Key histone chaperones have distinct roles in replisome progression and genomic stability

Replication-coupled (RC) nucleosome assembly is an essential process in eukaryotic cells in order to maintain chromatin structure during DNA replication. The deposition of newly synthesized H3/H4 histones during DNA replication is facilitated by specialized histone chaperones. Although the contribution of these histone chaperones to genomic stability has been thoroughly investigated, their effect on replisome progression is much less understood. By exploiting a time-lapse microscopy system for monitoring DNA replication in individual live cells, we examined how mutations in key histone chaperones including CAC1, RTT106, RTT109 and ASF1, affect replication fork progression. Our experiments revealed that mutations in CAC1 or RTT106 that directly deposit histones on the DNA, slowdown replication fork progression. In contrast, analysis of cells mutated in the intermediary ASF1 or RTT109 histone chaperones revealed that replisome progression is not affected. We found that mutations in histone chaperones including ASF1 and RTT109 lead to extended G2/M duration, elevated number of RPA foci and in some cases, increased spontaneous mutation rate. Our research suggests that histone chaperones have distinct roles in enabling high replisome progression and maintaining genome stability during cell cycle progression. Author Summary Histone chaperones (HC) play key roles in maintaining the chromatin structure during DNA replication in eukaryotic cells. Despite extensive studies on HCs, little is known regarding their importance for replication fork progression during S-phase. Here, we utilized a live-cell imaging approach to measure the progression rates of single replication forks in individual yeast cells mutated in key histone chaperones. Using this approach, we show that mutations in CAC1 or RTT106 HCs that directly deposit histones on the DNA lead to slowdown of replication fork progression. In contrast, mutations in ASF1 or RTT109 HCs that transfers H3/H4 to CAC1 or RTT106, do not affect replisome progression but lead to post replication defects. Our results reveal distinct functions of HCs in replication fork progression and maintaining genome stability.

G2/M elongation may stem from the activation of two mitotic checkpoints due to genomic instability accumulated during DNA replication [33]. These two checkpoints are the DNA damage checkpoint and the spindle assembly checkpoint [34,35]. Previously, mutation in CAC1 was shown to be implicated in the activation of the spindle checkpoint [36]. Specifically, it was shown that the deletion of MAD2, the master checkpoint protein of spindle assembly, suppressed the mitotic delay measured in cac1-and hir1-double deletion strain [36]. To examine whether the G2/M elongation observed in cac1-deletion cells (Fig. 2) was due to spindle checkpoint arrest, we deleted MAD2 in the background of cac1-deletion. We then measured replication times and G2/M duration in these mutant cells using the live-cell imaging system described above. In accordance with the previous study [36], we found that the elongation of G2/M duration due to CAC1 deletion is suppressed in the double mutant cells and is similar to the G2/M duration values measured for WT cells (Fig. 3).
To further support the importance of spindle checkpoint activation in cac1-deleted cells, we examined the growth of cac1-and mad2-double deleted cells using a spot assay. In agreement with the microscopy experiments, we found that the addition of MAD2 deletion suppressed the growth defects observed in cac1-deleted cells (Fig. S1). These results, further supports the role of CAC1 in ensuring mitotic spindle integrity prior to cell mitosis.

Separation of function of the WHD and PIP regions in Cac1
The elongation of replication times and G2/M duration of cac1-deleted cells can stem from the lack of Caf1 complex formation or from deficiencies in Cac1 specific interactions. Previously, Cac1 was shown to contain a WHD domain and a PIP region facilitating its direct interaction with the DNA and PCNA, respectively [23,25]. To examine the importance of these Cac1 regions for replisome progression and G2/M duration, we have generated strains for replication measurements containing K564E/K568E and F233A/F234G mutations in Cac1 WHD domain and the PIP region, respectively. These mutations in the WHD and PIP domains of Cac1 were previously shown to abolish binding to DNA and PCNA, respectively [23,25]. Next, we measured replication time and G2/M duration in individual cells containing these mutations as described above. We found that mutating Cac1 WHD domain leads to increased replication times with no effect on the G2/M duration, relative to WT cells (Fig. 4A). In contrast, we found that mutating the PIP region of the protein does not affect replication times but leads to a significant elongation of G2/M duration (Fig.   4B). These results demonstrate the separation of function between the WHD and PIP regions of Cac1, with DNA binding contributing to proper replisome progression and PCNA binding preventing replication stress. Next, we examined the level of nucleosome deposition on newly synthesized DNA in the WHD and PIP Cac1 mutant strains at the vicinity of ARS305 and ARS605 using chromatin immunoprecipitation (ChIP) following release from HU arrest. In accordance with the effect of these mutations on replisome progression rate and G2/M duration, we found a significant decrease in the level of nucleosome deposition during DNA replication in these mutant strains relative to the WT ( Fig. 4C-D and Fig. S2). To further examine the effect of HU replication stress on the two Cac1 mutants, we measured their replication times in presence of 20 mM HU. We found that both mutants exhibit more than 9 minutes longer replication times compared to WT (Fig. S3). These results highlight the importance of both PIP and WHD of Cac1 for nucleosome deposition and replication fork progression at early S-phase under DNA stress conditions.

Analysis of mutation in RTT106, RTT109 or ASF1 histone chaperones
To examine how mutations in additional H3/H4 histone chaperones affect replisome progression and G2/M duration, we have generated deletions in RTT106, RTT109 or ASF1 on the background of the lacO and tetO containing strain as described above. In addition, to examine how double deletion in CAC1 and RTT106 affect these properties we have generated this strain on the same background. Rtt106 was previously shown to interact with H3/H4 and the DNA to enable (H3/H4) 2 tetramer deposition on newly synthesized DNA [37]. Cac1 and Rtt106 were shown to physically interact suggesting that these histone chaperones cooperate to facilitate high level of nucleosome deposition on newly synthesized DNA [38]. Asf1 and Rtt109 are additional key histone chaperone and histone acetylase, respectively, acting prior to Cac1 and Rtt106 to facilitate H3K56 acetylation of newly synthesized histones [12,19]. Analysis of rtt106-deletion and the cac1-rtt106-double deletion cells using our assay showed significantly increased replication times, relative to WT cells.
In contrast, rtt109-or asf1-deletion did not lead to a significant change in replication times (Fig.   5A). Analysis of the G2/M duration in cac1-rtt106-double deletion, rtt109-or asf1-deleted cells was significantly extended while was not affected in rtt106-deleted cells relative to WT cells (Fig.   5B). Elongation of G2/M duration, observed in cac1-rtt106-double deletion, rtt109-or asf1-deleted cells, is in agreement with several studies showing checkpoint activation and replication stress in these strains [14,[19][20][21].

Increased G2/M duration in histone chaperone mutant cells is associated with ssDNA accumulation
Replication stress often lead to the accumulation of single-stranded DNA near active replication and repair sites [39]. This accumulation can be visualized as foci of the ssDNA-binding protein Replication Protein A (RPA) [40][41][42]. To examine whether long G2/M duration in histone chaperone mutant cells is associated with increased level of ssDNA accumulation, we monitored the levels of RPA foci in these mutant cells. For monitoring RPA foci in live cells, we used Rfa1, expressed under its native promoter, tagged with GFP at its N-terminus [43] and used fluorescent time-lapse microscopy to monitor foci in cells that started budding. To verify the nuclear localization of the RPA foci, we monitored their co-localization with a nuclear signal generated by simultaneous expression of NLS-tetR-tdTomato. Indeed, we found that RPA foci are identified within the cell nucleus following budding and entering S-phase (Fig. S4). Previously, the lack of PCNA ubiquitylation by Rad18 ubiquitin ligase was shown to significantly increase the level of RPA foci in mammalian cells [42]. Thus, to further validate our system, we generated rad18deletion strain and examined the level of RPA foci in mutant cells relative to WT. We found that the percentage of cells with RPA foci containing the rad18-deletion is significantly higher than WT cells (Fig. S5).
Next, we generated the different histone chaperone mutants on the background of GFP-Rfa1 strain and monitored the accumulation of RPA foci in mutant and WT cells (please see Fig. 6A for representative WT and asf1-deleted cells). We found an increase in the percentage of cells containing RPA foci in the histone chaperone mutant cells relative to WT cells indicating that these mutations lead to the accumulation of ssDNA [39,40] (Fig. 6B). Increase in RPA foci was most prominent in asf1-or rtt109-deleted strains, in which over 40% of the cell population contained at least one clear RPA focus. We found that the increased level of RPA foci in histone chaperone mutant strains is strongly correlated with increased G2/M duration, validating that the delayed G2/M phenotype seen in these strains is related to accumulation of DNA damage during S phase (Fig. S6).
Taken together, our results suggest that replication that is uncoupled to nucleosome deposition can lead to replication stress, accumulation of ssDNA, and significant delays in cell cycle progression ( Fig. 6 and Fig. S6).

Spontaneous mutation rate analysis of histone chaperone mutant strains
Replication fork stalling and accumulation of ssDNA during replication can lead to double strand breaks that can be detrimental to cell survival [44,45]. To avoid double strand breaks, cells employ a variety of mechanisms including the recruitment of translesion DNA polymerases (TLS) by the RAD6/RAD18 pathway [46,47]. This pathway, initiated by mono-ubiquitylation of PCNA on K164, can minimize fork stalling but may lead to increased mutation rates due to the inaccuracy of the TLS polymerases [46,47]. To examine whether mutation rates are elevated in the different histone chaperone mutant strains, we have utilized the CAN1 assay. We have previously utilized this assay to examine a variety of PCNA mutants leading to RAD6/RAD18 pathway activation [48]. In accordance with previous analysis, we found that asf1-or rtt109-single deletions lead to a mild increase in mutation rate of up to two-fold relative to the WT strain [49]. Single deletions of CAC1 or RTT106 lead to similarly mild increase. In contrast, we found that the cac1-rtt106-double deletion leads to a dramatic increase of ~7 fold in mutation rate, (Fig. 7 and Table 1). To examine whether the increased mutation rate in the cac1-rtt106-double deletion strain is due to RAD6/RAD18 pathway activation, we generated a cac1-rtt106-rad18-triple deletion strain and examined its mutation rate. Consistent with RAD6/RAD18 pathway activation, we observed that the triple deletion strain exhibits a dramatic reduction in mutation rate relative to the cac1-rtt106-double deletion strain (Fig. 7). In addition, we found that depletion of DNA polymerase ζ, a key TLS polymerase, in this strain leads to higher sensitivity to MMS, highlighting its possible activation upon DNA damage (Fig. S7). Taken together, our results with the cac1-rtt106-double deletion strain show that mutations in histone chaperones can lead to RAD6/RAD18 pathway activation and increased spontaneous mutation rates. However, results with the other histone chaperone mutant strains, exhibiting long G2/M duration and a mild increase in mutation rate (e.g. rtt109 or asf1 mutant strains), suggest that other pathways can be activated upon ssDNA accumulation during DNA replication (see Discussion section).

Discussion
DNA RC nucleosome assembly that connects DNA synthesis with nucleosome deposition and modification has been extensively studied in the past two decades [9,11,50]. These studies allowed the identification and characterization of histone chaperones that are important for nucleosome deposition on newly synthesized DNA. However, much less is known regarding how mutations in histone chaperones affect replisome progression during DNA replication. Here, we used our recently described direct approach to monitor replisome progression in individual yeast cells [28,29] that are mutated in different histone chaperone genes, including Caf1 subunits, RTT106, ASF1 and RTT109.
We found that mutation in Cac1 or Rtt106 that directly deposit H3/H4 histones on DNA, significantly slowdown replication fork progression ( Fig. 2-5). These results suggest that the direct interaction of these chaperones with the replication fork is essential for enabling high fork progression rate during S-phase. Interestingly, we found that mutations in histone chaperones which transfer H3/H4 on Cac1 or Rtt106 prior to their final assembly, including ASF1 or RTT109 histone acetyl transferase, do not affect replisome progression but significantly extend G2/M duration ( Fig.   5) highlighting the distinct and independent roles of histone chaperones in genome replication and stability. The normal replisome progression in asf1-deleted cells (Fig. 5) is in good agreement with previous studies showing that yeast budding and progression through S-phase is not significantly altered in this mutant strain [20,21].
Our system also allows the detection of elongations in G2/M duration by monitoring both fluorescence and morphological changes in individual cells. Our findings that single deletion of CAC1, RTT109 or ASF1 or double deletion of CAC1 and RTT106 (Fig. 5) leads to a significant increase in G2/M duration highlight the important role of these chaperones in preventing replication stress [39]. Our analysis of G2/M duration is probably an underestimation of the true G2/M length in these mutant cells since high percentage of the analyzed cells did not undergo mitosis during the experiment duration (Table S3). Our further examination of these strains revealed a dramatic increase in RPA foci formation following DNA replication (Fig. 6). We found that in the case of double deletion of CAC1 and RTT106 the elongated G2/M duration and RPA foci formation is also correlated with a significant increase in spontaneous mutation rate (Fig. 7). Previously, several studies have extensively examined different aspects of genome stability in histone chaperone mutant strains. An extensive study examining several single and double deletions of histone chaperones including RTT109, ASF1, CAC1 and RTT106 in yeast revealed that most mutations lead to increased recombination frequency, checkpoint activation, loss of replisome stability and sensitivity to DNA damaging agents [14]. In addition, RTT106 was shown to genetically interact with DDC1-MEC3-RAD17 that forms an alternative sliding clamp termed the 9-1-1 complex which is involved in checkpoint activation [51]. Furthermore, ASF1 was shown to directly interact with the downstream RAD53 effector kinase [52,53] suggesting that these checkpoint proteins are intimately connected to DNA RC nucleosome deposition. Our findings that histone chaperone mutants lead to high level of RPA foci formation highlight their importance for preventing the accumulation of ssDNA. It complements studies showing the interaction between histone chaperones and S-phase checkpoint activation where ssDNA accumulation sensed by RPA (Fig. 6) is transmitted to several proteins including the 9-1-1 complex [51], leading to Mec1 sensor kinase activation followed by Rad53 effector kinase activation [52,53]. The correlation between high level of RPA foci formation and G2/M duration (Fig. S6) highlights the detrimental effect of ssDNA accumulation for cell cycle progression. In contrast to WT cells in which RPA foci can be corrected during S-phase, we observed that for histone chaperone mutants, RPA foci can be maintained through the G2 phase following DNA replication (Fig. 6A).
Analysis of the different subunits of the Caf1 complex indicated that deletion of CAC1 exhibits the strongest effect on both replisome progression and G2/M duration (Fig. 2). Previously, Cac1 has been implicated in the MAD2-dependent mitotic spindle checkpoint [36] and our results further verify this study (Fig. 3). Further analysis of mutations in Cac1 WHD or PIP regions showed that while the WHD domain is important for replisome progression, the PIP region is important for preventing replication stress (Fig. 4). This separation of function highlights the importance of direct Cac1-DNA interaction, mediated by the WHD domain, for fast replisome progression. In contrast, elongation of G2/M duration in Cac1 PIP mutant highlights the importance of coordinating Cac1 activity with replisome progression for avoiding replication stress. Our further analysis of the WHD and PIP Cac1 mutants showed a significant reduction of nucleosome deposition of both mutants relative to WT strain as well as slowdown of replisome progression in the presence of HU (Fig. 4 and Fig. S3). Previously, several studies have examined the importance of Cac1 WHD and PIP regions for their biochemical and cellular functions including their importance for DNA binding following association with H3-H4 histones [27], nucleosome assembly in vitro [23] and sensitivity to DNA damaging agents [25]. Our results examining the function of these regions in live cells are in agreement with these studies and further enhance our understanding of the contribution of each interaction for efficient replisome progression and short G2/M duration.
Previously, mutations in histone chaperones were shown to increase recombination level and gross chromosomal rearrangement highlighting their roles in maintaining genome stability [14,16]. Asf1, Caf1 and Rtt109 were shown to be important for deactivation of damage checkpoint after double strand break repair [55,56]. However, much less is known regarding their contribution to DNA damage response pathways that include the activation of TLS polymerases for damage bypass. Our findings that mutations in histone chaperones lead to moderate increase in spontaneous mutation rate (Fig. 7) suggest that other pathways are dominant, including homologues recombination, to overcome replication stress.
Our findings are in good correlation with a previous study examining the level of spontaneous mutation rate in several deletion strains including the RTT109 or ASF1 single deletions [49]. The significant increase in spontaneous mutation rate in cells deleted in CAC1 and RTT106 (Fig. 7) may suggest that these chaperones link PCNA both to nucleosome deposition and activation of TLS polymerases through PCNA ubiquitylation by the RAD6/RAD18 pathway. Notably, these results are in agreement with the involvement of CAC1 in the RAD6/18 post-replication repair pathway [32].
Overall, our experiments reveal that histone chaperones exhibit distinct functions in replisome progression and maintaining genome stability. Our data showing the effects of cac1-or rtt106-deletion on replisome progression highlight the possible importance of direct interaction of these chaperones with H3/H4 and the DNA in facilitating fast replisome progression. In contrast, histone chaperones which are involved indirectly with the transfer of histones on DNA, including Asf1 and Rtt109 do not significantly affect replisome progression but are essential for preventing replication stress. Our system can be further utilized to investigate how additional histone chaperones, such as the FACT complex, Nap1, Vps75 [5,57], or combinations of mutants affect replication fork progression and G2/M duration.

Strain generation
Strains for replication time measurement were generated on the background of a W303 MATa Saccharomyces cerevisiae strain, expressing GFP-LacI and tetR-tdTomato fusion proteins in the nucleus. lacOx256 and tetOx224 arrays are located at chrIV:332960 and chrIV:352560 respectively, near ARS413 and with a mid-array distance of approximately 30.6 kb [28]. CAC1, CAC2, CAC3, RTT106, ASF1, RTT109 and MAD2 genes were replaced by natMX or hphMX antibiotic cassettes using the Lithium Acetate (LiAc) transformation method [58]. The cac1 WHD, cac1 PIP, and rtt106 KK245,246AA mutants were generated on the background of the cac1-or rtt106-deleted cells respectively, with a markerless CRISPR/Cas9 approach, by targeting either the natMX or hphMX with specific gRNAs and the respective mutant genes as DNA donors [59]. All replacements were validated with Polymerase Chain Reaction (PCR) and Sanger sequencing.

Microscopy and data analysis
All microscope experiments for the determination of replication time were conducted as previously described [28] with slight modifications. Briefly, yeast cultures were grown overnight to OD 600nm = 0.1-0.2 in synthetic complete (SC) medium containing 4% glucose at 30˚C. The cultures were then synchronized at G1 using 10 μg/ml α-factor (GenScript) for 2-3 hours. For hydroxyurea (HU) experiments, yeast cultures were incubated with 20 mM HU one hour before and during the imaging.
Cells were then immobilized on microscopy slide chambers (Ibidi) coated with 2 mg/ml concanavalin A (Sigma-Aldrich) and washed thoroughly from α-factor with warm SC medium containing 4% glucose prior to microscopy measurements. Live cell imaging of the cells was performed on a Marianas spinning-disk confocal microscope (3i) using an Evolve EM-CCD camera (Photometrics) with 1 min intervals for 2-8 hours at 28˚C, using a x63 oil objective (NA = 1.4) in 3D (12 z-sections at 0.7μm apart). GFP-LacI and TetR-tdTomato were excited with 488nm and 561nm lasers, respectively. For data analysis, time-lapse measurements were collected with SlideBook (3i) and analyzed in Matlab using a custom-made software ('DotQuant') which identifies, tracks and quantifies the fluorescent dots in each cell [28]. Statistical analysis of replication rate results was performed using Monte Carlo resampling with 1,000,000 iterations.
Mitosis events were identified manually as the timepoint when the dots split and move to the mother and daughter cell.
Exponentially growing cells were synchronized with 5 μg/ml α-factor for 3 hr at 25℃ and then released into fresh media containing 0.2 M HU for 45 min at 25℃. Samples were collected to perform ChIP assay using H3K56Ac and H3 (as a normalization control) antibodies. The ChIP DNA was analyzed by quantitative PCR using primers (ARS607/ARS607+14kb and ARS305/ARS305+12kb) amplifying replication origins as well as fragments downstream of replication origins. The mean and SD of three independent biological replicates were shown. The percentage of ChIP DNA relative to the total input DNA was calculated (Fig. 4), and the ratio of H3K56Ac ChIP signal to the total H3ChIP signal was also calculated (Fig. S2).

RPA foci analysis
Strains for RFA1 foci analysis were generated on the background of a BY4741 Saccharomyces cerevisiae strain which carries N-terminus GFP-tagged RFA1 expressed under control of its native promoter [43], as described above. To locate the cell nuclei, NLS-tetR-tdTomato was transformed into the ADE1 locus. For imaging, yeast cultures were grown overnight at 30˚C to OD 600nm = 0.2-0.6 in SC medium containing 4% glucose. Cells were fixed on microscopy slide chambers (Ibidi) coated with 2mg/ml concanavalin A (Sigma-Aldrich) and imaged using confocal microscopy as described above for 1-3h with 1 min intervals at 28˚C. GFP-Rfa1 and NLS-tetR-tdTomato were excited with 488 nm and 561 nm lasers, respectively. At least 60 cells from three independent experiments were analysed for each strain and scored for the presence of one or more RPA foci during S-phase. The percentage of cells with RPA foci was estimated by dividing the number of cells exhibiting one or more RPA foci in the nucleus by the total amount of cells that were taken into account in each experiment. Results were compared with one-way ANOVA and post-hoc analysis (Dunnett's test), after transforming the data with arcsin(sqrt(x)).

Can1 spontaneous mutation rate analysis
The determination of the mutation rate of the can1 gene was performed as previously described [60] based on the Lea and Coulson method [61] with some modifications. Strains for can1 mutation rate measurement were generated on the background of a BY4741 Saccharomyces cerevisiae strain as described above. Yeast cultures were grown overnight at 30˚C in YPD medium to OD 600nm ≈ 0.8-1.
Cells were then diluted 10 5 fold, plated as single cells on SC plates and grown for 4 days at 30˚C.
At least 16 single colonies were randomly chosen from each strain, resuspended in 1 ml of sterile ddH 2 O and diluted to OD 600nm ≈ 1.0. Next, 10 5 fold dilutions of each resuspended colony solution were plated on SC plates to define the number of viable cells and the rest of the solutions were plated on SC-Arg + 60mg/ml canavanine to obtain the number of canavanine-resistant cells. The median number of viable cells was estimated from the different colonies (replicates) and the mutation rates, the 95% Confidence Intervals (CI) and the p values were calculated by utilising the rSalvador package (v1.7) running on the R software (v3.4.3) [62].

Rev3 inducible degradation and spot assays
Strains for spot assays were generated on the background of a BY4741 Saccharomyces cerevisiae strain.
An auxin-inducible degron (AID) was inserted into the C'-terminus of rev3 as previously described [28]. Briefly, a PCR fragment containing the AID sequence, the Oryza sativa TIR1 gene and the hphMX resistance cassette as selection marker was generated with homologous ends to the 5' of the REV3 yeast gene. The presence of AID results in the degradation of the fused protein after incubation of yeast cells with 1-Napthaleneacetic acid (NAA) [63,64]. The viability of rev3-AID histone-chaperone mutants was tested with spot assays on agarose plates. Briefly, overnight exponentially-growing yeast cultures were diluted to OD 600nm = 0.3 and 4μl of 10-fold serial dilutions were spotted on YPD plates containing 200 μg/ml HYG, HYG+0.5mM NAA, HYG+0.005% MMS, or HYG+0.5mM NAA+0.005% MMS. Cells were incubated at 30˚C for 32 hours and representative pictures were captured.

Figure 1: Measuring replication rates in histone chaperone mutant yeast cells. (A)
Schematic illustration of the design for the real-time analysis of replication kinetics in live yeast cells containing mutations in histone chaperone genes. Strains are genetically engineered to contain lacO and tetO arrays located 20 Kb apart, downstream from an origin of replication. Binding of GFP-lacI and tetR-tdTomato lead to green and red fluorescent foci, respectively. During DNA replication, array duplication leads to recruitment of additional lacI-GFP and tetR-tdTomato proteins leading to an increase in fluorescence intensity. The newly-synthesized DNA is packed around new (orange) and pre-existing (blue) nucleosomes. Histone chaperones (HC, purple) deposit newly-synthesized histones as (H3/H4) 2 tetramers behind the replisome. The replication time of each locus is measured using time-lapse confocal microscopy. For simplicity, origin firing is shown only to the array direction. (B) Schematic display of the doubling of fluorescent intensity of the GFP-lacI and tetR-tdTomato foci in a single cell due to lacO and tetO array duplication during DNA replication. The time between replication of the arrays (replication time) is calculated using the mid-rise points of the GFP and tdTomato fluorescence intensities. The G2/M duration is estimated by the time delay between the mid-rise point of the tdTomato fluorescence intensity and the anaphase (C) Image of single cells from the yeast strains used in this study. Top -image following replication before anaphase, Bottom -after anaphase. Images scale: 2μm  Table  S1 and for (E) Table S2.   Table S1. (C-D). ChIP analysis of H3K56Ac deposition for WT and cac1 mutant strains. WT and cac1 mutant cells were arrested at G1 phase using α-factor at 25°C and then released into fresh medium containing 0.2 M HU. Equal numbers of cells were collected just prior to (G1, 0 min) and at 45 minutes following release into fresh medium containing HU. ChIP assays were performed using antibodies against H3K56Ac and H3 (as a normalization control). The ChIP DNAs were analyzed using four different primer pairs amplifying (C) ARS305, ARS305+12kb and (D) ARS607, and ARS607+14kb, using quantitative PCR. At least three independent biological repeats were performed, with similar trends in all cases. The mean and SDs of three independent biological replicates are shown. P values derived from two-way analysis of variance (ANOVA) (***P value ≤ 0.001). ChIP over input is shown here, results were similar after normalizing to total H3 ChIP for the same trend (Fig. S2), indicating that loss of H3K56Ac signal is not as a result of decreased total H3.  In the case of the WT cells, the RPA focus disappears in late S-phase. In the case of asf1-deleted cell, mitosis does not take place even after 120 min from initial budding and the RPA foci persists. (B) WT or histone chaperone mutant cells expressing N-terminal GFP-Rfa1 fusion were analyzed using time-lapse fluorescent microscopy. At least 60 cells from three independent experiments were analyzed for each strain and scored for the presence of one or more RPA foci during S-phase. Images scale: 2μm Figure 7: Increased spontaneous mutation rate in histone chaperone mutants is most significant in the cac1-rtt106-double deletion strain. Spontaneous mutations in this strain are dependent upon RAD6/RAD18 pathway since spontaneous mutation rate in the triple cac1-rtt106-rad18-deletion strain is dramatically reduced relative to the cac1-rtt106-double deletion strain. The data are rates of Can r mutation values within the 95% confidence limits of at least 16 independent analyses. The values of the spontaneous mutation rate of the different strains are presented in Table 1.  Figure S1: Growth defects of cac1-deletion strain is suppressed by mad2-deletion.
Growth was analyzed by a spot assay on YPD plates using a 10-fold serial dilution of yeast cultures spotted on the plate. Growth was analyzed following 24h (left) or 48h (right) of incubation.