Dynamics of RNA Polymerase II Pausing and Bivalent Histone H3 ...

Article

Dynamics of RNA Polymerase II Pausing and Bivalent Histone H3 Methylation during Neuronal Differentiation in Brain Development Graphical Abstract

Authors Jiancheng Liu, Xiwei Wu, Heying Zhang, Gerd P. Pfeifer, Qiang Lu

Correspondence [email protected]

In Brief Pol II pausing and chromatin bivalency are two mechanisms controlling gene transcription. Liu et al. reveal that Pol II pausing is associated with cell-typespecific functions in mouse neural progenitor cells and their daughter neurons, while change in H3K4me3/ H3K27me3 bivalency is associated with gene activation during mammalian neuronal differentiation.

Highlights d

Global patterns of Pol II and bivalent marks were characterized in NPCs and neurons

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Pol II pausing is associated with cell-type-specific functions in NPCs and neurons

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Bivalent marks prime neuronal specification genes for activation during differentiation

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Paused Pol II and H3K27me3 often co-exist in the promoterproximal regions

Liu et al., 2017, Cell Reports 20, 1307–1318 August 8, 2017 ª 2017 The Author(s). http://dx.doi.org/10.1016/j.celrep.2017.07.046

Accession Numbers GSE93011

Cell Reports

Article Dynamics of RNA Polymerase II Pausing and Bivalent Histone H3 Methylation during Neuronal Differentiation in Brain Development Jiancheng Liu,1,5 Xiwei Wu,2,5 Heying Zhang,1 Gerd P. Pfeifer,3,4 and Qiang Lu1,6,* 1Department

of Developmental and Stem Cell Biology of Molecular and Cellular Biology 3Department of Cancer Biology Beckman Research Institute of City of Hope, 1500 East Duarte Road, Duarte, CA 91010, USA 4Present address: Center for Epigenetics, Van Andel Institute, 333 Bostwick Ave. NE, Grand Rapids, MI 49503, USA 5These authors contributed equally 6Lead Contact *Correspondence: [email protected] http://dx.doi.org/10.1016/j.celrep.2017.07.046 2Department

SUMMARY

During cellular differentiation, genes important for differentiation are expected to be silent in stem/progenitor cells yet can be readily activated. RNA polymerase II (Pol II) pausing and bivalent chromatin marks are two paradigms suited for establishing such a poised state of gene expression; however, their specific contributions in development are not well understood. Here we characterized Pol II pausing and H3K4me3/H3K27me3 marks in neural progenitor cells (NPCs) and their daughter neurons purified from the developing mouse cortex. We show that genes paused in NPCs or neurons are characteristic of respective cellular functions important for each cell type, although pausing and pause release were not correlated with gene activation. Bivalent chromatin marks poised the marked genes in NPCs for activation in neurons. Interestingly, we observed a positive correlation between H3K27me3 and paused Pol II. This study thus reveals cell type-specific Pol II pausing and gene activationassociated bivalency during mammalian neuronal differentiation. INTRODUCTION Cellular differentiation is accompanied by global changes in gene expression, from patterns of maintaining stem/progenitor cells to patterns for supporting differentiation, controlled by key networks of transcription regulators and epigenetic mechanisms (Lagha et al., 2012; Lomvardas and Maniatis, 2016; Martynoga et al., 2012; Reik, 2007; Suzuki and Bird, 2008; Tee and Reinberg, 2014; Zhou et al., 2011). How such global changes of gene expression are orchestrated in development, however, is not well understood. The core enzyme in transcription, RNA polymerase II (Pol II), is at the center of multiple regulations. Recruitment of Pol II and formation of the pre-initiation complex

involving gene-specific transcription factors are known to be critical steps for initiating successful transcription. In addition to this key regulation, it was recently recognized that postrecruitment regulation has a crucial impact on whether or not Pol II can achieve a productive elongation of transcription (Adelman and Lis, 2012; Gaertner and Zeitlinger, 2014; Levine, 2011; Scheidegger and Nechaev, 2016). In this mechanism, Pol II could initiate transcription of a short product but then pause at about 30–50 nt downstream of the transcription start site (TSS). The balance between negative and positive regulatory factors determines whether Pol II is paused or released from pause into elongation (Zhou et al., 2012). Pausing of Pol II was originally discovered in the study of transcriptional regulation of heat shock genes in Drosophila (Gilmour and Lis, 1986; Rougvie and Lis, 1988), and it was later observed in a large number of genes (Core et al., 2008; Guenther et al., 2007; Muse et al., 2007; Nechaev et al., 2010; Rahl et al., 2010; Zeitlinger et al., 2007) in many different species, suggesting potential functions of pausing in developmental programs. Available data have documented the involvement of Pol II pausing to facilitate the response to changes in environmental conditions (such as the heat shock response), to coordinate synchronous expression of genes across cells of a tissue during development, and/or to maintain a basal level of gene expression in the cell (Adelman and Lis, 2012; Gaertner and Zeitlinger, 2014; Levine, 2011; Scheidegger and Nechaev, 2016). However, it remains to be further investigated to what extent pausing and pause release are involved in gene expression dynamics or contribute to specific developmental programs during cellular specifications. Bivalent histone H3 modification is another key feature implicated in the regulation of gene expression (Bernstein et al., 2006; Zhou et al., 2011). The histone H3 lysine 4 trimethylation (H3K4me3) is in general linked to genes with active expression, whereas the histone H3 lysine 27 trimethylation (H3K27me3) is associated with genes in a repressed state. Interestingly, genes containing overlapping H3K4me3 and H3K27me3 marks only express at a low level, which leads to the thought that such bivalent genes may be poised for activation or repression (or maintained in the bivalent silent state) during the progression of embryogenesis. In support of this idea, the bivalent H3K4me3

Cell Reports 20, 1307–1318, August 8, 2017 ª 2017 The Author(s). 1307 This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

and H3K27me3 chromatin domains are found to be preferentially enriched at developmental regulatory genes in mouse embryonic stem cells (ESCs) (Bernstein et al., 2006), germline cells (Sachs et al., 2013), pre-implantation embryos (Liu et al., 2016b), and several adult tissues (Cui et al., 2009, 2012; Hammoud et al., 2009; Kinkley et al., 2016; Mikkelsen et al., 2007). Therefore, bivalent histone H3 modification also presents a suitable regulatory mechanism for modulating gene expression levels during development. However, available data documenting the in vivo status of H3K4me3 and H3K27me3 bivalent chromatin during development is limited due to the technical difficulties associated with scarce amounts of embryonic tissues. It remains to be further examined whether the histone bivalency and a change of bivalency state are linked to major changes of gene expression during developmental progressions, such as during cell fate specifications. In the developing mouse cerebral cortex, neural progenitor cells (NPCs) first expand the progenitor cell pool by consecutive rounds of proliferation. Then NPCs initiate a transition from proliferation to differentiation and produce descendant neurons that populate different layers of a functional cortex (Go¨tz and Huttner, 2005; Custo Greig et al., 2013; Kriegstein and Alvarez-Buylla, 2009; Lodato and Arlotta, 2015; Rakic, 2009; Taverna et al., 2014). During neurogenesis, NPCs divide asymmetrically to generate daughter neurons directly or indirectly. At the same time, they undergo self-renewal to keep the neural progenitor pool. Young daughter neurons migrate outward to the superficial part of the cortex, while NPCs keep their location in the inner part of the cortex. This pattern of progression during neuronal differentiation provides an excellent system for characterizing the regulation of gene expression and epigenetic modifications during developmental transitions. We previously designed a dual reporter strategy by labeling NPCs with EGFP and differentiated neurons with monomer red fluorescent protein (mRFP) in the same transgenic animals to help alleviate the problem of carryover of reporter (EGFP in this case) from NPCs to progeny neurons, thus allowing effective simultaneous purification of NPCs and daughter neurons from the embryonic cortices (Wang et al., 2011). Using this system, we have performed comparative studies of transcriptomes (Liu et al., 2016a; Wang et al., 2011) and several key epigenetic marks (Hahn et al., 2013) between the purified NPCs and neurons. In this study, we sought to characterize global patterns of gene expression regulation during neuronal differentiation by studying genomic DNA-binding sites of Pol II as well as the status of bivalent chromatin domains using chromatin immunoprecipitation sequencing (ChIP-seq). Our data uncovered distinct functions of Pol II promoter-proximal pausing and bivalent promoters during neuronal differentiation. While Pol II pausing in NPCs and neurons displayed features associated with cell-type-specific functions, the change of bivalent promoter status between NPCs and neurons correlated with neuronal gene activation. RESULTS Profiles of Pol II Chromatin Binding in NPCs and Neurons We accumulated purified NPCs and their daughter neurons from the embryonic day 15.5 (E15.5) mouse cortices using a

1308 Cell Reports 20, 1307–1318, August 8, 2017

previously established dual transgenic reporter strategy (Wang et al., 2011). Because the amount of purified NPCs that could be obtained from the embryonic mouse brains was limited, we adopted a method of amplification for ChIPseq using purified cells for each experiment (see the Experimental Procedures). To characterize the genomic binding sites of Pol II, we used antibodies that recognize Pol II in different states: antibodies that recognize (1) the N terminus of the largest subunit of Pol II (N-20), (2) the hypophosphorylated state of the Pol II, (3) the phosphorylated C-terminal domain of Pol II at Ser5, and (4) the phosphorylated C-terminal domain of Pol II at Ser2. The first antibody could detect all forms of Pol II (Pol II-total) regardless of the phosphorylation status, whereas the second antibody could specifically bind to the hypophosphorylated form of Pol II (Pol II-nonP). The latter two antibodies could specifically recognize an early elongation form of Pol II (Pol II-Ser5P) that is often seen bound at promoter-proximal regions (Rahl et al., 2010) and an elongating Pol II (Pol II-Ser2P), respectively. ChIP by each of the abovementioned antibodies was done using 2.5 3 105 sorted cells (NPCs or neurons) per antibody, and amplified libraries were sequenced by next-generation sequencing (Table S1). Figure 1 and Figure S1 summarize the heatmaps and composite metagene analyses of ChIPseq data obtained from these antibodies comparing purified NPCs and neurons. In both NPCs and neurons, Pol II-total predominantly occupied promoter regions and peaked around the transcription start site (TSS) (Figures 1A and 1B). The binding signals extended from the TSS to the transcription end site (TES), reflecting Pol II in active elongation mode (Figure 1B). Genomic binding by Pol II-Ser5P (Figures 1A and 1B) or Pol II-Ser2P and Pol II-nonP (Figure S1) similarly showed higher signal intensity around the TSS and extended intensity along the gene body. In contrast to Pol II-total, the two antibodies to phosphorylated forms of Pol II appeared to show a noticeable difference in composite metagene profiles between NPCs and neurons (Figure 1; Figure S1). Particularly, binding by Pol II-Ser5P displayed a significant difference in signal intensity between NPCs and neurons around the TSS, suggesting more genes in NPCs having the early elongation form of Pol II accumulated to a promoter-proximal site than those in neurons. An independent ChIP-seq of purified NPCs and neurons with the Pol II-total and Pol II-Ser5P antibodies obtained similar results (Figure S2). In addition, there was an apparent shift between the Pol II-binding sites detected by the two antibodies (Figure 1C), where Pol II-Ser5P appeared to be located further downstream of Pol II-total from the TSS. Pol II-total signal peaked around nucleotide 30–40 downstream of the TSS, while Pol II-Ser5P signals centered around nucleotides 60–70. The separation of Pol II-total- and Pol II-Ser5P-binding peaks at the promoters suggested that the majority of Pol II engaged with promoters in NPCs and neurons might reflect a state of docking, as also observed in other systems (Maxwell et al., 2014). The binding of Pol II-Ser5P, on the other hand, might indicate a paused Pol II after the initiation of transcription, as suggested by previous studies (Nechaev et al., 2010; Rahl et al., 2010).

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Figure 2. ChIP-Seq of H3K4me3 and H3K27me3 in NPCs and Neurons

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Figure 1. ChIP-Seq of Pol II in NPCs and Neurons (A) Heatmaps of ChIP-seq data of two different forms of Pol II in NPCs and neurons. Rows were sorted by decreasing Pol II-total occupancy in the promoter regions (–2 kb to +2 kb of transcription start site) in NPCs. Scaled intensities are in units of log2 fold change between antibody-enriched sample and IgG control. (B) Metagene analyses of Pol II-total and Pol II-Ser5P on gene body region (upper panels) and on promoter region (lower panels) in NPCs (green lines) and neurons (red lines). TSS, transcription start site; TES, transcription end site. The y axis represents the log2 fold change between antibody-enriched sample and IgG control. (C) Metagene profiles of Pol II-total (yellow line) and Pol II-Ser5P (blue line) in NPCs. Arrows showed peak positions of profiles, indicating Pol II-Ser5P signals shifted a little more downstream of TSS compared with Pol II-total in NPCs.

Profiles of H3K4me3 and H3K27me3 Marks in NPCs and Neurons We next analyzed the H3K4me3 and H3K27me3 marks in purified NPCs and neurons using 2.5 3 105 sorted cells per antibody. Duplicate ChIP samples yielded consistent sequencing data (Figure S3A). The heatmaps (Figure 2A) and metagene analyses (Figure 2B) showed that, similar to Pol II, both chromatin marks also primarily peaked just downstream of the TSS region in NPCs and neurons. The promoter signal intensities of the two

(A) Heatmaps of ChIP-seq data of H3K4me3 and H3K27me3 in NPCs and neurons. Rows were sorted independently by H3K4me3 or H3K27me3 occupancy in the promoter regions (–2 kb to +2 kb of TSS) in each cell type. Scaled intensities are in units of log2 fold change between antibody-enriched sample and IgG control. (B) Metagene analyses of H3K4me3 and H3K27me3 on gene body region (upper panels) and on promoter region (lower panels) in NPCs (green lines) and neurons (red lines). TSS, transcription start site; TES, transcription end site. The y axis represents the log2 fold change between antibody-enriched samples and IgG control.

marks were markedly different between NPCs and neurons (Figure 2B), suggesting developmental changes of these marks associated with the neuronal differentiation process. Pol II Occupancy and Chromatin Marks in Relation to Gene Expression Level in NPCs and Neurons In both NPCs and neurons, the distribution of genes as a function of expression levels revealed a similar pattern (Figure 3A), which consisted of a bell-shaped curve of higher expressers (active genes) and a shoulder curve of lower expressers (inactive genes). We asked whether and how gene expression levels in NPCs and neurons were influenced by promoter Pol II binding or by the H3K4me3 and H3K27me3 marks. With respect to Pol II, our data showed that the promoter Pol II-total signal was positively correlated with gene expression levels in both NPCs and neurons (Figure 3B), whereas Pol II-Ser5P signal showed an apparent biphasic curve of correlation (Figure 3B), suggesting that promoter Pol II-total occupancy is more closely associated

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Figure 3. Promoter Pol II Occupancy and H3K4me3 and H3K27me3 Marks in Relation to Gene Expression Level (A) Global gene expression distribution patterns in NPCs (green) and neurons (red). (B) Promoter Pol II occupancy with reference to gene expression. Pol II occupancy was divided into six equal-sized groups from low signal to high signal.


with the overall strength of expression of the bound genes. With respect to the two chromatin marks, the promoter H3K4me3 signal was positively associated with gene expression levels in both NPCs and neurons (Figure 3C), consistent with previous studies of ESCs and other cell types. Interestingly, the promoter H3K27me3 signal displayed a biphasic relationship with gene expression level (Figure 3D), with lower H3K27me3 levels positively correlated with gene expression but higher H3K27me3 levels inversely correlated with gene expression. Further examination of the status of H3K4me3 in genes marked by H3K27me3 revealed that the biphasic curve of H3K27me3 was mainly attributed to the presence and contribution of H3K4me3 in these genes (Figure S3B). Pol II Pausing in NPCs and Neurons We next sought to characterize the group of paused genes in NPCs and neurons. For this purpose, we applied promoter occupancy by Pol II-Ser5P as an indicator of polymerase pausing and pause release (Nechaev et al., 2010; Rahl et al., 2010), using an operational definition in which a paused gene was defined by the following three features: (1) significant promoter Pol II-Ser5P signal (intensity R4), (2) pausing index (PI) or traveling ratio (TR) calculated by relative ratio of density of Pol II-Ser5P in the promoter-proximal region (30 to +300 bp) and the gene body (defined as from +300 to the TES) R1, and (3) transcript expression level defined by RNA sequencing (RNA-seq) R2. Using this operational definition, we identified 7,401 genes in NPCs and 2,212 genes in neurons showing these features of pausing. Within the groups of genes paused in NPCs and neurons, genes appeared to be expressed at a wide range of levels (Tables S2, S3, and S4), indicating that pausing did not predict a suppressed state (low expression) of gene expression. This was in agreement with previous observations that many paused genes were actively expressed in resting cells (Gilchrist et al., 2012). As 1,712 paused genes were shared by both NPCs and neurons, we further divided the paused genes into the following three groups (Figure 4A): genes specifically paused in NPCs (5,689 genes; Table S2), genes specifically paused in neurons (500 genes; Table S3), and genes paused in both NPCs and neurons (1,712 genes; Table S4). Database for Annotation, Visualization and Integrated Discovery (DAVID) bioinformatics analyses (DAVID 6.7) showed that the top biological processes in NPCspecific paused genes highlighted different aspects of catabolic processes and cell cycle (Figures 4B and 4E), functions that appear to be required for supporting the proliferative state of NPCs. Those of neuron-specific paused genes revealed enrichment of activities involving DNA damage response and repair and cellular stress responses (Figures 4C and 4F), functions important for maintaining the integrity of post-mitotic neuronal cells (Narciso et al., 2016; Pan et al., 2014). Interestingly, with respect to DNA damage response and repair, different groups

The normalized gene expression data were plotted for each promoter signal group. (C) Promoter H3K4me3 and H3K27me3 with reference to gene expression. H3K4me3 or H3K27me3 ChIP-seq signals were divided into six equal-sized groups from low signal to high signal. The normalized gene expression data were plotted for each promoter signal group.

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(A) Paused genes identified in NPCs and neurons. Pausing index was calculated by the average promoter Pol II-Ser5P signal (read counts in 30 bp to +300 bp of TSS) and average gene body Pol II-Ser5P signal (read counts in +300 bp of TSS to TES). 5,689 genes were paused in NPCs, but not in neurons, and 500 genes were paused in neurons, but not in NPCs, while 1,712 genes were paused in both NPCs and neurons. (B) Top biological processes of NPC-specific paused genes displayed features of proliferating cells, including different aspects of catabolic processes and cell cycle. The top 3,000 genes ranked by pausing index were analyzed by DAVID. (C) Top biological processes of neuron-specific paused genes showed enrichment of activities important for maintaining the integrity of postmitotic neuronal cells, involving DNA damage response and repair and cellular stress responses. (D) Top biological processes of paused genes shared by NPCs and neurons included common basic cellular functions, such as molecule transport, protein translation, and RNA processing. (E) Usp38 and Ppp1cb represent the group of NPC-specific paused genes. The numbers in square brackets represent track heights. Scale bars represent 5 kb for both genes. (F) Gtf2h5 and Esco2 represent the group of neuron-specific paused genes. The numbers in square brackets represent track heights. Scale bars represent 1 kb for Gtf2h5 and 2 kb for Esco2. (G) Hnrnpa2b1 and Ddx5 represent the group of paused genes shared by NPC and neuron. The numbers in square brackets represent track heights. Scale bars represent 1 kb for both genes.

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were closely associated with cell-typespecific functions. The top biological processes in paused genes shared by both NPCs and neurons included common basic cellular functions, such as molecule transport, protein translation, and RNA processing (Figures 4D and 4G).

H3K4me3 and H3K27me3 Marks in NPCs and Neurons Gtf2h5 Esco2 We next characterized the genes marked by the H3K4me3 and H3K27me3 modifiG [0-30] [0-40] NPC Pol II-total cations in NPCs and neurons, using the Neuron [0-30] [0-40] [0-20] [0-20] NPC promoter signals of the two chromatin Pol II-Ser5P [0-20] [0-20] Neuron modifications for selection of marked Ddx5 Hnrnpa2b1 genes. Figure 5A summarizes the dynamic changes of the chromatin modification state of four subgroups of of paused genes were identified between NPCs and neurons promoters from NPCs to neurons, including promoters con(Table S5). The NPC-specific paused group, but not the taining bivalent marks, prominent H3K4me3 alone, prominent neuron-specific group, was highly enriched with homologous H3K27me3 alone, or neither of the two marks. To assess the porecombination genes, which are particularly important for the tential involvement of the H3K4me3 and H3K27me3 marks in S/G2 phases of the cell cycle. This suggested that paused genes neuronal differentiation, we looked more closely at changes of


Figure 5. Dynamics of H3K4me3 and H3K27me3 in Neuronal Differentiation (A) Differentially marked genes in NPCs and neurons. Four different groups of promoters were categorized as follows: bivalent, promoters with both H3K4me3 and H3K27me3 peaks; K4me3 only, promoters with H3K4me3 but no H3K27me3 peaks; K27me3 only, promoters with H3K27me3 but no H3K4me3 peaks; and none, promoters without H3K4me3 and H3K27me3 peaks. (B) H3K4me3-marked genes in NPCs and neurons (upper panel) and bivalent genes in NPCs and neurons (lower panel). (C) Top biological processes of NPC-specific H3K4me3-marked genes (left panel) and bivalent genes (right panel). The numbers in the brackets represent gene number in each process. (D) Notch2 and Sox2 represent the group of H3K4me3-marked genes in NPCs, which contains many players crucial for maintaining the progenitor cell state and are downregulated in neurons. The numbers in square brackets represent track heights. Scale bar, 1 kb. (E) Tbr1 and Neurod2 represent the group of bivalent genes in NPCs, which includes many genes important for neuronal differentiation and lineage specification and are upregulated in neurons. The numbers in square brackets represent track heights. Scale bar, 1 kb. (F) Several oligodendrocyte progenitor cell-specific genes display bivalent marks in NPCs but show the H3K27me3 mark only in neurons. These genes are downregulated in neurons. The numbers in square brackets represent track heights. Scale bar, 1 kb.


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DNA metabolic process and cell cycle, reflecting prominent features of proliferating cells. This group of genes contained most NPC-specific genes whose protein products are known to be involved in supporting the progenitor cell state of NPCs, for instance, the Notch family (Notch 1, 2, and 3 and Hes1, 5, and 6), Sonic Hedgehog family (Gli2 and 3), Wnt family (Wnt7a and Tcf3 and 4), Hippo pathway genes (Tead1 and 2), TAM receptor tyrosine kinases (Axl and Tyro-3), Brca1, Ephrin-B1, Pax6, and Sox2 (Figure 5D; Table S6; Figure S4A). On the other hand, the top biological processes of NPC-specific bivalent genes (Figure 5C; Table S7) displayed embryonic organ development, embryonic morphogenesis, and neuron differentiation, reflecting functions related to differentiation and tissue patterning. These NPC-specific bivalent genes included many genes known to be critical for proper differentiation and lineage specification, migration, and/or function of cortical neurons, such as Tbr1, Fezf2, SatB2, NeuroD2, Trnp1, Crmp1, Nav2, Prdm8, Reln, and Rtn1 (Figure 5E; Table S7; Figure S4B). Most of these neuronal specification- and function-related bivalent genes showed a transition from a state of bivalency in NPCs to a state of more prominent H3K4me3 promoter abundance in neurons (Figure 5E; Figure S4B), which also well correlated with the upregulation of their gene expression levels (Table S7). Interestingly, multiple oligodendrocyte progenitor cell-specific genes, including Olig1, Olig2, and Pdgfra, contained bivalent H3K4me3 and H3K27me3 marks in NPCs (Figure 5F; Figure S4C). These genes were switched from bivalent to more abundant H3K27me3 mark in neurons, correlating with the downregulation of their expression.

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C on NP Neur Figure 6. Pausing and Pause Release or Dynamics of Bivalent Marks in Relation to Gene Activation from NPCs to Neurons (A) The changes of pause release ratio (PRR) of Pol II-Ser5P between NPCs and neurons (PRR_Neuron/NPC) were divided into 6 groups from low to high (x axis). Gene expression fold changes between NPCs and neurons were calculated and compared across groups. (B) Two defined gene groups: activated (555 genes) and non-activated (2,845 genes). (C) PRR (Neuron/NPC) changes between activated and non-activated gene groups. p value 3.1 3 105 (Welch two-sample t test). (D) Bivalent to H3K4me3 switch from NPCs to neurons accompanied with gene activation. p value 0.004 (Welch two-sample t test). (E) Activated gene groups enriched with bivalent to H3K4me3 switch genes during neuronal differentiation. p value p < 2.2e16 (Fisher’s exact test).

the H3K4me3 alone and bivalent groups of promoters (Figure 5B). The top biological processes of NPC-specific H3K4me3-marked genes (Figure 5C; Table S6) highlighted

Dynamic Changes of Pausing Status or Chromatin Marks in Relation to Gene Activation during Neuronal Differentiation The original discovery of polymerase pausing in Drosophila revealed that paused genes were likely highly inducible genes, such as heat shock genes, suggesting that pausing might prepare genes for quick induction in response to environmental or developmental stimuli, such as during cell fate specification. Similarly, bivalency of chromatin marks was also thought to reflect a poised state of genes for inducible expression during developmental progressions. We therefore asked whether polymerase pausing and pause release or changes of bivalency state are linked to gene activation during differentiation of NPCs into neurons during brain development. To assess whether pausing and pause release from NPCs to neurons might set the genes for activation during differentiation, we first looked at pause release in relation to gene expression change. Pause release ratio (PRR) (Chen et al., 2015), an inverse of PI or TR, was calculated by relative ratio of density of Pol II-Ser5P in the gene body and the promoter-proximal region. As shown in Figure 6A, with the increase of the PRR_Neuron/ NPC ratio, log2FC became more dynamic, but it did not change significantly from NPCs to neurons, suggesting that pause release is not indicative of gene activation. To further examine this, we tested if activated genes during neuronal differentiation were characterized by increased PRR. We defined the group of activated genes (gene expression in NPCs or neurons R2 and Log2FC-Neuron/NPC R 0.8) and the group of non-activated Cell Reports 20, 1307–1318, August 8, 2017 1313

Figure 7. Potential Interplay between H3K27me3 and Pol II Pausing (A) Pol II-total and Pol II-Ser5P promoter occupancy in different histone mark groups. (B) Association of H3K27me3, but not H3K4me3, with Pol II-Ser5P. Pol II-Ser5P signal at promoter region was divided into five groups from high to low both in NPCs and neurons (upper panels); H3K4me3- or H3K27me3-marked promoters were divided accordingly (middle and lower panels). (C) Reciprocal sequential ChIP analysis (Pol II-Ser5P - > K27me3 indicated first IP with anti-Pol II-Ser5P followed by second IP with anti-H3K27me3) showed co-presence of Pol II-Ser5P and H3K27me3 on the indicated promoters, which were among the top 20% genes showing strong positive correlation between H3K27me3 and Pol II-Ser5P in purified neurons (B). Assays were performed on FACS-purified E15.5 neurons. The sequential ChIP data are given as fold over IgG control in the second IP. Error bars represent SDs.

genes (gene expression in NPCs or neurons R2 and jLog2FCNeuron/NPCj % 0.1) (Figure 6B). Between these two gene groups, the ratio of PRR (neurons/NPCs) was lower in the activated genes than in the non-activated genes (Figure 6C), further suggesting that pause release is not inherently linked to gene activation. To assess whether changes of bivalency from NPCs to neurons might be associated with gene activation during differentiation, we analyzed the group of NPC-specific bivalent genes, particularly the subgroup of genes displaying a change from bivalency in NPCs to prominent H3K4me3 alone in neurons. This subgroup included many genes critical for proper neuronal differentiation and lineage specification (Figure 5E). We found that these genes showed significantly higher expression levels in neurons than in NPCs (Figure 6D), suggesting that a switch from the state of bivalency in NPCs to H3K4me3 in neurons was well correlated with gene activation during neuronal differentiation. Furthermore, we found that, within the group of activated genes (Figure 6B), there was a significantly higher ratio of genes (138/555) that showed a change from bivalency in NPCs to H3K4me3 in neurons, comparing to a much lower ratio of such genes (345/10,872) within the group of active genes in NPCs or neurons (gene expression level R2 in NPCs or neurons). Together, these data indicated that the bivalency to H3K4me3 switch is positively associated with gene induction during neuronal differentiation. Potential Interplay between H3K27me3 and Pol II Pausing The predominant domains of H3K4me3 and H3K27me3 around the TSS in both NPCs and neurons (Figure 2) raised an interesting question as to whether these histone modifications may function in connection with Pol II, which occupies a neighboring or somewhat overlapping domain (Figure 1). To address this issue, we compared promoter signals between Pol II and the chromatin marks in genes specifically enriched with H3K4me3, H3K27me3, or both marks (bivalent). Our data showed that Pol II-total was more abundant in H3K4me3 promoters in both NPCs and neurons, whereas the promoter signal of H3K27me3 displayed a significantly higher correlation with that of Pol II-Ser5P (Figure 7A), suggesting that H3K27me3 was positively correlated with paused Pol II. Furthermore, when sorted based on promoter Pol II-Ser5P signal, higher levels of H3K27me3 signal were associated with higher levels of Pol II-Ser5P around


the TSS region, whereas H3K4me3 did not appear to show such a correlation (Figure 7B). To address whether the observed correlation represented co-presence of H3K27me3 and Pol II-Ser5P in single cells rather than due to cellular heterogeneity, we further performed sequential ChIP of the two followed by qPCR analysis of several target genes of high correlation. Our results revealed that reciprocal sequential ChIP could detect co-enrichment of H3K27me3 and Pol II-Ser5P on these target genes (Figure 7C; Table S8), consistent with the idea of H3K27me3 and Pol II-Ser5P co-existing in the same cells. Together, these analyses showed that paused Pol II and H3K27me3 often co-exist in the promoter-proximal regions. DISCUSSION Developmental progression of stem/progenitor cell differentiation is accompanied by global changes of gene expression patterns in a precise temporal and/or spatial order, in which genes required for maintaining stem/progenitor cell state are tuned down while genes crucial for supporting the specification and function of differentiated cells are tuned up. Such an exquisite transition of global gene expression patterns requires a concerted action of various regulatory mechanisms working at different levels, including DNA methylation, chromatin modification, specific DNA-binding factors, etc. To enable this transition to proceed smoothly, it is conceivable that the cell-type-specific genes might be somehow placed in a standby state in stem/progenitor cells and can readily change transcriptional output upon being instructed to differentiate. Pol II pausing and bivalent chromatin modification are two mechanisms that are particularly suitable for such a poised state of gene induction; however, whether and how they are involved in controlling gene expression dynamics during cell fate specification are not clear. Combining purified cortical NPCs and their daughter neurons with ChIP-seq for a limited number of cells, we determined the global profile of genomic binding by Pol II and the H3K4me3 and H3K27me3 marks. Our data revealed some unique features of Pol II pausing and bivalent chromatin marks in the regulation of NPC differentiation during brain development. Paused genes have been characterized by both the global run-on sequencing (GRO-seq) method, which detects the functional Pol II engaged in RNA synthesis, and the ChIP-seq method, which detects promoter occupancy by Pol II. While GRO-seq would more accurately reveal the pausing status of Pol II, previous studies also indicated that the magnitude of GRO-seq signal correlated well with the density of Pol II in the promoter-proximal region based on ChIP-seq data (Core et al., 2012), suggesting that the abundance of Pol II, particularly the form of Pol II-Ser5P (Rahl et al., 2010), at the promoters could be one index to reflect pausing. Due to the limited number of cortical NPCs derived from embryonic mouse brains (Wang et al., 2011), we chose to use ChIP-seq of Pol II to examine paused genes in this study, as GRO-seq would have needed us to use significantly more (>40-fold more) purified cortical NPCs. Our ChIP-seq data revealed two interesting features: first, a global change of paused genes accompanied neuronal differentiation; and second, the change was highly correlated with cell-type-specific functional requirements for

NPCs or neurons. Thus pausing appeared to be a mechanism to globally orchestrate expression of thousands of genes in order for the cell to function properly for its cell state. The paused genes showed a wide range of expression levels. This perhaps reflected the distinct requirement of respective functions of individual genes at the given moment of the cell state. For instance, we would envision that NPCs in a faster or slower growing state may impose higher or lower expression levels of genes of nucleotide metabolic pathways, respectively. In this context, although pausing is not required for rapid gene induction as originally thought, it seems to be a crucial mechanism for maintaining or sustaining concerted expression of genes essential for the characteristics of a cell type and allowing fine-tuning of gene levels to suit the need of the state of the cell. Our ChIP-seq data on H3K4me3 and H3K27me3 revealed that dynamic changes of these two chromatin marks correlated well with cell fate determination during development. In particular, the H3K4me3 mark alone and the bivalent H3K4me3/ H3K27me3 marks showed close association with distinct molecular programs of cell fate specification in NPCs. Most genes known to be crucial for the maintenance of NPCs displayed abundant H3K4me3 promoter signals with little H3K27me3, suggesting a crucial function of H3K4me3 in promoting active expression of these genes in NPCs. After neuronal differentiation, these genes mostly lost the promoter H3K4me3 mark, which correlated with their silencing in neurons. In contrast, many neuronal lineage specification- and neuron functionrelated genes were characterized by co-existence of the promoter H3K4me3 and H3K27me3 signals in NPCs. Most of these genes showed a change of chromatin marks from the bivalency in NPCs to a more prominent H3K4me3 mark status in neurons during neurogenesis. This also correlated with a lower expression level of many of these genes in NPCs but an upregulated level in neurons. Thus, the H3K4me3 and H3K27me3 bivalent marks appeared to function to prepare the marked genes in NPCs to be induced for an active expression in neurons. Upon differentiation, the switch of bivalency to predominant H3K4me3 mark would maintain these genes in an active state to support the progression of neuronal differentiation. Whether Pol II pausing and the bivalent chromatin marks work independently or if they may coordinate with each other in transcriptional regulation is an intriguing question, given the observations that the paused Pol II and chromatin marks occupy neighboring or overlapping domains in the promoter-proximal region. Interestingly, our data revealed that H3K27me3 was positively correlated with Pol II-Ser5P in both NPCs and neurons. In this regard, it is intriguing to note that Drosophila PRC (Polycomb-repressive complex) could physically interact with paused Pol II (Tie et al., 2016) and was preferentially localized to paused promoters (Enderle et al., 2011). In addition, between the two classes of bivalent domains in ESCs (Ku et al., 2008), it was found that promoters bound by PRC2 alone could allow Pol II pausing (Min et al., 2011), although PRC1 and PRC2 co-occupied promoters were devoid of paused Pol II. Together, these observations suggest that H3K27me3 may not simply be a permissive mark but likely to either play an active role in Pol II pausing or coordinate with paused Pol II in controlling the


transcriptional output during development. Future studies are required to further investigate this potential interplay between paused Pol II and the H3K27me3 mark. In summary, our data indicate that Pol II promoter pausing and the bivalent H3K4me3 and H3K27me3 marks are two crucial but distinct regulatory mechanisms of transcriptional control for proper progression of neuronal differentiation during brain development. The bivalent marks are important for cell fate specification transitioning from NPC to neuron, while Pol II pausing is important for supporting the established state of the two cell types. EXPERIMENTAL PROCEDURES Animals Nestin-EGFP and Dcx-mRFP reporter mice were previously described (Hahn et al., 2013; Wang et al., 2011). Animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) and were conducted in accordance with NIH guideline and the Guide for the Care and Use of Laboratory Animals. All experiments were conducted with stage-matched embryos (E15.5), which contained both males and females. Purification of NPCs and Neurons Purification of E15.5 cortical cells using a dual reporter strategy was done as previously described (Hahn et al., 2013; Wang et al., 2011). RNA-Seq Purified E15.5 NPCs and neurons were accumulated to 0.5–1 million, and total RNAs were isolated using Trizol reagent (Invitrogen). Single-end libraries were prepared, size selected, gel purified, and sequenced using Illumina HiSeq2000 system. ChIP-Seq ChIP experiments were modified from previous protocols (Hahn et al., 2013) using the following antibodies: Pol II total (sc-899, Santa Cruz Biotechnology); Pol II nonP (ab817, Abcam); Pol II Ser2P (04-1571, EMD Millipore); Pol II Ser5P (ab5131, Abcam); H3K4me3 (39159, Active Motif); and H3K27me3 (07-449, EMD Millipore). Specifically, cells collected from fluorescence-activated cell sorting (FACS) were cross-linked by adding 37% formaldehyde to a final concentration of 1%. Cells were incubated on a rotator for 10 min at room temperature (RT), and formaldehyde was quenched by adding 1.25 M glycine to a final concentration of 125 mM. Cells were rocked for 5 min at RT, washed with cold PBS, and flash-frozen in liquid nitrogen. Cells were stored at 80 C and accumulated to a desired number. Cells were lysed in buffer containing 1% SDS, 5 mM EDTA, and 50 mM Tris-HCl (pH 8.0) with freshly added protease inhibitor (04693116001, Roche). The chromatin was fragmented to 200–500 bp with a Misonix S3000 Sonicator at 4 C. After centrifugation, around 100 mL supernatant from 2.5 3 105 cell lysate was diluted to 1 mL with a dilution buffer containing 1% Triton X-100, 2 mM EDTA, 150 mM NaCl, and 20 mM Tris-HCl (pH 8.0) with freshly added protease inhibitor. Antibody (1 mg) was added to chromatin and the mixture was incubated at 4 C overnight. Then 10 mL Dynabeads Protein A (10001D, Invitrogen) was added to the chromatin/antibody mixture and incubated for an additional 2 hr at 4 C. Beads were washed sequentially with 700 mL each of the following: washing buffer I (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 150 mM NaCl, and 20 mM TrisHCl [pH 8.0]), washing buffer II (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 500 mM NaCl, and 20 mM Tris-HCl [pH 8.0]), washing buffer III (0.25 M LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA, and 10 mM Tris-HCl [pH 8.0]), and TE buffer (10 mM Tris-HCl [pH 8.0] and 1 mM EDTA). The sample was eluted with 250 mL elution buffer containing 0.5% SDS, 25 mM Tris-HCl (pH 8.0), and 10 mM EDTA. The eluted sample was treated with 0.2 M NaCl and 1 mg/mL Protease K (P-2308, Sigma) at 65 C overnight. DNA was purified with phenol/chloroform and was precipitated with cold ethanol and glycogen. The dissolved DNA samples after ChIP were subjected to amplification with MicroPlex Library Preparation kit (AB-004-0012, Diagenode). Amplification was performed with a StepOnePlus Real-Time PCR System (Applied


Biosystem) and was stopped manually at the exponential phase. Library after amplification was purified twice with Agencourt AMPure XP beads (A63880, Beckman Coulter) and sequenced on a Hiseq 2500 sequencer. RNA-Seq Data Analysis Reads were aligned to the mouse reference genome mm9 using TopHat v2.0 with default settings. The expression levels of RefSeq genes were counted by matching the aligned reads to the coordinates of the RefSeq exons. Reads falling into exons that belong to multiple transcripts of the same genes were counted one time, and the reads from all exons of the same gene were combined to represent the gene level expression. Raw counts were then normalized by trimmed mean of M value (TMM) method implemented in the Bioconductor package edgeR. The normalized counts were then scaled by the gene length and log2 transformed to represent the expression value of each RefSeq gene model. ChIP-Seq Data Analysis Reads from ChIP-seq experiments were aligned to mouse genome assembly mm9 using Novoalign v3.02.07. Only reads aligned to a unique genome locus were reported. Peak calling was done by MACS v2.0 using the corresponding IgG control with the option of p = 0.001 and d = 200. All subsequent analysis was done using customized R scripts. Gene annotations of mm9 genome were downloaded from the RefSeq database. For metagene profile and heatmap at gene body or promoter regions, these regions were divided into equal number of bins for each gene, and fold enrichment of read counts within each bin between ChIP sample and IgG control sample was calculated. These data matrices were used to generate metagene profiles by plotting the average signal in each bin for all the genes in NPC and neuron. They were also used to generate heatmaps using Java TreeView. Promoters were defined as TSS ± 500 bp, and proximal promoters were defined as 30 bp upstream and 300 bp downstream of TSS. The H3K4me3 and H3K27me3 promoter signal was calculated by the fold enrichment of read counts at the promoter region between the ChIP sample versus the IgG control sample. Pausing index was calculated by the fold difference of read counts within the proximal promoter and gene body (TSS + 300 bp to transcription end). Sequential ChIP Real-Time PCR Around 16 million of FACS-purified neurons at E15.5 were collected, fixed, and fragmented as described in ChIP-seq. The sequential ChIP experiment was performed using a Re-ChIP-IT kit (Active Motif). Anti-Pol II-Ser5P (ab5131, Abcam) or anti-H3K27me3 (07-449, EMD Millipore) was used in the first immunoprecipitation, respectively. Then anti-H3K27me3 or anti-Pol II-Ser5P was added to precipitated chromatin in the second reaction. For control, a ChIP-grade IgG (Abcam) was used replacing anti-Pol II-Ser5P or antiH3K27me3 in the second reactions. DNAs eluted from the second immunoprecipitations were applied to real-time qPCR analysis using a DyNAmo flash SYBR green qPCR kit (Thermo Fisher) on a StepOnePlus real-time PCR system (Applied Biosystems). Statistical Analysis Statistical analyses were performed with R v3.0.2. When two groups were compared, we first performed a normality test. Parametric data were compared using a Welch t test. Enrichment analysis of gene lists was performed using Fisher’s exact test. Statistical significance was set at p < 0.05. The data are presented as mean ± SEM. ACCESSION NUMBERS The accession number for the ChIP-seq data of Pol II, H3K4me3, and H3K27me3 as well as RNA-seq data comparing NPCs versus neurons reported in this paper is GEO: GSE93011. SUPPLEMENTAL INFORMATION Supplemental Information includes four figures and eight tables and can be found with this article online at http://dx.doi.org/10.1016/j.celrep.2017.07.046.

AUTHOR CONTRIBUTIONS J.C.L. and Q.L. conceived of the study. J.C.L. and H.Y.Z. conducted the experiments. X.W.W. conducted bioinformatics analyses. J.C.L., X.W.W., G.P.P., and Q.L. analyzed the data. J.C.L., X.W.W., G.P.P., and Q.L. wrote the paper. All authors approved the final manuscript. ACKNOWLEDGMENTS We thank Donna Isbell and Cirila Arteaga for assistance with animal breeding and care, Lucy Brown and Jeremy Stark and their staff for helping with cell sorting, Jinhui Wang for performing next-generation sequencing, and Jeremy Stark for helpful discussion of DNA damage response and repair-related genes. This work was supported by NIH grants NS075393 from NINDS to Q.L. and MH094599 from NIMH to Q.L. and G.P.P. In addition, research reported in this study included work performed in the Analytical Cytometry Core and Integrated Genomics Core supported by the National Cancer Institute under award number P30CA033572.

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Received: March 8, 2017 Revised: June 17, 2017 Accepted: July 18, 2017 Published: August 8, 2017

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