Histone H1 depletion triggers an interferon response

Nucleic Acids Research, 2017 1 doi: 10.1093/nar/gkx746

Histone H1 depletion triggers an interferon response in cancer cells via activation of heterochromatic repeats Andrea Izquierdo-Bouldstridge1,† , Alberto Bustillos1,† , Carles Bonet-Costa1 , ´ 1 , Daniel Garc´ıa-Gomis1 , Marc Dabad2,3 , Anna Esteve-Codina2,3 , Patricia Aribau-Miralbes Laura Pascual-Reguant4 , Sandra Peiro´ 4 , Manel Esteller5,6,7 , Matthew Murtha5 , ´ ˜ 1 and Albert Jordan1,* Llu´ıs Millan-Ari no 1

Institut de Biologia Molecular de Barcelona (IBMB-CSIC), Barcelona, Catalonia 08028, Spain, 2 CNAG-CRG, Centre for Genomic Regulation (CRG), Barcelona Institute of Science and Technology (BIST), Barcelona, Catalonia 08028, Spain, 3 Universitat Pompeu Fabra (UPF), Barcelona, Catalonia 08003, Spain, 4 Vall d’Hebron Institute of Oncology, Barcelona, Catalonia 08035, Spain, 5 Cancer Epigenetics and Biology Program, Bellvitge Biomedical Research Institute (IDIBELL), L’Hospitalet, Barcelona, Catalonia 08028, Spain, 6 Physiological Sciences Department, School of Medicine and Health Sciences, University of Barcelona (UB), Catalonia 08028, Spain and 7 Institucio´ Catalana de Recerca i Estudis Avanc¸ats (ICREA), Barcelona, Catalonia 08028, Spain

Received May 26, 2017; Revised August 10, 2017; Editorial Decision August 11, 2017; Accepted August 15, 2017

ABSTRACT Histone H1 has seven variants in human somatic cells and contributes to chromatin compaction and transcriptional regulation. Knock-down (KD) of each H1 variant in breast cancer cells results in altered gene expression and proliferation differently in a variant specific manner with H1.2 and H1.4 KDs being most deleterious. Here we show combined depletion of H1.2 and H1.4 has a strong deleterious effect resulting in a strong interferon (IFN) response, as evidenced by an up-regulation of many IFN-stimulated genes (ISGs) not seen in individual nor in other combinations of H1 variant KDs. Although H1 participates to repress ISG promoters, IFN activation upon H1.2 and H1.4 KD is mainly generated through the activation of the IFN response by cytosolic nucleic acid receptors and IFN synthesis, and without changes in histone modifications at induced ISG promoters. H1.2 and H1.4 co-KD also promotes the appearance of accessibility sites genome wide and, particularly, at satellites and other repeats. The IFN response may be triggered by the expression of noncoding RNA generated from heterochromatic repeats or endogenous retroviruses upon H1 KD. In conclusion, redundant H1-mediated silencing of heterochromatin is im-

* To †

portant to maintain cell homeostasis and to avoid an unspecific IFN response. INTRODUCTION There are five major classes of histones that participate in the correct folding of eukaryotic DNA into chromatin: the four core histones H2A, H2B, H3 and H4 which form an octamer that constitutes the nucleosome core particle, and the linker histone H1, which binds nucleosomes near the entry/exit sites of linker DNA. Stabilization of the condensed states of chromatin is the function most commonly attributed to linker histone (1), in addition to its inhibitory effect in vitro on nucleosome mobility (2) and transcription (3). Unlike core histones, the H1 histone family is more evolutionary diverse and many organisms have multiple variants or subtypes. In humans, histone H1 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) (4–6). 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. 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. One of the major open questions in the field is whether different somatic H1 subtypes are mainly redundant or have specific functions, functions that are perhaps cell type

whom correspondence should be addressed. Tel: +34 402 0487; Fax: +34 403 4979; Email: [email protected]

These two authors contributed equally to this work as first authors. ˜ Integrated Cardio Metabolic Centre, Department of Medicine Huddinge, Karolinska Institutet, Stockholm, Sweden. Present address: Llu´ıs Mill´an-Arino,

C The Author(s) 2017. Published by Oxford University Press on behalf of Nucleic Acids Research. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact [email protected]

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specific. It is well established that H1 subtype composition varies through development and differentiation as well as between cell types and during disease associated processes such as neoplastic transformation (7–15). H1 subtypes are post-translationally modified, both at conserved and unique residues, and these modifications may modulate their interaction with an increasing number of proteins at the nucleoplasm or in chromatin (16,17). These interactions could explain some reported specific functions for certain H1 variants (18–23). Furthermore, genomic distribution of H1 variants is not absolutely redundant, with enrichment of different variants at certain chromatin types (reviewed in (6)). We have previously investigated the distribution of four H1 variants genome-wide in breast cancer cells, concluding that H1 variants are not distributed uniformly along the genome, H1.2 being the one showing the most specific pattern and strongest correlation with low gene expression (24,25). Previous studies on the effect of H1 depletion on global gene expression have found no effect on the vast majority of genes, but rather have detected variant specific up- or downregulation of small subsets of genes (26–29). However, it is not clear whether these effects are driven by variant specific roles to regulate particular promoters or play distinct roles depending on the cell type. For example, H1.2 has been reported to act as a coactivator by bridging between RNA polymerase II and components of the elongation apparatus in 293T cells, and to act as a corepressor by establishing a positive feedback loop with EZH2-mediated H3K27me3 deposition in MCF7 cells (30,31). Other H1 variants have been involved in chromatin compaction or gene silencing, such as H1.4 (21,22,32,33). In general, silent genes contain histone H1 at the promoter, and a ‘H1 valley’ appears upstream of transcription start site (TSS) upon gene activation (25,34,35). Altering the expression of H1 variants has been proven useful to study the contribution of individual variants to nuclear processes and to investigate the effects of global H1 decrease. In mice, single or double H1 variant knock-outs have no apparent phenotype due to compensatory up-regulation of other subtypes (36). These reports have favored the view that H1 variants are redundant. Knocking-out additional subtypes cannot be fully compensated by up-regulation of the remaining subtypes, resulting in embryonic lethality and demonstrating that the total amount of H1 is crucial for proper embryonic development (37). Despite embryonic lethality, triple knock-out H1.2–H1.3–H1.4 (TKO) mouse embryonic stem (ES) cells were obtained with a 50% global reduction in total H1 (27). Among the observed effects of TKO, a subset of imprinted genes are up-regulated that correspond to specific CpG regions becoming demethylated. Yang et al. demonstrated that some H1 subtypes specifically interact with the DNA methyltransferases DNMT1 and DNMT3B to promote methylation of imprinted genes (38). Major satellite repeats are also de-repressed in TKO (39). More recently, changes in the epigenetic signature of thousands of potential regulatory sites across the genome of TKO ES cells have been described, clustered to gene-dense topologically associating domains (TADs) (40). We have previously reported that depletion of single H1 subtypes by inducible RNA interference in cancer cells

produced a range of phenotypic effects, suggesting nonredundant functions for some of the histone H1 variants in somatic cells (26). Here, we further investigate consequences of depleting H1 variants by simultaneous depletion of several variants in breast cancer cells. We generate two new short-hairpin-RNAs (shRNAs), one for the specific depletion of H1.4, and the other affecting the expression of several H1 genes (multiH1sh), but mainly H1.2 and H1.4 at the protein level. While H1.2 and H1.4 single knock-downs (KDs) decrease proliferation similarly, multiple H1 KD has drastic consequences on cell growth and induces a strong transcriptional interferon (IFN) response. We observed the activation of IFN signaling transducers, participation of cytosolic nucleic acids receptors, IFN synthesis and up-regulation of IFN-stimulated genes (ISGs). In parallel, satellites and endogenous retroviruses are also upregulated and cytosolic RNA is increased. This is the first report of multiple H1 depletion in human cancer cells, and we show the importance of histone H1 to maintain heterochromatin integrity and to avoid a growth-inhibiting IFN response. We also show that pancreatic carcinomas characterized by a constitutively induced IFN response express low levels of several H1 variants. MATERIALS AND METHODS Cell lines, culturing conditions and treatments Breast cancer T47D-MTVL cells (carrying one stably integrated copy of luciferase reporter gene driven by the MMTV promoter) or derivative cells, were grown at 37◦ C with 5% CO2 in RPMI 1640 medium, supplemented with 10% FBS, 2 mM L-glutamine, 100 U/ml penicillin, and 100 ␮g/ml streptomycin, as described previously (26). HeLa cell line was grown at 37◦ C with 5% CO2 in DMEM GlutaMax medium containing 10% FBS and 1% penicillin/streptomycin. MCF7 cell line was grown at 37◦ C with 5% CO2 in MEM medium containing 10% FBS, 1% penicillin/streptomycin, 1% non-essential amino acids, 1% sodium pyruvate and 1% glutamine. Doxycycline (Sigma) was added at 2.5 ␮g/ml. IFNß (Sigma), Ruxolitinib (Selleckchem), BX795 (Selleckchem) and 2-aminopurine (Sigma) were added at indicated concentrations. Drug-inducible and constitutive RNA interference Inducible H1 knock-down cell lines were established from T47D-MTVL cells as described previously (26). Plasmids for the lentivirus vector-mediated drug-inducible RNA interference system (pLVTHM, ptTR-KRAB-Red, pCMCR8.91 and pVSVG) were provided by Didier Trono (University of Geneva) (41). The 71-mer oligonucleotides for shRNA cloning into Mlu/ClaI-digested pLVTHM were designed, annealed and phosphorylated as recommended by Didier Trono (http://tronolab.epfl.ch/). Target sequences are GTCCGAGCTCATTACTAAA for H1–4sh and GA ACAACAGCCGCATCAAG for multiH1sh. For the production of viral particles containing the lentiviral vector and infections, see Sancho et al. (26). The inducible knocked-down cell lines were sorted in a FACSCalibur machine (Becton Dickinson) for RedFP-positive and GFPpositive fluorescence after 3 days of doxycycline (Dox) treat-


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ment. For the constitutive depletion of TLR3, STING, MAVS, MDA5 and IFNAR, shRNA-expressing pLKO.1 vectors from the MISSION library (Sigma-Aldrich) were used. Viral particles production and infections were performed as described (26). Cells infected with the shRNAexpressing lentivirus were selected with 2 mg/ml puromycin (Sigma-Aldrich) 24 h after infection. Stable expression of HA-tagged H1 variants Generation of T47D derivative cells stably expressing HAtagged H1 variants using the lentiviral expression vector pEV833.GFP provided by Eric Verdin (Gladstone Institutes) was achieved as described previously (26). RNA extraction and reverse transcriptase (RT)-qPCR Total RNA was extracted using the High Pure RNA Isolation Kit (Roche). Then, cDNA was generated from 100 ng of RNA using the Superscript First Strand Synthesis System (Invitrogen). Gene products were analyzed by qPCR, 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. Specific qPCR oligonucleotide sequences are available as Supplementary Table S1. Histone and total protein extraction, gel electrophoresis and immunoblotting Histone H1 was purified by lysis with 5% perchloric acid for 1 h at 4◦ C. Soluble acid proteins were precipitated with 30% trichloroacetic acid overnight at 4◦ C, washed twice with 0.5 ml of acetone and reconstituted in water. For isolation of total histones, cell pellets were resuspended in 1 ml of hypotonic solution [10 mM Tris–HCl (pH 8.0), 1 mM KCl, 1.5 mM MgCl2 , 1 mM PMSF, 1 mM DTT] and incubated on ice for 30 min. The nuclei were pelleted at 10 000 × g for 10 min at 4◦ C. Sulfuric acid (0.2 M) was added to the pellet to extract the histones on ice for 30 min. The solution was centrifuged at 16 000 × g for 10 min at 4◦ C. TCA was added to the supernatant in order to precipitate histones and the precipitate was washed with acetone and finally resuspended in water. For isolation of total protein, cells were resuspended with Lysis Buffer [25 mM Tris-H–Cl (pH 7.5), 1% SDS, 1 mM EDTA (pH 8), 1 mM EGTA (pH 8), 20 mM B-glycerolphosphate] containing protease and phosphatase inhibitors, and boiled 20 min at 95◦ C. Cell lysates were obtained by centrifugation and protein concentration was determined by Micro BCA protein assay (Pierce). Purified histones or cell lysates were exposed to SDS-PAGE (12%), transferred to a PVDF membrane, blocked with Odyssey blocking buffer (LI-COR Biosciences) for 1 h, and incubated with primary antibodies overnight at 4◦ C and with secondary antibodies conjugated to fluorescence (IRDye 680 goat anti-rabbit IgG, Li-Cor) for 1 h 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 (26), are available from Abcam: H1.0 (ab11079),

H1.2 (ab17677), H1.3 (ab24174), H1-T146p (ab3596), H1.5 (ab24175) and antiH1X (ab31972). Other antibodies used were: H1 AE-4 (Millipore, 05-457), beta-tubulin (Sigma, no. T4026), H3K4me3 (Abcam, ab8580), H3K9me3 (Abcam, ab8898), H3K27me3 (Millipore, 07-449), H4K20me3 (Abcam, ab9053), HP1␣ (Active Motif, 39295). H1-T146p was used to identify H1.4 by immunoblot, as the immunogen was a synthetic peptide derived from within residues 100– 200 of human H1.4, phosphorylated at T146; this antibody could also recognize phospho T146 in H1.2, H1.3 (both 88% sequence identity with immunogen). To our knowledge this is the best available antibody to detect loss of H1.4 despite targeting a uniquely phosphorylated residue. To quantify H1 depletion with Coomassie staining, we load different amounts of total histones and choose the ones that showed the best linear range. After scanning with Image Gauge, individual histone H1 and H4 bands were quantified and H1 was expressed relative to H4, and relative to untreated cells. These were approximate quantifications due to the limitations of Coomassie staining of histone preps. Immunofluorescence One milliliter of hypotonic MAC buffer [50 mM glycerol, 5 mM KCl, 10 mM NaCl, 0.8 mM CaCl2 and 10 mM sucrose] was added to 0.5 million cells (resuspended in 1 ml RPMI 1640 medium) and incubated at room temperature for 5 min. 400 ␮l were spun down for 10 min at 500 rpm in a TermoShandon Cytospin 4 using a single-chamber Cytospin funnel as described in (42). Cells were air-dried for 1h at room tempearature. Then, fixed with 4% paraformaldehyde for 1h at room temperature. After three washes, they were permeabilized with 0.2% Triton X-100 for 15 min at room temperature and blocked with 3% bovine serum albumin for 1 h at room temperature. Then, the cells were incubated with primary antibody recognizing dsRNA (Scicons J2, English and Scientific Consulting Kft, Hungry) diluted in the blocking buffer, over-night at 4◦ C. After the pertinent washes, the secondary antibody Alexa Fluor 488 donkey anti-mouse (Life Technologies, A21202) was added for 1 h at room temperature in darkness. Nuclei were stained with DAPI, coverslips mounted using Mowiol, and samples were visualized by confocal laser scanning microscopy using a Leica TCS SP5 system. Cell proliferation analysis Cells of interest expressing an shRNA (GFP-positive) were mixed 1:1 with parental cells (GFP-negative) and cultured. Every three days, the cells were split and the percentage of cells that were GFP-positive was measured by FACS. MNase digestion Cells treated or not with Dox for six days were collected, resuspended and incubated for 10 min at 4◦ C in Buffer A [10 mM Tris–HCl pH7.4, 10 mM NaCl, 3 mM MgCl2 , 0.3 M Sacarosa and 0.2 mM PMSF], after that 0.2% of NP40 was added and incubated 10 min more at 4◦ C. After centrifugation, pellets were resuspended in Buffer A + 10 mM CaCl2 and 0.4 u of MNase per 5 millions of nuclei and incubated


4 Nucleic Acids Research, 2017 at 37◦ C. At different incubation times, MNase reaction was stopped by adding 10 mM EDTA pH 8. DNA was purified using EZNA Tissue DNA columns (OMEGA VWR) and run on a 2% agarose gel. Chromatin immunoprecipitation (ChIP) Immunoprecipitation of chromatin was performed according to the Upstate (Millipore) standard protocol. Briefly, cells were fixed using 1% formaldehyde for 10 min at 37◦ C, harvested and sonicated to generate chromatin fragments of 200–500 bp. Then, 30 ␮g of sheared chromatin was immunoprecipitated overnight with 2 ␮g of antibody. Immunocomplexes were recovered using 20 ␮l of protein A magnetic beads, washed and eluted. Cross-linking was reversed at 65◦ C 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 available on request. mRNA library preparation and sequencing (RNAseq) The library from total RNA was prepared using the TruSeq® Stranded Total Sample Preparation kit (Illumina Inc.) according to manufacturer’s protocol. Briefly, rRNA was depleted from 0.5 ug of total RNA using the Ribo-Zero Gold Kit followed by fragmentation by divalent cations at elevated temperature resulting into fragments of 80–450nt, with the major peak at 160nt. First strand cDNA synthesis by random hexamers and reverse transcriptase was followed by the second strand cDNA synthesis, performed in the presence of dUTP instead of dTTP. Blunt-ended double stranded cDNA was 3 adenylated and the 3 -‘T’ nucleotide at the Illumina indexed adapters was used for the adapter ligation. The ligation product was amplified with 15 cycles of PCR. Each library was sequenced using TruSeq SBS Kit v3-HS, in paired end mode with the read length 2 × 76 bp. We generated minimally 37 million paired end reads for each sample run in a fraction of a sequencing lane on HiSeq2000 (Illumina, Inc) following the manufacturer’s protocol. Images analysis, base calling and quality scoring of the run were processed using the manufacturer’s software Real Time Analysis (RTA 1.13.48) and followed by generation of FASTQ sequence files by CASAVA. RNAseq data analysis The mapping of the reads against the human reference genome (GRCh38) was done using the STAR program (version 2.5.1b) (43) and the quantification of genes was done with the RSEM program (version 1.2.28) (44). Differential expression analysis was done with DESeq2 (version 1.10.1) (45). Heatmaps, correlation scatter plot and boxplots were done using in-house R scripts. The expression values are those generated by DESeq2 using variance stabilizaing transformations. Genes differentially expressed by IFN treatment were selected from Interferome database

(version 2.01) (46) applying a filter of 2 < fold-change < –2 and those differentially expressed in more than one experiment. Enrichment of transcription factor binding DNA motifs was done using Homer software (47). Gene ontology enrichment analysis was performed using the R package ‘goseq’ (version 1.18.0) (48) and reduced by semantic similarity using REViGO software (49). Pathway enrichment analysis and figures were generated using ReactomeFIPlugin (50) for Cytoscape (51). To perform the enrichment analysis of repetitive sequences the RNA-seq reads were mapped to the Repbase database (version 21.02) (52) using Bowtie aligner (default parameters) (53). We only considered those reads which both paired-end reads were aligned to the same repetitive sequence. To test for significance a Fisher exact test was performed. Significantly enriched repetitive sequence were those with a adjusted Pvalue < 0,05 and not enriched in RDsh samples. Assay for transposase-accessible chromatin with highthroughput sequencing (ATAC-seq) ATAC experiment was performed as described (54). 75 000 cells treated were harvested and treated with transposase Tn5 (Nextera DNA Library Preparation Kit, Illumina). DNA was purified using MinElute PCR Purification Kit (Qiagen). All samples were then amplified by PCR using NEBNextHigh-Fidelity 2× PCR Master Mix (New Englands Labs.) and primers containing a barcode to generate the libraries. DNA was again purified using MinElute PCR Purification Kit and samples were sequenced using Illumina HiSeq 2500 system. Paired-end reads were first trimmed to 30 bp to remove adapter sequences. Then, aligned to the reference genome (hg19) using bowtie software (53). Duplicated reads and those mapping to the mitochondrial chromosome were removed. Accessibility peaks were obtained using MACS2 peak caller software (55). Average accessibility profile around the TSS of selected genes and pie charts representing the annotation of peaks was performed using CEAS software (56). For the repeats accessibility analysis, annotated repeats were taken from the RepeatMasker track from the UCSC and the number of peaks overlapping them computed. For the analysis of peaks overlapping upor down-regulated genes at the coding regions or promoter, or repeats, a permutation test (104 permutations) was performed using the R package regioneR (57). Expression data accession numbers RNAseq data is available in the Gene Expression Omnibus (GEO) database under the accession number GSE83277. Microarray data accession numbers are GSE11294 and GSE12299. ATACseq data accession number is GSE100762. Human H1 variant nomenclature The correspondence of the nomenclature of the human H1 variants with their gene names is as follows: H1.0, H1F0; H1.1, HIST1H1A; H1.2, HIST1H1C; H1.3, HIST1H1D; H1.4, HIST1H1E; H1.5, HIST1H1B; and H1X, H1FX.


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RESULTS Simultaneous depletion of multiple H1 variants in breast cancer cells impairs cell growth drastically We have previously reported the effects of the inducible depletion of six somatic histone H1 variants on cell growth and gene expression in breast cancer cells (24,26). H1.2 depletion caused cell-cycle arrest in G1 phase, while H1.4 KD caused cell death in T47D cells. Each variant KD altered the expression of small distinct subsets of genes. We have designed and assayed new shRNAs targeting H1. One of such shRNAs (sh120; H1.4sh) was specific for H1.4 (Figure 1A and B), while another (sh225; multiH1 or mH1sh from now on) reduced the expression of several H1 transcripts, although only H1.2 and H1.4 proteins were seen depleted consistently when tested by immunoblotting after 6 days of Doxycycline induced shRNA expression (Figure 1A and C, and Supplementary Figure S1). H1.3 was also seen partially depleted in some experiments. The two new shRNAs caused H1.0 up-regulation, both at the mRNA and protein level. This was previously observed for H1.2 KD, but not for the other H1 variants (26). Coomassie staining of histones extracted from multiH1 KD cells showed that the total content of H1 was reduced to ca. 70% despite of H1.0 up-regulation (Figure 1C and D). The intensity of protein bands containing H1.2 and H1.4 were decreased. In addition, multiH1sh expression caused a significant reduction in global nucleosome spacing as assessed by Micrococcal nuclease digestion of bulk chromatin (Figure 1E and Supplementary Figure S2). This was almost undetectable upon H1.4 KD, but was previously described for H1.2 depletion in breast cancer cells (26). Expression of both shRNAs impaired cell proliferation (Figure 1F). The effect of inhibiting simultaneously H1.2 and H1.4 in multiH1 KD cells on the growth rate was additive compared to single H1 depletion. H1 depletion increased the number of cells in G1 phase of the cell cycle and reduced cells in S phase, with the effects of multiH1sh being more pronounced than H1.4sh (data not shown). Transcriptome changes upon histone H1 variants knockdown We studied the consequences on global gene expression of multiple H1 KD and H1.4-specific depletion compared to the random shRNA using RNAseq. Data confirmed the efficacy of H1.4 and multiH1 shRNAs to inhibit the corresponding H1 genes without significant compensations other than some H1.0 up-regulation (Figure 2A). Results were compared with microarray data obtained before with the KD of the other variants (24,26). As previously described, depletion of individual H1 variants altered the expression of small subsets of genes, considerably specific for each variant, and containing both up- and down-regulated genes in different proportions. The inhibition of multiple H1s affected a larger number of genes, some already observed in single variant KDs (Figure 2B and Supplementary Table S2). In particular, 38% of genes affected in H1.4sh were also affected in multiH1sh (which also targets H1.4), while only 20–28% of the genes affected in the other single variant KDs

(H1.0, H1.2, H1.3 or H1.5) were significantly affected in the multiH1 KD. Among genes up-regulated in the multiH1 KD, there were many interferon-stimulated genes (ISGs) (Figure 2C). Comparison of our data with the Interferome database containing microarray data of IFN-treated cells, showed that 36% of the up-regulated and 18% of the down-regulated genes in the multiH1 KD were known to also respond to short- or long-term INF treatments. This is in comparison to the 6% of the total transcriptome known to be susceptible to IFN treatment (Figure 2C). Almost all the multiH1 KD up-regulated genes with the highest fold-change are ISGs. Moreover, fold-change gene induction upon multiH1 depletion was proportional (positive correlation) to the fold-change induction in response to IFN in multiple experiments deposited in the Interferome database (Figure 2D). Motif analysis in the promoters of multiH1sh deregulated genes found significant enrichment of interferon-stimulated response elements (ISRE), binding sites for interferonresponsive transcription factors (IRFs), but not in gammaactivated sites (GAS) (Figure 2E). ISRE are found in type I IFN-responsive genes, while GAS elements are present in type II IFN-stimulated genes. Down-regulated genes contained motifs for E2F, Sp1 and NFY transcription factors (q-value