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. 2021 Jan 5;10:e63589. doi: 10.7554/eLife.63589

Checkpoint inhibition of origin firing prevents inappropriate replication outside of S-phase

Mark C Johnson 1, Geylani Can 1, Miguel Monteiro Santos 1, Diana Alexander 1, Philip Zegerman 1,✉
Editors: Tim Formosa2, Kevin Struhl3
PMCID: PMC7806266  PMID: 33399537

Abstract

Checkpoints maintain the order of cell cycle events during DNA damage or incomplete replication. How the checkpoint response is tailored to different phases of the cell cycle remains poorly understood. The S-phase checkpoint for example results in the slowing of replication, which in budding yeast occurs by Rad53-dependent inhibition of the initiation factors Sld3 and Dbf4. Despite this, we show here that Rad53 phosphorylates both of these substrates throughout the cell cycle at the same sites as in S-phase, suggesting roles for this pathway beyond S-phase. Indeed, we show that Rad53-dependent inhibition of Sld3 and Dbf4 limits re-replication in G2/M, preventing gene amplification. In addition, we show that inhibition of Sld3 and Dbf4 in G1 prevents premature initiation at all origins at the G1/S transition. This study redefines the scope of the ‘S-phase checkpoint’ with implications for understanding checkpoint function in cancers that lack cell cycle controls.

Research organism: S. cerevisiae

Introduction

It is vitally important that in every cell division, the entire genome is replicated once and only once. In eukaryotes, this is achieved by linking DNA replication control to the cell cycle (Siddiqui et al., 2013). The first step in replication is the formation of the pre-replicative complex (pre-RC) at origins – a process called ‘licensing’. Licensing involves the origin recognition complex 1-6 (Orc1-6) and Cdc6-dependent loading of double hexamers of the Mcm2–7 helicase on double-stranded DNA. Licensing is restricted to late mitosis/early G1 phase by the activity of the APC/C (anaphase-promoting complex/cyclosome), which eliminates licensing inhibitors such as cyclin-dependent kinase (CDK) and geminin in this window of the cell cycle. In the budding yeast Saccharomyces cerevisiae, which lacks geminin, CDK inhibits licensing from late G1 phase until mitosis by multiple mechanisms, including direct phosphorylation of Orc2/Orc6 and nuclear exclusion of the Mcm2–7 complex, and by mediating SCFCDC4-dependent degradation of Cdc6 (Blow and Dutta, 2005; Nguyen et al., 2001).

Importantly, Mcm2–7 double hexamers loaded in late M/early G1 phase are inactive, and replication initiation can only occur after the inactivation of the APC/C at the G1/S transition. APC/C inactivation allows the accumulation of S-phase CDK and Dbf4-dependent kinase (DDK) activities (Labib, 2010). DDK directly phosphorylates the inactive Mcm2–7 double hexamers, generating a binding site for firing factors including Sld3/Sld7 and Cdc45, while CDK phosphorylates Sld3 and an additional initiation factor Sld2, which via phospho-interactions with Dpb11, results in replisome assembly by poorly understood mechanisms (Riera et al., 2017). This duality of function of CDK, both as an inhibitor of licensing and as an activator of the replisome, is critical to ensure once per cell cycle replication (Diffley, 2004).

In light of the importance of the linkage between DNA replication control and cell cycle progression, multiple checkpoints exist to regulate DNA synthesis and genome integrity before (G1 checkpoint), during (S-phase checkpoint), and after S-phase (G2/M checkpoint, Hartwell and Weinert, 1989; Kastan and Bartek, 2004). These checkpoints are mediated by the PI3 kinase superfamily checkpoint kinases ATM/ATR (Ataxia telangiectasia mutated/ATM and RAD3-related, called Tel1/Mec1 in budding yeast) and the effector checkpoint kinases Chk1/Chk2 (Chk1/Rad53 in budding yeast).

In G1 phase, DNA damage such as UV photoproducts causes checkpoint-dependent delays in the onset of DNA replication by inhibition of G1/S cyclin–CDK activity (Lanz et al., 2019; Shaltiel et al., 2015). In humans, the G1–S transition is delayed by ATM-Chk2-mediated stabilisation of p53, resulting in expression of the CDK inhibitor p21, as well as by checkpoint-dependent degradation of cyclin D and the CDK activator Cdc25A (Lanz et al., 2019; Shaltiel et al., 2015). In budding yeast, the delay in G1–S occurs in part by Rad53-dependent phosphorylation and inhibition of the Swi6 subunit of the transcriptional activator SBF (SCB binding factor) leading to reduced cyclin transcription (Sidorova and Breeden, 1997).

In S-phase, origin firing in unperturbed cells occurs as a continuum throughout S-phase, with some origins firing in the first half (early origins) or the second half of S-phase (late origins). When replication forks emanating from early firing origins stall, for example due to DNA lesions, activation of the S-phase checkpoint kinase response results in the dramatic slowing of replication rates (Painter and Young, 1980; Paulovich and Hartwell, 1995), which occurs in large part through inhibition of late firing origins (Yekezare et al., 2013).

Similar to G1 phase, checkpoint activation in S-phase in human cells can prevent origin firing through inhibition of CDK activity, for example by Chk1- and Chk2-mediated degradation of the CDK activator Cdc25A (Bartek et al., 2004). In budding yeast however, the checkpoint kinase Rad53 blocks late origin firing, not by inhibition of CDK activity, but by directly inhibiting two replication initiation factors: the DDK subunit Dbf4 and the CDK target Sld3 (Lopez-Mosqueda et al., 2010; Zegerman and Diffley, 2010). Rad53-dependent phosphorylation of Sld3 inhibits its interactions with various other replication factors including Dpb11 (Lopez-Mosqueda et al., 2010; Zegerman and Diffley, 2010), while recent work has shown that direct interaction between Rad53 and Cdc7-Dbf4 prevents DDK from binding to the Mcm2–7 complex (Abd Wahab and Remus, 2020). In higher eukaryotes, in addition to inhibition of CDK activity, the checkpoint can also inhibit origin firing through direct targeting of the Sld3 orthologue Treslin (Boos et al., 2011; Guo et al., 2015) and DDK (Costanzo et al., 2003; Lee et al., 2012). Therefore, multiple mechanisms exist to ensure the robust inhibition of replication initiation in S-phase by direct targeting of Sld3/Treslin and Dbf4-dependent kinase and by inhibition of CDK activity (Lanz et al., 2019). One function of such robust inhibition of origin firing during S-phase in the presence of DNA lesions is to prevent the exhaustion of essential factors, such as topoisomerase activities, by excessive numbers of replisomes (Morafraile et al., 2019; Toledo et al., 2017).

A key proposed feature of the DNA damage checkpoints is that the response is tailored to the cell cycle phase in which the DNA damage occurred (Shaltiel et al., 2015). Despite this, there is very little evidence to suggest that substrate specificity of the checkpoint kinases changes during the cell cycle. Indeed, in budding yeast, most forms of DNA damage and replication stress converge on the single effector kinase Rad53, but how different checkpoint responses in different cell cycle phases can be mediated by a single kinase is not known. In this study, we set out to explore the specificity of Rad53 towards the replication substrates Sld3 and Dbf4 across the cell cycle in the budding yeast, S. cerevisiae. We show that Rad53 phosphorylates both of these substrates throughout the cell cycle at the same sites as in S-phase. From this we hypothesised that although these substrates are deemed to be targets of the ‘S-phase checkpoint’, Rad53 may also prevent aberrant origin firing outside of S-phase. Indeed, we show that Rad53-dependent inhibition of Sld3 and Dbf4 limits re-initiation of replication in G2/M phase and also prevents premature firing of all origins, not just late origins, at the G1/S transition. This study overhauls our understanding of the cell cycle phase specificity of the ‘S-phase checkpoint’ and provides a novel mechanism that restricts replication initiation to a specific window of the cell cycle after DNA damage.

Results and discussion

Sld3 and Dbf4 are phosphorylated by Rad53 outside of S-phase

Since the DNA damage checkpoint response can be activated in all phases of the cell cycle, we addressed whether the replication factors Sld3 and Dbf4 could be targeted by Rad53 outside of S-phase in vivo in budding yeast. To test this, we first analysed the consequences of DNA damage in G1 phase cells arrested with the mating pheromone alpha factor (Figure 1A). These experiments were conducted with strains containing a null mutation in the alpha factor protease, bar1∆, to ensure that cells were fully arrested in G1 phase and had not started DNA replication. Addition of the UV mimetic drug 4-NQO to G1 phase cells resulted in robust Rad53 activation, as determined by the accumulation of the phospho-shifted forms of the kinase (Figure 1B,C). Importantly, we observed a dramatic increase in lower mobility forms of Sld3 when Rad53 was activated in G1 phase (Figure 1B), which was indeed Rad53 dependent (Figure 1—figure supplement 1B). In line with a previous study (Mattarocci et al., 2014), in G1-arrested cells in the absence of 4-NQO (Figure 1B) or in rad53∆ cells (Figure 1—figure supplement 1B), Sld3 can be seen to migrate as a doublet on phos-tag gels. This was shown to be caused by DDK phosphorylation of Sld3 (Mattarocci et al., 2014), but does not affect the DNA damage and Rad53-dependent phosphorylation of Sld3 described here (Figure 1B). For Dbf4, which is an APC/C substrate and partially degraded in alpha factor-arrested cells (Ferreira et al., 2000), we also observed a mobility shift in G1 phase coincident with Rad53 activation (Figure 1C).

Figure 1. Dbf4 and Sld3 are phosphorylated by Rad53 after DNA damage in G1 and G2 phase.

(A) Flow cytometry of strains arrested in G1 phase with the mating pheromone alpha factor. Strains were held in G1 phase, with or without the addition of 10 μg/ml 4-NQO for the indicated times. All strains are bar1∆ to maintain G1 arrest. (B) Western blot of Sld3 (anti-myc) and Rad53 phosphorylation from the experiment outlined in (A). Sld3 was resolved on a phos-tag sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) gel. (C) As (B), but for Dbf4. Both blots are from SDS–PAGE. (D) As (A), except strains were arrested in G2/M with nocodazole before the addition of 4-NQO. All strains are sml1∆, which is required for viability in cells lacking Rad53. (E) Western blot of Sld3 (anti-myc) and Rad53 phosphorylation as in (B) from the experiment outlined in (D). Sld3 was resolved on a phos-tag SDS–PAGE gel. *Cyclin-dependent kinase (CDK)-phosphorylated Sld3 (see Figure 1—figure supplement 1C). (F) As (E), but for Dbf4. Dbf4 is phosphorylated by other kinases in G2/M, resulting in residual phosphorylated forms remaining in rad53∆ cells *.

Figure 1.

Figure 1—figure supplement 1. Sld3 phosphorylation in G1 phase after DNA damage is Rad53 dependent.

Figure 1—figure supplement 1.

(A) Flow cytometry of strains arrested in G1 phase with the mating pheromone alpha factor. Strains were held in G1 phase, with the addition of 10 μg/ml 4-NQO for the indicated times. All strains are bar1∆ to maintain G1 arrest and also sml1∆. (B) Western blot of Sld3 and Rad53 phosphorylation from the experiment outlined in (A). Sld3 was resolved on a phos-tag sodium dodecyl sulfate–polyacrylamide gel electrophoresis gel. (C) Western blot of Sld3 from cells arrested in G2/M with nocodazole. These strains contain the cdc28-as1 allele that is inhibited by the addition of the ATP analogue 1-NM-PP1. As Cdc28 is the only catalytic CDK subunit in yeast, the addition of 1-NM-PP1 inhibits CDK activity, revealing that the lower mobility Sld3 band is CDK dependent. The first lane contains untagged Sld3, showing that all bands in this blot correspond to Sld3-13myc.
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To test whether Sld3 and Dbf4 could also be phosphorylated by Rad53 after DNA replication is complete, we performed the same experiment as in Figure 1A–C, except in cells arrested in G2/M phase with nocodazole (Figure 1D). Significantly, we observed a Rad53-dependent mobility shift in Sld3 and Dbf4, even in G2/M-arrested cells (Figure 1E,F). Sld3 and Dbf4 are phosphorylated by other kinases in G2/M, such as by CDK (Holt et al., 2009), giving rise to additional isoforms of Sld3/Dbf4 proteins even in Rad53 null cells (* Figure 1E,F). Note that the CDK-phosphorylated form of Sld3 is visible in Figure 1E as this is a phos-tag gel, and we confirmed that both of the bands in rad53∆ cells in Figure 1E are Sld3 dependent and the upper form is indeed CDK dependent (Figure 1—figure supplement 1C).

Previously, we have identified the serine and threonine residues in Sld3 and Dbf4 that are directly phosphorylated by Rad53 in S-phase (Zegerman and Diffley, 2010). We mapped 38 such phospho-sites in Sld3 and 19 sites in Dbf4. Mutation of these serine/threonine residues to alanine in Sld3- and Dbf4-generated alleles that are refractory to Rad53 phosphorylation in S-phase (Zegerman and Diffley, 2010) and are hereafter referred to as sld3-A and dbf4-19A, respectively. We reasoned that if the same sites in Sld3 and Dbf4 are phosphorylated by Rad53 throughout the cell cycle, then sld3-A and dbf4-19A should be defective in Rad53-dependent phosphorylation in G1 and G2/M as well. In G1 phase, the Sld3-A protein demonstrated a dramatic loss of Rad53 phosphorylation (Figure 2A), consistent with direct phosphorylation of Sld3 by Rad53 at the same sites as in S-phase. We also observed a similar result with the Dbf4-19A protein in G1 phase after Rad53 activation (Figure 2B). As in G1 phase, both Sld3-A and Dbf4-19A showed greatly reduced phosphorylation during Rad53 activation in G2/M phase (Figure 2C,D). Together, Figures 1 and 2 show that although Sld3 and Dbf4 are considered to be ‘S-phase checkpoint’ substrates of Rad53, they are phosphorylated at the same sites as in S-phase after DNA damage in G1 and G2 phase.

Figure 2. Rad53 phosphorylates Sld3 and Dbf4 in G1 and G2 phases at the same residues as in S-phase (A and B) as Figure 1B,C.

sld3-A and dbf4-19A refer to mutant alleles with Rad53 phosphorylation sites mutated to alanine (38 sites for Sld3 and 19 sites for Dbf4). (C) As Figure 1E, except this is western blot from an SDS–PAGE gel. (D) As Figure 1F.

Figure 2.

Figure 2—figure supplement 1. Dbf4-4A is expressed at similar levels to wild type and is also refractory to Rad53 phosphorylation in G1 phase.

Figure 2—figure supplement 1.

Western blot of strains arrested in G1 phase with the mating pheromone alpha factor. Strains were held in G1 phase, with the addition of 10 μg/ml 4-NQO for the indicated times. All strains are bar1∆ to maintain G1 arrest. As is from an asynchronous culture. The dbf4-4A mutant contains four serine/threonine to alanine mutations in the phospho-sites that are critical for Rad53-dependent inhibition of Dbf4. dbf4-4A is referred to as dbf4-A in the rest of this manuscript.
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The dbf4-19A mutant, although refractory to Rad53 phosphorylation, is also a hypomorph and is not fully functional for replication initiation (Zegerman and Diffley, 2010). We previously narrowed down the critical residues that are inhibited by Rad53 to four sites and generated a dbf4-4A allele, which still cannot be inhibited by Rad53, but is fully functional for its role in replication initiation (Zegerman and Diffley, 2010). In similarity to the 19A allele, Dbf4-4A was less Rad53-phosphorylated than wild type in G1 cells after DNA damage (Figure 2—figure supplement 1), suggesting that it is indeed the critical residues in Dbf4 that are phosphorylated by Rad53 outside of S-phase. As this 4A allele is fully functional for Dbf4’s role in replication initiation, from here on we only use this allele and refer to it as dbf4-A.

Rad53-dependent phosphorylation of Sld3 and Dbf4 reduces re-replication in G2 phase

We have previously shown that Rad53 phosphorylates Sld3 and Dbf4 in S-phase to inhibit origin firing (Lopez-Mosqueda et al., 2010; Zegerman and Diffley, 2010). As Sld3 and Dbf4 are phosphorylated at the same sites by Rad53 after DNA damage in both G1 and G2 phase (Figures 1 and 2), we wondered whether this phosphorylation could also be required to inhibit replication initiation outside of S-phase. DNA replication is tightly restricted to S-phase in large part by the action of CDK, which prevents licensing outside of late M/early G1 phase. As a result, transient reduction of CDK activity in G2/M phase is sufficient to induce re-replication (Dahmann et al., 1995). To test a role for Rad53 phosphorylation of Sld3/Dbf4 in re-replication control, we first combined the sld3-A/dbf4-A alleles, which cannot be inhibited by Rad53 (Zegerman and Diffley, 2010), with a hypomorphic mutant of the CDK catalytic subunit Cdc28 (cdc28-as1). This allele of Cdc28 is analogue sensitive and is inhibited by the addition of the ATP-competitive inhibitor 1-NM-PP1. Interestingly, we observed that the sld3-A/dbf4-A alleles are synthetically sick with cdc28-as1 in the presence of sub-lethal doses of 1-NM-PP1 (Figure 3—figure supplement 1A), suggesting that inhibition of Sld3 and Dbf4 by Rad53 is important in cells that have reduced CDK activity.

To specifically test whether the Rad53-dependent inhibition of origin firing is important to prevent re-replication, we combined the sld3-A/dbf4-A alleles with mutants that circumvent the CDK-dependent inhibition of licensing. Over-expression of Cdc6, forcing the nuclear localisation of the Mcm2-7 complex (through an Mcm7-2xNLS fusion) and mutation of the CDK phosphorylation sites in ORC is sufficient to induce re-replication in G2/M phase (Finn and Li, 2013; Nguyen et al., 2001) and has been shown to induce Rad53 activation (Archambault et al., 2005; Green and Li, 2005). Importantly, conditional over-expression of licensing mutants that cannot be inhibited by CDK combined with sld3-A and dbf4-A led to an increase in the total re-replication in nocodazole arrested cells (Figure 3A – compare flow cytometry overlay red vs. black).

Figure 3. Checkpoint-dependent inhibition of origin firing prevents re-replication in G2 phase.

(A) Flow cytometry of the indicated strains grown overnight in YPraffinose, then arrested in G2/M with nocodazole. After addition of fresh nocodazole, 2% galactose was added for the indicated times to express the licensing mutants. Right, overlay between the 0 and 2 hr time points for the licensing mutant strain with (red) or without (black) the sld3-A/dbf4-A alleles. (B) Copy number analysis of Chromosome III from the experiment in (A), after 4 hr in galactose. As in (A), the strains overexpress wild type Cdc6 and the indicated pre-replicative complex mutants from the galactose inducible promoter. DNA sequencing read depth per 1 kb bin was normalised to the 0 hr time point for each strain. Baseline was set at two as the strains are arrested in G2. The y-axis shows the DNA copy number after these steps. (C) Schematic diagram of re-replication-induced gene amplification (RRIGA) assay for gene amplification events. Re-replication of the split LEU2 gene from origin ARS317 results in tandem head to tail gene duplications, leading to a functional LEU2 gene. (D) RRIGA assay in (C) was performed with the indicated strains. Strains were grown overnight in YPraffinose then arrested in G2/M with nocodazole (pre-induction, blue time point). After addition of fresh nocodazole, 2% galactose was added for 3 hr to express the licensing mutants (red time point). Cells were plated on YPD (viable cell count) and SC-leu plates (Leu+ count) and the % of Leu+ colonies out of the viable cell population was plotted. N = 3, error bars are SD. *The p value was calculated as 0.0481 using an unpaired t-test.

Figure 3.

Figure 3—figure supplement 1. Checkpoint-dependent inhibition of origin firing prevents re-replication in G2 phase.

Figure 3—figure supplement 1.

(A) Growth assay of the indicated strains. (B) Analysis of mitochondrial DNA content by sequencing. (C) Comparison of DNA copy number at all origins between the strain over-expressing the pre-replicative complex mutants (x-axis) vs the same strain also containing the sld3-A/dbf4-A alleles (y-axis). Each dot represents a different origin of replication, and the origins of Chromosome III are red. The green line is the line of equal copy number between the two strains and therefore dots above the line are more re-replicated in the presence of the sld3-A/dbf4-A alleles. Note that the Chr III origins are the most efficiently re-replicated in both strains, with almost all of them showing more replication due to the sld3-A/dbf4-A alleles. (D) Schematic diagram of the RRIGA assay. Endogenous LEU2 was replaced with a split LEU2 marker, separated by the re-replication origin ARS317. The two non-functional halves contain 417 bp of identity*. Re-initiation at ARS317 followed by fork-breakage and strand annealing at the regions of LEU2 homology results in gene amplification and the generation of a functional LEU2 gene. (E) Verification of RRIGA by PCR using the amplicons as depicted in (A). The parental strain (which is Leu–) was induced to re-replicate in the presence of galactose and cells were plated on SC-leu plates. Ten independent Leu+ colonies were assayed for the presence of the duplication and all were positive.
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To analyse whether this shift in DNA content by flow cytometry was indeed due to genomic re-replication, we performed DNA sequencing and copy number analysis of the experiment in Figure 3A. While we did not detect any differences between the mitochondrial DNA content of the strains (Figure 3—figure supplement 1B), we observed a dramatic increase in DNA copy number largely for Chromosome III in strains that over-express Cdc6 and pre-RC mutants that cannot be inhibited by CDK (Figure 3B, Figure 3—figure supplement 1C). Previous studies have indeed shown that chromosome III is particularly susceptible to re-replication (Green et al., 2006). While we did not detect any re-replication in the sld3-A/dbf4-A strain alone (data not shown), we observed greater re-replication of chromosome III when the pre-RC mutants were combined with sld3-A and dbf4-A, which cannot be inhibited by Rad53 (Figure 3B). Together with Figure 3A, this suggests that Rad53-dependent inhibition of replication initiation can reduce inappropriate replication in G2 phase when licensing control is compromised.

One of the consequences of re-replication is the generation of head-to-tail tandem gene amplifications, a process termed re-replication-induced gene amplification (RRIGA) (Green et al., 2010). To examine whether the Rad53-dependent inhibition of origin firing helps to prevent RRIGA we adapted an assay to quantitatively assess gene amplification events in G2/M arrested cells (Finn and Li, 2013). Briefly, a marker gene (in this case LEU2, which allows growth on media lacking leucine) was split with some remaining homology across an origin that re-initiates when licensing control is lost (ARS317 – see for example Figure 3B). Re-initiation at ARS317 followed by fork-breakage and strand annealing at the regions of LEU2 homology results in gene amplification and the generation of a functional LEU2 gene (Figure 3C, Figure 3—figure supplement 1D,E). In this assay, as in Figure 3A, the re-replication mutants were induced only in G2/M-arrested cells. In contrast to wild-type yeast or the sld3-A/dbf4-A strain alone, expression of the licensing mutants by the addition of galactose resulted in a large increase in RRIGA events, as expected (Figure 3D). Importantly, RRIGA events were even greater when the mutants that allow licensing in the presence of CDK were combined with the sld3-A/dbf4-A alleles (Figure 3D). This assay demonstrates that the checkpoint kinase Rad53 indeed reduces gene amplification events after re-replication through inhibition of Sld3 and Dbf4, even in G2/M-arrested cells.

Rad53 prevents precocious origin firing after DNA damage in G1 phase

DNA damage in G1 phase delays the G1/S transition, which from humans to yeast, involves the checkpoint kinase-dependent down-regulation and/or inhibition of G1/S cyclin-CDK activity (Bertoli et al., 2013; Lanz et al., 2019; Shaltiel et al., 2015; Sidorova and Breeden, 1997). Here we have shown that DNA damage in G1 phase also results in the inhibitory checkpoint phosphorylation of two replication initiation factors, Sld3 and Dbf4 (Figures 1 and 2), suggesting that this might be an additional mechanism to prevent premature DNA replication at the G1/S transition (Figure 4A). To specifically analyse the consequences of DNA damage in G1 phase, we added 4-NQO to G1-arrested yeast cells and then released cells into S-phase in fresh medium without 4-NQO. Crucially, this approach resulted in robust Rad53 activation in G1 phase, such that cells enter S-phase with an already active checkpoint (Figure 4D, Figure 4—figure supplement 1). Rad53 activation in G1 phase resulted in the slowing of the G1/S transition as detected by the delay in budding (a G1 cyclin/CDK-mediated event) and the delay in DNA synthesis (Figure 4—figure supplement 1). Despite this, the sld3-A dbf4-A alleles caused little difference in S-phase progression after DNA damage in G1 phase, compared to the wild-type strain (Figure 4—figure supplement 1).

Figure 4. Checkpoint-dependent inhibition of origin firing prevents premature replication initiation at all origins at the G1–S transition.

(A) Activation of Rad53 in G1 phase can delay genome duplication by at least two mechanisms; by inhibition of the transcriptional activator Swi6, which is required for G1- and subsequently S-phase cyclin transcription and by inhibition of the origin firing factors Sld3 and Dbf4. (B) Flow cytometry of the indicated strains grown overnight in YPraffinose, then arrested in G1 phase with alpha factor. Cells were held in fresh alpha factor, while 2% galactose and 0.5 μg/ml 4-NQO was added for 30 min (0 time point) before washing and release from alpha factor arrest into fresh YPgal medium without 4-NQO. Swi4-t is a C-terminally truncated protein that cannot be inhibited by Rad53. (C) Budding index from the experiment in (B). Time point 0 refers to cells held in alpha factor + galactose + 0.5 μg/ml 4-NQO for 30 min. (D) Rad53 western blot from experiment in (B). (E) Copy number analysis of chromosome VII of the indicated strains 20 min after release as in (B–D). The y-axis ratio refers to the amount of DNA at the 20 min time point divided by the DNA copy number in G1 phase. Known origins are annotated above the replication profile and coloured according to their normal median replication time (Trep). (F) Box plots of the amount of replication at all origins, split into equal quintiles depending on their normal median firing time (Trep). Initiation occurs first at early origins in the swi4-t sld3-A dbf4-A strain, for example the yellow and green quintiles are significantly different at 20 min (*p value 0.044), but non-significantly different (ns) at 40 min. Arrows indicate that later firing origins also initiate by 40 min in the swi4-t sld3-A dbf4-A strain. P-values are from t-tests.

Figure 4.

Figure 4—figure supplement 1. 4-NQO addition in G1 phase delays the G1/S transition.

Figure 4—figure supplement 1.

(A) Flow cytometry of the indicated strains, which were grown overnight in YPraffinose and arrested in G1 phase with alpha factor (G1 time point). Strains were then held in G1 phase for an extra 30 min by the addition of fresh alpha factor plus galactose, with or without 0.5 μg/ml 4-NQO. Cells were washed into fresh YPgalactose medium to release into S-phase. Note that YPraffinose and galactose medium was used in this experiment to match the exact conditions that were used in the main Figure 4B with the PGAL-swi4-t strains, allowing a direct comparison, as in (D). (B) Budding index (a G1 cyclin/CDK-mediated event) from experiment in (A). (C) Rad53 western blot from the 4-NQO-treated samples in (A). (D) Overlay of budding profiles (from Figure 4C and this Figure B) between strains expressing or not expressing Swi4-t. Swi4-t, which cannot be inhibited by Rad53, causes earlier activation of G1 cyclin/CDK as shown here by budding.
Figure 4—figure supplement 2. Rad53 activation in G1 phase inhibits all origin firing through phosphorylation of Sld3 and Dbf4.

Figure 4—figure supplement 2.

Replication profile of chromosome VII as an example at 20, 30, and 40 min from the experiment in Figure 4B. Green * are examples of very early firing origins, which initiate at the 20 min time point in the PGAL-swi4-t sld3-A dbf4-A strain. Purple * are examples of late firing origins which do not initiate at the 20 min time point but are clearly firing by the 40 min time point in the PGAL-swi4-t sld3-A dbf4-A strain. This shows that the relative timing of origin firing is not affected in the PGAL-swi4-t sld3-A dbf4-A strain.
Figure 4—figure supplement 3. Genetic interactions of the sld3-A dbf4-A alleles with mutants that either accelerate or delay the G1/S transition.

Figure 4—figure supplement 3.

Scatter plot of the fitness of the yeast genome knock out collection grown in 0.5 μg/ml phleomycin with (y-axis) or without (x-axis) the sld3-A dbf4-A alleles. Each dot corresponds to a different gene deletion. The top 25% of gene deletions (yfg, your favourite gene) that significantly enhance (green) or suppress (red) the fitness of sld3-A dbf4-A are indicated. This data was originally published in Morafraile et al., 2019.
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As Rad53 is known to inhibit CDK activation through phosphorylation of the Swi6 subunit of the transcriptional activator SBF (Sidorova and Breeden, 1997), we wondered whether Rad53-dependent inhibition of both replication initiation factors and G1/S transcription might prevent precocious DNA replication after damage in G1 phase (Figure 4A). To test this, we over-expressed a truncated form of the SBF transcription factor Swi4 (Swi4-t), which lacks the C-terminus required for interaction with Swi6 and thus cannot be inhibited by Rad53 (Sidorova and Breeden, 1997). Over-expression of Swi4-t indeed resulted in faster progression through the G1/S transition in the presence of 4-NQO (Figure 4—figure supplement 1D) as expected (Sidorova and Breeden, 1997). Importantly, the combination of expression of Swi4-t together with sld3-A dbf4-A resulted in much faster S-phase progression after DNA damage in G1 phase compared to Swi4-t expression alone (Figure 4B). These differences in the onset of DNA replication between swi4-t with and without sld3-A/dbf4-A were not due to differences in the G1/S transition as these strains budded at the same time (Figure 4C) and both strains also exhibited similar levels of Rad53 activation (Figure 4D). Together this suggests that Rad53 activation in G1 phase prevents precocious DNA replication initiation by inhibiting not only G1/S transcription, but also Sld3 and Dbf4.

Activation of Rad53 during S-phase caused by fork stalling/DNA damage at early replicons results in inhibition of subsequent (late) origin firing, which in yeast is mediated by inhibition of Sld3 and Dbf4 (Lopez-Mosqueda et al., 2010; Zegerman and Diffley, 2010). We therefore wondered whether the accelerated S-phase we observe when we combine swi4-t with sld3-A dbf4-A is simply due to the canonical S-phase checkpoint inhibition of late origin firing or whether by activating Rad53 in G1 phase we are actually causing a delay in genome duplication from all origins. To assess this, we analysed the replication dynamics of the time course in Figure 4B–D by high-throughput sequencing and copy number analysis. At the earliest time point (20 min), while the swi4-t over-expressing strain alone had barely begun to replicate (Figure 4E, chromosome VII as an example), the swi4-t sld3-A dbf4-A strain showed peaks of replication initiation at the earliest firing origins (e.g., see *, Figure 4E), even though Rad53 is highly activated (Figure 4D). By analysis of initiation at all origins, split into quintiles according to their normal firing time, we observe much greater firing of early origins in the swi4-t sld3-A dbf4-A strain compared to swi4-t alone throughout the time course (Figure 4F, Figure 4—figure supplement 2). Over time, we also observe an increase in later firing origins in the swi4-t sld3-A dbf4-A strain (arrows Figure 4F, Figure 4—figure supplement 2), suggesting that the relative timing of origin firing is not affected. Together this demonstrates that activation of Rad53 and inhibition of Sld3 and Dbf4 in G1 phase contribute to the mechanism preventing the onset of DNA replication from all origins, not just late firing origins, in the presence of DNA damage (Figure 4A).

If the checkpoint-mediated inhibition of G1/S transcription and Sld3/Dbf4 contribute to prevent precocious S-phase entry, then we hypothesised that loss of both pathways should show synthetic lethality in the presence of DNA damage. We have previously conducted genetic interaction analysis of the sld3-A/dbf4-A alleles with the yeast whole genome gene knock-out collection in the presence of the DNA damaging agent phleomycin (Morafraile et al., 2019). Loss of function of genes that result in a delay in the G1/S transition, such as CLN2, SWI4, and BCK2 (Di Como et al., 1995), improved the growth of sld3-A/dbf4-A in the presence of phleomycin (suppressors, Figure 4—figure supplement 3), whereas loss of function of genes that would result in the acceleration of G1/S, such as WHI5 and SIC1 (Bertoli et al., 2013), were synthetic sick with the sld3-A/dbf4-A alleles (enhancers, Figure 4—figure supplement 3). These genetic interactions are consistent with an important role for Rad53-dependent inhibition of origin firing in preventing precocious replication after DNA damage in G1 phase.

Here we show that the two critical targets of the S-phase checkpoint-mediated inhibition of origin firing, Sld3 and Dbf4, are actually regulated by Rad53 after DNA damage throughout the cell cycle (Figures 1 and 2). This has important implications for understanding the consequences of inappropriate re-replication in human cells, where the role of the checkpoint differs depending on the cell cycle phase in which re-replication occurs (Klotz-Noack et al., 2012; Liu et al., 2007). By combining tight cell cycle arrests with the separation-of-function mutants, sld3-A dbf4-A, we show specifically that the checkpoint-dependent inhibition of origin firing limits further re-replication and gene amplifications in G2 phase when licensing control is compromised (Figure 3). This pathway is likely to be evolutionarily conserved, as checkpoint activation in S-phase in human cells also appears to limit re-replication through inhibition of Dbf4 (Lee et al., 2012). As tandem head-to-tail duplications are a prominent feature of many cancers (Menghi et al., 2018), knowledge of the pathways that prevent this form of structural variation may be important for understanding oncogenesis.

In addition to preventing re-replication in G2, we show that the checkpoint also inhibits the replication initiation factors Sld3 and Dbf4 to delay all origin firing after DNA damage in G1 phase (Figure 4). This likely increases the time for DNA repair to occur before replication begins and may also serve to increase the window of time where origin firing and licensing are mutually exclusive, preventing re-replication. It is interesting that failure to inhibit Sld3 and Dbf4 alone has little effect on the G1/S transition (Figure 4—figure supplement 1), probably because Sld3 (and Sld2) act downstream of CDK activation (Figure 4A). This study therefore provides evidence that two inter-connected pathways are important for restricting replication after DNA damage in G1: inhibition of G1/S CDK activity and direct Inhibition of replication initiation factors.

Mutations in genes such as Rb and p53 that control the G1/S transition and the G1 checkpoint response, respectively, are amongst the most common mutations in cancers (Malumbres and Barbacid, 2001; Massagué, 2004). Work from yeast to humans has shown that defects in the G1/S transition results in increased dependence on the checkpoint kinases for survival (Rundle et al., 2017; Sidorova and Breeden, 2002). The identification of the checkpoint inhibition of all origin firing as a failsafe mechanism that prevents precocious S-phase entry when the G1 checkpoint is compromised described here (Figure 4) may provide a potential mechanistic rationale for the selective targeting of p53/Rb mutant cancers using Chk1 and ATR inhibitors, which are currently in clinical trials (Bradbury et al., 2020).

Materials and methods

Key resources table.

Reagent type (species) or resource Designation Source or reference Identifiers Additional information
Antibody Anti-Rad53, rabbit polyclonal Abcam (ab104232) Antibody (1:5000)
Antibody Anti-c-myc, mouse monoclonal Merck Life Science (11667149001) Antibody (1:1000)
Peptide, recombinant protein Alpha-factor GenScript (RP01002) Peptide
Commercial assay or kit TruSeq Nano DNA Low Throughput Library Prep Kit Illumina (20015964) Commercial assay or kit
Chemical compound, drug Phos-tag Acrylamide AAL-107 Alpha Laboratories (304–93521) Chemical
Chemical compound, drug Nocodazole Merck Life Science (M1404) Chemical
Chemical compound, drug 4-Nitroquinoline N-oxide Merck Life Science (N8141) Chemical
Software, algorithm LocalMapper software Batrakou et al., 2020
Software, algorithm Repliscope software Batrakou et al., 2020
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Growth conditions and yeast strains

Cell growth, arrests, flow cytometry, and yeast protein extracts were performed as previously described (Zegerman and Diffley, 2010).

S. cerevisiae strains All the strains used in this work are derived from W303 (ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100, rad5-535).
SLD3 and DBF4 mutants that cannot be phosphorylated by RAD53 (sld3-38A and dbf4-4A) are abbreviated to sld3-A dbf4-A in the main text.
yPZ 1223 MATa SLD3-10his13myc::KanMX bar1∆::hisG
yPZ 1319 MATa sld3-38A-10his13myc::KanMX bar1∆::hisG
yPZ 1317 MATa DBF4-13myc::KanMX bar1∆::hisG
yPZ 705 MATa
yPZ 4076 MATa trp1::PGAL-swi4-t::TRP1
yPZ 4137 MATa dbf4-4A::HIS3 sld3-38A-10his13myc::KanMX trp1::PGAL-swi4-t::TRP1
yPZ 917 MATa dbf4-4A::HIS3 sld3-38A-10his13myc::KanMX
yPZ 1523 MATa sml1∆::URA3 rad53∆::LEU2 SLD3-10his13myc::KanMX bar1∆::hisG
yPZ 1522 MATa sml1∆::URA3 SLD3-10his13myc::KanMX bar1∆::hisG
yPZ 1198 MATa cdc28-as1 (F88G)
yPZ 1767 MATa cdc28-as1 (F88G) dbf4-4A::HIS3 sld3-38A-10his13myc::KanMX
yPZ 4014 MATa RRIGA-split LEU2 ChrIII (YCLWdelta5::HphMX-EU2-ARS317-LEU::NFS1), RAD5+ sld3-38A-10his13myc::KanMX dbf4-4A::HIS3
yPZ 4085 MATa RRIGA-split LEU2 ChrIII (YCLWdelta5::HphMX-EU2-ARS317-LEU::NFS1), RAD5+ trp1::orc6-4A-PGAL1-10-Mcm7-2xNLS::TRP1, ura3::PGAL-Cdc6-13myc::URA3
yPZ 4089 MATa RRIGA-split LEU2 ChrIII (YCLWdelta5::HphMX-EU2-ARS317-LEU::NFS1), RAD5+ trp1::orc6-4A-PGAL1-10-Mcm7-2xNLS::TRP1, ura3::PGAL-Cdc6-13myc::URA3 sld3-38A-10his13myc::KanMX dbf4-4A::HIS3
yPZ 125 MATa DBF4-13myc::KanMX
yPZ 170 MATa dbf4∆::TRP1 his3::PDBF4-dbf4-19A-13myc::KanMX::HIS3
yPZ 2 MATa sld3-38A-10his13myc::KanMX
yPZ 52 MATa SLD3-10his13myc::KanMX
yPZ 1471 MATa sml1∆::URA3 rad53∆::HphNT1 DBF4-13myc::KanMX bar1∆::hisG
yPZ 1473 MATa sml1∆::URA3 DBF4-13myc::KanMX bar1∆::hisG
yPZ 520 MATa sml1∆::URA3 SLD3-13myc::KanMX
yPZ 89 MATa sml1∆::URA3 rad53∆::LEU2 SLD3-13myc::KanMX
yPZ 519 MATa sml1∆::URA3 DBF4-13myc::KanMX
yPZ 228 MATa sml1∆::URA3 rad53∆::LEU2 DBF4-13myc::KanMX
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Yeast strain notes

dbf4-4A refers to the Rad53 site mutant. It has serine/threonine to alanine mutations at amino acids 518, 521, 526, 528.

dbf4-19A refers to the Rad53 site mutant. It has serine/threonine to alanine mutations at amino acids 53, 59, 188, 192, 203, 222, 224, 226, 228, 318, 319, 328, 374, 375, 377, 518, 521, 526, 528.

sld3-38A refers to the Rad53 site mutant. It contains serine/threonine to alanine mutations at amino acids 306, 310, 421, 434, 435, 438, 442, 445, 450, 451, 452, 456, 458, 459, 479, 482, 507, 509, 514, 519,521,524, 540, 541, 546, 547, 548, 550, 556, 558, 559, 565, 569, 582, 607, 653 and 654. 539 is mutated to arginine.

orc6-4A refers to CDK sites 106, 116, 123, and 146 mutated to alanine. Swi4-t refers to C-terminally truncated protein 1–814.

Western blot

Western blots were performed as previously described (Can et al., 2019). Rad53 was detected with ab104232 (Abcam, dilution 1:5000).

Replication profiles

Yeast genomic DNA was extracted using the smash and grab method (https://fangman-brewer.genetics.washington.edu/smash-n-grab.html). DNA was sonicated using the Bioruptor Pico sonicator (Diagenode), and the libraries were prepared according to the TruSeq Nano sample preparation guide from Illumina. To generate replication timing profiles, the ratio of uniquely mapped reads in the replicating samples to the non-replicating samples was calculated following Batrakou et al., 2020. Replication profiles were generated using ggplot and smoothed using a moving average in R. The values of Trep were taken from OriDB (Siow et al., 2012).

RRIGA assay using split LEU2 marker

Cells were pre-grown under permissive conditions in YPraff overnight 30°C. At 1 × 107 cells/ml, nocodazole (2 mg/ml in dimethyl sulfoxide) was added to a final concentration of 10 μg/ml. Cells were arrested for 90 min at 30°C (uninduced time point), and then galactose was added to a final concentration of 2% + fresh nocodazole for 3 hr (induced time point).

For each time point, a 0.5 ml sample was spun at 3.2 K for 1 min in benchtop centrifuge; cells were washed with 1 ml sterile water to remove YP, respun, and resuspended in 0.5 ml sterile water. Cells were sonicated briefly to ensure cells are separated and then a serial dilution was made into sterile water as follows: dilution 1: 10 μl cells + 990 μl water = approx 1 × 105 cells/ml = 1 × 102 cells/μl; dilution 2: 10 μl dilution 1 + 990 μl water = approx 1 × 103 cells/ml = 1 cell/μl.

One hundred microlitre of dilution 2 was plated on YPD plates in triplicate for the viability calculation. One hundred microlitre and 10 μl of undiluted cells were plated in triplicate on SC-leu plates to obtain the fraction of viable cells that are Leu+ before and after induction of re-replication. Plates were incubated at 25°C for 48 hr before colonies were counted. To calculate the percentage of cells in the population that were Leu+, the number of Leu+ colonies per ml was divided by the number of viable cells per ml.

Acknowledgements

We thank members of the Zegerman lab for critical reading of the manuscript. Work in the PZ lab was supported by AICR 10–0908, Wellcome Trust 107056/Z/15/Z, Cancer Research UK C15873/A12700 and Gurdon Institute funding (Cancer Research UK C6946/A14492, Wellcome Trust 092096). Part III undergraduate student DA was supported by the Department of Biochemistry. MS was funded by the BBSRC BB/M011194/1. GC was supported by a Turkish government grant and a Raymond and Beverley Sackler studentship.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Philip Zegerman, Email: paz20@cam.ac.uk.

Tim Formosa, University of Utah School of Medicine, United States.

Kevin Struhl, Harvard Medical School, United States.

Funding Information

This paper was supported by the following grants:

  • Worldwide Cancer Research AICR 10-0908 to Mark C Johnson, Geylani Can, Miguel Monteiro Santos, Diana Alexander, Philip Zegerman.

  • Wellcome Trust 107056/Z/15/Z to Mark C Johnson, Geylani Can, Miguel Monteiro Santos, Diana Alexander, Philip Zegerman.

  • Biotechnology and Biological Sciences Research Council BB/M011194/1 to Miguel Monteiro Santos.

  • Cancer Research UK C15873/A12700 to Mark C Johnson, Geylani Can, Miguel Monteiro Santos, Diana Alexander, Philip Zegerman.

  • Cancer Research UK C6946/A14492 to Mark C Johnson, Philip Zegerman, Miguel Monteiro Santos, Diana Alexander, Geylani Can.

  • Wellcome Trust 092096 to Mark C Johnson, Geylani Can, Miguel Monteiro Santos, Diana Alexander, Philip Zegerman.

Additional information

Competing interests

No competing interests declared.

Author contributions

Formal analysis, Methodology.

Formal analysis, Methodology.

Formal analysis, Methodology.

Formal analysis, Methodology.

Conceptualization, Formal analysis, Investigation, Methodology, Writing - original draft, Project administration.

Additional files

Transparent reporting form

Data availability

Sequencing data has been deposited in GEO under the accession code GSE159122 and GSE163571.

The following datasets were generated:

Johnson MC, Can G, Santos MM, Alexander D, Zegerman P. 2020. Checkpoint inhibition of origin firing prevents inappropriate replication outside of S-phase. NCBI Gene Expression Omnibus. GSE159122

Johnson MC, Can G, Santos MM, Alexander D, Zegerman P. 2020. Checkpoint inhibition of origin firing prevents inappropriate replication In G2/M phase. NCBI Gene Expression Omnibus. GSE163571

The following previously published dataset was used:

Morafraile EC, Hanni C, Allen G, Zeisner T, Clarke C, Johnson MC, Santos MM, Carroll L, Minchell NE, Baxter J, Banks P, Lydall D, Zegerman P. 2019. Genetic screens identify pathways that are important in the absence of checkpoint inhibition of origin firing. Supplemental_SuppTable3.xlsx. DC1

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Decision letter

Editor: Tim Formosa1
Reviewed by: Tim Formosa2, Bruce Stillman3, Karim Labib4

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

Johnson et al. describe roles for the "S-phase checkpoint" outside of S phase, extending the reach of the factors responsible for monitoring DNA integrity to other parts of the cell cycle. This careful analysis of the general roles of this checkpoint in yeast have obvious parallels for understanding checkpoint functions more generally, and the DNA damage response in higher eukaryotes that are less amenable to the types of analysis performed here.

Decision letter after peer review:

Thank you for submitting your article "Checkpoint inhibition of origin firing prevents inappropriate replication outside of S-phase" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Tim Formosa as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Kevin Struhl as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Bruce Stillman (Reviewer #2); Karim Labib (Reviewer #3).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, we are asking editors to accept without delay manuscripts, like yours, that they judge can stand as eLife papers without additional data, even if they feel that they would make the manuscript stronger. Thus the revisions requested below only address clarity and presentation.

Summary:

Johnson et al. use the yeast Saccharomyces cerevisiae system to show that oxidative damage triggers phosphorylation of DNA replication initiation factors Sld3 and Dbf4 by activated Rad53 kinase in cells arrested in either G1 or G2/M. They also show that regulating the activities of these factors outside of S-phase involves the same phosphorylation target sites that are used within S-phase. Finally, they show that the inappropriate replication that occurs in the presence of damage when the checkpoint is inactivated follows the normal timing program, with early origins activated first. These results show that the same checkpoint controls that monitor S phase progression are also used during other phases of the cell cycle. The effects during G1 provide a direct mechanism for inhibiting initiation, in addition to the previously studied parallel block to S phase entry produced by the effects of Rad53 on CDK activation. The effects in G2 limit re-replication, which was previously shown to be regulated by CDK activity as well. This work takes advantage of the high resolution available in the yeast system to concisely establish a broader role for the S-phase checkpoint system than had been previously assumed, and to cleanly demonstrate how multiple factors contribute to maintaining genomic stability in the presence of DNA damage.

Essential revisions:

1) The authors should submit their DNA sequence copy number data to a public DNA sequence database

2) Introduction: The authors should provide further support for the contention that S-phase checkpoint controls are currently assumed to act only within S phase.

3) Subsection “Rad53-dependent phosphorylation of Sld3 and Dbf4 reduces re-replication in G2 phase” and Figure 3B: The change in flow cytometry profiles seems small; have potential changes in cell size (FSC, SSC) and the variation in mitochondrial DNA associated with cell size been accounted for?

4) Introduction: Parallels to the human DNA damage checkpoints should be included here, especially the role of p21-CIP. In general, more discussion of the parallel contributions of phosphorylation by Rad53 to Sld3-Dbf4 regulation and CDK activity would strengthen the manuscript. The recent results from the Remus laboratory on the effects of Rad53 on the MCM complex would also provide additional context for the Sld3-Dbf4 results described here.

5) Figure 1B upper panel, Figure 1—figure supplement 1B upper panel, Figure 2A upper panel: Sld3 appears as a doublet in untreated G1 cells (0 minute sample). Similarly, in Figure 1E upper panel, the bottom band that is supposed to represent unphosphorylated Sld3 only decreases slightly upon DNA damage. The authors should demonstrate that the lower band is specific to Sld3 and explain these features of the data.

6) Figure 2B, D: The level of the dbf4-A mutant protein appears to be higher in cells compared to the Dbf4 WT protein. As Dbf4 is limiting for initiation and the effects of combining dbf4-A with swi4-t in Figure 4 are rather subtle, the authors should consider the possibility that the effects are due to overexpression of dbf4-A protein rather than to the lack of phosphorylation.

7) Subsection “Rad53-dependent phosphorylation of Sld3 and Dbf4 reduces re-replication in G2 phase” and Figure 3D: The statistical significance of the small change reported with 3 replicates suggests that further repeats would be needed to support the conclusion here and in Figure 4F.

8) The use of dbf4 alleles to establish which phosphorylation sites are important should be clarified. It is not always clear whether the 4A allele (mutations in the sites that are essential in S phase) or the 19A allele (mutations in all mapped phosphorylation sites) are being used in specific experiments. This could affect whether or not the conclusion that the same sites are important in S phase and G1/G2 is valid, so this needs to be clarified and the reasons for using different alleles justified.

eLife. 2021 Jan 5;10:e63589. doi: 10.7554/eLife.63589.sa2

Author response

Essential revisions:

1) The authors should submit their DNA sequence copy number data to a public DNA sequence database

The GEO accession number is GSE159122.

2) Introduction: The authors should provide further support for the contention that S-phase checkpoint controls are currently assumed to act only within S phase.

Here we make the general point that checkpoints are considered to be tailored to different cell cycle phases. To support this point, we have included the citation (Shaltiel et al., 2015) which states in the Abstract “despite the shared mechanisms of DNA damage detection throughout the cell cycle, the checkpoint and its reversal are precisely tuned to each cell cycle phase”. We think that our analysis of checkpoint inhibition of origin firing throughout the cell cycle shows that for at least this branch of the “S-phase” checkpoint there is no evidence of tailoring to a single phase.

3) Subsection “Rad53-dependent phosphorylation of Sld3 and Dbf4 reduces re-replication in G2 phase” and Figure 3B: The change in flow cytometry profiles seems small; have potential changes in cell size (FSC, SSC) and the variation in mitochondrial DNA associated with cell size been accounted for?

Cell size has been accounted for in the flow cytometer as FSC and SSC of all samples was the same. We decided that the best way to demonstrate genomic re-replication was to perform DNA sequencing of the experiment described in this comment by the reviewers. By over-expressing a subset of pre-RC mutants that are refractory to inhibition by CDK (Mcm7-NLS, Orc6-A, PGal-Cdc6) we observed re-replication mostly on Chromosome III (see new Figure 3—figure supplement 1C), which is in line with previous work (Green et al., 2006). This limited re-replication also underlines why the changes in DNA content by flow cytometry are modest even in the Mcm7-NLS, Orc6-A, PGal-Cdc6 strain. Importantly we observe greater re-replication of chromosome III when Sld3 and Dbf4 cannot be inhibited by Rad53 (new Figure 3B). From this sequencing we also observe that mitochondrial DNA content is equivalent between the strains (new Figure 3—figure supplement 1B). Together with the other data in Figure 3, we are confident that Rad53 can inhibit Sld3 and Dbf4 in G2 phase to limit the rate of re-replication.

4) Introduction: Parallels to the human DNA damage checkpoints should be included here, especially the role of p21-CIP. In general, more discussion of the parallel contributions of phosphorylation by Rad53 to Sld3-Dbf4 regulation and CDK activity would strengthen the manuscript. The recent results from the Remus laboratory on the effects of Rad53 on the MCM complex would also provide additional context for the Sld3-Dbf4 results described here.

We are very grateful for these comments and we have greatly expanded this section of the Introduction and included the points and references suggested. We then circle back to the parallel pathways of CDK and replication factor inhibition in the Discussion.

5) Figure 1B upper panel, Figure 1—figure supplement 1B upper panel, Figure 2A upper panel: Sld3 appears as a doublet in untreated G1 cells (0 minute sample). Similarly, in Figure 1E upper panel, the bottom band that is supposed to represent unphosphorylated Sld3 only decreases slightly upon DNA damage. The authors should demonstrate that the lower band is specific to Sld3 and explain these features of the data.

Sld3 has been shown to be phosphorylated by DDK in G1 arrested cells (Mattarocci et al., 2014) and we have included a sentence (subsection “Sld3 and Dbf4 are phosphorylated by Rad53 outside of S-phase”) to state that we consider it likely that the minor phospho-Sld3 band in the G1 arrested samples is due to DDK phosphorylation.

We are not sure why there is a persistent unphosphorylated fraction of Sld3 in nocodazole arrested cells (e.g. Figure 1E), which does not change after DNA damage. We include a new figure (Figure 1—figure supplement 1C) without tagged Sld3, with and without inhibition of CDK, which clearly identifies these bands as being specific to CDK phosphorylated and unphosphorylated Sld3.

6) Figure 2B, D: The level of the dbf4-A mutant protein appears to be higher in cells compared to the Dbf4 WT protein. As Dbf4 is limiting for initiation and the effects of combining dbf4-A with swi4-t in Figure 4 are rather subtle, the authors should consider the possibility that the effects are due to overexpression of dbf4-A protein rather than to the lack of phosphorylation.

In this study we have used two different mutants of Dbf4; 19A, which lacks all the Rad53 phospho-sites and 4A, which lacks the key Rad53 sites that inhibit Dbf4’s role in origin firing. (Note that the 4A sites are a subset of the 19A sites). Also, in light of point 8 raised by the reviewers, we realise that this is confusing and we did not explicitly state which allele was used in Figure 2, for which we apologise. For all phenotypic assays (including Figure 4) we always use Dbf4-4A, because this allele is fully functional for origin firing and the protein is expressed at wild type levels. Therefore, the expression level of Dbf4-4A is not an issue for Figure 4.

For Figure 2B and D (and only in these figures) we used Dbf4-19A, because it lacks all available Rad53 phospho-sites and therefore gives a more robust loss of phosphorylation by western blot. As pointed out by the reviewers, this 19A allele is more expressed than the wild type protein (and it’s a hypomorph, which may be why cells adapt by over-expressing it). We have rectified these issues as follows:

1) At the first mention of Dbf4 phospho-site mutants (subsection “Sld3 and Dbf4 are phosphorylated by Rad53 outside of S-phase”), we explain the difference between these 2 alleles. From then on, we refer to Dbf4-4A as Dbf4-A and Dbf4-19A as it is.

2) We clearly state that Dbf4-19A was used for Figure 2B and D in the figure, legend and the text.

3) We have included a new figure which shows that Dbf4-4A is expressed at very similar levels to wild type and also shows defects in Rad53 phosphorylation in G1 phase (new Figure 2—figure supplement 1).

7) Subsection “Rad53-dependent phosphorylation of Sld3 and Dbf4 reduces re-replication in G2 phase” and Figure 3D: The statistical significance of the small change reported with 3 replicates suggests that further repeats would be needed to support the conclusion here and in Figure 4F.

The statistical significance of these differences is less than 0.05, so the data supports our conclusions. More repeats would indeed increase the significance further.

8) The use of dbf4 alleles to establish which phosphorylation sites are important should be clarified. It is not always clear whether the 4A allele (mutations in the sites that are essential in S phase) or the 19A allele (mutations in all mapped phosphorylation sites) are being used in specific experiments. This could affect whether or not the conclusion that the same sites are important in S phase and G1/G2 is valid, so this needs to be clarified and the reasons for using different alleles justified.

See point 6.

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Data Citations

    1. Johnson MC, Can G, Santos MM, Alexander D, Zegerman P. 2020. Checkpoint inhibition of origin firing prevents inappropriate replication outside of S-phase. NCBI Gene Expression Omnibus. GSE159122 [DOI] [PMC free article] [PubMed]
    2. Johnson MC, Can G, Santos MM, Alexander D, Zegerman P. 2020. Checkpoint inhibition of origin firing prevents inappropriate replication In G2/M phase. NCBI Gene Expression Omnibus. GSE163571 [DOI] [PMC free article] [PubMed]
    3. Morafraile EC, Hanni C, Allen G, Zeisner T, Clarke C, Johnson MC, Santos MM, Carroll L, Minchell NE, Baxter J, Banks P, Lydall D, Zegerman P. 2019. Genetic screens identify pathways that are important in the absence of checkpoint inhibition of origin firing. Supplemental_SuppTable3.xlsx. DC1 [DOI] [PMC free article] [PubMed]

    Supplementary Materials

    Transparent reporting form
    elife-63589-transrepform.docx (66.5KB, docx)

    Data Availability Statement

    Sequencing data has been deposited in GEO under the accession code GSE159122 and GSE163571.

    The following datasets were generated:

    Johnson MC, Can G, Santos MM, Alexander D, Zegerman P. 2020. Checkpoint inhibition of origin firing prevents inappropriate replication outside of S-phase. NCBI Gene Expression Omnibus. GSE159122

    Johnson MC, Can G, Santos MM, Alexander D, Zegerman P. 2020. Checkpoint inhibition of origin firing prevents inappropriate replication In G2/M phase. NCBI Gene Expression Omnibus. GSE163571

    The following previously published dataset was used:

    Morafraile EC, Hanni C, Allen G, Zeisner T, Clarke C, Johnson MC, Santos MM, Carroll L, Minchell NE, Baxter J, Banks P, Lydall D, Zegerman P. 2019. Genetic screens identify pathways that are important in the absence of checkpoint inhibition of origin firing. Supplemental_SuppTable3.xlsx. DC1

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