Female mammals achieve dosage compensation by inactivating one of their two X chromosomes during development, a process entirely dependent on Xist, an X-linked long non-coding RNA (lncRNA). At the onset of X chromosome inactivation (XCI), Xist is up-regulated and spreads along the future inactive X chromosome. Contextually, it recruits repressive histone and DNA modifiers that transcriptionally silence the X chromosome. Xist regulation is tightly coupled to differentiation and its expression is under the control of both pluripotency and epigenetic factors. Recent evidence has suggested that chromatin remodelers accumulate at the X Inactivation Center (XIC) and here we demonstrate a new role for Chd8 in Xist regulation in differentiating ES cells, linked to its control and prevention of spurious transcription factor interactions occurring within Xist regulatory regions. Our findings have a broader relevance, in the context of complex, developmentally-regulated gene expression.
In mammals, sex is determined by the presence, number and combination of X and Y chromosomes1,2,3,4. Female mammals silence one of their two X chromosomes early in development in order to ensure X-linked gene dosage between females and males1,2. In mouse, an imprinted form of this process, where it is always the paternal X that is inactivated, is initiated in early development at the 2–4-cell stage (imprinted XCI). This form of XCI is then reversed during embryonic development at the blastocyst stage in the cells of the inner cell mass (ICM), where both X-chromosomes become active and subsequently have equal chances to be inactivated during the onset of gastrulation5. Xist lncRNA is the master regulator of XCI6. Its mono-allelic upregulation and in cis-spreading of Xist RNA triggers the initiation of the inactivation process by recruitment, directly or indirectly, of repressive chromatin and of DNA modifiers to silence gene-transcription6,7,8,9,10,11,12,13.
Differentiating female ES cells, carrying two X chromosomes, are a widely used model for the study of X chromosome inactivation as they closely recapitulate the early phases of random XCI4,14,15,16 in the embryo, and have been instrumental in allowing the dissection of the molecular pathways involved in Xist regulation. Previous work from the Avner laboratory has demonstrated that transcription of Xist and the Tsix lncRNA, its major antagonist, is precisely controlled by pluripotency factors17,18,19. Other laboratories have investigated the role of nearby genes encoding lncRNAs such as Jpx and Ftx and protein factors such as Rnf12 (also known as Rlim) and YY1/CTCF in the allelic regulation of Xist and Tsix20,21,22,23,24,25,26,27. The current model comprises a complex interaction network that is in a dynamic transcriptional equilibrium in undifferentiated ES cells28. In differentiating cells, Xist is strongly mono-allelically up-regulated, spreads onto the chromosome from which it is transcribed and silences genes in cis, including Tsix (reviewed in Van Bemmel et al. 29). It is known that the Xist promoter in undifferentiated ES cells is bivalent30, marked by both repressive and active histone modifications, and poised for activation. This state is resolved during differentiation at the up-regulated allele of Xist, which becomes actively transcribed and marked by a strong domain of H3K4me321,23. In the context of these changes at the chromatin level in the vicinity of the Xist promoter, we hypothesise that chromatin remodelers may have a role in the onset of Xist regulation during cell differentiation.
ATP-dependent chromatin remodelers play essential roles in chromatin biology, compacting or relaxing the chromatin threads and, in turn, restricting or facilitating access of transcription factors (TFs), RNA polymerase II (Pol II) and the basic transcriptional machinery to the underlying DNA sequences31. In mammals there are about 30 different types of ATP-dependent chromatin remodelers, belonging to four major remodeler families32: SWI/SNF, ISWI, NURD-CHD, and INO80. Such complexes are implicated in most cell activities, from regulation of differentiation, to pluripotency and gene-specific activities. Chromatin remodelers can form hundreds of distinct functional entities by complexing amongst themselves in the cell, although the true extent of this phenomenon and the issue of tissue specificity, remain for the most part unexplored. There are however indications that these complexes have relatively little or no functional redundancy as single-gene Knock-Outs (KO) of chromatin remodelers generally produce observable and often very severe phenotypes33. For example, the BAF250 family, belonging to the SWI/SNF complex, is composed of three members (BAF250A–B–C). BAF250-A null embryos die at ~E6.5 and the defect is not compensated for by either BAF250B or BAF250C34. Similarly, Chd2 mutants die before birth and other Chd-family members do not appear to compensate for such mutations35.
In order to focus our analysis on chromatin remodelers likely involved in the regulation of the initiation of XCI, we screened for candidate regulators, using the following criteria: (i) accumulation and binding to the X inactivation centre (XIC)28,36; (ii) robust expression during early mouse developmental stages37,38; (iii) a KO phenotype, where known, compatible with a role in XCI (MGI, www.informatics.jax.org/) or phenocopying XCI-defects. Based on these selection criteria and preliminary analysis, we decided to focus our immediate attention on the Chromodomain-Helicase-DNA-binding protein 8 (Chd8), which belongs to the CHD family. Chd8 contains two chromodomains, a SWI/SNF2-like ATP-dependent helicase domain, several SANT/CR domains and two terminal BRK domains31,39. Chd8 KO mice die in utero around E7.5–8 but appear to stop growing and start degenerating by E5.5, although sex-specific differences have not been reported40. A recent publication, which studied the role and the distribution of chromatin remodelers in ESCs, noted that different types of chromatin remodelers have specific histone substrate preferences, determining their genomic accumulation36. Chd8 seems to accumulate in open-chromatin regions marked by H3K4me3 and DNA hyper-sensitivity sites36. Chd8 has also been previously shown to regulate lncRNAs41 and autism-associated genes42. It is a key factor in correct neuronal differentiation, mutations which lead to autism spectrum disorders (ADS) phenotypes41,43,44,45. In our study, using undifferentiated and differentiating female ES cells and Chd8 knock-down (KD) and KO systems, we demonstrate that Chd8 has a critical role in Xist regulation. Chd8 regulates strong yet controlled bursts of Xist expression, which are necessary for its correct spreading across the chromatin and initiation of XCI by preventing the spurious binding of TFs at the Xist promoter.
Validation of experimental cell lines and conditions of differentiation
For our experiments we decided to use a previously developed and validated first-generation (F1) 129/castaneus hybrid ES cell line carrying the insertion of a premature termination site in the Tsix gene (Fa2L)46. As a consequence of this mutation, when these cells are differentiated, the chromosome carrying the mutation (the 129 allele) is forced to become the inactive X chromosome (Xi) in ~95% of cells46. Furthermore, the extensive polymorphism between the 129 and the castaneus genomes facilitate allelic analysis. Single nucleotide polymorphisms (SNPs) between the strains are expected every 80–100 bp, on average47.
As female XX ESCs are known to be prone to lose one of their two X chromosomes, after extended passaging, we subcloned this line (Fa2L sub-clone 4; Fa2L-S4 hereafter) and verified the presence of two X chromosomes by DNA FISH (Supplementary Fig. 1c). In order to assess the efficiency of X chromosome inactivation initiation, we differentiated these cells from 2i conditions (undifferentiated state) to the neuronal progenitor cell state (NPCs) for 3 days (see the “Methods” section and below). We monitored the efficiency of XCI and differentiation by means of H3K27me3 Immunofluorescence (IF) staining, as a surrogate marker for the inactive X chromosome (Xi)14,48 and qRT-PCR analysis of selected differentiation (Nestin/Rex1) and XCI (Xist/Tsix) markers17,19 (see also the “Methods” section).
We show that after 3 days after differentiation induction, Xist and the NPC-specific marker Nestin are up-regulated and the pluripotency marker Rex1 and Tsix (XCI marker) are correctly down-regulated. About 50% of the cells showed the presence of an inactive X chromosome by day 3 (H3K27me3 domains, Supplementary Fig. 1a, b). These data indicate that XCI has been robustly initiated by day 3 of differentiation, which was retained as the principal time point for subsequent analysis of Xist regulation during XCI initiation phase (see also Supplementary Fig. 4 and see the “Methods” section).
Chd8 accumulates at the Xist promoter in undifferentiated and differentiated cells
In order to determine the genomic distribution of CHD8 in our cell lines, we performed ChIP-seq experiments for CHD8 and H3K4me3, an active chromatin mark recognized by CHD8 chromo domains30. As expected, ~90% of CHD8 peaks overlap with H3K4me3 sites in undifferentiated conditions, and >95% in differentiating conditions (Fig. 1a). Genome-wide analysis of CHD8 peaks reveals that this protein localizes at transcriptional start sites (TSS), at promoters and at intergenic regions as previously reported49 (Fig. 1b, c and Supplementary Fig. 2). We went on to test whether Chd8 and H3K4me3 are enriched at the X inactivation center (XIC) with a particular focus on the Xist or Tsix promoters34. We note—as expected—that during differentiation there is a shift in H3K4me3 accumulation from the Tsix regulatory region (major promoter and DXPas34 tandem repeats) to the Xist promoter (Fig. 1d)50. We show that in ES cells (undifferentiated conditions) there is a CHD8 peak at the Xist promoter, whilst a slightly larger peak characterising this region under differentiating conditions (Fig. 1d). We reasoned that the CHD8 peaks might contribute to Xist regulation both in undifferentiated conditions and at the onset of XCI, potentially opening up the chromatin at the Xist promoter (see below).
Exploiting the SNP differences between 129 and castaneus, we performed allele-specific analysis of the H3K4me3 chromatin mark first, due to its abundance on chromatin, allowing for proper statistical analysis. We assigned ChIP-seq reads to the 129, castaneus (Cast) or reference genome (BL6) (see the “Methods” section). As expected, during differentiation, most of the H3K4me3 is associated with the 129 allele of the Xist promoter (the future Xi) after immunoprecipitation but not in the pre-IP materials (input) (Supplementary Fig. 3a, b)46. CHD8 also appears to be mostly associated with the 129 allele, although due to the known limited number of reads from Chd8 ChIP-seq experiments51,52, we do not have the power to detect statistical differences between the alleles (Supplementary Fig. 3a, b). It is possible that this robust gain of H3K4me3, upon differentiation, is necessary for recruiting Chd8 and/or other chromatin remodeler complexes to the Xist promoter through its chromodomains.
Chd8 KD affects Xist up-regulation but it does not majorly affects XCI initiation
To analyse the functional role of Chd8 in XCI initiation, we performed siRNA-mediated KD experiments in the parental cell line (Fa2L-S4). In order to choose the best time-point for siRNA delivery, we checked the dynamics of Chd8 expression relative to that of Xist during differentiation. During differentiation, Chd8 mRNA levels peak at day one and then decrease, concomitantly with an increase of Xist RNA (Supplementary Fig. 4a). Increases of Chd8 at the protein level were detected slightly later, at day 2/3 of differentiation (Supplementary Fig. 4b, c). In view of the expression kinetics of these genes, we decided to KD Chd8 at day 1 of differentiation, upon exit of the pluripotent state, when Chd8 mRNA is at its maximum level, by means of siRNA (for 48 h). Using a pool of 4 siRNAs we efficiently knocked down Chd8 at RNA level by ~50–60% at day 3 of differentiation (Fig. 2a). The progression of XCI and differentiation was analyzed by means of qRT-PCR and IF (see also “Methods” section) (Fig. 2 and see the “Methods” section). We noted that this partial Chd8 KD led to significantly reduced Xist expression (~30%) during differentiation (Fig. 2a), while Tsix down-regulation, cell differentiation markers (Rex1, Sox2, Nestin) and Xist-mediated gene-silencing processes were unaffected (Fig. 2b, c). We assessed whether the reduced Xist expression under Chd8 KD conditions during differentiation is associated with defective XCI initiation. XCI initiation was followed in differentiated control and Chd8 KD cells by means of H3K27me3 IF. Initiation of XCI was slightly reduced at day 3 of differentiation in KD vs. WT cells (Supplementary Fig. 5). As the H3K27me3 domains in Chd8 KD are indistinguishable from the parental line in size (Supplementary Fig. 5), this observation suggests that potentially fewer cells may initiate XCI in the Chd8 KD-treated cells. RNA-seq data, shows in Chd8 siRNA vs. Control siRNA (Diff condition), 11 differentially expressed genes (DEG, false discovery rate (FDR) ≤ 0.05, and absolute log2FC ≥ 1—Supplementary Data 1), of which 6 were upregulated and 5 downregulated.
This relatively minor variation in gene expression suggests that the differentiation process is not, per se, affected by partial Chd8 KD treatment (Supplementary Fig. 6a). Principal component analysis (PCA) reveals that the overall expression patterns of the Chd8 KD cell line, during differentiation, is very similar to that of the control cell line (Supplementary Fig. 6a). RNA-seq analysis confirmed Chd8 down-regulation (≥50%) was accompanied by only a small and statistically non-significant reduction in Xist expression (~17%). Differences in sensitivity and substrate preference between the qRT-PCR and the RNA-seq data likely explain such low-level discrepancies. Indeed, while qRT-PCR can capture both poly-Adenylated (PolyA+) and non-poly-Adenylated RNAs (PolyA−), our RNA-seq experiments were specifically designed to capture exclusively the cellular PolyA+ RNA pool (see the “Methods” section). Considering that a large proportion of Xist RNA is not polyadenylated53,54, these results are in line with our expectations. For these reasons, from this point, we will only refer to qRT-PCR analysis for Xist and Tsix paired quantification.
In order to assess the extent of Xist dependency on the levels of CHD8, we also generated two Chd8 KD stable cell lines by lentiviral transduction showing respectively mild and severe Chd8 KD (Chd8.1KD, Chd8.2KD, Chd8.1/Chd8.2 hereafter in the text) (Supplementary Fig. 7a, b, see also the “Methods”). In cell differentiation experiments, we noted that the mild Chd8 KD (Chd8.1), in agreement with the siRNA-mediated Chd8 KD, is associated with a reduced upregulation of Xist upon differentiation by qRT-PCR, although in this case, this variation was not statistically significant (p = 0.082). The discrepancies between the siRNA mediated and light shRNA Chd8 KD line (Chd8.1), might be due to differences in level of KD achieved in the siRNA vs the shRNA experiment (50–60% vs. ~40% RNA KD at the RNA level, respectively); or to the experimental set up (such as stable vs. transient Chd8 KD, see the “Methods” section). In the Chd8.1 cell line, qRT-PCR analysis, showed cell differentiation was not affected in this cell line at 3 days of differentiation in this cell line, using established markers (Supplementary Fig. 7c). We went on to test the role of mild CHD8 depletion during the XCI initiation phase by means of H3K27me3 IF in these stable KD cell lines. In the Chd8.1 cell line, we showed a modest reduction in the number of cells that initiate XCI (Supplementary Fig. 7a), similar to that seen for the siRNA-mediated KD (Supplementary Figs. 2 and 5). In the severe CHD8 KD (Chd8.2 ≥ 80% Chd8 KD), we show a modest but significant increase in Xist expression and a concomitant increase of Tsix repression, at the onset of XCI. Overall cell differentiation was not significantly affected, and no change in the number of cells initiating XCI at 3 days of differentiation was observed using H3K27me3 as marker of the Xi (Supplementary Fig. 7a, c).
RNA-seq analysis of these Chd8 KD lines (Supplementary Fig. 8, Supplementary Data 2), showed no major defects in cell differentiation and only a minor number of genes significantly deregulated for the Chd8.1 line (35 significantly deregulated genes, 8 of which upregulated and 27 downregulated; FDR ≤ 0.05, log2FC ≥ 1). RNA-seq analysis for the Chd8.2 line revealed more significantly deregulated genes, as expected from the higher level of CHD8 KD efficiency. We report 213 DEG in Chd8.2, roughly equally divided in upregulated and downregulated genes (108 downregulated and 105 upregulated; FDR ≤ 0.05, absolute log2FC ≥ 1).
These unexpected differences between light (siRNA/Chd8.1) and severe Chd8 KD (Chd8.2) (Supplementary Figs. 5 and 7) suggests the existence of a threshold effect in Chd8-dependent Xist regulation during XCI (see also Discussion).
Chd8 KO affects Xist regulation and XCI initiation
In our KD experiments, we achieved up to ~80% Chd8 RNA/protein KD efficiency, which would likely still leave a reasonable amount of protein available in the cell. In order to ablate the protein completely, we used CRISPR/Cas9-mediated gene editing. We designed guides targeting the chromo-domains and helicase-domains of Chd8, with the aim of inducing a frame-shift mutation upon CRISPR/Cas9-mediated non-homologous end-joining (NHEJ) repair (Fig. 3a, and see the “Methods” section). We obtained several clones carrying mutations in either or both domains. We decided to proceed with two sub-clones (C4.1 and C4.3; KO.1 and KO.2 hereafter; see the “Methods” section) carrying a 20 bp deletion/1 bp insertion in the helicase domain and 5 bp deletion in chromo domain (see the “Methods” section). Both mutations are predicted to generate frame-shift mutations. The absence of Chd8 protein was verified by sequencing and Western Blot (WB) analysis, respectively (see the “Methods” section) (Fig. 3b).
Expression of Xist in differentiated cells by qRT-PCR—normalised over the undifferentiated state — was revealed to be increased rather than decreased, mimicking, in part, the results seen with the severe Chd8 KD (Chd8.2). XCI initiation was revealed to be enhanced by up to ~1.5/2 fold by H3K27me3 IF analysis (Fig. 3e). The differentiation process per se using established markers, does not appear to be critically affected (Fig. 3c). We also report that undifferentiated C4.1 cells show a significant reduction in Xist basal expression (Fig. 3d). Equivalent results were found in the other Chd8 KO clone (KO.2).
RNA-seq analysis of the Chd8 KO lines over the parental line (Fa2L-S4), shows more widespread gene deregulation compared with the Chd8 si/shRNA KD (Supplementary Fig. 12). In particular, we report 2485 DEG genes, 857 of which are upregulated and 1628 downregulated (FDR ≤ 0.05, absolute log2FC ≥ 1, Supplementary Data 3). Noticeable, 160/213 (75.1%) deregulated genes from the severe Chd8 KD (Chd8.2) overlaps with the Chd8 KO-deregulated genes. The differentiation process, per se does not appear to be markedly compromised (see the “Discussion” section).
Chromatin accessibility at the XIC
We decided to assess whether Chd8 KO could affect local chromatin organization of the XIC, with a particular focus on the Xist and Tsix regulatory regions. To this end we performed ATAC-seq analysis on undifferentiated vs. differentiated Chd8 KO cells in clone C4.1 (KO.1) and C4.3 (KO.2) (Fig. 4). Whilst under undifferentiated conditions we saw no changes in chromatin accessibility at the Tsix major regulatory regions (e.g. Ex2/3 and DXPas34 microsatellite) nor at the Jpx promoter, we observed an increased accessibility at Tsix intron one and a small reduction in chromatin accessibility at the Xist promoter. Under differentiating conditions, we noted a marked increase of chromatin accessibility at the Xist promoter (Fig. 4). These results are in good agreement with the qRT-PCR and H3K27me3 IF data shown in Fig. 3.
In order to exclude potential cell-line specific artefacts, we decided to transfect the KO cell line (C4.1) with a plasmid expressing the full-length (FL) CHD8 protein. To this end, we generated, by bacterial recombineering, a FL Chd8 protein under the control of a constitutive CAG promoter (Fig. 5 and see the “Methods” section). Reconstituted CHD8 protein levels appear to be comparable with the parental cell line by WB analysis, a necessary condition for our analysis (Fig. 5). In brief, complemented cells were differentiated and XCI progression was assessed by H3K27me3 IF. As expected the FL form of CHD8 effectively rescued the defective KO phenotype, in the context of XCI initiation (see the “Methods” section and Fig. 5b, c). This indicates our results are unlikely to be majorly influenced by line-specific factors (see the “Discussion” and the “Methods” section), but rather linked to Chd8-specific regulation.
Candidate and unbiased TFs binding analysis at the Xist promoter and Chd8 mass-spectrometry
Taking into consideration the results obtained in Chd8 KD and KO conditions, we hypothesised a competitive binding at the Xist promoter between TFs and CHD8. In brief, one might expect that some factors would increase (or decrease) their binding at the Xist promoter, in the absence of CHD8, and contribute to the observed transcriptional deregulation that we observed, in Chd8 KO lines.
In order to determine whether in the absence of CHD8, another transcription factor binds at the Xist promoter in a non-physiological fashion (i.e. increase or decrease), we decided to use two complementary approaches. First, using known databases of ChIP-seq (https://chip-atlas.org/, see also the “Methods” section), we screened for all factors binding at the Xist promoter and expressed in ES and differentiating/differentiated conditions. On this basis, we selected YY1 and INO80 for further analysis, and excluded Nanog and Rex1 as they are not expressed in differentiated cells, are severely reduced in differentiating cells, and are known Xist repressors29. We also excluded from our analysis CTCF as its reduction during differentiation does not affect proper XCI initiation55. We also used a motif analysis approach (MEME, see the “Methods” section) in order to reveal, in an unbiased way, potential TFs competing with CHD8 at the Xist promoter. To this end, we extracted all the sequences underlying our CHD8 peaks (ChIP-seq analysis) and used these as bait to find potential consensus motifs shared with other TFs. The analysis revealed the enrichment of several known TF-binding motifs, including Egr1, SP1, SDPEF, and YY1 (Supplementary Fig. 9a), In particular, YY1 attracted our attention for its known role as a major Xist activator56, and we decided to further test this candidate (see below).
Finally, in order to have an unbiased candidate approach as well, we performed CHD8-IP followed by mass spectrometry analysis. We decided to do both native protein IP using two antibodies toward CHD8 and a tag-mediated (HA-tag) IP in order to recover only the strongest candidate. To this end, we decided to generate a Chd8 KI cell line carrying a C-term tag (3Flag-HA-TAG; 3FHT in short) (Fig. S10) to be used for the HA-mediated IP (for details about the generation and validation of this cell line, please refer to the “Methods” section). Interestingly, CHD8-mediated and HA-mediated IPs followed by mass spectrometry, revealed WDR5, a known CHD8-interacting protein both in mouse and human (STRING, see the “Methods” section) as main CHD8 partner (Supplementary Data 4 and Supplementary Fig. 9c). As WDR5 is a known component of the MLL complex and a strong activator of gene transcription and a Polycomb-repression antagonist57, we decided to include it in our analysis.
We tested these three factors by ChIP-seq or ChIP/Cut&Run qPCR analysis. In brief, by ChIP-seq analysis, we could not detect any change in INO80 occupancy at the Xist promoter and neighbouring regions, in the absence of CHD8. Hence, we excluded this factor as a potential candidate competing for the Xist promoter (Supplementary Fig. 11). Similarly, using Cut&Run qPCR analysis, we could not detect any enrichment of WDR5 in differentiating conditions, either in the parental WT line nor for the Chd8 KO line, at the Xist promoter. This suggests that it is unlikely that WDR5 is playing a major role in Xist upregulation during XCI initiation (Supplementary Fig. 11b). On the other hand, when we performed Cut&Run analysis for YY1, and we found an increase of this transcription factor at the Xist promoter (peaks 3–4, Fig. 5d), overlapping with the CHD8 peaks. Because YY1 is a known (strong) activator of Xist56, we reasoned that the elevated binding of YY1 at the Xist promoter, is very likely to be responsible of abnormal activation of Xist in the absence of Chd8 protein (see also below). In order to test YY1’s role in Xist transcription in the absence of Chd8, we decided to perform YY1 KD during differentiation in CHD8-depleted cells (Chd8 KO.1). As YY1 KD is known to affect cell/neuronal differentiation58,59, we decided to perform mild siRNA-mediated YY1 KDs (i.e. 20–30% reduction), not to affect overall cell differentiation (Fig. 5e, Supplementary Fig. 11c). qRT-PCR analysis of YY1 KD lines over control, revealed a minor reduction of Xist RNA (Fig. 5e). This reduction is, however, linked to a significant reduction of cells initiating XCI as previously reported56, but not to the abolishment of XCI initiation (Fig. 5e). These observations indicate that other factors (i.e. redundancy) are likely involved in the upregulation of Xist, other than Chd8 and YY160,61.
Whilst many aspects of Xist regulation have been intensely studied over the years17,18,19,20,21,22,23,24,62, little is known about the role of chromatin remodelers in the context of XCI. Here we show that Chd8 is a likely critical factor involved in the regulation of Xist expression.
Our data suggest the involvement of Chd8 as an activator of Xist in undifferentiated ES cells (Figs. 3 and 4) and in the fine-tuning Xist expression during differentiation (Figs. 1–4). We show by Chd8 KD experiments (Fig. 2, Supplementary Fig. 7), that correct Xist transcriptional activation appears to depend on the presence of WT-levels of Chd8 protein, recruited specifically to the Xist promoter via recognition of an H3K4me3 mark, which is specifically enriched on the allele chosen for X inactivation at the very onset of differentiation (Fig. 1 and Supplementary Fig. 3). In differentiating cells, Chd8 subsequently fine-tunes the amplitude of Xist expression (Figs. 3–5), we hypothesise with CHD8 binding at the Xist promoter, restricting alternative binding of transcriptional co-activators to its regulatory regions, via a competitive binding exclusion. This implies that in the absence of CHD8, co-activators and other TFs could bind to the Xist promoter and lead to transcriptional up-regulation (Fig. 6). In this context, known regulators of Xist transcription, such as YY1, might bind to the Xist promoter with greater avidity or more robustly. YY1 has been previously shown to bind to XCI escapees, including the Xist promoter63 and to be a strong activator of Xist transcription, competing with REX1 for binding at the Xist promoter56. Our analysis suggests that YY1 is one of the most-likely activators of Xist transcription in the absence of Chd8 protein since we observed a marked increase of YY1 recruitment at the Xist promoter in the absence of CHD8 (Fig. 5d, Supplementary Fig. 9). Indeed, our analysis of YY1 KD in Chd8 KO cells confirmed YY1’s role in the initiation of XCI and Xist regulation (Fig. 5e). YY1 KD in Chd8 null background, also supports the possibility that other factors are involved in Xist regulation61 (i.e. the absence of key regulatory proteins can be compensated by other proteins). The comparison of publicly available YY1 ChIP-seq datasets64 with our Chd8 ChIP-seq data in embryonic stem cells revealed ~75% of overlap between YY1 and Chd8 peaks. Furthermore, of the 2484 DFG in the Chd8 knockout (differentiated), 1203 promoters are bound by Chd8 and of those 391 also by YY1 (see the “Methods” section). This further suggests competition or cooperative effects in the regulation of target genes.
Other candidates competing for the Xist promoter that were tested in this study (INO80 and WDR5), did not show any occupancy change at the Xist promoter in Chd8 KO vs. WT conditions. These factors, therefore, are unlikely to play an important role at Xist promoter, under the tested conditions. Our experiments using different Chd8 KD lines, also suggest that CHD8 levels have a threshold effect on Xist regulation. Mild Chd8 KD downregulates Xist expression, supporting its role as a transcriptional activator65. Conversely, strong CHD8 depletion (KO) has the opposite effect, most likely through competition with TFs56 as discussed above.
Whilst our data fully supports a role for Chd8 as a transcriptional activator of gene-expression as previously shown49,65, Chd8 might also act as a repressor of transcription66. One possibility is that Chd8 regulates Xist expression under differentiating conditions by recruiting the histone H1, to moderate the amplitude of Xist up-regulation66. While our experiments cannot fully discriminate between the activating and/or repressive activities of Chd8, our data, and in particular the ATAC-seq experiments strongly support an activating role for CHD8 rather than a repressive role in Xist regulation. A possible complication in the comparison of WT vs. KO cells is that the cell differentiation in Chd8 KO cells could have a different differentiation kinetic from that of the matched parental cell line, with more differentiated cells at day 3, in line with previous reports44,52,67. We did not find any convincing evidence of such neuronal differentiation enhancement in Chd8 KO by RNA-seq analysis49,52 (Supplementary Fig. 12a, b). Finally, RNA-seq analysis of all the XIC factors28 in Chd8 null background (vs. control), indicates that Rnf12, Ftx, Xist main activators, and Chic1 are significantly downregulated (Supplementary Data 3). This further suggests that Xist upregulation is unlikely to be linked to an indirect effect mediated by these factors.
In summary, our experiments support the hypothesis that Chd8 fine-tunes Xist expression amplitude levels and regulates occupancy at the Xist promoter. In the absence of CHD8 other protein(s) are allowed to bind (or not to bind) at the Xist promoter inducing concomitant deregulation (Fig. 6). Our results have a broader applicability for the interpretation of the phenotypes observed in KO animals or cells. For example, the KO of a gene known to repress a given cadre of target genes, might result in the opposite effect (i.e. upregulation) in the case other TFs, such as an activator, is now allowed to engage with these regulatory sequences. Finally, our study emphasises our relative ignorance of the nature and importance of the interactions occurring between XCI regulation and chromatin remodeler activity suggesting the interest of additional studies on other remodelers in Xist regulation.
We purchased the FL Chd8 cDNA from Genescript and cloned it into the MreI and NotI sites of pCAGp292-Puro. The MreI-NotI Chd8-containing fragment from this plasmid was cloned into the same sites of the plasmid, pCAG-mCherry-modified (this work), leaving the CAG promoter intact to create pCAG-Chd8-Neo. This plasmid was cut with PvuI, then blunted, cut with EcoRV and re-ligated to create pCAG-Chd8-Neo-AmpS. To aid in identifying the correct recombinant plasmids, bacterial strains that produce Cas9 and a sgRNA against the sequence in question were made by recombineering. Sequencing confirmed that surviving plasmids contained the precisely designed deletion. Plasmid maps and sequences are available upon request.
KI generation and sub-cloning
Chd8 KI cell lines were generated by standard DNA-recombineering. About 20 µg of NotI-linearized plasmid was electroporated into the Fa2L recipient cell line, an F1 129/castaneus hybrid ES cell line carrying the insertion of a premature termination site in the Tsix gene (Fa2L)46. Two days after transfection, cells were selected with G418 at a concentration of 250 µg/ml for ~7 days followed by Gancyclovir at 2 µM concentration for 2–4 days. After 10–11 days, resistant colonies were picked and expanded. Using homologous recombination, we obtained several 3′ KI cell lines and selected clone 3B for further analysis (we will refer to this clonal cell line as Chd8KI hereafter) (Supplementary Fig. 10). We showed by Southern Blot (SB) and long-range PCR (LR-PCR) analyses the correct integration of our construct (Supplementary Fig. 10a, b). We verified, in this newly generated cell line, the presence of randomly selected 129/castaneus SNPs by Sanger sequencing (Supplementary Fig. 10c), the presence of two X-chromosomes by DNA FISH (Supplementary Fig. 10d), and the expression of the tagged allele by WB analysis (Supplementary Fig. 10e).
Characterization of differentiation of the Chd8KI line
In order to assess the efficiency of X chromosome inactivation initiation we differentiated the Chd8KI clone from 2i conditions to the NPCs for 3 days (see below). We monitored the progression of XCI and differentiation by H3K27me3 IF staining, as a surrogate marker for the inactive X chromosome (Xi) and used qRT-PCR for analysis of differentiation (Nestin/Rex1) and XCI (Xist/Tsix) markers.
Xist and NPC-specific Nestin were found to be correctly up-regulated, and the pluripotency marker Rex1 and Tsix properly down-regulated during differentiation with some 50–60% of the cells showing an inactive X chromosome by day 3 of differentiation (Supplementary Fig. 10f, g). These data indicate that XCI was robustly initiated by day 3 of differentiation.
We also sub-cloned the parental cell line (Fa2L) in order to obtain a homogeneous line of cells carrying two X chromosomes (Fa2L sub-clone 4; Fa2L-S4). Sub-cloning was achieved by plating cells at very-low density (500 cells/plate) in 10 cm tissue culture dishes then waiting 7–9 days before picking individual colonies and re-plating these in 2i medium Single colonies were expanded and characterised using routine procedures. The Fa2L-S4 line was characterized as described above for the Chd8 KI line (see also main text for details).
Cell culture conditions
Cells were grown in 2i conditions as previously published68. For differentiation experiments, the wells of a six-well plate or a 10 cm TC dish were pre-coated with gelatin (0.1%) containing laminin (1:1000) solution and incubated at 37 °C for 15 min. 150,000 cells per well in six-well plate (or 0.8 million for 10 cm dish) were plated in each well in neuronal differentiation medium (NDM) as previously published69. The cells were differentiated for 3–4 days and harvested for ChIP-seq, qRT-PCR work or IF/RNA-FISH analysis.
KD of Chd8 and other chromatin remodelers
In order to obtain Chd8 KD, we used Dharmacon Smart pool-of-four siRNAs (#67772) or single siRNA (YY1, from Thermo Fisher, Cat #AM16708) at a 50 nM final concentration, in accordance with the manufacturers protocol. Chd8 KD was initiated at day 1 of differentiation using lipofectamine transfection (RNAi-MAX Invitrogen).
In order to generate stable Chd8 KD lines, we used lentiviral-mediated KD using pre-packaged lentiviral particles (SIGMA). In brief, Fa2L-S4 cell line was infected either with a scrambled shRNA control (SHC002) or two independent Chd8-targeting constructs (SIGMA shRNA clones—CloneIDs: NM_201637.2-847s1c1 (Chd8.1); NM_201637.2-3342s21c1 (Chd8.2)). Two days after transfection, puromycin selection was applied for 7–9 days (2 µg/ml). Colonies were picked and expanded using standard protocols. Chd8 KD was assessed by conventional qRT-PCR analysis (see below).
Generation of Chd8 KO cell lines
Chd8 KO cells were generated by CRISPR-Cas9 mutagenesis. Briefly ~5 µg Cas9 expressing vector carrying Puromycin resistance and guides to the Chd8 helicase and chromo domains were co-transfected in Fa2L-S4 cells using Lipofectamine 3000 (Invitrogen). Two days after transfection, puromycin selection was applied for 7–9 days (2 µg/ml). Colonies were picked and expanded using standard protocols. Clones surviving selection were screened by PCR for indels in either the chromo, the helicase or both domains. Amplicons were topocloned and analysed by DNA sequencing. We selected sub-clones C4.1/C4.3 for further analysis (see also Fig. 3 and the “Results” section).
Complemented cell lines
We transfected Chd8 KO clones by lipofection (Lipofectamine 3000, Invitrogen) in accordance with the manufacturer’s protocol. Briefly 10 µg of construct expressing the FL form of Chd8 were transfected. Two days after transfection, Neomycin (G418) selection was applied for 7–9 days (400 µg/ml). Colonies were picked and expanded using standard protocols. Complementation was tested by means of WB analysis and immuno-fluorescence (IF) in the context of XCI initiation (see below and main text for details).
RNA extraction and qRT-PCR analysis
RNA was extracted using the RNeasy mini kit according to the producer’s manual (Qiagen). RNA was DNAse treated for 30 min at 37 °C and DNAse was then inactivated using a cDNA kit (Turbo DNase, Ambion). Reverse-transcription was carried out using a kit from Thermo Scientific (First Strand cDNA Synthesis Kit, Cat. K1612) in accordance with the manufacturer’s instructions (1–2 µg of total RNA). qRT-PCR was performed using Kapa BioSystem/Biorad SYBR-green reagent using an Applied Biosystem real-time system, following standard protocols. A wide panel of primers pairs was used to test cell differentiation and XCI progression, in the initial phases of the project. Selected primers were then used for all experiments. A complete list of primers used in this paper is available in Supplementary Data 5. Primer sequences used in preliminary analysis (nor shown) are available on demand.
Protein extraction and WB analysis
Whole lysate cell extracts were obtained after incubation on ice for 30′ in RIPA buffer with protease inhibitors, followed by centrifugation. WB analysis was done using standard SDS–PAGE protocols. CHD8 antibody(ab) was purchased from Novus biological (NB100-60417), HA ab (H6908), and Vin ab (V9264-200UL) both bought from Sigma, GAPDH ab bought from Millipore (CB1001), and RNA pol II ab obtained from Active Motif (#39097).
IF analysis and RNA-FISH were carried out as previously described48. H3K27me3 mono and polyclonal antibodies were purchased from Active Motif (#39155, #39536, respectively), Chd8 antibodies were bought from Novus (NB100-60417).
RNA-seq analysis was carried out as previously described8. Reads were trimmed with cutadapt and aligned to the genome with STAR70. The FeatureCounts software counted the number of reads per gene overlapping any exon71. Genes are defined as in Ensembl’s genome release Mus Musculus GRCm38.88 (https://www.ensembl.org/Mus_musculus/Info/Index). DFG were identified using the Bioconductor package DESeq2 in R version 3.6.2 requiring a FDR below 0.05 and an absolute log fold-change of at least 1 (https://www.bioconductor.org)72. Each analysis controlled for experimental batches and differences are reported as log2 fold-change with respect to the control group. Sequencing read counts were normalized using the regularized log transformation (rlog) in DESeq2 before PCA. The top 20 DFG (by P value) are shown in the heatmaps. To ensure comparability across genes, read counts were normalized by variance stabilizing transform (vst) in DESeq2 and mean normalized read counts per gene subtracted, before plotting the heatmap.
ChIP-seq procedure and bioinformatic analysis
ChIP-seq analysis was carried out as previously published73,74. CHD8 antibodies were bought from Novus and Bethyl (NB100-60417 and A301-224A, respectively), H3K4me3 antibodies (Active Motif, Cat number 39159); INO80 ab was purchased from Abcam (Cat. Number: ab105451). Reads were trimmed with cutadapt (http://cutadapt.readthedocs.io/en/stable/index.html) and aligned to the genome with bowtie2 using the ‘very sensitive’ alignment option75. Picard tools were used to remove duplicate reads and samTools to filter out reads with mapping quality below two (https://broadinstitute.github.io/picard/). Subsequently, coverage plots were generated with deepTools using a bin size of 50 (or 100b) and RPKM (reads per kilobase per million) normalization76. The software MACS2 called peaks with 0.05 as FDR cut-off and consensus peak sets were derived for the duplicated experiments using Bioconductor’s package DiffBind (http://liulab.dfci.harvard.edu/MACS/; https://www.bioconductor.org).
Comparison with other datasets
YY1 data was downloaded from the GEO repository (GSE92407). Bigwig files for YY1 (2i, 2 replicates) were converted to bedGraph. Peaks were called on the bedgraph files with MACS2. Consensus YY1 peaks were derived by intersecting the peaks called on the two replicates with bedtools. YY1 peaks were lifted from mm9 to mm10 using easyLift in R. YY1 and Chd8 peaks were overlapped using GenomicRanges in R. Peaks were annotated to genes (including distance to TSS) using the GREAT website (http://great.stanford.edu/public/html/). Promoter regions were defined as 0–2 kb upstream of the TSS, getting strand-specific information via biomaRt from the “m. musculus_gene_ensembl” dataset.
ChIP- and ATAC-seq allelic analysis
ATAC-seq reads were processed and aligned as described elsewhere77,78. The software SNPsplit (https://www.bioinformatics.babraham.ac.uk/projects/SNPsplit/) was used for allele-specific analysis. In addition, to the alignment to the reference genome the reads were aligned to a genome in which all SNPs have been masked. Subsequently, aligned reads were divided into reads which either match CAST_EiJ or 129S1_SvImJ SNPs or if not were considered unspecific. Downstream analysis was done as before separately for each class of reads.
This protocol was adapted from a CUT&RUN Protocol from EpiCypher (https://www.epicypher.com/content/documents/protocols/cutana-cut&run-protocol.pdf). Cells were harvested with accutase, counted and centrifuged at 600 × g for 3 min. Pellets were washed three times in 1.5 ml Wash buffer (20 mM HEPES pH 7.5; 150 mM NaCl; 0.5 mM Spermidine; 1× protease inhibitors EDTA-free) and resuspended in the right volume of Wash buffer in order to have 500,000 cells per 100 µl. 100 µl of the cell suspension were then transferred into an 8-stripe tube containing 10 µl of previously activated concanavalin A-coated magnetic beads (activating buffer: 20 mM HEPES pH 7.5, 10 mM KCl, 1 mM CaCl2, 1 mM MnCl2) (86057-3, BioMag®Plus Concanavalin A, Generon) and placed in a rotator for 10 min at room temperature. Cells were resuspended in 50 µl antibody buffer (5% (wt/vol) digitonin, 20 mM HEPES pH 7.5; 150 mM NaCl; 0.5 mM Spermidine; 1× protease inhibitors EDTA-free, 2 mM EDTA) and 0.5 µl antibody was added per sample and incubated at 4 °C overnight in a rotator. The following antibodies were used: YY1 (H-10): Santa Cruz sc-7341; YY1 Diagenode AB_2793763; WDR5: Diagenode C15310101. Cells were washed 3× in 250 µl Dig-wash buffer (5% (wt/vol) digitonin, 20 mM HEPES pH 7.5; 150 mM NaCl; 0.5 mM Spermidine; 1× protease inhibitors EDTA-free) to remove unbound antibody. Cells were resuspended in 50 µl cold Dig-wash buffer and pAG-MNase 1:200 was added and incubated at room temperature for 10 min At this stage, supernatant from samples incubated with IgG antibody was collected and stored to be used as input. Cells were washed 3× in 250 µl cold Dig-wash buffer to remove unbound pAG-MNase. Tubes were placed on ice and quickly mixed with 100 mM CaCl2 to a final concentration of 2 mM diluted in 50 µl Dig-wash buffer per sample. Cells were incubated for 2 h at 4 °C and the reaction was quenched by the addition of 33 µl 2× STOP buffer (340 mM NaCl; 20 mM 0.5 EDTA; 4 mM EGTA; 0.05% digitonin; 100 µg/mL RNAse A; 50 µg/mL glycogen). Cleaved fragments were liberated into the supernatant by incubating the cells at 37 °C for 10 min, and the supernatant was stored afterwards. DNA fragments were purified from the supernatant using the ChIP DNA Clean and Concentrator Kit (D5205, Zymo Research) and used for the construction of sequencing libraries.
MEME analysis and string
MEME analysis was done by extracting 100 and 500 bp windows underlying Chd8 peaks in undifferentiated and differentiating conditions. These sequences have been analysed using MEME motif discovery via the weblink (http://meme-suite.org/). In order to discover Chd8-interacting proteins, we used the online version of the STRING database (https://string-db.org/).
Chd8 IP-mass spectrometry
CHD8 and HA IP were done using Chd8 antibodies (abs) bought from Novus biological (NB100-60417) and Bethyl (A301-224A), HA ab was acquired from Sigma (H6908). For each sample 500 µl of extracted lysate was brought to 1 ml with RIPA buffer and precleared with 50 µl Protein G Dynabeads (Thermo) for 30 min on a rotating wheel at 4 °C. Beads were removed and 15 µl of antibodies was added to the cleared lysate and incubated at 4 °C for an hour on the rotating wheel, after which 50 µl of washed protein G beads were added and incubated for another hour. Beads were washed three times with 250 µl RIPA buffer and eluted in 20 µl Laemmle buffer by boiling for 5 min Samples were run on a 10% SDS–PAGE gel and analysed by mass spectrometry. Protein and RNA sequences were downloaded from UCSC genome browser and Uniprot, respectively (https://genome.ucsc.edu/; https://www.uniprot.org/).
Bands were cut from the gel and subjected to in-gel digestion with trypsin79. Peptides were extracted from the gel pieces by sonication for about 15 min, followed by centrifugation and the collection of the supernatant. A 50:50 solution of water:acetonitrile, 1% formic acid (2 × the volume of the gel pieces) was added for a second extraction. The samples were re-sonicated for 15 min, centrifuged and the supernatant pooled with the first extract. The pooled supernatants were processed using speed vacuum centrifugation. The samples were dissolved in 10 µL of reconstitution buffer (96:4 water:acetonitrile, 1% formic acid and analyzed by LC–MS/MS.
Peptides were separated using the nanoAcquity UPLC system (Waters) fitted with a trapping devise (nanoAcquity Symmetry C18, 5 µm, 180 µm × 20 mm) and an analytical column (nanoAcquity BEH C18, 1.7 µm, 75 µm × 200 mm). The outlet of the analytical column was coupled to an LTQ Orbitrap Velos (Thermo Fisher Scientific) using the Proxeon nanospray source. Solvent A was water, 0.1% formic acid and solvent B was acetonitrile, 0.1% formic acid. Samples were loaded with a constant flow of solvent A at 5 µL/min onto the trapping column, as routinely done. Trapping time was 6 min Peptides were eluted via the analytical column a constant flow of 0.3 µL/min. During the elution step, the percentage of solvent B was increased in a linear fashion. The peptides were introduced into the mass spectrometer instrument (Orbitrap Velos, Thermo) via a Pico-Tip Emitter 360 μm OD × 20 μm ID; 10 μm tip (New Objective) and a spray voltage of 2.2 kV was applied. The capillary temperature was set at 300 °C. Full scan MS spectra with mass range 300–1700 m/z were acquired in profile mode in the FT with a resolution of 30,000. The filling time was set at maximum of 500 ms with limitation of 1.0 × 106 ions. The most intense ions (up to 15) from the full scan MS were selected for sequencing in the LTQ. Normalized collision energy of 40% was used, and the fragmentation was performed after accumulation of 3.0 × 104 ions or after filling time of 100 ms for each precursor ion (whichever occurred first). MS/MS data was acquired in centroid mode. Only multiply charged (2+, 3+, 4+) precursor ions were selected for MS/MS analysis. The dynamic exclusion list was restricted to 500 entries with maximum retention period of 30 s and relative mass window of 10 ppm. In order to improve the mass accuracy, a lock mass correction using the ion (m/z 445.12003) was applied.
MS data analysis
The raw output files of IsobarQuant (protein.txt–files) were processed using the R programming language (ISBN 3-900051-07-0). Only proteins that were quantified with at least two unique peptides were considered for the analysis. Raw TMT reporter ion signals (signal_sum columns) were first cleaned for batch effects using the ‘removeBatchEffect’ function of the limma package80 and further normalized using vsn (variance stabilization normalization)81. Proteins were tested for differential expression using the limma package. A protein was annotated as a hit with a FDR smaller 5% and a fold-change of at least 100% and as a candidate with a fdr below 10% and a fold-change of at least 50%.
Conventional two-tailed t-test and Poisson regression fittings were used in R environment (https://www.r-project.org/) or JMP software (jmp.com) or excel (Microsoft). All experimental data points are clearly presented in the figure legends or in the figure the statistical test used and the error bar type is clearly indicated. No statistical test was performed on log-scaled values.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
Next-generation sequencing data has been deposited in GEO, Access numbers: GSE166858 and GSE166859. All source data underlying the graphs and charts presented in the main figures are available in Supplementary Data 6. Mass-spectrometry data has been deposited in the PRIDE database, accession number: PXD024155. All data is available upon request.
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We would like to thank all members of the Avner group and Claire Rougeulle, Joost Gribnau, Sara Buonomo and Kristina Havas for critical reading of the manuscript. Our thanks also go to Pedro Moreira, Philip Hublitz, Jim Sawiztke, Rabea Jesser, Alessandra Liverziani, Monica Di Giacomo, Eleftheria Ledaki for help in the initial phase of this project and for the generation of recombineering constructs and cell lines. We thank M. Gerard for providing the Chd8-KI construct and for sharing with us unpublished observations. We would like to thank Frank Stein and Mandy Rettel from the EMBL Proteomic Core facility for their help with the mass spectrometry experiments. This work was funded by EMBL recurrent funding to P.A. (50800) and a Rett Syndrome Research Trust and QMUL intramural support to A.C. and BARTS Charity grants to A.C. A.N.Y., A.B., G.M.S., M.A. (M. Arnold), MA (M. Ascolani), are/were funded by EMBL grants. S.J.M. is funded by an Edmond and Lily Safra Research Fellowship and a UK Dementia Research Institute Career Development Fellowship. The UK Dementia Research Institute receives its funding from UK DRI Ltd, funded by the UK Medical Research Council, Alzheimer’s Society and Alzheimer’s Research UK.
The authors declare no competing interests.
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Cerase, A., Young, A.N., Ruiz, N.B. et al. Chd8 regulates X chromosome inactivation in mouse through fine-tuning control of Xist expression. Commun Biol 4, 485 (2021). https://doi.org/10.1038/s42003-021-01945-1