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Spatial separation between replisome‐ and template‐induced replication stress signaling

2018; Springer Nature; Volume: 37; Issue: 9 Linguagem: Inglês

10.15252/embj.201798369

1460-2075

Néstor García‐Rodríguez, Magdalena Morawska, Ronald P.C. Wong, Yasukazu Daigaku, Helle D. Ulrich,

Genomics and Chromatin Dynamics

Article26 March 2018Open Access Source DataTransparent process Spatial separation between replisome- and template-induced replication stress signaling Néstor García-Rodríguez Néstor García-Rodríguez orcid.org/0000-0002-4049-1604 Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Magdalena Morawska Magdalena Morawska Institute of Molecular Biology (IMB), Mainz, Germany Cancer Research UK London Research Institute, Clare Hall Laboratories, Blanche Lane, South Mimms, UK Search for more papers by this author Ronald P Wong Ronald P Wong orcid.org/0000-0002-4652-514X Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Yasukazu Daigaku Yasukazu Daigaku Cancer Research UK London Research Institute, Clare Hall Laboratories, Blanche Lane, South Mimms, UK Search for more papers by this author Helle D Ulrich Corresponding Author Helle D Ulrich [email protected] orcid.org/0000-0003-0431-2223 Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Néstor García-Rodríguez Néstor García-Rodríguez orcid.org/0000-0002-4049-1604 Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Magdalena Morawska Magdalena Morawska Institute of Molecular Biology (IMB), Mainz, Germany Cancer Research UK London Research Institute, Clare Hall Laboratories, Blanche Lane, South Mimms, UK Search for more papers by this author Ronald P Wong Ronald P Wong orcid.org/0000-0002-4652-514X Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Yasukazu Daigaku Yasukazu Daigaku Cancer Research UK London Research Institute, Clare Hall Laboratories, Blanche Lane, South Mimms, UK Search for more papers by this author Helle D Ulrich Corresponding Author Helle D Ulrich [email protected] orcid.org/0000-0003-0431-2223 Institute of Molecular Biology (IMB), Mainz, Germany Search for more papers by this author Author Information Néstor García-Rodríguez1, Magdalena Morawska1,2,3, Ronald P Wong1, Yasukazu Daigaku2,4 and Helle D Ulrich *,1 1Institute of Molecular Biology (IMB), Mainz, Germany 2Cancer Research UK London Research Institute, Clare Hall Laboratories, Blanche Lane, South Mimms, UK 3Present address: Springer Nature, London, UK 4Present address: Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, Aoba-ku, Sendai, Japan *Corresponding author. Tel: +49 6131 3921490; E-mail: [email protected] The EMBO Journal (2018)37:e98369https://doi.org/10.15252/embj.201798369 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Polymerase-blocking DNA lesions are thought to elicit a checkpoint response via accumulation of single-stranded DNA at stalled replication forks. However, as an alternative to persistent fork stalling, re-priming downstream of lesions can give rise to daughter-strand gaps behind replication forks. We show here that the processing of such structures by an exonuclease, Exo1, is required for timely checkpoint activation, which in turn prevents further gap erosion in S phase. This Rad9-dependent mechanism of damage signaling is distinct from the Mrc1-dependent, fork-associated response to replication stress induced by conditions such as nucleotide depletion or replisome-inherent problems, but reminiscent of replication-independent checkpoint activation by single-stranded DNA. Our results indicate that while replisome stalling triggers a checkpoint response directly at the stalled replication fork, the response to replication stress elicited by polymerase-blocking lesions mainly emanates from Exo1-processed, postreplicative daughter-strand gaps, thus offering a mechanistic explanation for the dichotomy between replisome- versus template-induced checkpoint signaling. Synopsis Replisome-inherent problems activate the yeast Rad53 checkpoint via the replisome-associated mediator Mrc1. In contrast, checkpoint activation caused by replication-blocking template DNA damage appears to originate from Rad9 signaling induced by Exo1-processed daughter-strand gaps behind replication forks. Rad9 signaling is essential for damage resistance and Rad53 activation when damage bypass is delayed. Rad53 is required to ensure genome integrity during S phase, but not for postreplicative gap filling. Rad53-mediated inhibition of Exo1 and Pif1 maintains bypass competence during S phase. Damage signaling during S phase requires Exo1 activity at daughter-strand gaps. DNA lesions and replisome problems activate checkpoint signaling via distinct structures. Introduction Genome maintenance relies on checkpoint pathways that perceive DNA damage or replication problems and initiate an appropriate response. In eukaryotic cells, they are mediated by kinase cascades activated by distinct types of abnormal DNA structures (Nyberg et al, 2002). In vertebrates, damage signaling by the ATM kinase is initiated by DNA double-strand breaks (DSBs), whereas the related ATR kinase reacts to a variety of lesions and is activated mainly by single-stranded DNA (ssDNA). During S phase, cells are particularly vulnerable to conditions that challenge the progression of the replisome. In this situation, ssDNA is thought to accumulate at stalled replication forks by an uncoupling between helicase and polymerase movement or between leading and lagging strand synthesis. In budding yeast, the checkpoint response elicited by these structures is initiated by Mec1, the homologue of vertebrate ATR, which is responsible for activating an effector kinase, Rad53. Via phosphorylation of a large set of substrates, Rad53 mediates most aspects of the checkpoint response, including a stabilization of stalled forks, suppression of late origin firing, control of nucleotide levels, regulation of damage-induced transcription, and arrest of the cell cycle (Pardo et al, 2017). Intriguingly, checkpoint signaling in response to replication stress can be divided into two branches that both initiate from Mec1 and converge on Rad53, but differ in the mediator protein responsible for signal transmission: the DNA replication checkpoint and the DNA damage checkpoint (Pardo et al, 2017). Upon inhibition of ribonucleotide reductase by hydroxyurea (HU), Mec1 phosphorylates the replisome component, Mrc1, a homologue of claspin. In response to DNA damage, Mec1 cooperates with the 53BP1 homologue Rad9. This dichotomy has led to the speculation that a replication fork stalled by nucleotide depletion adopts a structure distinct from one that is stalled by a lesion in the template (Alcasabas et al, 2001; Nielsen et al, 2013). However, the basis for such difference remains unclear. Outside of S phase, ssDNA as a source of checkpoint activation can arise from nucleotide excision repair (NER) or from the resection of 5′-termini at DSBs or uncapped telomeres. In both situations, a 5′–3′ exonuclease, Exo1, contributes to Rad53 activation by widening NER gaps or processing DNA termini (Nakada et al, 2004; Dewar & Lydall, 2010; Giannattasio et al, 2010). At the same time, Exo1 is a downstream target of Rad53, which inhibits the nuclease by phosphorylation (Smolka et al, 2007; Morin et al, 2008). This results in a negative feedback that prevents excessive Exo1 activity. At stalled replication forks, Exo1 degrades abnormal structures and prevents fork reversal, but it does not contribute to damage signaling or replication restart and may even promote fork breakdown (Cotta-Ramusino et al, 2005; Segurado & Diffley, 2008). At collapsed replication forks, Exo1 activity is deemed to be mostly detrimental and is subject to checkpoint-mediated inhibition (Tsang et al, 2014). As an alternative to persistent fork stalling, re-priming of DNA synthesis downstream of a lesion can give rise to daughter-strand gaps behind the replication fork. This has been studied most extensively in bacterial systems (Heller & Marians, 2006), but there is good evidence for a "skipping" of DNA damage-induced lesions in eukaryotic cells as well (Lopes et al, 2006; Elvers et al, 2011). Ultimately, however, cell proliferation requires complete genome replication, necessitating the activity of DNA damage bypass pathways to copy the damaged DNA (Friedberg, 2005; Ulrich, 2009). Importantly, these pathways, initiated by the ubiquitylation of the replication factor PCNA (Hoege et al, 2002) and involving either translesion synthesis by specialized, damage-tolerant polymerases or a recombination-like process named template switching, are not necessarily coupled to replication fork progression. They can be delayed without major effects on genome stability until bulk genome replication is completed (Ulrich, 2009; Daigaku et al, 2010; Karras & Jentsch, 2010), although an impact on the transmission of epigenetic information has been reported (Sarkies et al, 2010). Under these conditions, daughter-strand gaps accumulate and give rise to a damage response, accompanied by a cell cycle arrest in G2/M phase (Lopes et al, 2006; Callegari et al, 2010; Daigaku et al, 2010). When damage bypass is re-activated at that point, the pathway mediates the filling of these gaps in a postreplicative manner (Daigaku et al, 2010; Karras & Jentsch, 2010). The significance of re-priming and daughter-strand gap formation for checkpoint signaling in WT cells is not well understood. A postreplication checkpoint that senses unreplicated DNA has been postulated (Callegari & Kelly, 2006), and Balint et al (2015) have described the assembly of a Mec1-activating complex distal to replication forks in response to DNA damage induced by the alkylating agent methyl methanesulfonate (MMS). However, the notion of postreplicative checkpoint activation contradicts the established concept of fork uncoupling, which invokes the stalled replication fork as the source of ssDNA that activates checkpoint signaling (Walter & Newport, 2000; Byun et al, 2005). In order to resolve this conflict, we made use of a genetic tool to delay damage bypass, thus causing a damage-dependent hyper-accumulation of daughter-strand gaps (Daigaku et al, 2010). In this setting, we identified an Exo1-dependent mechanism of Rad53 activation that in turn prevents erosion of gaps and an irreversible loss of viability largely attributable to the unrestrained activities of Exo1 and Pif1. Although reminiscent of the replication-independent action of Exo1 at DNA termini and NER gaps, this process required entry into S phase. Importantly, the same Exo1-dependent mechanism of Rad53 activation was observed in damage bypass-competent cells specifically during replication of damaged DNA, but not in response to nucleotide depletion or replisome problems. These findings explain the dichotomy between Mrc1- and Rad9-dependent Rad53 activation and suggest two distinct, spatially segregated mechanisms of how replication stress causes checkpoint activation: a fork-associated, Mrc1-dependent, Exo1-independent reaction in response to replisome-inherent problems and a gap-associated, Rad9- and Exo1-dependent process that predominates under conditions of template-induced polymerase stalling. We conclude that even in bypass-competent cells, regions of ssDNA left behind in the wake of replication forks and expanded by the action of processing factors such as Exo1, rather than stalled replication forks per se, constitute the predominant signal that leads to checkpoint activation in response to polymerase-stalling DNA lesions during S phase. Results Rad9-mediated checkpoint signaling is essential for damage resistance in the absence of damage bypass In order to systematically explore the relationship between checkpoint activation and damage bypass, we depleted Rad18, the ubiquitin ligase responsible for initiating the pathway (Hoege et al, 2002), thus enforcing hyper-accumulation of daughter-strand gaps during replication over lesions (Daigaku et al, 2010; Karras & Jentsch, 2010). In order to avoid the accumulation of suppressors, we used a regulable allele, Tet-RAD18, which conveys a rad18Δ-like phenotype only in the presence of doxycycline (Daigaku et al, 2010). We monitored the effects of Rad18 loss on defined checkpoint mutants with respect to three different types of genotoxic stress: the methylating agent MMS, which elicits a damage response primarily during replication, 4-nitroquinoline oxide (4NQO), which forms bulky adducts that are perceived in a replication-independent manner, and HU, which causes replication fork stalling by means of nucleotide depletion without inducing lesions in the replication template. Depletion of Rad18 strongly sensitized the checkpoint mutants mec1Δ, rad53Δ, and mrc1Δ rad9Δ toward MMS and 4NQO, confirming the importance of checkpoint signaling in the absence of damage bypass (Fig 1A). Synergism was also observable with rad9Δ alone, but not with mrc1Δ, suggesting that the DNA damage checkpoint—as opposed to the replication checkpoint—was responsible for the effect. In support of this model, depletion of Rad18 only mildly enhanced the sensitivity of any of the strains toward moderate concentrations of HU, indicating that replication fork stalling per se is not particularly detrimental in the absence of Rad18. A RAD18 deletion yielded comparable results (Fig EV1A). These findings imply a synergistic impact of damage bypass and specifically Rad9-dependent checkpoint signaling on the processing of DNA lesions. Figure 1. Contribution of checkpoint factors to DNA damage bypass DNA damage sensitivities of Tet-RAD18 strains carrying the indicated gene deletions, determined by growth assays in the presence (top) or absence (bottom) of Rad18. Experimental scheme for measuring recovery of viability after UV irradiation (20 J/m2) upon Tet-RAD18 induction at the indicated times after release into S phase (AS: asynchronous; αF: alpha-factor). For details, see Materials and Methods. Cell cycle profiles of the indicated strains at the time of plating. Survival of the indicated strains, relative to unirradiated controls. Error bars indicate SD derived from three independent experiments. Download figure Download PowerPoint Click here to expand this figure. Figure EV1. Contribution of checkpoint factors to DNA damage bypass DNA damage sensitivities of yeast strains carrying the indicated gene deletions, determined by growth assays. Re-expression of Rad18 (top) and HisPCNA ubiquitylation (bottom) after removing doxycycline in UV-irradiated cells of the indicated strains. Ubiquitylation of His6-tagged PCNA was detected as described previously (Daigaku et al, 2010). Experimental scheme for measuring viability after UV irradiation (20 J/m2) at the indicated times after release into S phase, performed as described in Fig 1B, but under conditions of continuous Tet-RAD18 expression (AS: asynchronous; αF: alpha-factor). Survival of the indicated strains, relative to unirradiated controls. Survival above 100% reflects cell division within 4 h. Recovery of viability is abolished in a catalytically inactive rad53 mutant. Magnification of selected panels from Fig 1D. Rad53 phosphorylation in the specified strains upon release into S phase after UV irradiation in the absence of Rad18, monitored by Western blotting. Data information: (D–F) Error bars indicate SD derived from at least three independent experiments. Source data are available online for this figure. Download figure Download PowerPoint Rad9-mediated checkpoint signaling maintains damage bypass competence during S phase Rad18 is a rate-limiting factor for PCNA ubiquitylation. Hence, the Tet-RAD18 allele allows us to modulate the activation of the damage bypass pathway at will in the course of a cell cycle. In this manner, we had previously shown that synchronized cells, treated in the G1 phase with low doses of ultraviolet (UV) radiation in the absence of Rad18, replicate the bulk of their genomes, but stall in G2/M phase with an activated checkpoint due to the hyper-accumulation of daughter-strand gaps (Daigaku et al, 2010). RAD18 re-expression at any time during or after genome replication allows them to recover, indicating that postreplicative gap filling can substitute for replication-associated damage bypass. We now used this approach to examine the mechanism of checkpoint activation under conditions of daughter-strand gap hyper-accumulation (Fig 1B): Alpha-factor (αF)-arrested G1 cells were UV-irradiated in the absence of Rad18 and subsequently released into S phase. RAD18 expression was then induced either immediately upon release, in mid-S phase, or after cells had reached G2/M phase (Fig 1C), and survival was determined by plating of aliquots. As previously reported, checkpoint-proficient (WT) cells recovered viability independently of the timing of RAD18 induction (Daigaku et al, 2010). In contrast, mec1Δ and rad53Δ mutants were completely unable to recover (Fig 1D). In the case of mec1Δ, the defect might be ascribed to a direct participation of the kinase in translesion synthesis by phosphorylation of Rev1 (Pages et al, 2009), but for rad53Δ this does not apply. Here, the defect was neither due to a failure to re-express RAD18 (Fig EV1B) nor caused by the UV sensitivity conferred by the rad53Δ mutation itself, as viability remained consistently higher when the assay was performed in the continuous presence of Rad18 (Fig EV1C and D). Hence, in rad53Δ cells even a temporary absence of Rad18 appears to cause a complete and irreversible loss of the capacity to productively use damage bypass. The ability to recover by RAD18 induction depended on the kinase activity of Rad53, as a catalytically deficient mutant, rad53-K227A, did not regain viability (Fig EV1E). As Rad53 can be activated either via the Rad9-dependent damage checkpoint or the Mrc1-dependent replication checkpoint (Pardo et al, 2017), we examined recovery of viability in rad9Δ and mrc1Δ mutants. As shown in Fig 1D, the mrc1Δ mutant fully recovered upon RAD18 re-expression, whereas deletion of RAD9 caused a significant loss of viability that grew successively more severe with a prolonged delay of RAD18 induction. In contrast, deletion of RAD9 or MRC1 in the presence of RAD18 had little effect on viability (Fig EV1D). This observation suggests that the Rad9-mediated damage checkpoint, rather than the Mrc1-dependent replication checkpoint, is essential to maintain damage bypass competence as cells progress through S phase. An mrc1Δ rad9Δ double mutant phenocopied rad53Δ mutants (Figs 1D and EV1F), indicating that Mrc1-mediated checkpoint signaling may partially compensate for the loss of Rad9 during early S phase. Consistent with this model, Rad53 phosphorylation was severely reduced upon RAD9 deletion, but completely abolished in the mrc1Δ rad9Δ double mutant (Fig EV1G). Hence, our findings suggest that Rad9-mediated activation of the Rad53 kinase becomes essential when damage bypass is delayed. Delay of damage bypass in rad53Δ mutants causes elevated homologous recombination and catastrophic chromosome fragmentation We next sought to elucidate how checkpoint mutants lost viability upon inhibition of damage bypass. Using pulsed-field gel electrophoresis (PFGE), we found that UV-irradiated rad53Δ cells grown in the absence of Rad18 failed to restore the pattern of intact chromosomes indicative of successful completion of genome replication (Fig 2A). Instead, we observed substantial chromosome fragmentation in the course of S phase (Figs 2A and EV2A). This was also observed in rad9Δ, but not in mrc1Δ cells (Fig EV2B). Consistent with the fragmentation pattern, rad53Δ cells in the absence of Rad18 accumulated strongly elevated numbers of recombination foci and exhibited aberrant chromosome segregation patterns (Figs 2B and C, and EV2C and D). Importantly, both chromosome breaks and hyper-accumulation of Rad52YFP foci were only observed in the absence of Rad18. From these observations, we conclude that when damage bypass fails, Rad9-mediated checkpoint signaling is essential to prevent massive chromosome fragmentation during S phase and—likely as a consequence of this—an elevated frequency of aberrant or failed divisions. Similar defects had been reported in rad18Δ cells in response to low doses of chronic damage (Hishida et al, 2009). Figure 2. Delay of damage bypass in rad53Δ mutants causes chromosome fragmentation, excessive recombination, and aberrant divisionWT and rad53Δ cells were grown in the presence or absence of Rad18, synchronized in G1, UV-irradiated, and released into S phase. Yeast chromosomes, analyzed by pulsed-field gel electrophoresis and ethidium bromide staining (top) or Southern blotting for chromosome V (middle). Replication intermediates accumulate in the wells. Cell cycle profiles are shown below the respective strains. Quantification of Rad52YFP recombination foci and representative images. Error bars indicate SD derived from three independent experiments. Scale bar = 5 μm. Analysis of mitotic aberrations. Cells were classified into cell cycle stages according to spindle morphology (see Fig EV2C and D for examples). Download figure Download PowerPoint Click here to expand this figure. Figure EV2. Consequences of delaying damage bypass in checkpoint mutants Pulsed-field gel electrophoresis and Southern blotting analysis of chromosome IV in WT and rad53Δ released into S phase after UV irradiation in the presence or absence of Rad18. Replication intermediates accumulate in the wells. Pulsed-field gel electrophoresis analyzed by ethidium bromide staining (top) and Southern blotting (middle) for chromosome IV in WT, mrc1Δ, and rad9Δ in the absence of Rad18, treated as above. Cell cycle profiles are shown at the bottom. Classification of cells into cell cycle stages according to spindle morphology (blue: DAPI: green: tubulin). Scale bar = 5 μm. Examples of cells undergoing aberrant divisions. Download figure Download PowerPoint Rad53 is required during S phase, but not for postreplicative gap filling The failure of rad53Δ mutants to reactivate damage bypass might be due to a direct requirement of Rad53 for the filling of daughter-strand gaps. However, the successive loss of chromosome integrity in the course of S phase suggested an essential function of checkpoint signaling already at the stage where the gaps emerge. In order to distinguish between these models, we used a previously characterized allele, rad53AID*−9myc, that encodes the kinase as a fusion with an auxin-inducible degron (Morawska & Ulrich, 2013). This allows depletion of the protein within < 1 h and confers a rad53Δ-like phenotype in the presence, but WT behavior in the absence of auxin (Morawska & Ulrich, 2013; Appendix Fig S1A). With this allele, we re-examined the recovery of viability under conditions where Rad53AID*−9myc was depleted either prior to the start of S phase (Fig 3A) or after passage through S phase, but before reactivation of Rad18 (Fig 3B). As expected, when Rad53AID*−9myc was removed prior to UV treatment, recovery was strongly compromised (Fig 3A). The defect was not as severe as in a rad53Δ strain, but this may have been due to residual protein even in the presence of auxin. Recovery was normal in the absence of auxin (Appendix Fig S1B), indicating that the failure to restore viability was indeed a consequence of the degradation of Rad53, and the AID*-tagged protein was functional under stabilizing conditions. Degradation of Rad53AID*−9myc after completion of S phase had no detrimental effect on viability, suggesting that Rad53 function is dispensable for damage bypass in G2/M (Fig 3B and Appendix Fig S1C and D). Figure 3. Rad53 is required during the S phase that precedes DNA damage bypass Loss of Rad53 before S phase: recovery assays upon RAD18 induction were performed as described in Fig 1B, but Rad53AID*−9myc degradation was induced by adding auxin during synchronization. Loss of Rad53 in G2/M phase: assays were performed as above, but Rad53AID*−9myc degradation was induced 4 h after release into S phase. Transient loss of Rad53: recovery was measured after Rad53AID*−9myc degradation during synchronization and re-expression together with RAD18 at the indicated times during the cell cycle. Data information: (A–C) Error bars indicate SD derived from at least three independent experiments. Download figure Download PowerPoint We then set up an experiment where Rad53AID*−9myc was temporarily degraded before release from G1 but re-expressed together with Rad18 at different times during the cell cycle (Fig 3C, -Aux). When rad53AID*−9myc was re-expressed before entry into S phase (0 h), cells recovered viability. However, if re-expression was postponed to mid-S (2 h) or G2 phase (4 h), loss of viability became irreversible, even though recovery of Rad53AID*−9myc protein levels resulted in a restoration of checkpoint signaling (assessed by the upregulation of the ribonucleotide reductase subunit, Rnr4; Appendix Fig S1E). Taken together, these data indicate that a transient loss of Rad53 during replication is sufficient to irreversibly prevent productive damage bypass. Rad53-mediated inhibition of Exo1 and Pif1 maintains bypass competence during S phase In order to identify the mechanism(s) by which Rad53 maintains damage bypass competence, we systematically examined possible contributions of Rad53's downstream targets. We found that upregulation of dNTP levels was required for efficient damage bypass, but not sufficient to restore viability in rad53Δ cells (Fig EV3A). Abolishing the suppression of late origin firing (Lopez-Mosqueda et al, 2010; Zegerman & Diffley, 2010) in a Rad53-proficient background strongly accelerated progression through S phase, but did not interfere with viability (Fig EV3B). Vice versa, delay of mitosis by nocodazole treatment did not rescue viability in the absence of Rad53 (Fig EV3C). We were also able to exclude a contribution of Rad53-induced gene expression controlled by the transcriptional co-repressor Nrm1 (de Bruin et al, 2006; Travesa et al, 2012; Fig EV3D) and an influence of histone gene dosage, which had also been shown to affect the damage sensitivity of rad53Δ mutants (Gunjan & Verreault, 2003; Fig EV3E). Finally, in order to assess whether elevated homologous recombination was the underlying cause of the problems or rather a reflection of (unsuccessful) attempts at repair, we analyzed the effects of various mutants defective in distinct stages of homologous recombination, such as mre11Δ, rad55Δ, mms4Δ, slx4Δ, yen1Δ, sgs1Δ, and srs2Δ. However, none of them restored Rad18-mediated survival in a rad53Δ background (Fig EV4). Click here to expand this figure. Figure EV3. Analysis of downstream targets of Rad53Recovery assays were performed according to the scheme in Fig 1B, by releasing UV-irradiated G1 cells into S phase in the absence of Rad18 and inducing Tet-RAD18 expression at the indicated times. In response to replication stress, Rad53 upregulates dNTP production through phosphorylation of Dun1, which in turn targets the transcriptional repressor Crt1 and the protein inhibitors Sml1 and Dif1, thereby relieving ribonucleotide reductase (RNR) inhibition and boosting dNTP levels (Hustedt et al, 2013) (left). Deletion of DUN1 reduces the capacity of cells to recover, and this effect is suppressed partially by deletion of SML1 and almost completely by a concomitant deletion of SML1 and CRT1 (middle). Hence, upregulation of dNTP levels is required for efficient damage bypass. However, sml1Δ and crt1Δ do not compensate for loss of RAD53 (right). Therefore, upregulation of dNTP levels is required, but not sufficient to maintain bypass competence. In response to replication stress, Rad53 inhibits late origin firing through phosphorylation of Dbf4 and Sld3 (Lopez-Mosqueda et al, 2010; Zegerman & Diffley, 2010) (left). Cells expressing non-phosphorylatable alleles of DBF4 and SLD3 (dbf4-4A sld3-A) (Zegerman & Diffley, 2010) recover virtually as efficiently as WT cells (middle), even though they progress through S phase as rapidly as the rad53Δ mutant (right). Hence, accelerated progression through S phase or firing of late origins does not preclude efficient postreplicative gap filling. Delay of mitosis by nocodazole treatment is insufficient to restore viability in rad53Δ. Middle: recovery assays performed with or without addition of nocodazole (added twice: 15 μg/ml at 0 h and 10 μg/ml at 2 h). Right: magnification of the graphs showing rad53Δ. Rad53 is responsible for induction of a set of MBF-regulated genes during S phase in response to DNA damage, mediated via inactivation of the transcriptional co-repressor Nrm1 (left) (Travesa et al, 2012). Hence, NRM1 deletion suppresses rad53Δ-associated lethality (de Bruin et al, 2006). However, nrm1Δ does not restore Rad18-dependent recovery of viability in rad53Δ (right), indicating that Nrm1 is not a relevant target of Rad53 in this context. Rad53 is required for degradation of excess histones upon DNA damage (Gunjan & Verreault, 2003) (left). Reduction in histone gene dosage therefore suppresses HU and MMS sensitivities of rad53Δ (Gunjan & Verreault, 2003). However, reducing histone dosage by deletion of HHT2 and HHF2 does not restore damage bypass competence in a rad53Δ background (right), thus excluding histone dosage as a critical factor. Data information: (A–E) Error bars indicate SD derived from three independent experiments.