1 (Fig. 2). Taken together, we propose that the majority of genome rearrangements occurring behind the Tus/Ter barrier in wild-type and sgs1 mutants are Shu1-dependent. The Substrate for Shu1-Dependent HRR Is an ssDNA Gap That Is Generated by Exo1. Because replication fork stalling at a Tus/

Ter barrier does not cause a double-strand break (DSB) in yeast (13), we investigated whether an ssDNA gap might be the primary lesion generated at a Tus/Ter-stalled fork. Exo1 has been demonstrated previously to create ssDNA gaps after DNA 4 of 6 | www.pnas.org/cgi/doi/10.1073/pnas.1706640114

replication perturbation (20, 21) and to suppress aberrant DNA replication-associated HRR events associated with loss of Sgs1 or Rqh1 (the RecQ orthologs in budding yeast and fission yeast, respectively) (22, 23). 2DGE analysis confirmed that loss of Exo1 did not noticeably affect the kinetics of replication fork stalling or resumption at the Tus/Ter barrier (Fig. S3A). Next, we analyzed URA3 mutation rates when EXO1 was deleted. We observed that the spontaneous mutation rate was elevated approximately fourfold in the exo1 mutant, which is in agreement with a previous study (24). We observed that induction of the Tus/Ter barrier did not noticeably elevate the URA3 mutagenesis rate in exo1 or exo1 sgs1 mutants (Fig. 4A). DNA sequence analysis of ura3 mutations in exo1 and exo1 sgs1 strains revealed that Tus/Ter-induced deletions and duplications were rare when Exo1 was deleted (Fig. 4B). We conclude, therefore, that Exo1, similar to Shu1 (Fig. 3), is required to generate the Tus/Ter-induced genome rearrangements that are detectable in wild-type and sgs1 strains. To examine whether an Exo1-generated ssDNA gap behind the stalled fork was the recombinogenic substrate for HRR, we analyzed ssDNA patterns around the Tus/Ter barrier, using chromatin immunoprecipitation of RPA bound to ssDNA (RPAChIP) (25). The level of RPA-associated ssDNA was quantified at various positions in/around the URA3 reporter gene, using qPCR (Fig. 4C). We observed a distinctive signature of RPA enrichment that was dependent on Tus expression (Fig. 4D). The major peak of RPA-associated ssDNA was detected in the region located 162–278 bp anterior to the first Ter site. Two smaller peaks of RPA enrichment were also detectable within the Ter array and the region 719–837 bp anterior to the first Ter site. We then compared the RPA profiles in wild-type, sgs1, exo1, and shu1 mutants (Fig. 4E). Similar levels of RPA enrichment were Larsen et al.

Discussion Using the E. coli Tus/Ter complex as an inducible replication fork barrier (13, 14, 27), we have analyzed the mutagenic events associated with replication fork stalling in a single-copy, and early-replicating, locus. We demonstrate that deletions and duplications of DNA sequences can occur via aberrant Shu1dependent HRR of an Exo1-generated ssDNA gap (Fig. 5). Hence, our data reveal the intrinsic error rate of a physiological process that is actively triggered in response to DNA replication perturbation. In the absence of Sgs1, the Tus/Ter-induced mutation rate is elevated, and the mutation spectrum is skewed toward increased levels of recurrent large deletions and complex mutations (clusters of multiple base errors within a 12-bp run). Nevertheless, the cellular responses to Tus/Ter-induced replication fork stalling, as well as the mutation types, were highly similar in wildtype and sgs1 strains. We propose, therefore, that similar DNA replication stress response activities are operational in wild-type and sgs1 cells, and that Sgs1 minimizes the error rate of Shu1dependent HRR (Fig. 5). Indeed, we detect unprocessed HRR intermediates and increased levels of aberrant HRR events when Sgs1 is absent, consistent with the reported roles for Sgs1 in promoting recombination fidelity (28, 29). It is also intriguing that complex mutations, which have mutagenic signatures that are reminiscent of Pol zeta-generated errors (3), are noticeably elevated in sgs1 mutants. Given that the increase in complex mutations in sgs1 mutants is also Exo1- and Shu1-dependent, we speculate that Pol zeta might assist Exo1/Shu1-dependent HRR by promoting D-loop extension in regions that are prone to form secondary structures during HRR-associated DNA synthesis. Degradation of nascent DNA at damaged replication forks has been observed in a number of different organisms and contexts, ranging from bacteria to humans (21, 30, 31). Controlled ssDNA generation after replication fork perturbation may serve a number of important functions, including ssDNA-mediated checkpoint signaling roles and the loading of the most appropriate repair/ replication factors required to overcome the DNA replication impediment. Interestingly, replication fork stalling at Tus/Ter is transient in yeast (13), and loss of Exo1 does not apparently affect the kinetics of fork stalling and resumption at the Tus/Ter barrier. A similar role for Exo1 in generating ssDNA at a stalled fork has also been demonstrated at the S. pombe RTS1 barrier, despite the different cellular responses and phenotypic outcomes to replication fork arrest at this barrier (6, 21). Therefore, the role of Exo1 in generating ssDNA at perturbed replication forks may be a universal, and nonpathological, default mechanism. The repair of ssDNA gaps during DNA replication is the most common use of HRR in E. coli (32). If the same is also true in eukaryotes, this suggests the repair of ssDNA gaps by HRR Larsen et al.

Fig. 5. Model for controlled ssDNA genesis and repair at a perturbed replication fork. After replication fork stalling at a Tus/Ter barrier, Exo1-mediated DNA degradation occurs behind the stalled replication fork to generate an ssDNA gap on the lagging strand template. Shu1 then promotes Rad51 loading onto the RPA-coated ssDNA gap (41). Fork remodeling may lead to small deletions and duplications via DNA polymerase slippage events (42). Alternatively, the ssDNA gap may be repaired by postreplicative HRR after replication resumption. Deletions and duplications may arise due to nonallelic HRR at regions of short homology or homeology. Complex genome alterations (manifesting as short clusters of base errors) can also occur after the strand-invasion step, possibly triggered by the deployment of Pol zeta at secondary structures that impede HRR-associated DNA synthesis. Sgs1 counteracts Shu1/Rad51-induced mutations by rejecting aberrant strand invasion events and disrupting the secondary structures that can impede HRR. In humans, an analogous process to this may contribute to CNV formation following DNA replication perturbation.

during S-phase could have profound effects on the highly repetitive genomes of proliferating somatic cells, particularly when BLM is mutated or overwhelmed (e.g., in precancerous cells exhibiting oncogene-induced DNA replication stress (7)). A number of studies have implicated the Shu complex in promoting HRR at specific types of DNA replication-blocking lesions. Importantly, this function appears to be conserved in eukaryotes, as loss of Shu orthologs in a number of organisms causes pronounced sensitivity to replicationblocking agents (e.g., MMS or crosslinking agents), but has only minor effects on DSB repair (16, 33–35). Our observations that Shu1-mediated HRR can generate genome rearrangements may therefore reveal an evolutionarily conserved mechanism by which CNVs can arise after DNA replication perturbation. Indeed, up to 12% of the human genome is estimated to be subject to CNV, and different organs and tissues within the same individual can show substantial genotypic differences (10). On the basis of our findings, we propose that the human Shu proteins (SWS1 and SWS1AP) could actively promote HRR-mediated CNVs after replication fork stalling in some circumstances. Furthermore, our proposed model (Fig. 5) may also explain why the replicative mechanisms involved in human CNV formation are error-prone (36). Given that an individual with a homozygous mutation in SWS1 has been identified (37), it will be of interest to examine precisely how SWS1-deficient cells respond to DNA replication stress. Methods Strains and Plasmids. The Tus-expression plasmids were described previously (13). All yeast strains used in this study are isogenic derivatives of BY4741,

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detected within the Ter array in all strains, suggesting this RPA signal is associated with Tus/Ter-mediated stalling of the replisome. Interestingly, the RPA-ChIP signal in the regions located at 162– 278 or 719–837 bp anterior to the first Ter site were diminished in the exo1 mutant. The profile of RPA enrichment was similar in wild-type, sgs1, and shu1 strains, suggesting ssDNA tracts arise with similar spatiotemporal kinetics in the absence of Sgs1 or Shu1. We also used a modified form of the quantitative amplification of single-stranded DNA (QAOS) (26) assay to analyze the strand bias of the ssDNA in the −278 to −162 amplicon. This analysis was consistent with the lagging strand template being the predominant source of the RPA signal (Fig. S3). Taken together, these data indicate that Exo1 degrades the nascent lagging strand behind a transiently stalled fork to generate an ssDNA gap that is subsequently repaired by Shu1-dependent HRR. Furthermore, we propose that aberrant Shu1-mediated HRR of this ssDNA gap is the principal mechanism that can also generate genome rearrangements behind a stalled replication fork.

which lack a 1.1-Kb segment of the URA3 gene on ChrV. The 14xTer modules, ORF deletion cassettes, and yeast marker-swap cassettes were all integrated into the yeast genome by targeted homologous recombination (38). The ChrVI and ChrIII targeting primers used in this study are indicated in Table S1. For the RPA-ChIP experiments, a Tus expression cassette was integrated into the ura3Δ0 locus on ChrV. Growth Conditions and Flow Cytometry. Yeast cultures were grown in YEP medium (Formedium) supplemented with 3% sodium DL-lactate solution. Cultures were synchronized in G1 with alpha-factor for 3 h (CASLO ApS). Tus expression was induced by adding 2% Galactose (final wt/vol) for the final 2.5 h of the G1-arrest. Release of cells from G1-arrest was achieved by centrifugation, washing, and resuspension of cells in fresh medium. Cell-cycle stage was determined by flow cytometry (38).

Chromatin Immunoprecipitation and QAOS. RPA-ChIP was performed as described previously (25). Real-time PCR was performed using a StepOnePlus Real-Time PCR System (Applied Biosystems) and Power SYBR Green Master Mix (ThermoFisher Scientific). The primers used for each amplicon are indicated in Table S1. The “Fold increase” for each amplicon was calculated as: Fold increase = (2−Ct Anti-RFA − 2−Ct BEADS)/(2-Ct Input × dilution factor). RPAChIP experiments were performed in triplicate, and average fold increase was calculated for each individual amplicon. All qPCR data were normalized to an averaged (across all strains and triplicate experiments) control region to the left of ARS607 that detects RPA levels at nonperturbed replication forks (Table S1). QAOS was also performed on the RPA-ChIP samples using the qPCR conditions described in Holstein and Lydall (26). The primers used for QAOS are indicated in Table S1.

Analysis of Mutation Rates and Types. Individual colonies picked from 2% raffinose plates were grown to saturation in nonselective medium containing 2% Galactose, and URA3 and CAN1 mutation rates were measured by fluctuation analysis (39, 40). Statistical analysis of differences in mutation rates was performed using a one-sided Mann-Whitney U test, and statistical significance in our assays was indicated when P < 0.01. For analysis of mutation types, cells growing on 2% raffinose plates were plated onto nonselective plates containing 2% Galactose. Plates were incubated at 25 °C for 4–5 d, and then replica plated onto plates containing 5-FOA. Individual colonies were confirmed as 5-FOA resistant, and the URA3 locus was sequenced using two different primers. Mutations were scored as events that were detectable in both sequence reads.

2D Gel Analysis of DNA Structures. The hexadecyltrimethylammonium bromide method of DNA extraction was used, and 20 μg DNA was analyzed by 2DGE, as described previously (38).

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ACKNOWLEDGMENTS. We gratefully acknowledge Lotte Bjergbæk for demonstrating the RPA-ChIP technique and Scott Lujan for advice about calculating mutation rates. We also thank Lotte Bjergbæk, Eva Hoffman, Michael Lisby, Simon Powell, Ralph Scully, and members of the I.D.H. laboratory for helpful discussions. Work in the authors’ laboratory is funded by the Danish National Research Foundation (DNRF115), the European Research Council, the Novo Nordisk Foundation, Fabrikant Einar Willumsens Mindelegat, the Danish Research Council, and the Nordea Foundation.

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