H3K36 Methylation as a Guardian of Epigenome Integrity

Domain: neuroscience
Published: 2025-12-11
Editorial outcome: major_revision

Abstract
H3K36 methylation is a key epigenetic mark with critical roles in development and disease. Here, we systematically dissect its functions using CRISPR-engineered mouse mesenchymal stem cells lacking combinations of the five H3K36 methyltransferases, culminating in quintuple knockout cells devoid of H3K36me2/3. We show that H3K36me2 influences enhancer activity, supports the expression of their target genes, and safeguards active genes from encroachment of the repressive marks, H3K27me2/3. In addition, we find that the loss of H3K36me triggers redistribution of large heterochromatic H3K9me3 domains into euchromatin, in part mediated by SUV39H1, leading to global epigenomic remodelling, constitutive heterochromatin erosion, and a collapse of 3D genome organization. Parallel analyses in human HNSCC cells overexpressing the H3K36M oncohistone reveal conserved disruptions to the epigenome and chromatin architecture. Together, these results establish H3K36 methylation as a pivotal regulator of chromatin state and genomic structure.

Introduction
DNA is wrapped around histones to form nucleosomes, the structural and functional unit of chromatin. Chromatin can be densely packed as heterochromatin or loosely packed as euchromatin. The extent of compaction, and thereby access to DNA, is in part regulated by chemical modifications to the histone tails, known as histone post-translational modifications (PTMs). These histone PTMs form a highly interdependent network, where euchromatic modifications facilitate access to DNA and heterochromatic modifications restrict access to DNA1–3. The different types of modifications at histone residues induce conformational changes that influence adjacent or interacting residues.

Methylation of histones plays pivotal roles in modulating transcriptional processes, primarily by influencing the accessibility of chromatin and serving as a substrate for a multitude of effector proteins to exert their functions. The lysine residues on the N-terminal tails of histones can receive up to three methylation marks, and the degree of methylation at specific residues is often associated with distinct transcriptional states and genomic features4,5. While the methylation states of certain residues coincide on the same histone tail and support similar processes, the methylation states of other residues are antagonistic, often due to steric bulk, and support opposing processes. In addition, the collective effects of histone methylation significantly influence the organization and 3D architecture of interphase chromatin6–11. Over the past decade, methylation of histone 3 lysine 36 (H3K36me) has increasingly emerged as a critical component of the epigenome. When aberrantly regulated, either through mutations affecting the genes of its writers or of histone 3 (H3), developmental disorders or cancer ensues, underscoring its functional significance12–21. H3K36me exists in mono- (H3K36me1), di- (H3K36me2), or tri-methylated (H3K36me3) states, and there are five well-established enzymes that catalyze H3K36 methylation: SETD2, NSD1, NSD2, NSD3, and ASH1L. Briefly, SETD2 tethers to the elongating RNAPII complex within actively transcribed genes to deposit nearly all H3K36me322,23. In contrast, the NSD family of lysine 36 methyltransferases (K36MTs) have established roles in the deposition of the lower methylation states, H3K36me1/2. NSD1 and NSD2 have been found to be responsible for the majority of global H3K36me2, in both IGRs and genes, and their individual contributions appear to vary by cell type and developmental stage3,14,18,24–26. In comparison, we have previously found that NSD3 has a more distinct role in the deposition of H3K36me2, primarily exerting its catalytic activity at active cis-regulatory elements (CREs)—most notably at enhancers and promoter flanking regions26. Finally, ASH1L is the most specific and least prolific of the K36MTs, primarily depositing H3K36me2 at specific developmentally important genes26.

Since H3K36me is generally associated with transcriptionally active, euchromatic regions of the genome, it is presumed to play a role in maintaining and fine-tuning transcription. It is one of the modifications broadly flanking CREs, whose regulatory states are frequently inferred by the enrichment profiles and co-localization of several histone modifications27. Notably, H3K4 methylation (H3K4me) and H3K27 acetylation (H3K27ac) are amongst the most well characterized modifications that distinguish active CREs. Functionally, H3K4me1 serves to fine tune enhancer activity by acting as a substrate for the binding of chromatin remodelers and other protein effectors28, while H3K27ac has a more direct effect on chromatin by neutralizing the positively charged histone tails, thereby weakening their interaction with DNA and establishing an environment permissive for transcription29. The co-presence of H3K4me1 and H3K27ac is frequently used to identify active enhancers27,30. In contrast, focal peaks of H3K4me3 and H3K27ac are hallmark chromatin signatures of active promoters27,31. H3K36me2 has been found to exist in broad domains at both active enhancers and promoters in most assayed cell types3,18,24–26. Loss of broad H3K36me2 domains, resulting from either pathological mutations or genetic ablation of its writer enzymes, has been shown to impact the distribution of H3K4me1/3 and H3K27ac, and is associated with altered gene expression18,25,32. While the general relationship between H3K36me2, CREs and gene expression has been documented, it has always been studied in the context of very broad, megabase scale domains, and the precise role of H3K36me2 in maintaining the activity of CREs and its influence on the deposition of other active histone marks in a localized setting has yet to be carefully examined.

Our understanding of the relationship between H3K36me and heterochromatin is also far from complete. On one hand, there exists a well-documented antagonism between H3K36me2/3 and the silencing marks H3K27me2/3. The PRC2 complex, which deposits all H3K27me, is allosterically inhibited from nucleosomes marked with H3K36me2/333,34. As a result, particularly H3K27me3 is excluded from H3K36me2/3 domains. While H3K27me2/3 are considered to be repressive marks, they are generally found in gene-rich regions and their presence is associated with facultative heterochromatin35. In comparison, H3K9me3 is distinctive of constitutive heterochromatin, and broad H3K9me3 domains generally span gene-poor regions containing tandem repeat elements, such as those found at telomeres and pericentromeric regions6,36,37. While broad genomic domains of H3K9me3 and H3K36me2 tend to be mutually exclusive, the relationship between H3K36me and H3K9me is currently unclear, largely because evidence for mechanistic links between the two marks is lacking.

One of the challenges associated with investigating the functional role of H3K36me is the broad nature of its distribution, which is exacerbated by the number of enzymes capable of depositing these modifications. We have previously used CRISPR-Cas9 gene editing to establish clonal cell lines harboring individual or multiple knockouts (KOs) of the K36MTs, for the purpose of deconstructing their individual and combined contributions to the H3K36me landscape in C3H10T1/2 mouse mesenchymal stem cells (mMSCs)26. Here, we employ these cell lines to investigate the effects of H3K36me in more specific contexts: we examine focal peaks of NSD3- and ASH1L-mediated H3K36me2 present at active CREs, and their subsequent erasure, to assess their functional influence on the local epigenome and transcriptome. We provide further mechanistic insights into the ability of H3K36me2/3 to exclude H3K27me2/3 from actively transcribed genes, and the downstream impact on other relevant epigenetic modifications in these regions. Unexpectedly, we uncover a relationship between H3K36me, H3K9me3 and the 3D architecture of the genome that appear to be conserved in both mouse and human cells. Overall, the progressive loss of H3K36me resulting from the sequential deletion of the K36MTs provides a unique opportunity to assess the role of H3K36me in maintaining the active state of CREs, transcription and genomic compartmentalization.

Results
NSD3-mediated H3K36me2 maintains the active state of enhancers and their target genes
We have previously found that knocking out NSD1/2 (DKO) in mMSCs results in a near total depletion of broad intergenic H3K36me238 (Fig. 1a). Further KO of SETD2 in the triple KO cells (TKO) results in the loss of nearly all genic H3K36me2 and H3K36me326 (Fig. 1a). The remaining focal H3K36me2 regions, deposited by NSD3, are primarily targeted to active CREs26. Upon subsequent KO of NSD3 in the NSD1/2/3-SETD2 quadruple KO cells (QKO), these broad peaks of H3K36me2 disappear (Fig. 1a). To further explore the role of H3K36me2 in these confined regions, we examined promoters and enhancers that remain accessible, as defined by the presence of an ATAC-seq peak in both the TKO and QKO conditions. Having previously established biological replicates for each cell line, here we used ChIP-seq (H3K36me2, H3K27ac, H3K4me3) and CUT&RUN (H3K4me1, H3K27me3) to profile the relevant active and repressive histone modifications (Supplementary Fig. 1a), ATAC-seq for chromatin accessibility, whole-genome bisulfite sequencing (WGBS) for DNA methylation (DNAme), and RNA-seq for transcriptional state to document the downstream changes following the depletion of H3K36me2 at CREs and their associated genes.

At active enhancers, the depletion of H3K36me2 is accompanied by slight decreases in chromatin accessibility (Fig. 1b). Concurrently, we also observe a decrease in the levels of other known active enhancer modifications, specifically H3K4me1/3 and H3K27ac (Fig. 1b). In contrast, the levels of H3K27me3 increase at enhancers that are depleted of H3K36me2 (Fig. 1b), which was expected given the known antagonistic relationship between these marks33,34. Previous studies have shown that H3K36me2/3 serve as templates for the deposition of de novo DNAme by DNMT3A/B, respectively38,39. Therefore, we expected that in the absence of H3K36me2/3, enhancers may experience substantial reductions in CpG methylation. Using WGBS, however, we find that DNAme is largely unchanged (Fig. 1b). We also interrogated strong enhancers, as defined by dense clusters of individual enhancers (Fig. 1a; Supplementary Fig. 1b), which are known to be essential for the maintenance and regulation of genes crucial to both normal development and cancer40. In TKO cells, most strong enhancers retain H3K36me2, which is subsequently depleted in the QKO condition, demonstrating that NSD3 is recruited to these regions (Fig. 1a, c). Here, we found similar yet more pronounced trends in the decrease of activating and increase of silencing modifications, further demonstrating that H3K36me2 is essential to the maintenance of enhancer activity (Fig. 1c).

Interestingly, the changes occurring at the promoters of NSD3 target genes appear to be considerably more attenuated, as compared to those at enhancers. The overall levels of NSD3-deposited H3K36me2 in TKO cells are lower at accessible promoters than at accessible enhancers (Fig. 1b–d). Thus, while the loss of H3K36me2 is nearly complete at all CREs, the relative decrease is smaller at promoters. Consequently, at promoters we observe no change to chromatin accessibility and H3K27ac, while the decrease of H3K4me1 and increase of H3K27me3 is lower than at enhancers (Fig. 1d). We do, however, observe a comparable twofold decrease in both promoter- and enhancer-marked H3K4me3 (Fig. 1d). Overall, these results provide further evidence that the presence of NSD3-associated H3K36me2 is particularly influential at enhancers.

We next examined the effects on gene expression following the depletion of CRE-associated H3K36me2. Genome-wide, we found no general trend towards downregulation, with more than twice as many genes going up in expression (289) than down (97) (Fig. 1e). However, many of these changes may be secondary, downstream effects from global H3K36me2 depletion and/or ablation of NSD3. In view of the more pronounced epigenetic changes occurring at enhancers, we hypothesized that the most direct effect on gene expression should be observed at genes whose expression is dependent on H3K36me2-marked enhancers. We identified a set of predicted H3K36me2-dependent genes based on their overlap with large H3K36me2 peaks that contained at least one annotated enhancer (from Ensembl and FANTOM databases). We also created a complimentary, randomized control set of H3K36me2-independent genes. The two sets were further matched for baseline gene expression levels by selecting only transcripts within the 9-12 FPKM range (Supplementary Fig. 1c). We found that following the loss of NSD3-dependent H3K36me2, genes associated with large H3K36me2 peaks become significantly more downregulated (150 down versus 69 up) compared to the control set of genes (108 down versus 111 up) (Fig. 1f). Our findings indicate that the depletion of NSD3-dependent H3K36me2 impacts gene expression primarily by reducing enhancer activation of target genes rather than directly influencing promoter activity.

We have previously examined the effects of the individual KO of NSD3 on the global H3K36me2 landscape, and we did not observe a significant depletion in the global abundance of H3K36me2, possibly because of compensation by the other K36MTs26. To investigate whether the effects on CREs are a result of the H3K36me2 catalytic product or driven by NSD3 itself, we analyzed the same set of enhancer-dependent NSD3 target genes identified in the TKO cells in the context of comparing parental to NSD3-KO cells. Although globally, we find that many more genes significantly increase in expression (172) than decrease (8) (Supplementary Fig. 1d), within the set of predicted H3K36me2-dependent genes derived from the TKO cells, we found a tendency towards downregulation (122) versus upregulation (76) (Supplementary Fig. 1e). This suggests that the compensation of H3K36me2 levels by the other K36MTs is not complete. Upon closer inspection, we observe a decrease of H3K36me2 at the promoters of this set of genes in NSD3-KO cells (Supplementary Fig. 1f) and an even greater decrease at the nearest enhancers of these promoters (Supplementary Fig. 1g). Hence, even in parental cells, where NSD3 is not the dominant K36MT, its absence is reflected in a slight, localized reduction of H3K36me2 and downregulation of enhancer-dependent genes.

Overall, our results support that while NSD3-mediated H3K36me2 is targeted to both enhancers and promoters, it is most influential at enhancers with regards to maintaining their active chromatin states, restricting the invasion of H3K27me3, and has discernible effects on maintaining the expression of their target genes.

Functional characterization of focal H3K36me2 at ASH1L target genes
In the QKO cells, we had previously identified 119 remaining H3K36me2 peaks, with the majority of these peaks straddling the transcription start sites (TSS) (n=60) or located within the gene bodies (n=24) of a subset of developmentally important genes26. Those peaks disappeared in the NSD1/2/3-SETD2-ASH1L-QuiKO (QuiKO) condition, following the ablation of ASH1L (Fig. 2a). Similar to the comparison between TKO and QKO cells, this provides a unique opportunity to study the functional consequences of H3K36me2 depletion in a very specific, as opposed to a global setting. We find that the majority of ASH1L target genes (62/84, p=0.0015, Wilcoxon test) decrease in expression following the loss of ASH1L (Fig. 2b, c). Furthermore, the change in expression is proportional to the size of the original H3K36me2 peaks: genes with the largest H3K36me2 peaks in the QKO cells have the largest fold changes in gene expression (Fig. 2d). In contrast, the same set of genes did not decrease in expression when comparing TKO to QKO cells, where H3K36me2 is unaffected at these promoters (Fig. 2b, c). Finally, it is possible that ASH1L itself may act as a transcriptional modulator, and may have an effect independent of its catalytic activity-as has been recently suggested in the case of NSD125. To distinguish whether the reduced expression of the 84 ASH1L-target genes may be due to the loss of ASH1L itself, or its catalytic product, H3K36me2, we looked at differences in expression between parental mMSCs and the individual ASH1L-KO cells. In the ASH1L-KO condition, the levels of H3K36me2 remain largely unchanged at the promoters of these genes, most likely as a result of compensation by the other K36MTs, which exhibit less specificity in their deposition of H3K36me2 (Supplementary Fig. 2a). Interestingly, we found no significant changes to the expression of ASH1L target genes, suggesting that the presence of H3K36me2, rather than the presence of ASH1L has a role in maintaining their expression (Fig. 2c).

Concurrently, we investigated changes to other epigenetic modifications at the remaining CREs marked with residual H3K36me2. Genome-wide, we found that chromatin accessibility increases in QuiKO cells compared to QKO, however, the CREs marked with residual H3K36me2 gain considerably less (Fig. 2e, f). This likely reflects the net negative effect of losing H3K36me2 at these specific loci in comparison to other genomic regions. Nevertheless, the depletion of ASH1L-mediated H3K36me2 is accompanied by a further decrease of H3K27ac and a corresponding increase of H3K27me3 at these CREs (Fig. 2e, f). Similar to the loss of NSD3 in QKO cells, we observe that the loss of H3K27ac is greater at enhancers than promoters (Fig. 2e, f), further supporting that the presence of H3K36me2 at CREs may be more influential at enhancers than promoters. Altogether, these analyses demonstrate that H3K36me2 contributes to maintaining the balance between activating and silencing epigenetic modifications at CREs, and that this has consequences for the expression of target genes.

The effect of H3K36 methylation on antagonizing H3K27me within genes
One of the presumed functions of H3K36me3 that is deposited within actively transcribed genes is to prevent the deposition of the silencing modifications at H3K2733,34. This effect may further be strengthened by the presence of H3K36me1/2. It has been shown that methylation at H3K36 antagonizes PRC2 and hinders the deposition of methylation at H3K2733,34,41. This is particularly true for the highest methylation levels, resulting in mutual exclusivity of H3K36me3 and H3K27me3 on the same histone tail34,41. Accordingly, H3K27me3 is excluded from actively transcribed genes, H3K27me2 is generally depleted, and only H3K27me1 invades gene bodies, where its distribution is similar to that of H3K36me3 (Fig. 3a). Using our panel of CRISPR-engineered mMSCs, we investigated the effects of H3K36me depletion on the presence of H3K27me in gene bodies. We used quantitative mass spectrometry (MS) to quantify the changes to the absolute abundance of each of the respective methylation marks, and ChIP-seq/CUT&RUN to profile changes to the global distributions of H3K27me states. Unexpectedly, in the SETD2 mutant which lacks nearly all H3K36me3 (Supplementary Fig. 3a), the increase of H3K27me3 in gene bodies is minimal, and primarily discernible within lowly transcribed genes (Fig. 3a; bottom row, middle box). However, the absence of H3K36me3 has a more pronounced effect on H3K27me2, allowing its deposition within gene bodies largely independent of transcriptional activity, and is accompanied by a corresponding reduction in H3K27me1 (Fig. 3a; middle row). Accordingly, these genic changes are reflected in the overall genome-wide abundance: H3K27me2 slightly increases, and this is accompanied by a proportional decrease of H3K27me1, while changes to H3K27me3 are minimal (Fig. 3b). These observations may be explained by the fact that the sole loss of SETD2 in the presence of the other intact K36MTs primarily depletes H3K36me3, and results in an increase of genic H3K36me2 due to fewer residues being upgraded to the higher methylation state26 (Supplementary Fig. 3a). Since the presence of H3K36me2 also hinders the activity of PRC2 to deposit higher orders of H3K27me33, these results suggest that the presence of H3K36me3 inhibits the deposition of both H3K27me2/3, while the presence of H3K36me2 may primarily inhibit the deposition of H3K27me3.

Therefore, we next focused specifically on the ability of H3K36me2 to protect gene bodies from methylation at H3K27. In NSD1/2-DKO cells, the levels of H3K36me2 are significantly reduced, with a moderate reduction of H3K36me3 (Supplementary Fig. 3a). This is accompanied by a significant increase in the global levels of H3K27me2/3 (Fig. 3b). Interestingly, the partial reduction of H3K36me2 has a more pronounced effect on genic H3K27me2/3 than what is observed with the near total depletion of H3K36me3 (Fig. 3a; middle and bottom rows). Although in vitro studies indicate a stronger antagonistic effect from H3K36me333, the greater observable effect of H3K36me2 depletion may reflect the generally higher absolute levels of H3K36me2 than H3K36me3 in gene bodies.

Finally, we used NSD1/2-SETD2-TKO cells to study the consequences of depleting both H3K36me2/3. Expectedly, in these cells we observe the strongest invasion of H3K27me2/3 into gene bodies (Fig. 3a; middle and bottom rows). Both H3K27me2/3 now appear free to populate genes, with little dependence on expression levels (Fig. 3a). Further depletion of H3K36me2 in our QKO and QuiKO cell lines leads to even greater invasion of H3K27me3 into gene bodies (Supplementary Fig. 3b). Interestingly, the cells overexpressing the H3K36M oncohistone (H3K36M-OE), which reduces both H3K36me2/3, show invasion of H3K27me that is intermediate between the DKO and TKO conditions (Fig. 3a, b).

To better illustrate the global changes to the distribution of H3K27me across the various conditions, we calculated H3K27me3 ‘peakiness scores’, which represents the average ChIP-seq/CUT&RUN signal in the top 1% of 1-kb bins over the total signal, confirming the broadening (lower peakiness score) of H3K27me3 following sequential removal of H3K36me (Supplementary Fig. 3c). The lower peakiness scores in the multiple-KO conditions demonstrates that as H3K36me is sequentially removed, the distribution of H3K27me3 shifts from a focused distribution in parental cells, to a broader distribution in cells lacking H3K36me. Lastly, more than 1.5-fold of genes with significant H3K27me3 invasion in their gene body decrease in expression (609 down-regulated compared to 391 upregulated), further supporting the hypothesis that the invasion of H3K27me3 into gene bodies contributes to suppressing gene expression (Fig. 3c).

Overall, while our results demonstrate that H3K36me plays a role in excluding H3K27me2/3 from gene bodies, additional factors are clearly responsible for preventing the full invasion of H3K27me. The two likely culprits are 1) nucleosome turnover, which results in replacement of modified with naïve nucleosomes during transcription42, and 2) DNAme, which may hinder the recognition of CpG-rich sequences, such as those found in exons, by PRC243,44.

The effect of H3K36 methylation on other epigenetic modifications within gene bodies
As previously described, H3K36me2/3 have been shown to serve as substrates for the localization of DNMT3A/B to deposit de novo DNAme38. Therefore, we expected that in the absence of H3K36me2/3, gene bodies may experience substantial reductions in CpG methylation. However, we find that even in the QuiKO cells, which lack nearly all H3K36me, the most highly expressed genes retain DNAme levels comparable to parental cells (Fig. 4a; third box). Methylation is visibly reduced in the lowest expression quantile, as well as in unexpressed genes–where the latter behaves similarly to inactive intergenic regions where DNA hypomethylation has previously been shown to be associated with the reduction of intergenic H3K36me2 (Fig. 4a; first and second box)3,18,38. It appears, however, that in highly expressed genes, H3K36me-mediated targeting of de novo DNMTs is unnecessary for the maintenance of high DNAme levels.

Within the gene bodies of the K36MT mutants, we detected no discernible changes in H3K27ac or H3K4me3, regardless of transcription activity (Supplementary Fig. 4a, b). In contrast, H3K4me1 levels increase substantially within gene bodies, with the degree of H3K36me2 depletion correlating with greater invasion of H3K4me1—even in previously silent genes and with minimal dependence on transcription (Fig. 4b). Consistent with these findings, we observe by MS that the global abundance of H3K4me1 increases in the multiple-KO conditions (Fig. 4c). Additionally, the distribution of H3K4me1 becomes increasingly diffuse across the multiple-KO conditions, as reflected by a progressive decrease in ‘peakiness’ scores (Supplementary Fig. 4c). Interestingly, this global increase in H3K4me1 is accompanied by alterations in the expression of the primary H3K4 methyltransferases: KMT2C and KMT2D. Specifically, KMT2C expression is reduced, while KMT2D expression is moderately upregulated in the multiple-KO mMSCs, raising the possibility that KMT2D may be responsible for the ectopic distribution of H3K4me1 (Supplementary Fig. 4d).

Surprisingly, we also found significant changes to the genic levels of the silencing-associated modification H3K9me3, whose relationship with H3K36me is currently unclear. In parental cells, H3K9me3 is generally excluded from transcribed genes, however, gradual removal of H3K36me results in an increase of H3K9me3 that appears to be correlated with both gene expression and the amount of residual H3K36me2/3 (Fig. 4d, Supplementary Fig. 3a). In DKO cells, where the loss of NSD1 and NSD2 substantially reduces H3K36me2, genic H3K9me3 increases, especially in lowly expressed genes (Fig. 4d; second box). In TKO cells, which also lack SETD2 and thereby most H3K36me3, the gene body levels of H3K9me3 increase further and appear to lose their dependence on transcription (Fig. 4d). These results intriguingly suggest that the presence of H3K36me2/3 within genes may antagonize the deposition of H3K9me3 in these regions.

Given the considerable increase in genic H3K9me3 in the TKO cells, we used MS to quantify the global abundance of H3K9me3 across conditions. Interestingly, we find that the bulk levels of H3K9me3 increase following the depletion of H3K36me3 in both SETD2-KO and TKO cells (Fig. 4e), suggesting that H3K36me3, in particular, may antagonize the deposition of H3K9me3 within genes. However, the genome-wide abundance of H3K9me3 in the QKO and QuiKO conditions returns to levels comparable to parental cells, and do not necessarily reflect the observed increases in active genes (Fig. 4e, Supplementary Fig. 4e). Overall, despite the increase of H3K9me3 in actively transcribed genes, the genome-wide abundance in the multiple-KO conditions does not significantly change. Thus, we hypothesized that there must be a corresponding decrease in other regions of the genome.

Loss of H3K36me leads to SUV39H1-mediated redistribution of H3K9me3 from heterochromatin to euchromatin
In parental mMSCs, the distribution of H3K9me3 follows two characteristic patterns: at a broad scale, it is deposited in large domains spanning several megabases (Fig. 5a), corresponding to highly heterochromatic, transcriptionally silent, lamina-associated domains6,45,46. At a finer scale, peaks of H3K9me3 are present at the promoters of certain silent genes and transposable elements (TEs) (Fig. 4d; ‘silent genes’). Remarkably, under H3K36me deficient conditions, these broad H3K9me3 domains entirely disappear (Fig. 5a). The depletion of heterochromatic domains is accompanied by increased H3K9me3 deposition in regions that are predominantly active, which, upon closer examination, are primarily gene bodies and TEs (Fig. 4d; Supplementary Fig. 5a). To identify and further characterize the genomic compartments with the greatest loss of H3K9me3, we subdivided the genome into 100-kb bins and compared profiles between parental and TKO cells, which are largely devoid of H3K36me. The H3K9me3 profiles of the genomic bins subdivide into two distinct clusters (Fig. 5b): cluster A (orange), comprising predominantly genic regions that gain H3K9me3 in TKO cells, and cluster B (blue) consisting mainly of intergenic regions that lose H3K9me3 in TKO cells.

To determine whether this redistribution of H3K9me3 in response to H3K36me loss is conserved in other cell types, we examined two human head and neck squamous cell carcinoma (HNSCC) cell lines: Cal27 and Detroit562. These HPV(-) HNSCC cell lines lack endogenous mutations affecting H3K36me18. In these cells, we overexpressed (OE) the H3K36M oncohistone—a mutation known to globally reduce all H3K36 methylation states and found in HPV(-) HNSCC tumors as well as other cancers (Supplementary Fig. 5b)14,24,47. As in mMSCs, parental Cal27 and Detroit562 cells display well-defined H3K9me3-marked heterochromatic domains (Fig. 5c, Supplementary Fig. 5c). Strikingly, overexpression of H3K36M in both cell lines resulted in marked depletion of these large H3K9me3 domains, corresponding to cluster B regions identified by our genome-wide density-based clustering analysis (Fig. 5b–d, Supplementary Fig. 5c–e). Furthermore, unlike in mMSCs, the loss of H3K9me3 domains in these cells was accompanied by a global reduction in H3K9me3 levels of approximately 35% (Supplementary Fig. 5b). These findings suggest that H3K36 methylation plays a conserved role in maintaining higher-order H3K9me3 organization across cell types.

Using the cluster A and B annotations, we next investigated the molecular mechanisms underlying H3K9me3 redistribution following H3K36me depletion. Specifically, we focused on SUV39H1, one of the primary H3K9 methyltransferases (K9MTs) which catalyzes H3K9me3 and plays a critical role in the propagation of H3K9me3 domains through a read-write mechanism, wherein its chromodomain recognizes existing H3K9me3 to enhance further methyltransferase activity48,49. Unlike SETDB1, which deposits focal peaks at repetitive elements, and SUV39H2, which is testis-enriched and minimally expressed in somatic cells, SUV39H1 has both the expression profile and enzymatic behavior consistent with widespread ectopic H3K9me3 deposition (Supplementary Fig. 5f)50,51. To profile the genomic distribution of SUV39H1, we overexpressed FLAG-tagged SUV39H1 in parental and TKO mMSCs and performed ChIP-Seq for FLAG-SUV39H1 (Supplementary Fig. 5g)52. In parental mMSCs, SUV39H1 exhibited enrichment patterns similar to H3K9me3 and was primarily localized to inactive cluster B regions (Fig. 5e, f; Supplementary Fig. 5h). In the TKO cells, however, SUV39H1 relocates to cluster A regions, with reduced occupancy in cluster B regions (Fig. 5e, f; Supplementary Fig. 5h). Additionally, SUV39H1 occupancy increased at genes that gained H3K9me3 in TKO cells (Fig. 5g). These results demonstrate that SUV39H1 relocalizes to the same genomic loci where ectopic H3K9me3 accumulates in the absence of H3K36me.

To test whether SUV39H1 contributes to both normal and ectopic H3K9me3 deposition, we knocked down (KD) SUV39H1 using shRNAs (Supplementary Fig. 5i). In parental cells, SUV39H1-KD resulted in a global H3K9me3 reduction of approximately 40%, primarily within cluster B regions (Fig. 5h; Supplementary Fig. 5j). Notably, SUV39H1-KD in parental cells preferentially depleted the broad heterochromatic domains, as evidenced by a reduction in the proportion of H3K9me3 signal within cluster B and a concomitant increase in cluster A (Fig. 5h, i). In contrast, in TKO cells, SUV39H1-KD reduced global H3K9me3 levels to a similar extent, but the depletion was comparable across both cluster A and B regions (Fig. 5h, i; Supplementary Fig. 5j). These findings support that SUV39H1 is primarily responsible for maintaining broad heterochromatic domains in parental cells, whereas in the absence of H3K36me, it redistributes genome-wide and mediates ectopic H3K9me3 deposition.

To further demonstrate that SUV39H1 becomes enriched in euchromatic cluster A regions following H3K36me loss and contributes to ectopic H3K9me3 deposition, we examined H3K9me3 distribution following SUV39H1 overexpression (OE) in mMSCs. In parental cells, SUV39H1-OE led to a modest increase in H3K9me3 within cluster A, but enrichment remained primarily restricted to inactive cluster B regions and silent genes (Fig. 5h; Supplementary Fig. 5k). By contrast, in TKO SUV39H1-OE cells, H3K9me3 levels increased substantially across both clusters A and B, with cluster A regions—including genes that gained H3K9me3—becoming especially enriched (Fig. 5h; Supplementary Fig. 5k). Collectively, these findings reveal a previously underappreciated mechanistic dependency between the H3K36 and H3K9 methylation pathways. Loss of H3K36me redirects SUV39H1 from transcriptionally silent cluster B regions into euchromatic cluster A regions, where it contributes to the aberrant deposition of H3K9me3.

Downstream epigenetic cascade following the depletion of H3K9me3 decorated heterochromatin
The loss of broad H3K9me3 domains in H3K36me-deficient cells suggests a fundamental disruption of constitutive heterochromatin. Therefore, we next asked how the chromatin landscape is remodeled in their absence. In normal cells, following the deposition of H3K9me3, heterochromatin protein 1 (HP1) binds to the methylated histone tail and promotes chromatin compaction and gene silencing by recruiting additional repressive factors and facilitating the spread of heterochromatin48,53. To assess changes in HP1 binding, we performed ChIP-seq. As expected, in parental mMSCs, HP1 was predominantly enriched in inactive cluster B regions (Fig. 6a, b). However, in TKO mMSCs, following SUV39H1-mediated H3K9me3 deposition in cluster A regions (Fig. 5e, f), we observed corresponding recruitment of HP1 to these euchromatic domains (Fig. 6a, b; Supplementary Fig. 6a), consistent with established H3K9me3-HP1 binding dynamics. Notably, HP1 accumulation in these regions was particularly enriched within gene bodies and TEs (Fig. 6c; Supplementary Fig. 6b).

Conversely, in cluster B regions where H3K9me3 is depleted and HP1 binding is lost, we observed the invasive broadening of H3K27me3 (Fig. 6a, b). In parental mMSCs, large heterochromatic, gene-poor regions are characterized by “convex” H3K9me3 domains flanked by reciprocal “concave” domains of H3K27me3 (Supplementary Fig. 6c). These regions are transcriptionally inactive and lack open chromatin sites or other active epigenetic marks (Supplementary Fig. 6c). In the absence of H3K36 methylation in TKO cells, H3K9me3 is lost, and many of these regions become occupied by H3K27me3, possibly as a compensatory silencing mechanism (Fig. 6a, b; Supplementary Fig. 6c). Thus, in addition to invading gene-rich, active regions (cluster A), H3K27me3 also invades gene-poor, inactive regions that lose H3K9me3 (cluster B), suggesting a shift from constitutive to facultative heterochromatin (Fig. 6b; Supplementary Fig. 6c, d). Similarly, in Cal27 and Detroit562 H3K36M-OE cells, H3K27me3 spreads into both euchromatic cluster A regions and heterochromatic cluster B regions depleted of H3K9me3 (Supplementary Fig. 6e, f). In both cell lines, these chromatin changes are accompanied by a global increase in H3K27me3 (Supplementary Fig. 6g) and a corresponding decrease in H3K36me and H3K9me3 (Supplementary Fig. 5b).

Building on these observations, we next examined whether the loss of H3K9me3 and the invasion of H3K27me3 are also associated with changes in chromatin accessibility and regulatory potential. We observed the emergence of newly accessible regions, as measured by ATAC-Seq, within cluster B regions under H3K36me-deficient conditions. In mMSCs, we identified 1914 new open chromatin regions in TKO compared to parental cells (Fig. 6d, e). Interestingly, most of these newly accessible regions are substantially smaller than typical open chromatin peaks found at active promoters and enhancers (Fig. 6d), suggesting they may represent focal disruptions of chromatin compaction rather than canonical regulatory elements. Similarly, in Cal27 and Detroit562 H3K36M-OE cells, we observed new open chromatin peaks in regions that lose H3K9me3, with 1008 new peaks identified in Cal27 (Fig. 6f, g) and 481 new peaks in Detroit562 (Supplementary Fig. 6j). In TKO mMSCs, the gain in chromatin accessibility is accompanied by a significant increase in H3K4me1, and in some regions, H3K27ac (Fig. 6h). This accumulation of histone marks associated with enhancer-like chromatin further supports the global increase of H3K4me1 observed by mass spectrometry (Fig. 4c). Notably, in addition to its previously observed accumulation within gene bodies under H3K36me-deficient conditions (Fig. 4b), H3K4me1 also becomes enriched in regions that lose H3K9me3 (Fig. 6h), suggesting a broader redistribution of enhancer-associated marks into formerly heterochromatic compartments. Furthermore, these newly accessible regions exhibit reduced DNAme, consistent with a release from constitutive repression (Supplementary Fig. 6j). RNA-Seq analysis further reveals increased transcriptional activity within these regions (Fig. 6h). Similarly, in Cal27 and Detroit562 H3K36M-OE cells, the newly accessible chromatin regions exhibit elevated H3K27ac levels and increased transcriptional output (Fig. 6i; Supplementary Fig. 6k), reinforcing that the loss of H3K36me triggers a conserved epigenomic response across cell types—one that includes the collapse of many constitutive heterochromatin domains and the acquisition of regulatory features characteristic of active chromatin.

Together, these findings reveal that loss of H3K36me leads to widespread epigenomic restructuring, characterized by the erosion of constitutive heterochromatin and the emergence of enhancer-like features in normally silenced regions. This includes a coordinated gain of chromatin accessibility, activation-associated histone marks, and transcriptional de-repression. These results underscore a critical role for H3K36me, and/or the respective K36MTs, in preserving heterochromatin integrity and highlight the remarkable plasticity of silent chromatin in response to disruptions to the balance of histone methylation marks.

H3K9me3 loss drives transcriptional activation despite compensatory H3K27me3 accumulation
Despite the compensatory accumulation of H3K27me3 in cluster B regions following H3K9me3 loss, these repressive changes appear insufficient to fully maintain transcriptional silencing. Partial correlation analysis reveals that although H3K27me3 levels increased within cluster B regions, this gain did not offset transcriptional activation and was not significantly associated with gene expression changes (Supplementary Fig. 7a). By contrast, H3K9me3 loss exhibited a strong correlation with gene upregulation in cluster B regions (Supplementary Fig. 7a), implicating this mark as the primary regulator of transcriptional changes at these loci. We next sought to characterize the genes undergoing H3K9me3 depletion. In TKO mMSCs, approximately 90% (2119/2354) of genes in these regions lose H3K9me3 (Fig. 7a). Among these, genes with the most pronounced loss of H3K9me3 become upregulated (157/179 genes) (Fig. 7b). A similar pattern is observed in Cal27 and Detroit562 H3K36M-OE cells, where genes within cluster B domains lose H3K9me3 and are significantly upregulated relative to an expression-matched random control gene set (Fig. 7a; Supplementary Fig. 7b).

To better understand the biological consequences of H3K9me3 depletion from cluster B regions, we next examined the subset of genes that lose H3K9me3 in both mMSC and HNSCC models to determine whether common regulatory targets or pathways are affected across species. Among the genes that lost H3K9me3 in cluster B regions in both mMSC TKO and Cal27 H3K36M-OE cells (Fig. 7a), 132 genes were shared across the two models (Fig. 7c). Remarkably, approximately 35% (46/132) of these genes encode olfactory receptors (ORs) (Fig. 7c), consistent with prior findings that OR gene clusters are densely marked by H3K9me3 and subject to tight epigenetic repression in the mouse genome54. Indeed, KEGG55 pathway analysis of the 132 genes indicates olfactory transduction to be the most significantly overrepresented pathway (Fig. 7d). Complementary Gene Ontology (GO)56 analysis confirmed this pattern, with smell-related terms ranking in the top enriched categories (Supplementary Fig. 7c). In addition to ORs, another gene family significantly enriched among the shared H3K9me3-depleted loci are gamma-aminobutyric acid (GABA) receptor genes (Fig. 7c, d; Supplementary Fig. 7c). Although traditionally associated with neuronal function, GABA receptor genes are increasingly recognized for their broader roles in cellular signaling, and there is emerging evidence that H3K9me3 contributes to their transcriptional silencing57. Notably, GABA receptor activity influences the expression of brain-derived neurotrophic factor (BDNF), which itself is negatively regulated by H3K9me3 in mouse models58.

In mMSCs, numerous TEs located in cluster B regions also become differentially expressed, with a strong bias toward upregulation (844 upregulated versus 86 downregulated) (Fig. 7e). Further analysis reveals that 44 TE subfamilies are upregulated, compared to only 9 that are downregulated, with the majority of upregulated TEs belonging to the endogenous retrovirus (ERV) family (Supplementary Fig. 7d). Interestingly, this widespread dysregulation of TEs is not observed in Cal27 or Detroit562 cells (data not shown). This discrepancy may reflect fundamental species-specific differences in TE content and regulation. The mouse genome harbors a higher proportion of young and transcriptionally competent ERVs, which rely heavily on H3K9me3 and SETDB1 for silencing, particularly in stem and progenitor cellular contexts59,60. In contrast, the majority of ERVs in the human genome are older, more degenerate, and often rendered transcriptionally inactive by fixed mutations, DNA methylation, or Polycomb-mediated H3K27me3, thereby reducing their reliance on H3K9me3 for repression61–63. These findings suggest that while gene de-repression following the loss of H3K9me3 is conserved across species, TE activation is a more prominent feature in murine cells.

Taken together, these results demonstrate that the loss of H3K9me3 contributes to focal derepression of select gene families and transposable elements in a species- and context-specific manner, and that the compensatory accumulation of H3K27me3 in cluster B regions is insufficient to restore transcriptional silencing.

H3K36me is essential for maintenance of nuclear architecture and long-range chromatin interactions
At a fundamental level, chromatin is spatially organized into active and inactive compartments, typically referred to as A and B compartments, respectively64. Given the widespread alterations in both euchromatic and heterochromatic histone modifications following H3K36me depletion, we examined whether chromatin conformation and nuclear compartmentalization were similarly affected. To test this, we performed Hi-C on parental and QuiKO mMSCs, which lack all H3K36me, as well as on parental and H3K36M-OE Cal27 and Detroit562 cells.

Parental mMSCs have distinct A/B compartments and euchromatin/heterochromatin structure, as illustrated by the strong checkerboard pattern at the chromosomal scale (Fig. 8a; upper diagonal). The inactive B compartments are highly correlated with H3K9me3 domains (Supplementary Fig. 8a) and display significant B-B compartment interactions (Fig. 8b), indicative of strong heterochromatin compaction. Notably, the frequency of B-B interactions is nearly double that of A-A interactions in parental cells (Fig. 8b). In contrast, QuiKO cells exhibit a drastic disruption in chromatin compartmentalization, coinciding with the loss of H3K9me3 domains (Fig. 8a; lower diagonal). This is reflected by a twofold reduction in both B-B and A-A interactions (Fig. 8b) and a homogenization of compartment scores resulting in the collapse of the bimodal nature of compartment identity, particularly affecting B-compartments (Fig. 8c). A similar loss of compartment integrity is observed in human HNSCC cell lines. In Cal27 and Detroit562, parental cells exhibit distinct A/B compartmentalization (Fig. 8d; upper diagonal and Supplementary Fig. 8b; upper diagonal), which becomes attenuated upon H3K36M-OE (Fig. 8d; lower diagonal and Supplementary Fig. 8b; lower diagonal). In Cal27 H3K36M-OE cells, B-B interactions decrease threefold, while A-A interactions are reduced by ~30% (Fig. 8b; second column). Similarly, Detroit562 H3K36M-OE cells show an ~40% reduction in B-B interactions and ~25% reduction in A-A interactions (Supplementary Fig. 8c). In both HNSCC cell lines, compartment scores shift toward zero, indicating a global weakening of compartmental identity (Fig. 8c; Supplementary Fig. 8d).

Across all three cell types, the total number of A and B compartments remains largely unchanged between parental and H3K36me-depleted conditions, although a modest shift from A to B compartments is observed (Fig. 8e; Supplementary Fig. 8e). Nonetheless, a more detailed analysis reveals that in mMSCs, long-range cis-interactions (>10 Mb) are particularly diminished (Supplementary Fig. 8f), suggesting weakened chromosomal compartmentalization at extended genomic distances. To evaluate additional aspects of genome architecture, we assessed topologically associating domains (TADs) and chromatin loops. Interestingly, the number and strength of TAD boundaries remain largely unaffected across all conditions and cell types (Supplementary Fig. 8g, h). However, both QuiKO and H3K36M-OE cells exhibit a greater than twofold reduction in the number of chromatin loops compared to their respective parental controls (1448 versus 701 in mMSC; 2379 versus 1037 in Cal27; 497 versus 367 in Detroit562) (Supplementary Fig. 8i). Notably, this reduction is more pronounced for long-range loops (>500 kb) than for short-range loops across all cell lines (Supplementary Fig. 8j). To assess whether disruptions to chromatin loops impact enhancer activity and target gene expression, we identified annotated enhancers located at the anchors of the affected loops, determined their corresponding target genes, and quantified changes in gene expression. Notably, we found that genes associated with these disrupted loops were preferentially downregulated (Supplementary Fig. 8k). Overall, these data demonstrate that H3K36me depletion leads to profound alterations in 3D genome architecture, characterized by a loss of long-range compartmental interactions, attenuation of heterochromatin compaction, and a marked reduction in chromatin loops—particularly those spanning large genomic distances. The preferential downregulation of genes linked to disrupted loops suggests that impaired chromatin topology may be associated with transcriptional dysregulation.

To visualize these structural alterations at the microscopic level, we performed immunofluorescence (IF) to examine the subnuclear distribution of H3K9me3 (Supplementary Fig. 8l). In parental mMSCs, H3K9me3 localizes to numerous discrete nuclear foci, likely corresponding to large heterochromatic domains (Fig. 8f; Supplementary Fig. 8l). Many of these foci are enriched at the nuclear periphery (Fig. 8f), consistent with interactions between heterochromatin and the nuclear lamina, a hallmark of spatial genome organization65,66. Similarly, Cal27 parental cells predominantly exhibit peripherally localized H3K9me3 foci (Fig. 8g). In contrast, QuiKO mMSCs show a markedly diffuse distribution of H3K9me3 with fewer defined foci and significantly reduced peripheral localization (Fig. 8f, h, i; Supplementary Fig. 8l). Instead, residual H3K9me3 signal is redistributed toward the nuclear interior. Notably, while H3K9me3 continues to co-localize with DAPI, which preferentially stains AT-rich, heterochromatic DNA, the DAPI signal itself also shifts toward the center of the nucleus in QuiKO cells (Fig. 8f), further supporting a large-scale reorganization of nuclear architecture. Similar alterations are observed in Cal27 H3K36M-OE cells (Fig. 8g–i; Supplementary Fig. 8m), indicating that this effect is conserved across species and cellular contexts.

Collectively, these results demonstrate that the loss of H3K36 methylation not only perturbs euchromatin but also fundamentally disrupts the spatial organization and compartmentalization of heterochromatin. The displacement of H3K9me3 from the nuclear periphery toward the interior may reflect a collapse in nuclear lamina-heterochromatin interactions, possibly contributing to the loss of long-range chromatin contacts observed by Hi-C. Given the central role of 3D genome organization in maintaining gene silencing and genome stability, our results suggest that H3K36 methylation is an upstream regulator of both chromatin topology and nuclear compartmentalization.

Methods
Cell culture
C3H10T1/2 mouse mesenchymal stem cells (ATCC) were cultured in DMEM (1X) (Invitrogen) with 10% fetal bovine serum (FBS) (Wisent) and supplemented with 1% GlutaMax and penicillin-streptomycin. C3H10T1/2 cell lines harboring individual knockouts of NSD1, NSD2, NSD3, SETD2, and ASH1L, as well as the NSD1/2/3-SETD2-QKO and NSD1/2/3-SETD2-ASH1L-QuiKO cell lines were previously established by our group using CRISPR-Cas9 RNP-mediated gene editing in Shipman and Padilla et al.26. C3H10T1/2 cells overexpressing the H3K36M mutation were provided by the lab of Dr. Chao Lu (Stanford University), and C3H10T1/2 NSD1/2-DKO and C3H10T1/2 NSD1/2-SETD2-TKO cell lines were provided by the lab of Dr. C. David Allis (Rockefeller University). Cal27 (ATCC, CRL-2095) and Detroit562 (ATCC, CCL-138) HNSCC cells were cultured in DMEM (F12) (Invitrogen) with 10% FBS and penicillin-streptomycin. HEK293T cells were cultured in DMEM (Invitrogen) supplemented with 10% FBS, 1% GlutaMax and penicillin-streptomycin. Drosophila S2 cells were cultured in Schneider’s Drosophila medium (Thermo Fisher) containing 10% FBS. All cell lines tested negative for mycoplasma contamination.

Generation of Cal27 and Detroit562 HNSCC cells overexpressing the H3K36M mutation
Epitope-tagged H3.3 was previously cloned into pCDH-EF1-MCS-Puro lentiviral vector and site-directed mutagenesis techniques were used to generate the H3K36M mutation14. The H3.3K36M pCDH-EF1-MCS-Puro construct was provided by Dr. Chao Lu (Stanford University). To produce lentivirus, 293T cells were transfected with the lentiviral vector containing H3.3K36M and helper plasmids (psPAX2 and pMD2.G). Supernatant containing lentivirus was collected after 48 h, filtered and used to infect Cal27 and Detroit562 cells. Transduced cells were grown under puromycin selection (1 μg/ml) for 48 h after transduction.

Generation of stable C3H10T1/2 SUV39H1 knockdown cell lines
To establish stable knockdown of SUV39H1, a collection of five different mouse SUV39H1 shRNA plasmids were obtained from the McGill University Life Science Complex High-Throughput Screening Facility (Sigma-Aldrich Mission shRNAs: TRCN0000097439, TRCN0000097440, TRCN0000097441, TRCN0000097442 and TRCN0000097443) (Supplementary Table 1). Lentiviral particles were produced by transiently transfecting individual lentiviral SUV39H1 or control shRNA vectors together with the helper plasmids psPAX2 and pMD2.G at a ratio of 5:4:1 respectively in HEK293T cells. Briefly, HEK293T cells were grown to 70% confluence in 60 mm cell culture dishes prior to transfection. Cells were transfected with the TransIT-LT1 transfection reagent following the manufacturer’s protocol. The lentivirus-containing supernatants were collected and filtered 72 h later using a 0.45 μm syringe filter and stored at –80°C. To maximize the knockdown of SUV39H1 expression, C3H10T1/2 parental and NSD1/2-SETD2 TKO cells were transduced simultaneously with all five SUV39H1 shRNA lentiviruses. Cells were grown to 70% confluence in a well of a 6 well plate prior to infection. Cells were infected in DMEM complete media supplemented with 5 mg/ml of polybrene and infected using 125 μl of pooled lentiviral particles. Media was refreshed 24 hours post-infection, and the cells were allowed to grow for an additional 48 hours prior to selection. The infected cells were selected with puromycin (1 µg/ml) for 7 days, and all surviving cells were pooled to generate stable cell lines with SUV39H1 knockdown. SUV39H1 expression was evaluated by Western blotting using an anti-SUV39H1 antibody (Supplementary Table 2).

Western blots
For each sample, 1 million cells were collected and counted using an automated Countess counter. Cell pellets were washed using PBS and resuspended in 100 ul of 1X RIPA buffer, 0.1 mM proteinase inhibitors cocktail, 0.1 mM PMSF, and incubated on ice for 1 hour. Samples were centrifuged for 10 minutes at 21,000 x g. Supernatants were transferred into new tubes. Protein quantification was performed using the BCA-Pierce Protein assay Thermo Scientific/Pierce (Cat# 23227) and the Infinite 200 Pro Tecan-iControl. For each sample, 40 µg of protein was mixed with 1X Laemmli buffer (0.35 M Tris HCl pH 6.8, 30% Glycerol, 10% SDS, 20% Beta-mercaptoethanol, 0.04% Bromophenol blue, H2O) and loaded into stain-free TGX 4%–15% gradient pre-cast gels (Bio-Rad, Cat# 4568084), and All-Blue (Bio-Rad, Cat# 1610373) and Unstained (Bio-Rad, Cat# 1610363) protein standards were mixed in equal amounts and loaded. Semi-dry electrotransfer onto a PVDF membrane was completed using the Trans-blot Turbo Transfer system (Bio-Rad, Trans-blot® Turbo RTA Mini LF PVDF Transfer Kit, 1704274). PVDF membranes were blocked with 5% BSA in TBSt, and probed overnight with the respective primary antibody (Supplementary Table 2). Following washes with TBSt, membranes were incubated for 1 hour with horseradish peroxidase-conjugated secondary antibody (anti-rabbit or goat anti-mouse) in 2% BSA-TBSt. Imaging was completed using the ECL Clarity or Clarity Max solutions (Bio-Rad, Cat# 1705060 and 1705062).

Crosslinking and Histone PTM ChIP-seq
Approximately 20 million cells per cell line were used. Crosslinking was performed in 150 mm cell culture plates using 1% formaldehyde (Sigma) at room temperature with gentle rocking for 10 minutes. The crosslinking reaction was quenched using 1.25 M Glycine with gentle rocking at room temperature for 5 minutes. Fixed cell preparations were washed with ice-cold PBS, scraped, and then washed twice more with ice-cold PBS. Crosslinked pellets were resuspended in 500 μl cell lysis buffer (5 mM pH 8.5 PIPES, 85 mM KCl, 1% (v/v) IGEPAL CA-630, 50 mM NaF, 1 mM PMSF, 1 mM phenylarsine oxide, 5 mM sodium orthovanadate, EDTA-free protease inhibitor tablet) and incubated for 30 minutes on ice. Samples were centrifuged at 12,000 x g for 10 minutes at 4°C, and pellets were resuspended in 500 μl of nuclei lysis buffer (50 mM pH 8.0 Tris-HCl, 10 mM EDTA, 1% (w/v) SDS, 50 mM NaF, 1 mM PMSF, 1 mM phenylarsine oxide, 5 mM sodium orthovanadate, EDTA-free protease inhibitor tablet) and then incubated for 30 minutes on ice. Sonication of lysed nuclei was performed using the BioRuptor UCS-300 at maximum intensity for 75-90 cycles (10s on, 20s breaks). Sonication efficiency to achieve fragments between 150-500 bp was evaluated using gel electrophoresis on a reversed-crosslinked and purified aliquot from each sample. After sonication, chromatin was diluted to reduce SDS levels to 0.1% and concentrated using Nanosep 10K OMEGA (Pall) columns. Prior to the chromatin immunoprecipitation (ChIP) reaction, 2% sonicated Drosophila S2 cell chromatin was spiked into each sample for quantification of total levels of histone marks. The ChIP reactions were performed using the Diagenode SX-8G IP-Star Compact and Diagenode automated iDeal ChIP-seq Kit for Histones. Dynabeads Protein A or G (Invitrogen) were washed, then incubated with specific antibodies (Supplementary Table 2), 1.5 million cells of sonicated cell lysate, and protease inhibitors for 10 hrs, followed by a 20 min wash cycle using the provided wash buffers (Diagenode Immunoprecipitation Buffers, iDeal ChIP-seq Kit for Histones). Reverse cross-linking was then performed using 5 M NaCl at 65°C for 4 h. ChIP samples were then treated with 2 μl RNase Cocktail at 65°C for 30 min followed by 2 μl Proteinase K at 65°C for 30 min. Samples were then purified with QIAGEN MinElute PCR purification kit (QIAGEN) as per the manufacturer’s protocol. In parallel, input samples (chromatin from about 50,000 cells) were reverse crosslinked and DNA was isolated following the same protocol. Library preparation was performed using the Kapa Hyper Prep library preparation reagents following the manufacturer’s protocol (Kapa Hyper Prep Kit, Roche 07962363001). ChIP libraries were sequenced by the McGill University Genome Centre using the Illumina HiSeq 4000 at 50 bp single reads or NovaSeq 6000 at 100 bp single reads.

Crosslinking and protein ChIP-seq
Approximately 40 million cells per cell line were used. Crosslinking was performed in 150 mm cell culture plates using 1% formaldehyde (Sigma) at room temperature with gentle rocking for 10 minutes. The crosslinking reaction was quenched using 1.25 M Glycine with gentle rocking at room temperature for 5 minutes. Fixed cell preparations were washed with ice-cold PBS, scraped, and then washed twice more with ice-cold PBS. Crosslinked pellets were resuspended in 500 μl cell lysis buffer (5 mM pH 8.5 PIPES, 85 mM KCl, 1% (v/v) IGEPAL CA-630, 50 mM NaF, 1 mM PMSF, 1 mM phenylarsine oxide, 5 mM sodium orthovanadate, EDTA-free protease inhibitor tablet) and incubated for 30 minutes on ice. Samples were centrifuged at 12,000 x g for 10 min at 4°C, and pellets were resuspended in 500 μl of nuclei lysis buffer (50 mM pH 8.0 Tris-HCl, 10 mM EDTA, 1% (w/v) SDS, 50 mM NaF, 1 mM PMSF, 1 mM phenylarsine oxide, 5 mM sodium orthovanadate, EDTA-free protease inhibitor tablet) and then incubated for 30 minutes on ice. Sonication of lysed nuclei was performed using the BioRuptor UCS-300 at maximum intensity for 75–90 cycles (10s on, 20s breaks). Sonication efficiency to achieve fragments between 150–500 bp was evaluated using gel electrophoresis on a reversed-crosslinked and purified aliquot from each sample. After sonication, chromatin was diluted to reduce SDS levels to 0.1% and concentrated using Nanosep 10K OMEGA (Pall) columns. Prior to the chromatin immunoprecipitation (ChIP) reaction, 2% sonicated Drosophila S2 cell chromatin was spiked into each sample for quantification. For each IP, Dynabeads Protein A or G (Invitrogen) were washed with RIPA buffer and then resuspended in RIPA buffer. Specific antibodies were added (Supplementary Table 2) and then incubated at 4°C for at least 3 hours on a rotator. Following antibody conjugation, Dynabeads were washed with RIPA buffer, and then resuspended in RIPA buffer, ~5 million cells of sonicated cell lysate, and protease inhibitors, followed by overnight incubation on a rotator at 4°C. Four successive washes were then performed using 1. RIPA buffer; 2. RIPA buffer with 500 mM NaCl (1% (v/v) Triton X-100, 0.10% SDS, 500 mM NaCl, 20 mM Tris-HCl pH 8.0, 2 mM EDTA, H2O); 3. LiCl buffer (1% NP-40, 250 mM LiCl, 10 mM Tris-HCl pH 8.0, 1 mM EDTA, H2O); 4. TE buffer (50 mM NaCl, 10 mM Tris-HCl pH 8.0, 1 mM EDTA, H2O). Samples were then resuspended in Elution buffer (50 mM Tris-HCl pH 8.0, 10 mM EDTA, 1% (v/v) SDS, H2O) and incubated at room temperature for 30 minutes. Reverse cross-linking was then performed using 5 M NaCl at 65°C for 4 h. ChIP samples were then treated with 2 μl RNase Cocktail at 65°C for 30 min followed by 2 μl Proteinase K at 65°C for 30 min. Samples were then purified with QIAGEN MinElute PCR purification kit (QIAGEN) as per the manufacturer’s protocol. In parallel, input samples (chromatin from about 50,000 cells) were reverse crosslinked and DNA was isolated following the same protocol. Library preparation was performed using the Kapa Hyper Prep library preparation reagents following the manufacturer’s protocol (Kapa Hyper Prep Kit, Roche 07962363001). ChIP libraries were sequenced by the McGill University Genome Centre using the Illumina HiSeq 4000 at 50 bp single reads or NovaSeq 6000 at 100 bp single reads.

CUT&RUN
CUT&RUN reactions were performed with the Epicypher CUTANA ChIC/CUT&RUN Kit Version 3 (cat# 14-1048) using 550,000 fresh C3H10T1/2 cultured cells and following the manufacturer’s protocol (Epicypher User Manual Version 3.5) for adherent cells. Prior to performing CUT&RUN, optimization of C3H10T1/2 cell permeabilization was performed following Appendix 1.1 (Epicypher User Manual Version 3.5). Cell permeabilization was found to be most optimal using a final working concentration of 0.01% Digitonin (Promega). Successful binding of cells to Concanavalin A beads was confirmed as recommended and outlined by the manufacturer. Manufacturer supplied H3K4me3 positive control and IgG negative control antibodies were used, and the reactions were performed in parallel with processed samples. Processed samples were incubated with specific antibodies (Supplementary Table 2). Manufacturer also supplied E.coli spike-in DNA, which was used in each reaction for sequencing normalization. Quantification of purified CUT&RUN DNA was performed using the Qubit fluorometer with the 1X dsDNA HS Assay Kit (Invitrogen). Library preparations were performed using the Epicypher CUTANA CUT&RUN Library Prep Kit Version 1 (cat# 14-1001) following the manufacturer’s protocol. Final CUT&RUN library concentrations were quantified as previously described, and enrichment of mononucleosome-sized fragments was assessed by Bioanalyzer (Agilent) prior to sequencing. CUT&RUN libraries were sequenced at a depth of 15 M reads per sample by the McGill University Genome Centre using the Illumina NovaSeq 6000 at 100 bp paired-end reads.

ATAC-Seq
ATAC-Seq library preparation was performed according to the Omni-ATAC protocol72. 50,000 C3H10T1/2 cultured cells were resuspended in 1 ml of cold ATAC-seq resuspension buffer (RSB; 10 mM Tris-HCl pH 7.4, 10 mM NaCl, and 3 mM MgCl2 in water). Cells were centrifuged at 500 x g for 5 min in a pre-chilled (4°C) fixed-angle centrifuge. After centrifugation, supernatant was aspirated and cell pellets were then resuspended in 50 μl of ATAC-seq RSB containing 0.1% IGEPAL, 0.1% Tween-20, and 0.01% digitonin by pipetting up and down three times. This cell lysis reaction was incubated on ice for 3 min. After lysis, 1 ml of ATAC-seq RSB containing 0.1% Tween-20 (without IGEPAL and digitonin) was added, and the tubes were inverted to mix. Nuclei were then centrifuged for 10 min at 500 x g in a pre-chilled (4°C) fixed-angle centrifuge. Supernatant was removed and nuclei were resuspended in 50 uL transposition mix (2x TD Buffer, 100 nM final transposase, 16.5 uL PBS, 0.5 uL 1% digitonin, 0.5 uL 10% Tween-20, 5 uL H2O) (Illumina Tagment DNA Enzyme and Buffer Small Kit, 20034197). Transposition reactions were incubated at 37°C for 30 min in a thermomixer with shaking at 1000 rpm. Reactions were cleaned up with DNA Clean and Concentrator 5 columns (Zymo Research). Illumina Nextera DNA Unique Dual Indexes Set C (Illumina, 20027215) were added and amplified (12 cycles) using NEBNext 2x Master Mix. Sequencing of the ATAC-Seq libraries was performed on the Illumina NovaSeq 6000 system using 100-bp paired-end sequencing.

RNA-Seq
Total RNA was extracted from approximately 1 million cells using the AllPrep DNA/RNA/miRNA Universal Kit (QIAGEN) and following the manufacturer’s protocol, including DNase treatment option. Library preparation was performed using the NEBNext® rRNA Depletion Kit v2 (Human/Mouse/Rat) (New England Biolabs) following the manufacturer’s protocol (including RNase H and DNase I digestions). 100-bp paired-end sequencing was performed on the Illumina HiSeq 4000 or NovaSeq 6000 platform.

Histone acid extraction, histone derivatization, and analysis of post-translational modifications by nano-LC-MS
4 million cells from each cell line were collected and frozen at -80°C. Thawed pellets were lysed in nuclear isolation buffer (15 mM Tris pH 7.5, 60 mM KCl, 15 mM NaCl, 5 mM MgCl2, 1 mM CaCl2, 250 mM sucrose, 10 mM sodium butyrate, 0.1% (v/v) beta-mercaptoethanol, commercial phosphatase and protease inhibitor cocktail tablets) containing 0.3% NP-40 alternative on ice for 5 min. Nuclei were subsequently washed twice in the same buffer without NP-40, and pellets were resuspended using gentle vortexing in chilled 0.4 N H2SO4, followed by a 3 h incubation while rotating at 4°C. After centrifugation, supernatants were collected and proteins were precipitated in 20% TCA overnight at 4°C, washed with 0.1% HCl (v/v) acetone once, followed by two washes with acetone alone. Histones were resuspended in deionized water. Acid-extracted histones (20 μg) were resuspended in 50 mM ammonium bicarbonate (pH 8.0), derivatized using propionic anhydride and digested with trypsin as previously described (15). After the second round of propionylation, the resulting histone peptides were desalted using C18 StageTips, dried using a centrifugal evaporator and reconstituted using 0.1% formic acid in preparation for LC-MS analysis. Nanoflow liquid chromatography was performed using a Thermo Fisher Scientific Vanquish Neo UHPLC equipped with an Easy-Spray™ PepMap™ Neo nano-column (2 µm, C18, 75 µm X 150 mm). Buffer A was 0.1% formic acid and Buffer B was 0.1% formic acid in 80% acetonitrile. Peptides were resolved using a room temperature with a mobile phase consisting of a linear gradient from 1 to 45% solvent B (0.1% formic acid in 100% acetonitrile) in solvent A (0.1% formic acid in water) over 85 mins and then 45 to 98% solvent B over 5 min at a flow rate of 300 nL/min. The HPLC was coupled online to an Orbitrap Exploris 240 (Thermo Scientific) mass spectrometer operating in the positive mode using a Nanospray Flex Ion Source (Thermo Fisher Scientific) at 1.9 kV. A full MS scan was acquired in the Orbitrap mass analyzer across 350–1050 m/z at a resolution of 120,000 in positive profile mode with an auto maximum injection time and an AGC target of 300%. Parallel reaction monitoring experiments were followed for monitoring the targeted peptides based on the inclusion list. Targeted ions were fragmented using HCD fragmentation. These scans typically used an NCE of 30, an AGC target standard, and an auto maximum injection time. Raw files were analyzed using EpiProfile 2.073 and Skyline74. The area for each modification state of a peptide was normalized against the total signal for that peptide to give the relative abundance of the histone modification. Calculated MS ratios (Supplementary Table 3).

Whole genome bisulfite sequencing
Whole genome sequencing libraries were generated from 1000 ng of genomic DNA extracted from approximately 4 million cells and fragmented to 300–400 bp using the Covaris focused-ultrasonicator E210. The NxSeq AmpFREE Low DNA Library Kit (Lucigen-CA10837-144) was then used. End repair, A-tailing, adapter ligation, and clean-up steps were performed as per the manufacturer’s recommendations. Bisulfite conversion was performed using the EZ-96 DNA Methylation-Gold™ MagPrep Kit (Zymo Research) according to the manufacturer’s protocol. Bisulfite-converted DNA was amplified by PCR using the KAPA Hifi HotStart Uracil+ Kit (Roche) according to the manufacturer’s protocol. The amplified libraries were then purified using AMPure XP Beads (Beckman Coulter). 150-bp paired-end sequencing of the WGBS libraries was performed using the Illumina HiSeq X system.

Hi-C
For each sample, two million cells were crosslinked and processed using the Arima-HiC+ kit and following the Arima Genomics HiC protocol for Mammalian Cell Lines (A510008). Proximally-ligated DNA was fragmented to an average size of 400 bp using the BioRuptor UCS-300 at low intensity for 10 cycles (30s ON, 90s OFF). Library preparation of fragmented proximally-ligated DNA was performed using the NEBNext® Ultra™ II DNA Library Prep Kit (E7645S), NEBNext Multiplex Oligos for Illumina (E6440S), and the Arima HiC+ Kit according to manufacturer protocols. Hi-C libraries were sequenced by the McGill University Genome Centre at 150 bp paired-end reads using the Illumina NovaSeq 6000.

Immunofluorescent staining and analysis
Cells were seeded on glass coverslips until between 60-80% confluent and fixed with a cold solution of methanol:acetone at 80:20 ratio for 15 minutes. The samples were blocked 1 h with 5% skim milk followed by 1 h with rabbit anti-H3K9me3 antibody (Supplementary Table 2) at 1:400 dilution and 1 h with Alexa Fluor-488 goat anti-rabbit secondary antibody, all in blocking solution. Samples were then counterstained with 4′,6-diamidino-2-phenylindole (DAPI) solution in PBS for 5 min. All the different treatments were separated by 3 washes with PBS. The coverslips were inverted onto permafluor mountant (epredia) on microscope slides, viewed and imaged on confocal LSM800 at the McGill University Advanced BioImaging Facility (ABIF). Z-stacks of 10 slices were taken, and an image projection (average setting) was built using Fiji75 v.2.15.1. All images were taken under the same conditions, but laser strength and exposure were slightly adjusted for each image so that there is no saturation and to ensure similar fluorescence intensity in each image. This allowed a proper comparison of the spatial distribution of the foci. Images were filtered using ImageJ76 v.1.54 for any nuclei that were dividing and any nuclei that were cut off at the border of the images. Analysis was performed using the open-source software CellProfiler77 v.4.2.6. A CellProfiler pipeline was designed to detect and count nuclei as well as the number of H3K9me3 foci within each nucleus using the same parameters for all images. For detecting nuclei, a manual thresholding method was applied with a threshold of 0.015 and only nuclei with a diameter from 50 to 350 were included. 130 nuclei per condition was included for analysis in mMSCs whereas 103 nuclei per condition was included in Cal27. For detecting the number of H3K9me3 foci within each nucleus, a manual thresholding method was applied with a threshold of 0.1, a smoothing filter of 11 and a minimum allowed distance of 6 (which distinguishes between clumped foci). The number of foci per nucleus in the H3K36me-deficient cells was normalized by multiplying the foci count per nucleus by the ratio of the mean surface area of the parental nuclei to the mean surface area of the H3K36me-deficient nuclei. Specifically, the foci count per nucleus in the QuiKO cells were normalized by the ratio of the mean nuclei surface area of the parental cells (13747.99 pixels) to that of the QuiKO cells (14394.36 pixels). Similarly, the foci count per nucleus in the H3K36M-OE cells were normalized by the ratio of the mean nuclei surface of the parental cells (6588.77 pixels) to that of the H3K36M-OE cells (6109.94 pixels). Lastly, the average distance of foci to the center of each nucleus was normalized by dividing by the radius.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Padilla et al studied effect of combined knock-out (KO) of the H3K36 lysine methyltransferases (KMTases) NSD1-3, ASHL1 and SETD2 on genome-wide distribution of H3K36me2, H3K27me1-3 and H3K9me3. They found H3K36me2 marked enhancers. Reduction of H3K36me2 reduced expression of some proportion of genes that supports the role H3K36 methylation on enhancer regulation of promoters and gene expression. Within the genic region, authors studied enrichment of H3K27me in the various KO mutants and reported H3K36me2 prohibit H3K27me2/3 accumulation. They also showed redistribution of H3K9me3 from heterochromatin to genic regions. Finally using Hi-C, they showed significant changes in chromatin compartmentalization associated with the loss of H3K36me2/3. The work is extensive and show interesting connection of H3K36me2 with higher order chromatin organization. If the work can be more connected and mechanistically explored, it would be a significant finding in the field.\nMajor Comments:\n1. The work currently composed of mostly three weakly connected parts: enhancer, H3K9me3 on heterochromatin and Hi-C compartmentalization study. More work needs to be done to flow these parts into a complete story.\n2. It is important to ensure the actual endogenous status of individual KMTases as their names imply (DKO, TKO, QKO and QuiKO). The authors should show immunoblots to demonstrate the level of knockdown of all individual KMTases (NSD1-3, SETD2 and ASH1L) in single KOs, DKO, TKO, QKO and QuiKO to demonstrate that each gene knockout did not ectopically change the levels of the other H3K36 KMTases and that the level of the unaffected KMTases are similar between different mutants.\n3. H3K36me profiles of single KMTase knockouts should be included as controls in Fig1a.\nProfile of H3K27ac should be included in bottom panel of QKO results in Fig 1a.\n4. In Figs 1b-c, the decrease in accessibility (20-26%), H3K27ac (30-40%) and increase in H3K27me3 (20-54%) were partial and significant amount of these still retained on the enhancers in the KO. The authors need to obtain quantitative functional evidence that such residual level is insufficient to support a full activity of these enhancers and the amount of enhancer activity is reduced in QKO relative to TKO as they expected.\n5. ChIP-seq of core and linker histones (H1, H2A, H2B, H3 and H4) should be performed to compare between TKO/QKO/QuiKO in Fig 1b-c and 2e-f to assess whether nucleosomal positioning and occupancy are affected by the KO.\n6. Fig 1e showed approx. 3x more genes upregulated than downregulated in QKO contradictory to what is expected from compromised enhancer function and authors attributed to “secondary, downstream effects from global H3K36me2 depletion and/or ablation of NSD3”. It is unclear why this was listed as the possible reason. Considering that H3K36me2 coordinates transcriptional elongation; its depletion is expected to downregulate transcription. It becomes even more confusing when the authors obtained a different number of differentially expressed genes by considering H3K36me2 enrichment. What is the proportion of genes that overlap between these two lists? Are all the 97 downregulated genes in Fig 1e found within that 150 in Fig 1f? The authors should explain why there is such a big discrepancy between these results.\n7. The authors referred to the function and regulation of the KMTases, but none of these proteins were studied in this work. Localization of the H3K36-KMTase proteins should be done in non- and KO cells, relative to the localization of PRC2 and SUV3-9 methyltransferase. Line 218 says “…demonstrating that NSD3 is recruited to these regions”. However, there is no evidence to support such claim, and authors should provide ChIP-seq data of NSD3 in TKO versus QKO. For this matter, the authors should perform whole genome ChIP-seq of the KMTases (NSD1-3, SETD2, ASH1L) in their PA and KO cells to correlate with the progressive loss of H3K36me2.\n8. SETD2 is mostly a H3K36 trimethylase. KO of SETD2 is expected to be opposite to the other K36 dimethylases (NSD1-3, ASH1L) by increasing the levels of H3K36me2 (instead of decreasing). If so, then TKO, QKO and QuiKO would contain higher H3K36me2 at some loci to confound and mask the effect of the dimethylases KO. The authors should show experimental evidence that this possibility is not the case, or at least do not significantly confound the effects of the dimethylases KO.\n9. The authors should perform whole genome analyses of RNA polymerase II as an independent confirmation of transcriptional changes. Does the invasion of H3K27me3 and H3k9me3 into the gene body correlate with the exclusion of RNA polymerase II from those localities?\n10. Why do enhancers retain H3K36me2 so much stronger over the other regions? Is this due to higher NSD3 localization at the enhancers? Do these enhancers contain sequence motifs to preferentially recruit NSD3? Alternatively, would NSD3 be bound preferentially to H3K27ac or H3K4me1 that are enriched at the enhancer sequences?\n11. The authors should explain why their results on DNA methylation was inconsistent with previous findings on the recruitment of DNMT3A/B by H3K36me2/3. The authors should check whether localization of DNMT3A/B are affected in their knockout cells.\n12. The authors should experimentally test whether the 3D organization disruption could abrogate long-range interaction in the KOs (presumably should be more severe in QuiKO > QKO > TKO) to cause downregulation of enhancer function.\n13. Line 323-324: Comparing the difference between ASH1L single KO and parental is unable to conclusively show whether transcriptional downregulation of the genes was due to the loss of catalytic activity. To achieve this aim, authors should instead perform RNA-seq in QuiKO cells ectopically expressed with non-mutated and catalytic site mutant of ASHL1.\n14. Line 333-334: “… Genome-wide, we found that chromatin accessibility increases in QuiKO cells…”. This increase looked rather marginal (20-30%), and it is unsure whether such small change could amount up to any significant impact to increase accessibility by RNA Pol II and other transcription machinery to affect the observed transcription changes. None of such molecular consequences were shown making the data only correlational.\n15. Is the increase in bulk level of H3K9me3 correlated with the increase in the expression level of histone H3 and/or H3K9 methyltransferases? The author should perform ChIP-seq to assess whether RNA polymerase II is excluded by the increase in H3K9me3 in regions not normally occupied by H3K9me3. They should also perform whole genome profiling of other heterochromatin binding proteins including HP1, HDAC, DNMT proteins as independent confirmation of heterochromatin invasion into those regions.\n16. The observation on the loss of compartmentalization and long-range chromatin interaction is very interesting that should be defined in more mechanistic details. The authors should perform ChIP-seq analyses in QuiKO on compartments and chromatin loops-forming condensin and cohesin proteins. They should also do ChIP-seq of CTCF to ascertain the loss of boundary function underlie the observed compartment dissolution.\n17. The Discussion section is mostly a repeat of the results section. The authors should instead put their results in the context of what is known in the field and comment on the novelty of their findings compared with previous publications. They should also discuss limitations of their studies. The can include a model figure to summarize the major findings of the work.\nMinor comments:\n(1) Catalogue numbers of the antibodies should be included into the supplementary table.\n(2) Line 46, although histone PTM is an epigenetic regulation, it is by no means the only epigenetic mark. Suggest remove “epigenetic or” in this sentence.\n(3) Line 50, what do “steric changes” refer to?\n(4) Legend of Supplementary Figure 1a is incongruent and needs rewriting.\n(5) Line 399, what does the authors mean by \"with only a weak dependence on transcription...\" Clarify from which data they discern such conclusion.\n(6) The authors should replace the word “spreading” with “broadening” (or similar), because spreading has a mechanistic connotation for heterochromatin expansion that involves the binding of silencing PTM by adaptors or KTMase that host PTM recognition motif. The lowering of peakiness score is simply descriptive and must not be confused with the mechanistic spreading event unless the authors can support with experiment evidence.\n(7) Title can be more specific.", "reviewer_2": "In this manuscript, the authors combine both new and previously published datasets with multiple new multiomic experiments and new analyses to examine the interplay between H3K36 methylation and other epigenetic modifications and gene expression. In doing so, they add further nuance to emerging data from both in vitro and in vivo systems to underscore the importance of H3K36 methylation in gene regulation. In particular, they reveal new observations regarding large effects on enhancers as well as on constitutive heterochromatin and long-range gene interactions and compartments.\nMore specifically, the authors make several interesting new observations including the role of NSD3 at enhancers and the redistribution of of H3K9me3 with loss of H3K36me2/3. That being said, there are some questions regarding their results that should be addressed:\n1) A major limitation of the current study is that virtually all of the experiments are in vitro results done in a single mouse cell line (apart from repeating the H3K9 result in the HNSCC cells). It would be helpful to now how applicable the other key findings are in other cells and contexts of H3K36me reduction/elimination.\n2) They observe a redistribution of H3K9me3. Do the authors see any changes in the expression of H3K9me3 histone methyltransferases? What about H3K4me1 methyltransferases (i.e. KMT2C, KMT2D) given the increases in H3K4me1 they report?\n3) In a similar fashion, while they report increases in H3K27me3, do they also observe a redistribution of this modification in their data with regions that also lose H3K27me3? This has been recently described by two manuscripts describing the effects of impaired H3K36 on cell fate (Ko, et al. Dev Cell. 2024; Hoetker, et al. Nat Cell Biol. 2023.), which should be cited here. Along these lines, the authors do not really discuss much regarding the classes or categories of gene expression that change with the different manipulations of H3K36me. This would be interesting to know given all of the associations of dysregulated H3K36 methylation leading to both cancer and altered cell fates. For example, RNA-seq from both the mouse mesenchymal stem cells and HNSCC cells could be compared to see if there are any common genes and pathways that are peturbed through the massive H3K9me3 redistribution.\n4) Given the dramatic changes in these repressive chromatin marks, can they use their datasets to determine which one is more associated with the concurrent changes in gene expression that they observe?\n5) Do the authors have any thoughts as to why they may not be seeing any change in DNA methylation levels despite their strong reduction of H3K36 methylation?", "reviewer_3": "The manuscript titled \"H3K36 Methylation - a Guardian of Epigenome Integrity\" by Padilla et al. explores the role of histone H3K36 methylation (H3K36me) in maintaining the integrity of the epigenome, particularly in the context of enhancer activity, gene expression regulation, and the interaction between active and repressive chromatin states. The authors achieved a systematic analysis of H3K36me function by knocking out all five methyltransferases targeting H3K36, individually and in various combinations, allowing the scrutiny of the functions of H3K36me2 and H3K36me3 separately. Their key findings include: 1) establishing the roles of H3K36me2 in sustaining enhancer activity; 2) providing further details on the antagonistic relationship between H3K36me H3K27me; 3) redistribution of H3K9me3 from large heterochromatic domains to euchromatic regions upon H3K36me depletion; the roles of H3K36me in maintaining large scale chromatin compartmentalization. Overall, the experiments described were well designed and executed. Data quality was generally good, and most data analyses are clear and thorough. The data and findings presented here offers new insights into the functions of H3K36me and provide valuable new avenues and resources for the community.\nMajor concerns:\n1. The authors speculate two possible mechanisms for redistribution of H3K9me upon loss of H3K36me without further testing either. This lack of further verification, validation, and mechanistic insights becomes a major apparent deficiency in supporting this novel discovery. ChIP-seq for the H3K9MTs SUV39H1/2 would significantly strengthen this study.\n2. The lack of intragenic DNA methylation change in highly expressed genes upon loss of H3K36me is a surprising finding. Given the gain of H3K9me in euchromatin and its direct link to DNA methylation, one could also speculate that the gain of H3k9me compensated for the loss of H3K36me in regulating genic DNA methylation. This scenario can be tested by knocking down/out H3K9MTs in the H3K36MT mutant background.\nMinor concerns:\n• The labels in some of the figures, especially in Figure 7a, are very small and hard to read. Increase the font size and/or improve figure image quality."}, "author_response": "Friday, June 6th, 2025\nDear Reviewers,\nWe sincerely thank you for your careful and constructive evaluation of our manuscript,\n“H3K36 methylation – a Guardian of Epigenome Integrity.” Your insightful\ncomments and suggestions have significantly strengthened the quality, depth, and\nclarity of our work.\nIn response to your feedback, we have undertaken a substantial number of new\nexperiments and analyses to address the key questions raised and expand the\nmechanistic scope of our findings. These efforts include, but are not limited to, the\nfollowing major additions:\n Mechanistic exploration of SUV39H1-mediated H3K9me3 redistribution,\nincluding both overexpression and knockdown of SUV39H1 in H3K36me-\ndeficient cells, paired with ChIP-seq and RNA-seq.\n ChIP-seq for HP1 to confirm the redistribution of heterochromatin-associated\nproteins.\n Expanded cross-species validation of our most novel findings using two\nadditional human HNSCC cell lines (Cal27 and Detroit562) overexpressing the\nH3K36M oncohistone. These analyses revealed conserved chromatin alterations\nin heterochromatin structure, transcriptional output, and 3D genome architecture.\n The integration of new multi-omic datasets, including ChIP-seq, CUT&RUN,\nATAC-seq, RNA-seq, and Hi-C.\nTogether, these additions provide a more comprehensive and mechanistically grounded\nview of how H3K36 methylation protects genome function by maintaining enhancer\nactivity, restricting repressive chromatin, and preserving higher-order nuclear structure.\nWe have revised the manuscript accordingly to reflect these new data, clarified key\ninterpretations, and expanded the discussion of implications, limitations, and future\ndirections.\nWe greatly appreciate the time and expertise you devoted to reviewing our work. In the\nfollowing pages, we provide detailed, point-by-point responses to each of your\ncomments, indicating how each concern has been addressed in the revised manuscript.\nSincerely,\nPadilla and Shipman et al.\nOn behalf of all authors\nTable of Contents\nI. Reviewer #1 comments and responses (pp. 3-35)\nII. Reviewer #2 comments and responses (pp. 36-48)\nIII. Reviewer #3 comments and responses (pp. 49-52)\nPadilla et al studied effect of combined knock-out (KO) of the H3K36 lysine\nmethyltransferases (KMTases) NSD1-3, ASHL1 and SETD2 on genome-wide\ndistribution of H3K36me2, H3K27me1-3 and H3K9me3. They found H3K36me2 marked\nenhancers. Reduction of H3K36me2 reduced expression of some proportion of genes\nthat supports the role H3K36 methylation on enhancer regulation of promoters and gene\nexpression. Within the genic region, authors studied enrichment of H3K27me in the\nvarious KO mutants and reported H3K36me2 prohibit H3K27me2/3 accumulation. They\nalso showed redistribution of H3K9me3 from heterochromatin to genic regions. Finally\nusing Hi-C, they showed significant changes in chromatin compartmentalization\nassociated with the loss of H3K36me2/3. The work is extensive and show interesting\nconnection of H3K36me2 with higher order chromatin organization. If the work can be\nmore connected and mechanistically explored, it would be a significant finding in the\nfield.\nMajor Comments:\n1. The work currently composed of mostly three weakly connected parts: enhancer,\nH3K9me3 on heterochromatin and Hi-C compartmentalization study. More work needs\nto be done to flow these parts into a complete story.\nWe thank the reviewer for their feedback and the opportunity to clarify the conceptual\nframework of our study. The central objective of our work is to understand the\ndownstream consequences of H3K36me depletion, a hallmark of various\ndevelopmental disorders and cancers. H3K36 methylation is a key mark of euchromatin\nthat influences transcription, antagonizes the spread of repressive chromatin, and—as\n3\nwe demonstrate in our current manuscript—contributes to higher-order genome\narchitecture. Our study builds upon this framework by systematically examining its loss\nat three interrelated levels:\n1. Local chromatin regulation at enhancers and promoters\n2. The effect on the epigenome—including redistribution of repressive\nhistone modifications, particularly H3K9me3—and transcription\n3. Global changes in 3D genome organization\nThese analyses are conceptually and mechanistically linked by their common\ndependence on the loss of H3K36me, rather than being independent lines of inquiry.\nWhile we do not claim that all observed downstream effects are directly causally related\nto one another, they are coordinated consequences of a shared upstream perturbation,\nand collectively reflect the multi-layered role of H3K36me in genome regulation. As the\nreviewer notes, H3K9me3-marked heterochromatin and compartmentalization are\nrelated phenomena—a connection that is well supported in the literature (e.g., Allshire &\nMadhani, Nat Rev Mol Cell Biol, 2018; Feng et al., Genome Biol, 2020; Bian et al.,\nPNAS, 2020; Nichols & Cortes, Cell Rep, 2021; Clyde, Nat Rev Genet, 2021; Spracklin\net al., Nat Struct Mol Biol, 2023; etc.). Our findings align with and build upon these\nstudies by showing that H3K36me depletion leads to ectopic distribution of H3K9me3\nand SUV39H1 in euchromatic regions, which correlates with erosion of chromatin\ncompartmentalization—a novel mechanistic link that expands the functional landscape\nof H3K36 methylation.\nindividual and multiple knockout mouse mesenchymal stem cell lines and their impact\non H3K36 methylation—have been published previously by our group and others (Lu et\nal., Science, 2016; Weinberg et al., Nature, 2019; Rajagopalan et al., PNAS, 2021;\nShipman et al., Genome Biol, 2024) and are appropriately cited. This allows the current\nmanuscript to focus on newly uncovered mechanistic relationships without reiterating\nprior findings. We respectfully disagree that the manuscript presents a disjointed\nnarrative. Reviewers #2 and #3 independently noted the coherence and significance of\nour study design, and we have further clarified the manuscript structure in the revised\ntext to help guide the reader through the logical progression from local to global\nconsequences of H3K36me loss. In addition, in the revised version of our manuscript\nwe have reproduced our most novel findings in two human cell lines and provided\nadditional mechanistic insights. We hope this better articulates the unifying theme and\nscientific value of the work.\n2. It is important to ensure the actual endogenous status of individual KMTases as their\nnames imply (DKO, TKO, QKO and QuiKO). The authors should show immunoblots to\ndemonstrate the level of knockdown of all individual KMTases (NSD1-3, SETD2 and\nASH1L) in single KOs, DKO, TKO, QKO and QuiKO to demonstrate that each gene\nknockout did not ectopically change the levels of the other H3K36 KMTases and that\nthe level of the unaffected KMTases are similar between different mutants.\nWe appreciate the reviewer’s concern regarding the specificity and validation of the\nknockout (KO) cell lines used in this study. As outlined in our response to major\n5\ncomment #1, all mouse mesenchymal stem cell (mMSC) lines employed here—\nincluding single, double (DKO), triple (TKO), quadruple (QKO), and quintuple\n(QuiKO) knockouts—were generated and thoroughly validated in previously\npublished studies. No new KO cell lines were generated for the present work. In\nparticular, we refer the reviewer to our recent publication, “Systematic perturbations of\nSETD2, NSD1, NSD2, NSD3, and ASH1L reveal their distinct contributions to H3K36\nmethylation” (Genome Biology, 2024), in which we directly addressed the potential for\ncompensatory expression changes among the H3K36 KMTases. As shown in\nSupplementary Figure S6 (also included below for convenience), expression profiling of\nall five KMTases across the KO lines demonstrated that the majority of knockouts do\nnot induce substantial changes in the expression levels of the remaining KMTases. The\none notable exception is a ~2-fold increase in NSD2 expression observed in the NSD1-\nKO line. However, this increase appears to be functionally insignificant, as NSD1-KO\ncells exhibit the most profound reduction in intergenic H3K36me2 among all individual\nKO lines—this suggests that compensatory effects are relatively minor. Taken together,\nthese data support the specificity of each KO and indicate that the loss of one KMTase\ndoes not broadly perturb the expression of the others in a manner that would confound\ninterpretation of the observed epigenomic and transcriptomic consequences.\n6\n3. H3K36me profiles of single KMTase knockouts should be included as controls in\nFig1a. Profile of H3K27ac should be included in bottom panel of QKO results in Fig 1a.\nWe thank the reviewer for these suggestions. In response, we have now included both\nH3K27ac and ATAC-seq profiles for the QKO sample in Fig. 1a of the revised\nmanuscript to provide a more comprehensive view of regulatory element activity and\nchromatin accessibility in the context of H3K36me loss. Regarding the inclusion of\nH3K36me profiles from individual KMTase knockouts: as previously noted, a detailed,\ngenome-wide comparative analysis of H3K36 methylation patterns in all individual KO\nlines, as well as their combined effects in multiple-KO contexts, has already been\nconducted and published in our recent study (Genome Biology, 2024). We specifically\nrefer the reviewer to Figure 4 of that study, which addresses the exact comparison\n7\nrequested. As our current manuscript builds directly upon this previous work, the\nemphasis here is not on characterizing differences among the individual KMTase\nknockouts per se, but rather on investigating the downstream consequences of global\nH3K36me depletion, particularly in the QKO and QuiKO models. These include changes\nto other epigenetic marks, chromatin organization, and transcriptional programs. We\nhave cited the previous publication throughout the current manuscript to clearly\ndistinguish between previous work and the new mechanistic insights presented in this\nstudy.\n4. In Figs 1b-c, the decrease in accessibility (20-26%), H3K27ac (30-40%) and increase\nin H3K27me3 (20-54%) were partial and significant amount of these still retained on the\nenhancers in the KO. The authors need to obtain quantitative functional evidence that\nsuch residual level is insufficient to support a full activity of these enhancers and the\namount of enhancer activity is reduced in QKO relative to TKO as they expected.\nWe respectfully disagree that additional functional assays are necessary to support the\nconclusion that enhancer activity is reduced. The observed reductions in H3K4me1 and\nH3K27ac—hallmarks of active enhancers—are widely accepted indicators of diminished\nenhancer function. H3K4me1 marks enhancers that are poised for activation, while\nH3K27ac distinguishes active enhancers by promoting an open chromatin environment\nthat facilitates transcription factor binding. Conversely, increased deposition of\nH3K27me3 is associated with reduced enhancer activity through promotion of chromatin\ncompaction and transcriptional repression (Calo et al., Mol. Cell, 2013; Creyghton et al.,\nPNAS, 2010; Heintzman et al., Nature, 2009). Our conclusions in Figures 1b and 1c are\n8\nsupported by multiple independent and complementary assays: CUT&RUN, which\nreveals a decrease in H3K4me1 and a reciprocal increase in H3K27me3; ChIP-seq,\nwhich demonstrates a reduction in H3K27ac levels; and ATAC-seq, which shows a\ndecrease in chromatin accessibility at enhancer regions. In addition, RNA-seq analysis\nfurther supports a functional consequence of these epigenomic changes, showing that\ngenes associated with the affected enhancers are downregulated. Specifically, among\ngenes proximal to strong enhancers described in Fig. 1c, 154 are downregulated while\nonly 88 are upregulated (provided again below for convenience), reinforcing the\nconclusion that enhancer activity is diminished. These findings are grounded in\nvalidated genomic annotations (see Methods), and are consistent with a broad body of\nliterature demonstrating that H3K36 methylation significantly influences enhancer\nactivity (Yuan et al., J. Biol. Chem., 2011; Streubel et al., Mol. Cell, 2018; Yuan et al.,\nCancer Discov., 2020; Farhangdoost et al., Cell Rep., 2021; Hoetker et al., Nat. Cell\nBiol., 2023; Barral and Déjardin, Nucleus, 2023; etc.). Our current study extends these\ninsights by examining this influence in a localized and mechanistically integrated\ncontext.\n9\n5. ChIP-seq of core and linker histones (H1, H2A, H2B, H3 and H4) should be\nperformed to compare between TKO/QKO/QuiKO in Fig 1b-c and 2e-f to assess\nwhether nucleosomal positioning and occupancy are affected by the KO.\nWe agree that quantifying changes to nucleosome dynamics in the absence of\nH3K36me is an intriguing direction for future investigation. However, we respectfully\nmaintain that this request is not necessary for supporting the current conclusions drawn\nfrom Figures 1b–c and 2e–f. These figures aim to demonstrate changes in enhancer\nand promoter activity through alterations in chromatin accessibility and histone\nmodification states—mechanisms that are already well-characterized by the assays we\nemployed, including ChIP-seq, CUT&RUN, ATAC-seq, RNA-seq, and Whole Genome\nBisulfite Sequencing. The reduction in H3K27ac and H3K4me1, combined with the gain\nof H3K27me3 and reduced ATAC-seq signal, robustly indicate decreased regulatory\n10\nelement activity and chromatin accessibility. These findings are consistent with\ncanonical interpretations of enhancer and promoter inactivation. As cited in our\nresponse to Major comment #4, H3K27me3 deposition is associated with facultative\nheterochromatin and chromatin compaction, while the loss of H3K27ac and H3K4me1\nreflects repression of enhancer activity. ATAC-seq directly measures chromatin\naccessibility, providing a functional readout of nucleosome positioning without the need\nfor core histone ChIP-seq.\nMoreover, the assumption that core histone occupancy data would provide\nsubstantially more insight in this context is not well supported. ChIP-seq for core\nhistones such as H3 or H2B generally provides a relatively uniform signal across the\ngenome and reflects bulk nucleosome presence and lacks the resolution or specificity to\ndistinguish between active and inactive regulatory states, especially when compared to\nthe epigenetic and transcriptional assays already included (Rando and Chang, Annu.\nRev. Biochem., 2009; Henikoff and Shilatifard, Trends Genet., 2011, etc.). Additionally,\nthe enrichment or depletion of histone marks already reflects changes to nucleosomal\ncontext and chromatin state. Overall, we believe our multimodal datasets provide\nsufficient evidence to support our conclusions regarding chromatin and transcriptional\nchanges upon H3K36me depletion, and we consider the additional experiments\nrequested by the reviewer beyond the scope and objectives of this study.\n6. Fig 1e showed approx. 3x more genes upregulated than downregulated in QKO\ncontradictory to what is expected from compromised enhancer function and authors\nattributed to “secondary, downstream effects from global H3K36me2 depletion and/or\n11\nablation of NSD3”. It is unclear why this was listed as the possible reason. Considering\nthat H3K36me2 coordinates transcriptional elongation; its depletion is expected to\ndownregulate transcription. It becomes even more confusing when the authors obtained\na different number of differentially expressed genes by considering H3K36me2\nenrichment. What is the proportion of genes that overlap between these two lists? Are\nall the 97 downregulated genes in Fig 1e found within that 150 in Fig 1f? The authors\nshould explain why there is such a big discrepancy between these results.\nTo clarify, although there is a positive correlation between H3K36me2 and gene\nexpression (see Additional File 1: Fig. S5b in Shipman et al., Genome Biology, 2024),\nglobal changes in gene expression do not necessarily mirror changes in H3K36me\nlevels. The ∼3-fold excess of upregulated versus downregulated genes in the QKO\nversus TKO comparison (Fig. 1e) represents a global transcriptomic response, not\none solely attributable to compromised enhancer function. Such global effects are also\nobserved in our H3K36M-overexpressing HNSCC models (Cal27 and Detroit562),\nwhere similar trends in transcriptional upregulation occur despite near-complete loss of\nH3K36 methylation (see volcano plots below). It is currently unclear what is driving the\ngenome-wide transcriptional activation. Instead, we emphasize the importance of\nfocusing on functionally relevant gene subsets—particularly those whose enhancers are\ndirectly impacted by H3K36me2 loss—to discern specific regulatory mechanisms rather\nthan genome-wide trends. In Fig. 1f, we identified predicted H3K36me2-dependent\ngenes based on proximity to large H3K36me2 peaks overlapping annotated enhancers,\nand compared them to randomized control genes with similar baseline expression.\nWithin this focused subset, the trend is reversed: 150 genes are downregulated versus\n12\n69 upregulated, directly linking the loss of NSD3-deposited H3K36me2 at enhancers to\ngene repression. The proportion of genes that overlap between the 97 downregulated\ngenes in Fig.1e and the 150 downregulated genes in Fig.1f is only 2%. However, to\ngenerate a control set of genes with similar gene expression, we only selected genes\nbetween 9 to 12 FPKM for Fig. 1f. Within the 97 downregulated genes in Fig. 1e, only 5\ngenes meet the criteria of being within 9 to 12 FPKM, with 3 of these genes found in the\ndownregulated genes in Fig. 1f. In the same section of the manuscript, we also\ncompared gene expression changes in NSD3-KO cells relative to parental controls and\nobserved a similar genome-wide trend toward transcriptional upregulation\n(Supplementary Fig. 1d). However, when we focused specifically on the same set of\npredicted enhancer-dependent NSD3 target genes identified in TKO cells, we again\nobserved a tendency toward downregulation (Supplementary Fig. 1e). This parallel\nresult supports our conclusion that loss of NSD3-deposited H3K36me2 at enhancers\ndirectly impacts the expression of specific target genes, even in the context of partial\ncompensation by other K36 methyltransferases. We also acknowledge in the\nmanuscript that it is important to note that NSD3 may have non-catalytic roles that could\ninfluence gene expression independently of H3K36 methylation. For example, NSD3\nhas been shown to interact with BRD4, and serve as a scaffolding protein in\ntranscriptional complexes (Shen et al., Molecular Cell, 2015; etc.). This might partially\nexplain the observation that both NSD3-KO (compared to parental) and QKO\n(compared to TKO) lead to more upregulation than downregulation, even though\nH3K36me2 levels are reduced.\n“coordinates transcriptional elongation.” To our knowledge, there is no direct evidence\nsupporting a role for H3K36me2 itself in coordinating transcriptional elongation. Most\navailable evidence attributes transcription elongation functions to H3K36me3, deposited\nby SETD2—which tethers to the elongating Pol II complex—and associated with\nrecruitment of factors for splicing and DNA repair fidelity (Molenaar and Leeuwen, Cell.\nand Mol. Life Sci., 2022; etc.). In contrast, H3K36me2 has primarily been linked to\nintergenic chromatin organization, enhancer regulation, and DNA methylation\ntemplating (Weinberg et al., Nature, 2019; etc). We would appreciate a specific citation\nif the reviewer is referring to a study suggesting a role for H3K36me2 in elongation.\n14\n7. The authors referred to the function and regulation of the KMTases, but none of these\nproteins were studied in this work. Localization of the H3K36-KMTase proteins should\nbe done in non- and KO cells, relative to the localization of PRC2 and SUV3-9\nmethyltransferase. Line 218 says “…demonstrating that NSD3 is recruited to these\nregions”. However, there is no evidence to support such claim, and authors should\nprovide ChIP-seq data of NSD3 in TKO versus QKO. For this matter, the authors should\nperform whole genome ChIP-seq of the KMTases (NSD1-3, SETD2, ASH1L) in their PA\nand KO cells to correlate with the progressive loss of H3K36me2.\nAs we have noted in response to earlier comments, we refer the reviewer to our\npreviously published manuscript, which contains most of the foundational and validation\nwork for these cell lines (Genome Biology, 2024). While we did not perform ChIP-seq\non the KMTases themselves, our experimental framework is specifically designed to\n15\ndissect their enzymatic contributions. The TKO cells lack NSD1, NSD2, and SETD2,\nand retain only a small set of H3K36me2 peaks. When NSD3 is ablated to generate the\nQKO line, most of the remaining peaks are entirely lost. This functional epistasis is\nstrong evidence that NSD3 is solely responsible for depositing the residual H3K36me2\nin the TKO background. To further confirm this, we performed NSD3 overexpression in\nQKO cells and observed restoration of H3K36me2 at active promoters and enhancers,\napproaching the levels seen in TKO cells (see heatmaps, below). These functional gain-\nand loss-of-function experiments establish a direct role for NSD3 in H3K36me2\ndeposition at these regulatory elements. Thus, our conclusion that “NSD3 is recruited to\nthese regions” reflects clear biochemical evidence derived from endogenous and\nectopic NSD3 perturbation experiments. We do not present the overexpression\nexperiment in our current manuscript as the individual and combined contributions of\nthe K36 KMTases were rigorously evaluated across several previous studies (see\ncitations in response to previous comments). Similarly, our previous study showed that\nASH1L knockout in the QKO background—where all other K36 KMTases are absent—\nabolishes all remaining H3K36me2 peaks, indicating that ASH1L is responsible for\ndepositing the final ~119 H3K36me2 peaks in the genome. These peaks localize to\ndevelopmentally regulated genes and are likely targeted via PBX2, as demonstrated in\nthat work. We reiterate that extensive functional and epigenomic validation of each\nKMTase has been performed and published, and the present study builds on that\nestablished foundation to address the regulatory consequences of H3K36me2 loss at\ncis-regulatory elements.\n16\n8. SETD2 is mostly a H3K36 trimethylase. KO of SETD2 is expected to be opposite to\nthe other K36 dimethylases (NSD1-3, ASH1L) by increasing the levels of H3K36me2\n(instead of decreasing). If so, then TKO, QKO and QuiKO would contain higher\nH3K36me2 at some loci to confound and mask the effect of the dimethylases KO. The\n17\nauthors should show experimental evidence that this possibility is not the case, or at\nleast do not significantly confound the effects of the dimethylases KO.\nWe appreciate the reviewer’s question but note that this concern has also been\naddressed in detail in our prior study (Genome Biology, 2024). As shown there, the\nindividual knockout of SETD2 does lead to an increase in H3K36me2 levels within gene\nbodies—where SETD2 normally deposits H3K36me3. However, this effect is dependent\non the presence of functional dimethylases. In the QKO and QuiKO cells, where NSD1,\nNSD2, and NSD3 are already ablated, global H3K36me2 is nearly eliminated: only\n0.03% and 0.009% of peptides, respectively, retain this mark (as shown by both mass\nspectrometry and Western blot in our previous publication, and provided again below for\nconvenience). Moreover, even with SETD2 knockout in the TKO cells, global\nH3K36me2 levels are significantly reduced compared to DKO (from 8.97% to 4.60% of\nH3K36me2-modified peptides), demonstrating that SETD2-KO does not obscure or\ncompensate for the loss of the dimethylases. Lastly, as also discussed in our prior work,\nSETD2 may contribute modestly to H3K36me2 deposition at some loci, but this\ncontribution is minor and does not alter our interpretation of NSD1/2/3 function.\nCollectively, these data rule out the possibility that SETD2-KO confounds or masks the\neffects of NSD-family KMTase ablation.\n18\n9. The authors should perform whole genome analyses of RNA polymerase II as an\nindependent confirmation of transcriptional changes. Does the invasion of\nH3K27me3 and H3k9me3 into the gene body correlate with the exclusion of RNA\npolymerase II from those localities?\nWhile determining the exact mathematical relationships between the levels of\nrepressive modifications, RNA polymerase II occupancy, and nascent transcript levels is\na worthy future goal, it is beyond the scope of our current study. Although we have not\ndirectly profiled RNA polymerase II occupancy, the well-established mutual exclusivity\n19\nbetween these repressive marks and Pol II (Barski et al., Cell, 2007; etc.) and the robust\ncorrelation between transcriptional output—as measured by RNA-seq—and Pol II\nactivity (Honkela et al., PNAS, 2015; Pretis et al., Genome Res., 2017; Zhu et al., Nat.\nPlants, 2018; Kaye et al. Nat. Comm., 2024) together support the inference that\naccumulation of H3K27me3 and H3K9me3 in gene bodies is accompanied by reduced\nPol II occupancy. Furthermore, the observed negative correlation between these\nhistone marks and RNA-seq signal in our datasets is consistent with such exclusion\n(see statistics for a multiple linear regression model below). Thus, we believe our RNA-\nseq-based conclusions sufficiently capture the underlying transcriptional repression\nassociated with these chromatin modifications.\nRNA-Seq log2 fold changes (TKO/PA) Coefficient Std. error t-value p-value\nΔH3K27me3 -1.034 0.056 -18.435 0.000\nΔH3K9me3 -0.236 0.107 -2.194 0.0283\nΔH3K27me3:ΔH3K9me3 -2.408 0.214 -11.216 0.000\nDF = 6425\nResidual standard error = 2.274\nP-value = 0.000\nR-squared = 0.0746\nAdjusted R-squared = 0.0742\n10. Why do enhancers retain H3K36me2 so much stronger over the other regions?\nIs this due to higher NSD3 localization at the enhancers? Do these enhancers contain\nsequence motifs to preferentially recruit NSD3? Alternatively, would NSD3 be bound\npreferentially to H3K27ac or H3K4me1 that are enriched at the enhancer\nsequences?\nIn our recently published paper (Shipman et al., 2024), we have shown that NSD3 is\npreferentially recruited to active enhancers and promoters (also see heatmaps in\nresponse to major comment #7), although this is only uncovered after knocking out\n(as determined by ATAC-Seq peaks) within NSD3-deposited H3K36me2 regions, we\nfound 50 significantly enriched TF binding sites. However, due to the large size of these\nremaining H3K36me2 peaks, we cannot confirm whether these are indeed NSD3\ntargets or simply enriched TF binding sites found at cis-regulatory elements. Below are\nthe top 10 significantly enriched TF binding sites at the remaining H3K36me2 peaks\nfound in TKO but absent in QKO. Those bZIP motifs are the most common general\ntranscription activator sequences that we find in many analyses and many cell types,\nsuggesting that NSD3 operates in a general open chromatin context. To avoid\nunnecessary speculation and for brevity, we prefer not to include this relatively\nuninformative analysis in the manuscript. Regarding chromatin context, we cannot\nascertain whether NSD3 recruitment is driven by co-occurrence with H3K27ac or\nH3K4me1, as these marks are commonly present together at active enhancers, and\nexperimentally resolving this lies beyond the scope of the present study. Overall, our\ndata suggest that NSD3 preferentially functions in transcriptionally permissive regions,\nbut further dissection of its recruitment determinants is beyond the scope of this study.\n21\n11. The authors should explain why their results on DNA methylation was inconsistent\nwith previous findings on the recruitment of DNMT3A/B by H3K36me2/3. The authors\nshould check whether localization of DNMT3A/B are affected in their knockout cells.\nWhile prior studies have shown that DNMT3A/B recruitment can be guided by\nH3K36me2/3 (Weinberg et al., 2019), our findings in mMSCs reveal a more nuanced\ncontext-dependent relationship. When examining intergenic regions (IGRs) within\ncluster A regions in our TKO samples, we observed a 15% decrease in DNA\nmethylation, which coincided with the spread of H3K9me3 and H3K27me3 (see below).\nIn lowly expressed genes, DNA methylation is reduced approximately 20% in TKO and\n15% in QKO and QuiKO cells (Fig. 4a; middle box). Highly expressed genes, on the\nother hand, retain high methylation levels despite complete loss of H3K36 methylation\n(Fig. 4a; rightmost box). To investigate this further, we have now created DNMT3A/B\ndouble-knockout mouse mesenchymal stem cells and found they phenocopied the\nmethylation loss observed in our multiple-KO models at lowly expressed genes, but had\nminimal impact at highly expressed loci (see below). This suggests that DNA\nmethylation maintenance at actively transcribed regions is largely independent of\nDNMT3A/B and H3K36me2/3, likely reflecting DNMT1-mediated maintenance\nmechanisms. We are currently investigating the contributions of de novo versus\n22\nmaintenance methyltransferases and their respective interactions with histone\nmodifications, but this project is beyond the scope of the current manuscript.\n12. The authors should experimentally test whether the 3D organization disruption\ncould abrogate long-range interaction in the KOs (presumably should be more severe in\nQuiKO > QKO > TKO) to cause downregulation of enhancer function.\nWe appreciate the reviewer's interest in the connection between chromatin architecture\nand enhancer function. While the requested experiments are beyond the current study’s\naims, our Hi-C data demonstrate that long-range interactions (loops with > 500 kb\ndistance from anchor to anchor) are significantly disrupted in both our mMSC and\nHNSCC models following H3K36me depletion. As shown in the barplots below, these\ndisruptions are more pronounced in QuiKO cells than in TKO cells. To analytically\nassess whether disruptions to chromatin loops affect enhancers and their associated\ntarget genes, we identified annotated enhancers located at the anchors of the affected\nloops. We then determined their corresponding target genes and quantified changes in\ngene expression. Notably, we found that genes associated with these disrupted loops\nexhibit preferential downregulation (TKO: 163 downregulated vs 123 upregulated;\nQuiKO: 173 downregulated vs 109 upregulated). These findings suggest that the loss of\nlong-range chromatin interactions is associated with reduced enhancer function,\n23\nconsistent with the observed downregulation of their target genes. These results have\nbeen incorporated into our revised manuscript (Supplementary Figure 8k).\n24\n25\n13. Line 323-324: Comparing the difference between ASH1L single KO and parental is\nunable to conclusively show whether transcriptional downregulation of the genes was\ndue to the loss of catalytic activity. To achieve this aim, authors should instead perform\nRNAseq in QuiKO cells ectopically expressed with non-mutated and catalytic site\nmutant of ASHL1.\nWe thank the reviewer for raising this point regarding the potential distinction between\ncatalytic and non-catalytic functions of ASH1L. While we agree that ectopic expression\nof wild-type versus catalytically inactive ASH1L in QuiKO cells would, in theory, allow for\na direct assessment, this experiment is not currently feasible due to the enormous size\nof the ASH1L protein (>300 kDa), which poses significant technical barriers to\noverexpression. Neither we, nor any of our collaborators, have been able to express\nexogenous ASH1L in any cell type. That said, we respectfully disagree with the\nreviewer’s assessment that our existing data are insufficient to support a catalytic role\nfor ASH1L. As described in the manuscript, we focused our analyses on a specific set of\n84 genes associated with the 119 residual H3K36me2 peaks deposited by ASH1L in\nQKO cells. When ASH1L is ablated (i.e., in QuiKO cell), these peaks—and their\nassociated H3K36me2 signal—are lost, and a majority of the associated genes are\nsignificantly downregulated. To further interrogate whether this transcriptional\ndownregulation could be attributed to the absence of ASH1L protein per se (rather than\nits catalytic activity), we examined gene expression in parental versus ASH1L single-KO\ncells. Importantly, in these cells, H3K36me2 is retained at the promoters of these target\ngenes, likely due to compensatory activity from other K36 methyltransferases. Under\nthese conditions, we observe no significant change in gene expression. This\n26\ncomparative analysis provides support that the catalytic product (H3K36me2), rather\nthan ASH1L protein alone, plays the predominant role in maintaining expression at\nthese loci. Finally, we note that our overall experimental design mirrors the logic\nemployed in our NSD3 analyses, in which we considered only genes associated with\nH3K36me2-marked enhancers, allowing us to more specifically attribute expression\nchanges to H3K36me2 depletion.\n14. Line 333-334: “… Genome-wide, we found that chromatin accessibility increases in\nQuiKO cells…”. This increase looked rather marginal (20-30%), and it is unsure whether\nsuch small change could amount up to any significant impact to increase accessibility\nby RNA Pol II and other transcription machinery to affect the observed transcription\nchanges. None of such molecular consequences were shown making the data only\ncorrelational.\nIn Section 2, our intention was not to imply that the modest genome-wide increase in\nchromatin accessibility observed in QuiKO cells directly facilitates transcriptional\nupregulation. Rather, we aimed to emphasize a contrasting and more functionally\nrelevant observation: that cis-regulatory elements previously marked with residual\nH3K36me2 in QKO cells exhibit a significantly attenuated gain in accessibility relative to\nthe genome-wide trend upon ASH1L loss in QuiKO cells. This disproportionate\nreduction in accessibility at these loci likely reflects localized functional consequences of\nH3K36me2 loss. Supporting this interpretation, we observed that these same regions\nconcurrently lose H3K27ac and gain H3K27me3 (Fig. 2e, f), consistent with a shift\ntoward a repressive chromatin state. Furthermore, genes associated with these residual\nH3K36me2-marked regions—which lose this mark in QuiKO—are significantly\n27\ndownregulated (Fig. 2c), reinforcing a mechanistic link between the local loss of\nH3K36me2 and transcriptional repression.\n15. Is the increase in bulk level of H3K9me3 correlated with the increase in the\nexpression level of histone H3 and/or H3K9 methyltransferases? The author should\nperform ChIP-seq to assess whether RNA polymerase II is excluded by the increase in\nH3K9me3 in regions not normally occupied by H3K9me3. They should also perform\nwhole genome profiling of other heterochromatin binding proteins including HP1, HDAC,\nDNMT proteins as independent confirmation of heterochromatin invasion into those\nregions.\nIn response to the first point: we examined the expression levels of major H3K9\nmethyltransferases (SUV39H1, SUV39H2, SETDB1) and heterochromatin protein 1\n(HP1; CBX5 in mouse) and found no significant changes in transcript levels between\nparental and TKO cells (see barplots below). This suggests that the changes in\nH3K9me3 distribution is not driven by increased expression of these enzymes. This has\nbeen included in the revised version of our manuscript.\nIn response to the suggestion to profile additional heterochromatin-binding\nproteins: we performed ChIP-seq for both HP1 and FLAG-tagged SUV39H1 in the\ncontext of H3K36me-deficient cells. These experiments revealed that SUV39H1\nredistributes from typically inactive (cluster B) regions to active (cluster A) chromatin\nfollowing H3K36me depletion (Fig. 5e, f). Similarly, HP1 binding increased at active\ngene bodies and transposable elements in cluster A (Fig. 6a, b; Supplementary Fig. 6a),\nmirroring the redistribution of H3K9me3. These results have been included in the\n28\nrevised manuscript and provide direct evidence that heterochromatin machinery invades\npreviously active chromatin domains in response to H3K36me loss.\nRegarding the recommendation to perform ChIP-seq for RNA polymerase II, HDACs,\nand DNMTs: we respectfully disagree that these experiments are necessary to support\nour conclusions. First, the exclusion of RNA Pol II from repressive chromatin marked by\nH3K9me3 is a well-established phenomenon (see citations from previous comments),\nand our RNA-seq data already show significant downregulation of genes within regions\nwhere H3K9me3 and HP1 are enriched. Thus, the functional consequence—\ntranscriptional repression—is already evident without needing Pol II occupancy data.\nSecond, the redistribution of SUV39H1 and HP1, along with increased H3K9me3 and\ntranscriptional repression, sufficiently demonstrate the expansion of heterochromatin\ninto previously active regions.\n16. The observation on the loss of compartmentalization and long-range chromatin\ninteraction is very interesting that should be defined in more mechanistic details. The\nauthors should perform ChIP-seq analyses in QuiKO on compartments and chromatin\n29\nloops-forming condensin and cohesin proteins. They should also do ChIP-seq of\nCTCF to ascertain the loss of boundary function underlie the observed\ncompartment dissolution.\nWe thank the reviewer for their interest in the mechanistic basis underlying the\nobserved disruption of 3D genome organization in H3K36me-deficient cells. In response\nto the suggestion to assess boundary and loop-forming factors, we performed ChIP-seq\nfor CTCF in both mMSCs (TKO and QuiKO) and Cal27 cells overexpressing H3K36M.\nIn all cases, we observed a genome-wide increase in CTCF binding, including at both\ncluster A and B regions as well as at compartment boundaries (see below). This\nsurprising—yet consistent—increase in CTCF occupancy suggests that the loss of\ncompartmentalization and long-range interactions is not due to diminished CTCF\nbinding or boundary element collapse. While this increased CTCF binding is highly\nintriguing, it does not explain the dissolution of compartment boundaries, and we don’t\nfeel that it would contribute additional clarity to the present observations on\ncompartmentalization. We are currently investigating the underlying causes, but\nresolving this question will require extensive additional experiments beyond the scope\nof this manuscript. Furthermore, we agree that further mechanistic insights into the role\nof loop extrusion factors such as cohesin and condensin would be valuable in future\nstudies, however, dissecting their redistribution or potential alterations is not necessary\nfor the results presented and beyond the current scope.\n30\n31\n32\n17. The Discussion section is mostly a repeat of the results section. The authors should\ninstead put their results in the context of what is known in the field and comment on the\nnovelty of their findings compared with previous publications. They should also\ndiscuss limitations of their studies. They can include a model figure to summarize\nthe major findings of the work.\nWe thank the reviewer for their feedback. We have revised our discussion to include\nstudy limitations, and a model to summarize the major findings (Fig. 9).\n(1) Catalogue numbers of the antibodies should be included into the supplementary\ntable.\nThe catalogue numbers for each antibody was included in the provided supplementary\ntable. A dedicated column detailing the catalogue numbers was made in the revised\nmanuscript for clarity (Supplementary Table #2; Additional Files).\n(2) Line 46, although histone PTM is an epigenetic regulation, it is by no means the only\nepigenetic mark. Suggest remove “epigenetic or” in this sentence.\nWe have removed “epigenetic or” as suggested in line 46.\n(3) Line 50, what do “steric changes” refer to?\n“Steric changes” refers to hindering or facilitating reactions via the physical or chemical\npresence of the PTM. In the revised manuscript, this has been reworded in the\nassociated text for clarity.\n(4) Legend of Supplementary Figure 1a is incongruent and needs rewriting.\nWe have revised the legend for Supplementary Figure 1a to include more\ncomprehensive details that precisely match the heatmap visualization. Specifically, the\nlegend has been rewritten to: “Correlation matrix heatmaps of genome-wide H3K27ac,\nH3K4me1, H3K27me3 and H3K4me3 signals, binned in 1 kb windows, demonstrate\nstrong reproducibility between replicates, with samples from each cell line clustering\ntogether. Pearson’s correlation coefficients were used.”\n34\n(5) Line 399, what does the authors mean by \"with only a weak dependence on transcription...\"\nClarify from which data they discern such conclusion.\nWhat we intended to convey is that the redistribution of H3K27me2/3 into gene bodies\nin H3K36me-deficient cells occurs largely independently of transcriptional activity.\nThis conclusion is based on the data shown in Figure 3a, where genes are stratified by\ntheir transcriptional activity (high, medium, low). In parental cells, the exclusion of\nH3K27me3 from gene bodies is more pronounced in highly expressed genes, as\nexpected due to the mutually exclusive relationship between transcription and PRC2-\nmediated repression. However, in the TKO and higher-order knockout conditions (QKO,\nQuiKO), H3K27me2 and H3K27me3 accumulate within gene bodies across all\nexpression levels—not just in lowly expressed genes.\nThis pattern indicates that the depletion of H3K36me permits H3K27me2/3 to spread\ninto gene bodies irrespective of transcriptional status. If the effect were strictly\ntranscription-dependent, one would expect invasion primarily in silent or weakly\ntranscribed genes, not in those with active transcription. Thus, the redistribution of\nH3K27 methylation into gene bodies shows only a weak dependence on\ntranscription, meaning the effect is not strongly modulated by transcriptional activity\nand occurs broadly when H3K36 methylation is lost. We have clarified this language in\nthe revised manuscript (now line 428).\n(6) The authors should replace the word “spreading” with “broadening” (or similar),\nbecause spreading has a mechanistic connotation for heterochromatin expansion that\ninvolves the binding of silencing PTM by adaptors or KTMase that host PTM recognition\n35\nmotif. The lowering of peakiness score is simply descriptive and must not be confused\nwith the mechanistic spreading event unless the authors can support with experiment\nevidence.\nWe have replaced “spreading” with “broadening”.\n(7) Title can be more specific.\nWe agree that the title is broad, but we would like it to remain intriguing and to invite the\nreader to read the entire manuscript to learn how various components of H3K36\nmethylation contribute to the integrity of the epigenome.\nIn this manuscript, the authors combine both new and previously published datasets\nwith multiple new multiomic experiments and new analyses to examine the interplay\nbetween H3K36 methylation and other epigenetic modifications and gene expression. In\ndoing so, they add further nuance to emerging data from both in vitro and in vivo\nsystems to underscore the importance of H3K36 methylation in gene regulation. In\nparticular, they reveal new observations regarding large effects on enhancers as well as\non constitutive heterochromatin and long-range gene interactions and compartments.\nMore specifically, the authors make several interesting new observations including the\nrole of NSD3 at enhancers and the redistribution of H3K9me3 with loss of H3K36me2/3.\nThat being said, there are some questions regarding their results that should be\naddressed:\n1) A major limitation of the current study is that virtually all of the experiments are in\nvitro results done in a single mouse cell line (apart from repeating the H3K9 result in the\nHNSCC cells). It would be helpful to now how applicable the other key findings are\nin other cells and contexts of H3K36me reduction/elimination.\nWe agree with the reviewer that generalizability beyond a single cell type is essential for\nbroader interpretation of our findings. In this revised version of the manuscript, we have\nsubstantially expanded the scope of our analyses by validating key findings in two\nadditional human HNSCC cell lines (Cal27 and Detroit562), which do not harbor\nendogenous mutations affecting H3K36 methylation. In these lines, we overexpressed\nthe H3K36M oncohistone, which reduces both H3K36me2 and H3K36me3 by more\nthan threefold. Importantly, this mutation is clinically relevant and has been implicated in\n37\nseveral cancer types. Thus, our use of the H3K36M model provides valuable\nmechanistic insight into the epigenetic dysregulation driven by this cancer-associated\nmutation, thereby broadening the biological and translational impact of our findings. To\ncomprehensively assess the effects of H3K36me depletion, we performed a full panel of\ngenome-wide assays in Cal27 and Detroit562 cells, including: Mass Spectrometry,\nChIP-seq (H3K9me3, and H3K36me2/3), CUT&RUN (H3K9me3), ATAC-seq, RNA-\nseq, and Hi-C. Across both human cell lines, these assays independently recapitulated\nour key findings in mouse mesenchymal stem cells (mMSCs), including:\n Loss of broad H3K9me3 domains and invasion of H3K27me3 into these\nregions (Figs. 5b-d and Supplementary Figs. 5b-e).\n Gain of chromatin accessibility and active chromatin marks at previously\nsilenced H3K9me3-marked loci (cluster B), as shown by ATAC-seq and RNA-seq\n(Figs. 6f,g,i and Supplementary Figs. 6e-i,k).\n Gene and TE expression changes at previously silenced H3K9me3-marked\nloci (cluster B), as shown by RNA-seq (Figs.7a,c,d and Supplementary Figs.\n7b,c)\n Decompartmentalization of nuclear architecture, with weakening of B-B\ninteractions, A-to-B compartment shifts, and widespread loss of long-range\nchromatin loops, as revealed by Hi-C (Figs. 8b-e, and Supplementary Figs. 8b-\nk).\n Disruption of nuclear H3K9me3 foci, with reduced numbers and more internal\nlocalization of these domains (Fig. 8g,h and Supplementary Fig. 8m), supporting\narchitectural destabilization.\nantagonism studies (Sections 3–4) in Cal27 or Detroit562 cells due to experimental and\ninterpretive constraints inherent to the H3K36M mutation:\n1. Sections 1–2 (Enhancer regulation): The effects studied in these sections rely\non localized removal of H3K36me2 in TKO (NSD3-specific) and QKO (ASH1L-\nspecific) cells. In contrast, H3K36M-OE causes widespread loss across\nmegabase-scale domains, making it infeasible to resolve the subtle distinctions in\nenhancer regulation at a resolution comparable to our CRISPR-based deletion\nmodels.\n2. Section 3 (H3K36me2/3 antagonism of PRC2): The H3K36M model depletes\nboth H3K36me2 and H3K36me3, which precludes distinguishing their relative\ncontributions, as we were able to do using SETD2-KO, NSD1/2-DKO, and\nNSD1/2-SETD2-TKO samples in mMSC lines.\n3. Section 4 (Progressive chromatin changes in gene bodies): The H3K36M-\nOE system does not allow for titrated depletion of H3K36 methylation. Thus, it is\nincompatible with our analysis of stepwise chromatin invasion and transcriptional\neffects across multiple engineered genotypes (mMSC cell lines).\nTogether, these new human cell line data confirm the reproducibility of our most salient\nfindings and highlight the conserved role of H3K36 methylation in maintaining\nheterochromatin architecture, 3D genome organization, and epigenetic insulation across\ndistinct biological systems.\n2) They observe a redistribution of H3K9me3. Do the authors see any changes in the\n39\nexpression of H3K9me3 histone methyltransferases? What about H3K4me1\nmethyltransferases (i.e. KMT2C, KMT2D) given the increases in H3K4me1 they report?\nWe thank the reviewer for their inquiry. In response, we performed differential\nexpression analysis of the key H3K9 and H3K4 methyltransferases to evaluate whether\ntranscriptional changes in these enzymes might contribute to the observed redistribution\nof H3K9me3 and H3K4me1, respectively. For H3K9me3, we analyzed the expression of\nSUV39H1, SUV39H2, SETDB1, and HP1 (CBX5 in mouse) in parental and TKO\nmMSCs, and also in Cal27 and Detroit562 HNSCC cells. None of these showed\nsignificant changes in expression, suggesting that the redistribution of H3K9me3 is not\ndriven by altered expression of its core methyltransferases (see barplots below). For\nH3K4me1, which we found to increase globally, we examined the expression of KMT2C\nand KMT2D, the primary H3K4me1 methyltransferases. Interestingly, KMT2C\nexpression was modestly reduced, whereas KMT2D—which is ~3-fold more highly\nexpressed than KMT2C in mMSCs—was significantly upregulated in TKO cells (see\nbarplots below). This trend was also observed across our other multiple-KO mMSC\nmodels. We have now incorporated these findings into the revised manuscript, including\nthe relevant expression barplots and discussion (H3K9 methyltransferases in\nSupplementary Fig. 5f and H3K4 methyltransferases in Supplementary Fig. 4d).\n40\n3) In a similar fashion, while they report increases in H3K27me3, do they also observe a\nredistribution of this modification in their data with regions that also lose H3K27me3?\nThis has been recently described by two manuscripts describing the effects of impaired\nH3K36 on cell fate (Ko, et al. Dev Cell. 2024; Hoetker, et al. Nat Cell Biol. 2023.),\n41\nwhich should be cited here. Along these lines, the authors do not really discuss much\nregarding the classes or categories of gene expression that change with the different\nmanipulations of H3K36me. This would be interesting to know given all of the\nassociations of dysregulated H3K36 methylation leading to both cancer and altered cell\nfates. For example, RNA-seq from both the mouse mesenchymal stem cells and\nHNSCC cells could be compared to see if there are any common genes and pathways\nthat are perturbed through the massive H3K9me3 redistribution.\nWe thank the reviewer for their suggestions. In contrast to our observations of\nH3K9me3, where we detect both gains and losses across distinct genomic regions,\nH3K27me3 shows a global gain with little evidence of regional loss in our TKO mMSCs.\nAs shown in the included scatterplot (please see below), where H3K27me3 ChIP-seq\nsignal was binned into 10 kb windows, there is an increase of H3K27me3 in both genic\nand intergenic regions in mMSC TKO compared to parental cells (coloured bins to the\nleft of the diagonal). This increase indicates a spreading rather than a redistribution of\nH3K27me3. In contrast, the H3K9me3 scatterplot (Fig. 5b) shows data points to the\nright of the diagonal (cluster B), reflecting regional H3K9me3 loss. Bins on the diagonal\nindicate similar levels of signal between the two conditions.\nH3K36M-OE cells (Fig. 7a), 132 were common to both cell lines. Approximately 35%\n(46/132) of these genes are olfactory receptor (OR) genes (see results below). This is\nconsistent with previous findings in which Magklara et al. (2011) found that OR genes\nare heavily marked by H3K9me3 in mice. Another notable group of genes we’ve found\nto be significantly enriched in these regions of H3K9me3 loss in both HNSCC cells and\nmMSCs are gamma-aminobutyric acid receptor (GABA) receptor genes. There is some\nindication in previous studies that H3K9me3 may directly repress GABA receptor\nexpression (Ionescu-Tucker et al., 2021). Furthermore, GABA receptor expression\n43\nmodulates brain-derived neurotrophic factor (BDNF), which has been shown to be\ninhibited by H3K9me3 in mice (Snigdha et al., 2016). Thus, in mMSCs and HSNCC,\nH3K9me3 may act on the upstream regulators of BDNF instead, which are the GABA\nreceptors. These findings have been included in our revised manuscript (Fig. 7c, d).\nFurthermore, we examined genes gaining H3K9me3 in cluster A regions that\nwere common between mMSC and Cal27. We identified 1,237 genes with elevated\nH3K9me3 at their gene bodies. These genes were enriched for mitochondrial and\nmetabolic gene ontology terms, such as the tricarboxylic acid cycle (see below).\n44\n45\n4) Given the dramatic changes in these repressive chromatin marks, can they use their\ndatasets to determine which one is more associated with the concurrent changes in\ngene expression that they observe?\nYes—using partial correlation analyses, we assessed how changes in H3K27me3 and\nH3K9me3 relate to gene expression. In cluster A regions, where most transcriptional\n46\nchanges occur, H3K27me3 changes were more strongly associated with gene\nexpression than H3K9me3, even though both marks are positively correlated (left plot\nbelow). In cluster B regions, which show H3K9me3 loss, gene upregulation correlated\nmost with H3K9me3 depletion. While H3K27me3 increased in these regions, it did not\noffset the transcriptional activation and was not significantly linked to expression\nchanges (right plot below). Overall, H3K27me3 is the main epigenetic correlate of\nexpression changes genome-wide, however, the loss of H3K9me3 plays a larger role\nwithin cluster B regions.\n47\n5) Do the authors have any thoughts as to why they may not be seeing any change\nin DNA methylation levels despite their strong reduction of H3K36 methylation?\nWe did observe modest but consistent reductions in DNA methylation following loss of\nH3K36 methylation. Specifically, in TKO cells, there was an ~20% decrease in DNAme\nat transcribed gene bodies (Fig. 4a, middle box). Additionally, intergenic regions within\ncluster A domains—where H3K27me3 and H3K9me3 spread—showed a ~15%\nreduction in DNAme (see plot below). Interestingly, in our multiple-KO cells, DNAme\nwas predominantly reduced at lowly expressed genes, while highly expressed genes\nretained high methylation levels, despite near-complete H3K36me depletion (Fig. 4a).\nTo explore this further, we generated DNMT3A/B double-knockout (DKO) mMSCs, as\nDNMT3A and DNMT3B are known to interact with H3K36me2/3 (Weinberg et al., 2019).\nThe DNMT3A/B-DKO cells showed a similar pattern to the multiple-KO cells: loss of\nDNA methylation at lowly expressed genes but no reduction at highly expressed genes.\nof H3K36 methylation, likely via DNMT1-mediated maintenance methylation.\nThe manuscript titled \"H3K36 Methylation - a Guardian of Epigenome Integrity\" by\nPadilla et al. explores the role of histone H3K36 methylation (H3K36me) in maintaining\nthe integrity of the epigenome, particularly in the context of enhancer activity, gene\nexpression regulation, and the interaction between active and repressive chromatin\nstates. The authors achieved a systematic analysis of H3K36me function by knocking\nout all five methyltransferases targeting H3K36, individually and in various\ncombinations, allowing the scrutiny of the functions of H3K36me2 and H3K36me3\nseparately. Their key findings include: 1) establishing the roles of H3K36me2 in\nsustaining enhancer activity; 2) providing further details on the antagonistic relationship\nbetween H3K36me H3K27me; 3) redistribution of H3K9me3 from large heterochromatic\ndomains to euchromatic regions upon H3K36me depletion; the roles of H3K36me in\nmaintaining large scale chromatin compartmentalization. Overall, the experiments\ndescribed were well designed and executed. Data quality was generally good, and most\ndata analyses are clear and thorough. The data and findings presented here offers new\ninsights into the functions of H3K36me and provide valuable new avenues and\nresources for the community.\nMajor concerns:\n1. The authors speculate two possible mechanisms for redistribution of H3K9me upon\nloss of H3K36me without further testing either. This lack of further verification,\nvalidation, and mechanistic insights becomes a major apparent deficiency in\nsupporting this novel discovery. ChIP-seq for the H3K9MTs SUV39H1/2 would\nsignificantly strengthen this study.\nbasis for H3K9me3 redistribution is critical to reinforcing our findings. Based on our\nRNA-Seq data from parental and multiple-KO mMSCs, we focused on SUV39H1 as the\nmost relevant H3K9 methyltransferase for further investigation. SUV39H2 is minimally\nexpressed in somatic cells and is primarily testis-enriched (supported by our own RNA-\nseq data, which indicates SUV39H2 is lowly expressed—Supplementary Fig. 5f), while\nSETDB1 predominantly targets repetitive elements and deposits focal H3K9me3 peaks.\nIn contrast, SUV39H1 displays robust expression in our mMSC models and is known to\nform broad H3K9me3 domains—making it a compelling candidate for mediating the\nobserved redistribution (Supplementary Fig. 5f in our revised manuscript). We initially\nattempted ChIP-Seq using multiple antibodies targeting endogenous SUV39H1;\nhowever, none yielded reliable or reproducible signal. As a solution, we overexpressed\nFLAG-tagged SUV39H1 and performed ChIP-seq using an anti-FLAG antibody in both\nparental and TKO mMSCs. Strikingly, SUV39H1 redistributes from heterochromatic\ncluster B regions to euchromatic cluster A regions upon H3K36me depletion in the TKO\ncell line, closely mirroring the redistribution pattern of H3K9me3 (Fig. 5e-h in our revised\nmanuscript). Additionally, consistent with the gain of H3K9me3 within actively\ntranscribed genes, we observed increased SUV39H1 occupancy in gene bodies in TKO\ncells (Fig. 5g). To further support this mechanism, we performed ChIP-Seq for\nendogenous HP1, a canonical reader of H3K9me3 (Figs. 6a-c; Supplementary Figs.\na,b). We found that HP1 similarly redistributes into the same euchromatic regions where\nSUV39H1 and H3K9me3 accumulate following H3K36me loss. These results reinforce\na model in which SUV39H1 mediates ectopic H3K9me3 deposition in the absence of\nregions. Finally, as noted in our response to Reviewer #2, endogenous SUV39H1\nexpression remains largely unchanged in TKO mMSCs (Supplementary Fig. 5f), further\nsupporting the notion that its redistribution, rather than altered expression, underlies the\nectopic H3K9me3 deposition. Together, these new ChIP-Seq data for SUV39H1 and\nHP1 provide mechanistic insight into the chromatin remodeling observed upon\nH3K36me loss and strengthen the evidence that SUV39H1 is the primary\nmethyltransferase responsible for the H3K9me3 redistribution we describe.\n2. The lack of intragenic DNA methylation change in highly expressed genes upon loss\nof H3K36me is a surprising finding. Given the gain of H3K9me in euchromatin and its\ndirect link to DNA methylation, one could also speculate that the gain of H3k9me\ncompensated for the loss of H3K36me in regulating genic DNA methylation. This\nscenario can be tested by knocking down/out H3K9MTs in the H3K36MT mutant\nbackground.\nWe agree that the preservation of intragenic DNA methylation in highly expressed\ngenes despite the loss of H3K36me is unexpected and appreciate the reviewer’s\nsuggestion regarding potential compensatory mechanisms. As shown in Fig. 4a, our\nmultiple-KO mMSCs exhibit a ~20% loss of DNA methylation at transcribed genes\noverall. However, stratification by gene expression revealed that this reduction is\nrestricted to lowly expressed genes, while highly expressed genes retain high\nmethylation levels despite complete depletion of H3K36me2/3. Given the known\nrecruitment of DNMT3A/B to gene bodies via H3K36me2/3 (Weinberg et al., 2019), we\n52\ngenerated DNMT3A/B double knockout (DKO) mMSCs to determine whether these\nenzymes deposit the residual methylation observed in the TKO mMSCs. The\nDNMT3A/B-DKO phenocopied the methylation loss seen in TKO cells at lowly\nexpressed genes, but—similar to TKO cells—had no effect on methylation at highly\nexpressed genes (see aggregate plots below). This suggests that residual methylation\nat highly expressed genes is maintained independently of both H3K36me and\nDNMT3A/B—potentially via DNMT1 maintenance mechanisms.\nMinor concerns:\n• The labels in some of the figures, especially in Figure 7a, are very small and hard to\nread. Increase the font size and/or improve figure image quality.\nThe font size has been increased and figure image quality has been improved, including\nin Figure 7a.\nReviewer #3 (Remarks on code availability):\nCompiled code has been included with revisions. Please access here:\nhttps://github.com/padilr1/H3K36me_guardian_epigenome_integrity_Padilla\nReviewer #2 (Remarks to the Author):\nOverall, the authors have made excellent progress in responding to the reviews and\nhave adequately answered my questions. As suggested in my initial review, I believe it\nwould be helpful to cite several important recent articles (listed below) that have looked\nat the functional effects of perturbed H3K36 methylation, and have shown a\nredistribution of heterochromatic modifications (namely H3K27me3) and how that leads\nto tissue-specific phenotypic changes. Given that the current manuscript has been\ncarried out all in in vitro experiments, citing these studies would help support the larger\nimportance of maintaining epigenetic homeostasis in vivo, and also offer an opportunity\nto suggest how their new findings may actually be playing a larger, yet still unexplored\nrole in these other systems where H3K9 methylation was not assessed.\nPashos ARS, Meyer AR, Bussey-Sutton C, O'Connor ES, Coradin M, Coulombe M,\nRiemondy KA, Potlapelly S, Strahl BD, Hansson GC, Dempsey PJ, Brumbaugh J.\nH3K36 methylation regulates cell plasticity and regeneration in the intestinal epithelium.\nNat Cell Biol. 2025 Feb;27(2):202-217. doi: 10.1038/s41556-024-01580-y. Epub 2025\nJan 8. PMID: 39779942.\nKo EK, Anderson A, D'souza C, Zou J, Huang S, Cho S, Alawi F, Prouty S, Lee V, Yoon\nS, Krick K, Horiuchi Y, Ge K, Seykora JT, Capell BC. Disruption of H3K36 methylation\nprovokes cellular plasticity to drive aberrant glandular formation and squamous\ncarcinogenesis. Dev Cell. 2024 Jan 22;59(2):187-198.e7. doi:\n10.1016/j.devcel.2023.12.007. Epub 2024 Jan 9. PMID: 38198888; PMCID:\nPMC10872381.\nHoetker MS, Yagi M, Di Stefano B, Langerman J, Cristea S, Wong LP, Huebner AJ,\nCharlton J, Deng W, Haggerty C, Sadreyev RI, Meissner A, Michor F, Plath K,\nHochedlinger K. H3K36 methylation maintains cell identity by regulating opposing\nlineage programmes. Nat Cell Biol. 2023 Aug;25(8):1121-1134. doi: 10.1038/s41556-\n023-01191-z. Epub 2023 Jul 17. PMID: 37460697; PMCID: PMC10896483.\nResponse:\nWe thank the reviewer for their helpful feedback and suggestions throughout the peer-\nreview process. We have incorporated the recommended citations to the manuscript\n(references #19-21 in the revised manuscript).\nReviewer #3 (Remarks to the Author):\nIn this revised manuscript, the authors have carried out additional experiments\naddressing some key questions raised by the reviewers. These new results, especially\nthe SUV39H1 ChIP-seq experiment offered new insights on H3K9me3 redistribution in\nthe H3K36 methyltransferase mutants. The overall quality and significance of the\nmanuscript is much improved. I recommend this manuscript be accepted for publication.\nReviewer #3 (Remarks on code availability):\nThe codes are available with sufficient documentations. No concerns.\nResponse:\nWe thank the reviewer for their recommendation for publication, and for their insightful\nsuggestions and feedback in strengthening the quality and impact of our manuscript.\nReviewer #4 (Remarks to the Author):\nI was asked to review the rebuttal and revised manuscript in place of reviewer one who\nwas unable to re-review this manuscript. As such, I had not previously seen or\nassessed the original draft. In this position I assessed the quality of the revised\nmanuscript and the author's responses to the reviewers comments (mainly focussing on\nreviewer one). Having done so I believe the revised manuscript is an interesting and\nrobust assessment of the regulation and regulatory capacity of H3K36me. It represents\na large amount of work that is impactful for the field of epigenetics and gene regulation,\nand explores important topics such as chromatin cross-talk and the impact of altered\nmodification state on higher order chromosomal organisation.\nWith regard to reviewer comments, the authors did directly address several key\nconcerns, but mainly elected to respectfully disagree with many of the points made.\nWorking through these points, I believe that the additions to the paper in response to all\nreviewers has strengthened their findings. Where comments were not addressed, I\nbelieve that the justification that these were not addressed are sound. Particularly as\nmany questions related to findings that have been previously published by the authors,\nadditional redundant assays to those already performed or simply (costly) experiments\nthat are beyond the scope of this already substantial study.\nThe results are impactful, critically appraised, robustly performed, interesting and\nsuitable for publication in Nature communications in their current form. I enjoyed\nreviewing this study.\nResponse:\nWe thank the reviewer for their thoughtful evaluation and positive assessment of our\nrevised manuscript. We appreciate their recognition of the scope, rigor, and impact of\nour study, as well as their acknowledgment that our responses and justifications\nappropriately addressed the prior reviewers’ comments. We are grateful for their\nsupportive recommendation for publication."}]

Decision letter