Decision letter: Histone deacetylase 1 maintains lineage integrity through histone acetylome refinement during early embryogenesis
Article Figures and data Abstract Editor's evaluation Introduction Results Discussion Materials and methods Data availability References Decision letter Author response Article and author information Metrics Abstract Histone acetylation is a pivotal epigenetic modification that controls chromatin structure and regulates gene expression. It plays an essential role in modulating zygotic transcription and cell lineage specification of developing embryos. While the outcomes of many inductive signals have been described to require enzymatic activities of histone acetyltransferases and deacetylases (HDACs), the mechanisms by which HDACs confine the utilization of the zygotic genome remain to be elucidated. Here, we show that histone deacetylase 1 (Hdac1) progressively binds to the zygotic genome from mid-blastula and onward. The recruitment of Hdac1 to the genome at blastula is instructed maternally. Cis-regulatory modules (CRMs) bound by Hdac1 possess epigenetic signatures underlying distinct functions. We highlight a dual function model of Hdac1 where Hdac1 not only represses gene expression by sustaining a histone hypoacetylation state on inactive chromatin, but also maintains gene expression through participating in dynamic histone acetylation–deacetylation cycles on active chromatin. As a result, Hdac1 maintains differential histone acetylation states of bound CRMs between different germ layers and reinforces the transcriptional program underlying cell lineage identities, both in time and space. Taken together, our study reveals a comprehensive role for Hdac1 during early vertebrate embryogenesis. Editor's evaluation In this revised and important study, the authors investigate the roles of histone deacetylases in the spatial epigenetic regulation of zygotic gene expression during embryo gastrulation. They provide convincing evidence for HDAC1 binding to genes around the timing of large-scale genome activation, and that inhibition of histone acetylation blocks gastrulation, blurring cell lineage integrity, tied to both positive and negative regulatory effects on transcription in space and time. The research reveals new insight on the role of histone acetylation-deacetylation in dynamics in epigenetic control of gene expression and cell fate determination during early tissue patterning in embryogenesis. https://doi.org/10.7554/eLife.79380.sa0 Decision letter Reviews on Sciety eLife's review process Introduction A fundamental question in early development is the mechanism of zygotic genome activation (ZGA), which requires the degradation of maternal mRNAs and the activation of embryonic transcription (Blitz and Cho, 2021, Tadros and Lipshitz, 2009). During ZGA, the embryonic genome undergoes a dramatic reprogramming of gene expression, which is also accompanied by remodeling of the embryonic epigenome. Post-translational modifications to histones are a major epigenetic regulation influencing chromatin structure and thus play a central role in ZGA. Histone acetylation appears during the onset of both minor and major ZGA waves in many species. In Drosophila, histone acetylation occurs at mitotic cycle 8 on a few early zygotic genes (Li et al., 2014). miR430, the first zygotically active gene, is marked by H3K27ac in 64-cell staged zebrafish embryos (Chan et al., 2019). Genome-wide H3K27ac is detected at mid-blastula shortly after the onset of ZGA in Xenopus (Gupta et al., 2014). In mice, the zygotic genome is increasingly marked by H3K27ac from immature and metaphase II oocytes to 2-cell-stage embryos (Dahl et al., 2016). Despite these findings, there remain several major questions. How is the interplay of enzymes regulating histone acetylation deployed in developing embryos? What is the role of observed histone acetylation on gene expression? How are the spatial and temporal patterns of histone acetylation established during ZGA? Histone acetylation occurs on the ε-amino group of the lysine residues within N-terminal tails of all four core histones (Inoue and Fujimoto, 1969; Seto and Yoshida, 2014). Acetylation is a reversible process that is directly catalyzed by opposing activities of histone acetyltransferases (HATs) and histone deacetylases (HDACs). In addition, HATs and HDACs can also regulate the acetylation of lysine residues on non-histone proteins (Choudhary et al., 2009). Histone acetylation is often associated with active gene transcription because the acyl groups neutralize the positive charge on the lysine residues, thereby reducing the affinity of histones to DNA (Wang et al., 2000; Anderson et al., 2001); it also serves as a binding platform for bromodomain (BRD) proteins which scaffold and stimulate the transcriptional machinery (Hassan et al., 2007; Filippakopoulos et al., 2012). The balance between HATs and HDACs directly shapes histone acetylation landscapes and subsequently affects transcriptomes. HDACs are critical epigenetic regulators because they reset chromatin states by returning acetylated lysine residues on histones to the basal state, which can subsequently be subjected to alternative modifications such as methylation. HDACs are grouped into four classes based on phylogenetic conservation. Class I (HDAC1, 2, 3, 8), Class II (HDAC4, 5, 6, 7, 9, 10), and Class IV (HDAC11) HDACs are zinc dependent and are related to yeast Rpd3, Had1, and Hos3, respectively; Class III (SIRT1, 2, 3, 4, 5, 6, 7) HDACs, also known as Sirtuins, are NAD+ dependent and are related to yeast Sir2 (Gregoretti et al., 2004; Milazzo et al., 2020). HDACs are well-characterized negative regulators of gene expression during development. For example, Hdac1 silences homeotic genes in cooperation with Polycomb group repressors in Drosophila (Chang et al., 2001). In zebrafish, Hdac1 represses Notch targets during neurogenesis (Cunliffe, 2004; Yamaguchi et al., 2005). In Xenopus, HDAC activity suppresses Vegt-induced ectopic mesoderm in ectoderm lineages (Gao et al., 2016), represses multi-lineage marker genes at blastula (Rao and LaBonne, 2018), and desensitizes dorsal Wnt signaling at late blastula (Esmaeili et al., 2020). Conversely, HDACs can also positively regulate gene expression. For instance, inhibition of HDAC activities rapidly down-regulates some genes in yeast, suggesting an activator function of HDACs (Bernstein et al., 2000). Genetic deletions or pharmacological application of HDAC inhibitors in cell lines results in both up- and down-regulation of genes (Reid et al., 2005; Zupkovitz et al., 2006; Meganathan et al., 2015). Furthermore, genome-wide studies showed that HDACs occupy genomic loci of active genes, and their binding correlates with gene activities (Kurdistani et al., 2002; Wang et al., 2002; Wang et al., 2009; Kidder and Palmer, 2012). These seemingly opposing functions of HDACs raise an important question as to the exact roles of HDACs on chromatin states and transcriptomes in developing embryos. In this study, we focus on the role of Hdac1 in regulating the zygotic epigenome and transcriptome during Xenopus germ-layer specification coinciding with ZGA. Current evidence in Xenopus as well as in other non-mammalian systems suggests that the early embryonic genome is rather naive and that major chromatin modifications occur at or after ZGA (Bonn et al., 2012; Vastenhouw et al., 2010; Gupta et al., 2014; van Heeringen et al., 2014; Hontelez et al., 2015). Thus, the system allows us to probe the earliest establishment of histone acetylation and dissect the link between actions of Hdac1, the zygotic histone acetylome, and zygotic transcriptome during the first cell lineage segregation event. Here, we report that the major Hdac1 binding to the embryonic genome occurs at blastula; the binding of Hdac1 during this stage is dependent on maternal factors. We highlight a dual function model for Hdac1. First, Hdac1 keeps inactive chromatin free of histone acetylation, preventing gene misactivation in respective germ layers. Second, Hdac1 participates in dynamic histone acetylation–deacetylation cycles on active chromatin, sustaining the expression of genes that are enriched in respective germ layers. Taken together, our study reveals a coordinated spatial and temporal regulation by Hdac1 during ZGA. Results Hdac1 binds to the genome progressively during blastula and onward To identify the major functional candidates of HDACs during the early Xenopus embryogenesis, we examined the temporal RNA expression profiles (Owens et al., 2016) of HDAC family members (Class I, II, and III HDACs) from the zygote to the beginning of the neurula stage in Xenopus tropicalis. The RNA expression level of hdac1 is the highest among all HDACs examined, followed by hdac2 (Figure 1—figure supplement 1A). Both Hdac1 and Hdac2 proteins are present in the unfertilized egg to the mid gastrula stage, and the overall expression levels of Hdac1 and Hdac2 are relatively constant during this period (Figure 1A). These data reveal that Hdac1/2 are the major maternally endowed HDACs functioning during this window of development. Figure 1 with 1 supplement see all Download asset Open asset Hdac1 binds to the genome gradually during early Xenopus development. (A) Western blot analyses showing Hdac1 and Hdac2 proteins over a time course of early development. β-Tubulin is used as a loading control. (B) Venn diagram comparing Hdac1 irreproducibility discovery rate (IDR) peaks among examined stages. The sums of peaks in st8 and st9 are smaller due to instances where more than one peak from st8 or st9 overlaps the same single st10.5 peak. (C) Clustered heatmap showing Hdac1 ChIP-seq signals at each stage over a window of 5 kb centered on the summit of all Hdac1 IDR peaks in descending order. (D) The expression levels (TPM, transcripts per million) of genes associated with Hdac1 clusters at different developmental periods. (E) Distributions of Hdac1 IDR peaks at each stage across five defined genomic features. The promoter is defined as −1 kb to +100 bp from TSS (transcription start site) while the TTS (transcription termination site) is defined as −100 bp to +1 kb from TTS. (F) Distributions of Hdac1 ChIP-seq signals within the intervals of 5 kb upstream of gene model 5′ ends, gene bodies (normalized for length), and 5 kb downstream of gene model 3′ ends at each stage. The signal of st9 input DNA ChIP-seq is used as a negative control. Y-axis values represent reads quantified by bins per million (BPM) at a bin size of 50 bp. (G) ChIP-qPCR showing Hdac1 enrichment on selected genomic regions (nine positive regions: alkbh2, fgf5, foxi4.2, gdnf, hhex, miR428a, not, snai1, and sp8; two negative regions: hspa4 and klf11) at indicated stages of early embryogenesis. Figure 1—source data 1 Western blot analyses showing Hdac1 and Hdac2 proteins during Xenopus embryogenesis. https://cdn.elifesciences.org/articles/79380/elife-79380-fig1-data1-v2.zip Download elife-79380-fig1-data1-v2.zip Since Hdac1 modulates various aspects of transcriptional regulation and the chromatin landscape, we examined genome-wide Hdac1-binding patterns during early germ-layer development (Figure 1—figure supplement 1B) using chromatin immunoprecipitation (ChIP) assays followed by sequencing (ChIP-seq). A set of high-confidence peaks at each stage were obtained using irreproducibility discovery rate (IDR) analysis (Li et al., 2011) from two biologically independent samples (Figure 1—figure supplement 1C). Hdac1 binds to 1340 regions at the mid-blastula (st8), 5136 regions at the late blastula (st9), and 22,681 regions at the early gastrula (st10.5) stages (Figure 1B). Overall, a minority of Hdac1 peaks are unique to each of the blastula stages (Clusters a and b) and a majority of Hdac1 peaks are present across multiple stages (Cluster c) and at the early gastrula stage (Cluster d) (Figure 1C, Figure 1—figure supplement 1D). The expression levels of zygotic genes associated with Cluster c Hdac1 peaks are higher than those associated with Clusters a, b, and d, suggesting that the genes associated with sustained Hdac1 binding are more active during gastrulation (Figure 1D). Similarly, Hdac2 gradually accumulates on the embryonic genome (Figure 1—figure supplement 1E). Interestingly, 99% of Hdac1 peaks at stage 9 and 97% of Hdac1 peaks at stage 10.5 overlap with Hdac2 peaks (Figure 1—figure supplement 1F, G). When the levels of Hdac2 peaks associated with Hdac1 were compared to Hdac2 peaks , Hdac1 peaks display a significantly higher Hdac2 peak enrichment (Figure 1—figure supplement 1H). Together, we revealed that Hdac1/2 are progressively directed to the genome during early development. Since a majority of Hdac1-bound peaks are similarly bound by Hdac2, we focus on Hdac1 peaks in subsequent analyses. To investigate the differences of Hdac1 genomic occupancy across early stages, we examined various genomic features of regions bound by Hdac1 at each stage. The majority of Hdac1 peaks are found in either intergenic or intronic regions; a minor fraction of Hdac1 peaks is present within exons or transcriptional termination sites. A notable observation is the increased Hdac1 binding to more promoter regions over developmental times (Figure 1E). We then analyzed the distribution of Hdac1 across three stages along the gene bodies of genes bound by Hdac1. A higher enrichment of Hdac1 binding is located near the promoters of genes as development proceeds (Figure 1F). Importantly, the timing of Hdac1-binding accumulation coincides with the mid-blastula stage (st8) (Figure 1G), suggesting that Hdac1 is involved in the epigenetic regulation of these genes during ZGA. Blastula Hdac1 binding is maternally instructed Since Hdac1 binds progressively to the genome during ZGA, we investigated the contribution of zygotic factors to the recruitment of Hdac1 at the late blastula stage. We injected α-amanitin, which blocks both transcription initiation and elongation (Chafin et al., 1995), to block embryonic transcription (Figure 2—figure supplement 1A). A high Pearson correlation between α-amanitin-injected and uninjected embryos is observed genome-wide or at all Hdac1 IDR peaks (Figure 2A), which is higher than the Pearson correlation representing the batch effect when compared to a different batch of same staged embryos (WT) (Figure 2—figure supplement 1B). There is no significant signal difference between α-amanitin-injected and uninjected embryos at all Hdac1 IDR peaks (Figure 2—figure supplement 1C, H). We conclude that Hdac1 binding is independent of zygotic transcription at the late blastula stage, suggesting the importance of maternal factors in Hdac1 recruitment. Figure 2 with 1 supplement see all Download asset Open asset Maternal factors instruct Hdac1 recruitment during blastula stages. (A) Pairwise Pearson correlation analyses comparing ChIP-seq signals of st9 Hdac1 irreproducibility discovery rate (IDR) peaks between α-amanitin-injected and uninjected embryos across the genome (left) and among Hdac1 IDR peaks (right). (B) Motif analyses of st9 Hdac1 peaks (500 bp centered on IDR peak summit). Motif sequence to the corresponding factor is retrieved from JASPAR. Hey1 is an example of TF motif with low significance. (C) Clustered heatmap depicting st9 Foxh1 and Sox3 ChIP-seq signals in a window of 10 kb centered on st9 Hdac1 IDR peaks with descending order. (D) ChIP-seq signal enrichment of Hdac1, Foxh1, and Sox3 within the intervals of 5 kb upstream of gene model 5′ ends, gene bodies (normalized for length), and 5 kb downstream of gene model 3′ ends of st9 Hdac1 associated genes. The signal of st9 input DNA ChIP-seq is used as a negative control. Y-axis values represent reads quantified by bins per million (BPM) at a bin size of 50 bp. (E) St9 sequential ChIP-qPCR analyses for Foxh1 and Hdac1 co-bound regions and Sox3 and Hdac1 co-bound regions. anti-HA is used as a negative control. The error bars represent the variation from two technical replicates. (F) ChIP-qPCR analysis of Hdac1 peaks that are also Foxh1/Sox3 co-bound after Foxh1 and/or Sox3 depletion. Genomic loci associated with klf11 (no Hdac1 signals), erfl and insm2 (Hdac1 signals without Foxh1 or Sox3 signals) are negative controls. The error bars represent the variation from two technical replicates. Student’s t-test is used to calculate p-values over Hdac1 enrichment of uninjected embryos. * is for p< 0.05** is for p< 0.01. To identify maternal factors that facilitate the recruitment of Hdac1 to the genome, we performed a de novo motif search of the DNA sequences of 5136 st9 Hdac1peaks. Sox, Foxh1, and Pou motifs are found to be the most frequent maternal TF motifs (Figure 2B, Supplementary file 1). We thus compared the genomic binding profiles of Hdac1 to two maternal TFs, Foxh1 (Charney et al., 2017) and Sox3. A majority of Hdac1-bound regions (Cluster 1) overlaps with both Foxh1- and Sox3-bound regions while only a small fraction of Hdac1-bound regions (Cluster 2) overlaps the binding of either Foxh1, or Sox3, or neither (Figure 2C). More than 80% of Hdac1 peaks overlap with Foxh1 or Sox3 peaks (Figure 2—figure supplement 1D, E). A positive correlation between Hdac1 binding and Foxh1/Sox3 binding is observed at all Hdac1 IDR peaks (Figure 2—figure supplement 1F, G). We noted frequent overlapping binding of Hdac1 with Foxh1/Sox3 (Figure 2—figure supplement 1I, J), and highly enriched signals of Hdac1, Foxh1, and Sox3 present around promoters of genes (Figure 2D). All these observations suggest a role for Foxh1 and Sox3 in Hdac1 recruitment. We confirmed the co-occupancy of Hdac1 with each of Foxh1 and Sox3 TFs on the same DNA molecules using sequential ChIP-qPCR (Figure 2E, Figure 2—figure supplement 1K, L). Depletion of Foxh1, Sox3, or both TFs by morpholino injections showed a reduced binding of Hdac1 around Foxh1/Sox3 co-occupied genomic regions (Figure 2F). Hence, we propose that Foxh1 and Sox3 maternal TFs facilitate Hdac1 recruitment during ZGA. Hdac1 binds to genomic regions with distinct epigenetic signatures To further characterize regions bound by Hdac1 across early germ-layer development, we examined epigenetic signatures (Gupta et al., 2014; Hontelez et al., 2015; Charney et al., 2017) on Hdac1 peaks across various stages. Ep300 binding (a HAT), which catalyzes the acetylation of histone, is observed in Hdac1 peaks (Clusters b–d) from the late blastula and onward where RNA polymerase II signals also emerge (Figure 3—figure supplement 1A). This indicates that Hdac1 and Ep300 share similar binding profiles on many transcriptionally active genes. We next surveyed several histone acetylation modifications in regions bound by Hdac1 (Figure 3A). Consistent with the overlapping binding of Ep300, Hdac1 peaks display signals of H3K9ac (Clusters c and d), H3K18ac (Clusters b–d), H3K27ac (Clusters b–d), and pan-H3 lysine acetylation (pan-H3Kac) (Clusters b–d). We then examined several histone methylation modifications that are associated with gene activation (Figure 3—figure supplement 1B). H3K4me1, a primed enhancer mark (Creyghton et al., 2010), and H3K4me3, an active promoter mark (Heintzman et al., 2007), display signals at Hdac1-bound regions (Clusters c and d); H3K36me3, a transcription elongation mark (Kolasinska-Zwierz et al., 2009), displays minimal signals at any Hdac1-bound regions. These observations reveal that Hdac1 binds to genomic regions with active epigenetic signatures. Figure 3 with 1 supplement see all Download asset Open asset Hdac1 binds to cis-regulatory modules (CRMs) containing functionally distinct histone modifications. Clustered heatmaps showing signals from several stages of various (A) histone acetylation marks and (B) repressive histone methylation marks on Hdac1 peaks. Each cluster corresponds to the same regions present in Figure 1C. (C) Venn diagram illustrating Hdac1 peaks overlapping with H3K27me3 and H3K27ac peaks from both st9 and st10.5 combined. (D) Clustered heatmaps depicting signals of H3K27me3 and H3K27ac on combined st9 and st10.5 Hdac1 peaks. Clusters denote the same genomic regions in C. Numbers on the right side indicate the total number of regions in each cluster. The signals are shown in a window of 5 kb centered on the summits of Hdac1 peaks presented in descending order of track signal intensities within each cluster. Hdac1 resets the state of acetylated histones by removing acetyl groups, and thus can facilitate the formation of repressive chromatin. Here, we analyzed several histone methylation modifications associated with gene repression (Figure 3B). Hdac1 binds to genomic regions mostly free of H3K9me2, H3K9me3, or H4K20me3. All three modifications are known to mark constitutive heterochromatin denoting gene-poor areas consisting of tandem repeats (Richards and Elgin, 2002). In addition, a fraction of Hdac1-bound regions (Clusters c and d) display signals of H3K27me3 (Figure 3B), which is known to mark facultative heterochromatin consisting of developmental-cue silenced genes (Trojer and Reinberg, 2007). These observations suggest that Hdac1 binds to facultative heterochromatic regions facilitating the repression of genes. Since the majority of Hdac1 peaks (Clusters b–d) are marked by both active and repressive epigenetic signatures, we wonder how Hdac1 functions in epigenetically distinct genomic loci. We compared peaks between two functionally opposing histone modifications H3K27ac and H3K27me3 to Hdac1 peaks, then subdivided these Hdac1 peaks into four clusters representing functionally distinct (Figure Figure 3—figure supplement 1C). Cluster I Hdac1 peaks marked by both H3K27ac and H3K27me3 (Figure that both H3K27ac and H3K27me3 are on the same lysine we that these regions are marked in space due to cell present in the Hdac1 Cluster I peaks are to as Cluster II Hdac1 peaks marked by only H3K27me3 that these regions are associated with inactive developmental genes (Figure Hdac1 Cluster II peaks represent repressive Cluster III Hdac1 peaks marked by only suggesting that these are active Cluster IV Hdac1 peaks with neither H3K27ac H3K27me3 modifications (Figure genomic loci marked with two distinct modifications (Figure 3—figure supplement the expression levels of genes bound by Hdac1 II, and are higher than that of genes and (Figure 3—figure supplement suggesting that Hdac1 binding correlates with transcriptional activity of the genes. Together, we show that Hdac1-bound CRMs are to distinct epigenetic which differential HDAC activity is for differential germ-layer histone A major function of HDACs is to the of acetyl groups from We that Hdac1 regulates histone acetylation of four different Hdac1 Clusters (Clusters in Figure To this we embryos with a used A et al., beginning at the stage and followed the development to stages. with are at gastrula (Figure We observed the of dorsal suggesting that the but not the initiation of gastrulation is A Class I HDAC et al., also a similar (Figure supplement 1A). We first showed that levels of Hdac1 and Hdac2 are not by (Figure supplement 1B). Hdac1 ChIP-seq display high Pearson on embryos with control or at st9 (Figure supplement and st10.5 (Figure supplement which is further by higher Pearson than the batch effect (Figure supplement differential peak analysis on Hdac1 IDR peaks in embryos with control or showed any differential Hdac1 signals peaks or at st9 (Figure supplement and st10.5 (Figure supplement 1H). These results suggest that of early Xenopus embryos not the recruitment of Hdac1 to the To the of HDAC activity inhibition by we surveyed histone acetylation modifications by increased levels of and H3K27ac are observed in embryos (Figure and LaBonne, 2018), and are not detected during this stage of development. These data indicate that HDAC activity is to the level of histone acetylation during gastrulation. Figure with 3 see all Download asset Open asset Hdac1 maintains differential between germ layers. (A) with A gastrulation the dorsal side containing the early (B) Western blot analyses showing various histone acetylation modifications by HDAC is used as a loading control. (C) Clustered heatmap depicting signals of at st10.5 in embryos and The signals are shown in a window of 5 kb centered on the summits of combined and peaks presented in descending order within each cluster. (D) representing of signals found at Hdac1 cis-regulatory clusters in Figure A Hdac1 peak is to it overlaps with either or peaks (Cluster or in a Hdac1 peak is to it overlaps with peaks between and (Cluster A in not overlap with any peak. (E) signals across Hdac1 clusters in Figure in or the differences of signals between and genomic regions are used as the negative control. (F) of signals at Hdac1 clusters in Figure in or the level of (G) of signals in Cluster I of Hdac1 clusters in Figure for each spatial effect size by Figure data 1 Western blot analyses showing various histone acetylation modifications after HDAC Download that Hdac1 Clusters (Clusters in Figure are subjected to both active and repressive epigenetic modifications in different germ we compared the of the (pan-H3Kac) between two distinct germ the and the We performed ChIP-seq at early gastrula (st10.5) on and from embryos with either or control (Figure supplement A of high-confidence peaks from each reveals that a majority of are marked in and germ layers (Figure Figure supplement We observed the signal of in genomic regions (Cluster A in Figure is significantly higher than those of or genomic regions (Clusters and in Figure which also coincides with higher gene expression levels (Figure supplement 2C). To the differential states and the differential gene expression profiles between the two germ we high enrichment peaks to the genes within 10 kb and compared the expression levels of these genes in each germ (Blitz et al., the expression level of genes enriched with are higher in than (Figure supplement and the expression levels of genes enriched with are higher in than (Figure supplement genes with expression signals between germ layers (Figure supplement 2F). These results that the histone acetylation coincides with the and expression of To the role of Hdac1 in regulating histone acetylation states of clusters (Clusters in Figure we first examined the distribution of in these Hdac1 Consistent with both CRMs (Cluster I, both H3K27ac and and active CRMs (Cluster only but not repressive CRMs (Cluster II, only are marked by of marks on or active CRMs are either or (Figure suggesting that these CRMs are active in germ layers. We then
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