Establishment of developmental gene silencing by ordered polycomb complex recruitment in early zebrafish embryos

  1. Graham JM Hickey
  2. Candice L Wike
  3. Xichen Nie
  4. Yixuan Guo
  5. Mengyao Tan
  6. Patrick J Murphy
  7. Bradley R Cairns  Is a corresponding author
  1. Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, United States
  2. Department of Biomedical Genetics, Wilmot Cancer Center, University of Rochester School of Medicine, United States

Abstract

Vertebrate embryos achieve developmental competency during zygotic genome activation (ZGA) by establishing chromatin states that silence yet poise developmental genes for subsequent lineage-specific activation. Here, we reveal the order of chromatin states in establishing developmental gene poising in preZGA zebrafish embryos. Poising is established at promoters and enhancers that initially contain open/permissive chromatin with ‘Placeholder’ nucleosomes (bearing H2A.Z, H3K4me1, and H3K27ac), and DNA hypomethylation. Silencing is initiated by the recruitment of polycomb repressive complex 1 (PRC1), and H2Aub1 deposition by catalytic Rnf2 during preZGA and ZGA stages. During postZGA, H2Aub1 enables Aebp2-containing PRC2 recruitment and H3K27me3 deposition. Notably, preventing H2Aub1 (via Rnf2 inhibition) eliminates recruitment of Aebp2-PRC2 and H3K27me3, and elicits transcriptional upregulation of certain developmental genes during ZGA. However, upregulation is independent of H3K27me3 – establishing H2Aub1 as the critical silencing modification at ZGA. Taken together, we reveal the logic and mechanism for establishing poised/silent developmental genes in early vertebrate embryos.

Editor's evaluation

This manuscript uses genomics tools and pharmacological treatment to study the chromatin landscape change in early-stage Zebrafish embryos, at the critical stage from using maternally deposited transcripts to actively turning on embryotic gene expression. In particular, this work addresses how key chromatin factors coordinate in regulating distinct groups of genes during early vertebrate development and should be of interest to researchers in chromatin biology and developmental biology fields.

https://doi.org/10.7554/eLife.67738.sa0

Introduction

Early vertebrate embryos initiate embryonic/zygotic transcription, termed zygotic genome activation (ZGA), and must distinguish active housekeeping genes from developmental genes, which must be temporarily silenced, but kept available for future activation (Chan et al., 2019; Lindeman et al., 2011; Murphy et al., 2018; Potok et al., 2013; Vastenhouw et al., 2010). Developmental gene promoters in early embryos are packaged in ‘active/open’ chromatin – which can involve a combination of histone variants (e.g. H2A.Z), open/accessible chromatin (via ATAC-seq), permissive histone modifications (e.g. H3K4me1/2/3, H3K27ac), and (in vertebrates such as zebrafish) focal DNA hypomethylation (Akkers et al., 2009; Bogdanovic et al., 2012; Chan et al., 2019; Jiang et al., 2013; Lindeman et al., 2011; Murphy et al., 2018; Potok et al., 2013; Vastenhouw et al., 2010). As H3K9me3 and H3K27me3 are very low or absent at ZGA in zebrafish, it remains unknown how developmental gene silencing occurs at ZGA within an apparently permissive chromatin landscape, and how subsequent H3K27me3 is established at developmental genes during postZGA stages.

We addressed these issues further in zebrafish, which conduct full ZGA at the tenth synchronous cell cycle of cleavage stage (~3.5 hr post fertilization (hpf), ~ 2000 cells) (Schulz and Harrison, 2019). Prior to ZGA (preZGA), zebrafish package the promoters and enhancers of housekeeping genes and many developmental genes with chromatin bearing the histone variant H2afv (a close ortholog of mammalian H2A.Z, hereafter termed H2A.Z(FV)), and the ‘permissive’ modifications H3K4me1 and H3K27ac (Murphy et al., 2018; Zhang et al., 2018); a combination termed ‘Placeholder’ nucleosomes – as they hold the place where poising/silencing is later imposed (Murphy et al., 2018). Curiously, at ZGA in zebrafish (and also in mice and humans), silent developmental gene promoters also contain H3K4me3, a mark that normally resides at active genes (Dahl et al., 2016; Lindeman et al., 2011; Liu et al., 2016; Vastenhouw et al., 2010; Xia et al., 2019; Zhang et al., 2016). After ZGA, developmental genes progressively acquire H3K27me3 via deposition by polycomb repressive complex 2 (PRC2) (Lindeman et al., 2011; Liu et al., 2016; Vastenhouw et al., 2010; Xia et al., 2019). Here, we address the central issues regarding how developmental genes bearing Placeholder nucleosomes and H3K4me3 are transcriptionally silenced during preZGA and ZGA stages in the absence of H3K27me3, and how subsequent H3K27me3 is focally established during postZGA (Figure 1—figure supplement 1A).

Results

H2Aub1 is present in preZGA zebrafish embryo chromatin

First, we sought a repressive histone modification that might explain how developmental genes are silenced at ZGA. We examined zebrafish embryos at preZGA (2.5 hpf, ~256 cells), ZGA (3.5 hpf ~2000 cells), and postZGA (4.3 hpf, >4 K cells) – and confirmed very low-absent H3K27me3 (Figure 1—figure supplement 1B; Lindeman et al., 2011; Vastenhouw et al., 2010) and H3K9me3 absence (Laue et al., 2019) during preZGA and ZGA, but revealed the presence of histone H2A monoubiquitination at lysine 119 (termed hereafter H2Aub1), a repressive mark deposited by the polycomb repressive complex 1 (PRC1) (Figure 1A and B, one of three biological replicates is displayed) (de Napoles et al., 2004; Kuroda et al., 2020; Wang et al., 2004b). Notably, zebrafish sperm lacked H2Aub1 whereas oocytes displayed H2Aub1 (Figure 1B; Figure 1—figure supplement 1C; Figure 1—figure supplement 2A). Current antibodies were not designed to distinguish H2A.Z(FV)ub1 from H2Aub1, so hereafter we refer to the epitope as H2Aub1.

Figure 1 with 3 supplements see all
Polycomb repressive complex 1 (PRC1) occupancy and activity precedes H3K27me3 establishment at promoters.

(A) Detection of H2Aub1 and histone H3 (control) by western blot, prior to (2.5 hr post fertilization [hpf]), during (3.5 hpf), following (4.3 hpf) zygotic genome activation (ZGA), and 24 hpf. (B) Nuclear H2Aub1 immunofluorescence in zebrafish sperm, oocytes (egg), and embryos prior to (2.5 hpf), during (3.5 hpf), and following (4.3 hpf) ZGA. Dashed square: field of view in upper panels. (C) K-means clustering of DNA methylation (DNAme) and chromatin immunoprecipitation (ChIP)-seq (histone modifications/variant) at promoters (UCSC refseq). For DNAme, red color indicates regions that lack DNAme. (D) Genome browser screenshots of ChIP-seq enrichment at representative genes from the clusters in panel (C). (E) Transcription factor motif enrichment from HOMER (Heinz et al., 2010) at promoter clusters from (C).

Developmental promoters acquire Placeholder, Rnf2, and H2Aub1 during preZGA

To localize H2Aub1 we conducted chromatin immunoprecipitation (ChIP) experiments at preZGA, ZGA, and postZGA (replicate structures for Figure 1—figure supplement 2B-D), and examined promoters (Figure 1) and enhancers (Figure 2). For all ChIP experiments, two to three biological replicates were conducted, which involved isolating different batches of zebrafish embryos. To more finely examine ZGA, we conducted additional ChIP profiling of Placeholder nucleosomes at ZGA (3.5 hpf), which complemented our prior profiling at preZGA (2.5 hpf) and postZGA (4.3 hpf) (Murphy et al., 2018) (replicate structure for Figure 1—figure supplement 2E, F). Interestingly, we found H2Aub1 highly co-localized at gene promoters and enhancers with Placeholder nucleosomes, H3K27ac (Zhang et al., 2018; Figure 1C), and ATAC-seq sensitive chromatin (Figure 1—figure supplement 2L, two biological replicates) during preZGA and ZGA stages. However, during postZGA, high levels of H2Aub1 overlap with only a portion of Placeholder-bound loci, an observation explored further, below. For our comparisons to DNA methylation (DNAme), we note that DNAme patterns are reprogrammed between fertilization and the preZGA (2.5 hpf) timepoint (Potok et al., 2013; Jiang et al., 2013), but remain static in zebrafish embryos from 2.5 hpf (preZGA) to 4.3 hpf (postZGA). Therefore, for brevity we chose to display only a single timepoint for DNAme data in subsequent figures, which is representative of all developmental stages examined by the genomics approaches in this work.

Figure 2 with 1 supplement see all
Polycomb repressive complex 1 (PRC1) occupancy and activity precedes H3K27me3 at enhancers.

(A) K-means clustering of whole genome bisulfite sequencing (WGBS) (for DNAme) and chromatin immunoprecipitation (ChIP)-seq at enhancers (postzygotic genome activation [postZGA] H3K4me1 peak summits located outside of promoters). DNAme heatmap displays WGBS fraction-methylated scores (note: red color indicates regions that lack DNAme). ChIP-seq heatmaps display log2(ChIP/input) scores. (B) Genome browser screenshots of ChIP-seq enrichment at representative loci from the indicated K-means clusters in (A). (C) Transcription factor motif enrichment from HOMER (Heinz et al., 2010) at enhancer clusters from (A). (D) Features of an enhancer cluster with exceptionally high H3K27ac and Nanog binding. K-means clusters generated in (A) were utilized to plot heatmaps of Nanog and H3K27ac. (E) A genome browser screenshot depicting Nanog and H3K27ac ChIP enrichment at a DNA-methylated enhancer from cluster 5 in (D).

Rnf2 is the sole zebrafish ortholog of Ring1a and Ring1b/Rnf2, the mutually exclusive E3 ligases within mammalian PRC1, which adds monoubiquitin (ub1) to H2A and H2A.Z (de Napoles et al., 2004; Le Faou et al., 2011; Wang et al., 2004b). Rnf2 ChIP-seq at preZGA, ZGA, and postZGA revealed striking coincidence with H2Aub1, and clustering by Rnf2 occupancy revealed five promoter chromatin types, which differed in Rnf2, H2Aub1, and H3K27me3 enrichment, and gene ontology (Figure 1C; Figure 1—figure supplement 2G-K). Regions with high H2Aub1 and Rnf2 involve broad clustered loci encoding developmental transcription factors (TFs) (cluster 1, e.g. hoxaa) or narrow solo/dispersed developmental TFs (cluster 2, e.g. vsx2) (Figure 1C and D; GO analysis for Figure 1—figure supplement 2K). In counter distinction, loci bearing Placeholder and low-moderate levels of H2Aub1 and low Rnf2 largely constitute housekeeping/metabolic genes, with either broad (cluster 3, e.g. prdx2) or narrow (cluster 4, e.g. gtf3ab) H3K4me3 and H3K27ac occupancy at postZGA (Figure 1C and D; Figure 1—figure supplement 2K). Notably, ‘minor wave’ ZGA genes (genes transcribed at 2.5 hr), including those for pluripotency (e.g. nanog, pou5f3/oct4), bear marking similar to housekeeping genes, and the robustly transcribed mir430 locus appears markedly enriched in H3K27ac at preZGA (Figure 1—figure supplement 2M; Chan et al., 2019). Finally, cluster 5 promoters contain DNAme, and lack Placeholder, H2Aub1, and Rnf2. Thus, over the course of ZGA, loci with Placeholder nucleosomes resolve into two broad classes of loci: developmental genes with high PRC1 and H2Aub1 (clusters 1 and 2), and housekeeping (or ‘minor wave’) genes that lack substantial PRC1 and H2Aub1, but contain high H3K4me3 and H3K27ac at postZGA (clusters 3 and 4) (Figure 1C and D; Figure 1—figure supplement 2J, K).

H3K27me3 establishment occurs during postZGA, and only at locations pre-marked with high Rnf2 and H2Aub1

We find robust H3K27me3 deposition occurring during postZGA at promoters marked during preZGA with high Rnf2 and H2Aub1, specifically at clusters 1 and 2 (Figure 1C and D). Furthermore, as embryos transition from preZGA to postZGA, H3K27ac diminishes at developmental loci, whereas housekeeping genes (clusters 3 and 4 Figure 1C and D) retain strong H3K27ac and become active. During postZGA, developmental genes acquire the combination of low-moderate H3K4me3 and high H3K27me3, termed ‘bivalency’ (Azuara et al., 2006; Bernstein et al., 2006). Interestingly, promoters that become bivalent postZGA involve those pre-marked with higher relative Rnf2 and H2Aub1, whereas promoters with high H3K27ac, high H3K4me3, and low-absent H3K27me3 postZGA involve those pre-marked with lower relative Rnf2 and H2Aub1 (Figure 1C and D; Figure 1—figure supplement 2J). This observation raised the possibility that high H2Aub1 levels may help recruit PRC2 to subsequently deposit H3K27me3 at developmental genes. Notably, analysis of Rnf2, H2Aub1, and H3K27me3 at LINE, LTR, and satellite repeats revealed no ChIP-seq enrichment at these genomic elements during preZGA, ZGA, or postZGA (Figure 1—figure supplement 3A-C), reinforcing that this modification axis is focused on the marking of developmental loci.

To identify candidate TFs that might bind selectively at the promoters of particular clusters, we analyzed the DNA sequences flanking the transcription start site (TSS) (500 bp) at each cluster using the motif finding program, HOMER (Figure 1E; Heinz et al., 2010). Largely non-overlapping motifs were identified for TF-binding sites at clusters linked to developmental vs. housekeeping genes (Figure 1E, clusters 1 and 2 vs. 3 and 4; partitioned by H2Aub1/Rnf2 levels). Here, the strong enrichment of motifs for homeodomain-containing TFs (and other families) in clusters 1 and 2 provides candidate factors that may help recruit PRC1 to developmental loci. DNA-methylated promoters at ZGA (cluster 5) represent a large and heterogeneous set of genes, which are activated in particular cell types later in development.

Enhancer poising parallels features and factors at promoters

Analysis of enhancer regions revealed features that were similar to those at developmental promoters (Figure 2). Specifically, enhancers with high H2Aub1 and Rnf2 during preZGA and ZGA acquired robust H3K27me3 during postZGA (Figure 2A and B; Figure 2—figure supplement 1A). Enhancers with low/absent Rnf2 and H2Aub1 failed to attract robust H3K27me3 at postZGA, instead bearing high levels of H3K27 acetylation (Figure 2A and B; Figure 2—figure supplement 1A). Candidate TF-binding sites were enriched at enhancers (Figure 2C), and these sites overlapped partly with those enriched at promoters, consistent with the expectation that the factors that recruit histone modifiers to promoters and enhancers partially overlap. Taken together, enhancers acquire H3K27me3 during postZGA in proportion to their levels of Rnf2 and H2Aub1 during preZGA and ZGA, consistent with our observations at promoters.

A unique enhancer class with high H3K27ac and DNA methylation

Curiously, enhancer cluster 5 (Figure 2A) was unique at postZGA – displaying high H3K4me1, very high H3K27ac, and open chromatin (via ATAC-seq analysis; Figure 2—figure supplement 1C,D) – but bore DNA methylation – an unusual combination given the typical strong correlation between high H3K4me1 and DNA hypomethylation. Notably, Nanog-binding sites were highly enriched solely at cluster 5, and Nanog ChIP-seq during ZGA and postZGA (Xu et al., 2012) showed Nanog occupancy highly and selectively enriched at cluster 5 relative to other enhancer clusters (Figure 2C–E). Thus, cluster 5 enhancers may utilize Nanog and H3K27ac to open and poise these DNA-methylated enhancers for later/subsequent transition to an active state. Consistent with this notion, GO analysis of cluster 5 enriches for terms related to developmental and signaling processes (p-value; 5.1E-18) (Figure 2—figure supplement 1B).

The PRC2 component Aebp2 is coincident with Rnf2 and H2Aub1 at developmental loci

The pre-marking of developmental genes with H2Aub1 and Rnf2-PRC1 prior to H3K27me3 establishment raised the possibility of a ‘non-canonical’ (nc) mode of recruitment, involving ncPRC1 action (H2Aub1 addition) followed by the recruitment of PRC2 (ncPRC2) – via H2A/Zub1 recognition – to deposit H3K27me3. This mode and order of recruitment has precedent in Drosophila and in mammalian embryonic stem (ES) cell cultures, with the Aebp2 and Jarid2 protein components of ncPRC2 recognizing H2Aub1 and both targeting and facilitating H3K27me3 addition (Blackledge et al., 2014; Blackledge et al., 2020; Cooper et al., 2014; Cooper et al., 2016; Kalb et al., 2014; Kasinath et al., 2021; Tamburri et al., 2020). We then addressed whether establishment of H3K27me3 during postZGA is mediated by the non-canonical Aebp2-Jarid2-PRC2 complex at loci pre-marked with H2Aub1. Interestingly, Aebp2 protein levels were very low during preZGA and ZGA stages, but robustly detected postZGA (Figure 3A), without a large increase in aebp2 transcript levels (Figure 3—figure supplement 1A). Aebp2 ChIP-seq during postZGA revealed a remarkably high coincidence of Aebp2 with H2Aub1, Rnf2, and H3K27me3 at promoters (Figure 3B and C; Figure 3—figure supplement 1B-D) and at enhancers (Figure 3D and E, Figure 3—figure supplement 1E). Taken together, these results suggest that translational upregulation and/or protein stability enables Aebp2 protein accumulation postZGA – enabling the ‘reading/binding’ of H2Aub1, and H3K27me3 deposition during postZGA by PRC2.

Figure 3 with 2 supplements see all
Aebp2-polycomb repressive complex 2 (PRC2) mediates de novo H3K27me3 at loci pre-marked by H2Aub1.

(A) Nuclear Aebp2 detection by immunofluorescence in postzygotic genome activation (postZGA) zebrafish embryos (4.3 hr post fertilization [hpf]). No Aebp2 staining was detected at preZGA (2.5 hpf) or ZGA (3.5 hpf). Bottom row: The dashed square indicates the field of view in upper panels. One of three biological replicates is shown. (B) Aebp2 binding at promoters during postZGA overlaps and scales with occupancy of Rnf2, H2Aub1, and H3K27me3. Promoter clusters from Figure 1C were utilized to plot heatmaps. (C) Genome browser screenshots of chromatin immunoprecipitation (ChIP)-seq at representative promoter loci from clusters in (B). (D) Aebp2 binding at enhancers during postZGA overlaps with occupancy of Rnf2, H2Aub1, and H3K27me3. Enhancer clusters from Figure 2A were utilized to plot heatmaps. (E) Genome browser screenshots of ChIP-seq enrichment at representative enhancer loci from clusters in (D).

To investigate the possible additional contribution of Jarid2 to ‘reading/binding’ of H2Aub1 (leading to H3K27me3 deposition), we performed ChIP-seq of Jarid2 during postZGA (Figure 3—figure supplement 2A). Here, profiling of Jarid2 occupancy by ChIP-seq resulted in chromatin maps with only modest enrichment and dynamic range, providing 448 bound promoters (Figure 3—figure supplement 2B). Comparison of Aebp2 and Jarid2 occupancy at promoters revealed that the majority of Jarid2-binding sites (295/448) overlap with Aebp2-bound sites (Figure 3—figure supplement 2B). Only a minority of Aebp2-bound promoters overlapped with Jarid2 – an observation that may reflect our modest ChIP efficiency, but could also reflect the presence of Aebp2-bound promoters that lack Jarid2 binding. Notably, promoters bound by both Aebp2 and Jarid2 had enrichment of GO-term categories corresponding to developmental genes (Figure 3—figure supplement 2C, D). Conversely, promoters bound solely by Jarid2 were not associated with developmental functions and instead enriched for ribosomal genes (Figure 3—figure supplement 2C). Analysis of Aebp2 and Jarid2 occupancy at enhancer loci revealed similarities to our analysis at promoters. Here, the majority of Jarid2-bound enhancers (84/163) were also bound by Aebp2 (Figure 3—figure supplement 2E, F). However, this overlap accounted for only a minority of the Aebp2-bound enhancers. As our Aebp2 ChIP-seq exhibited greater robustness and dynamic range than Jarid2, we hereafter utilized Aebp2 occupancy as the primary functional marker for non-canonical PRC2 complex in the remainder of our analyses.

Loss of H2Aub1 via Rnf2 inhibition prevents Aebp2 localization and H3K27me3 deposition

To functionally test whether H2Aub1 recruits Aebp2-PRC2 for de novo establishment of H3K27me3 at developmental genes, we utilized the RNF2 inhibitor, PRT4165 (Chagraoui et al., 2018; Ismail et al., 2013; Zhu et al., 2018). PRT4165 is a small molecule inhibitor of Rnf2 that has previously been shown to strongly reduce H2Aub1 modification, but not to affect the activity of related H2A E3 ligases such as Rnf8 and Rnf168 (Chagraoui et al., 2018; Ismail et al., 2013; Zhu et al., 2018). In each of three biological replicates, PRT4165 treatment (150 μM) from the one-cell stage onward largely eliminated H2Aub1 by 4 hpf (ZGA) (Figure 4A and B), and conferred a developmental arrest that resembled untreated 4 hpf embryos (Figure 4—figure supplement 1A). To determine whether the loss of H2Aub1 conferred loss of Aebp2 genomic targeting, we performed ChIP experiments on Aebp2 in PRT4165-treated and DMSO-treated embryos (three biological replicates per condition). Remarkably, developmental loci that normally display high Aebp2 in untreated or DMSO-treated embryos lost Aebp2 binding following PRT4165 treatment (Figure 4C, clusters 3 and 4; Figure 4—figure supplement 1B, C). Curiously, PRT4165 treatment also conferred many new/ectopic Aebp2 peaks (Figure 4C, clusters 1 and 2), however our profiling of H3K27me3 following PRT4165 treatment (three biological replicates) revealed that new/ectopic Aebp2 sites did not acquire H3K27me3 (Figure 4C, clusters 1 and 2; Figure 4—figure supplement 2B), consistent with prior observations that other modifications (such as H2Aub1; Kalb et al., 2014) may be needed to stimulate H3K27me3 addition by PRC2. Importantly, treatment with PRT4165 eliminated or strongly reduced H3K27me3 at virtually all loci normally occupied by Aebp2 and H3K27me3 (Figure 4C, clusters 3 and 4, Figure 4—figure supplement 1D), a conclusion supported by our use of a ‘spike-in’ control involving Drosophila nuclei bearing H3K27me3-marked regions in all three replicates per condition. Finally, new ectopic H3K27me3 peaks were very rare in the PRT4165 treatment, and none of these loci were bound by Aebp2 (Figure 4—figure supplement 2B). Taken together, H2Aub1 deposition by Rnf2 during preZGA is required for the recruitment of Aebp2 and subsequent de novo deposition of H3K27me3 postZGA at developmental loci.

Figure 4 with 3 supplements see all
Catalytic activity of polycomb repressive complex 1 (PRC1) is required for aebp2 binding, H3K27me3 establishment, and transcriptional repression of developmental genes.

(A) Experimental design of drug treatments to inhibit Rnf2 activity. Embryos at the one-cell stage were added to media containing either PRT4165 (150 μM) or DMSO and raised until 4 hr post fertilization (hpf). (B) PRT4165 treatment of embryos confers bulk loss of H2Aub1 at 4 hpf. Left: Western blot for H2Aub1 in 4 hpf embryos treated with DMSO (vehicle) or 150 μM PRT4165 (Rnf2 inhibitor). Right: Quantification western blot in left panel. (C) Impact of Rnf2 inhibition on Aebp2 genomic localization and H3K27me3. K-means clustering of Aebp2 and H3K27me3 chromatin immunoprecipitation (ChIP)-seq enrichment at all loci with called peaks in any of the datasets plotted. Embryos were treated from the one-cell stage with either DMSO or 150 μM PRT4165 and harvested at 4 hpf for ChIP analysis. H3K27me3 ChIP-seq from untreated embryos at 4.3 hpf is plotted as a comparitor. (D) Impact of Rnf2 inhibition on gene expression. Volcano plot of RNA-seq data from PRT4165-treated vs. untreated embryos (4 hpf). Green and red data points signify transcripts with p-values < 0.01 and at least a 3-fold change (increase or decrease) in expression, respectively. Marquee upregulated genes encoding developmental transcription factors are labelled. (E) Genome browser screenshots of representative developmental genes which, upon Rnf2 inhibition, lose Aebp2 binding and H3K27me3 marking, and become transcriptionally active.

To determine whether H2Aub1 impacts transcriptional repression of developmental genes at postZGA, we performed RNA sequencing (three biological replicates per condition) on 4 hpf embryos that were either vehicle-treated (DMSO) or PRT4165-treated from the one-cell stage onward (Figure 4—figure supplement 1E). Here, we identified and characterized both up- and downregulated genes (Figure 4E). PRT4165-upregulated and -downregulated genes (>3-fold, p-value < 0.01) were both enriched in developmental factors, but the number of genes associated with upregulated GO-terms was substantially greater than downregulated GO-terms (Figure 4—figure supplement 1F). Here, ~16.6% of H2Aub1-marked protein coding genes were upregulated, which may reflect the availability at postZGA of an opportunistic activator, following H2Aub1 loss (Figure 4—figure supplement 2C). Affected genes include those in clustered loci (e.g. Hox genes) where the effect was moderate, as well as non-clustered/solo formats where the effect of PRT4165 was more pronounced (Figure 4E and F; Figure 4—figure supplement 2A).

Having observed precocious developmental gene upregulation in response to PRT4165 treatment during postZGA (4 hpf), we were curious whether their upregulation could also be observed during the preZGA and ZGA stages. To test, we repeated our PRT4165 treatment regimen and isolated embryos at preZGA (2.5 hpf) and ZGA (3.5 hpf) stages, and performed RNAseq (with biological triplicates; Figure 4—figure supplement 3). Here, analysis of upregulated genes meeting our threshold criteria (fold change ≥1.5; p-value ≤ 0.01) by GO-term analysis uncovered no enrichment of developmental genes upon RNF2 inhibition during preZGA or ZGA. Instead, enriched GO-terms from upregulated transcripts corresponded to genes encoding RNA-binding proteins and ribosomal proteins. Thus, chromatin de-repression (via H2Aub1 removal) does not cause transcriptional activation of large numbers of developmental genes during preZGA or ZGA – not even the developmental genes that are activated postZGA following Rnf2 inhibition. Here, we note that general/housekeeping transcription does not occur until ZGA; only the miRNA-430 locus and a very limited number of genes are transcribed during preZGA. Therefore, H2Aub1 removal does not, by itself, lead to the activation of developmental genes that are normally marked by H2Aub1 during preZGA. Taken together, these results strongly suggest that H2Aub1 represses developmental genes during preZGA and ZGA, independent of H3K27me3, and that the absence of H2Aub1 renders developmental genes susceptible to precocious activation following ZGA, which confers developmental arrest prior to gastrulation.

Discussion

Our work reveals that developmental gene silencing in early zebrafish embryos is established through sequential recruitment and activity of PRC1 and PRC2 complexes, respectively, to otherwise open/permissive loci bearing Placeholder nucleosomes (Figure 5). Placeholder nucleosomes containing H3K4me1 and the histone variant H2A.Z(FV) are installed by the chromatin remodeler SRCAP during preZGA (Murphy et al., 2018), and are focally pruned to small regions by the chaperone Anp32e (Kobor et al., 2004; Krogan et al., 2003; Mao et al., 2014; Obri et al., 2014; Wang et al., 2016). Here, our reanalysis of published data confirms that H3K27ac is an additional component of Placeholder nucleosomes during preZGA (Zhang et al., 2018). Functional studies reveal that Placeholder nucleosomes prevent DNAme where they are installed and are utilized to reprogram the DNAme patterns during cleavage stage. Therefore, from a ‘permissive’ Placeholder platform during preZGA, two very different chromatin/transcriptional states are attained at ZGA: active or poised (Murphy et al., 2018).

Model for polycomb-mediated establishment of developmental gene silencing during zebrafish embryogenesis.

Prior to zygotic genome activation (ZGA), Rnf2-PRC1 is recruited by transcription factors (TFs) to promoters (shown) and enhancers (not shown) of developmental genes bearing Placeholder nucleosomes (H2A.Z(FV), H3K4me1, H3K27ac). Rnf2-PRC1 deposition of H2Aub1 recruits Aebp2-PRC2 to catalyze H3K27me3 addition. H2Aub1 ablation (via Rnf2 inhibition) eliminates Aebp2-PRC2 recruitment and prevents H3K27me3 establishment. Notably, H2Aub1 loss causes precocious transcription of certain developmental genes after ZGA, identifying H2Aub1 as a critical component of silencing at ZGA.

Our work suggests that the initial poised state at developmental genes and enhancers involves the imposition of polycomb-based silencing upon the permissive states established by Placeholder nucleosomes. Specifically, we observe H2Aub1 addition by Rnf2/PRC1 during preZGA and ZGA to confer initial transcriptional silencing at developmental loci – which is subsequently read by the Aebp2/PRC2 complex to add H3K27me3 after ZGA (Figure 5). Prior work in ES cells has provided an in vitro parallel in which PRC1 activity (H2Aub1 addition) can occur independently of PRC2-mediated recruitment at certain loci (Blackledge et al., 2014; Blackledge et al., 2020; Cooper et al., 2014; Cooper et al., 2016; Kalb et al., 2014; Tamburri et al., 2020; Tavares et al., 2012), which has been termed ‘non-canonical’ order of recruitment, to contrast with prior data showing the reverse/canonical order (Wang et al., 2004a). Furthermore, and consistent with our work, human AEBP2 and JARID2 have recently been shown to directly bind H2Aub1 and stimulate PRC2 activity in the presence of H3K4 methylation (Kasinath et al., 2021), and a similar mechanism may be utilized to establish bivalency after ZGA in zebrafish.

Our work also clarifies and extends prior work in zebrafish which showed that an incross of zebrafish heterozygous for an rnf2 loss-of-function truncation mutation yielded a pleitropic terminal phenotype at 3 days post fertilization (dpf), coincident with the loss of rnf2 RNA. Notably, loss of rnf2 at 3 dpf was associated with partial upregulation of certain developmental genes, but H3K27me3 remained fully present, showing that Rnf2 is not required for H3K27me3 maintenance (Chrispijn et al., 2019). However, progression of rnf2 mutants to 3 dpf may have relied on maternally inherited WT rnf2 RNA or protein to provide the initial establishment of gene silencing and H3K27me3, raising the possibility that rnf2 is actually essential at a much earlier developmental stage. Our use of the Rnf2 inhibitor, PRT4165, during preZGA and ZGA stages reveals the necessity for Rnf2 activity for the establishment of developmental gene silencing, for subsequent H3K27me3 addition, and for progression of zebrafish development beyond the ZGA stage. Importantly, developmental gene upregulation is not attributable to H3K27me3 loss, as maternal zygotic ezh2 mutant zebrafish embryos do not precociously activate developmental genes during ZGA, and they progress through gastrulation without H3K27me3 (Rougeot et al., 2019; San et al., 2016; San et al., 2019). Notably, although upregulation of developmental genes in the presence of PRT4165 is clear, this involves ~16.6% of the developmental gene repertoire occupied by H2Aub1/Rnf2. Here, we suggest that tissue-specific activators for the majority of developmental genes are not present at ZGA.

Maternal loading of mRNA and proteins present challenges for functional experiments during early embryonic stages of development. Our use of the Rnf2 inhibitor, PRT4165, enabled us to test whether H2Aub1 impacts gene expression at ZGA, and whether H2Aub1 is required for the recruitment of Aebp2-PRC2 and for the subsequent establishment of H3K27me3 at developmental loci during postZGA. The observable impact of PRT4165 includes the loss of H2Aub1, the loss of Aebp2 recruitment to all loci that formerly bore H2Aub1, and the loss of H3K27me3 deposition at all developmental loci that are normally marked by H3K27me3 during postZGA. Our interpretation of these observations is that they are linked to, and dependent on, Rnf2 inhibition. However, it is important to note that PRT4165 may have off-target effects aside from inhibition of Rnf2 which may contribute to our observations. While we cannot formally account for possible off-target effects of PRT4165, we note that ZGA (which involves the activation of thousands of genes) progresses relatively normally, except for the upregulation of a cohort of developmental genes which were formerly H2Aub1 marked. This provides a measure of confidence that the upregulation of developmental genes is largely a direct consequence of H2Aub1 loss in the early embryo. However, as PRT4165 was the only approach we employed that successfully removed the vast majority of H2Aub1, future studies involving orthogonal approaches to elicit Rnf2 loss in zebrafish will be needed to further validate the consequences of H2Aub1 loss in early zebrafish embryos.

While our work was in review, work on H2Aub1 and H3K27me3 dynamics in the preimplantation mouse embryo was reported by two separate groups (Chen et al., 2021; Mei et al., 2021). Both studies in mouse demonstrated that H2Aub1 temporally precedes H3K27me3 at developmental genes during early embryonic stages, and both studies also lowered H2Aub1 via one effective approach: either through maternal genetic loss of non-canonical PRC1 subunits (Pcgf1/6) or through overexpression of an H2A de-ubiquitinase (PR-DUB). Here, both approaches led to precocious transcription of a moderate subset of developmental genes at or shortly after ZGA. Thus, these sets of results align well with our observations in zebrafish.

Notably, each method of H2Aub1 perturbation in mice was unique (with attendant advantages and caveats) and conferred different impacts on embryogenesis and establishment of H3K27me3 at developmental loci. In mice, zygotic overexpression of PR-DUB resulted in rapid and significant erasure of H2Aub1, precocious transcription of certain developmental genes, and growth arrest at the four-cell stage. However, reduction of H2Aub1 by zygotic PR-DUB overexpression had minimal impact on H3K27me3, likely due to the fact that H3K27me3 is very low at the four-cell stage. In contrast, Mei et al. disrupted H2Aub1 via maternal genetic ablation (paternal genes remained intact) of two genes, Pcgf1/6, which encode subunits of noncanonical PRC1. Here, Pcgf1/6 mutant oocytes were fertilized with wild-type sperm to produce maternal-null Pcgf1/6 embryos. In contrast to PR-DUB overexpression, maternal-null Pcgf1/6 embryos displayed global reductions in both H2Aub1 and H3K27me3 during cleavage stages – a finding that aligns well with our findings. Notably, the reductions of H2Aub1 and H3K27me3 from maternal Pcgf1/6 loss largely returned to wild-type levels by the morula stage, possibly due to zygotically (and paternally) expressed Pcgf1/6. Here, we postulate that the more severe loss of H3K27me3 from Pcgf1/6 deletion may stem from the temporal difference of disrupting H2Aub1 in the female germline vs. the early mouse embryo, or potentially the developmental stage in which H3K27me3 was assayed ( two cell vs. four cell). A major issue not addressed in these recent papers in mouse is how mechanistically H2Aub1 directs H3K27me3. Notably, our study addresses this mechanism by demonstrating that non-canonical PRC2 complex containing the H2Aub1 ‘reader’ Aebp2 occupies all loci with high H2Aub1 during postZGA, which then receive H3K27me3.

Interestingly, Chen et al. and Mei et al. utilized hybrid mouse strains to trace parental asymmetries of H3K27me3 and H2Aub1 in the early embryo. Here, both groups uncovered rapid erasure of H2Aub1 from the paternal genome shortly after fertilization which is re-established by the two-cell stage. Our data also demonstrate that the paternal genome is initially devoid of H2Aub1, and rapidly acquires H2Aub1 shortly after fertilization. However, zebrafish sperm are entirely devoid of H2Aub1 (Figure 1B; Figure 1—figure supplement 1C), whereas mouse sperm bear H2Aub1 coincident with H3K27me3. Thus, the paternal genome in both mammals and zebrafish attain a transient state lacking H2Aub1 soon after fertilization – but arrive at that state by alternative routes.

Here, we observe that developmental or housekeeping gene promoters attract either a high- or low-level PRC1 binding, respectively. How ncPRC1 is recruited to CpG islands in ES cells is partially understood, as the ncPRC1 subunit Kdm2b helps recruit ncPRC1 to CpG islands of developmental genes. In this context, Kdm2b binds to hypomethylated CpGs via the CxxC motif (Blackledge et al., 2014; Farcas et al., 2012; He et al., 2013; Wu et al., 2013). However, ncPRC1 is recruited robustly to only a minority of Kdm2b-bound CpG islands, implying that additional factors are needed to specify recruitment of ncPRC1 to CpG islands of developmental genes. Here, we speculate that particular TFs, such as the candidates in Figures 1E and 2C, are likewise utilized in cooperation with zebrafish Kdm2b to enable strong focal recruitment of Rnf2-PRC1 to Placeholder-occupied developmental loci. In contrast, the candidate TFs at housekeeping genes would recruit MLL complexes to implement H3K4 methylation, and not Rnf2-PRC1. Thus, future work will explore which particular TFs recruit SRCAP (for Placeholder/H2A.Z installation), PRC1 (for H2Aub1 addition at developmental genes), or MLL complexes (for H3K4me3 modification at housekeeping genes) to define which specific genes are subjected to silencing or activation at ZGA.

One curiosity arising from our work is why zebrafish utilize the non-canonical PRC1 complex, which adds monoubiquitination to H2A/H2A.Z(FV), rather than canonical PRC1 complex, which compacts chromatin for initial developmental gene silencing. Here, we speculate that the rapid (~16 min) cell cycles that characterize the preZGA cleavage state – coupled to the need for continual DNA replication during cleavage stage – are not compatible with the compaction conferred by canonical PRC1 (Gao et al., 2012; Grau et al., 2011; Lau et al., 2017). Furthermore, the necessary substrate to recruit canonical PRC1 to chromatin, H3K27me3, is absent in preZGA embryos. Instead, the use of the repression modes conducted by non-canonical PRC1 addition of H2Aub1 – which antagonizes RNA Pol II transcriptional initiation or bursting – may help confer silencing without conferring a compaction that might impede DNA replication (Dobrinić et al., 2020; Stock et al., 2007). However, once embryos exit cleavage stage, the cell cycle greatly lengthens, and Aebp2-PRC2 complexes add H3K27me3 to loci – which may then enable canonical PRC1 to localize to H3K27me3-marked loci, and conduct compaction. Indeed, prior work has shown that later stages of zebrafish development utilize canonical PRC complexes (Rougeot et al., 2019). Here, future studies may reveal more precisely the logic and timing underlying the transition from non-canonical to canonical utilization of PRC complexes in zebrafish development.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Gene (Danio rerio)Zebrafish GenomeUCSCZv10
AntibodyAnti-H2Aub1(rabbit monoclonal)Cell Signaling TechnologyCat# 8240; RRID:AB_10891618WB (1:1000)ChIP (1:100)IF (1:500)
AntibodyAnti-rnf2(rabbit monoclonal)Cell Signaling TechnologyCat# 5694; RRID:AB_10705604(1:100)
AntibodyAnti-H2A.Z(rabbit polyclonal)Active MotifCat# 39113; RRID: AB_2615081(5 μl)
AntibodyAnti-H3K4me1(rabbit polyclonal)Active MotifCat# 39297; RRID: AB_2615075(10 μl)
AntibodyAnti-aebp2(rabbit monoclonal)Cell Signaling TechnologyCat# 14,129 S; RRID: AB_2798398ChIP (1:100)IF (1:500)
AntibodyAnti-H3K27me3(rabbit polyclonal)Active MotifCat# 39155; RRID: AB_2561020ChIP (5 μl)IF (1:500)
AntibodyAnti-H3(mouse monoclonal)Active MotifCat# 39763; RRID: AB_2650522(1:2000)
AntibodyAnti-Jarid2rabbit polyclonal)NovusCat# NB100-2214; RRID:AB10000529(5 μl)
Cell line (Danio melanogaster)Cell line S2: S2-DRSCATCCCat# CRL-1963 RRID:CVCL_Z232
Chemical compound, drugPRT4165TocrisCat# 5047
Commercial assay or kitNEBNext ChIP-Seq Library Prep Master Mix Set for IlluminaNew England BiolabsCat# E6240.
Commercial assay or kitIllumina TruSeq Stranded mRNA Library Prep KitIlluminaCat# RS-122–2101, RS-122–2102
Software, algorithmNovoalignNovocraftRRID:SCR_014818
Software, algorithmSamtoolsLi et al., 2009RRID:SCR_002105
Software, algorithmdeepToolsRamírez et al., 2016RRID:SCR_016366
Software, algorithmMACS2Zhang et al., 2008RRID:SCR_013291
Software, algorithmUCSC Exe UtilitiesUCSC Genome Browserhttp://hgdownload.soe.ucsc.edu/downloads.html#source_downloads
Software, algorithmIGVThorvaldsdóttir et al., 2012RRID:SCR_011793
Software, algorithmDAVIDHuang et al., 2009https://david.ncifcrf.gov
Software, algorithmRR Development Core Team, 2020RRID:SCR_001905
Software, algorithmR StudioRStudio Team, 2021RRID:SCR_000432
Software, algorithmChipseekrYu et al., 2015https://www.bioconductor.org/packages/release/bioc/html/ChIPseeker.html
Software, algorithmBio-ToolBoxTim Parnell of the Huntsman Cancer InstituteParnell, 2021b; https://github.com/tjparnell/biotoolbox
Software, algorithmMulti-Replica Macs ChIPSeq WrapperTim Parnell of the Huntsman Cancer InstituteParnell, 2021a; https://github.com/HuntsmanCancerInstitute/MultiRepMacsChIPSeq
Software, algorithmHOMERHeinz et al., 2010RRID:SCR_010881
Software, algorithmSTARDobin et al., 2013RRID:SCR_015899
Software, algorithmfeatureCountsLiao et al., 2013RRID:SCR_012919
Software, algorithmDESeqAnders and Huber, 2010RRID:SCR_000154

Zebrafish husbandry

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Wild-type Tübingen zebrafish were maintained as described (Westerfield, 2007). All experiments involving zebrafish were approved by University of Utah IACUC (Protocol 20–04011). Embryos were scored for developmental staged as described (Kimmel et al., 1995).

Acid extraction of nuclear proteins

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Two-hundred embryos were added to 1.5 ml tubes and washed twice with cold PBS; 800 μl of Mild Cell Lysis Buffer (10 mM Tris-HCl pH 8.1, 10 mM NaCl, 0.5 NP-40, 2× protease inhibitors) was applied to embryos and incubated on ice for 5 min. Embryos were homogenized by passing through a 20-gauge syringe several times. Samples were briefly centrifuged to bring down chorions. Supernatants were transferred to fresh tubes and centrifuged at 1300× g for 5 min at 4°C. Pelleted nuclei were washed twice with cold Mild Cell Lysis Buffer. Nuclei were resuspended to a final volume of 800 μl in cold Mild Cell Lysis Buffer and supplemented with 10 μl of sulfuric acid (18.4 M). Samples were sonicated for 10 s (1 s ON, 0.9 s OFF) at 30% output using a Branson sonicator. Proteins were extracted for 30 min at 4°C on a rotator, and 160 μl of 100% trichloroacetic acid was added and proteins were allowed to precipitate for 30 min on ice. Samples were centrifuged at 13,000 rpm for 5 min at 4°C. Protein pellets were washed with 800 μl of cold acidified acetone, and centrifuged again at 13,000 rpm for 5 min at 4°C. Protein pellets were washed with 800 μl of cold acetone, and centrifuged again at 13,000 rpm for 5 min at 4°C. Supernatant was discarded and pellets were dried at 37°C for 5 min. Dried protein pellets were resuspended in 2× Laemmli sample buffer and boiled for 8 min. Samples were then used for western blotting. Bands were quantified in ImageJ (Schneider et al., 2012).

Immunohistochemistry and DAPI staining

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Standard protocol for immunohistochemistry was followed as described (Fernández and Fuentes, 2013; Zhang et al., 2018). Three biological replicates were performed for each immunohistochemistry experiment. Briefly, 30 embryos were collected at appropriate timepoints and fixed with fresh 4% paraformaldehyde (Electron Microscopy, Cat# 50980487) in 1× PBS at room temperature for 12 hr. Droplets of glacial acetic (100%, Merck, Cat# 1000560001) or DMSO (final concentration 0.5%, Sigma) were added 5–10 s after initiation of the fixation. Chorions were manually removed from fixed embryos with forceps and dechorionated embryos were dehydrated in methanol and stored at –20°C. For immune-staining, embryos were rehydrated into PB3T (1× PBS with 0.3% TritonX-100, and then incubated in blocking agent 1% BSA, 0.3 M glycine in PB3T). Embryos were incubated with primary antibodies diluted in blocking agent overnight at 4°C. Primary antibodies were removed and embryos were washed extensively with PB3T. Embryos were next incubated with appropriate secondary antibodies in the dark followed by extensive washes in PB3T. Primary antibodies used for immune-staining are listed below. Secondary antibodies used were donkey α-rabbit IgG-488 at 1:500 (Life Technologies, Cat# A-21206). DAPI was used at 1:1000 as a nuclear counterstain. The yolk cells were removed from embryo and embryo was mounted on glass slide with ProLong Gold Antifade mounting media (Thermo Fisher, Cat# P-36931) and a 2.0 mm square coverslip sealed with nail polish. Samples were stored at 4°C until imaged.

Imagining of zebrafish embryos

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Images were acquired on a Leica SP8 White Light laser confocal microscope. Image processing was completed using Nikon NIS-Elements multi-platform acquisition software with a 40×/1.10 Water objective. Fiji (ImageJ, V 2.0.0-rc-69/1.52p) was utilized to color DAPI channel to cyan, GFP color remained green. Confocal images are max projections of Z stacks taken 0.5 μm apart for a total of the embryo ~7–12 μm.

Primary antibodies

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The following antibodies were utilized in for the present study: anti-H2Aub1 (Cell Signaling Technology Cat# 8240; RRID:AB_10891618), anti-Rnf2 (Cell Signaling Technology Cat# 5694; RRID:AB_10705604), anti-H2A.Z (Active Motif Cat# 39113; RRID:AB_2615081), anti-H3K4me1 (Active Motif Cat# 39297; RRID:AB_2615075), anti-Aebp2 (Cell Signaling Technology Cat# 14129; RRID: AB_2798398), anti-H3K27me3 (Active Motif Cat# 39155; RRID: AB_2561020), anti-H3 (Active Motif Cat# 39763; RRID:AB_2650522), and anti-Jarid2 (Novus Cat# NB100-2214; RRID:AB10000529).

ChIP-seq in zebrafish embryos

Embryo fixation

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Approximately 1.5 million cells were used for each ChIP replicate. Embryos were allowed to progress to the desired developmental stage and then transferred to 1.5 ml microcentrifuge tubes (~200 embryos per tube). Chorions were removed enzymatically by treatment with pronase (1.25 mg/ml in PBS). Dechorionated embryos were gently washed twice with PBS to remove pronase. Samples were fixed with 1% formaldehyde (Electron Microscopy Sciences, Cat# 15712) for 10 min at room temperature with end over end rotation. Fixation was quenched with 130 mM glycine for 5 min at room temperature. Samples were centrifuged for 5 min at 500× g at 4°C. Supernatant was discarded and cell pellets were washed twice with ice-cold PBS. Cell pellets were frozen with liquid nitrogen and stored at –80°C.

Nuclei isolation and lysis

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One ml of Mild Cell Lysis Buffer (10 mM Tris-HCl pH 8.1, 10 mM NaCl, 0.5 NP-40, 2× proteinase inhibitors) was applied to cell pellets from 1000 embryos and rotated at 4°C for 10 min. Samples were centrifuged at 1300× g for 5 min at 4°C. Supernatant was discarded and nuclei pellets were resuspended in 1 ml Nuclei Wash Buffer (50 mM Tris-HCl pH 8.0, 100 mM NaCl, 10 mM EDTA, 1% SDS, 2× protease inhibitors) and rotated at room temperature for 10 min. Samples were centrifuged at 1300× g for 5 min at 4°C to pellet nuclei. Supernatant was discarded and nuclei pellets were resuspended in 100 µl of Nuclei Lysis Buffer (50 mM Tris-HCl pH 8.0, 10 mM EDTA, 1% SDS, 2× proteinase inhibitors). Samples were incubated on ice for 10 min; 900 µl of IP Dilution Buffer (16.7 mM Tris-HCl pH 8.1, 167 mM NaCl, 1.2 mM EDTA, 0.01% SDS, 1.1% Triton X-100, 2× proteinase inhibitors) was added to samples.

Chromatin sonication

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Nuclear lysates were sonicated with a Branson Digital Sonifier with the following settings: 10 s duration (0.9 s ON, 0.1 s OFF), 30% amplitude. Seven sonication cycles were performed. Samples were placed in an ice bath for at least 1 min between each sonication cycle. Sonicated samples were centrifuged at 14,000 rpm, 4°C, for 10 min to pellet insoluble material. Supernatants were transferred to new tubes. A portion of the sample was set aside to confirm optimal chromatin shearing by agarose gel electrophoresis.

Preclear

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Twenty µl of Dynabeads (Invitrogen) were blocked with 0.5 mg/ml BSA in PBS. Blocked Dynabeads were subsequently applied to each sonicated sample and rotated for 1 hr at 4°C. Samples were placed on a magnet stand for 1 min and precleared supernatant was transferred to a new tube; 5% of the sample was removed and stored at –80°C as input. Antibody and fresh 1× protease inhibitors were added to each sample. Samples were rotated overnight at 4°C.

Pulldown

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Samples were centrifuged at 14,000 rpm, 4°C, for 5 min to pellet insoluble material. Supernatants were transferred to new tubes; 50 µl of Dynabeads (Invitrogen) were blocked with BSA 5 mg/ml in PBS. Blocked Dynabeads were subsequently applied to each sample and rotated for 6 hr at 4°C.

Stringency washes

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All wash buffers were kept ice cold during stringency washes. Samples were washed eight times with RIPA Buffer (10 mM Tris-HCl pH 7.5, 140 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1% SDS, 0.1% sodium deoxycholate, 2× protease inhibitors), two times with LiCl Buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 250 mM LiCl, 0.5% NP-40, 0.5% sodium deoxycholate, 2× protease inhibitors), two times with TE Buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 2× protease inhibitors).

Elution and reversing crosslinks

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One-hundred µl of Elution Buffer (10 mM Tris-HCl pH8.0, 5 mM EDTA, 300 mM NaCl, 0.1% SDS) was added to beads. Two µl of RNase A (Thermo Fisher, Cat#EN531) was added to each ChIP and input sample and incubated at 37°C for 30 min with gentle agitation. Ten µl of Proteinase K (Thermo Fisher, Cat# 25530049) was added to each sample and incubated at 37°C for 1 hr with gentle agitation. Crosslinks were reversed overnight at 65°C with gentle agitation. ChIP DNA was purified with a Qiagen MinElute PCR Purification kit (Cat#28004).

ChIP-seq library construction and sequencing

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ChIP-seq libraries were prepared using NEBNext ChIP-Seq Library Prep Reagent Set for Illumina (New England BioLabs, Cat# E6240). High-throughput sequencing was performed on Illumina HiSeq 2500 for single-end 50 bp reads or Illumina NovaSeq 6000 for paired-end 50 bp reads.

ChIP-Rx-seq in zebrafish embryos

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ChIP-Rx was adapted from Orlando et al., 2014, for H3K27me3 ChIP in zebrafish embryos treated with DMSO or PRT4165. Crosslinked Drosophila melanogaster S2 cells (ATCC Cat# CRL-1963) were spiked into resuspended zebrafish embryo pellets at a ratio of 5:1 (zebrafish cells: S2 cells). ChIP-Rx was subsequently performed in the same way as described above.

ChIP in zebrafish sperm

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ChIP in zebrafish sperm was conducted as described (Murphy et al., 2018).

qPCR was carried out using 2× SsoAdvanced Universal SYBR Green Supermix (Biorad Cat# 1725270) and a Biorad CFX real-time thermal cycler.

Oligonucleotides

See Table 1 for oligonucleotides used for ChIP-qPCR.

Table 1
Oligonucleotide sequences used for ChIP-qPCR for amplifying promoter regions.
Target promoterDirectionSequence (5’→3’)
pax6aForwardctccggatccgaatcacaaaactagtcc
pax6aReversecaaaggggtttgcaatctctcacaacc
vsx1Forwardcccgtcatggtggcagtttc
vsx1Reversegacagtgggatgatctgctggt
isl1Forwardgtctcccatgtcaagaaagtaaggcg
isl1Reversegccactttcccaccttcacagat
idh3gForwardcagcaagcgaacactgaccttgt
idh3gReversegcagttgggaaatacagcaaaggtacg
pcf11Forwardcgatcgtttcagagcagccaataag
pcf11Reversegtccgtcgtactttagcagagactg
lman2Forwardcccgtccgttatatctgaatatacggaag
lman2Reversectcgtaaaatgccggtgtgtcac

RNA-seq

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Total RNA was harvested from zebrafish embryos with a Qiagen Allprep kit (Cat# 80204). The Invitrogen DNA-free DNA removal kit (Cat# AM1906) was subsequently used to remove contaminating DNA from RNA samples. Intact poly(A) RNA was purified from total RNA samples (100–500 ng) with oligo(dT) magnetic beads and stranded mRNA sequencing libraries were prepared as described using the Illumina TruSeq Stranded mRNA Library Preparation Kit (RS-122–2101, RS-122–2102). Purified library quality was assessed on an Agilent Technologies 2200 TapeStation using a D1000 ScreenTape assay (Cat# 5067–5582 and 5067–5583). The molarity of adapter-modified molecules was defined by qPCR using the Kapa Biosystems Kapa Library Quant Kit (Cat# KK4824). Individual libraries were normalized to 5 nM and equal volumes were pooled in preparation for Illumina sequence analysis. High-throughput sequencing for RNAseq was performed on an Illumina HiSeq 2500. RNA-seq data displayed in Figure 3—figure supplement 1A was collected from http://www.ebiac.uk/gxa/experiments/E-ERAD-475. We would like to thank the Busch-Nentwich lab for providing RNA-seq data used in Figure 3—figure supplement 1A.

ChIP-seq analysis

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ChIP-seq Fastq files were aligned to Zv10 using Novocraft Novoalign with the following settings: -o SAM -r Random. SAM files were processed to BAM format, sorted, and indexed using Samtools (Li et al., 2009). ChIP-seq replica correlation was assessed with deepTools (Ramírez et al., 2016). Briefly, BAM files were read normalized with deeptools bamCoverage with the --normalizeUsingRPKM flag. Deeptools multiBigwigSummary bins and plotCorrelation were used to generate genome-wide correlation matrices for assessing replica correlation. ChIP-seq peak calling was accomplished using MACS2 with the following settings: callpeak -g 1.4e9 -B -q 0.01 -SPMR (Zhang et al., 2008). ChIP-seq peak calling for comparisons between Aebp2 and Jarid2 as well as ChIP-seq involving drug treatments was performed by utilizing the Multi-Replica Macs ChIPSeq Wrapper. Called peaks were annotated in R with the ChIPseeker package (R Development Core Team, 2020; Yu et al., 2015). Output bedgraph files from MACS2 were processed into bigwig files with UCSC Exe Utilities bedGraphToBigWig (Kent et al., 2010). Resulting bigwig files were loaded into IGV for genome browser snapshots of ChIP-seq enrichment (Thorvaldsdóttir et al., 2012). Heatmaps of ChIP-seq enrichment at promoter and enhancer regions were made with deepTools (Ramírez et al., 2016). A bed file of Zebrafish zv10 UCSC RefSeq genes from the UCSC Table Browser was utilized for plotting heatmaps of ChIP enrichment at promoters (Karolchik et al., 2004). Genes residing on unmapped chromosomal contigs were excluded. Enhancer heatmaps utilized a bed file of postZGA H3K4me1 ChIP-seq (Bogdanovic et al., 2012) peak summits that had been filtered to exclude promoters and unmapped chromosomal contigs. Gene ontology analysis was performed with DAVID (Huang et al., 2009).

Violin plot of ChIP-seq enrichment at promoters

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Log2(ChIP/input) data for promoter regions (±1 Kb from TSS) of interest was collected from processed bigwig files by utilizing the program Bio-ToolBox ‘get_datasets.pl’. Collected data was plotted in violin format. Unpaired t-tests with Welch’s correction were utilized to determine statistical differences in ChIP enrichment between promoter K-means clusters. Violin plot and statistical analysis (Figure 1—figure supplement 2J) were performed in GraphPad Prism version 8.3.1 using GraphPad Prism version 8.3.1 for MacOS, GraphPad Software, San Diego, CA, https://www.graphpad.com.

DNA motif analysis

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HOMER was utilized for identifying putative TF-binding motifs present at promoters and enhancers (Heinz et al., 2010). The following parameters were used on bed files of promoters and enhancers of interest: findMotifsGenome.pl danRer10 -size –250,250. Known motifs (as opposed to de novo motifs) from HOMER were presented in Figures 1 and 2.

ChIP-seq analysis involving drug treatments

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Analysis of ChIP-seq experiments involving drug treatments was performed by utilizing the Multi-Replica Macs ChIPSeq Wrapper.

RNA-seq analysis

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RNA-seq fastq files were aligned to Zv10 using STAR (Dobin et al., 2013) with the following settings: --runMode alignReads --twopassMode Basic --alignIntronMax 50000 --outSAMtype BAM SortedByCoordinate --outWigType bedGraph --outWigStrand Unstranded --clip3pAdapterSeq AGATCGGAAGAGCACACGTCTGAACTCCAGTCA. The resulting sorted BAM files were subsequently indexed using Samtools (Li et al., 2009). FeatureCounts was utilized to collect count data for zv10 genes via the following command: -T 16s 2 –largestOverlap (Liao et al., 2013). Count data for all replicates across experimental conditions were combined into a single count matrix in R (R Development Core Team, 2020). This count matrix was subsequently used to identify differentially expressed genes with the R package DESeq (Anders and Huber, 2010). RNA-seq replica correlation was assessed with deepTools (Ramírez et al., 2016). Briefly, BAM files were read normalized with deeptools bamCoverage with the --normalizeUsingRPKM flag (Ramírez et al., 2016). Deeptools multiBigwigSummary bins and plotCorrelation were used to generate genome-wide correlation matrices for assessing replica correlation (Ramírez et al., 2016).

Reprocessed ChIP-seq datasets

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PreZGA H2Az and preZGA H3K4me1 ChIP-seq data (Murphy et al., 2018) (GEO: GSE95033), postZGA H3K4me1 ChIP-seq data (Bogdanovic et al., 2012) (GEO: GSE32483), preZGA and postZGA H3K27ac ChIP-seq data (Zhang et al., 2018) (GEO: GSE114954), postZGA H3K4me3 and H3K27me3 ChIP-seq data (Zhang et al., 2014) (GEO: GSE44269), and Nanog ChIP-seq (Xu et al., 2012) (GEO: GSE34683) were downloaded from the Gene Expression Omibus and reprocessed as described above.

Whole genome bisulfite sequencing analysis

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Whole genome bisulfite sequencing (WGBS) from Potok et al., 2013 (DRA/SRA: SRP020008) was processed as described (Murphy et al., 2018).

Data access

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All sequencing datasets generated in this study have been deposited at the Gene Expression Omnibus under the accession number GSE168362.

Drug treatments

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PRT4165 (Tocris Cat#5047) was dissolved in DMSO at a concentration of 50 mM. PRT4165 was further diluted to a working concentration of 150 µM in embryo water and mixed vigorously. Zebrafish embryos were collected and immediately placed in embryo water containing 150 µM PRT4165 (or DMSO) and allowed to develop to the desired developmental stage.

Data availability

Sequencing data have been deposited in GEO under accession code GSE168362.

The following data sets were generated
    1. Hickey G
    2. Cairns BR
    (2021) NCBI Gene Expression Omnibus
    ID GSE168362. Establishment of Developmental Gene Silencing by Ordered Polycomb Complex Recruitment in Early Zebrafish Embryos.
The following previously published data sets were used
    1. Murphy P
    2. Cairns BR
    (2017) NCBI Gene Expression Omnibus
    ID GSE95033. 'Placeholder' nucleosomes underlie germline-to-embryo DNA methylation reprogramming.
    1. Bogdanović O
    2. Gómez-Skarmeta JL
    (2012) NCBI Gene Expression Omnibus
    ID GSE32483. Dynamics of enhancer chromatin signatures mark the transition from pluripotency to cell specification during embryogenesis.
    1. Zhang B
    2. Xie W
    (2018) NCBI Gene Expression Omnibus
    ID GSE114954. Widespread enhancer dememorization and promoter priming during parental-to-zygotic transition.
    1. Zhang Y
    2. Vastenhouw NL
    3. Liu XS
    (2014) NCBI Gene Expression Omnibus
    ID GSE44269. Canonical Nucleosome Organization at Promoters Forms During Genome Activation.
    1. Xu C
    2. Zon LI
    (2012) NCBI Gene Expression Omnibus
    ID GSE34683. Nanog-like Regulates Endoderm Formation through the Mxtx2-Nodal Pathway.
    1. Murphy PJ
    2. SF Wu
    3. James CR
    4. Wike CL
    5. Cairns BR
    (2013) NCBI Sequence Read Archive
    ID SRP020008. Reprogramming the Maternal Zebrafish Genome after Fertilization to Match the Paternal Methylation Pattern.

References

  1. Software
    1. R Development Core Team
    (2020) R: A Language and Environment for Statistical Computing
    R Foundation for Statistical Computing, Vienna, Austria.
  2. Book
    1. Westerfield M
    (2007)
    The Zebrafish Book: 5th Edition: A Guide for the Laboratory Use of Zebrafish
    University of Oregon Press.

Decision letter

  1. Xin Chen
    Reviewing Editor; Johns Hopkins University, United States
  2. Richard M White
    Senior Editor; Memorial Sloan Kettering Cancer Center, United States
  3. Charles K Kaufman
    Reviewer; Washington University School of Medicine, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Establishment of Developmental Gene Silencing by Ordered Polycomb Complex Recruitment in Early Zebrafish Embryos" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Richard White as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Charles K Kaufman (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1. This study first reported chromatin landscape changes in early zebrafish embryos at three distinct stages at preZGA, ZGA and postZGA stages, using a series of ChIP-seq experiments. This part of the results is of high quality, rigorously analyzed and clearly presented. The study then moved on and used a single drug to inhibit the enzyme with the hope to reduce ubH2A/Z mark. However, using the drug could lead to pleiotropic effect and cause general or secondary phenotypes. For example, this treatment leads to developmental arrest of embryogenesis at very early stage with low to no H3K27me3 mark, according to other data included in this work. Therefore, based on current results, it is hard to draw the conclusion for the causal connection from inhibiting RNF2 enzymatic function to the lack of ubH2A/Z, and subsequently to the lack of H3K27me3. In addition, due to the close interactions between these two Polycomb complexes (PRC1 and PRC2), the actual order could be reversed. Without a genetic experiment or more specific disruption assay, the conclusions should be carefully revised to be more balanced and reflect the caveats with the current results.

2. This sequential function of PRC1 and PRC2 has been reported in several other systems, including X inactivation and early embryogenesis in mice. The conceptual advance of this study could be extending this finding to another vertebrate system. However, due to the complications to draw conclusions (see #1 point), the Abstract and Discussion should be revised. Comparisons among these systems and research strategies should be further discussed.

3. H3K27me3 pattern should also be analyzed before ZGA. The authors detected a low level of H3K27me3 before ZGA using immunostaining. However, previous ChIP-seq analysis demonstrated that H3K27me3 are present on promoters before ZGA in zebrafish (e.g. PMID: 22137762).

4. It is unclear whether Aebp2 is the sole subunit for the recruitment of PRC2 to ubH2A/Z-marked regions. Paralleled analysis of the changes for Aebp2 and H3K27me3 upon RNF2 inhibitor treatment should be performed. And Aebp2-dependent and -independent regions should be separately classified for data analysis.

5. The role of PRC1 on the temporal regulation of gene expression needs to be better elucidated. The authors only analyzed the RNA-seq results for RNF2 inhibitor treated embryos postZGA. Therefore, it is currently not clear if the role of PRC1 in transcriptional repression is restricted to this stage. RNA-seq analysis of RNF2 inhibitor treated embryos on other stages are also warranted.

Reviewer #2 (Recommendations for the authors):

Some specific points:

Page 8, line 159-169. I'm confused by the authors' description of the enhancer cluster 5. By visual inspection, this cluster apparently only shared very moderate H3K4me1 and H3K27ac levels. The authors should provide a quantitative analysis similar to Figure 1S2J.

Figure 1 and 2. In Figure 1A and 2A. WGBS data of the preZGA stage embryo was shown. Yet in Figure 1B and 2B, the WGBS data was designated to postZGA embryo. Please confirm that the labeling is correct.

Page 10, line 209. Only a representative genomic region is shown in Figure 4S2B. Global analysis of the distribution changes for H3K27me3 needs to be performed in parallel with Aebp2 ChIP-seq analysis in Figure 4C.

Figure 3 Only colocalization was observed, yet functional links are not established in this study. Therefore, it is justified to claim that Aebp2 mediates H3K27me3 deposition at the current stage. Overall, the contribution of Jarid2 was not examined in the current stage.

Reviewer #3 (Recommendations for the authors):

To push the approach further, and not required for the current study, I wonder if another perturbation to test the model could be to delete key example genomic loci (or, perhaps even better, mutate Figure 1E candidate TF binding sites in the region?) to test for loss of proper Placeholder histone deposition. Or, engineer transgenic fish with developmental enhancer/promoter sequences in an ectopic location (or profile an existing transgenic around the time of ZGA) to show that they can drive establishment of the set of Placeholder chromatin/DNA marks that, in their model, delineate developmental gene loci that are silenced at the ZGA but will later be expressed.

https://doi.org/10.7554/eLife.67738.sa1

Author response

Essential revisions:

1. This study first reported chromatin landscape changes in early zebrafish embryos at three distinct stages at preZGA, ZGA and postZGA stages, using a series of ChIP-seq experiments. This part of the results is of high quality, rigorously analyzed and clearly presented. The study then moved on and used a single drug to inhibit the enzyme with the hope to reduce ubH2A/Z mark. However, using the drug could lead to pleiotropic effect and cause general or secondary phenotypes. For example, this treatment leads to developmental arrest of embryogenesis at very early stage with low to no H3K27me3 mark, according to other data included in this work. Therefore, based on current results, it is hard to draw the conclusion for the causal connection from inhibiting RNF2 enzymatic function to the lack of ubH2A/Z, and subsequently to the lack of H3K27me3. In addition, due to the close interactions between these two Polycomb complexes (PRC1 and PRC2), the actual order could be reversed. Without a genetic experiment or more specific disruption assay, the conclusions should be carefully revised to be more balanced and reflect the caveats with the current results.

We thank the reviewers for their overall positive view of the work and examination of the chromatin landscape changes via ChIP approaches. The issues raised here include: 1) whether the inhibition of Rnf2 is appropriate functional evidence, and 2) whether we are confident in the order of H2Aub1 (PRC1 action) preceding H3K27me3 (PRC2 action).

First, regarding our inhibition experiments – our work, like the other two papers published in Nature Genetics (Mei et al., 2021; Chen et al., 2021), each used one approach to lower H2Aub1. We agree that our approach could have off target effects, and now state that in the Discussion. We note, however, that the other two mouse papers likewise each took one approach to lower H2Aub1: one employed BAP overexpression, which arrested embryos at the 4-cell stage (when H3K27me3 is extremely low) and therefore could not assess whether H3K27me3 was impaired when H2Aub1 was lowered. The second employed maternal, but not paternal, genetic disruption of PRC1, which showed a transient lowering of H2Aub1 and H3K27me3, but restoration by the morula stage. Therefore, each approach has its advantages and limitations, which we now discuss in the revised Discussion. However, together, all three papers provide complementary evidence for the causal relationship.

We respectfully disagree that current evidence supports the notion that “the actual order (of PRC1/2 complexes) could be reversed”. Our evidence of PRC1/H2Aub1 preceding PRC2/H3K27me3 does not rely solely on the PRT4165 inhibitor experiment – it is fundamentally supported by the temporal relationships of the modifications. In preZGA embryos we clearly detect H2Aub1 by western and IF approaches, and importantly detect robust ChIP specifically at thousands of developmental genes. In contrast, we and almost all other investigators detect no H3K27me3 by Western or IF, nor can we and almost all others detect it at any genes. As detailed in point #3 below – the one paper that has claimed preZGA H3K27me3 (Lindemann et al., 2011) showed low H3K27me3 at only 83 genes during preZGA through ZGA, not at the thousands of developmental genes that bear H2Aub1 during preZGA through ZGA. Thus, the vast majority of the evidence points strongly to H3K27me3 being virtually absent during preZGA, whereas we show the clear presence of H2Aub1, and its residence at large numbers of developmental genes during preZGA.

We will note here (and briefly in the Discussion) that the lack of H3K27me3 has no measurable effect on the progression of zebrafish embryos through gastrulation, and through 24hpf, at which they display a normal body plan and organogenesis (though fail by 48hrs). This was determined in work published by L. Kamminga and colleagues (Rougeot et al., Development 2019; on which we were co-authors) using maternal zygotic ezh2 null mutations. Thus, H3K27me3 appears to play only minor roles until late stages of development, where it acts in a canonical manner.

We agree with the reviewers that a complementary approach would help to validate our results with the inhibitor, and we expended significant effort on this issue during revision. Regarding alternative inhibitors, PRT4165 is currently the only commercially available inhibitor of Rnf2. In the absence of alternative inhibitors, we attempted genetic approaches, beginning with the expression of a minimal version of the H2A deubiquitinase complex, PR-DUB in early zebrafish embryos. Here, single-cell stage zebrafish embryos were microinjected with mRNA encoding the two essential subunits of PR-DUB, bap1 and asxl2, to disrupt H2A/Zub1. As a negative control we utilized a point mutant of bap1 that abolishes catalytic activity. Injected embryos were allowed to develop to 4hpf and subsequently subjected to western blot analysis to assay H2Aub1 abundance. Unfortunately, this approach yielded no appreciable change in bulk levels of H2Aub1 compared to control, despite attempting multiple different doses of the bap1 and asxl2 mRNAs. Importantly, we validated (by an installed V5 tag) by Western analysis that the proteins were each expressed. Here, we reason that additional factors (beyond the two we expressed) might be needed to create a fully active (or properly targeted) BAP/ASXL2 complex and/or that inhibitors of BAP1 might be present that prevented enzymatic activity. Regardless, the inability to greatly lower H2A/Zub1 precluded further analyses.

As a tertiary approach to lower H2A/Zub1, we micro-injected single-cell stage zebrafish embryos with a mutant h2af/z mRNA encoding and form of H2A.Z that cannot be ubiquitinated. We also injected wild-type h2af/z as a negative control. Unfortunately, our Western analyses showed that we were not able to arrive at protein levels derived from the injected species that approached the large stores of WT H2A.Z protein present in the embryo, so competition was not achieved, and H2A/Zub1 was not lowered.

Taken together, our work parallels the work earlier this year from two groups who each took one effective approach to lower H2AUb1: ours with PRT4165 and theirs with BAP1/ASXL1 overexpression or a PRC1 maternal-specific mutant approach. Each approach has advantages and caveats, and the reviewer rightly points out that our approach can have off target affects. Therefore, we have included this caveat and how it may affect our interpretations accordingly in the revised paper (see Discussion, and also point #2, below). Specifically, in the Discussion of the revised manuscript we state:

“However, it is important to note that PRT4165 may have off-target effects aside from inhibition of Rnf2 which may contribute to our observations.” (page 16, line #336-337)

and

“However, as PRT4165 was the only approach we employed that successfully removed the vast majority of H2Aub1, future studies involving orthogonal approaches to elicit Rnf2 loss in zebrafish will be needed to further validate the consequences of H2Aub1 loss in early zebrafish embryos” (page 16 and line #342-345).

2. This sequential function of PRC1 and PRC2 has been reported in several other systems, including X inactivation and early embryogenesis in mice. The conceptual advance of this study could be extending this finding to another vertebrate system. However, due to the complications to draw conclusions (see #1 point), the Abstract and Discussion should be revised. Comparisons among these systems and research strategies should be further discussed.

There is indeed precedent for that order in X chromosome inactivation in ES cells, and we have cited that work in the revision. In addition, the work published earlier this year in Nature Genetics that determines the order for the X chromosome and developmental genes in mouse embryos is now discussed in the revised version (pages 16-18, lines 346-383, and see below). However, at the time when we submitted our work, the order of polycomb complex action at developmental genes was not known. Our manuscript was submitted and progressed to full review at eLife (and was posted on the BioRxiv) prior to the work in mice being published in Nature Genetics. Therefore, we hope that the novelty and conceptual advance of the work will be assessed based on our date of submission.

The recent mouse work (in Nature Genetics) on H2AUb inhibition in mice strongly supports our work in zebrafish, and we have now included those papers in the Discussion of our revised paper (pages 16-18, lines 346-383). Indeed, their methods for reducing H2AUb1 in mouse embryos led to upregulation of developmental genes, in keeping with our observations. Regarding novelty, our work goes much further, as only our paper addresses the mechanism by which H2AUb1 is ‘read’ in the early embryo by a non-canonical PRC2 complex (bearing Aebp2) to place H3K27me3 on developmental genes. Therefore, we believe our work has conceptual advances both in the areas of overlap, and also in unique aspects related to PRC2 targeting.

3. H3K27me3 pattern should also be analyzed before ZGA. The authors detected a low level of H3K27me3 before ZGA using immunostaining. However, previous ChIP-seq analysis demonstrated that H3K27me3 are present on promoters before ZGA in zebrafish (e.g. PMID: 22137762).

The paper referred to (Lindeman et al., 2011; PMID 22137762) is an interesting and influential paper in the field, however some of the interpretations have required revisiting. The Lindeman paper does claim that H3K27me3 can be found on developmental promotes during preZGA. However, the key questions are – how many promoters, what fraction of promoters, and how robust is the marking? The Lindeman paper and follow-up work by us and others (Zhang et al., Genome Research 2014; Zhang et al., Molecular Cell, 2018), agree that there are >3500 developmental genes marked by H3K27me3 in postZGA. However, the Lindeman data claims that only 201 genes are marked by H3K27me3 in preZGA embryos (DocumentS2, Table S1; Venn Diagrams in Lindeman et al., 2011; PMID 22137762). Also, one expects that if the purpose of that H3K27me3 involves gene poising, it should be stable and last into postZGA – and when one queries their data and asks how many genes bear H3K27me3 from preZGA through postZGA, the number drops to 83 genes. These 83 genes are extremely few compared to the >3,500 genes marked in postZGA embryos by H3K27me3 in the Lindeman data and Zhang data. Also, these 83 H3K27me3 genes show very low median ChIP signal during preZGA (less than 2-fold enrichment) and only show clear (>4-fold enrichment) during postZGA (see Author response image 1).

Author response image 1

Importantly, we emphasize that H3K27me3 ChIP experiments from the Schier lab (Vastenheow et al., Nature, 2010), Wei Xie’s lab (Zhang et al., Molecular Cell, 2018) and our lab (described as ‘data not shown’ in Murphy et al., Cell 2018) did not reveal any peaks for H3K27me3 during preZGA. Furthermore, both our IF data (this work; Figure 1, FigureS1, Panel B) and Zhang et al., 2018 (Supplemental Figure S1A) fail to show H3K27me3 signal preZGA. Taken together, work from several labs supports the view that there is extremely low H3K27me3 in the genome at preZGA, which has resulted either in a lack of ChIP peaks preZGA (Schier, Xie and Cairns labs) or detection of very low H3K27me3 at a very small number of loci (~83-201; Lindeman et al). However, all groups agree that after ZGA, H3K27me3 increases dramatically, enabling clear detection at >3500 genes. Therefore, the much larger questions – addressed in our work – are how >3500 developmental genes are silenced at ZGA (we show it is H2AUb1, not H3K27me3), and how those >3500 obtain high H3K27me3 (we show it is by non-canonical PRC2 binding to H2AUb1).

4. It is unclear whether Aebp2 is the sole subunit for the recruitment of PRC2 to ubH2A/Z-marked regions. Paralleled analysis of the changes for Aebp2 and H3K27me3 upon RNF2 inhibitor treatment should be performed. And Aebp2-dependent and -independent regions should be separately classified for data analysis.

We thank the reviewers for raising these important points, which we have addressed. Previous data including the very nice recent cryo-EM structure of the non-canonical PRC2 complex bound to H2Aub1+ nucleosomes from Eva Nogales’ group (Kasinath et al., Science 2021), indicates that Aebp2 is not the sole subunit involved in recruitment of PRC2 to H2A/Zub1. Rather, Aebp2 and Jarid2 subunits of ncPRC2 both contribute to recruitment of PRC2 via direct contacts with H2A/Zub1. However, we note that the PRC2 complex can be isolated from cells or produced in a recombinant form with Aebp2, but lacking Jarid2 (Cao et al., Science 2002; Ciferri et al., eLife 2012) – suggesting that these proteins are not together in all PRC2 complexes, and that Aebp2 assembles well into PRC2 without Jarid2. Accordingly, we have modified the text of our manuscript to clarify that Aebp2 is not the sole PRC2 subunit involved in recruitment to H2A/Zub1+ nucleosomes.

To investigate the overlap between Aebp2 and Jarid2 concept further, we performed ChIP-seq of Jarid2 in postZGA embryos. We have included our Jarid2 ChIP profiling into our manuscript in Figure 3 —figure supplement 2. Although the ChIP signal and dynamic range were modest, the majority of Jarid2 sites overlap with Aebp2-bound sites, supporting the notion that these two proteins may work together at a portion of PRC2-bound sites. However, given the modest ChIP signal of Jarid2, we do not believe that highly quantitative statements about binding selectivity can be made.

Next, as requested – we reanalyzed our ChIP-seq data for Aebp2 and H3K27me3 upon Rnf2 inhibition in a paralleled manner. This was an important addition to the paper. This paired Aebp2/H3K27me3 analysis has been incorporated into new Figure 4 and reflected in the text of the manuscript. The clear result is that the same places that lose Aebp2 binding (following the loss of H2AUb1) also lose H3K27me3. We believe this revised way of presenting the data conveys the result much clearer than the previous version of Figure 4.

5. The role of PRC1 on the temporal regulation of gene expression needs to be better elucidated. The authors only analyzed the RNA-seq results for RNF2 inhibitor treated embryos postZGA. Therefore, it is currently not clear if the role of PRC1 in transcriptional repression is restricted to this stage. RNA-seq analysis of RNF2 inhibitor treated embryos on other stages are also warranted.

The reviewers raise an interesting question, which also addresses whether the removal of H2AUb from developmental genes might enable their activation during preZGA, which would confer to developmental genes the status/privilege of ‘minor wave’ genes. To address this question, we performed RNA-seq at preZGA and ZGA timepoints with embryos treated with the RNF2 inhibitor (PRT4165) or vehicle control (DMSO) and examined whether precocious transcription of developmental genes occurred prior to post ZGA. This work has been incorporated into our revised manuscript in Figure 4 —figure supplement 3. GO-term analysis of genes meeting our threshold criteria (p-value ≤0.01, fold change ≥1.5) uncovered no enrichment of developmental gene activation upon RNF2 inhibition prior to postZGA. Instead, enriched GO-terms of upregulated transcripts correspond to RNA-binding and ribosomal protein genes. Thus, chromatin de-repression (via H2Aub1 removal) does not cause precocious transcriptional activation of developmental genes until postZGA (4hpf).

Here, we reason that the observed precocious transcription of developmental genes at postZGA in embryos treated with PRT4165 relies on binding of opportunistic transcriptional activators. However, these activators may be temporally constrained during preZGA by: 1) translation of mRNA encoding these factor(s) and/or 2) chromatin remodeling events needed to expose binding sites for opportunistic activators.

Reviewer #2 (Recommendations for the authors):

Some specific points:

Page 8, line 159-169. I'm confused by the authors' description of the enhancer cluster 5. By visual inspection, this cluster apparently only shared very moderate H3K4me1 and H3K27ac levels. The authors should provide a quantitative analysis similar to Figure 1S2J.

We apologize for the confusion, which also raised a comment from Reviewer #1. There are two points of clarification to make here. First, we now clarify in the manuscript that DNAme patterns are essentially identical/static from 2.5hpf (preZGA) to 4.3hpf (postZGA). On Page 5, line 100 we now state:

“For our comparisons to DNA methylation (DNAme), we note that DNAme patterns are reprogrammed between fertilization and the preZGA (2.5hpf) timepoint (Potok et al., 2013; Jiang et al., 2013), but remain static in zebrafish embryos from 2.5 hpf (preZGA) to 4.3 hpf (postZGA). Therefore, for brevity we chose to display only a single timepoint for DNAme data in subsequent figures, which is representative of all developmental stages examined by the genomics approaches in this work”.

Second, our reference to H3K27ac being very high refers specifically to enhancer cluster 5 in the postZGA state. Thus, H3K27me3 is very high, and these loci are DNA methylated (DNAme shown for preZGA, but identical/methylated in postZGA).

Figure 1 and 2. In Figure 1Aand2A. WGBS data of the preZGA stage embryo was shown. Yet in Figure 1B and 2B, the WGBS data was designated to postZGA embryo. Please confirm that the labeling is correct.

We thank the reviewer for their comment. The labeling is correct. As noted above, the DNA methylation state remains constant at enhancers and promoters from preZGA to postZGA.

Page 10, line 209. Only a representative genomic region is shown in Figure 4S2B. Global analysis of the distribution changes for H3K27me3 needs to be performed in parallel with Aebp2 ChIP-seq analysis in Figure 4C.

We agree and have addressed: Figure 4 has been largely revised to examine the changes in localization of Aebp2 and H3K27me3 in parallel, as also requested in the list of unified comments.

Figure 3 Only colocalization was observed, yet functional links are not established in this study. Therefore, it is justified to claim that Aebp2 mediates H3K27me3 deposition at the current stage. Overall, the contribution of Jarid2 was not examined in the current stage.

Here, our functional link is that when H2Aub1 is prevented (via PRT4165), Aebp2 fails to occupy all sites that formerly had H2Aub1, and H3K27me3 is not deposited at those same loci – which is now much more clearly displayed through the revised analyses present in Figure 4.

The reviewer raises an interesting issue in their request to examine Jarid2. Here, although the Jarid2 antibody and resulting ChIP signal was not as robust as with our potent Aebp2 antibody (limiting highly quantitative conclusions) the majority of Jarid2 targets were also Aebp2 targets – consistent with Aebp2 and Jarid2 working together for H2Aub1 recognition (Figure 3 —figure supplement 2).

Reviewer #3 (Recommendations for the authors):

To push the approach further, and not required for the current study, I wonder if another perturbation to test the model could be to delete key example genomic loci (or, perhaps even better, mutate Figure 1E candidate TF binding sites in the region?) to test for loss of proper Placeholder histone deposition. Or, engineer transgenic fish with developmental enhancer/promoter sequences in an ectopic location (or profile an existing transgenic around the time of ZGA) to show that they can drive establishment of the set of Placeholder chromatin/DNA marks that, in their model, delineate developmental gene loci that are silenced at the ZGA but will later be expressed.

We appreciate the reviewer’s insightful comments. In the future we will use Crispr-CAS9 mediated targeting of transcription factors and histone modifications to test these interesting questions.

https://doi.org/10.7554/eLife.67738.sa2

Article and author information

Author details

  1. Graham JM Hickey

    Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9665-839X
  2. Candice L Wike

    Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    Contribution
    Formal analysis, Investigation, Methodology, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
  3. Xichen Nie

    Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  4. Yixuan Guo

    Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    Contribution
    Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1940-1931
  5. Mengyao Tan

    Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  6. Patrick J Murphy

    1. Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    2. Department of Biomedical Genetics, Wilmot Cancer Center, University of Rochester School of Medicine, Rochester, United States
    Contribution
    Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  7. Bradley R Cairns

    Howard Hughes Medical Institute, Department of Oncological Sciences and Huntsman Cancer Institute, University of Utah School of Medicine, Salt Lake City, United States
    Contribution
    Conceptualization, Funding acquisition, Methodology, Project administration, Resources, Supervision, Writing – original draft, Writing – review and editing
    For correspondence
    brad.cairns@hci.utah.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-9864-8811

Funding

Howard Hughes Medical Institute (Cairns)

  • Bradley R Cairns

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Brian Dalley and the Huntsman Cancer Institute High-Throughput Sequencing Shared Resource for sequencing data. We thank Tim Parnell, Chongil Yi, and Jingtao Guo for helpful discussions regarding bioinformatic analysis. We thank David Grunwald and Rodney Stewart for providing helpful feedback during manuscript preparation. We thank all current and former members of the Cairns lab for their perspectives on this project. BRC is an investigator with the Howard Hughes Medical Institute. CW was funded by the T32 Developmental Biology Training Grant (59202072), 4DNucleome (NIH Common Fund) NBR – 92275293 S9001779, and HFSP RGP0025/2015. Imaging was performed at the University of Utah Microscopy Core (1S10RR024761-01). Financial support was received from the Howard Hughes Medical Institute and the Huntsman Cancer Institute core facilities (CA24014).

Ethics

All of our work on zebrafish is authorized and overseen by our institutional animal care and use committee (IACUC). This work is authorized by the current IACUC Protocol #IACUC protocol 20-04011 'Germ Cell Epigenetics in Zebrafish'. We have no surgical procedures, and followed AVMA Guidelines involving a 2-step euthanasia for embryos and larvae which includes rapid chilling followed by immersion in a dilute sodium hypochlorite solution (discussed on page 89 of the 2020 Guidelines).

Senior Editor

  1. Richard M White, Memorial Sloan Kettering Cancer Center, United States

Reviewing Editor

  1. Xin Chen, Johns Hopkins University, United States

Reviewer

  1. Charles K Kaufman, Washington University School of Medicine, United States

Publication history

  1. Received: February 21, 2021
  2. Preprint posted: March 16, 2021 (view preprint)
  3. Accepted: December 29, 2021
  4. Accepted Manuscript published: January 4, 2022 (version 1)
  5. Version of Record published: January 19, 2022 (version 2)

Copyright

© 2022, Hickey et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Graham JM Hickey
  2. Candice L Wike
  3. Xichen Nie
  4. Yixuan Guo
  5. Mengyao Tan
  6. Patrick J Murphy
  7. Bradley R Cairns
(2022)
Establishment of developmental gene silencing by ordered polycomb complex recruitment in early zebrafish embryos
eLife 11:e67738.
https://doi.org/10.7554/eLife.67738

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    Jakub Gemperle et al.
    Tools and Resources

    CRISPR technology has made generation of gene knock-outs widely achievable in cells. However, once inactivated, their re-activation remains difficult, especially in diploid cells. Here, we present DExCon (Doxycycline-mediated endogenous gene Expression Control), DExogron (DExCon combined with auxin-mediated targeted protein degradation), and LUXon (light responsive DExCon) approaches which combine one-step CRISPR-Cas9-mediated targeted knockin of fluorescent proteins with an advanced Tet-inducible TRE3GS promoter. These approaches combine blockade of active gene expression with the ability to re-activate expression on demand, including activation of silenced genes. Systematic control can be exerted using doxycycline or spatiotemporally by light, and we demonstrate functional knock-out/rescue in the closely related Rab11 family of vesicle trafficking regulators. Fluorescent protein knock-in results in bright signals compatible with low-light live microscopy from monoallelic modification, the potential to simultaneously image different alleles of the same gene, and bypasses the need to work with clones. Protein levels are easily tunable to correspond with endogenous expression through cell sorting (DExCon), timing of light illumination (LUXon), or by exposing cells to different levels of auxin (DExogron). Furthermore, our approach allowed us to quantify previously unforeseen differences in vesicle dynamics, transferrin receptor recycling, expression kinetics, and protein stability among highly similar endogenous Rab11 family members and their colocalization in triple knock-in ovarian cancer cell lines.

    1. Chromosomes and Gene Expression
    Sarah Lensch et al.
    Research Article

    In mammalian cells genes that are in close proximity can be transcriptionally coupled: silencing or activating one gene can affect its neighbors. Understanding these dynamics is important for natural processes, such as heterochromatin spreading during development and aging, and when designing synthetic gene regulation circuits. Here, we systematically dissect this process in single cells by recruiting and releasing repressive chromatin regulators at dual-gene synthetic reporters, and measuring how fast gene silencing and reactivation spread as a function of intergenic distance and configuration of insulator elements. We find that silencing by KRAB, associated with histone methylation, spreads between two genes within hours, with a time delay that increases with distance. This fast KRAB-mediated spreading is not blocked by the classical cHS4 insulators. Silencing by histone deacetylase HDAC4 of the upstream gene can also facilitate background silencing of the downstream gene by PRC2, but with a days-long delay that does not change with distance. This slower silencing can sometimes be stopped by insulators. Gene reactivation of neighboring genes is also coupled, with strong promoters and insulators determining the order of reactivation. Our data can be described by a model of multi-gene regulation that builds upon previous knowledge of heterochromatin spreading, where both gene silencing and gene reactivation can act at a distance, allowing for coordinated dynamics via chromatin regulator recruitment.