Abstract
It is important to accurately regulate the expression of genes involved in development and environmental response. In the fission yeast Schizosaccharomyces pombe, meiotic genes are tightly repressed during vegetative growth. Despite being embedded in heterochromatin these genes are transcribed and believed to be repressed primarily at the level of RNA. However, the mechanism of facultative heterochromatin formation and the interplay with transcription regulation is not understood. We show genome-wide that HDAC-dependent histone deacetylation is a major determinant in transcriptional silencing of facultative heterochromatin domains. Indeed, mutation of class I/II HDACs leads to increased transcription of meiotic genes and accumulation of their mRNAs. Mechanistic dissection of the pho1 gene where, in response to phosphate, transient facultative heterochromatin is established by overlapping lncRNA transcription shows that the Clr3 HDAC contributes to silencing independently of SHREC, but in an lncRNA-dependent manner. We propose that HDACs promote facultative heterochromatin by establishing alternative transcriptional silencing.
Introduction
Heterochromatin is critical to eukaryotic cells and exists in two functionally distinct forms: constitutive and facultative heterochromatin. Constitutive heterochromatin is found mainly at gene-poor regions such as centromeres and telomeres, and is important for chromosome segregation during cell division. In contrast, facultative heterochromatin is found in gene-dense regions of chromosomes and controls silencing of developmentally and environmentally regulated genes (including meiotic genes1–4 and genes regulated by phosphate availability, such as pho15). The mechanisms underlying the formation and maintenance of stable constitutive heterochromatin have been extensively studied in the fission yeast Schizosaccharomyces pombe (S. pombe), which has proven a powerful model for understanding heterochromatic silencing. In contrast, the mechanisms mediating silencing of reversible facultative heterochromatin are not well understood.
Chromatin modifying complexes play a central role in heterochromatin formation. These include the Clr4/Suv39h-methyltransferase complex (CLRC) that methylates Lys9 of histone H3 (H3K9me), as well as the class I (Clr6 and Hos2), class II (Clr3) and class III (Sir2, NAD+-dependent, ‘sirtuins’) histone deacetylase complexes (HDACs). Hypoacetylation of histones often depends on HDAC recruitment via low level non-coding (nc) transcription occurring within these regions. Heterochromatic ncRNAs are converted into dsRNAs through the action of the RNA-directed RNA polymerase complex (RDRC) and subsequently cleaved by the ribonuclease III Dicer (Dcr1) to produce small interfering RNAs (siRNAs) of 21-23 nucleotides6–8. These siRNAs are converted into single-stranded siRNAs and, together with Argonaut proteins, form the RNA-induced transcriptional silencing (RITS) complex. siRNAs direct the other RITS components, as well as Clr4, to heterochromatin by complementary base pairing with nascent transcripts6,9,10. The Clr4 H3K9me mark, in turn, acts as a landing platform for the HP1-like proteins Swi6 and Chp27. These proteins target class I and II HDAC complexes, leading to further compaction of heterochromatin and transcriptional gene silencing11. The class I HDAC Clr6 removes acetyl groups from H3K14 and K9, and H4K5, K8, K12 and K1612. The class II HDAC Clr3 is part of SHREC (Snf2/Hdac [histone deacetylase]-containing Repressor Complex) that deacetylates H3K14. In addition to Clr3, SHREC is composed of four other factors – Clr1, Clr2 and the ATPase and chromatinremodelling subunit Mit111. SHREC has been shown to be recruited to pericentromeric heterochromatin by the conserved protein Seb113, which harbours RNA- and Pol II C-terminal-domain (CTD) binding modules (RNA-Recognition-Motif, RRM and CTD-Interacting-Domain, CID).
Unlike at constitutive heterochromatin, H3K9me at facultative heterochromatin does not induce transcriptional silencing. Instead, it has been proposed that meiotic transcripts are primarily silenced post-transcriptionally, with the RNA-binding proteins Mmi1 (which recognises DSR (Determinant of Selective Removal) sequences on RNA14) and Zinc-finger protein Red1 recruiting the RNA degradation machinery. Mmi1 and Red1 do also recruit RNAi and establish H3K9me but transcriptional silencing does not appear to be induced as a consequence. Despite all this, we previously demonstrated that the acid phosphatase-encoding gene pho1, on which we found facultative heterochromatin, is repressed at the transcriptional level by the DSR-containing lncRNA prt (pho1-repressing transcript). prt is produced in the presence of phosphate from a promoter located upstream of pho15,15–17. Under such conditions, expression of prt leads to transcriptional repression of pho1. Co-transcriptional recruitment of Mmi1 to prt initiates the deposition of H3K9me by Clr4, formation of transient facultative heterochromatin across the gene and degradation of prt by the exosome complex. In the absence of non-coding transcription (either when the non-coding promoter is deleted or when cells are shifted to media without phosphate) pho1 transcription is fully activated. Surprisingly though, in contrast to the complete loss of silencing observed upon deletion of the prt non-coding promoter, deletion of Clr4 had a minor effect on pho1 expression and instead affected only induction kinetics5. This led us to speculate that there could be another, Clr4-independent mechanism of repression that is also mediated by prt non-coding transcription.
In Saccharomyces cerevisiae (S. cerevisiae) it has been shown that meiotic genes are repressed at the level of transcription by HDAC activity18,19. Repression of euchromatic genes by non-coding transcription has also recently been shown to require the class I HDAC Rpd3S (Rpd3 Small complex) and Set3/Rpd3L (Rpd3 Large complex)20,21. HDAC activity within these complexes is tightly linked to methylation of either H3K36 or H3K4 by, respectively, Set2 and Set1 methyltransferases. Through interacting with serine 5 phosphorylated CTD Set1 methylates H3K4 at gene 5’ ends22. It establishes a zone of H3K4me3 near transcription start sites followed by a zone of H3K4me2 downstream23,24. Set2 on the other hand establishes H3K36 methylation later during transcription and interacts with CTD phosphorylated at serines 2 and 525–27. H3K36me can negatively affect transcription by targeting histone deacetylation by Rpd3S28–30. Both the Set2-Rpd3S and Set1-Rpd3L pathways have been shown to be important for modulating transcriptional dynamics during certain environmental changes in S. cerevisiae. Interestingly, while steady-state mRNA levels are not affected, gene induction occurs more rapidly upon carbon source shift in the absence of Set2. This is reminiscent of the situation in S. pombe where we also see faster pho1 induction in a Clr4 mutant. Here, homologous HDAC activities are provided by Clr6 and Hos2 complexes but, unlike budding yeast Rpd3, Clr6 is essential for cell viability31. A possible reason for these differences might be due to the fact that Clr6 also functions in transcriptional silencing at constitutive heterochromatin together with Clr3 and the class III HDAC Sir2.
In this study we set out to investigate whether HDACs contribute to transcriptional silencing at regions of facultative heterochromatin. Using the pho1 locus as a model of facultative heterochromatin, we demonstrate that transcription is repressed upon inhibition of class I/II HDACS using trichostatin A (TSA). This seems to be due to the action of Clr3 as its deletion has the same effect on pho1 levels. Strikingly, Clr3-dependent repression of pho1 is mediated via the lncRNA prt, which is needed for Clr3 recruitment to the loci. Interestingly, loss of Set1 and Set2 methyltransferases also leads to loss of transcriptional silencing, with the Set1 deletion having the most striking effect, suggesting that the role of Set1 in facilitating recruitment of HDACs via non-coding transcription is conserved in budding and fission yeast. In contrast to constitutive heterochromatin13, Clr3 is likely to mediate pho1 silencing by a mechanism that acts in addition to Clr4. Strikingly, class I/II HDAC inhibition results in loss of transcriptional silencing of meiotic genes as analysed by NET-Seq and RNA-Seq. We conclude that HDACs have an important function in transcriptional silencing at facultative heterochromatin.
Results
The class II HDAC Clr3 contributes to pho1 silencing
We previously reported that transcriptional repression of the acid phosphatase pho1 in response to extracellular inorganic phosphate relies on the transcription of the overlapping ncRNA prt5. We showed that the H3K9 methyltransferase Clr4 contributes to this effect but, since its deletion does not lead to complete loss of silencing, we speculated that other players must be involved. In addition to H3K9me, hypoacetylation of histones by HDACs has been linked to transcriptional repression. To then investigate whether HDACs are involved in repression at pho1, we treated cells with a class I and II HDAC inhibitor, trichostatin A (TSA). Strikingly, a substantial increase in pho1 mRNA is observed by Northern blot when cells are transiently treated with TSA (Figure 1a). At the same time, no change in the expression level of the house-keeping gene adh1 is observed upon TSA treatment.
In order to identify which of the four S. pombe HDACs is responsible for the repression of pho1, we analysed RNA levels from the following HDAC mutant strains: clr3Δ, clr6-1, hos2Δ and sir2Δ (Figure 1b). Of the TSA-sensitive HDACs, Clr3 was the only one that had an effect on pho1 expression, with its deletion resulting in increased pho1 mRNA levels (Figure 1b). In contrast, deletion of TSA-insensitive sir2, which contributes to transcriptional silencing at constitutive heterochromatin, had no effect on pho1 expression (Figure 1b, lane 5). Furthermore, no additional increase was observed when clr3Δ cells were treated with TSA suggesting that the increased pho1 levels observed upon TSA treatment are solely due to Clr3 inhibition (Figure 3e, lanes 5 and 6). Previously, we showed that pho1 levels are altered in RNA decay mutants (rrp6Δ and mmi1Δ) due to increased expression of prt5. In contrast, clr3Δ does not affect levels of prt (Figure 1b), suggesting a different mechanism of action here.
To further understand the role of Clr3 in repression of pho1, chromatin immunoprecipitation (ChIP) followed by qPCR was performed to determine whether Clr3 is recruited to the locus. This revealed that Clr3 indeed localizes to the gene, particularly at the non-coding region (Figure 1c). A recent study proposed a mechanism of Clr3 recruitment via the DNA and RNA binding activity of the non-catalytic SHREC subunit Clr232. In order to determine whether Clr3 is recruited and functions at pho1 as part of SHREC or independently of the complex, the effects of clr1, clr2, and mit1 deletion on RNA levels were analysed by Northern blot (Figure Supplement 1a). Compared to wild-type pho1 mRNA levels, a slight accumulation was detected for all strains. However, this is less pronounced than in clr3Δ, indicating that Clr3 is unlikely to entirely depend on Clr2 or other components of SHREC for recruitment to pho1. This is consistent with previous reports showing that Clr3 can also localize to several sites independently of other SHREC components11.
Lack of Clr3 coincides with increased H3K14 acetylation and transcription at pho1
Having established that Clr3 is recruited to the non-coding prt region, we wanted to investigate how it exerts its silencing effect on pho1. Previous studies at the mat locus have proposed that Clr3 can restructure the chromatin environment in such a way as to restrict Pol II access33. In order to address if this is the case at the pho1 region as well, the occupancy of Pol II at the locus was determined by ChIP-qPCR (Figure 1d). We found that loss of Clr3 leads to increased Pol II levels upstream of the pho1 promoter and particularly across the gene body. This is consistent with increased transcription of pho1 mRNA.
Since Clr3 possesses histone deacetylase activity, we expected changes in H3K14 acetylation (H3K14ac) levels at the pho1 locus in clr3Δ. Indeed, an increase in the levels of H3K14ac could be detected by ChIP-qPCR (Figure 1e); similar to what has been shown previously at pericentromeric heterochromatin11. Concomitantly, the biggest increase in H3K14ac occurred over the non-coding region to which Clr3 is recruited. Taken together these data suggest that Clr3 functions in silencing the pho1 mRNA by a mechanism that depends on its HDAC activity.
Clr3 and Clr4 function independently to silence pho1
It has previously been suggested that Clr3, as part of SHREC, promotes H3K9me at pericentromeric heterochromatin and acts in the same pathway as the Clr4 methyltransferase13. Consequently, we next tested whether this is also the case at the pho1 locus. In order to address this, we generated the double mutant clr3Δclr4Δ. Remarkably, this mutant showed an additive accumulation of pho1 mRNA levels (Figure 2a) and displayed a slow growth phenotype considerably more severe than either of the single mutants, clr3Δ or clr4Δ (Figure 2b). These data support the idea that Clr3 and Clr4 can act independently of each other to promote gene silencing at facultative heterochromatin, in contrast to the mechanism proposed at constitutive heterochromatin.
Non-coding transcription is required for Clr3 recruitment
To further our understanding of how Clr3 drives transcriptional silencing, we investigated whether non-coding transcription is needed for its function. We found that, in the absence of non-coding transcription in a strain lacking the prt promoter (ncproΔ)5, Clr3 recruitment to pho1 was reduced (Figure Supplement 2a). H3K14ac levels were also increased in this strain (Figure Supplement 2b), similar to that seen in clr3Δ (Figure 1e). Moreover, we reasoned that if Clr3 recruitment can only occur in the presence of non-coding transcription, a clr3ΔncproΔ double mutant should not have an additive effect on pho1 expression compared to the respective single mutants. To test this, we analysed pho1 levels in these strains and, as expected, Northern blot analysis revealed no obvious additive effect compared to the ncproΔ single mutant (Figure Supplement 2c). This indicates that Clr3 is likely to operate downstream of prt transcription.
To test which specific region(s) of the prt ncRNA other than its Mmi1 binding site are involved in pho1 silencing, a series of five different 189 bp deletions within the 5’ proximal region of prt were generated (prt-1Δ (-1197- to -1008), prt-2Δ (-1008- to -819), prt-3Δ (-819 to -630), prt-4Δ (-441 to -252), and prt-5Δ (-252 to -63) (pho1 ATG=1)) (Figure 3a). We were unable to generate a deletion of the region -630 to -441. Interestingly, elevated pho1 levels were observed in prt-1Δ, prt-2Δ, and prt-3Δ (Figure 3b, lanes 1-4) suggesting that element(s) responsible for pho1 silencing are located within the 5’ part of prt. No change in pho1 expression could be detected for either prt-4Δ or prt-5Δ (Figure 3b, lanes 5 and 6). In the case of prt-3Δ, this phenotype is likely a result of lost Mmi1 recruitment since this mutant lacks the DSR motifs we previously mapped (5,34, Figure 3a, Mmi1 CRAC). We tested Mmi1 recruitment to the prt locus in this mutant and it is indeed defective (data not shown). We next wanted to examine whether inhibition of Clr3 HDAC activity by TSA has any additive effect on pho1 levels in the prt mutants. In prt mutants prt-1Δ, prt-2Δ, prt-4Δ, and prt-5Δ, pho1 levels were identical to the wild-type upon TSA treatment (Figure 3b, lanes 7, 8, 9, 11, 12). In contrast, pho1 levels in prt-3Δ (which lacks DSRs) were noticeably increased in TSA-treated compared to untreated cells (Figure 3b, compare lanes 4 and 10) or any other TSA treated samples. Taken together, these data are in agreement with a model in which Clr3 recruitment is independent of H3K9me and the Mmi1/Clr4/H3K9me pathway acts in parallel to Clr3/H3K14ac.
To understand further how transcription of prt mediates recruitment of Clr3 we wanted to test whether other histone methylation marks are involved in pho1 repression. Interestingly, deletion of the H3K4 methyltransferase Set1 leads to derepression of pho1 (Figure 3c). In contrast, deletion of the H3K36 methyltransferase Set2 only had a minor effect on pho1 derepression, suggesting that the mechanism underlying pho1 silencing primarily relies on histone methylation by Set1. In S. cerevisiae, Set1 mediates recruitment of the Rpd3L complex for repression of metabolic genes. However, we show that loss of the Rpd3 homologues Clr6 or Hos2 does not have an effect on pho1 levels (Figure 1b). Instead, our data suggest that Set1 is likely to function via a different HDAC (Clr3) in transcriptional repression in fission yeast.
We next examined whether co-transcriptional recruitment of Clr3 is RNA dependent. To this end we performed ChIP experiments on chromatin extracts that were treated with RNases. As a control, we tested recruitment of the known prt-binding protein Mmi1 and, as expected, binding was lost upon RNase treatment (Figure Supplement 3a). We found the Clr3 signal at the pho1 locus to be reduced suggesting that its recruitment is indeed dependent on RNA (Figure Supplement 3b). At the same time, RNase treatment did not affect the Pol II profile over pho1 (Figure Supplement 3c). Interestingly, Set1 has recently been demonstrated to interact with nascent RNA within promoter–proximal regions via its RRM domains35–37. These studies proposed that RNA binding might be important for Set1’s function in transcriptional repression. It is possible that both Set1 and Clr3 interact with RNA. Alternatively, the observed loss of Clr3 from chromatin upon RNase treatment might be connected to loss of the methyltransferase, which is consistent with the model where HDAC targeting depends on Set1.
Given that budding yeast Set1 has no RNA sequence specificity, it is not entirely clear to us why deletion of the promoter-proximal regions of prt but not the distal regions leads to loss of pho1 silencing (Figure 3b). Budding yeast Set1 was proposed to rely on Pol II phosphorylation (specifically on serine 5 within the CTD) for promoter-proximal targeting38. It is of course possible that the yeast proteins differ, and that S. pombe Set1 in fact has some preference either for nascent RNA sequence or secondary structure. Otherwise, specificity might be provided by another RNA or DNA-binding protein. Previous studies have shown that Clr3 recruitment can be mediated via the transcription factor Atf133,39,40. However, the absence of the well characterized Atf1 binding site ATGACGT in prt suggests that Atf1 is unlikely to be involved in Clr3 recruitment to this locus. Indeed, no increase in pho1 levels was observed in an atf1Δ strain (Figure 3d, lane 5). Altogether, these data suggest that HDAC-mediated silencing is dependent on Set1 and non-coding transcription and acts in addition to Mmi1/Clr4/H3K9me.
HDACs act independently of RNA processing and degradation on pho1
To determine at which stage of RNA synthesis Clr3 mediates transcriptional silencing, we asked whether there is any genetic interaction between histone deacetylation and Seb1-dependent read-through or Mmi-dependent RNA degradation. We previously demonstrated that Seb1 is a key factor that regulates 3’ end cleavage and transcription termination of Pol II transcribed genes41. Accordingly, read-through transcription is observed in seb1 mutants from the pho84 gene located upstream of prt (41, Figure 3a), potentially interfering with its expression and leading to activation of pho1. Furthermore, consistent with Clr3 acting independently at this locus, an additive effect on pho1 accumulation is observed when Clr3 HDAC activity is inhibited by TSA in strains either carrying mutation in seb1 (seb1-1) or deletion of the Seb1 binding site (prt-bsΔ) (Figure 3e, compare lanes 3 to 4 and 9 to 10). Similarly, mmi1Δ shows an additive effect with TSA (Figure 3e, compare lane 7 to 8), which is also consistent with that seen when prt-3Δ, the strain harbouring a deletion spanning DSR elements (Figure 3a), is treated with TSA (Figure 3b, compare lanes 4 to 10). From these data we conclude that Clr3 is likely to act in addition to Seb1 and Mmi1 to repress pho1.
Phosphate-response genes are regulated differently
To explore whether other genes involved in phosphate metabolism are regulated by HDACs, we studied the expression of a gene encoding transporter for glycerophosphodiester 1 (tgp1). Similar to pho1, tgp1 is repressed in response to inorganic phosphate, via non-coding transcription16, and induced upon growth in the absence of phosphate (Figure Supplement 4a, lanes 9-11). The nc-tgp1 RNA is highly unstable and accumulates upon deletion of the exosome subunit Rrp6 (16, Figure Supplement 4a, lane 12) or the exosome specificity factor Mmi116,34. We found accumulation of the tgp1 transcript in the seb1-1 mutant compared to wild-type (Figure Supplement 4a, compare lanes 1 and 2). In addition, we previously placed the seb1 gene under the control of the thiamine regulated nmt promoter, where seb1 expression is repressed upon switch from medium lacking to medium containing thiamine. Coincident with loss of Seb1, tgp1 mRNA levels start to accrue (Figure Supplement 4a, lane 13). Seb1’s involvement in regulation of tgp1 likely relates to failed termination of nc-tgp1 in the two mutants, similar to the demonstrated role of its homologue in S. cerevisiae42,43. Unlike the case at pho1, we found that deletion of clr3 has no discernible induction effect on tgp1. No detectable increase in either tgp1 or nc-tgp1 was observed in sir2Δ and two strains harbouring a clr6 mutation (Figure Supplement 4a, lanes 6, 7 and 8) or upon treatment with TSA (Figure Supplement 4b). Thus, it appears that HDACs, or at least those sensitive to TSA, are not implicated in the repression of tgp1 expression. Consistent with a clr3 deletion strain having no effect on tgp1 mRNA levels, recruitment of the protein, or indeed any other SHREC subunit, could not be detected at this locus (data not shown). Similarly, clr4Δ, or the double mutant clr3Δclr4Δ (Figure Supplement 4a, lane 4 and 5), revealed no change in mRNA expression levels. These results indicate that unlike pho1, tgp1 repression is not reliant on Clr3, other class I/II HDACs, nor Clr4's methyltransferase activity.
Inhibition of HDACs induces dramatic changes in global Pol II transcription
We next asked how global expression of genes is regulated by the action of HDACs. To examine the contribution of class I and II HDACs on gene expression, we performed RNA-seq to systematically analyse the consequences of HDAC inhibition upon treatment with TSA. We used S. cerevisiae for spike-in to normalize for changes in overall RNA levels and compared cells before and after addition of TSA44–46. In contrast to our expectation considering the result at pho1, analysis of the data revealed that inhibition of HDAC activity results in more transcripts being decreased by at least 2-fold (3525) than increased (289) (Figure 4a and Figure Supplement 5a). Northern blot analysis confirmed decreased levels of SPAC2E1P3.05c and ctr4 transcripts (Figure 4b) upon TSA treatment. A possible explanation for such a large amount of transcripts being decreased may relate to the role HDACs have been shown to have in suppressing overlapping antisense transcription in both fission and budding yeast31,47. Indeed, one study demonstrated that 74% of 23 transcripts positively regulated by the Rpd3L show overlapping antisense non-coding transcription47. However, our data did not show a strong correlation with antisense transcription for genes down-regulated upon inhibition of HDACs (data not shown). This suggests that suppression of antisense transcription via HDACs may not be a major mechanism to positively control transcription in fission yeast.
In order to test whether the observed changes in transcript level are due to changed transcription or post-transcriptional RNA stability, as is the case in mammals48, we performed calibrated NET-seq. This technique allows for the genome-wide assessment of Pol II distribution at single-nucleotide resolution and in a strand specific manner49. We saw the same general trend of a greater number of genes being down-regulated than up-regulated (Figure Supplement 5a). Remarkably, about 80% of genes that are more than 2-fold up- or down-regulated show the same response to TSA in NET-Seq as observed by RNA-seq (Figure 4a). This suggests that the majority of changes in gene expression induced by HDAC inhibition occur at the transcriptional level.
Increased histone acetylation correlates with transcription activation upon TSA treatment
To determine the direct effect of TSA treatment on gene expression and histone acetylation levels, we performed ChIP-seq experiments to measure histone H3K14ac and histone H3 levels genome-wide. While H3K14ac levels show large differences between TSA treated and untreated samples, the overall levels of H3 do not seem to be affected (Figure Supplement 5b). Consistent with previous results50 we identified a peak of H3K14ac at gene 5’ ends (150 nt before to 150 nt after gene TSS) (Figure Supplement 5c). Increased H3K14ac in this region correlates with up-regulation in RNA levels, which is in good agreement with a positive role for acetylation on gene transcription and a direct role for HDACs in repression of these genes (Figure 4c). Majority of the genome shows an increase in acetylation (Figure Supplement 5d). As expected this is true at centromeres and, as a consequence, we see higher RNA levels (Figure 4d). Consistently, down-regulated transcripts show lower H3K14ac levels compared to genes whose expression is not significantly affected by TSA treatment (Figure 4c). However, this effect is less important than the observed increase of H3K14ac for two-fold up-regulated genes, suggesting that lack of H3K14ac alone does not play a main role in transcription repression. It is possible that deacetylation at other lysines by Clr6 could be primarily responsible for a repressive effect on gene expression. Indeed, a strong decrease in the levels of the transcripts SPAC2E1P3.05c and ctr4 can be seen upon mutation of Clr6 (Figure 4b). While deletion of Clr3 results in down-regulation of SPAC2E1P3.05c RNA this is not the case for ctr4 suggesting that the observed effect upon TSA treatment is due to a different HDAC than Clr3. However, this does not seem to be Hos2 as no decrease in RNA levels is observed in a mutant of this HDAC.
Transcriptional silencing of meiotic genes requires HDAC activity
Interestingly GO enrichment analysis of the genes up-regulated in TSA and with increased H3K14ac, revealed meiotic transcripts (Supplementary Table 1 and 2 and Figure 5a, b and c), and multiple membrane proteins including efr3, pdh1, SPCC1235.18/17, SPAC977.06, ght5, ght6 and ght8 (Supplementary Table 1 and 2 and Figure Supplement 6 and 7). Regions encoding for repetitive selfish elements also appear to depend on HDACs for transcriptional silencing. An interesting example is a group of so called selfish wtf genes (Figure Supplement 7). These proliferate due to meiotic drive, killing gametes that do not carry the genes and causing infertility51. Similar to pho1 (Figure 5d), meiotic genes depend on Clr3 for transcriptional silencing. In the case of meu31 some redundancy can be seen with Clr6 and Hos2 also contributing to the repression of transcription (Figure 5c). In agreement with a previously demonstrated role for the nuclear exosome complex in the degradation of meiotic transcripts during mitosis, we observe increased levels of meu19 and meu31 in rrp6Δ (Figure 5b and c). However, mug14 is not affected in this exosome mutant, suggesting that the gene is likely to be regulated only at the transcriptional level (Figure 5a). In support of repression stemming from both the transcriptional and RNA level, serial dilutions onto TSA plates reveal a strong synthetic growth defect for rrp6Δ (Figure 5e). In summary, our data suggest that, in addition to post-transcriptional regulation, regions of facultative heterochromatin around prt/pho1 and meiotic genes are repressed at the transcriptional level via the action of HDACs.
pho1, an archetype of lncRNA/HDAC-mediated gene regulation?
Finally, we wanted to examine whether there are other loci that, similar to pho1, are controlled via unstable non-coding transcripts originating from intergenic regions. To first identify non-coding transcripts that are regulated by the exosome, we performed high resolution calibrated RNA-seq comparing wild-type with exosome deficient rrp6Δ. Our spike-in normalized results show up-regulation of multiple transcripts that were previously reported to be degraded by the exosome complex. We were also able to annotate new transcripts whose levels are controlled by the exosome (Supplementary Table 1). One such transcript is the novel intergenic cryptic unstable transcript (CUT) between the SPAC27D7.09c/11c loci (Figure Supplement 6). Interestingly, similar to meiotic genes, these CUTs are also up-regulated upon inhibition of HDAC activity. Interestingly, at least for some of the transcripts that overlap with ncRNA, we observe increases in H3K14 acetylation and Pol II transcription (Supplementary Table 3 and Figure Supplement 6 and 7). These transcripts include genes encoding for membrane transporters, such as the calcium transporter cta3, and hexose transporters, such as ght8. This suggests that other loci might be regulated via HDAC activity dependent on non-coding transcription, possibly via the activity of the Rpd3 (Clr6 and Hos2) complexes described in S. cerevisiae.
We conclude that class I and II HDACs play a significant role in shaping the fission yeast transcriptome by regulating transcription both positively and negatively. We have demonstrated that regions of facultative heterochromatin, including meiotic genes and genes regulated via non-coding transcription, such as pho1, require HDAC activity for their transcriptional repression.
Discussion
The ability of cells to invoke rapid changes in gene expression is critical for the execution of developmental programs, adaptation to environmental stress, and during the fate decision of stem cells52. Many meiotic mRNAs are repressed in mitotic cells and derepressed during meiosis when the function of the proteins they encode is needed. Similarly, expression of the genes encoding for metabolic enzymes such as pho1 is also dynamically regulated in response to nutrient availability5. Despite these genes being rooted in a chromatin environment that has the canonical features of heterochromatin (such as H3K9me and marks associated with transcriptional repression at centromeric and telomeric regions), loss of H3K9me upon deletion of Clr4 does not lead to up-regulation of their transcription4,5,16. Instead, it was believed that they were repressed primarily at the level of RNA stability. Here, we demonstrate that pho1 and meiotic genes are in fact repressed at the level of transcription by HDACs. This repression works in concert with RNA degradation and H3K9me (Figure 6). Indeed, we see a synthetic growth defect when suppression of HDACs (TSA treatment) is combined with inhibition of RNA degradation (rrp6Δ).
How HDACs control transcriptional repression of only specific regions of the genome is not fully understood. Recent studies have proposed that the pattern of histone methylation at promoters of protein-coding genes could provide the specificity. Histone H3 methylation by Set1 at promoter proximal regions controls gene expression either positively or negatively depending on the H3K4me2/me3 ratio23,24. A characteristic chromatin environment featuring high levels of H3K4me2 has been proposed to recruit the Rpd3L HDAC to repress transcription in budding yeast20, whereas H3K4me3 positively affects transcription. H3K4me2 peaks downstream of H3K4me3 and represses initiation at cryptic promoters to promote the production of full-length mRNA47. According to this model, unless H3K4me2 is introduced through the act of upstream overlapping non-coding transcription, genuine promoters should be resistant to Set1-dependent inhibition. We demonstrate that transcriptional silencing within facultative heterochromatin, in fission yeast, is established via non-coding transcription and relies on a Set1/Clr3 mechanism. Multiple non-coding transcripts recently reported to be produced from regions proximal to or overlapping meiotic transcripts53–55 might be responsible for mediating the effect of Set1 and Set2 in recruitment of class I/II HDACs to these genes. However, it is not entirely clear how widespread the role of non-coding transcription is in repressing regions of facultative heterochromatin derepressed by TSA. We show examples of protein-coding and non-coding transcripts that do not have any overlapping non-coding transcription and whose promoters are likely to be directly regulated by Set1. While it is not clear how Set1 represses genuine promoters one possibility could be that the ratio of H3K4me2/me3 is higher on these genes due to different Set1 residency times. Indeed, recent studies have shown that residency time on promoters can be regulated via Set1’s interaction with the nascent RNA. It was proposed that this could, in turn, affect the H3K4me2/me3 ratio36,37. This suggests an exciting possibility that Set1 binding to RNA is regulated at these promoters to create a specific chromatin environment that is not permissive for transcription. Alternatively, H3K4me2/me3 ratio could be regulated via demethylation activities targeted to these promoters.
Pervasive antisense transcription occurring as a result of Set2 deletion has been shown to modulate expression of highly regulated metabolic genes56, even though global antisense transcription does not show correlation with sense transcription57. Set2 deletion has also been shown to contribute to the regulation of tgp1 and pho116, although we show that Set2’s contribution at pho1 is minor compared to Set1’s. We have shown that Rpd3 complexes and other class I/II HDACs are not involved in tgp1 repression, suggesting that Set2 may repress this gene through a different mechanism. In contrast to constitutive heterochromatin where HDAC recruitment is dependent on H3K9me and sequence-specific DNA binding proteins (such as ATF/CREB family transcription factors Atf1 and Pcr139), at facultative heterochromatin Clr4-dependent deposition of H3K9me depends on the sequence-specific RNA-binding protein Mmi134. It has been proposed that Set1 can also be recruited to select euchromatic loci via Atf140. While our data suggest that the prt promoter proximal region is important for pho1 repression, we did not observe any effect on pho1 in an atf1Δ strain, arguing against a role for Atf1 in providing the specificity for recruitment of Set1. Furthermore, transcripts whose expression was reported to be increased upon deletion of Atf1 differ from the transcripts we report to be derepressed upon HDAC inhibition.
It has previously been proposed that SHREC promotes H3K9me13. The mechanistic details of how SHREC achieves this are unclear but it has been suggested that either deacetylation of H3 or of a non-histone substrate by Clr3 is necessary for Clr4-mediated methylation of H3K9. In contrast to the case at constitutive heterochromatin, we show that Clr3 and Clr4 proteins act in parallel, and that H3K14 deacetylation plays a much more prominent role than H3K9 methylation in transcriptional repression. In mammals, developmental genes are believed to be primarily silenced by H3K27me3 positioned by Polycomb Repressive Complex 2 (PRC2) in proliferating cells. However, this has been challenged by a recent study demonstrating that H3K27me-independent histone deacetylation plays an important role in transcriptional repression of developmental genes in mouse cardiomyocytes, and is important for normal heart function58. This implies that the role of HDACs in transcriptional silencing of developmentally regulated genes is likely conserved in mammals and therefore it will be even more important to understand how these complexes function.
This study significantly advances our understanding of the molecular mechanisms underlying repression of meiotic genes. However, many questions are still left unanswered. It remains to be seen how HDACs are targeted to regions of facultative heterochromatin and why it is that many genes are down-regulated upon loss of HDAC activity. Future studies addressing some of these unresolved questions will provide further insight into the function of HDACs in the regulation of transcription.
Materials and methods
Yeast strains and manipulations
S. pombe strains were grown in either YES medium, or EMMG with 10 mM KH2PO4 to an OD600 of 0.4 - 0.7 before harvesting59. Strains and oligonucleotides used in this study are listed in Supplementary Tables 4 and 5.
Standard PCR-based methodology was used for epitope tagging60. The pop-in, pop-out method for allele replacement was used to introduce deletions of the pho1 lncRNA61. DNA fragments carrying the desired deletions were generated by 2-step PCR. Fragments were confirmed by sequencing, and cloned into pCR blunt II TOPO (Life Technologies), according to the manufacturer’s instructions, before subcloning into pKS-URA460 by SpeI/NotI digestion and ligation. BseRI was used to linearize the plasmid before transformation into S. pombe cells lacking the ura4 locus. Ura+ colonies were selected on EMMG without uracil and genotyped by DNA sequence analysis of PCR products. Positive clones were plated onto 5-FOA to select cells in which the ura4+ gene had been ‘popped-out’. Clones were verified by colony PCR and sequencing of products.
Northern blotting
Northern blot experiments were essentially performed as described62. Gene-specific PCR-generated fragments were used as probes using oligonucleotides listed in Supplementary Table 5.
Chromatin immunoprecipitation (ChIP)
ChIP was performed as previously described5. Immunoprecipitations (IPs) were conducted with either rabbit IgG agarose (Sigma) or antibodies against H3 (Abcam, 1791, RRID:AB_302613), H3K14ac (Millipore, 07-353, RRID:AB_310545), Rpb1 (Millipore, 8WG16, RRID:AB_492629) or c-Myc (Santa Cruz, SC-40, RRID:AB_627268) coupled to protein G dynabeads (Life Technologies). Primers for qPCR analysis are listed in Supplementary Table 5.
RNase ChIP
RNase ChIP was performed as described above with the following exceptions: (i) formaldehyde cross-linking time was reduced from 20 min to 5 min (ii) Chromatin was prepared in FA lysis buffer with 0.05% SDS and (iii) cross-linked chromatin was treated with either 7.5 U of RNase A (Thermo Scientific) and 300 U of RNase T1 (Thermo Scientific) or an equivalent volume of RNase storage buffer (50 mM Tris-HCl (pH 7.4) and 50% (v/v) glycerol). After incubation at room temperature for 30 min, immunoprecipitations were performed as usual.
ChIP-seq
Chromatin was prepared as above. After washing and eluting bound material from the beads, protein was removed by incubation with 0.4 mg pronase for 1 hr at 42°C, followed by overnight incubation at 65°C. RNA was degraded by incubating samples with 0.02 mg RNase A (Roche) for 1 hr at 37°C. DNA was purified using ChIP DNA Clean & Concentrator kit (Zymo Research, USA) according to the manufacturer’s instructions. A sequencing library was constructed using NEBNext Fast DNA Library Prep Set for Ion TorrentTM Kit (NEB, USA). Libraries with different barcodes were pooled together and loaded onto the Ion PITM Chip v3 using the Ion ChefTM Instrument (Life Technologies, USA). Library sequencing was carried out on the Ion Torrent Proton. The resulting sequences were trimmed to remove low quality reads (less than Phred score 20) and reads shorter than 20 nt using Trimmomatic (version 0.36)63.
RNA sequencing
RNA-seq was performed in duplicates as described previously34. Libraries were prepared and sequenced by the High-Throughput Genomics Group at the Wellcome Trust Centre for Human Genetics on the Illumina HiSeq 2500 platform. Quality trimming was performed using Trimmomatic (Galaxy Version 0.32.3, RRID:SCR_011848)63.
For identification of CUTs, total RNA was extracted from biological duplicates of WT and rrp6Δ cells, using standard hot-phenol procedure. ERCC RNA Spike-In (2 μl of 1:100 dilution; Ambion) was added to 1 μg of total RNA, then rRNAs were depleted using the RiboMinus Eukaryote System v2 (Life Technologies). Strand-specific total RNA-Seq libraries were prepared using the TruSeq Stranded Total RNA Sample Preparation Kit (Illumina). Paired-end sequencing (2x50 nt) of the libraries was performed on a HiSeq 2500 sequencer.
NET-seq
Biological duplicates of Rpb3-Flag S. pombe cells were treated for 2.5 hours with 20 μg/ml of the HDAC inhibitor trichostatin A (TSA). After lysis, each S. pombe lysate was mixed with an aliquot of a lysate of Rpb3-Flag S. cerevisiae cells (spike), in a 50:1 ratio.
Libraries were constructed according to the previously published protocol49, with minor modifications, starting from 1L of exponentially growing cells. Ligation of DNA 3’-linker was performed as described64, starting with 2-3 μg of purified nascent RNA. Ligated nascent RNA was then submitted to alkaline fragmentation for 20 minutes at 95°C. Single-end sequencing (50 nt) of the libraries was performed on a HiSeq 2500 sequencer.
After removal of the 5’-adapter sequence using cutadapt, reads were uniquely mapped to the S. pombe and S. cerevisiae reference genomes using version 2.2.5 of Bowtie, with a tolerance of 1 mismatch. Bioinformatics analyses used uniquely mapped reads. Tags densities were normalized either on the total number of uniquely mapped reads (IP & Input samples), or the signal for S. cerevisiae ribosomal proteins-coding genes.
Processing and bioinformatic analysis of genome-wide data
Reads were aligned to the S. pombe genome (ASM294v2.28) and the S. cerevisiae genome (sacCer3) separately either with TopHat (Galaxy Version 0.9, RRID:SCR_013035)65 in the case of RNA-Seq, or with Bowtie2 v2.2.6 (RRID:SCR_005476)66 in the cases of NET-Seq and ChIP-Seq. Reads that were present in both alignments were removed using the Picard Tool Suite (http://broadinstitute.github.io/picard) to keep only reads unique for one of the genomes. All remaining S. cerevisiae reads were counted and used for the normalization of S. pombe samples. In the case of ChIP-Seq, differences in the input was also taken into account when calculating normalization values and H3K14ac was normalized to H3 density as determined by H3 ChIP-Seq. For NET-Seq, reads were trimmed to leave only the 3’ nucleotide corresponding to the nucleotide in the active centre of Pol II at the time of the experiment. For RNA-Seq, only properly aligned pairs were kept and all other reads were removed using samtools67. Spearman correlation matrices and metagene plots were calculated using deepTools68. For differential expression analysis, indicated regions were counted using R for ChIP-Seq and NET-Seq while HTSeq was used for RNA-Seq data69. The analysis for all data sets was performed with the R package DESeq2 (RRID:SCR_000154)70 using manual normalization to S. cerevisiae spike-in counts determined as described above. All other data analysis was also performed with R using in-house scripts featuring Bioconductor packages71,72.
For CUTs annotation, reads were mapped to the S. pombe reference genome using version 2.0.6 of TopHat, with a tolerance of 3 mismatches and a maximum size for introns of 5 kb. Tags densities were normalized on the ERCC Spike-In signals. Segmentation was performed using the ZINAR algorithm73. CUTs were defined as ≥200 nt segments showing a >2-fold enrichment in rrp6Δ vs WT, with a P-value (adjusted for multiple testing with the Benjamini-Hochberg procedure) <0.05 upon differential expression analysis using DESeq74.
Data access
Raw (fastq) and processed sequencing data can be downloaded from the NCBI Gene Expression Omnibus repository (http://www.ncbi.nlm.nih.gov/geo, accession number GSE104713 (RNA-seq), GSE104712 (NET-seq).
Contributions
B.R.W. and L.V. conceived and designed experiments. B.R.W. performed all experiments except rrp6Δ RNA-seq which was performed by M.W. and analysed by C.G. S.W. analysed ChIP-seq, RNA-seq, PAR-CLIP and NET-seq datasets. K.K and D-H.H also contributed to genome-wide analyses. B.R.W. and L.V. wrote the paper and all authors edited the manuscript.
Acknowledgements
We thank H.D Madhani, and the National BioResource Project (NBRP) for strains. We are grateful to Sofia Battaglia and Patrick Cramer for contributions not shown here. This work was supported by the Wellcome Trust Research and Career Development and Wellcome Trust Senior Research fellowships to L.V. (WT088359MA and WT106994MA). High throughput sequencing was performed by the High-Throughput Genomics Group at the Wellcome Trust Centre for Human Genetics (Wellcome Trust grant 090532/Z/09/Z). This work has benefited from the facilities and expertise of the High-Throughput Sequencing platform of Institut Curie (Paris), supported by the Agence Nationale de la Recherche (ANR-10-EQPX-03, ANR10-INBS-09-08). The BBSRC provided support to B.R.W. and S.W. was supported by a studentship from the MRC.