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. Author manuscript; available in PMC: 2014 Sep 26.
Published in final edited form as: Cell. 2013 Sep 26;155(1):10.1016/j.cell.2013.08.061. doi: 10.1016/j.cell.2013.08.061

Hira-dependent histone H3.3 deposition facilitates PRC2 recruitment at developmental loci in ES cells

Laura A Banaszynski 1,#, Duancheng Wen 2,#, Scott Dewell 3, Sarah J Whitcomb 1, Mingyan Lin 4, Nichole Diaz 1, Simon J Elsässer 1, Ariane Chapgier 5, Aaron D Goldberg 1, Eli Canaani 6, Shahin Rafii 2, Deyou Zheng 4,7, C David Allis 1,*
PMCID: PMC3838450  NIHMSID: NIHMS525479  PMID: 24074864

SUMMARY

Polycomb repressive complex 2 (PRC2) regulates gene expression during lineage specification through trimethylation of lysine 27 on histone H3 (H3K27me3). In Drosophila, polycomb binding sites are dynamic chromatin regions enriched with the histone variant H3.3. Here we show that in mouse embryonic stem cells (ESCs) H3.3 is required for proper establishment of H3K27me3 at the promoters of developmentally regulated genes. Upon H3.3 depletion, these promoters show reduced nucleosome turnover measured by deposition of de novo synthesized histones, and reduced PRC2 occupancy. Further, we show H3.3-dependent interaction of PRC2 with the histone chaperone, Hira, and that Hira localization to chromatin requires H3.3. Our data demonstrate the importance of H3.3 in maintaining a chromatin landscape in ESCs that is important for proper gene regulation during differentiation. Moreover, our findings support the emerging notion that H3.3 has multiple functions in distinct genomic locations that are not always correlated with an “active” chromatin state.

INTRODUCTION

Chromatin in embryonic stem cells (ESCs) is characterized by an open state thought to allow for rapid transcription changes that accompany the transition from pluripotency to differentiated cell states (Gaspar-Maia et al., 2011; Meshorer et al., 2006). ESC chromatin displays an overall less condensed structure with relative absence of constitutive heterochromatin and genome-wide low-level transcriptional activity. The chromatin landscape of ESCs reflects this genome activity, displaying abundant levels of histone post-translational modifications associated with active transcription, such as trimethylation of lysine 4 on histone H3 (H3K4me3) and acetylation of histones H3 and H4.

Actively transcribed regions of metazoan genomes are also enriched with the histone H3 variant H3.3. H3.3 contains only 4-5 amino acid changes from canonical H3 isoforms H3.1 and H3.2 (Figure S1A), yet displays distinct properties in regulation and timing of expression and mode of incorporation into chromatin (Filipescu et al., 2013). While canonical H3 is expressed only during S phase, showing replication-dependent incorporation by its chaperone, CAF-1, H3.3 is expressed throughout the cell cycle and shows distinct patterns of genomic enrichment mediated by at least two independent chaperone systems, with Hira mainly facilitating genic deposition and Atrx/Daxx responsible for repeat regions. While H3.3 deposition has long been associated with gene activation, surprisingly, our recent genome-wide profiling demonstrated that H3.3 is also enriched at the promoters of developmentally regulated genes in ESCs (Goldberg et al., 2010), which are typically silenced or transcribed at low levels in ESCs.

Promoters of developmentally regulated genes in ESCs are defined by the presence of both activation-associated H3K4me3 and repression-associated H3K27me3. These dually modified regions, dubbed “bivalent domains” (Bernstein et al., 2006), are proposed to be poised for activation upon differentiation. Promoters of bivalent genes are bound by RNA polymerase II phosphorylated on serine 5 (RNAPII S5p) associated with transcriptional initiation and recruitment of the H3K4 methyltransferase (Brookes and Pombo, 2009; Weake and Workman, 2010). These promoters are also bound by polycomb repressive complex 2 (PRC2), which contains the enzymatic activity responsible for H3K27me3 incorporation into chromatin (Margueron and Reinberg, 2011). In Drosophila, H3.3 is enriched at polycomb response elements (PREs), regions of PRC2 binding associated with highly dynamic chromatin (Mito et al., 2007). That, coupled with our own finding that H3.3 localizes to developmental promoters in ESCs that are not highly transcriptionally active, suggests that H3.3 may have a role at lowly transcribed genes.

Here, we show that H3.3 promotes establishment of the bivalent chromatin landscape in ESCs, with H3.3 depletion leading to reduced levels of H3K27me3 enrichment at the promoters of developmentally regulated genes, altered ESC developmental potential, and misregulation of bivalent genes upon differentiation. Our data suggest that rather than acting as a preferred substrate for PRC2 activity, H3.3 facilitates a dynamic chromatin environment that allows for optimal PRC2 binding and activity. Our studies underscore the intriguing possibility that chromatin dynamics themselves play an important role in the establishment of chromatin landscapes, with subsequent effects on differentiation and developmental processes.

RESULTS

H3.3 Depletion Results in Globally Reduced Chromatin Dynamics at Promoters and Reduced H3K27me3 Enrichment at Promoters of Developmentally Regulated Genes

In mammals, histone H3.3 is encoded by two different genes (h3f3a and h3f3b) whose transcription results in identical protein products (H3.3A and H3.3B) (Wellman et al., 1987). We used a designed zinc finger nuclease (ZFN) to knock out both alleles of h3f3b, along with two independent short hairpin RNAs (shRNA) targeted against the h3f3a transcript, to generate two H3.3-depleted ESC lines (H3.3 KD1, H3.3 KD2; Figures 1A and 1B, Figures S1B and S1C). As a control, we generated an H3.3B-HA epitope tagged ESC line carrying a nontargeting shRNA sequence.

Figure 1. H3.3 Depletion Results in Global Loss of Nucleosome Turnover and Reduced H3K27me3 Enrichment at Promoters of Developmentally Regulated Genes.

Figure 1

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A) H3.3 transcript levels in control and H3.3-depleted ESCs (n = 3 ± s.d.). B) Immunoblot of H3.3 and total H3 in nuclear extracts from control and H3.3-depleted ESCs. Direct blue stain as loading control. C) H3.3 enrichment profiles at the transcription start site (TSS) of genes classified by bivalency (Mikkelsen et al., 2007). D) Box plots represent CATCH-IT normalized to H3gen ChIP-seq at promoters in control and H3.3 KD ESCs. The bottom and top of the boxes correspond to the 25th and 75th percentiles, and the internal band is the 50th percentile (median). The plot whiskers correspond to 1.5× interquartile range. P-values were calculated using Wilcoxon tests. ChIP-qPCR analyses of (E) H3K4me3 and (F) H3K27me3 at representative housekeeping and bivalent genes in control and H3.3-depleted ESCs. Error bars represent s.d. from one experiment (n = 3). Data are representative of at least 3 independent ChIP experiments. Genome browser representations of H3.3, H3K4me3, H3K27me3, and H3gen profiles in control and H3.3 KD1 ESCs for (G) two housekeeping genes, Actin (left) and Rps19 (right), and (H) two developmentally regulated bivalent genes in ESCs, Tbx3 (left) and Lhx5 (right). Read counts normalized to total number of reads for each data set. I) ChIP-seq profiles of H3K4me3 (left) and H3K27me3 (right) at the TSS of bivalent genes in control and H3.3 KD1 ESCs. Data are represented as reads per mapped million per 50 bp bin. J) Ratio of H3K27me3 peak heights in control vs. H3.3 KD1 ESCs at promoter (±2 kb from TSS), promoter-distal (−50 kb from TSS to +5 kb from transcription end site (TES) excluding promoter peaks), and intergenic (all other) peaks, defined by distance from nearest annotated genes. X-axis values >1 indicate reduced H3K27me3 enrichment upon H3.3 depletion. See also Figure S1 and Table S1.

As studies from Henikoff et al have linked regions of high nucleosome turnover in Drosophila to H3.3 deposition (Mito et al., 2007), and our own previous studies demonstrated enrichment of H3.3 at bivalent loci in ESCs as defined by Mikkelsen et al (Goldberg et al., 2010; Mikkelsen et al., 2007), we first tested whether bivalent loci were areas of active turnover in mammalian systems, and whether chromatin dynamics were altered in H3.3-depleted ESCs. We first confirmed the presence of H3.3 at bivalent loci in ESCs (Figure 1C). To measure nucleosome turnover, we used CATCH-IT (covalent attachment of tags to capture histones and identify turnover) (Deal et al., 2010). CATCH-IT measures nucleosome dynamics (i.e., incorporation of newly synthesized histones) via a pulse of metabolite labeling, capture of labeled, chromatinized H3/H4 histones, and sequencing of associated DNA fragments. The CATCH-IT profile in control ESCs was similar to that of H3.3 enrichment and correlated well with gene expression (Figure S1D). Consistent with the presence of H3.3 at bivalent loci, bivalent promoters showed nearly identical nucleosome dynamics to promoters modified only with H3K4me3 (Figure 1D) after normalizing for total number of non-duplicate mapped reads and nucleosome occupancy levels (as determined by ChIP-seq using a general H3 antibody). Analysis of chromatin dynamics in the absence of H3.3 showed reduced nucleosome turnover at both active and bivalent promoters (Figure 1D).

To assess the chromatin landscape at promoters in ESCs, we performed chromatin immunoprecipitation using antibodies recognizing H3K4me3 and H3K27me3, followed by qPCR analysis (ChIP-qPCR) of selected housekeeping and developmentally regulated genes in both control and H3.3-depleted ESCs. As H3.3 has traditionally been associated with actively transcribed chromatin enriched with H3K4me3, we expected to observe a decrease in H3K4me3 enrichment at promoters of housekeeping genes in the absence of H3.3. Surprisingly, we observed no difference in H3K4me3 enrichment at highly expressed housekeeping genes such as beta-actin (Actin) and ribosomal protein S19 (Rps19) in the absence of H3.3 (Figure 1E). This unexpected finding is in agreement with our previous observation that knockout of Hira, the chaperone responsible for genic H3.3 deposition, resulted in minimal transcriptional changes in ESCs (Goldberg et al., 2010). Additionally, H3.3 depletion had little effect on H3K4me3 levels at developmentally regulated bivalent genes (Figure 1E). In contrast, our ChIP-qPCR analysis of H3K27me3 at bivalent genes showed a clear reduction of enrichment in the absence of H3.3 (Figure 1F, see Figures S1E and S1F for antibody specificity). This result was reproducible over independent experiments using bulk or clonally selected knockdown cells and a second H3K27me3 antibody, and required depletion of both H3.3A and H3.3B (Figures S1G and S1H).

We next used ChIP followed by massively parallel DNA sequencing (ChIP-seq) to determine whether H3K27me3 enrichment was reduced genome-wide at developmentally regulated genes in the absence of H3.3. Comparison of ChIP-seq enrichment at housekeeping genes Actin and Rps19 (Figure 1G) and bivalent genes Tbx3 and Lhx5 (Figure 1H) confirmed our ChIP-qPCR observations in that while H3K4me3 enrichment was unperturbed, H3K27me3 enrichment at bivalent genes was strikingly reduced upon H3.3 depletion. Reduced H3K27me3 enrichment cannot be attributed to reduced nucleosome occupancy upon depletion of H3.3, as overall levels of H3 (H3gen) showed little variation at either housekeeping genes or bivalent genes (Figures 1G and 1H).

To identify all bivalent genes in ESCs, we used MACS (Zhang et al., 2008) to call H3K4me3 and H3K27me3 peaks from our control data sets and then grouped genes according to enrichment at the promoter region (±2 kb of the transcription start site (TSS)). We identified 15,670 genes in control ESCs containing promoters enriched with H3K4me3 only and 2,938 genes enriched with both H3K4me3 and H3K27me3 (Table S1). The vast majority of genes enriched with H3K27me3 were also enriched with H3K4me3 (23 genes were classified as enriched with H3K27me3 only). H3K4me3 enrichment at promoters of bivalent genes was nearly identical in control and H3.3 KD1 ESCs (Figure 1I, left panel). In agreement with our ChIP-qPCR data, genome-wide analysis of promoters of these genes showed a decrease in H3K27me3 enrichment upon H3.3 depletion (Figure 1I, right panel, p = 2.2×10−16) with no loss of nucleosome occupancy (Figure S1I). Decreased H3K27me3 enrichment in H3.3 KD1 ESCs resulted in a reduction of bivalent domains, with only 1,588 such classified genes upon H3.3 depletion (Table S1).

We further studied the effect of H3.3 knockdown by analyzing H3K27me3 in control and H3.3 KD1 ESCs. Among the 3,125 H3K27me3 promoter peaks identified in both control and H3.3 KD1 ESCs, 2,081 (~67%) showed greater than 1.5-fold reduction of H3K27me3 enrichment upon H3.3 depletion (Figure 1J). By comparison, promoter-distal and intergenic regions enriched for H3K27me3 (See Figure 1 legend, 3,583 and 5,534 regions, respectively) showed little perturbation upon depletion of H3.3 (Figure 1J and Figure S1J). Differences in global H3K27me3 levels could not be distinguished by immunoblotting (Figure S1K), perhaps because promoter-proximal peaks account for only ~25% of total H3K27me3 peaks identified. Additionally, we observed increased enrichment of H3K27me3 at short tandem (AT)n repeat regions in H3.3 KD1 ESCs, along with a small increase of enrichment on the Y chromosome (Figure S1L). Gene ontology analysis using the Genomic Regions Enrichment of Annotation Tool (GREAT) (McLean et al., 2010) showed that genes displaying at least a 1.5-fold reduction of H3K27me3 at their promoters upon H3.3 depletion are enriched for neural tube mesenchyme and ear development and are associated with structural and neuronal ear defects in mouse knock-out studies (Figure S1M), suggesting that in vivo this early lineage may be particularly susceptible to disruptions in chromatin architecture in the absence of H3.3. In fact, a recent study has shown that a mutation preventing H3.3 incorporation into chromatin in zebrafish results in craniofacial abnormalities (Cox et al., 2012).

Depletion of H3.3 Allows Upregulation of Transcription Factors Essential for the Trophectoderm Lineage in Mouse ESCs

To determine whether H3.3-dependent reduction of H3K27me3 enrichment at bivalent genes resulted in their global transcriptional upregulation, we performed gene expression profiling by RNA-seq for both control and H3.3-depleted ESCs. Analysis of these data revealed modest changes in gene expression upon H3.3 depletion, with only 84 genes upregulated (Figure 2A) and 9 genes downregulated (Figure 2B) in both H3.3-depleted cell lines, as compared to control shRNA-treated ESCs (p < 0.05 with multiple testing correction, Table S2). After treatment with either shRNA, h3f3a was one of the most downregulated genes, confirming efficiency of knockdown.

Figure 2. Transcription Factors Essential for Trophectoderm Specification are Upregulated upon H3.3 Depletion.

Figure 2

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RNA-seq data sets were obtained from control and H3.3 KD ESC poly(A)-RNA, and fragments per kilobase per million mapped reads (FPKM) were calculated for all mouse Ensembl genes. Differentially expressed genes were identified using Cuffdiff with FDR < 0.05. FPKMs for genes (A) upregulated or (B) downregulated in both H3.3 KD1 and H3.3 KD2 (Venn diagrams) with respect to control are represented by box plots. The bottom and top of the boxes correspond to the 25th and 75th percentiles, and the internal band is the 50th percentile (median). The plot whiskers correspond to the 10th and 90th percentiles. P-values were calculated using Wilcoxon tests. C) Gene ontology biological process categories for genes significantly upregulated upon H3.3 knockdown from analysis performed with DAVID software (Huang et al., 2009). D) RT-qPCR of pluripotency and selected lineage marker genes in control and H3.3 KD ESCs. Data are represented as mean expression relative to Gapdh with control normalized to 1. Error bars represent standard deviation (s.d.) for an experiment performed in biological triplicate and technical duplicate, ***p<0.001. E) Immunoblot of control ESCs and H3.3 KD1 ESCs exogenously expressing C-terminally Flag-HA tagged H3.2 or H3.3. F) RT-qPCR of trophectoderm-specific transcription factors in control ESCs and H3.3 KD1 ESCs exogenously expressing either H3.2 or H3.3. Data are represented as mean expression relative to Gapdh ± s.d. (n = 3). Box plots representing ChIP-seq read counts at promoters of genes upregulated by RNA-seq (RNA-seq up) and bivalent promoters for (G) H3K4me3, (H) H3K27me3, and (I) H3K27ac in both control and H3.3 KD1 ESCs. Plots as in panels A and B, except plot whiskers correspond to 1.5× interquartile range. See also Figure S2 and Table S2.

Consistent with minimal transcriptional changes, we found that H3.3 is not required for maintenance of the mouse ESC state. H3.3-depleted ESCs showed colony morphology identical to control ESCs and normal proliferation under growth conditions promoting self-renewal (Figures S2A and S2B). In addition, protein levels of key pluripotency transcription factors Oct4 and Sox2 were unaltered (Figure S2C). After more than one month in culture, H3.3-depleted ESCs maintained a normal diploid karyotype (data not shown), suggesting that H3.3 is not required for proper chromosome segregation in this cell type, despite its proposed role as an S-phase place marker for the centromeric histone variant CENP-A (Dunleavy et al., 2011) and defective chromosome segregation observed in h3f3b−/− mice (Bush et al., 2013).

Gene ontology analysis of upregulated transcripts by RNA-seq revealed the most significant overlap with genes encoding glycosylated proteins (p < 10−4) and genes expressed in the placenta (p < 10−4) (Figure 2C). This is intriguing given that ESCs are typically restricted from differentiation into extraembryonic trophectoderm that gives rise to the placenta (Rossant, 2008). Upregulated transcripts were validated by quantitative PCR with reverse transcription (RT-qPCR) (Figures 2D and S2D) and included transcription factors essential for trophectoderm establishment (e.g., Cdx2, Hand1, Dlx3, and Ets1) and multiple imprinted genes (e.g., Meg3, Peg10, and Igf2), many of which function in extraembryonic tissues (Hudson et al., 2010). We observed no upregulation of endoderm (Gata4, Gata6, Sox17), mesoderm (Nodal, Brachury), or ectoderm (Nestin, Fgf5) lineage markers (Figure 2D). Upregulation of Cdx2 and Hand1 in the absence of a decrease in expression of pluripotency factors was confirmed using an independent clonal knockout of h3f3b (Figure S2E) and required loss of both H3.3A and H3.3B (Figure S2F). Importantly, exogenous expression of H3.3, but not H3.2, was able to partially reduce trophectoderm-specific transcription factor expression in H3.3 KD1 ESCs (Figures 2E and 2F).

To understand the susceptibility of this small subset of genes to loss of H3.3, we examined the chromatin landscape of these genes both in the presence and upon depletion of H3.3. For all 84 transcripts upregulated upon H3.3 depletion, we observe a gain of H3K4me3 (p = 0.006) and a loss of H3K27me3 (p = 0.01) enrichment at the promoter (Figures 2G and 2H, Figures S2G and S2H). Recent studies have shown that global loss of H3K27me3 results in concomitant increase in acetylation of H3 on lysine 27 (H3K27ac), with loci-specific analyses suggesting that these modifications are mutually exclusive (Lau and Cheung, 2011; Pasini et al., 2010; Reynolds et al., 2012; Tie et al., 2009). We therefore profiled H3K27ac in our control and H3.3-depleted ESCs (Figure 2I and Figure S2I). Analysis of the promoters of genes upregulated upon H3.3 depletion revealed an increase in H3K27ac (p = 0.0001). Likewise, we observed a significant H3K27ac increase upon depletion of H3.3 at all bivalent promoters (p = 2.2×10−16). Interestingly, H3K27ac levels in promoters of genes upregulated upon H3.3 depletion as compared to all bivalent promoters were significantly elevated even in the control sample (p = 1.4×10−5), while H3K27me3 levels were decreased (p = 2.2×10−16). These results suggest that the balance of H3K27ac versus H3K27me3 is perturbed upon depletion of H3.3, possibly contributing to aberrant gene expression.

H3.3 Depletion Alters the Differentiation Potential of ESCs

Our RNA-seq results suggest that H3.3-depleted ESCs may be able to access the extraembryonic trophectoderm lineage. To assess the developmental potential of H3.3-depleted ESCs, we injected cells subcutaneously into immunodeficient mice and allowed them to give rise to teratomas, tumor-like formations containing tissues from all three embryonic germ layers. This assay can identify mutations in ESCs that allow contribution to trophectoderm (Koh et al., 2011; Lu et al., 2008).

Histological analysis showed that H3.3-depleted ESCs successfully differentiated into the three embryonic germ layers, with contribution to endoderm, ectoderm and mesoderm similar to teratomas from control ESCs (Figure 3A and Figure S3A). In addition, multiple hemorrhagic placental-like structures were observed in all teratomas derived from H3.3-depleted ESCs (Figure 3A, Figures S3A and S3B), whereas only one small area was observed in one teratoma derived from control ESCs (Control, n = 12; H3.3 KD1, n = 12; H3.3 KD2, n = 3). These foci contained many giant cells with large nuclei and multinucleated cells (indicated by arrows in Figure 3A, right panel), indicative of extra-embryonic trophoblast giant cells. Additionally, these foci stained positively for Troma-I, an antibody that detects keratin 8 expressed by trophectoderm (Figure S3B). As keratin 8 expression is not exclusive to trophectoderm (see epithelial endoderm staining present in H3.3 KD1 and H3.3 KD2 teratomas in Figure S3B which was also present in control teratomas), we isolated RNA from control and H3.3 KD1 teratomas (Figure 3B). RT-qPCR showed upregulation of Cdx2 and Hand1, two transcription factors essential for formation of trophoblast, in teratomas derived from H3.3-depleted ESCs. We also observed upregulation of Dlx3, a homeobox transcription factor whose deletion results in fetal death due to abnormal placental development (Morasso et al., 1999), and placental cadherin (Cdh3). Together, these histological and expression analyses demonstrate that H3.3-depleted ESCs can differentiate towards the typically restricted trophectoderm lineage.

Figure 3. H3.3-Depleted ESCs Show Altered Potential and Misregulation of Lineage-Specific Factors Upon Differentiation.

Figure 3

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A) Histology of teratomas from control (left panel) and H3.3 KD1 (right panel) ESCs. Sections were stained with Hematoxylin and Eosin (H&E). Trophoblast focus outlined and representative giant trophoblast cells indicated by arrows (right panel). B) RT-qPCR of placental markers in control and H3.3 KD1 teratomas. Data are represented as mean expression relative to Gapdh ± s.d. for an experiment performed in biological and technical triplicate. *p<0.05, **p<0.01, ***p<0.001. C-E) RT-qPCR of indicated lineage markers in control and H3.3 KD1 ESCs subject to differentiation. Data are represented as mean expression relative to Gapdh ± s.d. (n = 3) at time points indicated, normalized to control ESCs. F) Fold-enrichment over ESC of genes defined as ESC-bivalent found differentially regulated in H3.3-null MEFs. For all comparisons, p<1x10−6. See also Figure S3.

In H3.3-depleted ESCs, global reduction of H3K27me3 at promoters does not result in global upregulation of developmentally poised genes (see the modest changes in gene expression shown in Figures 2A and 2B). We therefore considered that H3.3-dependent reduction of H3K27me3 enrichment at developmentally regulated promoters (Figure 1E, Figures S3C and S3D) might result in gene misregulation upon differentiation rather than deregulation in the pluripotent state. To test this, control and H3.3-depleted ESCs were subject to embryoid body formation by cell aggregation and withdrawal of leukemic inhibitory factor (LIF) from the culture medium. As expected, both Cdx2 and Hand1 were upregulated in H3.3-depleted cells compared to control cells over the course of the differentiation experiment (Figure 3C). In addition, we observed upregulation of Gata4 and Gata6, primitive endoderm markers (Figure 3D), as well as Brachury and Fgf8, mesoderm markers (Figure 3E), at distinct time points after LIF withdrawal. We observed no delay in the downregulation of pluripotency factors (data not shown) and no upregulation of the ectoderm markers Nestin and Pax6 in H3.3-depleted cells (Figure S3E).

To assess the effect of H3.3 depletion on gene expression in a more differentiated cell type, we injected H3.3-null ESCs into blastocysts and isolated mouse embryonic fibroblasts (MEFs) at greater than 90% purity from the resulting chimeras (Figures S3F and S3G). RNA-seq showed 1,287 misregulated genes, with nearly even distribution of up- (635) and down-regulated (642) genes. Analysis of all genes differentially expressed (DE) in MEFs showed that 24% were bivalent in ESCs, a 2-fold enrichment over total (12% of all genes were defined as bivalent in ESC) (Figure 3F, p<2.2×10−16). When DE genes in MEFs were further binned into 2-fold up- or down-regulated, fold-enrichment over total was 1.5 and 2.7, respectively. A more stringent threshold of 5-fold up- or down-regulation resulted in fold-enrichment of 1.9 and 3.2, respectively. DE genes in MEFs had H3.3 levels in ESCs that were 20% higher than genes that showed no change in expression between control and H3.3-null cells (p<2.2×10−16). Overall, our data suggest that expression of at least a subset of bivalent genes showing reduced H3K27me3 enrichment is perturbed upon H3.3 depletion once factors supporting self-renewal are removed.

H3.3-dependent Chromatin Dynamics Contribute to PRC2 Binding and Activity at Developmentally Regulated Bivalent Promoters in ESCs

We next wanted to understand how loss of H3.3 deposition leads to changes in H3K27me3 levels at developmentally regulated promoters. We first tested whether PRC2 preferred H3.3 over canonical H3 as a substrate. However, an in vitro histone methyltransferase assay using the minimal recombinant PRC2 complex (Eed, Ezh2, Suz12, and RbAp48) showed similar activity towards both H3.2- and H3.3-containing recombinant mononucleosomes (Figures S4A-C). Loss of H3K27me3 at promoters could not be attributed to globally reduced levels of PRC2 in H3.3-depleted ESCs (Figures S4D-E). Additionally, we did not observe expression changes in UTX and Jmjd3, the H3K27me3 demethylases (Figure S4F), nor were Jmjd3 or UTX enrichment at target loci perturbed in the absence of H3.3 (Figures S4G-I).

We finally tested whether H3.3 depletion reduced PRC2 enrichment at the promoters of developmentally regulated genes in ESCs. Similar to what we observed with H3K27me3 enrichment, enrichment of PRC2 complex members Suz12 and Jarid2 at promoter-distal and intergenic H3K27me3 peaks showed minimal differences in the absence of H3.3 (Figure S4J). However, H3.3 knockdown reduced PRC2 enrichment at bivalent promoters (Figure 4A), suggesting that reduction of H3K27me3 is due at least in part to decreased PRC2 enrichment at its target promoters. H3.3 levels at developmental loci in Suz12−/− ESCs were not reduced, suggesting that H3.3 incorporation is upstream of PRC2 recruitment (Figure 4B).

Figure 4. H3.3 Contributes to PRC2 Binding at Bivalent Promoters.

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A) ChIP-qPCR of Suz12 (top) and Jarid2 (bottom) at representative housekeeping and bivalent genes in control and H3.3-depleted ESCs. Error bars represent s.d. (n = 3). Data confirmed by multiple biological replicates. B) H3.3 enrichment profile at the TSS of bivalent genes in wild type and Suz12−/− ESCs. Data are represented as reads per mapped million per 50 bp bin. C) RT-qPCR of trophectoderm-specific transcription factors in control ESCs and H3.3 KD1 ESCs exogenously expressing H3.2, H3.3, or H3.3 mutated at K27. Data are represented as mean expression relative to Gapdh ± s.d. (n = 3). Data from control and H3.2/H3.3 addback are also shown in Figure 2F. See also Figure S4.

Because H3.3 depletion reduced H3K27me3 and PRC2 levels at bivalent promoters, we next tested whether methylation of Lys27 on H3.3 was required to prevent upregulation of trophectoderm transcription factors in ESCs (see Figure 2F). We introduced point mutations of H3.3 at Lys27 (H3.3K27A and H3.3K27R) into H3.3 KD1 ESCs (Figure 4C). Surprisingly, we found that expression of either H3.3K27A or H3.3K27R in H3.3-depleted ESCs was sufficient to prevent upregulation of Cdx2 and Hand1, suggesting that methylation of H3.3 itself is not required to rescue expression phenotypes in ESCs. This unexpected result suggests that instead of acting as the primary substrate for PRC2, H3.3 facilitates the proper chromatin environment for PRC2 occupancy at its target regions and/or for PRC2 activity towards H3, regardless of isoform (i.e., H3.1/2, or H3.3).

Hira-dependent Deposition of H3.3 Facilitates Establishment of the Bivalent Chromatin Landscape at Developmentally Regulated Genes in ESCs

The histone chaperone Hira is associated with replication-independent deposition of H3.3 into chromatin (Tagami et al., 2004). Recent studies have refined our understanding of H3.3 incorporation into chromatin by demonstrating that Hira is responsible for most if not all H3.3 deposition in genic regions, including deposition within the promoters of bivalent genes (Figure S5A), while an alternate chaperone system, Atrx/Daxx, facilitates H3.3 deposition at many non-genic repeat regions (Filipescu et al., 2013).

To determine whether the chromatin landscape at developmentally regulated genes was dependent upon either Hira- or Atrx/Daxx-dependent H3.3 deposition, we performed ChIP-qPCR experiments in both Hira−/− and Daxx-depleted ESCs (Figures S5B and S5C). Loss of Hira had no effect on H3K4me3 levels at housekeeping genes Actin and Rps19 (Figure 5A), in agreement with our previous observation that Hira-dependent H3.3 deposition is not required for maintenance of gene expression in pluripotent cells (Goldberg et al., 2010). Similarly, no consistent pattern of change in H3K4me3 enrichment was obvious at developmentally regulated genes in Hira−/− ESCs (Figure 5A); while we observe an ~2-fold increase in H3K4me3 enrichment at the Tbx3 promoter, RT-qPCR shows Tbx3 upregulation in Hira−/− ESCs (data not shown). Daxx depletion had no effect on H3K4me3 enrichment at either housekeeping or developmentally regulated genes (Figure 5B), unsurprising given its role in H3.3 deposition at repeat elements. Finally, we performed ChIP-qPCR to analyze H3K27me3 enrichment at bivalent promoters in the absence of either Hira or Daxx. In support of our observations in H3.3-depleted ESCs, we observe reduced H3K27me3 enrichment at bivalent promoters in Hira−/− ESCs (Figure 5C), whereas Daxx knockdown did not affect H3K27me3 enrichment (Figure 5D). As observed in H3.3-depleted ESCs, Hira−/− ESCs also showed reduced enrichment of the PRC2 components Suz12 and Jarid2 at bivalent promoters (Figures 5E and 5F) not coupled to changes in protein expression level (Figure S5D); while perturbation of Suz12 at promoter-distal or intergenic H3K27me3 peaks was minimal, we did observe reduced enrichment of the low levels of Jarid2 at these regions (Figure S5E). Overall, these data are in agreement with Hira-dependent deposition of H3.3 at the promoters of developmentally regulated genes (Figures S5F-H).

Figure 5. Maintenance of H3K27me3 Levels at Bivalent Promoters Requires Hira-Dependent H3.3 Deposition.

Figure 5

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A-D) ChIP-qPCR of (A, B) H3K4me3 and (C, D) H3K27me3 at representative housekeeping and bivalent genes in (A, C) control and Hira−/− ESCs and (B, D) control and Daxx-depleted ESCs. ChIP-qPCR of (E) Suz12 and (F) Jarid2 at representative housekeeping and bivalent genes in control and Hira−/− ESCs. For panels A-F, error bars represent s.d. (n=3) with data confirmed by 3 biological replicates. G) Box plots represent CATCH-IT normalized to H3gen ChIP-seq at promoters in control and Hira−/− ESCs. The bottom and top of the boxes correspond to the 25th and 75th percentiles, and the internal band is the 50th percentile (median). The plot whiskers correspond to 1.5× interquartile range. P-values were calculated using Wilcoxon tests. H) RT-qPCR of trophectoderm markers in control and Hira−/− ESCs subject to differentiation. Data are represented as mean expression relative to Gapdh ± s.d. (n = 3) at the time points indicated, normalized to control ESCs. See also Figure S5.

In accordance with the loss of newly incorporated histone at promoters in the absence of H3.3 (Figure 1D) and the requirement of Hira for H3.3 deposition at promoters (Figure S5A), Hira−/− ESCs show reduced nucleosome turnover at both active (p < 2.2×10−16) and bivalent loci (p = 8.1×10−12) (Figure 5G). The lack of change in gene expression or H3K4me3 enrichment observed in the absence of H3.3 or Hira, conditions which reduce nucleosome turnover, supports that expression of active genes and maintenance of their associated chromatin states do not require incorporation of newly synthesized histones. In contrast, Hira-dependent H3.3 deposition associated with nucleosome turnover facilitates H3K27me3 enrichment at bivalent loci.

To test whether Hira deletion and H3.3 depletion have the same effect on gene expression, we subjected wild type and Hira−/− ESCs to in vitro differentiation as described above. In agreement (Figures 2D and 3C), the trophoblast-specific transcription factors Cdx2 and Hand1 are upregulated both in the pluripotent state as well as all time points of differentiation in the absence of Hira (Figure 5H).

Hira Co-localizes with Promoter-proximal RNA Polymerase II and the PRC2 Complex at the Promoters of Developmentally Regulated Genes in ESCs in an H3.3-dependent Manner

Because Hira−/− ESCs recapitulated the loss of H3K27me3 and PRC2 at bivalent promoters, we next sought to determine whether Hira localized to these regions. We generated an ESC line stably expressing Hira C-terminally tagged with the 3xFlag epitope (Figure S6A; Hira3xFlag) and performed ChIP-seq using a Flag-specific M2 antibody. Chromosome-wide analysis showed that Hira and H3.3 display similar enrichment profiles (Pearson correlation coefficient of 0.78, p<2.2×10−16), with enrichment observed at genic regions (Figure 6A). We next aligned our ChIP-seq data to the TSS of genes binned by expression level. In agreement with Hira’s role as a chaperone for genic H3.3 deposition and H3.3′s association with transcription, as well as a recent Hira ChIP-seq data set (Pchelintsev et al., 2013), enrichment of Hira and H3.3 were both associated with gene expression levels (Figure S6B).

Figure 6. Hira Co-localizes with Promoter-proximal RNAPII and PRC2 at Bivalent Promoters in an H3.3-dependent Manner.

Figure 6

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A) Genome browser representation of input, H3.3, and Hira profiles for chromosome 11. Read counts are normalized to total number of reads obtained for each data set. B) ChIP-seq density heat maps for RefSeq genes classified as H3K4me3 only (n = 15,670), bivalent (n = 2,938), or H3K27me3 only (n = 23). Plots show ±5 kb centered on the TSS. H3K4me3 only genes are rank-ordered by H3K4me3 enrichment, and bivalent and H3K27me3 only genes are rank-ordered by H3K27me3 enrichment, based on FPKM for each ChIP-seq data set. Data are represented as reads per mapped million per 50 bp bin. Color intensity represents tag counts, standardized across gene groupings for each ChIP-seq data set. C) Immunoblotting of Hira immunoprecipitated from wild type or H3.3-null nuclear extracts showing coimmunoprecipitation with Ubinuclein and H3.3 (* non-specific band) (5% input), and H3.3-dependent coimmunoprecipitation with RNAPII S5p, Ezh2, Suz12, and Jarid2 (0.5% input). D) Hira enrichment profile at the TSS of genes classified by chromatin modification state in wild type (solid lines) and H3.3 KO2 (dashed lines) ESCs. Data are represented as reads per mapped million per 50 bp bin. E) Model of H3.3 in modulation of PRC2 recruitment and activity in ESC. Recruitment of Hira and H3.3 to PRC2 targets are co-dependent and upstream of PRC2 recruitment. Either Hira or H3.3 directly, or the high frequency of free DNA afforded by their presence, allow for optimal PRC2 recruitment and activity at target loci. See also Figure S6.

We next assessed the enrichment of various chromatin-related factors at genes classified by their histone post-translational modification status (Figure 6B) using published data sets for PRC2 (Peng et al., 2009) and RNAPII (Brookes et al., 2012). Promoters marked by H3K4me3 only were also associated with Hira and H3.3, but not PRC2 (H3K27me3, Ezh2, Suz12, and Jarid2). H3K4me3 only promoters were enriched with RNAPII S5p consistent with promoter-proximal pausing of active genes, as well as RNAPII S2p (Figure 6B). As expected, promoters with both H3K4me3 and H3K27me3 were enriched with PRC2 and RNAPII S5p, albeit at lower levels than observed in genes marked by H3K4me3 only. Consistent with our initial observation that bivalent promoters are enriched with H3.3 (Figure 1C), we observed similar patterns, albeit lower levels, of Hira enrichment at this class of genes (Figure 6B and Figure S6C). Together, these data support the presence of Hira-dependent H3.3 deposition at both active genes and targets of PRC2 regulation in ESCs.

We next wanted to determine whether Hira physically associated with PRC2. Using nuclear extracts prepared from both wild type and Hira3xFlag ESCs, we found that Hira coimmunoprecipitated its known complex member Ubinuclein and its histone substrate H3.3 (Figure 6C and Figure S6D). In support of our observed colocalization of Hira and RNAPII and published studies demonstrating their association (Ray-Gallet et al., 2011), we found that Hira coimmunoprecipitated both promoter-proximal RNAPII S5p and elongating RNAPII S2p (Figure 6C and Figure S6D). In addition, Hira coimmunoprecipitated PRC2 complex members Ezh2, Suz12, and Jarid2 (Figure 6C and Figure S6D), but not Atrx (Figure 6C), demonstrating specificity of the interaction.

To determine whether Hira-PRC2 interaction required H3.3, we used nuclear extracts from H3.3-null ESCs to immunoprecipitate Hira-associated factors. While the Hira-Ubinuclein interaction was maintained in the absence of H3.3, association with both RNAPII and PRC2 was lost (Figure 6C). In agreement with loss of association with interacting factors, we found that Hira was not enriched at its target promoters in the absence of H3.3 (Figure 6D). Overall, our data suggest that while the nuclear Hira complex is maintained in the absence of H3.3, Hira and H3.3 localization to chromatin are co-dependent and upstream of PRC2 in their recruitment to bivalent promoters (Figure 6E).

DISCUSSION

A growing literature demonstrates that embryonic stem cells display an open chromatin state (Gaspar-Maia et al., 2011). One signature of the pluripotent state is the presence of bivalent domains within the promoters of developmentally regulated genes, marked by the presence of both H3K4me3 and H3K27me3 (Bernstein et al., 2006); most of these genes are expressed at marginally detectable to low levels in ESCs (mean FPKM in our data set of ~3). The bivalent chromatin landscape is thought to contribute to the plasticity of ESCs, facilitating their ability to rapidly initiate changes in gene expression programs upon receipt of differentiation stimuli. Our data demonstrate the importance of the histone variant H3.3 at bivalent promoters and suggest that chromatin dynamics may play a key role in maintaining this chromatin landscape.

Given its close association with active transcription, it is surprising that an absolute requirement for H3.3 in the maintenance or activation of gene expression profiles has not been identified (Cui et al., 2006). In our study, ESCs lacking H3.3 are functionally pluripotent as they maintain the ability to self-renew and contribute to all three germ layers. Others have found that Drosophila embryos are able to develop to adulthood in the absence of H3.3, challenging the importance of this variant for active transcription (Hodl and Basler, 2009; Sakai et al., 2009). Additionally, our work has shown that the absence of H3.3 deposition does not affect the chromatin landscape of actively transcribed regions (e.g., H3K4me3, H3K36me3). We therefore conclude that although H3.3 is found co-localized with these activation-associated modifications, the enzymes responsible for their installation show no preference towards either canonical H3.1/2 or the variant H3.3 as a substrate (Loyola et al., 2006) and no dependence upon H3.3 for their recruitment or activity.

Despite the association of H3K27me3 with gene repression, reduction of this histone post-translational modification at bivalent developmentally regulated promoters upon H3.3 depletion does not correlate with global upregulation of gene expression under conditions promoting self-renewal. H3.3 depletion does lead to upregulation of a small number of developmentally regulated genes, in particular a number essential for establishment of the extraembryonic trophectoderm lineage. Placental-like foci identified in teratomas derived from H3.3 KD ESCs constitute a small percentage (<5%) of the resulting tumors, leading us to conclude that only a subset of H3.3-depleted ESCs express trophectoderm markers at sufficient levels to drive this lineage formation, and that H3.3-depleted ESCs do not, in general, represent a more totipotent cell state.

The expression data presented herein are somewhat in contrast to observations of ESCs lacking PRC2 components (Surface et al., 2010). While able to maintain the pluripotency transcription program, these cells exhibit low-level de-repression of lineage-specific markers when cultured under conditions supporting self-renewal. When subject to differentiation in vitro, they fail to fully activate lineage-specific genes and maintain aberrant expression of pluripotency markers. The discrepancy between PRC2 null versus H3.3-depleted ESCs could result from incomplete abrogation of PRC2 activity on chromatin upon H3.3 depletion; compared to H3.3-depleted ESCs, PRC2 null cells contain drastically reduced global and local levels of H3K27me3. Alternatively, these discrepancies may arise from a function of H3.3 unrelated to Hira-dependent deposition into chromatin, or perhaps a function of PRC2 that is not abrogated by the loss of H3.3 and is responsible for the PRC2 null phenotype.

In our current study, ontology analysis of the bivalent promoters most affected by H3.3 depletion predicts that H3.3 may play a critical role in neural tube mesenchyme. Indeed, zebrafish expressing a mutant of H3.3 preventing chromatin incorporation exhibit craniofacial skeletal defects (Cox et al., 2012). Several additional studies suggest that H3.3 may play an essential role in development. Hypomorphic h3f3a gene trap mice display partial neonatal lethality, stunted growth, neuromuscular deficits, and male sub-fertility (Couldrey et al., 1999). Deletion of Hira in mice (Roberts et al., 2002) and depletion of H3.3 or Hira in Xenopus laevis (Szenker et al., 2012) result in pleiotropic gastrulation defects and embryonic lethality, due in part to an inability to maintain nucleosome occupancy in the absence of H3.3 (Ray-Gallet et al., 2011). We do not detect decreased nucleosome occupancy in H3.3-depleted ESCs. However, ESCs spend ~70% of their cell cycle in S phase with access to abundant levels of canonical H3; perhaps maintenance of nucleosome occupancy in the absence of H3.3 would play a greater functional role upon exit from pluripotency and especially upon terminal differentiation.

While we did not detect decreased nucleosome occupancy, we did observe altered chromatin dynamics in H3.3-depleted ESCs. As rescue of gene misregulation due to depletion of H3.3 did not require Lys27, we hypothesize that PRC2 can utilize either H3.3 or canonical H3 as a substrate, but that H3.3 might provide a dynamic chromatin environment beneficial to PRC2 localization and/or activity at the promoters of developmentally regulated genes. Our CATCH-IT data reveal lower levels of nucleosome turnover upon depletion of H3.3 at both active and bivalent genes; it remains unclear why the chromatin landscape and transcriptional output of active loci remain unaffected, while the chromatin landscape established by PRC2 is compromised.

Interestingly, Jarid2 is able to bind free DNA with a bias towards GC-rich regions (Li et al., 2010), facilitating PRC2 recruitment to target loci in ESCs (Margueron and Reinberg, 2011). Promoters of developmentally regulated genes tend to be CpG-rich regions, typically depleted of nucleosomes and enriched for H3.3. We hypothesize that the dynamic environment associated with H3.3 incorporation results in these regions existing in nucleosome-free states at higher frequency than other regions, facilitating Jarid2’s role in recruitment of PRC2 to CpG-rich bivalent promoters. Intriguingly, Jarid2 is highly expressed in ESCs under control of the pluripotency transcription machinery and is downregulated upon differentiation. In addition, our analysis in MEFs demonstrates enrichment of H3.3 at both H3K4me3 and bivalent promoters, but not at the thousand promoters modified by H3K27me3 only (data not shown). We therefore propose that the requirement for H3.3 and Jarid2 to fine-tune PRC2 activity at bivalent promoters might be unique to the pluripotent state, with alternate PRC2 recruitment mechanisms taking precedence in differentiated cells. This hypothesis suggests evolutionary or developmental divergence in PRC2 recruitment mechanisms from Drosophila, where H3.3 is enriched at PREs in adult somatic cells.

We report a previously unappreciated role for H3.3, a histone variant associated with gene activation, in establishment of the bivalent chromatin landscape at lowly expressed developmental genes in ESCs. Our data supports the emerging notion that H3.3 is not required for active transcription, but instead may be necessary for establishing a chromatin landscape allowing for proper gene expression upon differentiation. Intriguingly, point mutations of H3 and H3.3, specifically at Lys27, have recently been discovered in pediatric glioblastomas (Schwartzentruber et al., 2012; Wu et al., 2012). In light of the work presented herein, this finding is particularly exciting given emerging links between development and oncogenesis, with certain types of cancers thought to be due to either regression to a less differentiated state or an initial failure to establish differentiation. Future studies will address these and other roles of this unique H3 variant in normal development and abnormal development leading to pathological conditions.

EXPERIMENTAL PROCEDURES

ESC culture

ESCs (C57Bl/6J background) were cultured under standard conditions. Mouse h3f3b ZFNs and targeting were previously described (Goldberg et al., 2010), and H3.3A shRNA constructs were purchased from OpenBiosystems (TRCN0000012023, TRCN0000012027). See Supplementary Experimental Procedures for more details.

Chromatin immunoprecipitation (ChIP) and antibodies

Native and crosslinking ChIP with approximately 2×107 ESCs per experiment were preformed as previously described (Goldberg et al., 2010). All antibodies (see Supplementary Experimental Procedures) were previously reported as suitable for ChIP and immunoblotting.

ChIP-seq

ChIP-seq libraries were prepared according to the Illumina protocol and sequenced with either the Genome Analyzer II or HiSeq 2000. Reads were mapped to the mouse genome (build 37, mm9) using the Bowtie alignment software (Langmead et al., 2009). Unique reads mapped to a single best-matching location with no more than two mismatches were kept for peak identification and profile generation. MACS (Zhang et al., 2008) was used for peak calling in each replicated sample and the resulting peaks were filtered by peak height to reduce false positives. We computed the total number of reads in individual promoters for comparing histone modifications in different groups of promoters. Read depths were used to normalize peak heights and promoter modification levels across samples.

Wiggle files, generated by extending each read 100 bp from its 3′ end, were used to generate average ChIP-seq profiles using the Cistrome analysis pipeline (http://cistrome.dfci.harvard.edu/ap/). Numbers of reads used for analysis: H3.3 (22 million), H3K4me3 (25 million), H3K27me3 (25 million), H3 general (100 million), H3K27ac (20 million), Hira3xFlag (13.5 million), Hira (23 million), UTX (10 million), Jmjd3 (10 million).

RNA-seq

RNA was isolated using Qiagen RNeasy. Libraries were prepared according to the Illumina TruSeq protocol and were sequenced on the HiSeq 2000. Resulting reads (101 nt) were aligned to the mouse genome (mm9) using TopHat (Trapnell et al., 2009). Gene expression level measured as Fragments Per Kilobase of exons per Million mapped fragments (FPKM) was determined by the maximum likelihood estimation method implemented in the Cufflinks software package with annotated transcripts as references. Differential expression was analyzed using the Student’s t-test in the program Cuffdiff (Trapnell et al., 2010) with p-values corrected for multiple testing.

Immunoprecipitation

5 μg antibody bound to 25 μL Dynabeads was incubated with 1 mg of ESC nuclear extract (see Supplementary Experimental Procedures) for 3 hrs at 4 °C. Beads were washed four times with 1 mL buffer (20 mM HEPES pH 7.9, 300 mM KCl, 0.01% NP-40, 5 mM 2-mercaptoethanol, 0.4 mM PMSF) and eluted in 1x SDS loading buffer.

Supplementary Material

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02
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Research Highlights.

  • Developmental promoters in embryonic stem cells are dynamic loci marked by H3.3

  • H3.3 promotes H3K27me3 levels in embryonic stem cells

  • H3.3 facilitates PRC2 recruitment to developmental promoters

  • The H3.3 chaperone Hira and PRC2 interact in an H3.3-dependent manner

Acknowledgements

This work was supported by The Rockefeller University Fund (to C.D.A), Tri-Institutional Stem Cell Initiative (to C.D.A. and S.R.), US National Institute of Mental Health (MH087840 and MH073164 to D.Z.), Damon Runyon Cancer Research Foundation (L.A.B.), and The Rockefeller University Women & Science Initiative (L.A.B.). We thank I. Falciatori for help with the teratoma assay, Y. Zhang for purified recombinant PRC2 complex, and S. Chen and Y. Shi for the Jmjd3 antibody. The Troma-I hybridoma developed by P. Brulet and R. Kemler was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biology, Iowa City, IA 52242. We thank L. Baker, E. Korb, E. Duncan, and C. Lu for critical reading of this manuscript, and J. Wysocka and members of the Allis lab for helpful discussions.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Accession Numbers. Our ChIP-seq and RNA-seq data sets have been deposited in the GEO data base with accession numbers GSE42155.

Supplemental Information. Supplemental information includes Supplemental Experimental Procedures, six figures, and two tables and can be found with this article online.

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Associated Data

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

Supplementary Materials

01
NIHMS525479-supplement-01.pdf (34.4MB, pdf)
02
NIHMS525479-supplement-02.xls (1.7MB, xls)
03
NIHMS525479-supplement-03.xls (42.5KB, xls)

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