Genome Distribution of Replication-independent Histone H1 Variants Shows H1.0 Associated with Nucleolar Domains and H1X with RNA Polymerase II-Enriched Regions*

Unlike core histones, the linker histone H1 family is more evolutionarily diverse and many organisms have multiple H1 variants or subtypes. In mammals, the H1 family includes seven somatic H1 variants, H1.1 to H1.5 being expressed in a replication-dependent manner, whereas H1.0 and H1X are replicationindependent. Using ChIP-seq data and cell fractionation, we have compared the genomic distribution of H1.0 and H1X in human breast cancer cells, in which we previously observed differential distribution of H1.2 compared to the other subtypes. We have found H1.0 to be enriched at nucleolus-associated DNA repeats and chromatin domains, whereas H1X is associated with coding regions, RNA polymerase II-enriched regions and hypomethylated CpG islands. Further, H1X accumulates within constitutive or included exons and retained introns, and towards the 3’ end of expressed genes. Inducible H1X knockdown does not affect cell proliferation but dysregulates a subset of genes related to cell movement and transport. In H1X-depleted Genomic Distribution of Replication-independent H1 Variants 2 cells, the promoters of up-regulated genes are not occupied specifically by this variant, have a lower than average H1 content and, unexpectedly, do not form an H1 valley upon induction. We conclude that H1 variants are not distributed evenly across the genome and may participate with some specificity in chromatin domain organization or gene regulation. INTRODUCTION There are five major classes of histones that participate in the correct folding of eukaryotic DNA into chromatin: the core histones H2A, H2B, H3, and H4 that form an octamer and constitute the nucleosome core particle, and the linker histone H1, which binds to the nucleosomes near the entry/exit sites of linker DNA. Stabilization of the condensed states of chromatin is the function most commonly attributed to the linker histone (1,2), in addition to its inhibitory effect in vitro on nucleosome mobility (3) and transcription (4). Histone H1 in humans is a family of closely related, single-gene encoded proteins, including seven somatic subtypes (from H1.1 to H1.5, H1.0 and H1X), three testis-specific variants (H1t, H1T2 and HILS1) and one restricted to oocytes (H1oo) (5,6). Among the somatic histone H1 variants, H1.1 to H1.5 are expressed in a replication-dependent manner, while H1.0 and H1X are replication independent. The H1.1 to H1.5-encoding genes are clustered in a region of chromosome 6 together with the core histone genes, while H1X and H1.0 genes are on chromosomes 3 and 22, respectively. H1.2 to H1.5 and H1X are ubiquitously expressed, H1.1 is restricted to certain tissues, and H1.0 accumulates in terminally differentiated cells. There are few studies characterizing the most recently identified and distantly related human variant H1X, and its specific function in the cell remains unknown. Like H1.0, it has been suggested that H1X is enriched in a less accessible region of chromatin, but expression of the two variants is regulated differently (7). It has been shown previously that H1X accumulates in nucleoli in G1 and is distributed across the entire nucleus in the S phase (8). The same year, Takata et al. found that H1X was preferentially located at the chromosome periphery in mitosis, and they observed defects in chromosome alignment and segregation after H1X knock-down (KD) (9). Taken together, these findings indicate that H1X may have functions that differ from those of the other variants. As it participates in the formation of higher-order chromatin structures, H1 is seen as a structural component related to chromatin compaction and inaccessibility to transcription factors and to RNA polymerase. Nonetheless, it has also been suggested that histone H1 plays a more dynamic and gene-specific role, participating in the regulation of gene expression. Previous studies on the effect of H1 depletion on global gene expression have found no effect on the vast Genomic Distribution of Replication-independent H1 Variants 3 majority of genes but rather have detected upor down-regulation of small groups of genes (10-13). It is not clear whether the different variants have specific roles or regulate specific promoters. In mice, single or double H1 variant knock-outs have no apparent phenotype due to compensatory upregulation of other subtypes (14). These reports have favored the view that H1 variants are redundant. On the other hand, we reported that depletion of single H1 subtypes by inducible RNA interference in breast cancer cells produced a range of phenotypic effects (10), suggesting different functions for the various H1 variants in somatic cells. Furthermore, H1 subtypes can be posttranslationally modified and these modifications modulate their interaction with various other proteins. This could explain some reported specific functions for certain H1 variants (15-24). Moreover, H1 subtypes have cell typeand tissuespecific expression patterns and their expression is regulated over the course of differentiation and development (25-30). Different H1 subtypes have also been differentially related to cancer processes (31-34). To fully understand the function of histone H1 and its variants, several studies have explored the genomic distribution of H1 in vivo. Initial biochemical and microscopy-based approaches suggested a non-uniform distribution of H1 in the cell nucleus and found differences between variants (35-37). However, due to the lack of specific ChIP-grade antibodies for most H1 variants, it has been challenging to identify the precise mapping of H1 variants in the genome until recently. Two reports, using ChIP of tagged H1 variants in mouse embryonic stem cells and DamID technology in human IMR90 cells, respectively, showed depletion of H1c and H1d from guanine-cytosine (GC)and gene-rich regions, as well as an overrepresentation in major satellites (38), and depletion of H1.2 to H1.5 from CpG-dense and regulatory regions, only H1.1 having a distinct profile (39). Moreover, it has previously been shown that when a gene is transcriptionally active, there is depletion of H1 (an H1 valley) at the transcription start site of its promoter (40). Using variant-specific antibodies against H1 and hemagglutinin (HA)-tagged recombinant H1 variants expressed in breast cancer cells, we investigated the distribution of six H1 variants in promoters (ChIP-chip) and genome-wide (ChIPseq), including H1.0 and H1X for the first time (41). In short, we reported that histone H1 is not uniformly distributed across the genome and there are differences between variants, H1.2 showing the most specific pattern and strongest correlation with low gene expression. H1.2 is enriched at chromosomal domains with low GC content, and is associated with gene-poor chromosomes, intergenic DNA and lamina-associated domains (LADs). Meanwhile, other variants are associated with higher GC-content, CpG islands and geneGenomic Distribution of Replication-independent H1 Variants 4 rich domains. Overall, the distribution of H1.2 along chromosomes differed from that of other variants including H1.0 and H1X, the two variants most structurally distant within the somatic H1 family. In this new work, we have further analyzed the distribution of H1 variants in other genomic regions including repetitive DNA, nucleolusassociated chromatin domains (NADs), and ribosomal DNA (rDNA), and their association with methylated CpG sites and RNA polymerase II-­‐enriched regions. This analysis has revealed that H1.0 and H1X are enriched at particular regions compared to the other variants. H1.0 is the variant that is most abundant at NADs, rDNA and certain satellite repeats related to nucleolus organizer regions (NORs). The association of H1.0 with nucleolar chromatin has been confirmed by immunoblotting on fractionated cellular extracts. In contrast, H1X is associated with RNA polymerase II-­‐enriched sites, coding regions and hypomethylated CpG islands. Notably, the H1X content at coding regions is higher at active genes, especially towards the 3’ end of genes, and more abundant at exons and intron-exon junctions than within introns themselves. We have also further investigated the functionality of H1X by testing the effect of an inducible KD of this H1 variant on cell proliferation and global gene expression. EXPERIMENTAL PROCEDURES Cell lines and culture conditions–T47D-MTVL (carrying one stably integrated copy of luciferase reporter gene driven by the MMTV promoter) (42), and MCF7 breast cancer cells were separately grown at 37oC with 5% CO2. T47Dderivative cells were grown in RPMI 1640 medium, supplemented with 10% FBS, 2 mM Lglutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. MCF7 cells were grown in MEM medium containing 10% FBS, 1% penicillin/streptomycin, 1% glutamine and 1% sodium pyruvate. Doxycycline (Sigma) was added at 2.5 μg/ml when required. Drug-inducible RNA interference–H1X KD cell lines were established from T47D-MTVL and MCF7 breast cancer cells. Plasmids for the lentivirus vector-mediated drug-inducible RNA interference system (pLVTHM, ptTR-KRAB-Red, pCMC-R8.91 and pVSVG) were provided by Dr. D. Trono (University of Geneva) (Wiznerowicz et al., 2003). After testing five shRNAs against H1X from the MISSION library (Sigma-Aldrich), the 21-mer H1X-specific target sequence 5’CAACGGTTCCTTCAAGCTCAA-3’ was chosen to generate the inducible system. The 71-mer oligonucleotides for shRNA cloning into Mlu/ClaI-digested pLVTHM were designed, annealed and phosphorylated as recommended by Dr. Trono (http://tronolab.epfl.ch/). For the production of viral particles containing the lentiviral vector and infections see Sancho et al. (10). The inducible knocked-down cell lines were Genomic Distribution of Replication-independent H1 Variants 5 sorted in a FACSCalibur machine (Becton Dickinson) for RedFP-positive and GFP-positive fluorescence after 3 days of doxycycline (Dox) treatment. Then, cells were amplified in the absence of Dox until an experiment was performed. Over a 6-day treatment with Dox, cells were passaged on day 3. When required, serumcontaining media was replaced with serum-free media on day 4 to arrest growth. Histone H1 extraction, gel electrophoresis and immunoblotting–Histone H1 was purified by lysis with 5% perchloric acid for 1 hour at 4oC. Soluble acid proteins were precipitated with 30% trichloroacetic acid overnight at 4oC, washed twice with 0.5 ml of acetone and reconstituted in water. Protein concentration was determined with the Micro BCA protein assay (Pierce). Purified histones were exposed to SDS-PAGE (10%), transferred to a PVDF membrane, blocked with Odyssey blocking buffer (LI-COR Biosciences) for 1 hour, and incubated with primary antibodies overnight at 4oC and with secondary antibodies conjugated to fluorescence (IRDye 680 goat antirabbit IgG, Li-Cor) for 1 hour at room temperature. Bands were visualized in an Odyssey Infrared Imaging System (Li-Cor). Polyclonal antibodies specifically recognizing human H1 variants, including those generated in our laboratory (10), are available from Abcam: H1.0 (ab11079), H1.2 (ab17677), H1.3 (ab24174), H1.4-T146p (ab3596), H1.5 (ab24175) and rabbit antiH1X (ab31972). Mouse antiH1X was obtained from Sigma (SAB1400328). Other antibodies used were: beta-tubulin (Sigma, nrT4026), nucleophosmin (Abcam, ab15440), nucleolin (Abcam, ab22758), H3K4me3 (Millipore, 07-473), and H3K9me3 (Abcam, ab8898). Cell fractionation for nucleoli purification–Cell fractionation was performed as described by Andersen et al. (43). Briefly, 30 million cells were resuspended in 1 ml of Buffer A (10 mM HEPESKOH pH 7.9; 1.5 mM MgCl2; 10 mM KCl; 0.5 mM DTT and protease inhibitors: 1 mM phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, 0.1 U/ml aprotinin, 1 mM orthovanadate, and 50 mM NaF), and incubated for 10 min on ice. Then, the cell pellet was homogenized, by passing the cell suspension through a 23-gauge needle 15 times and 25-gauge needle 10 times. From this, we collected the total protein fraction. The homogenized suspension was pelleted at 228g for 5 min at 4oC, and the supernatant was taken as the cytoplasmic fraction. The remaining pellet was resuspended in Buffer B (0.25 M sucrose; 10 mM MgCl2 and protease inhibitors), and was homogenized again, by passing the suspension through a 23-gauge needle 10 times. Then, it was centrifuged at 1430g for 5 min at 4oC on a sucrose cushion (Buffer C: 0.35 M sucrose; 0.5 mM MgCl2 and protease inhibitors). The remaining pellet was resuspended with Buffer C and sonicated for 6 cycles of 10 seconds on ice. The sonicated sample was centrifuged at 2800g for 10 min at 4oC on a sucrose pillow (Buffer D: 0.88 Genomic Distribution of Replication-independent H1 Variants 6 M sucrose; 0.5 mM MgCl2 and protease inhibitors). The supernatant was collected as the nucleoplasm fraction. The nucleoli pellet was washed with Buffer C and centrifuged at 200g for 2 min at 4oC. Then, it was resuspended with Lysis Buffer (SDS 2%; 67 mM Tris-HCl, pH 6.8). Protein concentration in all fractions was determined with the Micro BCA Protein Assay Kit (Pierce). Fractionated extracts were exposed to SDS-PAGE (10%), transferred to a PVDF membrane, and immunoblotted as described above. Immunoblot band intensities were measured using ImageJ (Version 1.48) software and normalized by Coomassie staining. Immunostaining–Cells were grown over coverslips, washed twice with PBS, and fixed with 4% formaldehyde for 15 min at room temperature. After three washes, they were permeabilized with Triton-X-100 for 15 min at room temperature and blocked with bovine serum albumin for 1 h. Then, the cells were incubated with primary antibodies diluted with bovine serum albumin for 1 h at room temperature in darkness. After the pertinent washes, the secondary antibodies Alexa-555 and Alexa-647 were added for 1 h at room temperature in darkness. The nucleus was stained with DAPI. The coverslips were mounted on the glass slides using Mowiol mounting medium. The samples were visualized by confocal laser scanning microscopy using a Leica TCS SPE system. Cell cycle analysis–Cells were washed with cold PBS 1x, fixed in 70% ethanol and stained with Analysis solution: 3% Ribonuclease A (Sigma) 10 mg/ml, and 3% solution A (38 mM sodium citrate, 500 μg/ml propidium iodide) in PBS 1X. Samples were analyzed with a FACSCalibur machine, using CellQuest Pro Analysis software (both from Becton Dickinson) and ModFit LT software (Verity Software House). Chromatin immunoprecipitationImmunoprecipitation of chromatin was performed according to the Upstate (Millipore) standard protocol. Briefly, cells were fixed using 1% formaldehyde for 10 min at 37oC, harvested and sonicated to generate chromatin fragments of 200 to 500 bp. Then, 20 μg of sheared chromatin was immunoprecipitated overnight with 2 μg of antibody. Immunocomplexes were recovered using 20 μl of protein A magnetic beads, washed and eluted. Cross-linking was reversed at 65oC overnight and immunoprecipitated DNA was recovered using the PCR Purification Kit from Qiagen. Genomic regions of interest were identified by real-time PCR (qPCR) using SYBR Green Master Mix (Invitrogen) and specific oligonucleotides in a Roche 480 Light Cycler machine. Each value was corrected by the corresponding input chromatin sample. Oligonucleotide sequences used for the amplifications are shown in Table 1. RNA extraction, reverse transcriptase qPCR and expression microarrays–Total RNA was extracted using the High Pure RNA Isolation Kit (Roche). Then, cDNA was generated from 100 ng of RNA Genomic Distribution of Replication-independent H1 Variants 7 using the Superscript First Strand Synthesis System (Invitrogen). Gene products were analyzed by qPCR, again using SYBR Green Master Mix (Invitrogen) and specific oligonucleotides in a Roche 480 Light Cycler machine. Each value was corrected by human GAPDH and represented as relative units. Each experiment was performed in duplicate. Gene-specific oligonucleotide sequences are shown in Table 1. The procedures for microarray hybridization using an Agilent platform (SurePrint G3 Human Gene Expression 8x60K v2) and data analysis are described elsewhere (41). Gene ontology analysis was performed using the DAVID software (Database for Annotation, Visualization and Integrated Discovery). Analysis of ChIP-seq data–As there are a limited number of H1-variant specific ChIP-grade antibodies (only H1.2 and H1X being available to us), we developed T47D-derived cell lines stably expressing hemagglutinin (HA)-tagged versions of each of the five somatic H1 variants expressed in most cell types (H1.0, H1.2, H1.3, H1.4 and H1.5) (10). Therefore, in addition to using H1.2 and H1X antibodies to pull down these variants in parental T47D cells, an anti-HA antibody was used to specifically pull down H1-associated chromatin fragments in cells expressing H1-HAs. ChIP-chip and ChIP-seq data on the occurrence of H1 variants at promoters and genome-wide in T47D-derivative cells, respectively, was generated in previous research and the analysis reported elsewhere (41). Briefly, ChIP-seq libraries were prepared with the ChIP-seq Sample Preparation Kit (Illumina) and sequencing was performed with Illumina HiSeq 2000 system. Read mapping and peak detection methods have been described before (41). Other types of analysis used are as follows. Publicly available genome-wide location data analysis: genomic locations of CpG islands and LADs (in hg18) were taken from the UCSC database (44), genomic locations of NADs from Nemeth et al. (45) and RNAPII binding sites from Ballaré et al. (46). Further, acromeric satellite 1 (ACRO1) genomic locations (in hg 18) were taken from the UCSC database. Repetitive sequences were taken from RepBase database (47). The mean methylation levels at individual CpG islands were calculated, by assessing the overlap between the methylation levels from Vanderkraats et al. (48) and the CpG islands using BedTools (49). The genomic locations of hyperand hypomethylated regions from the T47D cell line were re-calculated as in (50), the source of the raw data. RNA-Seq data from T47D cell line was taken from Vanderkraats et al. (48). Reads were mapped to the hg18 genome using the TopHat algorithm (version 2.0.12) (51). Next, we extracted the database of “cassette” exons and retained introns included in the MISO software package (52). The inclusion levels (Ψ) of alternatively spliced exons (ASEs) and retained introns were estimated using the MISO algorithm (52) with default parameters. Genomic Distribution of Replication-independent H1 Variants 8 Exons with inclusion level Ψ ≥ 0.9 were considered to be included ASEs and those with a Ψ ≤ 0.1 excluded ASEs. Retained introns with a Ψ ≥ 0.9 were considered to be retained in the T47D cell line. H1 occupancy at genomic features: Inputsubtracted normalized average H1 variant read density was calculated at each location enriched in CpG islands, NADs, hyperand hypomethylated regions, ACRO1, repetitive elements, exons, introns, ASEs, retained introns and RNAPII, and represented in box plots using in-house R scripts. As a control, a random sample of bulk genomic windows with equal width was used to perform the significance test (Kolmogorov-Smirnov test). In addition, for H1 abundance at ACRO1, a second method was used, namely, mapping to sequences in RepBase (47) with the bowtie aligner (53) allowing for multiple positions. ChIP signals around the center of RNAPII binding sites were calculated using normalized input subtracted-average tag numbers in each 50-bp bin in a set window. Relative distances of each tag from the aforementioned positions and average signals were determined using the ‘Sitepro’ script from the CEAS package (54) and plotting using R. Continuous ChIP signal profile distribution of reads along the meta-gene, exons and introns were performed using CEAS (54). Correlation analysis between NAD content and H1.0 abundance on individual chromosomes was performed using inhouse R scripts. H1 occupancy at individual chromosomes: Occupancy of H1 variants at all human chromosomes is measured in terms of the mean of the input-subtracted ChIP-seq signal in 50-bp windows. LAD and NAD occupancy at all chromosomes was calculated as the number of bases coinciding with LADs or NADs divided by the length of the chromosome. Expression on each chromosome is the mean of the expression of all genes in that particular chromosome. Heatmaps and dendrograms were created with in-house R scripts. H1 occupancy at rDNA: The abundance of H1 variants on rDNA was assessed as in (55). In short, as the rDNA sequence is not included in the reference genome, a custom hg18 assembly was constructed with the bowtie-build tool (53) adding a human rDNA repeat (GenBank accession no. U13369). Alignment was carried out with the bowtie aligner (53) allowing up to two mismatches and only unique hits were kept. The inputsubtracted ChIP-Seq signal in the rDNA sequence was calculated in reads per kilobase per million mapped and plotted using in-house R scripts. Overlap analysis of H1 islands: The number of enriched and depleted H1 islands which overlapped NADs, and RNAPII binding sites was calculated using BedTools (49) and plotted using in-house R scripts. Box plots showing the methylation levels of CpG islands overlapping enriched and depleted islands were calculated in Genomic Distribution of Replication-independent H1 Variants 9 the same way. Features were considered to overlap if the genomic intervals shared at least one base. Chromatin states based on the combined presence of H1 variant-enriched regions were calculated with a multivariate hidden Markov model using the chromHMM software (56). Human H1 variant nomenclature–The correspondence of the nomenclature of the human H1 variants with their gene names is as follows: H1.0, HIF0; H1.1, HIST1H1A; H1.2, HIST1H1C; H1.3, HIST1H1D; H1.4, HIST1H1E; H1.5, HIST1H1B; and H1X, HIFX. Accession numbers—The data sets are available in the Gene Expression Omnibus (GEO) database under the accession numbers GSE49345 and GSE62766. RESULTS Human H1 variants are differentially associated with NADs and repetitive DNA—To further explore whether the distribution of H1 variants is heterogeneous along several genome features or chromatin domains, we used our previously reported ChIP-seq data on endogenous H1.2, H1X, H3 and HA-tagged H1.0, H1.2 and H1.4 (41). DNA sequences associated with the human nucleolus have recently been identified and used to define NADs by Németh et al. (45). Different gene families and certain satellite repeats are the major building blocks of NADs, which constitute about 4% of the genome. Using the input-subtracted ChIP-seq signal, we investigated the occupancy of H1 variants within NADs. H1.0 was significantly enriched at NADs (Fig. 1A). Furthermore, H1.0 was the variant that had the largest number of H1-enriched regions overlapping NADs (Fig. 1B). A large part of chromosome 19 is associated with the nucleolus and is reported to be located in central regions of the interphase nucleus, being close to the nucleoli (45). We have previously reported that H1.0 is highly enriched at this chromosome (41). Correlation analysis between H1 variant ChIP-seq signals and NAD content at each chromosome confirmed that H1.0 is the most abundant variant at chromosomes with a higher NAD content, while H1.2 signals were negatively correlated with NAD content (Fig. 1C-D). As predicted, there was a negative association between the content of NADs and LADs at chromosomes, the former being located within the inner part of the nucleus and the latter at the periphery. We previously reported that H1.2 overlaps with LADs (41). Ribosomal DNA encoding the 45S single transcription unit giving rise to the 18S, 5.8S and 28S rRNA, and flanked by non-transcribed spacers, is present as repetitive DNA at the short arms of acrocentric chromosomes, called NORs, within NADs. We aligned the input-subtracted H1 variant ChIP-seq signal to the rDNA complete repeating unit and found that H1.0 was enriched in the rDNA, mostly in the non-transcribed spacers Genomic Distribution of Replication-independent H1 Variants 10 (Fig. 2A). Instead, at the single transcription unit, H1X was locally enriched. H1.0 was also enriched at 5S ribosomal RNA subunit (encoded in tandem arrays, the largest one on chromosome 1), whereas H1X was slightly enriched at microRNAs and small nucleolar RNAs, compared to other variants (Fig. 2B). Next, we aligned the input-subtracted H1 variant ChIP-seq signal to many repetitive DNA categories found in RepBase. One of the few categories that showed differential occupancy was ACRO1 (a 147-bp satellite found in the short arm of acrocentric chromosomes, where NORs are located), which presented H1.0 enrichment (Fig. 2C). H1.0 was also enriched at SINE-VNTR-Alus (SVAs, non-autonomous, hominid-specific nonLTR retrotransposons) and telomeric satellites (Fig. 2D). In summary, H1.0 is found to be enriched at DNA associated with nucleoli, including NADs, rDNAs, acrocentric and telomeric satellites, suggesting that it could be involved in the stabilization of perinucleolar late-replicating heterochromatin. In contrast, H1X is over-represented in the coding region of non-coding RNAs, such as 45S rRNA, miRNA and snoRNA, possibly related to the association of this variant with transcribed genes