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. 2026 Feb 16;31(2):e70092. doi: 10.1111/gtc.70092

A Temperature Increase Induces Atf1/Pcr1‐Dependent Rapid Depletion of Histones in Transcriptionally Activated Gene Bodies in Fission Yeast

Arata Hoshino 1, Takuya Kajitani 2, Masaya Oki 2, Tetsuji Kakutani 1,✉, Soichi Inagaki 1,✉
PMCID: PMC12909096  PMID: 41698824

ABSTRACT

Transcriptional regulation is central to organisms' response and adaptation to environmental changes. However, the coordination of transcriptional machinery with chromatin when rapid transcriptional changes are needed is largely unknown. Here, we studied the rapid response of histone modifications upon heat treatment of fission yeast Schizosaccharomyces pombe. We observed a significant eviction of H3 in the transcribed regions of upregulated genes, named Hasty Overheat‐Triggered Differentially Occupied Genes (HOT‐DOGs), including Heat Shock Protein‐encoding genes. The loss of H3 in a third of the HOT‐DOGs was dependent on the Atf1‐Pcr1 transcription factor complex. Furthermore, the H3K9 methyltransferase Clr4 likely modulates the rate of transcription elongation and helps prevent transcriptional read‐through beyond polyadenylation sites in these genes. Our results unveiled a novel function of Clr4 in regulating the dynamics of transcription within gene bodies and provided insights into how cells cope with rapid environmental changes while preserving chromatin integrity.

In the fission yeast Schizosaccharomyces pombe, the transcription factor complex Atf1‐Pcr1 is activated by phosphorylation upon heat stress to initiate a rapid cellular response. This activated complex facilitates the significant eviction of histone H3 from the transcribed regions of specific genes termed HOT‐DOGs. RNAP II, depicted as delivery trucks, then conducts high‐speed transcription elongation within these gene bodies. To regulate this process, the H3K9 methyltransferase Clr4 modulates the rate of elongation to ensure proper transcription dynamics. Furthermore, Clr4 prevents transcriptional read‐through beyond polyadenylation sites, acting as a barrier to maintain genomic stability. Together, these factors allow the cell to adapt to sudden environmental changes while effectively preserving chromatin integrity.

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1. Introduction

In eukaryotic cells, external stimuli cause transcriptional reprogramming; some genes are transcriptionally activated and others are repressed. Transcription takes place on chromatin, which is a highly organized, dense structure. Chromatin is composed of the basic unit, nucleosome, where ~147 bp DNA is wrapped around a histone octamer consisting of two copies each of core histone proteins H2A, H2B, H3, and H4. Transcription by RNA polymerase II (RNAPII) complex on chromatin temporarily disassemble nucleosomes and therefore chromatin is an intrinsic obstacle for the transcription process (Kujirai and Kurumizaka 2020; Petesch and Lis 2012; Teves et al. 2014). How cells realize rapid transcriptional induction in response to environmental changes while preserving chromatin structure is largely unknown.

Chromatin structure can determine the accessibility of transcription factors and RNAPII. Post‐translational modifications, such as methylation, acetylation, and ubiquitination, of histone proteins affect the chromatin structure and transcriptional activity of the underlying DNA. The active histone mark, such as lysine acetylation on H3 or H4, facilitates access of transcription factors and RNAPII through making chromatin open, and activates the transcription (Brownell et al. 1996; Li et al. 2007; Nightingale et al. 1998). Actively transcribed genes are also marked by methylation of H3 lysine‐4 (H3K4): monomethylation (H3K4me1), dimethylation (H3K4me2), and trimethylation (H4K4me3). In budding yeast and humans, the H3K4 methyltransferase Set1 (in yeast) and SETD1A/B complex (in humans) interact with the RNA polymerase II (RNAPII), thus depositing H3K4me3 on the transcribed regions of genes during transcription (Bae et al. 2020; Ebmeier et al. 2017; Ng et al. 2003). Cooperation between H3K4 methyltransferase/demethylase and the RNAPII complex was also reported in other species, such as plants (Inagaki et al. 2021; Mori et al. 2023; Oya et al. 2022). However, the roles of transcription‐coupled modulation of histone modifications during normal development and during rapid environmental changes remain largely unknown.

In contrast to these active marks, histone H3 lysine‐9 methylation (H3K9me) is distributed mostly in condensed and transcriptionally repressed heterochromatin regions (Allshire and Madhani 2018; Janssen et al. 2018; Padeken et al. 2022; Zhang et al. 2018). In the fission yeast Schizosaccharomyces pombe, which has been a model to study heterochromatin regulation, H3K9me is conferred by the sole H3K9 methyltransferase, Clr4 (Jih et al. 2017; Nakayama et al. 2001; Zhang et al. 2008). Loss of Clr4 causes transcriptional activation of genes in heterochromatin regions, indicating that Clr4 represses transcription through H3K9 methylation. H3K9me provides a binding platform for the heterochromatin protein 1 (HP1) family of proteins which play central roles in transcriptional repression by forming compact and closed chromatin structures to restrict the access of RNAPII. (Hall et al. 2002; Iglesias et al. 2020; Motamedi et al. 2008; Noma et al. 2001; Sugiyama et al. 2007).

In fission yeast, the mitogen‐activated protein kinase (MAPK) Sty1/Spc1 and its downstream factors Atf1 and Pcr1, the basic zipper (bZIP)‐containing transcription factors, are activated by external stimuli and promote the transcriptional activation of stress responsive genes (Chen et al. 2003; Eshaghi et al. 2010; Lawrence et al. 2007; Wilkinson et al. 1996). Furthermore, the change of nucleosome distribution is observed in the gene body of a subset of Atf1‐regulated genes upon oxidative stress, and the histone acetyltransferase Gcn5 controls H3K9 and H3K14 acetylation and chromatin remodeling in these genes (Sansó et al. 2011). Paradoxically, Atf1 and Pcr1 also contribute to heterochromatin assembly at the mating‐type locus by binding to cis‐elements in this region and recruiting Clr4 (Jia et al. 2004; Wang and Moazed 2017). Thus, Atf1/Pcr1 is located at the center of the regulation of transcription, chromatin structure and epigenetic modifications upon external stimuli in fission yeast.

To respond appropriately to environmental fluctuations, cells must reprogram transcription and chromatin structure rapidly. However, genome‐wide observations of chromatin dynamics during rapid environmental responses have been limited. Furthermore, the changes in epigenetic modifications that accompany transcriptional reprogramming during short‐term environmental responses remain largely unexplained. In this study, we analyzed the genome‐wide methylation of H3K4 and nucleosome dynamics (represented as H3) within a few minutes after heat treatment to capture the short‐term response of chromatin modifications to environmental changes in S. pombe. We found that H3 is rapidly depleted in the whole gene body of the transcriptionally activated gene loci. We explored the genetic mechanism of this substantial reduction in histones in transcriptionally activated genes. Our results revealed an unexpected role of Clr4 in controlling transcription elongation and termination of heat‐inducible genes, and we discussed the implication of this mechanism in preserving chromatin integrity during rapid environmental changes.

2. Results

2.1. Heat‐Induced Transcriptional Upregulation Accompanies H3 Eviction in Gene Bodies

We investigated histone modification dynamics over a short time scale by analyzing H3K4 mono‐, di‐, and trimethylation (H3K4me1, 2, 3), and H3 (as control) along with the transcriptome at 0 (non‐treated control), 5, and 25 min after heat (37.5°C) treatment of S. pombe (Figure 1A,B). We observed two changes within the transcribed regions of a subset of genes with upregulated expression levels in response to heat, such as heat‐shock protein 16 (hsp16), hsp104, and a hsp70 family gene, ssa1. (1) H3 levels significantly decreased 5 and 25 min after heat treatment (Figure 1B,C). (2) H3‐normalized H3K4me1/2/3 levels significantly increased 5 and 25 min after heat treatment (Figure 1C). Although it has been reported that nucleosomes get loose in highly transcribed genes (Sansó et al. 2011; Vega et al. 2023), we did not expect them to be evicted over the whole gene body in such a short time. We named these genes with increased expression and decreased H3 occupancy after heat treatment as “Hasty Overheat‐Triggered Differentially Occupied Genes (HOT‐DOGs)” (red dots in Figure 1C and Table S1) and further analyzed them in this study. Subsequently, we analyzed H3 and the transcriptome at earlier time points after heat treatment (Figure 2A). H3 started to decrease in the HOT‐DOGs within 2.5 min after heat treatment (Figure 2B−D).

FIGURE 1.

FIGURE 1

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A subset of heat‐induced genes lose H3 and gain H3K4me upon heat. (A) The scheme of time‐course experiments. Dots indicate the timing of sampling. (B) Browser views showing H3, H3K4me, and mRNA profiles around heat‐inducible HOT‐DOGs. HOT‐DOGs are highlighted in red. The numbers in brackets indicate the scale of normalized values (per million mapped reads). (C) Relationships between H3 or H3‐normalized H3K4me changes and mRNA changes at 5 and 25 min compared to 0 min after heat treatment. Black dots, all protein‐coding genes; red dots, HOT‐DOGs. The kernel density plots on the top and side of each panel indicate the distribution of the log2 ratio for protein‐coding genes and HOT‐DOGs, painted in black and red, respectively. In each panel, the ChIP‐seq data (vertical axis) represents the average of the two biological replicates and the mRNA‐seq data (horizontal axis) represents the average of the three biological replicates.

FIGURE 2.

FIGURE 2

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HOT‐DOGs are transcriptionally activated and lose H3 immediately after heat treatment. (A) The scheme of time‐course experiments. Dots indicate the timing of sampling. (B) Browser views showing H3, mRNA, and nascent RNA (PRO‐seq) profiles around heat‐inducible HOT‐DOGs. The HOT‐DOGs are highlighted in red. The numbers in brackets indicate the scale of normalized values (per million mapped reads). (C) Relationships between H3 changes and mRNA or nascent RNA changes at 2.5 and 5 min compared to 0 min after heat treatment. Black dots, all protein‐coding genes; red dots, HOT‐DOGs. The kernel density plots on the top and side of each panel indicate the distribution of the log2 ratio for protein‐coding genes and HOT‐DOGs, painted in black and red, respectively. In each panel, the ChIP‐seq data (vertical axis) and the PRO‐seq data (horizontal axis) represent the average of the two biological replicates, and the mRNA‐seq data (horizontal axis) represents the average of the three biological replicates. (D) Averaged profiles of H3 (top), PRO‐seq (middle), and mRNA‐seq (bottom) around all protein‐coding genes (left) and HOT‐DOGs (right). The line plots indicate the mean and the shaded areas indicate the standard error. TSS, transcription start site; PAS, poly‐A site. (E) Gene Ontology terms enriched in HOT‐DOGs and –log10 (p‐value) of Fisher's exact test for enrichment of the terms.

We performed Precision Run‐On Sequencing (PRO‐seq) before and after (2.5 and 5 min) heat treatment to determine whether the increased transcript levels of HOT‐DOGs upon heat treatment were owing to transcriptional activation (Figure 2A). PRO‐seq maps nascent RNA synthesis by RNA polymerases with base‐pair resolution (Mahat et al. 2016). The HOT‐DOGs showed increased PRO‐seq signals within 2.5 min after heat treatment along the whole gene body, as seen in the mRNA‐seq (Figure 2B–D), supporting the idea that heat promotes the transcription of HOT‐DOGs. We also performed PRO‐seq and ChIP‐seq after returning the culture condition to 30°C to analyze the dynamics after heat stress (Figure S1A). As a result, transcriptional upregulation and the loss of H3 rapidly diminished after returning to 30°C (Figure S1B). Gene Ontology (GO) analysis of the HOT‐DOGs showed that GO terms such as “protein folding” and “response to heat” were significantly enriched (Figure 2E). Together, our results revealed that transcriptional activation accompanied the loss of H3 over the whole gene body at early time points after heat treatment.

Next, we investigated whether the loss of H3 is specific to rapidly upregulated genes, (i.e., HOT‐DOGs upon heat), or is common to highly transcribed genes. Using the PRO‐seq data, we selected 100 genes with the highest or lowest transcription levels under non‐stressed conditions (0 min), called “top 100 genes” or “bottom 100 genes,” respectively, and compared the signals of PRO‐seq, mRNA‐seq, and H3 ChIP‐seq at each timepoint (Figure 3A). Although the H3 levels are almost the same in the top 100 genes compared to other classes of genes, a drastic loss of H3 is observed only in HOT‐DOGs after heat treatment (Figure 3A,B). Even after heat treatment, the transcription levels of HOT‐DOGs are comparable or lower than the top 100 genes as shown by PRO‐seq (Figure 3A). Therefore, the loss of H3 is associated with rapid upregulation rather than a constantly high level of transcription. In addition, the loss of H3 occurred not only around TSS, but also over the whole gene body in HOT‐DOGs upon heat (Figure 3C), while the top 100 genes showed H3 depletion at TSS and upstream regions, likely reflecting open chromatin at transcription factor‐binding sites in promoters. These results suggest that the loss of H3 in the whole gene body is not always associated with frequent transcription, but triggered by hasty transcription.

FIGURE 3.

FIGURE 3

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Loss of H3 in gene body is specific to rapidly induced genes. (A) Kernel density plots for PRO‐seq (left), mRNA‐seq (middle), and H3 ChIP‐seq (right) signals showing distribution of all genes (black), HOT‐DOGs (red), top 100 genes (green), and bottom 100 genes (blue). RPKM, read per kilobase per million mapped reads; TPM, transcripts per million. (B) Kernel density plots showing distribution of the ratio between 0 min and 2.5 min (top) or 5 min (bottom) after heat treatment. (C) Averaged profiles of H3 level around all protein‐coding genes, top 100, top 100–200, top 200–300 genes, and HOT‐DOGs. The line plots indicate the mean and the shaded areas indicate the standard error.

2.2. Heat‐Induced H3 Depletion Is Dependent on Atf1 and Pcr1 in Some HOT‐DOGs

In S. pombe, a basic zipper (bZIP)‐containing transcription factor Atf1 primarily regulates the transcription of stress responsive genes (Chen et al. 2003; Shiozaki and Russell 1996; Wilkinson et al. 1996). Indeed, the atf1Δ mutant exhibits hyper‐sensitivity to several stressors, such as osmotic, oxidative and heat stresses (Kon et al. 1997; Paulo et al. 2014; Quinn et al. 2002; Sansó et al. 2008). To investigate whether Atf1 is involved in the regulation of transcription in HOT‐DOGs, we performed mRNA‐seq in the atf1Δ mutant before and after heat treatments. As expected from the Atf1 function, the expression of subsets of HOT‐DOGs, such as gpd1, rtc3 and hsp9, decreased in atf1Δ compared to WT upon heat (Figure 4A). Atf1 forms a heterodimeric complex with Pcr1 and functions in stress‐responsive transcriptional regulation (Chen et al. 2003; Eshaghi et al. 2010; Kon et al. 1997). Therefore, we also tested whether Pcr1 is involved in the upregulation of HOT‐DOGs during heat treatment. The pcr1Δ strain showed decreased expression of these HOT‐DOGs like in atf1Δ upon heat treatment (Figure 4A). Further transcriptome analyses in atf1Δ and pcr1Δ before and after heat treatment revealed that 26 and 27 genes out of 78 HOT‐DOGs showed attenuated transcriptional induction upon heat in atf1Δ and pcr1Δ compared with WT, respectively (Figure 4B,C and Table S1). These Atf1‐ and Pcr1‐regulated HOT‐DOGs also showed an attenuation of H3 loss upon heat in atf1Δ and pcr1Δ (Figures 4A,D,E and S2A–D). Of the 27 genes in Pcr1‐regulated HOT‐DOGs, 26 genes overlapped with Atf1‐regulated HOT‐DOGs (Figure S4E). We called these genes “Atf1/Pcr1‐regulated HOT‐DOGs” in subsequent analysis.

FIGURE 4.

FIGURE 4

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Atf1 and Pcr1 are required for transcriptional activation and H3 eviction in a subset of HOT‐DOGs. (A) Browser views showing H3 and mRNA profiles around heat‐inducible HOT‐DOGs in WT, atf1Δ, and pcr1Δ mutants. The HOT‐DOGs are highlighted in red. The numbers in brackets indicate the scale of normalized values (per million mapped reads). (B, C) Volcano plots showing the comparison of mRNA (TPM) between WT and atf1Δ (B) or pcr1Δ (C) mutant at 0 and 5 min. Atf1‐regulated HOT‐DOGs and Pcr1‐regulated HOT‐DOGs (log2 Fold‐change < −1 and FDR < 0.05 at 5 min) are highlighted in blue and green, respectively, and other HOT‐DOGs are shown in red. (D, E) Relationships between H3 changes and mRNA changes in atf1Δ (D) or pcr1Δ (E) mutants at 5 min compared to 0 min after heat treatment. Black dots, all protein‐coding genes; blue dots in (D) or green dots in (E) mutants, Atf1‐ or Pcr1‐regulated HOT‐DOGs, respectively; red dots, the other HOT‐DOGs. The Kernel density plots on the top and side of each panel indicate the distribution of the log2 ratio for protein‐coding genes, Atf1‐ or Pcr1‐regulated HOT‐DOGs, and the other HOT‐DOGs, painted in black, blue in (D) or green in (E), and red, respectively. In each panel, the ChIP‐seq data (vertical axis) represents the average of the two biological replicates and the mRNA‐seq data (horizontal axis) represents the average of the three biological replicates. (F) ChIP‐seq profiles for Atf1 (left) and Pcr1 (right) in all genes, Atf1‐ or Pcr1‐regulated HOT‐DOGs, and other HOT‐DOGs, based on the previously reported data in (Skribbe et al. 2025).

We reanalyzed recently published ChIP‐seq data for the Atf1 and Pcr1 proteins (Skribbe et al. 2025), and found that Atf1 and Pcr1 strongly accumulated at the upstream of TSS of the Atf1‐ and Pcr1‐regulated HOT‐DOGs, respectively (Figure 4F). These results support the idea that both Atf1 and Pcr1 directly regulate the transcription and nucleosome eviction at the subsets of HOT‐DOGs.

2.3. H3K9 Methyltransferase Clr4 Influences Transcription Dynamics Upon Heat Treatment

The Atf1‐Pcr1 heterodimer is also known to have a role for heterochromatin silencing at the mating type locus in S. pombe (Jia et al. 2004; Wang and Moazed 2017). We wondered whether there was a link between the downstream factors of Atf1‐Pcr1 involved in heterochromatin formation and Atf1/Pcr1‐regulated HOT‐DOGs. To investigate the role of Clr4, a histone H3 lysine 9 methyltransferase, which is an essential factor for heterochromatin formation, in the transcription regulation of Atf1/Pcr1‐regulated HOT‐DOGs, we performed mRNA‐seq and PRO‐seq in WT and clr4Δ before and after heat treatment (2.5 and 5 min) (Figure 5A). “Heterochromatin genes” that are embedded within H3K9me2‐accumulating regions and are repressed by Clr4 in normal conditions (including genes in the subtelomere and non‐coding transcription units in pericentromeric regions) showed clear increases in PRO‐seq signals along the transcription units in clr4Δ compared to WT before and after heat treatment (Figure S3A–C and Table S2). Additionally, the signals of mRNA‐seq and PRO‐seq significantly increased after heat treatment within Atf1/Pcr1‐regulated HOT‐DOGs in WT and clr4Δ (Figures 5B,C and S3D). These results confirmed that our mRNA‐seq and PRO‐seq data represent transcriptional dynamics in WT and clr4Δ. The mRNA levels of Atf1/Pcr1‐regulated HOT‐DOGs significantly increased before and after heat treatment in clr4Δ compared to WT (Figure 5C,D), suggesting that the presence of Clr4 negatively impacts transcription of these genes. To analyze the function of Clr4 in nucleosome dynamics after heat treatment, we compared the H3 levels in WT and clr4Δ before and after heat treatment. As a result, the loss of H3 was less conspicuous in clr4Δ compared to WT (Figure S4A). Therefore, increased mRNA levels of HOT‐DOGs in clr4Δ are not attributed to accelerated eviction of histones.

FIGURE 5.

FIGURE 5

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H3K9 methyltransferase Clr4 regulates transcription dynamics of HOT‐DOGs. (A) The scheme of time‐course experiments. Dots indicate the timing of sampling. (B) Browser view showing H3, mRNA (WT and clr4Δ), and PRO‐seq (WT and clr4Δ) profiles around an Atf1/Pcr1‐regulated HOT‐DOG (gpd1), highlighted in red. The numbers in brackets indicate the scale of normalized values (per million mapped reads). (C) Averaged profiles of the signals for mRNA‐seq (left) and PRO‐seq (right) around Atf1/Pcr1‐regulated HOT‐DOGs. The line plots indicate the mean, and the shaded areas indicate the standard error. (D, E) Kernel density plots showing the distribution of the log2 ratio of mRNA‐seq (D) and PRO‐seq (E) signals within the gene bodies in the clr4Δ mutant to WT for all protein‐coding genes, Atf1/Pcr1‐regulated HOT‐DOGs, and other HOT‐DOGs, painted in black, blue, and red, respectively. The p‐values are based on two‐sample Kolmogorov–Smirnov tests comparing against all protein‐coding genes. (F) Definitions of the gene body and read‐through regions used in (G). (G) Kernel density plots showing the distribution of the log2 ratio of PRO‐seq signals within the gene bodies to the read‐through in the clr4Δ mutant to WT. The p‐values are based on two‐sample Kolmogorov–Smirnov tests comparing against all protein‐coding genes.

Interestingly, although clr4Δ showed an increased PRO‐seq signal within Atf1/Pcr1‐regulated HOT‐DOGs after heat treatment (Figure 5C), the intragenic PRO‐seq signal was lower in clr4Δ than in WT (Figure 5C,E). The decreased PRO‐seq signal indicated that the elongation rate of transcription was accelerated in Atf1/Pcr1‐regulated HOT‐DOGs in clr4Δ compared to WT since mRNA‐seq signals after heat treatment within the same set of genes increased in clr4Δ compared with WT. This result contrasts with the significantly increased mRNA‐seq and PRO‐seq signals within heterochromatin genes in clr4Δ compared to WT (Figure S3A–C). Therefore, Clr4 differentially affected the transcription of heterochromatin genes and Atf1/Pcr1‐regulated HOT‐DOGs. Furthermore, the read‐through signals beyond the polyA site showed a significant increase in clr4Δ compared to WT (Figure 5C) and the ratio of read‐through signal to gene‐body signal was significantly higher in clr4Δ than in WT after heat treatment (Figure 5F,G). The clr4Δ‐induced increase of read‐through signal by PRO‐seq was observed not only for Atf1/Pcr1‐regulated HOT‐DOGs but generally for all genes (Figures 5G and S3A), suggesting that the Clr4‐controlled transcription elongation and termination is a general mechanism. These analyses were also performed on HOT‐DOGs that are not Atf1/Pcr1‐regulated (referred to as other HOT‐DOGs), and it was revealed that both the increase in mRNA levels in gene body and the decrease in intragenic PRO‐seq signals in clr4Δ upon heat were smaller than those observed for Atf1/Pcr1‐regulated HOT‐DOGs (Figure 5D,E). Furthermore, the ratio of read‐through to gene‐body after 5 min heat treatment showed no significant difference compared to protein‐coding genes (Figure 5G), suggesting that Clr4 has a weaker influence on the regulation of transcription elongation in the other HOT‐DOGs. The transcription kinetics of HOT‐DOGs during and after heat treatment was not affected by clr4Δ (Figure S4B,C). Taken together, our results suggest that Clr4 negatively regulates the transcription elongation rate, which likely prevents transcription read‐through beyond the transcription termination sites.

Since Clr4 possesses H3K9 methylation (H3K9me) enzyme activity, we wondered whether H3K9me is part of the Clr4‐mediated regulation of transcription elongation in HOT‐DOGs. To investigate this, we performed H3K9me2 ChIP‐seq analysis in WT and clr4Δ with spike‐in control. The ratio of H3K9me2 level in clr4Δ to those in WT was very low in heterochromatin genes, indicating that Clr4 is responsible for H3K9me2 in these regions. On the other hand, the clr4Δ/WT ratio for H3K9me2 in Atf1/Pcr1‐regulated or other HOT‐DOGs was comparable to all protein‐coding genes before and after heat treatment (Figure S4D), suggesting that HOT‐DOGs are not specifically targeted by Clr4 for H3K9 methylation. Therefore, the role of Clr4 in regulating transcription of heat‐induced HOT‐DOGs could be separated from its role as H3K9 methyltransferase.

3. Discussion

This study demonstrated that a loss of H3 across the whole gene body was specifically observed in genes whose transcription was rapidly induced after heat treatment. Considering several previous reports, H3 depletion likely represents nucleosome eviction (Sansó et al. 2011; Vega et al. 2023; Zhao et al. 2005). While H3 levels are low around TSS of highly transcribed genes, the loss of H3 across the gene body was not associated with a constantly high level of transcription but triggered by hasty transcription upon heat as in HOT‐DOGs (Figure 3). Furthermore, transcriptional induction is not always associated with a loss of H3 (Figures 1C and 2C). Therefore, hasty transcription‐triggered nucleosome eviction is likely a regulated process rather than an inevitable consequence of transcription. Similar nucleosome eviction at transcriptionally induced gene loci is reported in Drosophila, budding yeast, and fission yeast (Kristell et al. 2010; Petesch and Lis 2008; Teves and Henikoff 2011; Zhao et al. 2005). It would be interesting to see if transcription‐coupled nucleosome eviction triggered by stimuli is a common phenomenon in other eukaryotic species.

The Atf1‐Pcr1 heterodimer complex plays two distinct roles. One is as a transcription factor that is activated by the mitogen‐activated protein kinase (MAPK) Sty1 upon receiving environmental stimuli and induces the transcription of downstream genes (Chen et al. 2003; Eshaghi et al. 2010; Lawrence et al. 2007). The other is as a heterochromatin factor that binds to the cis‐acting repressor element III (REIII site) at the mat locus and recruits Clr4 to introduce H3K9 methylation and induce transcriptional repression (Jia et al. 2004; Sun et al. 2024). Switching of the two distinct functions of Atf1 is coordinated by its phosphorylation state (Salat‐Canela et al. 2017; Sun et al. 2024). In our study, Atf1 and Pcr1 were required for transcriptional activation and the hasty transcription‐induced depletion of H3 in a subset of HOT‐DOGs (Figure 4), and Atf1 and Pcr1 bind to the promoters of these genes (Figure 5). Furthermore, these Atf1/Pcr1‐regulated HOT‐DOGs were regulated by Clr4 at the level of transcription elongation (Figure 5). Previous studies reported that Atf1 interacts with the Heterochromatin Protein1 (HP1) protein Swi6, and Swi6 interacts with and recruits Clr4 at heterochromatin regions (Haldar et al. 2011; Sun et al. 2024), suggesting that Clr4 could be specifically recruited to Atf1/Pcr1‐regulated HOT‐DOGs via Atf1‐interacting Swi6. Taken together, our results suggest that the Atf1‐Pcr1 complex and Clr4 regulate transcriptional initiation and elongation of Atf1/Pcr1‐regulated HOT‐DOGs, respectively, and Atf1‐Pcr1 and Clr4 could be functionally connected via Swi6. The coordination of these mechanisms between heterochromatin formation and stress‐induced transcription will be an interesting topic for future studies. Upon heat, the atf1Δ and pcr1Δ mutants still deplete nucleosomes in two thirds of HOT‐DOGs, suggesting that other factors may also function to upregulate transcription and nucleosome eviction for other HOT‐DOGs. Furthermore, clr4Δ also affects transcription dynamics of other HOT‐DOGs (Figure 5D–G), with the effect extending even to transcription read‐through across all genes (Figure 5G). Therefore, our results illuminate a previously unappreciated function of the heterochromatin factor Clr4 as a transcription regulator of protein‐coding genes.

Genome‐wide transcription analyses in WT and clr4Δ suggested that Clr4 has an impact on the transcription dynamics of Atf1/Pcr1‐regulated HOT‐DOGs by slowing down the transcription elongation rate of RNAPII and preventing transcription read‐through beyond polyadenylation sites (Figure 5). Increased transcription velocity by expressing mutant RNAPII induces transcription read‐through (Cortazar et al. 2019; Fong et al. 2015; Leng et al. 2020). Therefore, regulating the transcription elongation rate by Clr4 in Atf1/Pcr1‐regulated HOT‐DOGs may be required for the appropriate control of transcription termination. Additionally, the elongation of RNAPII drives nucleosome turnover in the body of highly transcribed genes (Deal et al. 2010; Teves and Henikoff 2011) and the nucleosome can act as a barrier to transcription elongation (Kujirai and Kurumizaka 2020; Petesch and Lis 2012; Teves et al. 2014). Indeed, the elongation rate of RNAPII is extremely slow on a chromatin template compared to that of naked DNA in vitro (Izban and Luse 1992). These reports suggest that transcription elongation is a destructive process that pushes away interfering nucleosomes. In particular, when it is necessary to rapidly and massively express stress‐responsive genes under environmental stress, the absence of nucleosomes in the gene‐body region is advantageous for fast transcription, but it could also cause chromatin fragility. Considering these previous studies, Clr4 may compensate for the loss of nucleosomes to coordinate the elongation rate of transcription and preserve chromatin integrity. Given that Clr4 possesses the enzyme activity of H3K9 methylation, it is possible that H3K9me is involved in the regulation of transcription elongation and the compensation of the nucleosome loss in these genes. Furthermore, previous studies have shown that mutations defective in RNAi‐dependent heterochromatin formation upregulate expression of stress responsive genes including a subset of HOT‐DOGs (Kajitani et al. 2017; Woolcock et al. 2012), further connecting H3K9 methylation with stress responsive transcription regulation. However, H3K9me2 did not accumulate specifically in HOT‐DOGs upon heat, leaving the involvement of H3K9me in the regulation of transcription dynamics of HOT‐DOGs unclear. Future studies are required to determine how Clr4 regulates transcription elongation and affects chromatin structure in challenged conditions.

4. Experimental Procedures

4.1. Yeast Strains and Normal Growth Conditions

All strains used in this study are described in Table S3. Cells were grown at 30°C in 100 mL of YES medium, which includes 0.5 g of yeast extract (Biokar diagnostics), 3 g of glucose (Sigma‐Aldrich), and 0.1125 g of 5S supplement mix (Adenine, Lysine, Leucine, Histidine and Uracil; Sigma‐Aldrich). Wild‐type (WT; SFY1) and clr4Δ (SFY30) were kindly gifted from the Murakami laboratory at Hokkaido University. The other strains were generated by inserting the cassette of an antibiotics‐resistant gene for kanamycin (KanMX6) into the ORF of the target genes using homologous recombination (Bähler et al. 1998). The transformation was performed with electroporation.

4.2. Cell Culture and Sampling for ChIP‐Seq and mRNA‐Seq

When cells reached a density between 2.16 and 3.05 × 106 cells mL−1, untreated (0 min) cells were collected immediately. The flasks for heat‐treated samples were moved from a 30°C water bath (TAITEC) to a 37.5°C water bath and incubated for the specified time (2.5, 5, or 25 min) with shaking. For the experiments of ChIP‐seq in Figures S1A,B and S4B,C, the flasks were returned to a 30°C water bath after the 5 min incubation at 37°C, and cultured for the specified time (2.5, 5, or 10 min) with shaking. At each time point, 900 μL of cell culture medium was dispensed for mRNA‐seq into a 1.5 mL Eppendorf tube and was immediately frozen with liquid nitrogen and stored at −80°C. The remaining cell culture medium was cross‐linked with 1% formaldehyde (Nacalai) and incubated for 15 min with continuous shaking. Then, fixed cells were centrifuged at 4°C, 3000 g for 2 min to remove the supernatant, and the pellet of those cells was washed with 20 mL of PBS buffer once. Then, the samples were centrifuged at 4°C, 3000 g for 2 min, the supernatant was discarded, and the pellet was frozen with liquid nitrogen and stored at −80°C.

4.3. ChIP‐Seq

ChIP‐seq for histone modification was performed as described previously (Sorida et al. 2019) with some modifications. Briefly, the stored samples were resuspended by 400 μL of ChIP buffer I (50 mM HEPES‐KOH (pH 7.5), 140 mM NaCl, 1% Triton X‐100, 1 mM EDTA, 0.1% sodium deoxycholate) containing cOmplete protease inhibitor cocktail (Merck), transferred to 2 mL screw cap tubes containing 0.5 mm diameter of Zirconia/Silica beads (BioSpec), and homogenized by multi beads shocker (Yasui Kikai) at 2°C, 2700 rpm for 60 s ON and 60 s OFF; 10 cycles. Then, the bottom of the tubes was poked with an 18G needle (Terumo) burned by a gas burner, and the tubes were set into 5 mL collection tubes and centrifuged at 4°C, 200 g for 3 min to collect the homogenized solution. The solution was sonicated with Picoruptor (Diagenode), and centrifuged at 4°C, 20,000 g for 5 min. Then, the sonication debris was removed and the supernatant (chromatin solution) was collected.

The chromatin solution was filled up to 500 μL with ChIP buffer I containing cOmplete protease inhibitor cocktail and incubated with 0.1 μg of antibody for α‐H3 (ab1791; Abcam), α‐H3K4me1 (ab8895; Abcam), α‐H3K4me2 (ab3256; Abcam), and α‐H3K4me3 (ab8580; Abcam), and 0.2 μg for α‐H3K9me2 (MABI0317; Wako). Incubation with antibody was performed overnight at 4°C with rotation. For spike‐in ChIP‐seq, one‐fiftieth of the chromatin DNA from Arabidopsis thaliana compared to that of S. pombe was added to the sonicated chromatin solution, and the resulting spiked chromatin was divided into two tubes; one is for H3 ChIP and the other is for H3K9me2 ChIP. After the incubation, each sample was mixed with 20 μL of Dynabeads M‐280 sheep anti‐mouse IgG (Thermo Fisher) for H3K9me2 antibody or 10 μL of Dynabeads Protein G (Thermo Fisher) for the other antibodies used in this study and incubated at 4°C with rotation. Then, the Dynabeads were washed with each 1 mL of ChIP buffer I containing cOmplete protease inhibitor cocktail, ChIP buffer II (50 mM HEPES‐KOH (pH 7.5), 500 mM NaCl, 1% Triton X‐100, 1 mM EDTA, 0.1% sodium deoxycholate), ChIP buffer III (10 mM Tris–HCl (pH 8.0), 0.25 M LiCl, 0.5% NP‐40, 0.5% sodium deoxycholate, 1 mM EDTA), and TE buffer sequentially, rotating at 4°C for 10 min in each wash step. The washed Dynabeads were resuspended with 100 μL of ChIP elution buffer (50 mM Tris–HCl (pH 8.0), 10 mM EDTA, 1% SDS) and incubated at 65°C for approximately 9 h. Then, the samples were incubated at 37°C with 2 μL of RNaseA (10 mg mL−1; Nippongene) for 30 min and 4 μL of Proteinase K (20 mg mL−1; Thermo Fisher) for 2 h sequentially. Finally, the chromatin solution was purified with the Monarch PCR & DNA cleanup kit (NEB) and eluted in 10 μL.

The libraries for Illumina sequencing were created using the ThruPLEX DNA‐Seq Kit (Clontech) and purified with the SPRIselect beads (Beckman Coulter). The 150 bp paired‐end sequencing was performed with the HiSeqX or NovaSeqX Plus sequencer (Illumina). Two independent biological replicates were performed for each experiment.

Sequenced reads were trimmed using Trimmomatic (Bolger et al. 2014) under the condition; SE, LEADING:30, TRAILING:30, ILLUMINACLIP:TruSeq3‐SE.fa:2:10:10, SLIDINGWINDOW:5:20, MINLEN:30. Trimmed reads were mapped on the S. pombe ASM294v2 genome and the A. thaliana TAIR10 genome (for spike‐in ChIP‐seq) using Bowtie (Langmead et al. 2009). The read count for each transcription unit was calculated using the “coverage” function of BEDTools (Quinlan and Hall 2010). Meta plots were drawn using the ngs.plot program (Shen et al. 2014). For spike‐in ChIP‐seq, the spike‐in control ratio was calculated for each sample by dividing the total mapped reads of S. pombe by those of A. thaliana . The ratio for H3K9me2 was divided by the ratio for H3 to calculate the spike‐in normalized H3K9me2/H3 for each sample. The H3K9me2 signals were multiplied by these spike‐in normalized H3K9me2/H3 ratios as spike‐in normalization.

The HOT‐DOGs were selected by the following criteria: the ratio of H3 signals, log2(RPKM[5 min]/RPKM[0 min]), was less than −0.1 and p‐value was less than 0.05 in four biological replicates, and the ratio of mRNA signals, log2(TPM[5 min]/TPM[0 min]), was more than 1 and p‐value was less than 0.05 in three biological replicates.

The Atf1‐ and Pcr1‐regulated HOT‐DOGs were selected by the following criteria: among the HOT‐DOGs, the ratio of mRNA signals in atf1Δ and pcr1Δ 5 min after heat treatment compared to those in WT, log2(TPM[atf1Δ]/TPM[WT]) and log2(TPM[pcr1Δ]/TPM[WT]), was less than −1 and FDR was less than 0.05 in three biological replicates, respectively.

The heterochromatin genes were selected by the following criteria: the ratio of spike‐in normalized‐H3K9me2 signals in clr4Δ compared to WT before heat treatment, log2(RPKM[clr4Δ]/RPKM[WT]), was less than −2 on average across two biological replicates, and the ratio of mRNA signals in clr4Δ compared to WT before heat treatment, log2(TPM[clr4Δ]/TPM[WT]), was greater than 2 on average across three biological replicates.

4.4. mRNA‐Seq

The total RNA extraction method was described previously (Iyer and Struhl 1996) with some modifications. Briefly, the stored samples were thawed and centrifuged at 4°C, 3000 g for 20 s. The supernatant was removed, and the pellet of those samples was resuspended with 400 μL of lysis buffer (10 mM Tris‐Cl (pH 7.5), 10 mM EDTA, 0.5% SDS) and an equal volume of water‐saturated phenol by vortexing for 3 min, incubated at 65°C for 60 min with occasional vortexing and kept on ice for 10 min. Then, the samples were centrifuged at 4°C, 13,000 g for 10 min, and the aqueous phase of the samples was collected. An equal volume of citric‐acid‐saturated phenol (pH 4.2): chloroform (5:1) was added to the collected aqueous phase solution and vigorously vortexed, and the mixture was centrifuged at 4°C, 15,000 g for 5 min, and the aqueous phase was collected. This operation was repeated once again. Then, the collected aqueous phase was mixed with 0.1 vol of 3 M sodium acetate and 2 vol of ethanol and kept at −80°C for more than 30 min. After the incubation, the samples were centrifuged at 4°C, 15,000 g for 30 min, and washed with 1 mL of 80% ethanol to make a total RNA pellet. Finally, the pellet was dried up and resuspended in 25 μL of RNase‐free water.

600 ng of the total RNA was used to create mRNA‐seq libraries by KAPA mRNA HyperPrep Kit (Kapa Biosystems). Three independent biological replicates were analyzed for each genotype used in this analysis. The libraries were 150 bp paired‐end sequenced by the HiSeqX or NovaSeqX Plus sequencer (Illumina). Three independent biological replicates were performed for each experiment.

Sequenced reads were trimmed using Trimmomatic (Bolger et al. 2014) under the condition; SE, CROP:50, ILLUMINACLIP:TruSeq3‐SE.fa:2:10:10, SLIDINGWINDOW: 5:20, MINLEN:30. Trimmed reads were mapped on the S. pombe ASM294v2 genome using STAR (Dobin et al. 2013) with –outSAMtype, BAM SortedByCoordinate; ‐‐outFilterType, BySJout; ‐‐alignSJoverhangMin, 8; ‐‐alignSJDBoverhangMin, 1; ‐‐quantMode, GeneCounts parameters.

4.5. Cell Culture and Sampling for PRO‐Seq

The details of PRO‐seq were described previously (Mahat et al. 2016). Briefly, when cells reached a density between 2.16 and 3.05 × 106 cells mL−1, untreated (0 min) cells were collected immediately. The flasks for heat‐treated samples were moved from a 30°C water bath to a 37.5°C water bath and incubated for the specified time (2.5 or 5 min) with shaking. After the 5 min incubation, the flasks were returned to a 30°C water bath from a 37.5°C water bath and cultured for the specified time (2.5, 5, or 10 min) with shaking. At each time point, 10 mL of cell culture medium was dispensed into a 15 mL conical tube and was immediately dipped into ice‐cold water. The harvested cells were centrifuged at 4°C, 3000 g for 5 min to remove the supernatant, resuspended in 5 mL of ice‐cold permeabilization buffer (0.5% N‐Lauroylsarcosine sodium salt, 0.5 mM DTT, 20 units RNase inhibitor (TaKaRa) and cOmplete protease inhibitor cocktail) and kept on ice for 20 min. After the incubation, the samples were centrifuged at 4°C, 1000 g for 5 min. The supernatant was discarded by decantation, and the pellet was resuspended in 100 μL of storage buffer (10 mM Tris‐Cl (pH 8.0), 25% glycerol, 5 mM MgCl2, 0.1 mM EDTA and 5 mM DTT). Finally, the samples in the storage buffer were frozen with liquid nitrogen and stored at −80°C.

4.6. PRO‐Seq

PRO‐seq was performed as described previously (Mahat et al. 2016) with some modifications. Briefly, permeabilized cells were thawed on ice and centrifuged at 4°C, 20,000 g for 5 min. The supernatant was discarded, and the pellet was resuspended with 300 μL of transcription buffer (20 mM Tris‐Cl (pH 7.7), 5 mM MgCl2, 200 mM KCl, 2 mM DTT, 60 U of RNase inhibitor (Invitrogen), 0.5% N‐Lauroylsarcosine sodium salt) containing each 3.5 μL of 1 mM biotin‐rCTP (Perkin‐Elmer), 1 mM biotin‐rUTP (Perkin‐Elmer), 1 mM rATP (NEB), and 1 mM rGTP (NEB). The samples were incubated at 30°C for 5 min and placed on ice immediately after the incubation. Then, the samples were centrifuged at 4°C, 12,000 g for 5 min, and the transcription buffer was removed. The pellets were quickly resuspended with 500 μL of Phenol/Chloroform/Isoamyl alcohol (PCI), and then an equal volume of AE buffer (50 mM sodium acetate (pH 5.2), 10 mM EDTA) and 50 μL of 10% SDS were added and mixed by pipetting to rupture cells. The mixtures were incubated at 65°C for 7 min with inverting the tubes every minute and then kept on ice for 5 min. After the incubation, 120 μL of chloroform was added to the samples and mixed by inverting and centrifuged at 4°C, 20,000 g for 5 min. The aqueous phase was mixed with 500 μL of PCI by vortexing and centrifuged at 4°C, 20,000 g for 5 min. This step was repeated once again. Then, the aqueous phase was transferred to a new tube, mixed with 0.1 vol of 3 M sodium acetate and 1 vol of 2‐propanol, and placed at −80°C overnight.

The samples were centrifuged at 4°C, 20,000 g for 30 min, and the supernatant was removed. The pellets were washed with 1 mL of 75% ethanol and dried up and then resuspended with 20 μL of diethylpyrocarbonate (DEPC)‐treated water. The RNA solutions were incubated at 65°C for 40 s and placed on ice for 2 min. 5 μL of 1 N NaOH was added to each sample and incubated on ice for 10 min, and then 25 μL of 1 M Tris‐Cl (pH 6.8) was immediately added for quenching. The solutions were transferred into the P‐30 column (Bio‐rad) and dropped off by centrifugation at 4°C, 1000 g for 4 min. 40 U of RNase inhibitor (TOYOBO) was added to the RNA solutions, filled up to 50 μL with binding buffer (10 mM Tris‐Cl (pH 7.4), 300 mM NaCl, 0.1% Triton X‐100), and mixed with an equal volume of Dynabeads M‐280 streptavidin (Thermo Fisher). The solutions containing biotinylated RNA and streptavidin beads were incubated at room temperature for 20 min with rotation. Then, the beads binding to biotinylated RNA were washed sequentially with each 500 μL of High Salt Wash buffer (50 mM Tris‐Cl (pH 7.4), 2 M NaCl, 0.5% Triton X‐100), binding buffer, and Low Salt Wash buffer (5 mM Tris‐Cl (pH 7.4), 0.1% Triton X‐100). The supernatant was discarded, and the beads were resuspended with 300 μL of TRIzol reagent (Ambion) and incubated at room temperature for 3 min. 60 μL of chloroform was added to the solutions, mixed thoroughly by vortexing, and placed for 3 min. The solutions were centrifuged at 4°C, 20,000 g for 5 min, and the aqueous phase was collected. The purification by TRIzol was repeated once again. Then, the aqueous phase was collected, mixed with 0.1 vol of sodium acetate and 1 vol of 2‐propanol, and kept at −20°C for more than 20 min. After the incubation, the samples were centrifuged at 4°C, 20,000 g for 30 min, and the RNA pellets were washed with 1 mL of 75% ethanol. Then, the RNA pellets were dried up and resuspended with 4 μL of 3′ RNA adaptor solution. The solutions containing RNA and adaptor were incubated at 65°C for 20 s, mixed with 6 μL of ligation mix (16.7% PEG8,000, 1.67 mM ATP, 20 U of RNase inhibitor (TOYOBO), 1 μL of T4 RNA ligase reaction buffer (NEB), 1 μL of T4 RNA ligase I (NEB)), and incubated at 20°C for 4 h.

After the incubation, the ligation products were filled up to 50 μL with the binding buffer, mixed with an equal volume of Dynabeads M‐280 streptavidin, and incubated at room temperature for 20 min with rotation. The beads binding to biotinylated RNA were washed sequentially with each 500 μL of the High Salt Wash buffer, the binding buffer, and the Low Salt Wash buffer. Then, the buffer was removed, and the beads were resuspended with 300 μL of TRIzol reagents and RNA was purified as described above. Then, the aqueous phase was collected and mixed with 0.1 vol of sodium acetate and 1 vol of 2‐propanol. The mixtures were kept at −20°C for 20 min and centrifuged at 4°C, 20,000 g for 30 min to remove the supernatant. The pellets were washed with 1 mL of 75% ethanol, dried up, and resuspended with 10 μL of DEPC‐treated water. The RNA solutions were incubated at 65°C for 20 s and mixed with 10 μL of the reaction mix (2 μL of RppH (NEB), 2 μL of NEBuffer2 (NEB), 20 U of RNase inhibitor (TOYOBO)). The mixtures were incubated at 37°C for 60 min. Then, the RNA solutions were mixed with 80 μL of PNK mix (2.5 μL of PNK (NEB), 10 μL of PNK buffer (NEB), 1.25 mM ATP, 50 U of RNase inhibitor (TOYOBO)) and incubated at 37°C for 60 min. After the incubation, a purification by TRIzol was performed as described above, and the collected aqueous phase was mixed with 0.1 vol of sodium acetate and 1 vol of 2‐propanol and incubated at −80°C for more than 20 min. The RNA solutions were centrifuged at 4°C, 20,000 g for 30 min to remove 2‐propanol and washed with 1 mL of 75% ethanol, and then the RNA pellets were dried up. The pellets were resuspended with 4 μL of 5′ RNA adaptor solution and incubated at 65°C for 20 s. Then, the solutions were mixed with 6 μL of the ligation mix and incubated at 20°C for 4 h.

After the incubation, filling up with the binding buffer, beads addition, beads washing, and purification by TRIzol were performed as described above. Then, the aqueous phase was mixed with 0.1 vol of sodium acetate and 1 vol of 2‐propanol, kept at −20°C for 20 min, and centrifuged at 4°C, 20,000 g for 30 min. The supernatant was discarded, and the RNA pellets were washed with 1 mL of 75% ethanol and dried up. The dried pellets were resuspended by 5 μL of DEPC‐treated water with RNA PCR primer for reverse transcription. The RNA solutions were incubated at 65°C for 5 min, mixed with 5 μL of reverse transcription mix (2 μL of First strand synthesis buffer (Invitrogen), 20 mM DTT, 1.25 mM each of dNTPs, 10 U of RNase inhibitor (TOYOBO), 1 μL of SuperScriptIII RTase (Invitrogen)), and following thermal cycling (45°C for 10 min, 50°C for 40 min, 55°C for 10 min, 70°C for 15 min, sequentially) was performed. 1 μL of the reverse transcription product was mixed with PCR reaction mix (0.2 mM each of dNTPs, 10 μL of Phusion HF buffer (NEB), 0.5 μL of Phusion DNA polymerase (NEB), and PCR primers for Illumina sequencing), and amplified by following thermal cycling (1 cycle of 1st step; 98°C for 30 s, 5 cycles of 2nd step; 98°C for 10 s, 56°C for 30 s and 72°C for 30 s, 6 cycles of 3rd step; 98°C for 10 s, 65°C for 30 s and 72°C for 30 s, 1 cycle of 4th step; 72°C for 5 min). Finally, the DNA sequencing libraries were purified by AMPure beads (Backman) and eluted by 15 μL of MilliQ‐H2O. The 150 bp paired‐end sequencing was performed with the HiSeqX sequencer (Illumina). Two independent biological replicates were performed for each experiment.

Reads were mapped on the S. pombe ASM294v2 genome using Bowtie (Langmead et al. 2009). The read count for each transcription unit was calculated using the “coverage” function of BEDTools (Quinlan and Hall 2010). Meta plots were drawn using the ngs.plot program (Shen et al. 2014).

Author Contributions

Arata Hoshino: conceptualization, formal analysis, investigation, methodology, visualization, writing – original draft, writing – review and editing. Takuya Kajitani: investigation, methodology, resources, writing – review and editing. Masaya Oki: resources, project administration, writing – review and editing. Tetsuji Kakutani: conceptualization, funding acquisition, supervision.Soichi Inagaki: conceptualization, validation, visualization, funding acquisition, supervision, project administration, writing – original draft, writing – review and editing.

Funding

This work was supported by Japan Society for the Promotion of Science, JP20H05913, JP22H02299, JP24K21926, JP25K02252, JP21H04977, JP23H00365; Precursory Research for Embryonic Science and Technology, JPMJPR17Q1; Mitsubishi Foundation, 202110005.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Transcription and histone dynamics during and after heat treatment.

Figure S2: Patterns of the H3 ChIP‐seq and RNA‐seq in atf1Δ and pcr1Δ.

Figure S3: PRO‐seq patterns in heterochromatin genes and HOT‐DOGs.

Figure S4: Histone dynamics and transcription kinetics of HOT‐DOGs during and after heat treatment.

Table S1: HOT‐DOGs list.

Table S2: Heterochromatin genes list.

Table S3: Yeast strains used in this study.

GTC-31-0-s001.pdf (640.6KB, pdf)

Acknowledgments

We thank Yota Murakami for the fission yeast strains and helpful discussion, Jun‐ichi Nakayama for helpful discussion, and Mami Hirayama and Mayumi Takahashi for technical assistance. The computations were partially performed on the NIG supercomputer at the National Institute of Genetics, Japan. This study was supported by grants from Japan Science and Technology Agency (JST) PRESTO (no. JPMJPR17Q1 to S.I.), Japan Society for the Promotion of Science (JSPS) (nos. JP20H05913, JP22H02299, JP24K21926, and JP25K02252 to S.I., JP21H04977 and JP23H00365 to T.K.), and Mitsubishi Foundation (no. 202110005 to S.I.).

Hoshino, A. , Kajitani T., Oki M., Kakutani T., and Inagaki S.. 2026. “A Temperature Increase Induces Atf1/Pcr1‐Dependent Rapid Depletion of Histones in Transcriptionally Activated Gene Bodies in Fission Yeast.” Genes to Cells 31, no. 2: e70092. 10.1111/gtc.70092.

Transmitting Editor: Hitoshi Kurumizaka

Contributor Information

Tetsuji Kakutani, Email: tkak@bs.s.u-tokyo.ac.jp.

Soichi Inagaki, Email: soinagak@bs.s.u-tokyo.ac.jp.

Data Availability Statement

The raw and processed next‐generation sequencing data have been deposited in NCBI's Gene Expression Omnibus (GEO) and are accessible through GEO Series accession numbers GSE265755.

References

  1. Allshire, R. C. , and Madhani H. D.. 2018. “Ten Principles of Heterochromatin Formation and Function.” Nature Reviews. Molecular Cell Biology 19: 229–244. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Bae, H. J. , Dubarry M., Jeon J., et al. 2020. “The Set1 N‐Terminal Domain and Swd2 Interact With RNA Polymerase II CTD to Recruit COMPASS.” Nature Communications 11: 1–10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bähler, J. , Wu J. Q., Longtine M. S., et al. 1998. “Heterologous Modules for Efficient and Versatile PCR‐Based Gene Targeting in Schizosaccharomyces pombe .” Yeast 14: 943–951. [DOI] [PubMed] [Google Scholar]
  4. Bolger, A. M. , Lohse M., and Usadel B.. 2014. “Trimmomatic: A Flexible Trimmer for Illumina Sequence Data.” Bioinformatics 30: 2114–2120. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Brownell, J. E. , Zhou J., Ranalli T., et al. 1996. “Tetrahymena Histone Acetyltransferase A: A Homolog to Yeast Gcn5p Linking Histone Acetylation to Gene Activation.” Cell 84: 843–851. [DOI] [PubMed] [Google Scholar]
  6. Chen, D. , Toone W. M., Mata J., et al. 2003. “Global Transcriptional Responses of Fission Yeast to Environmental Stress.” Molecular Biology of the Cell 14: 214–229. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Cortazar, M. A. , Sheridan R. M., Erickson B., et al. 2019. “Control of RNA Pol II Speed by PNUTS‐PP1 and Spt5 Dephosphorylation Facilitates Termination by a “Sitting Duck Torpedo” Mechanism.” Molecular Cell 76: 896–908.e4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Deal, R. B. , Henikoff J. G., and Henikoff S.. 2010. “Genome‐Wide Kinetics of Nucleosome Turnover Determined by Metabolic Labeling of Histones.” Science 328: 1161–1164. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Dobin, A. , Davis C. A., Schlesinger F., et al. 2013. “STAR: Ultrafast Universal RNA‐Seq Aligner.” Bioinformatics 29: 15–21. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Ebmeier, C. C. , Erickson B., Allen B. L., et al. 2017. “Human TFIIH Kinase CDK7 Regulates Transcription‐Associated Chromatin Modifications.” Cell Reports 20: 1173–1186. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Eshaghi, M. , Lee J. H., Zhu L., et al. 2010. “Genomic Binding Profiling of the Fission Yeast Stress‐Activated MAPK Sty1 and the bZIP Transcriptional Activator Atf1 in Response to H2O2 .” PLoS One 5: e11620. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Fong, N. , Brannan K., Erickson B., et al. 2015. “Effects of Transcription Elongation Rate and Xrn2 Exonuclease Activity on RNA Polymerase II Termination Suggest Widespread Kinetic Competition.” Molecular Cell 60: 256–267. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Haldar, S. , Saini A., Nanda J. S., Saini S., and Singh J.. 2011. “Role of Swi6/HP1 Self‐Association‐Mediated Recruitment of Clr4/Suv39 in Establishment and Maintenance of Heterochromatin in Fission Yeast.” Journal of Biological Chemistry 286: 9308–9320. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Hall, I. M. , Shankaranarayana G. D., Noma K.‐I., Ayoub N., Cohen A., and Grewal S. I. S.. 2002. “Establishment and Maintenance of a Heterochromatin Domain.” Science 297: 2232–2237. [DOI] [PubMed] [Google Scholar]
  15. Iglesias, N. , Paulo J. A., Tatarakis A., et al. 2020. “Native Chromatin Proteomics Reveals a Role for Specific Nucleoporins in Heterochromatin Organization and Maintenance.” Molecular Cell 77: 51–66.e8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Inagaki, S. , Takahashi M., Takashima K., Oya S., and Kakutani T.. 2021. “Chromatin‐Based Mechanisms to Coordinate Convergent Overlapping Transcription.” Nature Plants 7: 295–302. [DOI] [PubMed] [Google Scholar]
  17. Iyer, V. , and Struhl K.. 1996. “Absolute mRNA Levels and Transcriptional Initiation Rates in Saccharomyces cerevisiae .” Proceedings of the National Academy of Sciences of the United States of America 93: 5208–5212. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Izban, M. G. , and Luse D. S.. 1992. “Factor‐Stimulated RNA Polymerase II Transcribes at Physiological Elongation Rates on Naked DNA but Very Poorly on Chromatin Templates.” Journal of Biological Chemistry 267: 13647–13655. [PubMed] [Google Scholar]
  19. Janssen, A. , Colmenares S. U., and Karpen G. H.. 2018. “Heterochromatin: Guardian of the Genome.” Annual Review of Cell and Developmental Biology 34: 265–288. [DOI] [PubMed] [Google Scholar]
  20. Jia, S. , Noma K.‐I., and Grewal S. I. S.. 2004. “RNAi‐Independent Heterochromatin Nucleation by the Stress‐Activated ATF/CREB Family Proteins.” Science 304: 1971–1976. [DOI] [PubMed] [Google Scholar]
  21. Jih, G. , Iglesias N., Currie M. A., et al. 2017. “Unique Roles for Histone H3K9me States in RNAi and Heritable Silencing of Transcription.” Nature 547: 463–467. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Kajitani, T. , Kato H., Chikashige Y., et al. 2017. “Ser7 of RNAPII‐CTD Facilitates Heterochromatin Formation by Linking ncRNA to RNAi.” Proceedings of the National Academy of Sciences of the United States of America 114: E11208–E11217. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Kon, N. , Krawchuk M. D., Warren B. G., Smith G. R., and Wahls W. P.. 1997. “Transcription Factor Mts1/Mts2 (Atf1/Pcr1, Gad7/Pcr1) Activates the M26 Meiotic Recombination Hotspot in Schizosaccharomyces pombe .” Proceedings of the National Academy of Sciences of the United States of America 94: 13765–13770. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Kristell, C. , Orzechowski Westholm J., Olsson I., Ronne H., Komorowski J., and Bjerling P.. 2010. “Nitrogen Depletion in the Fission Yeast Schizosaccharomyces pombe Causes Nucleosome Loss in Both Promoters and Coding Regions of Activated Genes.” Genome Research 20: 361–371. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Kujirai, T. , and Kurumizaka H.. 2020. “Transcription Through the Nucleosome.” Current Opinion in Structural Biology 61: 42–49. [DOI] [PubMed] [Google Scholar]
  26. Langmead, B. , Trapnell C., Pop M., and Salzberg S. L.. 2009. “Ultrafast and Memory‐Efficient Alignment of Short DNA Sequences to the Human Genome.” Genome Biology 10: R25. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Lawrence, C. L. , Maekawa H., Worthington J. L., Reiter W., Wilkinson C. R. M., and Jones N.. 2007. “Regulation of Schizosaccharomyces pombe Atf1 Protein Levels by Sty1‐Mediated Phosphorylation and Heterodimerization With Pcr1.” Journal of Biological Chemistry 282: 5160–5170. [DOI] [PubMed] [Google Scholar]
  28. Leng, X. , Ivanov M., Kindgren P., et al. 2020. “Organismal Benefits of Transcription Speed Control at Gene Boundaries.” EMBO Reports 21: e49315. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Li, B. , Carey M., and Workman J. L.. 2007. “The Role of Chromatin During Transcription.” Cell 128: 707–719. [DOI] [PubMed] [Google Scholar]
  30. Mahat, D. B. , Kwak H., Booth G. T., et al. 2016. “Base‐Pair‐Resolution Genome‐Wide Mapping of Active RNA Polymerases Using Precision Nuclear Run‐On (PRO‐Seq).” Nature Protocols 11: 1455–1476. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Mori, S. , Oya S., Takahashi M., Takashima K., Inagaki S., and Kakutani T.. 2023. “Cotranscriptional Demethylation Induces Global Loss of H3K4me2 From Active Genes in Arabidopsis.” EMBO Journal 42: e113798. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Motamedi, M. R. , Hong E.‐J. E., Li X., et al. 2008. “HP1 Proteins Form Distinct Complexes and Mediate Heterochromatic Gene Silencing by Nonoverlapping Mechanisms.” Molecular Cell 32: 778–790. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Nakayama, J. , Rice J. C., Strahl B. D., Allis C. D., and Grewal S. I.. 2001. “Role of Histone H3 Lysine 9 Methylation in Epigenetic Control of Heterochromatin Assembly.” Science 292: 110–113. [DOI] [PubMed] [Google Scholar]
  34. Ng, H. H. , Robert F., Young R. A., and Struhl K.. 2003. “Targeted Recruitment of Set1 Histone Methylase by Elongating Pol II Provides a Localized Mark and Memory of Recent Transcriptional Activity.” Molecular Cell 11: 709–719. [DOI] [PubMed] [Google Scholar]
  35. Nightingale, K. P. , Wellinger R. E., Sogo J. M., and Becker P. B.. 1998. “Histone Acetylation Facilitates RNA Polymerase II Transcription of the Drosophila hsp26 Gene in Chromatin.” EMBO Journal 17: 2865–2876. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Noma, K. , Allis C. D., and Grewal S. I.. 2001. “Transitions in Distinct Histone H3 Methylation Patterns at the Heterochromatin Domain Boundaries.” Science 293: 1150–1155. [DOI] [PubMed] [Google Scholar]
  37. Oya, S. , Takahashi M., Takashima K., Kakutani T., and Inagaki S.. 2022. “Transcription‐Coupled and Epigenome‐Encoded Mechanisms Direct H3K4 Methylation.” Nature Communications 13: 4521. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Padeken, J. , Methot S. P., and Gasser S. M.. 2022. “Establishment of H3K9‐Methylated Heterochromatin and Its Functions in Tissue Differentiation and Maintenance.” Nature Reviews. Molecular Cell Biology 23: 623–640. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Paulo, E. , García‐Santamarina S., Calvo I. A., et al. 2014. “A Genetic Approach to Study H2O2 Scavenging in Fission Yeast—Distinct Roles of Peroxiredoxin and Catalase: H2O2 Scavenging in Fission Yeast.” Molecular Microbiology 92: 246–257. [DOI] [PubMed] [Google Scholar]
  40. Petesch, S. J. , and Lis J. T.. 2008. “Rapid, Transcription‐Independent Loss of Nucleosomes Over a Large Chromatin Domain at Hsp70 Loci.” Cell 134: 74–84. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Petesch, S. J. , and Lis J. T.. 2012. “Overcoming the Nucleosome Barrier During Transcript Elongation.” Trends in Genetics 28: 285–294. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Quinlan, A. R. , and Hall I. M.. 2010. “BEDTools: A Flexible Suite of Utilities for Comparing Genomic Features.” Bioinformatics 26: 841–842. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Quinn, J. , Findlay V. J., Dawson K., et al. 2002. “Distinct Regulatory Proteins Control the Graded Transcriptional Response to Increasing H(2)O(2) Levels in Fission Yeast Schizosaccharomyces pombe .” Molecular Biology of the Cell 13: 805–816. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Salat‐Canela, C. , Paulo E., Sánchez‐Mir L., et al. 2017. “Deciphering the Role of the Signal‐ and Sty1 Kinase‐Dependent Phosphorylation of the Stress‐Responsive Transcription Factor Atf1 on Gene Activation.” Journal of Biological Chemistry 292: 13635–13644. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Sansó, M. , Gogol M., Ayté J., Seidel C., and Hidalgo E.. 2008. “Transcription Factors Pcr1 and Atf1 Have Distinct Roles in Stress‐ and Sty1‐Dependent Gene Regulation.” Eukaryotic Cell 7: 826–835. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Sansó, M. , Vargas‐Pérez I., Quintales L., Antequera F., Ayté J., and Hidalgo E.. 2011. “Gcn5 Facilitates Pol II Progression, Rather Than Recruitment to Nucleosome‐Depleted Stress Promoters, in Schizosaccharomyces pombe .” Nucleic Acids Research 39: 6369–6379. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Shen, L. , Shao N., Liu X., and Nestler E.. 2014. “Ngs.Plot: Quick Mining and Visualization of Next‐Generation Sequencing Data by Integrating Genomic Databases.” BMC Genomics 15: 284. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Shiozaki, K. , and Russell P.. 1996. “Conjugation, Meiosis, and the Osmotic Stress Response Are Regulated by Spc1 Kinase Through Atf1 Transcription Factor in Fission Yeast.” Genes & Development 10: 2276–2288. [DOI] [PubMed] [Google Scholar]
  49. Skribbe, M. , Soneson C., Stadler M. B., et al. 2025. “A Comprehensive Schizosaccharomyces pombe Atlas of Physical Transcription Factor Interactions With Proteins and Chromatin.” Molecular Cell 85: 1426–1444.e8. [DOI] [PubMed] [Google Scholar]
  50. Sorida, M. , Hirauchi T., Ishizaki H., et al. 2019. “Regulation of Ectopic Heterochromatin‐Mediated Epigenetic Diversification by the JmjC Family Protein Epe1.” PLoS Genetics 15: e1008129. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Sugiyama, T. , Cam H. P., Sugiyama R., et al. 2007. “SHREC, an Effector Complex for Heterochromatic Transcriptional Silencing.” Cell 128: 491–504. [DOI] [PubMed] [Google Scholar]
  52. Sun, L. , Liu L., Song C., Wang Y., and Jin Q.‐W.. 2024. “Heat Stress‐Induced Activation of MAPK Pathway Attenuates Atf1‐Dependent Epigenetic Inheritance of Heterochromatin in Fission Yeast.” eLife 13: e90525. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Teves, S. S. , and Henikoff S.. 2011. “Heat Shock Reduces Stalled RNA Polymerase II and Nucleosome Turnover Genome‐Wide.” Genes & Development 25: 2387–2397. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Teves, S. S. , Weber C. M., and Henikoff S.. 2014. “Transcribing Through the Nucleosome.” Trends in Biochemical Sciences 39: 577–586. [DOI] [PubMed] [Google Scholar]
  55. Vega, M. , Barrios R., Fraile R., et al. 2023. “Topoisomerase 1 Facilitates Nucleosome Reassembly at Stress Genes During Recovery.” Nucleic Acids Research 51: 12161–12173. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Wang, X. , and Moazed D.. 2017. “DNA Sequence‐Dependent Epigenetic Inheritance of Gene Silencing and Histone H3K9 Methylation.” Science 356: 88–91. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Wilkinson, M. G. , Samuels M., Takeda T., et al. 1996. “The Atf1 Transcription Factor Is a Target for the Sty1 Stress‐Activated MAP Kinase Pathway in Fission Yeast.” Genes & Development 10: 2289–2301. [DOI] [PubMed] [Google Scholar]
  58. Woolcock, K. J. , Stunnenberg R., Gaidatzis D., et al. 2012. “RNAi Keeps Atf1‐Bound Stress Response Genes in Check at Nuclear Pores.” Genes & Development 26: 683–692. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Zhang, H. , Lang Z., and Zhu J.‐K.. 2018. “Dynamics and Function of DNA Methylation in Plants.” Nature Reviews. Molecular Cell Biology 19: 489–506. [DOI] [PubMed] [Google Scholar]
  60. Zhang, K. , Mosch K., Fischle W., and Grewal S. I. S.. 2008. “Roles of the Clr4 Methyltransferase Complex in Nucleation, Spreading and Maintenance of Heterochromatin.” Nature Structural & Molecular Biology 15: 381–388. [DOI] [PubMed] [Google Scholar]
  61. Zhao, J. , Herrera‐Diaz J., and Gross D. S.. 2005. “Domain‐Wide Displacement of Histones by Activated Heat Shock Factor Occurs Independently of Swi/Snf and Is Not Correlated With RNA Polymerase II Density.” Molecular and Cellular Biology 25: 8985–8999. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Figure S1: Transcription and histone dynamics during and after heat treatment.

Figure S2: Patterns of the H3 ChIP‐seq and RNA‐seq in atf1Δ and pcr1Δ.

Figure S3: PRO‐seq patterns in heterochromatin genes and HOT‐DOGs.

Figure S4: Histone dynamics and transcription kinetics of HOT‐DOGs during and after heat treatment.

Table S1: HOT‐DOGs list.

Table S2: Heterochromatin genes list.

Table S3: Yeast strains used in this study.

GTC-31-0-s001.pdf (640.6KB, pdf)

Data Availability Statement

The raw and processed next‐generation sequencing data have been deposited in NCBI's Gene Expression Omnibus (GEO) and are accessible through GEO Series accession numbers GSE265755.

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