High-Resolution Mapping of Chromatin Packaging
in Mouse Embryonic Stem Cells and Sperm
Benjamin R. Carone,1Jui-Hung Hung,2,3Sarah J. Hainer,4,5Min-Te Chou,2Dawn M. Carone,6Zhiping Weng,3
Thomas G. Fazzio,4,5and Oliver J. Rando1,*
1Department of Biochemistry and Molecular Pharmacology, University of Massachusetts Medical School, Worcester, MA 01605, USA 2Institute of Bioinformatics and System Biology, National Chiao Tung University, 75 Bo-Ai Street, Hsin-Chu, Taiwan 300
3Program in Bioinformatics and Integrative Biology, University of Massachusetts Medical School, Worcester, MA 01605, USA 4Program in Gene Function and Expression, University of Massachusetts Medical School, Worcester, MA 01605, USA 5Program in Molecular Medicine, University of Massachusetts Medical School, Worcester, MA 01605, USA
6Department of Developmental and Cell Biology, University of Massachusetts Medical School, Worcester, MA 01605, USA *Correspondence:[email protected]
http://dx.doi.org/10.1016/j.devcel.2014.05.024
SUMMARY
Mammalian embryonic stem cells (ESCs) and sperm
exhibit unusual chromatin packaging that plays
important roles in cellular function. Here, we extend
a recently developed technique, based on deep
paired-end sequencing of lightly digested chromatin,
to assess footprints of nucleosomes and other
DNA-binding proteins genome-wide in murine ESCs and
sperm. In ESCs, we recover well-characterized
fea-tures of chromatin such as promoter nucleosome
depletion and further identify widespread footprints
of sequence-specific DNA-binding proteins such as
CTCF, which we validate in knockdown studies. We
document global differences in nuclease
accessi-bility between ESCs and sperm, finding that the
majority of histone retention in sperm preferentially
occurs in large gene-poor genomic regions, with
only a small subset of nucleosomes being retained
over promoters of developmental regulators. Finally,
we describe evidence that CTCF remains associated
with the genome in mature sperm, where it could play
a role in organizing the sperm genome.
INTRODUCTION
Eukaryotic genomes are packaged into a nucleoprotein complex known as chromatin, whose repeating subunit consists of147 base pairs (bp) of DNA wrapped around an octamer of histone proteins. Nucleosomes affect essentially all DNA-templated pro-cesses; as a result, there has been a great deal of interest over the past decade in characterizing nucleosome positions across the genomes of a variety of organisms (Jiang and Pugh, 2009; Radman-Livaja and Rando, 2010). In mammals, genome-wide maps have been reported for nucleosome positions in a number of cell types and tissues, including several immune cell types (Schones et al., 2008; Valouev et al., 2011); liver (Li et al., 2012); and embryonic stem cells (ESCs), neural precursor cells, and embryonic fibroblasts (Li et al., 2012; Teif et al., 2012). These genome-wide maps reveal characteristic features that are
conserved throughout eukaryotes, such as nucleosome deple-tion at promoters and other regulatory elements.
Several cell types exhibit unusual chromatin states that appear to be linked to biological function. ESCs, which are pluripotent, are characterized by ‘‘hyperdynamic’’ chromatin in which his-tone proteins exchange rapidly on and off of the genome (Meshorer et al., 2006). This has been proposed to contribute to a generally permissive chromatin state in which genes impor-tant for differentiation are accessible for rapid transcriptional activation. In contrast, mammalian sperm exhibit a highly un-usual chromatin state that is vastly different from that of other cell types (Ooi and Henikoff, 2007); most of the histone proteins are lost during spermatogenesis, first replaced by transition pro-teins, and eventually replaced by small basic proteins termed protamines. However, not all histones are lost (murine sperm retain 2% of their histones), and recent studies on histone retention in human and mouse sperm suggest that there is a bias for promoters of genes expressed early during development to be specifically packaged in histones (Arpanahi et al., 2009; Brykczynska et al., 2010; Erkek et al., 2013; Gardiner-Garden et al., 1998; Hammoud et al., 2009). These findings contrast with several lines of evidence suggesting that histone retention in sperm primarily occurs over repeat elements; small-scale cloning of DNA released by nuclease digestion of sperm revealed primarily repeat elements such as long and short inter-spersed nuclear element (LINE and SINE, respectively) se-quences (Pittoggi et al., 1999) and pericentric repeats (Govin et al., 2007), while immunostaining studies on mature sperm reveal colocalization of histone proteins with the repeat-enriched sperm chromocenter (Govin et al., 2007; van der Heijden et al., 2006). The discrepancies between these views of the sperm chromatin landscape remain unresolved.
In general, genome-wide nucleosome mapping relies on the characterization of the products of micrococcal nuclease (MNase) digestion of chromatin; MNase preferentially cleaves the linker DNA between nucleosomes, leaving nucleosomal DNA relatively intact as 147 bp footprints. Genome-wide characterization of MNase digestion products has proceeded rather rapidly from early studies using1 kb resolution micro-arrays to higher resolution tiling micromicro-arrays, to the modern era of deep sequencing (Radman-Livaja and Rando, 2010). Most recently, the Kent and Henikoff groups reported a significant advance in chromatin mapping (Henikoff et al., 2011; Kent
et al., 2011) by carrying out paired-end deep sequencing of an entire MNase digestion ladder (as opposed to using size-selected mononucleosomal DNA from such a ladder). These maps reveal not only nucleosome footprints of120–150 bp (fragments shorter than 147 bp result from MNase ‘‘nibbling’’ on the ends of nucleosomal DNA) but also shorter (<80 bp) foot-prints of other DNA-binding proteins such as transcription fac-tors (TFs).
Here, we use this method to analyze the chromatin structure of murine ESCs and sperm. We find that nucleosome positioning in ESCs is consistent with that of many other cell types, with broadly conserved features such as promoter nucleosome depletion that scales with transcription rate. More interesting, we confirm that this protocol yields footprints of a wide variety of sequence-specific DNA-binding proteins, including pluripo-tency factors such as Oct4, as well as the genomic insulator CTCF. We confirm that short MNase footprints over CTCF bind-ing sites correspond to bona fide CTCF bindbind-ing events by showing that these footprints are lost on CTCF knockdown in ESCs. Furthermore, we show that sperm nuclease accessibility is broadly different than that of ESCs. In contrast to prior genome-wide studies, we demonstrate that the majority of nu-cleosomes in sperm are retained over gene-poor regions, and we independently confirm this result microscopically in mature sperm. Nonetheless, using a specific subset of MNase digestion conditions, we are also able to recapitulate previous reports of
histone retention at CpG-rich promoters, although we show that this signal can only account for a minority of the total levels of histone retention in sperm. Last, we unexpectedly identify widespread CTCF footprints in sperm, suggesting that this factor may play a role in organizing the sperm genome for fertilization or for embryonic genome activation.
RESULTS
We set out to thoroughly characterize the chromatin structure in murine ESCs and sperm, using a protocol recently developed by the Henikoff group (Henikoff et al., 2011). As is common in nucleosome mapping studies, cells were crosslinked with form-aldehyde, lysed, and subjected to MNase digestion. Purified DNA after MNase digestion from ESCs is visualized inFigure 1A, revealing the expected nucleosome ladder. Following digestion, protected genomic DNA was isolated, and the entire nucleo-somal ladder was subject to paired-end deep sequencing. In this protocol, there is no gel purification of mononucleosome-sized fragments, although several steps during sequencing library preparation select against DNA fragments longer than 300 bp. After mapping sequencing reads back to the mouse genome, we calculated insert sizes for each fragment. Both libraries exhibited strong mononucleosome-sized protected fragments; 144 bp for ESCs and 157 bp for sperm (Figure 1B). For both libraries, subnucleosomal size peaks are also present,
Figure 1. Average Promoter Architecture in ESCs
(A) Gel electrophoresis of MNase digestion ladders for murine ESCs subject to EGFP KD. Characteristic nucleosomal laddering is evident in underdigested ES lanes. ESC digestion patterns are shown here for unfractionated (unspun) sam-ples; identical results are obtained for the super-natant of low-speed centrifugation (‘‘spun,’’ data not shown). Asterisk indicates the titration step used for paired-end deep sequencing library construction.
(B) Fragment size distribution for sequenced MNase digestion products. After deep sequencing of the indicated libraries, paired sequences were mapped back to the mouse genome, and insert size was calculated from genomic distance be-tween reads. ES ‘‘spun’’ indicates that MNase digestion was fractionated by centrifugation, and the library was constructed from supernatant material, whereas the ‘‘unspun’’ library was con-structed from the entire MNase digestion. PE, paired end.
(C) Normalized deep sequencing data plotted relative to TSSs of all mouse mm9 refGene tran-scripts for 1 to 80 bp MNase-protected fragments, 135 to 165 bp fragments, DNaseI hypersensitivity (GSE40869), and RNAPII ChIP-seq (indicated as Pol2), as indicated. The y axis represents the average value, for all genes, in parts per million reads.
(D) Comparison between cell types, and compar-ison to liver data from (Li et al., 2012). MNase footprints (135–165 bp) were selected for the indicated libraries, and occupancy was averaged for all TSS-aligned genes, shown as log2of the enrichment relative
to the genome-wide average read depth. Two key features are apparent here. First, sperm cells exhibit a strong NDR on average. Second, the unspun ESC library exhibits a relatively positioned +1 nucleosome on average, whereas the supernatant ‘‘spun’’ library exhibits the 30-extended promoter NDR observed in several prior studies (Teif et al., 2012; Li et al., 2012), indicating that 50nucleosomes are relatively insoluble under typical MNase digestion conditions.
Developmental Cell
with one predominant peak of60 bp in sperm (Montellier et al., 2013) and several smaller peaks in ESCs corresponding to 124, 104, 94, and 68 bp. The successive 20 bp reductions in peak size for ESCs likely represent the products of endonuclease cleavage at sharp bends in the nucleosomal DNA path where DNA interac-tions with the histone octamer are diminished (Crick and Klug, 1975; Noll and Kornberg, 1977).
We first focused on nucleosome-length (135–165 bp) MNase-protected fragments, expecting to recapitulate prior studies on ESC chromatin structure (Li et al., 2012; Teif et al., 2012). Basic characteristics of nucleosome positioning observed here are consistent with prior observations in a multitude of species, including mammalian cell types (Radman-Livaja and Rando, 2010). Promoters typically exhibit a nucleosome-depleted region (NDR) that coincides with genomic hypersensitivity to DNase I, and both of these features are centered just upstream of both the annotated transcriptional start site (TSS) and the peak of RNA polymerase II (RNAPII) chromatin immunoprecipitation (ChIP) signal (Yildirim et al., 2011) (Figure 1C; discussed later). In a prior study on ESC nucleosomes, Teif et al. reported an un-usual extension of promoter nucleosome depletion downstream of the TSS (Teif et al., 2012). We do not find such a 30NDR exten-sion when deep sequencing unfractionated MNase-digested chromatin (Figure 1C), but this unusual NDR behavior was reca-pitulated in another data set in which we centrifuged the MNase digestion and isolated relatively soluble chromatin prior to gener-ating our deep sequencing library (Figures 1B and 1D). These re-sults suggest that nucleosomes located at the extreme 50end of coding regions, such as the +1 nucleosome, may have reduced solubility due to their interactions with transcriptional machinery or other large protein complexes, which is consistent with prior studies on variability in nucleosomal solubility following nuclease digestion in flies (Henikoff et al., 2009). Our results using unfrac-tionated MNase digests thus show that chromatin organization in ESCs is consistent with chromatin organization in other cell types and species, at least at the level of nucleosome posi-tioning/occupancy.
Chromatin Organization at the TSS
In addition to nucleosome-sized DNA fragments (135–165 bp), we were also able to recover a wide range of digestion fragments, including subnucleosomal-sized DNA fragments. In yeast, such MNase footprints (<80 bp) are particularly abundant over the binding locations of sequence-specific DNA-binding proteins such as TFs (Henikoff et al., 2011; Kent et al., 2011). To extend these observations to a mammalian system, we aver-aged data for 1 to 80 bp fragments for all TSS-aligned genes. As shown inFigure 1C, these footprints were strongly enriched at the 50 end of the NDR, consistent with these footprints corre-sponding to TFs or the basal transcription machinery.
We further explored the connection between MNase footprints and promoter architecture using ‘‘V-plots’’ introduced by Henik-off et al. (2011). Briefly, in these plots, sequencing reads are aligned by a specific genomic position (anchor); in this case, the annotated TSSs of all genes in the mouse genome. The x axis represents the distance from the anchor along the genome, while the y axis shows the sequencing fragment size. Enrichment or depletion of footprints is shown in red or green, respectively, in a heatmap.Figure 2A shows V-plots for TSS-aligned data in
ESCs for three sets of genes grouped according to expression level. Examining nucleosome-sized DNA fragments (size range, 135–165 bp) reveals the expected behavior: nucleosomes are dramatically depleted from the promoters of highly expressed genes, with modest nucleosome depletion occurring at moder-ately expressed genes and minimal nucleosome depletion at silent genes (Figure 2A). At highly expressed genes, we observe a well-positioned +1 nucleosome with a slightly enlarged (175 bp) footprint, potentially representing a nucleosome engaged either with RNAPII or with some component of the basal transcription machinery. In contrast, short DNA fragments (1–80 bp) reveal the converse behavior, with footprints being observed most strongly at promoters with the highest expres-sion in ESCs. This is consistent with the notion that these foot-prints result from TFs or the basal transcription machinery, as previously suggested in flies (Teves and Henikoff, 2011). Finally, we note that intermediate-sized footprints (100–130 bp) are depleted surrounding the TSS in all three groups of genes. These footprints presumably result from MNase digestion of partially unwrapped or less stably bound nucleosomes and show little correspondence with transcription rate.
Next, we compared these TSS plots in ESCs to sperm ( Fig-ure 2B). Most dramatically, in contrast to the strong TF signal at ES promoters, sperm promoters were strongly depleted of these subnucleosomal footprints. This lack of promoter TF signal is consistent with the global, or nearly global, absence of active transcription in mature sperm (Kierszenbaum and Tres, 1975; Ward and Zalensky, 1996). Curiously, while no, or very little, tran-scription occurs in mouse sperm (expression values for sperm TSS plots were generated based on gene expression during the round spermatid stage), significant depletion of nucleo-some-sized fragments remains evident at promoters (Figure 1D andFigure 2B). We speculate that this lack of apparent nucleo-some occupancy at promoters could potentially be a remnant of transcriptional activity in round spermatids rather than the result of active repositioning or selective removal of histones during chromatin compaction in spermatogenesis (discussed later). In addition to the lack of a strong signature of the basal tran-scription machinery (discussed earlier), and the depletion of nucleosome footprints observed at promoters, we also noted a broadening of mononucleosomal footprint width in the sperm V-plots (Figure 2B; see alsoFigure 1B). The cause of this broad-ening is currently unclear.
The conclusions drawn from the averaged data were further confirmed on a gene-by-gene basis for ESCs.Figure 2C shows how the three size class fragments described earlier correlate with gene expression level: for each size class, all genes are ordered by ESC messenger RNA (mRNA) abundance from high (top) to low (bottom). Together, our ESC data validate our exper-imental approach and extend our understanding of ESC chro-matin to include a broad atlas of DNA-binding proteins.
Sperm Histone Retention
Having demonstrated the utility of this protocol in ESCs, we next sought to understand the unusual chromatin state of mammalian sperm by comparing our sperm and ESC data sets in more detail. First, it is notable that, in our protocol, MNase digestion of sperm results in a nucleosome ladder very similar to that observed for somatic cells (discussed later). While this was
expected for ESCs, the ladder observed for sperm is intriguing, given that only2% of histones are retained in mature mouse sperm. If sparse nucleosomes were randomly retained at 2% of genomic loci, then adjacent nucleosomes producing a dinucleosomal band would occur quite rarely; instead, the finding of a nucleosomal ladder indicates that nucleosome reten-tion in sperm generally occurs in blocks of multiple adjacent nucleosomes.
Turning to the deep sequencing data, we noted in initial sur-veys of genome browser tracks that nucleosome occupancy was fairly uniform across the genome in ESCs but was substan-tially more variable at the megabase scale in murine sperm, with long blocks exhibiting2- to 3-fold variation in average nucleo-some occupancy (Figures S1A and S1B available online). When comparing the enrichment of nucleosome-sized fragments (135– 165 bp) between mouse ES and sperm cells using a 2 Mb sliding window, we observed that differential nucleosome occupancy was strongly correlated with gene density (Figure 3A). Specif-ically, nucleosomes appear to be preferentially retained in gene-poor regions in sperm relative to ESCs. To quantitate this trend genome-wide, we binned the genome into 2 Mb bins and
plotted total nucleosome-sized sequencing reads versus the number of genes within each window. As expected, ESC chro-matin exhibits uniform nucleosome occupancy across the genome, whereas in sperm, the number of nucleosome-length fragments for a given window is strongly anticorrelated with the number of genes in that window (Figure 3B). Notably, the low nucleosome occupancy over promoters (Figure 1D) and gene-rich regions (Figure 3B) inferred from nucleosome-sized MNase footprints was also observed using low-coverage anti-histone H3 ChIP sequencing (ChIP-seq) (Figures S1C and S1D), validating the hypothesis that the150 bp MNase foot-prints (Figure 4A) represent bona fide histone retention in sperm. Several previous studies examining histone retention in murine and human sperm reported strong retention of nucleosomes in the promoter regions of genes implicated in early development; these regions generally are also characterized by high CpG density and low DNA methylation (Erkek et al., 2013; Hammoud et al., 2009). This can be observed in our reanalysis of those published data sets, as TSS-aligned aggregation plots of read coverage confirms strong retention of promoter nucleosomes under the reported conditions (Figure 4B). In contrast,
Figure 2. Relationship between Transcript Abundance and MNase Footprinting
(A and B) Enrichment of reads around TSSs is shown in a heatmap (red, enriched; green, depleted) for genes grouped according to mRNA abundance in E14 mouse ESCs: High (top 5%), Medium (middle 5%), and Low (bottom 5%). For each panel, the x axis shows distance from the TSS, and the y axis shows size of the MNase footprint. Data for ESC MNase digestion are shown in (A), and data for mature sperm are shown in (B). Note that sperm ‘‘expression levels’’ derive from mRNA abundance from round spermatids (Namekawa et al., 2006) as mature sperm are transcriptionally inactive.
(C) Gene-resolution heatmap of deep sequencing reads from ESC MNase digestion, aligned by TSS for three prominent size classes: <80 bp, 100–130 bp, and 135–165 bp. Each row is a single gene, and rows are sorted from high to low mRNA abundance.
Developmental Cell
nucleosome-sized MNase footprints in our data set were gener-ally depleted from promoters, including developmental pro-moters such as theHox promoters (Figures 2B,4C, andS1B). We speculated that these dramatic discrepancies might result from differences in the extent of MNase digestion; in yeast, several studies have reported a subset of so-called ‘‘fragile nucleosomes’’ that are particularly susceptible to extensive MNase digestion (Weiner et al., 2010; Xi et al., 2011). Deep sequencing of mononucleosome-sized DNA released from sperm by varying levels of MNase digestion revealed that nucle-osome depletion at promoters was also observed at relatively low MNase concentrations, arguing against promoter nucleo-somes being fragile (Figure 4C). In contrast, after extensive MNase digestion of sperm, we observed a150 bp footprint becoming enriched over promoters (Figure 4D), particularly over the ‘‘high CpG’’ class of promoters that are associated with developmental regulators (Figure S2A).
In addition to the extent of MNase digestion, our protocol differs from the Peters and Cairns protocols in that we digest formaldehyde-crosslinked sperm—we could not generate nucleosome-sized fragments from murine sperm in the absence of crosslinking—whereas both the Peters and Cairns groups carried out MNase digestions on uncrosslinked sperm that were first swelled in water/dithiothreitol (DTT) prior to MNase digestion. Notably, we found that, after centrifuging the sperm
MNase digests to separate primarily histone-associated DNA (supernatant) from the more abundant protamine-associated genome in the pellet, promoter retention of150 bp footprints was somewhat stronger in the pellet than in the supernatant ( Fig-ures S2B and S2C). Thus, we conclude that the previously re-ported histone retention at developmental promoters represents a specific subset of all histone retention in sperm in which promoter-bound nucleosomes are revealed only by extensive nuclease digestion—presumably, ‘‘overdigestion’’ eventually degrades the more abundant nucleosomes found over gene deserts, revealing the more stable promoter nucleosomes— and may furthermore be partially lost during centrifugation due to formaldehyde crosslinking to some insoluble scaffold.
Our hypothesis that histones associated with developmental promoters represent a small subset of the nucleosomes retained in sperm is also supported by elementary arithmetical consider-ations: human sperm retain10% of the nucleosome comple-ment of a somatic cell (Hammoud et al., 2009), yet even if histones were penetrantly retained in 100% of sperm across 1 kb of sequence at all 30,000 promoters, this would only account for 1% of the haploid genome. Nonetheless, we sought to independently assess our conclusion that most histone retention occurs in gene-poor regions in murine sperm ( Fig-ure 3B). Given the sensitivity to nuclease levels documented earlier for digestion-based assays, we used fluorescence
Figure 3. Preferential Histone Retention Occurs at Gene Deserts in Sperm
(A) Sperm nucleosomes are retained in gene-poor regions compared to ESC nucleosomes. Normalized mononucleosomal (135–165 bp) MNase footprints were averaged for 2 Mb bins, and the log2of the relative enrichment for ESCs versus sperm (y axis) is shown for a typical stretch of chromosome 8 (genomic coordinate
on x axis).
(B) Scatterplot of gene density (x axis) versus number of mononucleosome-sized MNase sequencing reads (y axis) for ESCs and sperm. The small number of points in the lower left corner depleted of sequencing reads correspond to 2 Mb bins composed of largely unmappable or unannotated sequence. Low-depth ChIP-seq using anti-H3 in sperm also reveals this anticorrelation between gene density and H3 signal, strongly supporting the claim that mononucleosome-sized fragments released by MNase digestion of sperm indeed correspond to nucleosomes.
in situ hybridization (FISH) and immunofluorescence as a completely independent assay for histone localization in mature mouse spermatozoa. Mammalian sperm cells contain a highly organized nucleus with a DAPI-dense heterochromatic center surrounded by a euchromatic periphery that exhibits more
modest DAPI staining (Li et al., 2008; van der Heijden et al., 2006). To determine the location within the sperm nucleus of CpG-rich developmental promoters, we generated fluorescent probes from 50 to 500 bp size-selected DNA fragments of HpaII-digested genomic DNA. Cot-1 DNA probes were used to
Figure 4. A Small Subset of Nucleosomes Is Retained at Promoters in Sperm
(A) Gel electrophoresis of soluble versus insoluble MNase digestion products from matureM. musculus sperm. Lane with asterisk designates level of digestion
used for most analyses. Pellet fraction contains the majority of protamine protein, while supernatant carries the majority of histone proteins. (B) Aggregation plot of data fromHammoud et al. (2009)(H. sapiens) andErkek et al. (2013)data sets (M. musculus) aligned by TSS.
(C) Aggregation plot of MNase sequencing reads for 1 U and 10 U MNase digestion levels [digestions shown in (A)]. These libraries were generated from the supernatant of the MNase digestion material.
(D) Aggregation plots of MNase sequencing data for unspun, not size-selected, digestion series of formaldehyde-crosslinkedM. musculus sperm. A peak of
promoter nucleosome occupancy is revealed only after extensive MNase digestion [note that MNase levels are not directly comparable to those in (C) as fewer sperm were used in this digestion] and is more prominent when MNase digestions are not subject to centrifugation.
(E) Representative images for FISH with HpaII probe (poorly methylated CpG islands; seeExperimental Procedures) and Cot-1 probe (gene-poor heterochro-matic regions), as well as immunofluorescence using antibodies to TH2B (testis-specific H2B) and bulk H4, demonstrating that the majority of anti-histone staining overlaps the DAPI-rich chromocenter that also stains with Cot-1 probe.
See alsoFigure S2.
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confirm the physical location of repetitive heterochromatic se-quences. Signal from the HpaII probe is prominent in the nuclear periphery, while Cot1 signal overlaps almost perfectly with the chromocenter, as expected (Figure 4E). We then used immuno-fluorescence to characterize the localization of histones H4 and TH2B, finding that the majority of histone staining occurs in the central DAPI-dense chromocenter, which is consistent with prior studies (Figure 4E) (Govin et al., 2007; Meyer-Ficca et al., 2013; van der Heijden et al., 2006). Taken together, our data confirm that a small subset of histone retention occurs at CpG-rich pro-moters but demonstrate that the vast majority of histone reten-tion in mammalian sperm occurs in gene deserts.
Short MNase-Protected Fragments Coincide with DNA-Binding Protein Footprints
We finally turn to more detailed analysis of short MNase foot-prints in sperm and ESCs. As shown elsewhere (Kent et al., 2011), short MNase-protected footprints can reveal binding sites for DNA-binding proteins such as TFs. In order to determine whether our data revealed the locations of specific DNA-binding proteins, we sought sequence motifs enriched in short MNase-protected fragments relative to the background of longer (nucleosome or dinucleosome) footprints (see Experimental Procedures). In ESCs, significantly enriched motifs included sequence matches for a number of TFs that play key roles in the pluripotency transcriptional network, such as Klf4, as well as a strong match to the binding site for key insulator and locus control regulator CTCF. Plotting short MNase footprints around in vivo binding sites for a variety of TFs reveals a range of foot-printing behaviors for TFs (Figure S3). Despite the lack of active transcription in sperm, we were also able to identify a number of significantly enriched motifs among the short MNase footprints in sperm. Sperm footprints did not carry strong matches to the Klf4 binding site or most other pluripotency TFs but instead were enriched for a subset of sequence motifs from the ESC motif list such as Stat3, RelA, and, most notably, CTCF (Figures 5A andS3). In contrast, a number of TFs, including key pluripo-tency factors Oct4, Nanog, Sox2, and Klf4, were associated with strong 1 to 80 bp footprints specifically in ESCs but not in sperm. Notably, we did not identify any motifs specifically protected in sperm but not in ESCs.
To further explore MNase footprinting patterns around TF binding sites, we focused on Klf4 and CTCF, both of which have previously been mapped in murine ESCs (Chen et al., 2008). Using matches to the relevant sequence motif located within previously identified ChIP-seq peaks as anchors (n = 15,657 for CTCF, n = 1,813 for Klf4), we generated V-plots for ESC MNase footprints (Figure 5B). As previously shown for yeast general regulatory factors such as Reb1 (Henikoff et al., 2011), three characteristics are apparent in these plots. First, strong enrichment in the 30 to 60 bp size range is observed centered on the TF motif, indicating presumed protection of DNA from MNase by the bound TF. Second, a V-shaped depletion of frag-ments results from MNase cleavage immediately adjacent to the TF. Finally, enriched 140 to 160 bp fragments located outside this V are consistent with well-positioned nucleosomes occur-ring flanking the TF binding sites, as previously reported for CTCF (Fu et al., 2008; Teif et al., 2012). Similar, albeit weaker, results were observed over Klf4 anchors.
In contrast to the extensive literature on ESC TF binding, little is known about the localization, or even presence, of sequence-specific DNA-binding factors in mature sperm. However, as noted earlier, we identified a number of sequence motifs en-riched among short MNase footprints in mature sperm, most notably, the CTCF motif (Figure 5A). We repeated the V-plot analyses for CTCF and Klf4 binding sites for sperm MNase data (Figure 5C). Consistent with our motif searches, we found little evidence for Klf4 binding in mature sperm, but we confirm enrichment for <60 bp fragments at CTCF binding sites (Figures 5C and 5E). The modest enrichment observed here relative to the ESC peak is explained by the fact that only a subset (40%) of the CTCF motifs that are bound in ESCs are associated with MNase footprints in mature sperm (Figure 5D).
Subnucleosomal MNase Footprints Represent Bona Fide CTCF Binding Events
The presence of sequence motifs underlying short MNase foot-prints provides us with strong hypotheses regarding TF binding events, which make several additional testable predictions. First, the extent of footprinting over CTCF motifs should correlate with in vivo occupancy levels as assayed by ChIP-seq. Second, these footprints should be eliminated by CTCF knockdown (KD). To test these predictions, we carried out ChIP quantitative PCR for six loci predicted to be CTCF binding sites in ESCs, as well as three CTCF motifs only exhibiting MNase footprints in sperm. All nine loci exhibited the expected CTCF binding in ESCs, and this binding was lost on CTCF KD (Figure S4A). More globally, we made use of published CTCF ChIP-seq localization data generated using ESCs (Chen et al., 2008). Enrichment of short MNase fragments over CTCF binding motifs was indeed corre-lated with in vivo CTCF occupancy, consistent with the idea that these short footprints correspond to CTCF protected fragments (Figure 6A). It is interesting that nucleosome phasing surrounding CTCF binding sites also correlated with CTCF binding levels (Figure 6B), supporting the hypothesis that CTCF acts as a positioning barrier to establish nucleosome arrays (Radman-Livaja and Rando, 2010).
Finally, we directly assessed whether short MNase footprints over CTCF binding sites resulted from bona fide CTCF binding (as opposed to MNase sequence biases or binding by a factor other than CTCF, such as its paralog CTCFL/BORIS) by repeating our MNase mapping experiment in ESCs following CTCF KD. Overall, we achieved a90% KD of CTCF as illus-trated by both western blotting and mRNA abundance (Figures 6C and 6D). Genome-wide, results for this experiment were strongly correlated with results from control or enhanced green fluorescent protein (EGFP) knockdown ESCs, with similar frag-ment size distributions, nucleosome depletion at promoters, and short transcription-correlated promoter footprints (data not shown). However, as predicted, short fragments over CTCF binding sites were greatly diminished in the CTCF KD cells ( Fig-ure 6F;Figures S4B and S4C). The remaining short fragments could result either from residual CTCF following incomplete (90%) knockdown or from the CTCFL/BORIS protein that binds a similar sequence motif to CTCF (Jelinic et al., 2006). Nucleosome phasing was also lost surrounding CTCF binding motifs in the knockdown cells (Figure 6F), providing strong evidence that CTCF binding (rather than some aspect of DNA
’’programming’’ of chromatin structure) is required for nucleo-some positioning at these locations.
Finally, we asked whether MNase footprints in sperm were likely to result from CTCF itself or from CTCFL/BORIS, which binds to the same sequence and plays important roles in male reproduction (Jelinic et al., 2006; Loukinov et al., 2002). First, western blotting confirms the presence of CTCF protein in mature sperm (Figure S4D), which is consistent with prior reports (Tang and Chen, 2006). Despite the presence of this protein in sperm, to date, we have been unable to successfully recover ChIP mate-rial; this is likely due to the unique crosslinking and fragmentation
challenges presented by the highly compact sperm nucleus. As an indirect approach to the question of the CTCF footprinting pro-tein, we focused on CTCF-specific and BORIS-specific binding events recently described in ESCs (Sleutels et al., 2012) and asked whether these sites were occupied by short MNase DNA fragments from our sperm data set. We find that <60 bp MNase footprints over CTCF motifs in sperm preferentially occur over sites bound exclusively (in ESCs) to CTCF, while they are completely absent over known CTCFL-only binding sites ( Fig-ure S5). These results support our hypothesis that CTCF itself is associated with the genome in mature murine sperm.
Figure 5. Enrichment of Short MNase Fragments over Binding Sites for Sequence-Specific TFs
(A) Short MNase footprints in sperm occur over CTCF binding sites. MNase digestion products (<60 bp) from sperm were searched for overrepresented sequence motifs, and the top motif hit is shown alongside the published sequence motif for CTCF.
(B and C) V-plots (as inFigure 2A, but showing all digestion products as points rather than enrichment/depletion) for ESC (B) or sperm (C) MNase digestion products, anchored by CTCF and Klf4 binding sites.
(D) Short footprints in sperm are associated with a subset of the CTCF binding sites identified in ESCs. Heatmap shows enrichment of <60 bp DNA fragments from a sperm data set, centered on CTCF motifs that are empirically CTCF-bound in ESCs (Chen et al., 2008). Sites are sorted from high to low signal intensity over 100 bp surrounding the binding site.
(E) Examples of short (<80 bp) footprints averaged over various TF motifs. See alsoFigure S3.
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DISCUSSION
By characterizing a wide size spectrum of MNase digestion products, we have profiled the chromatin architecture of two unusual mouse cell types, ESCs and sperm, illuminating nucleo-some occupancy and TF binding in each cell type and providing strong evidence for the presence of bound TFs such as CTCF in mature sperm.
ESC Chromatin
In ESCs, our analysis of mononucleosome-sized footprints confirms well-known aspects of chromatin biology, such as a strong relationship between promoter nucleosome depletion and transcription level. We therefore focused on analysis of features only appreciable in non-size-selected libraries. Extend-ing our analysis to short MNase footprints reveals a strong cor-relation between these footprints at the 50 ends of promoters and transcription rate, thus supporting the hypothesis (Teves and Henikoff, 2011) that these MNase footprints reflect the pres-ence of the basal transcription machinery (among other DNA-binding proteins).
Short MNase footprints were enriched for sequence motifs associated with a subset of key pluripotency regulators, such as Klf4. It is interesting that we did not observe equally strong
Figure 6. Nuclease-Resistant Footprints over CTCF Motifs Represent Bona Fide CTCF Binding in ESCs
(A) Abundance of short MNase footprints over CTCF binding sites in ESCs correlates with CTCF ChIP-seq enrichment. CTCF motifs are split into quintiles according to ChIP-seq signal (Chen et al., 2008).
(B) As in (A), but for mononucleosome-length footprints. Flanking nucleosomes are more strongly positioned when CTCF ChIP-seq signal is highest.
(C) Western blots of CTCF and EGFP esiRNA KD in mouse ESCs, probed with anti-CTCF andb-actin antibodies.
(D) Quantitative RT-PCR of CTCF mRNA abun-dance.
(E) Aggregation plot for <80 bp and 135–165 bp digestion products in EGFP KD ESCs, aligned using CTCF motifs.
(F). KD of CTCF in ESCs results in loss of <80 bp footprint enrichment over CTCF motifs, with an associated increase in nucleosome occupancy over the CTCF motif and loss of surrounding nucleosome positioning.
See alsoFigures S4andS5.
footprinting at experimentally validated binding sites for all TFs (Figure S3). This was previously observed in yeast studies, in which an unusual subset of TFs (largely corresponding to the abundant ‘‘general regulatory factors’’) showed far stronger footprinting than the majority of TFs (Henikoff et al., 2011). The absence of strong MNase footprinting at certain TF binding sites (such as for Esrrb in ESCs) could reflect weak crosslinking to DNA, low average occupancy, or inability of the DNA-bound TF to interfere with MNase digestion. It is interesting in this regard that the binding sites associated with strong footprints, such as CTCF and Klf4 binding sites, tended to be associated with relatively well-positioned flanking nucleosomes (Figure 5B). One potential explanation for this correlation would be that TF footprints are strongest for factors associated with particular chromatin re-modeling complexes.
We tested the hypothesis that these short MNase footprints report on bona fide TF binding in vivo, finding that CTCF foot-prints exhibited strong correlation with CTCF ChIP-seq data and further showing that MNase footprints over CTCF binding sites are greatly reduced on CTCF KD. Finally, we extended the prior observation that CTCF binding is associated with strong nucleosome phasing (Fu et al., 2008) by showing that the level of CTCF binding corresponds to the strength of nucle-osome positioning, supporting the hypothesis that TFs can function as positioning barriers. Moreover, loss of CTCF binding on KD leads to a loss of nucleosome phasing around this site. Taken together, these results lend further support to the hy-pothesis that TFs are capable of directing nucleosome positioning.
Analysis of Sperm Histone Retention
The unusual packaging state of mammalian sperm, which retain 2%–15% of the histone load of somatic cells (Ooi and Henikoff, 2007), has been proposed to play roles in programming early embryonic gene expression (Brykczynska et al., 2010; Gardiner-Garden et al., 1998; Hammoud et al., 2009) and in epigenetic inheritance of cytosine methylation states (Nakamura et al., 2012). The specialized subset of genomic loci associated with retained histones in sperm is, therefore, of great interest.
In principle, retention of nucleosomes over 2% of the genome in sperm could occur stochastically (e.g., each sperm cell retains nucleosomes over a different 2% of the genome), deterministi-cally (all sperm cells retain nucleosomes over specific loci), or somewhere in between. Prior reports on sperm histone retention in humans and mice, using nuclease sensitivity as a key assay (Arpanahi et al., 2009; Brykczynska et al., 2010; Hammoud et al., 2009), came to somewhat different conclusions. Two studies isolated solubilized mononucleosome-sized fragments after MNase digestion of human sperm, finding enrichment over the promoters of key developmental regulators, with a very strong correlation with GC content (Brykczynska et al., 2010; Hammoud et al., 2009; Vavouri and Lehner, 2011). Some-what different results were obtained via tiling microarray analysis of DNA released after light restriction enzyme or MNase diges-tion of human and mouse sperm, with gene and promoter-rich regions flanked by CTCF sites very broadly overrepresented in much longer (megabase-scale) domains (Arpanahi et al., 2009; Saida et al., 2011).
Here, we confirm that nucleosome-sized fragments can be found over CpG-rich promoters but find that these nucleosomes are unusual in that they are only released by extensive MNase digestion and appear to be crosslinked by formaldehyde into a somewhat insoluble complex. Moreover, we show that these nucleosomes represent a small fraction of all nucleosomes retained in mature sperm; preferential histone retention over gene deserts is observed robustly across a range of MNase digestion levels, and fluorescence microscopy independently confirms that the majority of histone protein is localized in a distinct chromocenter occupied by repeat elements rather than the CpG-rich fraction of the genome associated with develop-mental promoters (Figure 4E). Whether either of these classes of histone-bound loci serves any function in early embryonic pro-cesses (seeIhara et al., 2014)—and how these functions might differ between the broad blocks of nucleosomes occurring over repeat elements (e.g., gene deserts) and the more focal nucleosomes found over CpG islands—is currently un-clear. Notably, recent studies have shown that retention of H3K9me2-marked histones plays a role in protecting a fraction of the paternal genome from Tet3-dependent cytosine demethy-lation (Nakamura et al., 2012). Preliminary low-coverage anti-H3K9me2 ChIP-seq shows preferential localization of this mark to gene-poor regions of the genome (data not shown). Given that we found histone retention primarily occurring over gene de-serts, our results suggest a model in which the sperm packaging state preferentially protects epigenetic silencing marks from removal at repeat elements. Advances in editing the epigenome should allow targeting histone eviction or covalent modifications at specific loci in gametes to definitively determine the role for sperm histones in zygotic biology.
Finally, we provide evidence that a subset of CTCF binding sites is associated with strong MNase footprints in mature sperm. This suggests the hypothesis that mature sperm carry chromatin-associated CTCF at specific genomic locations. CTCF protein has previously been reported to be present in mature mouse sperm (Tang and Chen, 2006), although it was not shown to associate with the sperm genome. CTCF is intri-cately involved in epigenetic control of a number of well-studied imprinted genes (Phillips and Corces, 2009), raising the intriguing possibility that CTCF associated with the paternal genome plays a regulatory role in gene expression during early development. Future studies using conditional alleles of CTCF will be needed to assess if and how chromatbound CTCF protein in sperm in-fluences gene expression on zygotic genome activation and whether inheritance of CTCF-bound sperm nuclei is essential for developmental gene regulation.
EXPERIMENTAL PROCEDURES Tissue Culture and Sperm Isolation
ESC lines used in this study were mouse E14 lines cultured using standard conditions. Briefly, E14 ESCs were cultured on gelatin-coated dishes without feeder cells in standard media containing serum and leukemia inhibitory factor at 37C with 5% CO2. ESCs were passed by washing with PBS and
dissoci-ating with trypsin. Sperm cells were isolated from the caudal epididymis and vas deferens of 10-week-old C57/6J mice by lacerating tissue in 1 ml of 37C M2 medium (Sigma). Sperm cells were allowed to swim up for 1 hr, washed 13 in water, 13 PBS, crosslinked at 37C with 1% formaldehyde for 15 min, quenched with 250 mM glycine for 5 min, and frozen in liquid nitro-gen. The majority of figures were generated using data from control EGFP KD ES cultures, with the exception ofFigures 4E andS3, which used untreated ESC data that had higher levels of short MNase footprints. Control and EGFP KD data yielded nearly identical results; EGFP KDs were used simply based on the higher sequencing depth for these libraries. Animals were maintained on site in accordance with approved Institutional Animal Care and Use Committee protocols.
Sperm Permeabilization
Frozen sperm cells were briefly thawed at 37C, pelleted 4,0003 g for 4 min at room temperature (RT), and washed 13 in 1 ml PBS; then, the pellet was resuspended in 1 ml sperm permeabilization buffer (SPB) (0.025% trypsin/ EDTA; 10 mM DTT; 5 mM Tris, pH 7.5; 5 mM NaCl; 1.5 mM MgCl2; 0.25%
NP40; 1mM phenylmethylsulfonyl fluoride [PMSF]). Sperm cells were incu-bated in SPB for 5 min at 37C. After 5 min, 10ml of fetal bovine serum was added to inactivate trypsin, and the sample was incubated for an additional 5 min at 37C. Sperm cells were pelleted 4,0003 g for 4 min at RT, washed with PBS 23, resuspended in 500 ml PBS + 2 mM CaCl2, and incubated
5 min at 37C. RNAi-Mediated KD
RNAi using endoribonuclease III small interfering RNAs (esiRNAs) was per-formed as previously described (Fazzio et al., 2008), transfecting E14 mouse ESCs using Lipofectamine 2000. Briefly, templates for esiRNA production were produced using a two-step PCR procedure followed by in vitro transcrip-tion and dicing of double-stranded RNA using RNase III. KDs were performed for 48 hr to achieve effective KD.
MNase Digestion
Permeabilized ES and sperm nuclei were treated with 10 U per 106
cells and 1 U per 108
cells of MNase (Worthington), respectively, for 5 min at 37C. Reaction was stopped with the addition of 10 mM EGTA on ice. For ESCs, nuclei were incubated for 4 hr at 4C with rotation, pelleted at 5,0003 g for 5 min, and separated into supernatant and pellet (spun) or were treated the same without centrifugation and separation (unspun). After MNase digestion of sperm nuclei, digestion was centrifuged for 5,000 3 g for 5 min: the
Developmental Cell
supernatant from this spin contained the vast majority of histone protein, whereas the vast majority of protamine protein was located in the pellet (data not shown). All sperm data except inFigures 4D,S2A, and S2C were generated using the supernatant fraction; the sperm data inFigures 4D and
S2A were generated using libraries constructed without any centrifugation step. ForFigures S2B and S2C, libraries were prepared from sperm digested with a range of MNase concentrations—1, 5, 20, and 100 U—for 5 min at 37C, where digestion was stopped with EGTA on ice, and incubated for 4 hr at 4C. Supernatant and pellet fractions were separated by centrifugation at 5,0003 g for 5 min, and the150 bp mononucleosome-sized DNA was used to generate deep sequencing libraries. In all cases, DNA was isolated by phenol:chlorofor-m:isoamyl alcohol and EtOH precipitation.
Sperm ChIP-Seq
Mouse sperm cells were isolated as described earlier. Chromatin isolated from six mice was used for the immunoprecipitation experiments. Crosslinked sperm cells were washed with 13 with PBS, resuspended in 600 ml of 20 mM Tris with 20% glycerol and 10 mM PMSF and subjected to bead-beating (0.5 mm glass beads). SDS was added to a final concentration of 1%, and chromatin was sheared to300 bp with Bioruptor. Samples were kept on ice at all stages. ChIP was performed overnight at 4C with Pan-H3 antibody (abcam, ab1791).
Deep Sequencing
To characterize the entire range of MNase digestion products, we generated paired-end libraries of DNA fragments from both ES and sperm digestions and performed sequencing using Illumina HiSeq technology. Libraries were prepared as described byHenikoff et al. (2011). Reads were mapped to the mouse genome (mm9) using Bowtie2, and uniquely mapped reads were used for further analysis. For library insert size distributions, seeFigure 1B. For most figures, reads were normalized to parts per million reads. InFigures 4,6, andS2, data are normalized to the average genome-wide coverage and represented as log2of the fold difference from the genomic average.
The majority of analyses were carried out using an ESC (unspun) data set comprising 388,389,043 total reads obtained from aggregating two biological replicates, an ESC (spun) data set with 384,462,519 reads, and a sperm data set comprising 389,791,625 reads obtained from aggregating three biological replicate data sets.
Motif Analysis
We first trimmed sequencing adaptors, considering reverse complementarity of the paired-end reads, to recover reads sequenced from short DNA frag-ments. Then, all reads were aligned to mouse genome mm9 by Bowtie2 with default parameters (end-to-end alignment mode and random selection for multiple mappers). Only concordantly aligned read pairs were retained, and the length of the sequenced DNA fragments was calculated according to their mapped coordinates. Reads representing DNA fragments of mononucleoso-mal MNase (135–165 bp), subnucleosomononucleoso-mal MNase (100–130 bp), and TF-sized footprints (<80 bp) were segregated for several analyses, as indicated. We per-formed peak calling from TF-sized reads against other two groups (as back-ground inputs) using the Model-based Analysis for ChIP-seq. We further selected the top 1,000 most significant peaks and refined their borders by ex-tending 20 bp both up- and downstream of the peak summits, which are the points with highest read coverage within peaks. We then searched for enriched motifs by using MEME (Multiple EM for Motif Elicitation) among the genomic sequences of the refined peaks. Enriched de novo motifs were compared to known motifs using TOMTOM with the motif frequency models from Jaspar. FISH
Sperm cells were collected by swim-up as previously described, and slides were prepared as described byvan der Heijden et al. (2006). Sperm cells placed on coverslips were permeabilized in CSK buffer, 5% triton, and vanadyl ribonucleoside complex for 3 min; then, they were fixed in 4% paraformalde-hyde for 10 min and stored in 70% EtOH. Antibodies used for immunohisto-chemistry were Pan-H4 (Upstate, catalog number 05-858, lot number 27171) and TH2B (Abcam, catalog number ab23913, lot number 825930). Primary an-tibodies were detected with anti-rabbit 488 (Jackson ImmunoResearch). Slides were incubated with 1:250 primary antibody in 1% BSA, 13 PBS for
1 hr at 37C and then washed and immunodetected using a 1:500 dilution of conjugated (Alexa Fluor 488 or Alexa Fluor 594, Invitrogen) secondary (anti-goat, -mouse, or -rabbit) antibody in 13 PBS with 1% BSA. HpaII probe was prepared by performing an overnight restriction enzyme digest on purified mouse sperm DNA, followed by gel purification of 50 to 500 bp DNA fragments, ligation of adapters, and PCR amplification in the presence of biotin-dCTP. Mouse CoT-1 DNA (Invitrogen) probe was nick-translated using biotin-11-dUTP (Roche). Two hundred nanograms of HpaII biotinylated probe or 50 ng CoT-1 biotin nick-translated probe was hybridized to fixed and dena-tured cells.
ACCESSION NUMBERS
Data will be available via the Gene Expression Omnibus database under accession ID GSE58101.
SUPPLEMENTAL INFORMATION
Supplemental Information includes five figures and can be found with this article online athttp://dx.doi.org/10.1016/j.devcel.2014.05.024.
ACKNOWLEDGMENTS
We thank K. Ahmad and members of the O.J.R. laboratory for insightful discussions and comments on the manuscript. O.J.R. is supported in part by grant HD080224 from the Eunice Kennedy Shriver National Institute of Child Health and Human Development (NICHD), and by the Harold and Leila Mathers Charitable Foundation. T.G.F. is supported in part by grant HD072122 from NICHD and is a Pew Scholar in the Biomedical Sciences. Z.W. is supported by National Science Foundation grant DBI-0850008. B.R.C. is supported by a T32HD007439 training grant. S.J.H. is supported by a T32CA130807 training grant and by a Leukemia and Lymphoma postdoctoral fellowship. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
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