Analyses of POL30 (PCNA) reveal positional effects in transient repression or bi-modal active/silent state at the sub-telomeres of S. cerevisiae
Epigenetics & Chromatin volume 16, Article number: 40 (2023)
Classical studies on position effect variegation in Drosophila have demonstrated the existence of bi-modal Active/Silent state of the genes juxtaposed to heterochromatin. Later studies with irreversible methods for the detection of gene repression have revealed a similar phenomenon at the telomeres of Saccharomyces cerevisiae and other species. In this study, we used dual reporter constructs and a combination of reversible and non-reversible methods to present evidence for the different roles of PCNA and histone chaperones in the stability and the propagation of repressed states at the sub-telomeres of S. cerevisiae.
We show position dependent transient repression or bi-modal expression of reporter genes at the VIIL sub-telomere. We also show that mutations in the replicative clamp POL30 (PCNA) or the deletion of the histone chaperone CAF1 or the RRM3 helicase lead to transient de-repression, while the deletion of the histone chaperone ASF1 causes a shift from transient de-repression to a bi-modal state of repression. We analyze the physical interaction of CAF1 and RRM3 with PCNA and discuss the implications of these findings for our understanding of the stability and transmission of the epigenetic state of the genes.
There are distinct modes of gene silencing, bi-modal and transient, at the sub-telomeres of S. cerevisiae. We characterise the roles of CAF1, RRM3 and ASF1 in these modes of gene repression. We suggest that the interpretations of past and future studies should consider the existence of the dissimilar states of gene silencing.
Positional effect variegation is caused by the proximity of a gene to a heterochromatin boundary and is characterised as a stochastic Active/Silent mode of expression. In Drosophila melanogaster, this effect is readily observed by the variegated expression of the whiteV gene . Seemingly similar stochastic Active/Silent states of the expression of reporter genes has been characterised at the telomeres of S. cerevisiae and other model organisms .
The Proliferating Cell Nuclear Antigen (PCNA, POL30) forms a homo-trimeric sliding clamp which acts at the core of the eukaryotic replisome . PCNA is highly conserved and interacts with numerous proteins, including the leading and lagging strand DNA polymerases, DNA repair polymerases, DNA ligase, and the Fen1p flap-endonuclease. The function of PCNA in DNA replication and repair had been the focus of outstanding reviews [2,3,4] and will not be discussed here. Both human and yeast PCNA directly interact with Chromatin Assembly Factor-1 (CAF-1), a histone chaperone required for replication-coupled chromatin assembly in vitro [7, 8]. In S. cerevisiae, mutations in POL30 lead to loss of heterochromatin-mediated gene silencing [9,10,11,12], thus revealing a direct role in the maintenance and transmission of chromatin structure. Three POL30 mutations, pol30-6, pol30-8 and pol30-79 [9, 13], were pivotal in providing details about this role of PCNA. These three alleles have double alanine substitutions at residues Asp 41 and Asp 42 in pol30-6, residues Arg 61 and Asp 63 in pol30-8, and residues Leu 126 and Ile 128 in pol30-79. On the three-dimensional trimeric structure of PCNA, the pol30-6 mutation is located at the central loop, pol30-8 is located at the tip of a domain bulge, and pol30-79 is located at the interdomain connecting loop .
At the VIIL telomere, the pol30-6, pol30-8 and pol30-79 alleles lead to a significant loss of gene silencing as detected by the insertion of URA3 and the measurement of the sensitivity of the cells to 5-Fluoroorotic acid (5-FOA) [9, 10, 14]. At the HMR and HML mating type loci, these mutations cause a minor loss of silencing that can be detected by the highly sensitive CRASH (Cre-Reported Altered States of Heterochromatin) assay [12, 15]. In both assays, the deletion of CAC1 (it encodes the largest subunit of CAF-1) synergistically reduces gene silencing in the pol30-6 and pol30-79 mutants, but not in pol30-8 [9, 10, 12]. Conversely, in pol30-8 cells the deletion of the histone chaperones ASF1 and HIR1 further reduce gene silencing at the telomeres while having little effect in pol30-6 and pol30-79 . ASF1 acts at the advancing replication forks, but such a role for HIR1 has not been established [16, 17]. Interestingly, in pol30-6, pol30-8 and pol30-79 strains, passage of the replication fork through HMR is not required for the establishment of silencing . These observations prompt questions on the precise mechanisms by which PCNA affects chromatin assembly and/or maintenance.
Loss of gene silencing could be attributed to a slow recovery and reassembly of heterochromatin in the wake of the replication fork or by a full epigenetic conversion of the silenced to the active state. The highly sensitive 5-FOA resistance and CRASH assays do not distinguish which one of the two underlying events leads to the loss of silencing. In this study we revisited the silencing effects of the three pol30 mutants with a routine 5-FOAR assay and a novel drug-free low-sensitivity assay [18, 19]. Our results reveal positional differences in the effects of the pol30 mutations that lead to transient de-repression or complete epigenetic conversions at the VIIL sub-telomere of S. cerevisiae.
pol30 mutations lead to transient de-repression of URA3 at the VIIL sub-telomere
The irreversible 5-FOA resistance and CRASH assays [12, 20] do not distinguish between events of transient destabilization of gene silencing or a complete Silent→Active (S→A) conversion of a gene. Recently, we produced two dual adh4-URA3-HTB1→yEGFP-tel and adh4-URA3-yEGFP←HTB1-tel reporters for the analyses of gene silencing at the VIIL telomere [18, 19]. These reporters contain the URA3 gene followed by fusion HTB1-yEGFP reporter driven by the Histone H2B promoter (HTB1) in the orientation towards or away from the telomere, respectively (Additional file 1: Fig. S1). The proportion of cells with silenced URA3 or HTB1-yEGFP can be determined as the percentage of 5-FOA resistant cells (FOAR, high sensitivity) or GFP-negative cells (low sensitivity), respectively . Loss of gene silencing is revealed by the decrease of the percentage of 5-FOAR or the proportion of GFP-negative cells in the mutant relative to wild-type isogenic strains. These two reporters had reproduced previously documented effects of various mutants in replication factors or histone chaperones [18, 19].
Using the 5-FOA resistance assay, we compared the impact of the pol30-6, pol30-8 and pol30-79 alleles on the expression of URA3 by the adh4-URA3-tel reporter , which had been used to characterize the pol30 mutants in prior publications, and our dual reporters (Fig. 1). Two concentrations of 5-FOA were applied. We found that the proportions of FOAR in the cells harboring the dual reporters were about 5 times lower that in the cells harboring adh4-URA3-tel (Fig. 1A–C). We attribute these differences to the longer distance between the telomere and URA3 and of the presence of the HTB1 promoter in-between (Additional file 1: Fig. S1). Importantly, all three reporters produced similar reductions of the percentage of FOAR cells in the various pol30 mutants (Fig. 1D, E). These results were in agreement with previous analyses of the pol30 mutants with the FOAR or the CRASH assays [9, 10, 12]. Taken together, we conclude that the dual reporters faithfully capture the effects of the pol30 mutations.
The higher concentration of 5-FOA revealed a larger difference in the percentage of FOAR-cells between the mutants and the POL30 strain (Fig. 1C). This outcome suggested that the two concentrations of 5-FOA detect a gradient of cellular URA3 expression and not a distinct bi-modal Active/Silent state of this gene. While it was apparent that the gradient was shifted towards higher expression of URA3 in the pol30 mutants, the mechanism that causes this shift was not clear. We attempted to resolve this issue by flow cytometry analyses of the expression of the juxtaposed HTB1-yEGFP reporters.
Lack of bi-modal active/silent expression of yEGFP←HTB1-tel
We used an isogenic POL30 strain with no yEGFP reporter to set the upper threshold for the GFP-negative cells, which are the functional equivalent of the 5-FOAR cells. Based on this criterium, the calculated percentage of GFP-negative cells harboring the adh4-URA3-yEGFP←HTB1-tel construct in the pol30 mutants displayed results consistent with the results with this construct in the FOAR assay; however, the proportions of GFP-negative cells were in the 75–98% range compared to 0.2–6% range of FOAR cells (compare Figs. 1C and 2A). Consequently, the calculated differences between the POL30 and the mutant pol30 strains followed a similar trend but were significantly smaller compared to the differences observed with the FOAR assay (compare Figs. 1D, E and 2B). Hence, there was a notable discrepancy in the detection of silenced reporters by the high sensitivity and low sensitivity assays. Importantly, the flow cytometry analyses revealed little effect of the pol30 mutations on the silencing of the yEGFP←HTB1-tel reporter (Fig. 2C). Equally important, there was no evidence for a bi-modal Active/Silent state of HTB1-yEGFP expression in the POL30-0, pol30-6 and pol30-79 strains, and a tiny population of cells with elevated GFP signal in the pol30-8 strain (Fig. 2C). These results supported the idea that the pol30 mutations cause a transient de-repression and not a conversion to a distinct active state of yEGFP←HTB1-tel reporter. This transient de-repression cannot be detected by a low sensitivity assay.
Bi-modal active/silent expression of HTB1→yEGFP-tel
We readily observed two distinct populations of GFP-positive and GFP-negative cells when the flow cytometry assays were performed with the adh4-URA3-HTB1→yEGFP-tel reporter (Fig. 2D). In the POL30 strain, the percentage of the GFP-negative cells was 60% and decreased to about 40% in the mutants. The differences between POL30 and the mutants were far smaller when compared to the differences detected by the FOAR assay with URA3, again supporting the idea that the FOAR assay is detecting transient de-repression and not Silent→Active epigenetic conversions. Importantly, the comparison between the adh4-URA3-HTB1→yEGFP-tel and adh4-URA3-yEGFP←HTB1-tel reporters pointed to a difference in the expression of HTB1-yEGFP depending on the position and orientation of the reporter and independent of the effect of the pol30 mutations (Fig. 2A, and B).
We noticed a higher spread of the GFP-positive population across the FSC-A axis. We performed a Pearson correlation analysis between FSC-A and EGFP-A compensated values for both orientations of the fragment. A linear positive correlation was observed in all strains, with strains harboring the HTB1→yEGFP-tel fragment showing stronger correlation (Additional file 1: Fig. S2A). Using microscopy, we followed up by measuring the size of the cells harboring no GFP or the HTB1-yEGFP fragments in both orientations. This analysis showed that all pol30 mutants have an increased cell size as compared to the wild-type counterparts (Additional file 1: Fig. S2B). However, we did not observe any statistically significant difference between the strains with and without HTB1-yEGFP. These analyses suggest that although higher GFP signal and FSC-A are associated, the cell size and the GFP signal statistical associations are not caused by the HTB1 promoter. Equally importantly, the single or bi-modal modes of expression of the reporters are not related to the size of the cells.
Analyses of the expression of HTB1-yEGFP by fluorescent microscopy
We followed with analyses of the expression of HTB1-yEGFP using fluorescent microscopy (Fig. 3A, B). The measurement of the intensity of GFP fluorescence in individual cells demonstrated that the adh4-URA3-HTB1→yEGFP-tel construct produced, on average, 3.5 times higher signals compared to the adh4-URA3-yEGFP←HTB1-tel construct across all strains (Fig. 3C, D). Both constructs produced higher signals in the pol30 mutants relative to the isogenic POL30-0 strain with pol30-8 cells showing the strongest effect (Fig. 3C, D). The percentage of GFP-positive cells was determined as follows. We averaged the intensity of signals in ROI (Region of Interest) with no cells and postulated a threshold of 1.25% higher signal for GFP-positive cells. By these criteria, in the POL30-0 strain the two constructs produced 60% and 94% GFP-negative cells, respectively (Fig. 3E). The reduction of GFP-negative cells in the pol30 mutant strains mirrored the effects observed by flow cytometry (Figs. 2C and 3E). Again, the pol30-8 mutation showed the strongest statistically significant effect regardless of the construct used, while only the adh4-URA3-HTB1→yEGFP-tel construct produced statistically significant reduction of silencing in the pol30-6 and pol30-79 strains (Fig. 3D). These analyses were in good agreement with the data obtained by flow cytometry and 5-FOA-resistance assays (Figs. 1 and 2).
Genetic interactions of pol30 mutants with CAC1 and ASF1
Previous studies have shown that the deletion of CAC1 has a synergistic negative effect on the gene silencing at the VIIL sub-telomere in pol30-6 and pol30-79, but not pol30-8 mutants [9, 10]. Conversely, in pol30-8 cells, the deletion of the histone chaperone ASF1 further reduced silencing at the sub-telomeres while having little effect in pol30-6 and pol30-79 . These studies were performed with the 5-FOA-resistance assay and did not distinguish between transient de-repression and/or epigenetic conversions of URA3. Therefore, we asked if the deletions of ASF1 and CAC1 in the pol30 mutants would present evidence for transient de-repression or a bi-modal Active/Silent state of the HTB1-yEGFP reporter.
First, we confirmed that URA3 in these reporters can also reproduce the previously reported observations of the pol30 mutations in cac1∆ and asf1∆ genetic backgrounds. The analyses were performed with the adh4-URA3-HTB1←yEGFP-tel construct, which produced slightly higher percentage of FOAR cells (Fig. 1C) and can more reliably detect the reduction of silencing in the cac1∆ and asf1∆ strains. We found that at 0.5 × 5-FOA the pol30 mutations had no statistically significant effect in the cac1∆ background (Fig. 4A, B). At the higher 1 × 5-FOA concentration, pol30-6 and pol30-79, but not pol30-8, exacerbated the silencing defects at URA3 in cac1∆ cell (Fig. 4C, D). In the asf1∆ background, the pol30-8 mutation markedly reduced the silencing of URA3 at both concentrations of 5-FOA (Fig. 4A, C), while pol30-6 and pol30-79 showed statistically significant loss of silencing only in 1 × concentration (Fig. 4C, D). Hence, our construct can reproduce the effects of cac1∆ and asf1∆ in the context of the three pol30 mutations [9, 10, 14]. Notably, the effects of cac1∆ and asf1∆ have been reproduced at different concentrations of 5-FOA, thus reiterating that subtle differences in the concentrations of 5-FOA could lead to different interpretations in different studies.
Next, we tested how the deletions of CAC1 and ASF1 affect the expression of HTB1-yEGFP in the two reporters. As previously shown in Fig. 2, the adh4-URA3-HTB1→yEGFP-tel reporter displayed a distinct bi-modal Active/Silent state of the expression of the HTB1-yEGFP (Fig. 5A, Additional file 1: Fig. S3A, B). The deletion of CAC1 decreased the proportion of GFP-negative cells in all four strains, with stronger effects in the pol30-6 and pol30-79 strains (Fig. 5C). The deletion of ASF1 had a stronger effect in the pol30-8 mutants (Fig. 5C).
The analyses with the adh4-URA3-yEGFP←HTB1-tel construct revealed a different picture. In Fig. 5B, we show that the deletion of CAC1 in the context of POL30 and the pol30 mutant backgrounds decreased the overall proportions of GFP-negative cells as demonstrated by the upward shift of the signals in the density plots. However, a clear bi-modal distribution of GFP-positive and GFP-negative cells was observed only in the pol30-8 mutant (Fig. 5B, Additional file 1: Fig. S3A, B). This, along with the data in Fig. 4, suggests that the deletion of CAC1 only leads to transient de-repression of both genes in this dual reporter. In contrast, the deletion of ASF1 produced a small but distinct proportion of GFP-positive cells in POL30 cells, which was increased in the pol30 mutants with pol30-8 having a more significant effect (Fig. 5B, C, Additional file 1: Fig. S3C, D). We conclude that at this position of the HTB1-yEGFP reporter the loss CAF1 function is leading to transient de-repression and that this effect is similar to the one caused by the deficiency of POL30 function. Conversely, the deficiency of Asf1p leads to a bi-modal Active/Silent state of the reporter. In both genetic backgrounds, the mutations in POL30 quantitatively exacerbate these effects, but have distinct qualitative effects.
Genetic interactions of pol30 mutants with RRM3
The replication forks frequently pause in the sub-telomeric regions of the chromosomes. The events of pausing are exacerbated in rrm3∆ mutants, but the exact positions of the pausing are not known [21, 22]. RRM3 encodes a DNA helicase necessary for the restart of paused replication forks . We considered the possibility that the positional effects observed with our dual reporters are linked to the pausing of the fork. If this assumption is correct, the deletions of RRM3 would have different effects on the expression of URA3 and HTB1-yEGFP in the two reporters.
In Fig. 6A, B, we show that the deletion of RRM3 in the POL30-0 and pol30-79 strains reduced the percentage of FOAR cell threefold to tenfold at 0.5 × or 1 × 5-FOA concentration, respectively. The deletion of RRM3 in the pol30-6 and pol30-8 strains did not show a significant additive effect (Fig. 6A, B, Additional file 1: Fig. S4A). We followed up by flow cytometry to measure the expression of HTB1-yEGFP. In both constructs, deletion of RRM3 produced minor, but statistically significant decreases in the percentage of GFP-negative cells in POL30-0 and pol30-79 strains and not in pol30-6 and pol30-8 strains (Fig. 6C, D Additional file 1: Fig. S4B, C). Hence, it seems unlikely that the observed positional effects in the expression of HTB1-yEGFP are caused by the pausing of the fork. At the same time, it is noteworthy that RRM3 synergistically interacts with POL30-0 and pol30-79, but not with the pol30-6 and pol30-8 alleles.
Physical interactions of the pol30 mutants with Cac1p and Rrm3p
Cac1p and Rrm3p contain a PIP (PCNA Interacting Peptide) and both are known to directly interact with Pol30p (PCNA) proteins in vitro [7, 8, 23,24,25]. We tested if these two proteins differentially interact with the mutant Pol30p proteins by a two-hybrid interaction assay as described previously . Cac1p and Rrm3p were used as baits and the wild type and mutant Pol30p as prey. In this assay, the three pol30 mutations reduced the binding to Cac1p to the levels observed with the negative cac1PIP∆ control in which the PCNA-Interacting-Peptide (PIP) sequence was destroyed (Fig. 7A). The interaction with Rrm3p was reduced by the pol30-6 and pol30-8 mutations, but not by the pol30-79 mutation.
We followed up by co-immunoprecipitation assays with Cac1p-FLAG and Rrm3p-Myc tagged proteins (Fig. 7B, C). These were expressed from low copy plasmids in the POL30-0 and the mutant pol30 strains and the immuno-precipitates were tested for the presence of Pol30p. In agreement with the two-hybrid assay and previous reports , the three Pol30p mutant proteins were found to bind poorly to Cac1p-FLAG (Fig. 7B, D). The interaction of Rrm3p was reduced fivefold by the pol30-6 mutation, less than twofold by pol30-8 and was not affected by pol30-79 (Fig. 7C, D). We conclude that all three mutations in POL30 affect the association with Cac1p, while pol30-6 and pol30-8 reduce the association with Rrm3p.
Transient loss of gene silencing in POL30 mutants
Earlier studies have reported that mutations in POL30 lead to loss of gene silencing at the sub-telomeric and mating type loci of S. cerevisiae and that different pol30 mutations have distinct, synergistic effects with the deletion of the histone chaperones CAF-1 and Asf1p [9, 10, 12]. In this study, we used the strains from  and conducted a detailed analysis with three different reporter constructs and three alternative assays. In all three constructs, we tested the expression of URA3 and reproduced the previously reported effects. However, the application of increasing concentrations of 5-FOA revealed significant differences in the proportions of 5-FOA-resistant and 5-FOA-sensitive cells (Fig. 1). These observations pointed to the possibility that the mutations in POL30 cause transient de-repression of URA3. This possibility was further supported by the comparison of FOAR assays data to the parallel analyses of the expression of the HTB1-yEGFP at two different positions in the VIIL sub-telomere. Indeed, the mutations in POL30 that lead to a dramatic reduction of the percentage of 5-FOA-resistant cells reduced the percentage of GFP-negative cells at a much lower magnitude (Fig. 2). Considering the higher sensitivity of the 5-FOA resistance assay, we favor the idea that the mutations in POL30 lead to transient de-repression of URA3 and HTB1-yEGFP and not to a higher incidence of conversions from silent to active state. This outcome is in agreement with the recent analyses of the silencing of the HML mating type locus by CRASH assay . Importantly, our results and interpretation add a significant detail to earlier studies [9, 12] and indicate that the most likely effects observed were through transient de-repression and not through the shifting of a bi-modal variegated state as previously thought.
Position effects in sub-telomeric gene silencing
We observed qualitative differences in the expression of HTB1-yEGFP when the promotor of the reporter was positioned distal or proximal to the telomere. In the HTB1→yEGFP-tel orientation the reporter displayed a clear bi-modal population of GFP+/GFP− cells, while in the yEGFP←HTB1-tel orientation, the reporter produced a single population of cells with GFP signals spreading below and above the threshold of signals from cells with no reporter cells. The pol30 mutations moderately reduced the silencing of both constructs (Fig. 2) but did not lead to a qualitative shift to a bi-modal expression in the yEGFP←HTB1-tel orientation (Figs. 2 and 3). Similarly, the deletions of CAC1 or RRM3 did not qualitatively shift the expression of yEGFP←HTB1-tel to a bi-modal state (Figs. 5, 6, Additional file 1: Figs. S3, S4), while the deletion of ASF1 did (Fig. 5, Additional file 1: Fig. S3).
These results clearly indicate that distinct modes of repression of epigenetically silenced genes can operate at different positions. Numerous studies had shown that a gene can be robustly and constitutively silenced. An example for such constitutive silencing would be the HML and HMR mating type loci in S. cerevisiae . Under the state of constitutive repression, various defects in cis-acting silencing factors or in the progression of replication forks can lead to transient de-repression or a bi-modal state, depending on the nature of the mutation [12, 27, 28]. A second mode of epigenetic silencing would be a normal bi-modal Active/Silent state. In this situation, defects in many genes engaged in chromatin maintenance and/or epigenetic transmission would affect the proportion of cells with Active or Silent state. An example for such bi-modal mode of repression would be the classical whiteV locus in Drosophila  or some, but not all, sub-telomeric genes in S. cerevisiae. Finally, here we present evidence for a state in which genes are transiently de-repressed but do not display a clearcut bi-modal Active/Silent state.
The distinction between these modes of silencing is important for the interpretations of different sets of results. For example, earlier studies unveiled major discrepancies in the magnitude of sub-telomeric silencing in S. cerevisiae when these were assessed by genome-wide array analyses or by 5-FOA-resistance assays [30,31,32]. It is possible that these analyses were detecting a mixture of bi-modal and transient effects, leading to mixed conclusions on the role of various genes involved in gene silencing. Similarly, discrepancies are observed when the silencing of the mating type loci in S. cerevisiae is assessed by the 5-FOA-resistance or the more sensitive CRASH assay.
At this point, we do not know what mechanisms underlay the modes of silencing and the roles of various silencing factors in them. However, based on our results (Figs. 4, 5 and 6), it is tempting to speculate that CAC1, POL30 and RRM3 are all involved in the prevention of excessive transient de-repression in the wake of the replication forks. One possibility is that the activity of these three genes is necessary for the timely maturation of replicated heterochromatin. On the other hand, ASF1 has a distinct role, and its loss can lead to bi-modal mode of expression.
Relevance to telomere position effect on native telomeres
Our synthetic reporters are devoid of sub-telomeric elements that are normally present at the native telomeres. Hence, our assays have their limitations and do not directly address the specific mechanisms of TPE at native telomeres. Nevertheless, our findings at an engineered telomeric locus are still bearing relevance to gene silencing at native telomeres, as well as other silenced loci in the genome of this and other organisms. They also challenge multiple earlier conclusions on the effects of various mutations on gene silencing in S. cerevisiae and add significant details on the stability of epigenetic state, its transmission and the perceived on/off variegated transcription at the heavily studied VIIL telomere. Equally importantly, our manuscript corrects a substantial body of work on the effects of POL3, CAC1 and ASF1 mutations. These prior studies were conducted using highly sensitive irreversible assays and do not necessarily address the stability of the silenced state. Finally, we complement previous work on the differences in the repression of genes at different positions in the sub-telomeres of S. cerevisiae [33,34,35] with the notion that it remains unclear if these differences are at the level of stability or at the level of on/off bi-modal expression.
Is the pausing of replication forks responsible for the observed position effects?
Prior studies have demonstrated that replication forks frequently pause in the sub-telomeric regions of the chromosomes, but the exact positions are not known [22, 36]. It is also known that the pausing of forks in the sub-telomeres is exacerbated by the deletion of RRM3 [22, 36]. We reasoned that a specific pause event can contribute to the different modes of repression of HTB1-yEGFP. If this is the case, the deletion of RRM3 in cells harboring the two constructs would have different effects. However, the deletion of RRM3 did not lead to a distinct silencing of the reporters in the wild type and pol30 mutant strains (Fig. 6). Therefore, we suggest that the pausing of the fork is unlikely to contribute to the differences in the repression of HTB1-yEGFP at the two positions.
Physical interactions of Pol30p(PCNA) with Cac1p and Rrm3p
Previous studies have demonstrated that Pol30p directly interacts with the Cac1p subunit of CAF-1 and with Rrm3p . It had also been reported that the in vitro interaction between Pol30p and Cac1p is severely diminished by the pol30-6 and pol30-79 mutations and to a lesser extent by the pol30-8 mutation . However, it is not known how these mutations affect the interaction with Rrm3p. We addressed this question by two-hybrid interaction and immunoprecipitation assays and reproduced the results of  (Fig. 7). In these assays, we obtained evidence that the pol30-6 and pol30-8, but not the pol30-79, mutations reduce the affinity to Rrm3p (Fig. 7). On the other hand, the deletion of RRM3 in the pol30-79, but not in the pol30-6 and pol30-8, had a synergistic negative effect on gene silencing (Fig. 6). These results are consistent with the interpretation that the loss of silencing phenotype in pol30-6 can be partially attributed to the impaired interaction with Rrm3p. At this point, we cannot comment on how the pol30-8 allele relates to RRM3. Rrm3p remains capable of physically interacting with Pol30-8p, albeit poorly (Fig. 7), yet the deletion of RRM3 in the pol30-8 strain had little effect on silencing (Fig. 6). We suspect that a complex interplay between Pol30p interacting factors is responsible for this lack of effect. The pol30-79 mutation is located on inter-domain connecting loop and has been shown to be defective in replication with POL32 (Polδ) . A recent structural analysis study has shown that all three pol30 mutations have structural anomalies at a common surface 10 Å away from PIP-binding site . The retention of Rrm3p by Pol30-79p and the additive silencing defect observed upon deletion of RRM3 in pol30-79 strain pose the question if Rrm3p binds to a non-canonical PIP binding site and plays an important role in maintaining fork stability in these defective strains.
Concluding remarks and significance
In this study, we have described two distinct modes of gene repression at the telomeres of S. cerevisiae: transient repression and a bi-modal Active/Silent state. We have demonstrated that mutations in POL30(PCNA) or the deletions of CAC1 and RRM3 affect the transient mode of repression, while the deletion of ASF1 leads to the conversion from transient mode to the Active/Silent state. Future analyses of gene silencing in this and other organisms should consider these two modes of repression.
Materials and methods
Yeast strains, plasmids, and primers
The POL30 and the mutant pol30 strains are identical to the ones published in . The derivatives of these strains were produced by PCR-directed deletion of the target genes as described in , and were PCR confirmed. The reporter fragments were inserted at the VIIL telomere via electroporation and were confirmed by PCR. The adh4-URA3-tel construct was identical to the one described in . The adh4-URA3-HTB1→yEGFP-tel and adh4-URA3-yEGFP←HTB1-tel reporters were described in [18, 19]. All strains are listed in Additional file 2: Table S1. Primers used for knocking out genes are listed in Additional file 2: Table S2. The recombinant strains were routinely maintained in YPD medium (1% yeast extract, 2% tryptone, 2% glucose) at 30 °C. Flow cytometry and fluorescent microscopy experiments were conducted in Synthetic Complete (SC) media. Plasmids used in two hybrid assay and co-immunoprecipitation experiments are listed in Additional file 2: Table S3. Generation of the expression plasmids are described in . pol30 mutations were introduced in pBL240-PCNA-Gal4AD prey plasmid using site directed mutagenesis. Primers used for site-directed mutagenesis are listed in Additional file 2: Table S2. The plasmid was amplified with partially overlapping primers harboring pol30-6, pol30-8 and pol30-79 mutations using PaCeR™ HP™ Master Mix (GeneBioSystems#PCR-002-01). The PCR products were digested with Dpn1 and transformed in E. coli DH5α. Plasmids extracted from E. coli was sequenced to confirm insertion of the mutations and were transformed in yeast strain W303 using lithium acetate transformation. Plasmids utilized in co-immunoprecipitation experiment were introduced in POL30 and mutant pol30 strains using electroporation. Strains for co-immunoprecipitation and two hybrid experiments were grown in Synthetic Drop-out (SD) media, as appropriate.
5-FOA sensitivity assay
Cells were grown in YPD at 30 °C overnight. Saturated cultures were serial-diluted by 1:10 and 5 μL of each dilution was spotted on YPD, SC-Ura, 0.5 × 5-FOA and 1 × 5-FOA plates. Final 5-FOA concentrations were 5 mg/mL in 0.5 × 5-FOA plates and 10 mg/mL in 1 × 5-FOA plates (BioBasic#703-95-7). All plates were incubated at 30 °C for 3–5 days. The colonies were counted with a Gallenkamp colony counter and the percentage of 5-FOA resistant cells were calculated and plotted using Microsoft Excel®.
Cells were grown in synthetic complete media up to OD600 = 1 and washed with phosphate buffer saline. Resuspended cells were sonicated for two cycles of 30 s ON/10 s OFF with a Mandel Scientific ultrasonic sonicator at 50% output to disperse cell clusters. 488 nm laser of Sony SH800z flow cytometer was used to detect GFP, and LESH00SZFCPL™ software was used to generate the density plots and to analyze data. 100,000 events were screened for each strain, and GFP negative gate was set based on an isogenic strain lacking GFP. Data from at least three independent experiments were pooled in Microsoft Excel® to create %GFP negative graph and calculate standard deviation.
Cells were grown in SC media and 2 µL of the cell suspension were analyzed on 8-well microscope slides. Images were captured by Leica DM600B with brightfield and green fluorescent channels at 40× magnification. Volocity™ software was used to quantify GFP intensity and number of GFP positive cells. The intensity of the GFP signal in individual cells was determined as follows. The pixel values of 100 cell-free ROI (regions of interest) were averaged and deemed background. The pixel values in ROI over at least 100 cells were then measured and plotted in distribution graph. The evaluation of the percentage of GFP+ cells was determined as described in . A negative threshold was set at 125% of average background intensity and cells with intensity scores greater than the set threshold was counted as GFP positive. Images from three independent experiments were used to show the GFP distribution, calculate % GFP negative population and standard deviation.
Yeast two-hybrid assay
Cells containing reporter, bait and prey plasmids were grown in SC/Ura−/Leu−/His−/2% glucose media at 30 °C up to OD600 0.8–1. Cells were then collected, washed with cold H2O, and incubated in SC/Ura−/Leu−/His−/2% galactose/1% raffinose media at 30 °C for 4 h. Cells were then harvested and washed with ice-cold H2O and resuspended in 3 mL of buffer P (50 mM sodium phosphate, pH 7.7, 300 mM sodium acetate, 1 mM 2-Mercaptoethanol, 500 nM DTT, 10% v/v glycerol, 1 mM PMSF and 1% v/v protease inhibitor cocktail (Sigma#P8215)). The cells were split in three aliquots, pelleted, and resuspend in 200 µL Buffer P to extract protein in three replicates. 200 µL of 0.55 mm glass beads (Cole-Parmer BioSpec#11079105) were added to each tube and vortexed at 30 s on/10 s off pulse for 15 min at 4 °C. The cell lysates were centrifuged to remove debris and 50 µL of the extracts were aliquoted to measure total protein concentration. On a 96-well plate, extracts from each replicate were serially diluted by 1:10 with Buffer P and 4 mg/mL ortho-Nitrophenyl-β-galactoside (ONPG) (Sigma #N1127-1G) was added to each well. The plate was incubated at 30 °C and a timer was started immediately to record the time it takes to develop a medium-yellow color in the positive control wells. Thermo Scientific Multiskan GO plate reader was used to record the absorbance of the wells at 420 nm and 550 nm, and the time of reading was recorded. The absorbance readings were taken at three different timepoints. The units of β-galactosidase were calculated using the following formula:
Total protein concentrations were measured using Bio-Rad protein assay (#500-0006) and used to normalize the β-galactosidase signal as units of β-galactosidase/mg of protein. Data from three independent experiments were pooled to calculate averages and standard deviations.
Cells harboring Cac1p-FLAG and Rrm3p-Myc expressing plasmids were grown in SC/Leu−/Lys−/2% glucose at 30 °C up to OD600 0.8–1. The cells were harvested, washed with ice-cold H2O and resuspended in 300 µL IP Buffer (150 mM NaCl, 50 mM Tris (pH7.5), 2 mM EDTA, 5 mM NaF, 5 mM β-Glycerophosphate, 0.1 mM NaVO3, 1% v/v protease inhibitor cocktail (Sigma#P8215) and 1 mM PMSF). 300 µL of 0.55 mm glass beads (Cole-Parmer BioSpec#11079105) were added to each tube and vortexed at 30 s on/10 s off pulse for 15 min at 4 °C. The cell lysates were centrifuged to remove debris and 50 µL of the extracts were aliquoted to load as input protein. The remaining lysate was split in half and used for α-FLAG and α-Myc pulldowns using Rat α-DYKDDDDK (Cell Signaling#14793S) and Mouse α-Myc (9E10) antibodies, respectively. The lysates were precleared with α-Rabbit/α-Mouse Ig agarose and nutated with 1ug of respective antibodies for 2 h at 4 °C. α-Flag antibody was pulled down with Protein A Sepharose beads (Invitrogen#101041), α-Myc antibody was pulled down with Protein G Sepharose beads (SinoBiological Inc#13103-PNAE-RN). The beads were washed three times with IP buffer + 0.2% Triton X100 + 0.01% SDS and eluted with 2µg of synthetic FLAG peptide (SinoBiological Inc# PP101274) or Myc peptide (SinoBiological Inc# PP100029).
Extracts and elutes from co-immunoprecipitation were boiled in 2X Laemmli loading buffer and resolved in SDS-10% polyacrylamide gels. Proteins were subsequently transferred on polyvinylidene difluoride (PVDF) membrane using Bio-Rad Trans-Blot® SD semi-dry transfer cell at 18 V for 18 min. The membranes were blocked in TBST + 5% skim milk, cut in half, and incubated in parallel with α-FLAG/α-Myc and α-PCNA primary antibodies followed by α-rabbit/α-mouse horseradish peroxidase (HRP) conjugated secondary antibodies as appropriate. The blots were visualized using Enhanced Chemiluminescence (ECL) substrate (Bio-Rad #1705061) and imaged using Bio-Rad ChemiDoc™ XRS + with Image Lab™ software. Quantification of the signals were performed using ImageJ. PCNA band intensities in load and elute were normalized against the corresponding Cac1p/Rrm3p band intensities, and then elute intensities were expressed as a factor of corresponding input intensities. Data from at least two biological replicates were pooled to create bar graph and calculate standard deviation.
Data analysis and statistics
Microsoft Excel® was used to calculate averages, standard deviations and to generate graphs from all experiments. Statistical analysis was performed using IBM SPSS. P values were determined by one-way ANOVA followed by post hoc Dunnett’s test for multiple comparisons, and by independent sample T test for pair-wise comparisons. P values < 0.05 were considered significant.
Availability of data and materials
All data generated and analysed in this study are included in this article and its additional files.
Riddle NC, Shaffer CD, Elgin SC. A lot about a little dot - lessons learned from Drosophila melanogaster chromosome 4. Biochem Cell Biol. 2009;87(1):229–41.
Yankulov K. Dynamics and stability: epigenetic conversions in position effect variegation. Biochem Cell Biol. 2013;91(1):6–13.
Moldovan GL, Pfander B, Jentsch S. PCNA, the maestro of the replication fork. Cell. 2007;129(4):665–79.
Choe KN, Moldovan GL. Forging ahead through darkness: PCNA, still the principal conductor at the replication fork. Mol Cell. 2017;65:380.
Collins SR, Miller KM, Maas NL, Roguev A, Fillingham J, Chu CS, et al. Functional dissection of protein complexes involved in yeast chromosome biology using a genetic interaction map. Nature. 2007;446(7137):806.
Costa A, Diffley JFX. The initiation of eukaryotic DNA replication. Ann Rev Biochem. 2022;91:107.
Gopinathan Nair A, Rabas N, Lejon S, Homiski C, Osborne MJ, Cyr N, et al. Unorthodox PCNA binding by chromatin assembly factor 1. Int J Mol Sci. 2022;23(19):11099.
Shibahara K, Stillman B. Replication-dependent marking of DNA by PCNA facilitates CAF-1-coupled inheritance of chromatin. Cell. 1999;96(4):575–85.
Zhang Z, Shibahara K, Stillman B. PCNA connects DNA replication to epigenetic inheritance in yeast. Nature. 2000;408(6809):221–5.
Sharp JA, Fouts ET, Krawitz DC, Kaufman PD. Yeast histone deposition protein Asf1p requires Hir proteins and PCNA for heterochromatic silencing. Curr Biol. 2001;11(7):463–73.
Miller A, Yang B, Foster T, Kirchmaier AL. Proliferating cell nuclear antigen and ASF1 modulate silent chromatin in Saccharomyces cerevisiae via lysine 56 on histone H3. Genetics. 2008;179(2):793–809.
Brothers M, Rine J. Mutations in the PCNA DNA polymerase clamp of Saccharomyces cerevisiae reveal complexities of the cell cycle and ploidy on heterochromatin assembly. Genetics. 2019;213(2):449–63.
Krishna TS, Kong XP, Gary S, Burgers PM, Kuriyan J. Crystal structure of the eukaryotic DNA polymerase processivity factor PCNA. Cell. 1994;79(7):1233–43.
Jeffery DC, Kakusho N, You Z, Gharib M, Wyse B, Drury E, et al. CDC28 phosphorylates Cac1p and regulates the association of chromatin assembly factor I with chromatin. Cell Cycle. 2015;14(1):74–85.
Janke R, King GA, Kupiec M, Rine J. Pivotal roles of PCNA loading and unloading in heterochromatin function. Proc Natl Acad Sci U S A. 2018;115(9):E2030–9.
Stewart-Morgan KR, Petryk N, Groth A. Chromatin replication and epigenetic cell memory. Nat Cell Biol. 2020;22:361.
Rowlands H, Dhavarasa P, Cheng A, Yankulov K. Forks on the run: can the stalling of DNA replication promote epigenetic changes? Front Genet. 2017;8:86.
Mano Y, Kobayashi TJ, Nakayama J, Uchida H, Oki M. Single cell visualization of yeast gene expression shows correlation of epigenetic switching between multiple heterochromatic regions through multiple generations. PLoS Biol. 2013;11(7): e1001601.
Shaban K, Sauty SM, Fisher A, Cheng A, Yankulov K. Evaluation of drug-free methods for the detection of gene silencing in Saccharomyces cerevisiae. Biochem Cell Biol. 2023;101(1):125.
Gottschling DE, Aparicio OM, Billington BL, Zakian VA. Position effect at S. cerevisiae telomeres: reversible repression of Pol II transcription. Cell. 1990;63(4):751–62.
Ivessa AS, Zhou JQ, Zakian VA. The Saccharomyces Pif1p DNA helicase and the highly related Rrm3p have opposite effects on replication fork progression in ribosomal DNA. Cell. 2000;100(4):479–89.
Makovets S, Herskowitz I, Blackburn EH. Anatomy and dynamics of DNA replication fork movement in yeast telomeric regions. Mol Cell Biol. 2004;24(9):4019–31.
Rolef Ben-Shahar T, Castillo AG, Osborne MJ, Borden KL, Kornblatt J, Verreault A. Two fundamentally distinct PCNA interaction peptides contribute to chromatin assembly factor 1 function. Mol Cell Biol. 2009;29(24):6353–65.
Schmidt KH, Derry KL, Kolodner RD. Saccharomyces cerevisiae RRM3, a 5′ to 3′ DNA helicase, physically interacts with proliferating cell nuclear antigen. J Biol Chem. 2002;277(47):45331–7.
Wyse B, Oshidari R, Rowlands H, Abbasi S, Yankulov K. RRM3 regulates epigenetic conversions in Saccharomyces cerevisiae in conjunction with chromatin assembly factor I. Nucleus. 2016;7(4):405–14.
Rusche LN, Kirchmaier AL, Rine J. The establishment, inheritance, and function of silenced chromatin in Saccharomyces cerevisiae. Annu Rev Biochem. 2003;72:481–516.
Fouet M, Rine J. Limits to transcriptional silencing in Saccharomyces cerevisiae. Genetics. 2023;223(2): iyac180.
Saxton DS, Rine J. Nucleosome positioning regulates the establishment, stability, and inheritance of heterochromatin in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A. 2020;117(44):27493.
Henikoff S. Gene Silencing in Drosophila. In 1995. p. 193–208.
Ellahi A, Thurtle DM, Rine J. The chromatin and transcriptional landscape of native Saccharomyces cerevisiae telomeres and subtelomeric domains. Genetics. 2015;200(2):505.
Rossmann MP, Luo W, Tsaponina O, Chabes A, Stillman B. A common telomeric gene silencing assay is affected by nucleotide metabolism. Mol Cell. 2011;42(1):127–36.
Takahashi YH, Schulze JM, Jackson J, Hentrich T, Seidel C, Jaspersen SL, et al. Dot1 and histone H3K79 methylation in natural telomeric and HM silencing. Mol Cell. 2011;42(1):118–26.
Pryde FE, Louis EJ. Limitations of silencing at native yeast telomeres. EMBO J. 1999;18(9):2538–50.
Fourel G, Revardel E, Koering CE, Gilson E. Cohabitation of insulators and silencing elements in yeast subtelomeric regions. EMBO J. 1999;18(9):2522–37.
Power P, Jeffery D, Rehman MA, Chatterji A, Yankulov K. Sub-telomeric core X and Y’ elements in S cerevisiae suppress extreme variations in gene silencing. PLoS ONE. 2011;6(3): e17523.
Ivessa AS, Zhou JQ, Schulz VP, Monson EK, Zakian VA. Saccharomyces Rrm3p, a 5’ to 3’ DNA helicase that promotes replication fork progression through telomeric and subtelomeric DNA. Genes Dev. 2002;16(11):1383–96.
Eissenberg JC, Ayyagari R, Gomes XV, Burgers PM. Mutations in yeast proliferating cell nuclear antigen define distinct sites for interaction with DNA polymerase delta and DNA polymerase epsilon. Mol Cell Biol. 1997;17(11):6367.
Kondratick CM, Litman JM, Shaffer KV, Washington MT, Dieckman LM. Crystal structures of PCNA mutant proteins defective in gene silencing suggest a novel interaction site on the front face of the PCNA ring. PLoS ONE. 2018;13(3):e0193333–e0193333.
We thank Dr. J. Rine for the donation of the pol30 mutant strains used in this study; the MCB Imaging Facility, University of Guelph, for the technical support; Jakob Esch for help with Co-IP strain generation; Ashley Fisher and Andrew Dolson for helpful discussions.
Research in Yankulov lab is supported by a Discovery grant from NSERC to KY (RGPIN-2015-06727). SMS is supported by a bursary from the College of Biological Science at the University of Guelph.
Ethics approval and consent to participate
Consent for publication
The authors declare no competing interest.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Figure S1. Schematic of the reporter fragments used in this study. Figure S2. Cell size analysis. Figure S3. GFP signal distribution by flow cytometry. Figure S4. Genetic interaction of RRM3 with pol30 mutants.
Table S1. List of strains used in this study. Table S2. List of primers used in this study. Table S3. List of plasmids used in this study.
About this article
Cite this article
Sauty, S.M., Yankulov, K. Analyses of POL30 (PCNA) reveal positional effects in transient repression or bi-modal active/silent state at the sub-telomeres of S. cerevisiae. Epigenetics & Chromatin 16, 40 (2023). https://doi.org/10.1186/s13072-023-00513-7