Crosstalk between chromatin structure, cohesin activity and transcription

Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 https://doi.org/10.1186/s13072‑019‑0293‑6 RESEARCH Crosstalk betweenchromatin structure, cohesin activity andtranscription Douglas Maya‑Miles1†, Eloísa Andújar2†, Mónica Pérez‑Alegre2†, Marina Murillo‑Pineda1,3, Marta Barrientos‑Moreno1, María J. Cabello‑Lobato1,4, Elena Gómez‑Marín1, Macarena Morillo‑Huesca1 and Félix Prado1* Abstract Background: A complex interplay between chromatin and topological machineries is critical for genome architec‑ ture and function. However, little is known about these reciprocal interactions, even for cohesin, despite its multiple roles in DNA metabolism. Results: We have used genome‑wide analyses to address how cohesins and chromatin structure impact each other in yeast. Cohesin inactivation in scc1‑73 mutants during the S and G2 phases causes specific changes in chromatin structure that preferentially take place at promoters; these changes include a significant increase in the occupancy of the − 1 and + 1 nucleosomes. In addition, cohesins play a major role in transcription regulation that is associated with specific promoter chromatin architecture. In scc1‑73 cells, downregulated genes are enriched in promoters with short or no nucleosome‑free region (NFR) and a fragile “nucleosome − 1/RSC complex” particle. These results, together with a preferential increase in the occupancy of nucleosome − 1 of these genes, suggest that cohesins promote transcription activation by helping RSC to form the NFR. In sharp contrast, the scc1‑73 upregulated genes are enriched in promoters with an “open” chromatin structure and are mostly at cohesin‑enriched regions, suggesting that a local accumulation of cohesins might help to inhibit transcription. On the other hand, a dramatic loss of chromatin integrity by histone depletion during DNA replication has a moderate effect on the accumulation and distribution of cohesin peaks along the genome. Conclusions: Our analyses of the interplay between chromatin integrity and cohesin activity suggest that cohesins play a major role in transcription regulation, which is associated with specific chromatin architecture and cohesin‑ mediated nucleosome alterations of the regulated promoters. In contrast, chromatin integrity plays only a minor role in the binding and distribution of cohesins. Keywords: Chromatin, Cohesin, Transcription, Scc1 © The Author(s) 2019. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creat iveco mmons .org/licen ses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creat iveco mmons .org/ publi cdoma in/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Background The first level of structural organization of chromosomes is the chromatin fiber, in which DNA is assembled into regularly spaced nucleosomes through the coordinated activity of histone chaperones, chromatin assembly factors and nucleosome modifiers and remodellers. Assembly of replicated DNA is coupled to the replication fork and occurs by the deposition of both newly synthesized and parental histones, which are distributed randomly between the sister chromatids [1]. In addition, replication-independent mechanisms of nucleosome assembly reset the chromatin changes induced by processes like transcription or DNA repair during the cell cycle [2, 3]. Chromatin fiber permits several levels of compaction of DNA into the nucleus, depending on the nuclear functional state or the cell cycle phase. Although the exact molecular structure behind compaction is still Open Access Epigenetics & Chromatin *Correspondence: [email protected] †Douglas Maya‑Miles, Eloísa Andújar and Mónica Pérez‑Alegre contributed equally to this work 1 Department of Genome Biology, Andalusian Molecular Biology and Regenerative Medicine (CABIMER), CSIC‑University of Seville‑ University Pablo de Olavide, Seville, Spain Full list of author information is available at the end of the article Page 2 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 unclear, current models suggest that the packaging of chromatin requires dynamic and regulated interactions between irregularly folded fibers of nucleosomes, leading to a meshwork of intramolecular contacts [4]. The compaction, dynamics and regulation of this network rely mostly on the coordinated activities of topological machineries: cohesins, condensins and topoisomerases [5]. Whereas topoisomerases change the DNA supercoiling by cutting DNA molecules and re-sealing them in a new topological state [6], cohesins and condensins are ring-shaped molecules able to hold together distant chromatin fragments, thus compacting chromosomes [7]. In budding yeast, cohesins are loaded at chromatin by the cohesin loader complex Scc2/Scc4 during G1 and early S phase in a non-stable conformation that promotes topological and non-topological interactions (depending on whether or not cohesin entraps chromosomal DNA inside its ring) through dynamic turnover [8–10]. After loading, cohesins are moved away by the transcriptional machinery and accumulate preferentially at intergenic regions (IGRs) [11–14]. A fraction of cohesins becomes cohesive during S phase by topologically entrapping the two sister chromatids [15–18] and remains stably bound until its degradation in mitosis [19] (reviewed in [20, 21]). Although essential, sister chromatid cohesion is not the only function of cohesins, and multiple roles in DNA compaction, transcription regulation, DNA repair and DNA replication have been revealed in the past few years [22–25]. Remarkably, Scc2/Scc4 has been reported to have a role in RSC-mediated chromatin remodeling and transcription [26]. Specifically, the chromatin remodeling complex RSC recruits Scc2/Scc4 to specific promoters, where the cohesin loader helps to maintain the nucleosome-free region (NFR) for transcription activation [26]. RSC also interacts with and is required for cohesin loading [27] through a mechanism that involves direct interactions between RSC and both cohesin and Scc2/ Scc4 [28]. However, cohesins and cohesin loaders accumulate at non-overlapping peaks along the genome after their loading [12]. Therefore, it is unclear whether or not cohesins take part in chromatin remodeling and/or transcription in yeast. Nucleosomes seem to have an inhibitory effect on cohesin binding. The Snf2-related complexes RSC and Irc5 in yeast, and SNF2h in humans are required for cohesin binding to chromosomes [27, 29, 30]. Importantly, although the chromatin remodeling activity of RSC is not necessary for cohesin loader recruitment, an ATPase dead RSC complex is defective for cohesin loading; accordingly, nucleosomes interfere with cohesin loading invitro [28]. The distribution of cohesins is also associated with NFRs [31], although this preference might be an indirect consequence of their accumulation at IGRs [11, 13, 32], which contain NFRs [33, 34]. To address the connection between chromatin structure and cohesin function, we have now studied whether disrupting one affects the other. For this, we allow cells to progress from G1 to mitosis under conditions of histone depletion and/or lack of cohesin activity. Defective chromatin positioning by histone depletion had little or only region-specific effects on the accumulation and distribution of the major cohesin peaks. In contrast, the lack of cohesin activity affected the primary structure of chromatin at specific genomic regions, preferentially promoters. Critically, genome-wide analyses revealed a major role for cohesins in transcription regulation that is associated with the promoter chromatin architecture and location of the regulated genes. Results Cohesins contribute tostructuring chromatin To analyze what impact, if any, cohesins have on chromatin structure, we performed high-throughput sequencing of MNaseI-digested chromatin (MNase-seq) followed by dynamic analysis of nucleosome position and occupancy by sequencing (DANPOS) [35]. This approach allows nucleosomes to be mapped along the whole genome and categorizes the altered ones as changed in occupancy (measure of nucleosome density), position shift and fuzziness (degree of nucleosome deviation from its preferred position in a population). The experiment was performed with cells that express a thermosensitive allele of SCC1 (scc1-73). The α-kleisin subunit Scc1 forms the tripartite ring-like structure with the SMC (structural maintenance of chromosomes) subunits Smc1 and Smc3 in the cohesin complex. The scc1-73 allele encodes a mutant protein (S525N) that loses its ability to interact with Smc1/Smc3 at 37°C and thereby causes cohesin inactivation [8, 36]. Cells were grown and synchronized in G1 at 26 °C, released at 37°C and arrested in metaphase with nocodazole. This strategy ensures analysis of cells that have completed replication without cohesin activity and that have accumulated at the same cell cycle stage. The global profile of nucleosomes was similar in both scc1-73 and wild-type cells (Fig.1a). Accordingly, the lack of cohesin activity did not affect the distribution of nucleosome fuzziness scores, neighboring distances or occupancy periodicities (Fig. 1b). However, some changes were observed when nucleosome occupancy was aligned for all genes relative to the transcription start site (TSS). Thus, although the pattern of nucleosome positioning at both sides of the promoter-associated NFR was apparently unaffected (Fig.1c), the lack of cohesin activity caused a significant increase in the occupancy of nucleosomes − 1 and + 1, but not in the occupancy of the nucleosomes in Page 3 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 the gene body (+ 2 to + 5) (Fig.1d, asterisks). In addition, DANPOS analysis revealed an elevated number of altered nucleosomes in scc1-73 cells as compared to wild-type cells (~ 1.1% of total nucleosomes; Table1). Most alterations affected the nucleosome occupancy, with a similar number of nucleosomes displaying either higher or lower c d b a scc1-73 t::HHF2 scc1-73wt t::HHF2 scc1-73 t::HHF2 t::HHF2 scc1-73 wt t::HHF2 RRP7 HIS4 BIK1 RNQ1FUS1 FRM2 ARS 306 HBN1 scc1-73 t::HHF2 scc1-73 wt nucleosome profiles t::HHF2 Chr. III (65-76 kb) 210 120 60 30 240 150 90 270 000 210 120 60 30 240 150 90 270 0 150 100 50 25 175 125 75 225 200 150 100 50 25 175 125 75 225 200 occupancy (reads) occupancy (reads) genes -500 0 500 1000 1500 -500 0 500 1000 1500 -500 0 500 1000 1500 -500 0 500 1000 1500 1 kb scc1-73 wt distance to TSS (bp) 100 60 40 120 140 80 160 -500 0 500 1000 1500 ** * * * * * distance to TSS (bp) 100 60 40 120 140 80 160 -500 0 500 1000 1500 t::HHF2 wt distance to TSS (bp) 100 60 40 120 140 80 160 -500 0 500 1000 1500 distance to TSS (bp) distance to TSS (bp) distance to TSS (bp) distance to TSS (bp) distance to TSS (bp) 100 60 40 120 140 80 160 -500 0 500 1000 1500 fuzziness (bp) 0 7565554535 85 1 2 3 4 nucleosomes (%) distance (bp) 0 220200180160140120100 240 15 20 10 5 25 nucleosomes (%) scc1-73 t::HHF2 scc1-73 wt t::HHF2 occupancy (reads) 0 4003002001000 6 8 4 2 nucleosomes (%) Fig. 1 Genome‑wide nucleosome profiles in cohesin and histone deposition mutants. a Representative nucleosome profile by MNase‑seq of wild type, scc1‑73, t::HHF2 and t::HHF2 scc1‑73 cells synchronized in G1 and released until metaphase in nocodazole‑containing (15 μg/ml) medium under conditions of restrictive temperature (37 °C) and histone depletion. b Histone depletion affects nucleosome fuzziness, distance and occupancy. Distribution of nucleosome fuzziness scores, neighboring distances and occupancy periodicities were determined by DANPOS analyses. c and d Heat map (c) and occupancy profile (d) of nucleosomes for all yeast genes aligned relative to the transcription start site (TSS). The occupancy of nucleosomes − 1 and + 1 was specifically increased in scc1‑73 cells. Statistically significant differences in occupancy at the peak of nucleosomes − 2 to + 5 between the wild type and the scc1‑73 mutant are shown (paired two‑tailed Student’s t test; *, p < 0.05; ***, p < 0.001) Page 4 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 occupancy than in wild-type cells. Further analysis of the altered nucleosome distribution in the genome showed that they were enriched in IGRs, tRNA genes and telomeres; averagely distributed in the pericentric chromatin and rDNA; and had a less than average distribution in the ORFs (Additional file1: TableS1 and Additional file2: TableS2). We conclude that cohesins contribute to establish the primary structure of chromatin at specific DNA regions, preferentially promoters. Lack ofcohesin activity hasamajor effect intranscription thatislinked tospecific chromatin structure atpromoters As the effect of scc1-73 was more prominent on the chromatin of IGRs and the occupancy of nucleosomes − 1 and + 1, we asked if those chromatin changes are associated with alterations in the pattern of transcription regulation. We synchronized wild-type and scc1-73 cells in G1 and then released them into fresh medium until metaphase under restrictive conditions, to compare their genome-wide transcription profiles by RNA-seq under the same conditions that were used to study their chromatin structure. The absence of cohesin activity from G1 to metaphase in the scc1-73 mutant yielded an elevated number of misregulated genes, with 445 downregulated genes and 569 upregulated genes (q value < 0.01; 1.41- fold cutoff relative to wild type); this represents ~ 15% of all yeast genes. (Noncoding RNAs were not affected.) Of these, 255 genes (~ 4%; 91 downregulated and 164 upregulated) showed a > 2-fold change (Fig.2a and Additional file3: TableS3). These genes showed a significant overrepresentation of gene ontology biological process terms related to RNA and amino acids metabolism and stress responses (Additional file3: TableS3). Next, we asked whether gene regulation by cohesins is associated with chromatin alterations. Misregulated genes upon cohesin inactivation in the scc1-73 mutant were not associated with DANPOS-defined altered nucleosomes, indicating that these alterations were not caused by defective transcription. The only exception was a slight enrichment in nucleosomes with higher occupancy at the promoters of the downregulated genes (Fig.2b). In line with this, the increase in the occupancy of nucleosome − 1 remained significant in the scc1-73 downregulated (but not upregulated) genes, even though it was lost in control groups (chromosomes VII and XII) with a similar number of genes (Fig.2c). (Note that the number of genes analyzed in Fig.1d was ~ 10 times higher than that in Fig.2c.) The downregulated genes in scc1-73 cells were also characterized by significant changes at the nucleosomes downstream of the promoter, which might result from defective transcription. In contrast to these changes, the increase in the occupancy of nucleosome + 1 was also detected in control groups of transcriptionally unaffected genes (Fig.2c, chromosomes VII and XII), indicating that it was transcription independent. We next addressed whether the effects that a lack of cohesin activity had on transcription were associated with the promoter chromatin architecture. In yeast, gene promoters can be grouped according to their chromatin structure. One of these classifications distinguishes genes with occupied-proximal nucleosome (OPN) and depleted-proximal nucleosome (DPN) promoters, depending on the absence or presence of an NFR just upstream of nucleosome + 1 [37]. A similar analysis defines a higher number of genes as “open” or “closed” depending on the presence or absence of a clear NFR at the promoter [38]. Notably, scc1-73 downregulated genes were highly enriched in OPN and “closed” genes (Table2). More recent and exhaustive studies involving deep sequencing of MNaseI-treated DNA at different degrees of digestion uncovered two groups of promoters, defined according to the stability of nucleosome − 1: (i) promoters with a nuclease-resistant nucleosome (stable nucleosome, SN), which have a constitutive NFR of less than 150 bp, and (ii) promoters with a nuclease-sensitive nucleosome (fragile nucleosome, FN), which is removed through the action of transcription factors; these factors, together with the chromatin remodeling complex RSC, lead to an NFR of more than 150bp upon transcription activation [39, 40]. Notably, scc1-73 downregulated genes are highly enriched in FN promoters (Table2) and accordingly in promoters with RSC/nucleosome complexes [41] (Table2). Finally, we asked if scc1-73 downregulated genes were preferentially associated with Scc2/Scc4-regulated genes, as RSC-mediated chromatin remodeling and transcription activation requires Scc2/ Scc4 promoter binding [26]. We did not find a preferential association between Scc2-binding genes and Table 1 Number of altered nucleosomes (in position, fuzziness and/or occupancy) upon histone depletion (in t::HHF2 cells) or cohesin inactivation (in scc1-73 cells) as compared to the profile of nucleosomes in wild-type cells * The percentage of altered nucleosomes between t::HHF2 cells (6.68%) and scc1‑73 t::HHF2 cells (9.28%) was statistically different according to a two‑tailed Chi‑square test (p < 0.001) t::HHF2 scc1-73 t::HHF2 scc1-73 Altered nucleosomes 4431* 768 6151* Position shift 319 35 389 Fuzziness change 1516 77 2093 Occupancy change 3989 703 5618 Occupancy increase 1304 375 2080 Occupancy decrease 2685 328 3538 Page 5 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 scc1-73-misregulated genes (Table 2). Therefore, transcription activation by cohesins seems to be linked to specific chromatin structure at promoters. These results, together with a preferential increase in the occupancy of nucleosome − 1 of scc1-73-downregulated genes (Fig. 2c), suggest that cohesins promote transcription activation by helping RSC to form the NFR. Approximately 20% of yeast genes contain a TATA box [42]. This element is enriched in the OPN genes and underrepresented in the DPN genes [37], and accordingly, the TATA box was highly enriched among the downregulated genes in scc1-73 cells (Table 2). Early studies analyzing steady-state RNA levels showed that TATA-containing genes preferentially use the SAGA complex rather than the general transcription factor TFIID [42]. Similar studies have revealed a predominant role for TFIID (~ 90% of genes) and a more restricted role for SAGA (~ 10% of genes) [43]. More recent analyses have also shown that the Mediator complex is particularly important for the expression of TATA-containing, SAGA-dominated genes, whereas the histone acetyltransferase complex NuA4 is preferentially associated with TFIID [44, 45]. As expected from these associations, the set of downregulated genes in scc1-73 cells were dominated by the Mediator and SAGA and not by NuA4 and TFIID; in addition, they were enriched in SWR1- dominated genes [46] (Table 2). In marked contrast with scc1-73-downregulated genes, scc1-73-upregulated genes were enriched in genes with “open” promoters and dominated by TFIID and NuA4 (Table2). Even though ac b up-regulated genes down-regulated genes log2 (fold change) -log10 (q-value) 6 3.5 3 2.5 2 1.5 1 0.5 0420-2-4-6 scc1-73 up-regulated genes (569) scc1-73 wt distance to TSS (bp) 100 60 40 120 140 80 160 -500 0500 1000 1500 scc1-73 wt distance to TSS (bp) 100 60 40 120 140 80 160 -500 0500 1000 1500 control genes chrom. VII (720) scc1-73 wt distance to TSS (bp) 100 60 40 120 140 80 160 -500 0500 1000 1500 control genes chrom. XII (497) scc1-73 wt distance to TSS (bp) 100 60 40 120 140 80 160 -500 0500 1000 1500 scc1-73 down-regulated genes (445) 14 scc1-73 down-regulated genes (445) p=0.01 obs/exp: 1.5 up-nuc at promoters in scc1-73 (114) * * * * ** * * * * * * * ** * * * * * * * Enrichment of scc1-73-altered nucleosomes at scc1-73-down-regulated genes occupancy (reads) occupancy (reads) Fig. 2 Transcription downregulation in scc1‑73 cells is associated with chromatin changes. a Volcano plot of altered genes in scc1‑73 cells relative to wild‑type cells (1014 out of 6692 genes; ~ 15%). Changes in gene expression (log2) are plotted against q‑values (–log10). Genes with a q‑value < 0.01 are shown in green for downregulated (445) and in red for upregulated (569). b Overlap between downregulated genes, and genes with promoters that contain nucleosomes with higher occupancy (up‑nucleosomes), in scc1‑73 cells as compared to wild‑type cells. The number of genes in each case is indicated in parentheses. The probability of producing the given overlap based on random distributions was generated using a hypergeometric test. The rate between the observed and expected frequencies of common genes is also shown. c Nucleosome occupancy profiles of the scc1‑73 up‑ and downregulated genes, aligned relative to their TSS. Non‑affected genes from chromosomes VII and XII were used as control groups. The number of genes of each group is indicated in parentheses. Statistically significant differences in occupancy at the peak of nucleosomes − 2 to + 5 between the wild‑type cells and the scc1‑73 mutants are shown (paired two‑tailed Student’s t test, whereby one, two and three asterisks indicate p values of < 0.05, < 0.01 and < 0.001, respectively) Page 6 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 Table 2 Down- andupregulated genes inscc1-73 cells are associated withaspecific chromatin structure atpromoters andcohesin peaks, respectively The number of genes in each case is indicated in parentheses. The probability of producing the given overlapping if their distributions were random was generated using a hypergeometric test. The rate between the observed and expected frequencies of common genes is also shown. Significantly higher (underlined) and lower (italics) differences(p value < 0.001)are highlightedin bold. *Only downregulated genes are analyzed. See Additional file4: TableS4 for list of genes and references OPN promoter (544) DPN promoter (494) Closed promoter (1646) Open promoter (3586) FN promoter (1953) SN promoter (3066) RSC- binding promoter (239) Scc2- binding promoter (199) TATA- containing (1090) SAGA- dominated (577) TFIID- dominated (5130) Mediatordominated (315)* NuA4- dominated (500)* SWR1- dominated (462) Scc1- binding promoter (2417) scc1‑73 Up‑reg. (569) Overlap. 57 51 149 373 222 300 15 16 85 50 467 29 92 41 298 Obs/exp (p val) 1.2 (0.015) 1.2 (0.021) 1.06 (0.026) 1.22 (1.3e−10) 1.34 (3.5e−8) 1.15 (9e−5) 0.74 (0.045) 0.95 (0.1) 0.92 (0.03) 1.02 (0.06) 1.07 (2.1e−4) 1.08 (0.07) 2.25 (1e−14) 1.04 (0.06) 1.45 (4.9e−17) scc1‑73 Down‑reg. (445) Overlap. 93 27 153 246 212 202 36 15 166 103 307 53 25 48 136 Obs/exp (p val) 2.56 (8.8e−19) 0.83 (0.017) 1.4 (4.3e−7) 1 (0.03) 1.63 (7.7e−18) 0.99 (0.039) 2.26 (1.5e−6) 1.13 (0.095) 2.29 (4.2e−29) 2.68 (2e−22) 0.9 (2.7e−5) 2.53 (9.8e−11) 0.78 (0.02) 1.56 (5e−4) 0.85 (1.6e−3) Page 7 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 these associations may reflect different mechanisms of gene regulation, they connect the role of cohesin in gene expression with specific configurations of transcriptional regulators at promoters. The lack ofcohesin activity andhistone depletion leads tosimilar occupancy changes inasubset ofnucleosomes Our group has previously reported that the increased accessibility to MNaseI of the centromeric chromatin at CEN3 that can be observed in histone-depleted cells is partially suppressed by cohesin inactivation, suggesting that cohesins could contribute to the loss of chromatin integrity associated with defective histone deposition [47]. To further understand the function of cohesins in chromatin organization, we analyzed the genome-wide effects of the scc1-73 allele in cells that express histone H4 from the doxycycline (dox)-regulatable tet promoter (t::HHF2) [48]. For this, t::HHF2 and t::HHF2 scc1-73 cells were released together with scc1-73 and wild-type cells from G1 under conditions of histone depletion and restrictive temperature (Fig.1). Cells depleted of histone H4 during S phase displayed a severely altered nucleosome profile in metaphase (Fig. 1a), which was especially enriched in nucleosomes with reduced occupancy (Table1). Histone depletion caused a global increase in fuzziness and a wider distribution of both nucleosome occupancy and distance between adjacent nucleosomes (Fig. 1b). The overall loss of nucleosome positioning became particularly evident when nucleosome occupancy was aligned for all genes relative to the TSS, where the precise pattern of nucleosome positioning at both sides of the promoter-associated NFR was lost following histone depletion (Figs.1c, d). These results are in accordance with a previous study showing that defective histone supply after H3 depletion strongly affects chromatin integrity genome-wide [49]. The double-mutant t::HHF2 scc1-73 displayed a large number of chromatin perturbations (Fig.1a, d). This loss of chromatin integrity at t::HHF2 cells was confirmed at three different regions by indirect end labeling of MNaseI-treated cells (Additional file8: Fig. S1). Importantly, the lack of Scc1 activity increased the number of genome-wide altered nucleosomes in histone-depleted cells (Table1; compare t::HHF2 scc1-73 with t::HHF2), consistent with an additive effect of the lack of histones and cohesin activity. Thus, the partial suppression of nucleosome alterations at CEN3 described previously was specific for that locus [47]. We could not check these changes in our MNase-seq experiments because the centromeric nucleosomes were not detected (as previously reported in similar studies [49, 50]). A comparative analysis showed that both the profile of genomic regions with altered nucleosomes and the class of changes in nucleosome occupancy in these genomic regions coincided in many cases in scc1-73 and t::HHF2 (Additional file1: TableS1 and Additional file2: TableS2). This observation prompted us to study if nucleosomes with altered occupancy lie at the same genomic regions in both mutants. Statistical analyses of IGRs with altered nucleosomes showed a highly significant number of common elements (Fig.3a). Importantly, overlapping IGR shared nucleosomes with the same change—either increased or decreased—in occupancy (Fig.3b). Similar results were obtained from the analysis of ORF with altered nucleosomes (Fig.3c, d). A deeper analysis showed that this overlap was due to the fact that t::HHF2 and scc1-73 share a highly significant number of altered nucleosomes with the same change (Fig.3e, f), especially in nucleosomes with increased occupancy (8 to 10 times more than expected). Moreover, manual inspection of these nucleosomes showed that the change in the double-mutant t::HHF2 scc1-73 was similar to that displayed by the single mutants for 92% of nucleosomes (see representative examples, Fig. 3e, f). Altogether, these results suggest that part of the chromatin changes induced by defective nucleosome assembly and cohesin inhibition occurs through the same mechanism (~ 17% of scc1-73-altered nucleosomes). Mutant scc1-73 cells donotdisplay defects inthedeposition ofnewly synthesized histones duringDNA replication The epistatic effect of scc1-73 and t::HHF2 on the integrity of a subset of nucleosomes might reflect a role for cohesins in histone deposition. Indeed, our RNA-seq analysis showed that the scc1-73 mutant was specifically affected for expression of histones H4 and Htz1, as well as of histone chaperones Nap1 and Chz1 (Additional Fig. 3 A lack of cohesin activity and histone depletion lead to similar occupancy changes in a subset of nucleosomes. a, c Overlap between IGR (a) or ORF (c) with altered nucleosomes in t::HHF2 or scc1‑73 cells. b, d Overlap between IGR (b) or ORF (d) with either up‑ (increased occupancy) or down‑nucleosomes (decreased occupancy) in t::HHF2 and scc1‑73. e, f Overlap between up‑ (increased occupancy) or down‑nucleosomes (decreased occupancy) in IGR (e) and ORF (f) in t::HHF2 and scc1‑73 cells. Representative examples are shown; up‑ and down‑nucleosomes are marked in green and red, respectively. The number of genomic regions (a–d) or nucleosomes (e, f) in each case is indicated in parentheses. The probability of producing the given overlap if their distributions were random was generated using a hypergeometric test. The rate between the observed and expected frequencies is also shown (See figure on next page.) Page 8 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 a b c d e p=2e-10 17 33 IGR with down-nuc in t::HHF2 (700) IGR with down-nuc in scc1-73 (151) IGR with altered nuc in scc1-73 (286) p=4.1e-7 p=1.6e-16 37 108 IGR with up-nuc in scc1-73 (129) p=0.002 24 IGR with up-nuc in t::HHF2 (504) IGR with altered nuc in t::HHF2 (1180) IGR with altered nucleosomes in both t::HHF2 and scc1-73 Common altered nucleosomes in t::HHF2 and scc1-73 (at IGR) Common altered nucleosomes in t::HHF2 and scc1-73 (at ORF) IGR with similar and opposite nucleosome occupancy changes in t::HHF2 and scc1-73 ORF with similar and opposite nucleosome occupancy changes in t::HHF2 and scc1-73 ORF with altered nucleosomes in both t::HHF2 and scc1-73 ORF with altered nuc in scc1-73 (358) p=3.2e-14 201 ORF with altered nuc in t::HHF2 (2154) f p=0.03 p=2.3e-17 28 down-nuc in IGR in t::HHF2 (826) down-nuc in IGR in scc1-73 (154) p=1.6e-6 21 down-nuc in ORF in t::HHF2 (2164) down-nuc in ORF in scc1-73 (214) p=4.6e-3 17 up-nuc in IGR in scc1-73 (134) up-nuc in IGR in t::HHF2 (530) p=3e-19 27 up-nuc in ORF in scc1-73 (183) up-nuc in ORF in t::HHF2 (762) p=1.5e-21 30 63 ORF with down-nuc in t::HHF2 (1587) ORF with down-nuc in scc1-73 (206) p=9e-5 80 ORF with up-nuc in scc1-73 (157) p=0.05 48 ORF with up-nuc in t::HHF2 (647) p=0.03 YLR046C FRE8SPT7 tR(UCU)B scc1-73 wt t::HHF2 t::HHF2 scc1-73 SLX8 LGE1 LEE1SPR6 scc1-73 wt t::HHF2 t::HHF2 scc1-73 BUD5 MRPL8 scc1-73 wt t::HHF2 t::HHF2 scc1-73 RRP14 YCL068C scc1-73 wt t::HHF2 t::HHF2 scc1-73 obs/exp: 2.12 obs/exp: 1.49 obs/exp: 1.29obs/exp: 3.57 obs/exp: 1.41obs/exp: 1.11obs/exp: 2.31 obs/exp: 7.74 obs/exp: 3.23 obs/exp: 1.88 obs/exp: 9.87 obs/exp: 3.37 obs/exp: 1.75 obs/exp: 1.5 Page 9 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 file3: TableS3). To determine whether cohesins regulate histone deposition, we analyzed the incorporation into chromatin of newly synthesized histones at replicating DNA regions by following acetylated H3 at lysine K56 (H3K56ac) [51]. Although only a few nucleosomes were affected in scc1-73 cells, we speculated that recycling parental histones could mask a major defect in the deposition of newly synthesized histones. H3K56ac accumulation at replicating DNA regions from origin ARS305 was analyzed in cells synchronized in G1 and released into S phase in the presence of hydroxyurea, which reduces the pool of dNTPs and causes replication forks to be stalled in the proximity of the origin [52]. As expected, H3K56Ac accumulated around the origin (2.6kb from ARS305) but not at an unreplicated DNA region (18Kb from ARS305). Importantly, the absence of cohesin activity in scc1-73 cells did not affect H3K56ac incorporation (Fig.4), suggesting that cohesins do not have a major role in depositing new histones during DNA replication. Histone depletion hasmodest effects ontheaccumulation anddistribution ofcohesin peaks To further understand the interplay between cohesins and chromatin structure we studied the importance of chromatin integrity in cohesin binding and distribution. For this, cells were synchronized in G1 and released into fresh medium until G2/M under conditions of histone depletion at 30°C, and the association of Scc1 was followed by a high-density ChIP-on-chip analysis (5-bp resolution). This analysis showed an overall similarity of cohesin peaks in t::HHF2 and wild-type cells (Fig.5a, b and Additional file9: Fig. S2a), with cohesins preferentially bound to the pericentromeric chromatin, rDNA, tRNA genes and the IGR of convergently transcribed genes (Fig.5a–c) [11–13]. However, the amount of cohesins at rDNA, tRNA genes and telomeres relative to the genome average was higher in histone-depleted cells than in wild-type cells (Fig.5d and Additional file5: TableS5). As the ChIP-on-chip analysis was not normalized with an internal control, no comparison between absolute amounts of signal between wild-type and mutant could scc1-73wild type G1 time (min): 30 45 G1 30 45 0 20 30 10 40 H3K56ac enrichment ARS305 + 2.6 kb ARS305 + 18 kb Fig. 4 Cohesins are not involved in the deposition of newly synthesized histones. ChIP analysis of the incorporation of newly synthesized histone H3 (acetylated in lysine 56) in wild‑type and scc1‑73 cells that have been synchronized in G1 and released into S phase in the presence of 200 mM HU for 45 min. H3K56c enrichment both at the proximity of ARS305 and at an unreplicated region (2.6 and 18 kb from the origin, respectively) was calculated as the amount of DNA immunoprecipitated with an antibody against H3K56ac relative to that obtained with antibody against total histone H3. The average and range from two independent experiments are shown. IgG‑treated cells were used as an internal control to confirm the specific enrichment at each region Fig. 5 Histone depletion has a moderate effect on cohesin binding and distribution. a Cohesin distribution at chromosome I in wild‑type and t::HHF2 cells synchronized in G1 and released into fresh medium until G2‑metaphase, as determined by ChIP‑on‑chip analysis against Scc1‑HA. b Cohesin distribution at the ribosomal DNA locus in wild‑type and t::HHF2 cells. c Number of IGRs classified according to the orientation of the flanking genes that overlap with cohesins (by at least 1 bp) in wild‑type and t::HHF2 cells. d Relative amount of cohesins in t::HHF2 cells relative to wild‑type cells at the indicated genomic regions. The total amount of cohesins at each region was calculated considering the sum of positive signals (relative to the untagged strain) with a p < 0.05. The ratio between the mutant and the wild‑type at each region was normalized to that obtained for the whole genome, which was taken as 1. The proportion of cohesins between mutant and wild‑type cells at each genomic region relative to the genome average was statistically different according to a two‑tailed Chi‑square test (p < 0.001). e Cohesin enrichment in t::HHF2 cells relative to wild‑type cells at different genomic regions, as determined by ChIP and qPCR analyses against HA‑Scc1 in cells grown as in a. IGR and tRNA genes are indicated in Methods section. Cohesin enrichment was calculated as the ratio between immunoprecipitated DNA and input in t::HHF2 cells relative to the same value in the wild‑type cells. The average and SEM from 3 to 4 independent experiments are shown. An untagged strain was used as an internal control to confirm the specific enrichment at each region. The amount of Scc1 relative to Pgk1 is shown on the left. No significant differences were observed between wild‑type and mutant cells from four independent measurements. f Number of IGRs, ORF, tRNA, ARS and telomeres that overlap with cohesins (by at least 1 bp) in wild‑type and t::HHF2 cells. g Probability that scc1‑73 alters cohesin‑associated nucleosomes if scc1‑73‑altered nucleosomes were randomly distributed, as determined by a hypergeometric test. The rate between the observed and expected frequencies of common nucleosomes is also shown. The number of nucleosomes in each case is indicated in parentheses. h Comparison of the percentages of altered nucleosomes by histone depletion (showing occupancy, fuzziness and position shift) in the regions with cohesins (9596 nucleosomes) relative to the whole genome (66,278 nucleosomes) in t::HHF2 cells. A hypergeometric test was used to determine the probability of obtaining the indicated percentages of altered nucleosomes at regions with cohesin if their distributions were random (See figure on next page.) Page 16 of 18 Maya‑Milesetal. Epigenetics & Chromatin (2019) 12:47 Comparative analyses ofgenomic elements Analyses of the genome distribution and overlapping of dynamic nucleosomes, cohesin peaks and gene elements were performed with the tools of the Galaxy web platform (usegalaxy.org) [76]. Other statistical analyses were performed with the GraphPad Prism software. Additional files Additional file1: TableS1. Comparison of the number and percentage of nucleosomes altered after histone depletion (in t::HHF2 cells) or cohesin inactivation (in scc1‑73 cells) in the indicated genomic regions relative to the whole genome. A hypergeometric test was used to determine the probability to obtain the indicated percentages of altered nucleosomes at the indicated genomic regions if their distribution along the genome were random. Significantly higher (green) and lower (red) percentages are highlighted. *rDNA is analyzed as a single copy. Additional file2: TableS2. Comparison of the number and percent‑ age of nucleosomes altered after histone depletion (in t::HHF2 cells) or cohesin inactivation (in scc1‑73 cells) in the indicated genomic regions relative to the whole genome. A hypergeometric test was used to deter‑ mine the probability of obtaining the indicated percentages of altered nucleosomes at the indicated genomic regions if their distribution along the genome were random. Significantly higher (green) and lower (red) percentages are highlighted. *rDNA, analyzed as a single copy. Additional file3: TableS3. Misregulated genes in ssc1‑73. Lists of up‑ and downregulated genes are shown. Additional file4: TableS4 List of misregulated genes from additional studies used for comparative analyses in Table 2. Additional file5: TableS5. Cohesin distribution in wild‑type and histone‑ depleted cells at the indicated genomic regions. A peak of Scc1 is defined as a DNA fragment with continuous Scc1 signals that are both positive (relative to the untagged strain) and with a p < 0.05. A peak signal was calculated as the sum of these positive signals. Genomic regions with Scc1 and peaks of Scc1 at a particular genomic region are defined by at least 1 bp overlapping. Additional file6: TableS6. The effect of cohesin inactivation on chroma‑ tin structure is more severe in cohesin‑associated regions. The probability that scc1‑73 alters nucleosomes in Scc1‑binding IGR or Scc1‑binding ORF if the altered genomic regions in scc1‑73 cells were randomly distributed was determined by a hypergeometric test. The rate between the observed and expected frequencies of common genomic regions is also shown. The number of regions in each case is indicated in parenthesis. Additional file7: TableS7. Saccharomyces cerevisiae strains (a) and oligos (b) used in this study. Additional file8: Fig. S1. Effect of cohesin inactivation in scc1‑73 cells on the chromatin structure of three different loci of histone‑depleted cells. Nucleosome positioning analyses are shown for the indicated loci after MNase I digestion and indirect‑end labeling of the indicated strains synchronized in G1 and released into fresh medium for 1 h at 37 °C until G2/M. Additional file9: Fig. S2. Effect of histone depletion on cohesin binding and distribution. a Cohesin distribution at different regions of chromo‑ somes II and V in wild‑type and t::HHF2 cells that have been synchronized in G1 and released into fresh medium until G2/M, as determined by ChIP‑ on‑chip analysis against HA‑Scc1. b Probability that IGR that had either lost or gained cohesins after histone depletion would overlap with IGR with altered nucleosomes in t::HHF2 cells or in both t::HHF2 and scc1‑73 cells if they were randomly distributed, as determined by a hypergeo‑ metric test. The rate between the observed and expected frequencies of common IGR is also shown. The number of IGR in each case is indicated in parenthesis. nd, not determined. Additional file10: Fig. S3. Preparation and analysis of the nucleosomal DNA used for MNaseI‑seq. a Generation of mononucleosomes in the indicated strains after partial digestions with MNase I. The DNA purified for DNA‑seq is marked in red. b, c Electrophoretic (b) and electropherogram (c) analyses of the purified nucleosomal DNA. Abbreviations ARS: autonomous replication sequence; DANPOS: dynamic analysis of nucleo‑ some position and occupancy by sequencing; DPN: depleted‑proximal pro‑ moter; FN: fragile nucleosome; IGR: intergenic region; NFR: nucleosome‑free region; OPN: occupied‑proximal promoter; ORF: open reading frame; rDNA: ribosomal DNA; SN: stable nucleosome; tDNA: transfer DNA; TSS: transcription start site. Acknowledgements We thank José Carlos Reyes for critically reading the manuscript. Authors’ contributions DM, MM‑P, MB‑M, MJC‑L, EG‑M and MM‑H performed the experiments. DM, EA, MP‑A, MM‑P, MB‑M and FP analyzed the results. FP conceived the project and wrote the manuscript. All authors read and approved the final manuscript. Funding This work was supported by the Spanish Ministry of Economy and Competi‑ tiveness (BFU2012‑38171 and BFU2015‑63698‑P) and the Andalusian Govern‑ ment (P12‑CTS‑2270). Availability of data and materials Genome‑wide data have been deposited at the MIAME‑compliant Gene Expression Omnibus (GEO). The remaining data and material are available from the corresponding author on request. Ethics approval and consent to participate Not applicable. Consent for publication Not applicable. Competing interests The authors declare that they have no competing interests. Author details 1 Department of Genome Biology, Andalusian Molecular Biology and Regen‑ erative Medicine (CABIMER), CSIC‑University of Seville‑University Pablo de Ola‑ vide, Seville, Spain. 2 Genomic Unit, Andalusian Molecular Biology and Regen‑ erative Medicine Center (CABIMER), CSIC‑University of Seville‑University Pablo de Olavide, Seville, Spain. 3 Present Address: Department of Biochemistry, University of Oxford, Oxford, UK. 4 Present Address: Division of Cancer Sciences, Manchester Cancer Research Center, University of Manchester, Manchester, UK. Received: 22 March 2019 Accepted: 10 July 2019 References 1. Prado F, Maya D. Regulation of replication fork advance and stability by nucleosome assembly. Genes (Basel). 2017;8:49. 2. Polo SE, Almouzni G. Chromatin dynamics after DNA damage: the legacy of the access‑repair‑restore model. DNA Repair. 2015;36:114–21. 3. Venkatesh S, Workman JL. Histone exchange, chromatin structure and the regulation of transcription. 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