Site-specific programming characterizes dynamic post-translational acetylation of histone H2B lysine 108 in mouse embryonic stem cells

Fangni Chai; Qin Huang; Li Zhou; Yijia Ren; Yanping Zhong; Wei Cheng; Zhihong Xue; Haiyan Ren

doi:10.1038/s41467-026-77439-7

. 2026 Sep 3;17:10467. doi: 10.1038/s41467-026-77439-7

Site-specific programming characterizes dynamic post-translational acetylation of histone H2B lysine 108 in mouse embryonic stem cells

Fangni Chai ^1,^#, Qin Huang ^1,^#, Li Zhou ^1,^#, Yijia Ren ², Yanping Zhong ¹, Wei Cheng ¹, Zhihong Xue ^2,^✉, Haiyan Ren ^1,^✉

PMCID: PMC13635217 PMID: 42830307

Abstract

Dynamic histone modifications are critical for regulating stem cell fate. However, generalized strategies for site-specific programming of histone modifications profiling on unperturbed chromatin in embryonic stem cells (ESCs) remain challenging. In this study, we established a genetic code expansion-based platform in mouse ESCs (mESCs) for site-specific proteomic mapping and functional analysis within native chromatin. Using this platform, we revealed Rps19bp1 promotes Sirt1-mediated deacetylation of histone H2B lysine 108 acetylation (H2B-K108ac). Moreover, Site-specific-AcK revealed that H2B-K108ac induces the formation of H2B puncta and modulates chromatin accessibility. Functionally, elevation of H2B-K108ac via genetic code expansion and the loss of Rps19bp1 cause significant alterations in the expression of genes associated with mESC differentiation. Furthermore, Rps19bp1 deficiency promotes neural differentiation in mESC-derived teratoma. Our study develops a robust platform for linking specific histone acetylation with chromatin dynamics and cell fate determination, which lays a foundation for future exploration of additional histone PTMs in ESCs.

Subject terms: Post-translational modifications, Embryonic stem cells

The authors present a genetic code expansion-based platform in embryonic stem cells, revealing the functional significance of H2B-K108 acetylation during mESC differentiation.

Introduction

Histone post-translational modifications (hPTMs) critically regulate chromatin architecture and gene expression, enabling rapid cellular adaptation to physiological cues and directing cell fate decisions^1,2. Dysregulated hPTMs have been linked to diverse diseases, including cancer, inflammatory disorders, metabolic syndromes, and neuropsychiatric conditions^2–6, highlighting the therapeutic relevance of decoding their spatiotemporal dynamics. However, understanding how individual, site-specific hPTMs function within native chromatin microenvironments remains elusive, largely due to the lack of generalizable tools for site-specific programming of dynamic modifications^7–12. This gap dramatically limits the ability to harness epigenetic plasticity for regenerative medicine and targeted therapies.

The ESCs, capable of differentiating into all somatic cell types, serve as powerful models for dissecting hPTM dynamics. They possess unique histone modification and chromatin signature landscapes that are distinctly different from those of other cell types^13–16. However, mechanistic studies are severely limited by current methodological limitations. Conventional approaches, such as reader-dependent tools (e.g., antibodies), synthetic chromatin systems, mutagenesis, or enzyme manipulation^10,17–19, suffer from low histone accessibility, scarcity of high-quality readers, disruption of chromatin structure, and poor residue specificity of modifying enzymes. Additionally, conventional mutagenesis strategies fail to recapitulate the dynamic nature and physiological properties of native PTMs. In vitro histone peptide probes and nucleosome-based tools suffer from the loss of native chromatin context or restrictions to histone tail regions^7–11. Although recently developed nucleus-targeted photoaffinity probes enable profiling in native living cells, they cannot characterize structurally constrained histone domains¹². Therefore, innovative technologies that enable precise investigation of site-specific hPTMs in living ESCs are urgent.

To bridge this gap, we developed Site-specific toolkits in mESC based on genetic code expansion (GCE) technology. We fundamentally re-engineered the GCE machinery to overcome genomic instability and transgene silencing—thereby establishing a generalizable site-specific toolkit in ESCs. In principle, the GCE system enables site-directed incorporation of non-canonical amino acids (ncAAs) with tailored chemical or physical properties, such as reactive groups or mimicking PTMs (e.g., acetylation), into any defined histone position while preserving native chromatin integrity. Using this strategy, we achieved covalent capture and interactome mapping, as well as functional analysis of specific histone residues within native chromatin contexts in ESCs. Building on these analyses, we additionally identified an unreported biological function and mechanistic cascade through which core histone modifications remodel chromatin accessibility and orchestrate stem cell differentiation.

We particularly focused on hPTMs located within histone globular domains, which serve as key interfaces for DNA wrapping and chromatin protein engagement. For instance, histone acetylation within these regions can reduce its interaction with DNA and provide docking sites for chromatin readers^1,20–23. Despite their significance, the functional characterization of globular domain modifications remains technically challenging, primarily due to their buried location within the nucleosome core and limited accessibility to conventional detection tools, such as antibodies.

Using the GCE-based platform, we successfully captured the chromatin-interacting proteome associated with H2B-K108 and identified its readers and regulators. Moreover, we demonstrated that H2B-K108 acetylation promotes the formation of subnuclear H2B puncta and modulates chromatin accessibility. Notably, alternations of H2B-K108ac levels or its regulator Rps19bp1 lead to obvious changes of mESC commitment.

Together, we establish a versatile and robust framework for linking site-specific hPTMs to cell fate control, offering valuable molecular insights into the epigenetic regulation of stem cell differentiation.

Results

Develop the site-specific toolkits in mESCs

To elucidate the molecular mechanisms and functions of site-specific modifications in mESCs, we developed Site-specific toolkits for precision engineering (Fig. 1a). This platform is built on GCE technology, which uses engineered translation systems to incorporate ncAAs bearing diverse functional groups at defined sites, thereby enabling precise manipulation and analysis of PTMs (Fig. 1a). Genetic code expansion relies on engineered orthogonal aminoacyl-tRNA synthetase/tRNA pairs. In this study, we employed the orthogonal pyrrolysyl-tRNA synthetase (PylRS, Supplementary Fig. 1a) and multicopy tRNA^Pyl (PylT, Supplementary Fig. 1a) to decode the amber stop codons (TAG) in proteins of interest (POI-X-TAG, Supplementary Fig. 1b), which has been widely used in mammalian cells. While conventional designs employ identical promoters to drive multiple tRNA copies^24–27, we recognized that exogenous genetic elements, particularly those containing repetitive or retroviral-like sequences, are prone to transcriptional silencing in ESCs due to robust epigenetic repression mechanisms^28–30. To overcome this limitation, we constructed a multi-promoter expression system for optimized PylT^M15 variants³¹ (Supplementary Fig. 1c) and incorporated CCA trinucleotides (PylT^M15_CCA, Fig. 1b) to improve tRNA stability³²; meanwhile, endogenous Elongation Factor 1-alpha (EF1α) promoter was used instead of silencing-prone viral promoters to optimize PylRS and POI expression (Supplementary Fig. 1c, d).

Fig. 1 — a Schematic of the GCE-based Site-specific toolkits in mESCs. PylRS: Methanosarcina mazei tRNA^Pyl synthetase; PylT: tRNA^Pyl_CUA; AAs: amino acids; ncAAs: non-canonical amino acids; GCE: genetic code expansion. b Sequence alignment of tRNAs. c Chemical structure of the (3-(3-methyl-3H-diazirine-3-yl)-propaminocarbonyl-Nɛ-l-lysine (DiZPK). d Strategy for generating stable mESC lines harboring the Site-specific toolkits. Hygro B: hygromycin B; EF1α: constitutive EF1α promoter; hU6: constitutive human U6 promoter; mU6: constitutive mouse U6 promoter; sU6: chimeric mouse and human U6 promoters; INS: insulator. e Successful incorporation of DiZPK into EGFP-Y182TAG in mESC^R26_DiZPK cells. f Successful incorporation of DiZPK into EGFP-Y182TAG in mESC^PB_DiZPK cells. g Structural formulas of Nɛ-acetyl-L-lysine (AcK). h Successful incorporation of AcK into histones in mESC^R26_AcK cells. i DiZPK- and UV-dependent crosslinking in mESC^R26_DiZPK cells. j Mass spectrometric analysis confirming AcK incorporation into H3.3-K56TAG in mESC^R26_AcK cells (n = 3 samples). Intensity of K56ac-modified peptides from the H3.3-K56AcK group (gray) were normalized to those of the H3.3-WT group (greige). Quantitative experiments represented the mean ± SEM from three independent experiments. P-value was determined by a two-tailed Student’s t-test. ** p < 0.01, p = 0.0058. For western blots shown in e, f, h, i, representative images were from three independent experiments with similar results. Samples were derived from the same experiment and all blots were processed in parallel. Source data are provided as a Source Data file.

To validate the functional integrity of our engineered system, we first constructed a Site-specific-Link toolkit based on the orthogonal DiZPK-RS/PylT^M15_CCA pair designed for precise incorporation of the photoactivatable crosslinker methyl-diazirin-L-lysine (DiZPK) (Fig. 1c). This system enables covalent capture of protein interaction partners via site-specifically integrated DiZPK residues (Supplementary Fig. 1e). We evaluated amber suppression efficiency using an EGFP-Y182TAG reporter, where the tyrosine 182 codon was replaced with TAG. In HEK293T cells, the DiZPK-RS/PylT^M15_CCA system exhibited DiZPK-dependent amber suppression, as evidenced by full-length EGFP expression (Supplementary Fig. 1f). In mESCs, despite optimization of transfection parameters, including suspension-based delivery, which significantly improved transfection efficiency (Supplementary Fig. 1g), transient transfection of the PylRS/PylT^M15_CCA components failed to achieve detectable TAG suppression in ES-E14TG2a (E14) mESCs (Supplementary Fig. 1f).

To overcome this limitation, we implemented two genomic integration strategies: 1) PiggyBac transposon-mediated random integration and 2) Clustered Regularly Interspaced Short Palindromic Repeats-associated protein 9 (CRISPR/Cas9) -directed site-specific integration at the Gt(ROSA)26Sor (ROSA26) safe harbor locus (Fig. 1d). These approaches generated monoclonal lines mESC^PB_DiZPK (random integration, Supplementary Fig. 1h) and mESC^R26_DiZPK (ROSA26 integration, Supplementary Fig. 1i), respectively. Both lines showed efficient DiZPK incorporation into the EGFP-Y182TAG reporter (Fig. 1e, f and Supplementary Fig. 1f). Given the lower genomic perturbation associated with targeted integration, the mESC^R26_DiZPK line, which exhibited optimal DiZPK incorporation efficiency, was selected for subsequent studies.

Histone acetylation plays important roles in stem cell fate determination. By neutralizing the positive charge of lysine side chains, acetylation weakens ionic interactions between the basic lateral surface of the histone octamer and the negatively charged DNA backbone. Although lysine acetylation in histone tails has been extensively characterized, the functional and mechanistic implications of lateral surface acetylation remain poorly understood. Current approaches, such as histone tail deletion, non-acetylated lysine-to-arginine mutations, and acetylation-mimetic lysine-to-glutamine substitutions, lack both dynamic and biochemical fidelity. The engineered Site-specific-AcK system, which continuously incorporates N_ɛ-acetyl-L-lysine (AcK, Fig. 1g) into newly synthesized target protein using a reprogrammed PylRS variant AcK-RS^33,34, addresses these limitations. Both PiggyBac-generated (mESC^PB_AcK) and ROSA26-targeted (mESC^R26_AcK) monoclonal stem cell lines supported efficient amber suppression, as confirmed by full-length EGFP expression in cells harboring the EGFP-Y182TAG reporter (Supplementary Fig. 1j).

Site-specific system mediate ncAAs incorporation into histones in living mESCs

To establish the utility of our platform for histone studies in a native chromatin context, we introduced ncAAs at defined lysine residues in histones H2A.X, H2B, H3.3, and H4 in living mESCs. Successful incorporation of both DiZPK and AcK were achieved at multiple positions in H2A.X, H2B, and H3.3 (Fig. 1h and Supplementary Fig. 1k). In line with previous reports, H4 failed to allow ncAAs integration (Fig. 1h and Supplementary Fig. 1k). The failure to incorporate ncAAs into H4 likely arises from multiple factors, including intrinsic low solubility, poor mRNA stability, structural constraints of H4, and incompatibility with the GCE system^35–38. Further validation revealed that H3.3-K56DiZPK formed robust UV-dependent (365 nm irradiation) covalent complexes, confirming the photo-crosslinking capability of DiZPK and the feasibility of the GCE platform (Fig. 1i).

To assess the site-specificity and biochemical stability of AcK incorporation, we expressed H3.3-K56AcK using the Site-specific-AcK toolkit and extracted the protein via acid extraction^39,40. Subsequent mass spectrometry (MS) analysis verified the precise AcK incorporation at H3.3 K56, with the acetylation persisting in histone isolated from cells up to 48 hours post-transfection (Fig. 1j and Supplementary Fig. 1l), due to the continuous incorporation of AcK into newly synthesized H3.3 by the GCE system.

Collectively, these results demonstrate our platform's ability to install and track dynamic histone modifications with high spatiotemporal precision in living mESCs.

Site-specific-Link revealed Rps19bp1 as a direct interacting partner of H2B-K108DiZPK in mESCs

Mass spectrometry analysis of isolated nucleic H2B from mESCs covered 12 acetylated lysine sites (Supplementary Fig. 2a), most of which have been identified in previous studies with limited functional characterization^41–49. To investigate how these acetylation sites are recognized and regulated, we employed the site-specific crosslinking strategy to systematically replace individual lysine (represented with X) with DiZPK (H2B-KX-DiZPK) (Fig. 2a), where the diazirine moiety in DiZPK enables covalent capture of the proximal partners upon 365 nm UV activation (Fig. 2b). Notably, H2B-K108DiZPK with UV activation generates obvious high-molecular-weight (HMW) adducts on denaturing SDS-PAGE gel (Fig. 2c), indicating covalent complexes with H2B-K108DiZPK, whereas no detectable crosslinking in other H2B variants (Fig. 2c, Supplementary Fig. 2b). Importantly, the covalent complexes did not harbor histone H2B, H3, H4, H3.3 and H2A.X (Supplementary Fig. 2b–e). These findings suggest that H2B-K108DiZPK may specifically captures regulators, readers, or erasers.

H2B-K108 located on the outermost surface of the nucleosome^50,51, and its acetylation has been suggested to promote activity of Dot1L in vitro⁵², an enzyme known to mediate H3K79 methylation and activate transcription⁵³. Although H2BK108 acetylation has been observed, its specific functions in cells remain unclear. To investigate the function of H2B-K108ac, we employed the Site-specific-AcK system to mimic constitutive H2B-K108ac in mESCs. MS analysis verified site-specific incorporation of AcK into H2B at K108 (H2B-K108AcK, Supplementary Fig. 2f, g). Fractionation analysis confirmed successful integration of H2B-K108AcK into chromatin within mESCs (Supplementary Fig. 2h). Bulk RNA sequencing (RNA-seq) revealed widespread transcriptional changes upon H2B-K108 acetylation, with 520 genes up-regulated and 362 genes down-regulated (p < 0.05, Fig. 2d, e). Gene Ontology Biological Process (GO_BP) enrichment analysis showed that these differentially expressed genes (DEGs) were predominantly involved in key developmental processes (Fig. 2f). These findings suggest that H2B-K108 acetylation may serve as an important epigenetic modulator that modulates the differentiation potential of mESCs.

To systematically profile the interaction proteome of the H2B-K108DiZPK, we employed a tandem purification workflow coupled with mass spectrometry analysis^54,55. Candidate interactors were filtered according to thresholds: positive detection in at least two out of three biological replicates and fold enrichment (FC) > 1.5 relative to wild-type H2B controls. Applying these criteria, we robustly identified 39 significantly upregulated interaction partners (UIPs) (Fig. 2g). GO analysis revealed enrichment of UIPs in chromatin organization and gene transcription (Fig. 2h). Notably, two mechanistically significant interactors, Rps19bp1, a direct regulator of deacetylase Sirt1, and Smarcd1, a key component of the SWI/SNF chromatin remodeling complex, were significantly enriched (Fig. 2g and Supplementary Fig. 2i).

Mass spectrometry identified five potential interaction candidates with molecular weights ranging from 13 to 25 kDa (Supplementary Fig. 2j), exhibiting the possibility to form corresponding crosslinked complexes with H2B-K108DiZPK. To validate, we co-expressed H2B-K108DiZPK-HA with Flag-tagged candidates in living cells. Immunoblotting with an anti-Flag antibody detected UV-dependent crosslinking adducts exclusively in cells co-expressing H2B-K108DiZPK-HA and Rps19bp1-Flag (Fig. 2i). Further validation through co-immunoprecipitation (Co-IP) confirmed that Rps19bp1 interacts with H2B in living HEK293T cells (Supplementary Fig. 2k) and mESCs (Fig. 2j). In vitro GST pulldown assays demonstrated direct binding between Rps19bp1 and H2B (Supplementary Fig. 2l). These results suggested that Rps19bp1 as a direct interactor of H2B.

Rps19bp1 promotes deacetylation of H2B K108 in a Sirt1-dependent manner

Rps19bp1 has been reported to regulate the Lys deacetylase activity of Sirt1, an NAD⁺-dependent class III HDAC^56,57. Site-directed mutagenesis of K108 to arginine (Arg, R) significantly reduced the H2B-Rps19bp1 interaction (Supplementary Fig. 3a). To further investigate the effect of K108 acetylation on Rps19bp1 binding, we expressed H2B-K108AcK using the Site-specific-AcK toolkit. Notably, Rps19bp1 showed stronger binding ability with H2B-K108AcK than WT H2B (Fig. 2k–l). Lysine-to-glutamine (K-to-Q) substitution is a well-established acetylation-mimetic mutation⁵⁸. Accordingly, the K108Q mutation markedly strengthens the H2B–Rps19bp1 interaction (Fig. 2m).

Rps19bp1 exhibits context-dependent regulatory effects that vary according to substrate specificity and cellular conditions. Previous studies have shown that it enhances Sirt1-mediated deacetylation of P53 both in vitro and in vivo^57,59,60. Other investigations, however, have shown that Rps19bp1 activates Sirt1-dependent P53 deacetylation during cell stress^61,62, while exhibiting inhibitory effects in vitro⁶³. Factors including substrates, biological microenvironment, and buffer formulations can modulate the regulatory activity of Rps19bp1⁶⁴. To precisely study Rps19bp1’s role in modulating H2B-K108 acetylation, we knocked down (KD) Rps19bp1 in mESCs. The successful depletion of Rps19bp1 was validated by quantitative real-time PCR (RT-qPCR) and immunoblotting analysis (Supplementary Fig. 3b, c). To further assess the role of Rps19bp1 in regulating H2B-K108 acetylation, we performed mass spectrometry analysis in Rps19bp1 knockout (KO) cells. Notably, Rps19bp1 KO elevates endogenous acetylation at the H2B-K108 residue (Fig. 2n), suggesting its function in promoting deacetylation of H2B-K108ac in living cells. Subsequent Co-IP experiments showed that H2B-K108ac exhibits enhanced binding affinity with Sirt1-Rps19bp1 in live cells (Fig. 2o).

To confirm these findings, we purified recombinant proteins, including H2B, Rps19bp1, Sirt1, H2B-K108Q and site-specifically acetylated H2B-K108 (Supplementary Fig. 3d–h). Pull-down assays demonstrated that K108 acetylation stabilizes the Sirt1-Rps19bp1-H2B complex in vitro (Supplementary Fig. 3i). We next quantified the impact of H2B-K108ac on the binding affinity between H2B and Rps19bp1. Microscale thermophoresis (MST) analysis showed that H2B-WT interacts with Rps19bp1 at an equilibrium dissociation constant (K_d) of 1.31 ± 0.06 μM. By comparison, the acetylation-mimicking H2B-K108Q mutant displayed markedly stronger binding, yielding a K_d of 122 ± 13 nM (Supplementary Fig. 3j, k). Quantitative analysis confirmed that mimic of H2B K108 acetylation strengthens the H2B–Rps19bp1 interaction by approximately 10.7-fold (p < 0.0001, Supplementary Fig. 3l). Additionally, in vitro deacetylation assays demonstrated that Sirt1-dependent deacetylation of H2B-K108ac dose-dependently on Rps19bp1 (Fig. 2p, Supplementary Fig. 3m).

Overall, these results established that Rps19bp1 negatively regulates H2B-K108 acetylation in a Sirt1-dependent manner in cells and in vitro.

The Rps19bp1-H2B-K108ac cascade plays important roles in mESC differentiation

Transcriptome analysis suggested significant roles of H2B-K108ac in regulating mESC differentiation (Fig. 2f). To investigate the potential role of Rps19bp1, we first performed bulk RNA-seq in Rps19bp1 KD mESCs (Fig. 3a, b). Compared with WT controls, Rps19bp1 KD cells exhibited robust transcriptional reprogramming, with 2,092 significantly upregulated and 461 downregulated genes (p < 0.05) (Fig. 3a). The striking preference for gene upregulation indicated that Rps19bp1 functions mainly as a transcriptional repressor in mESCs. GO enrichment analysis of these DEGs revealed strong associations with developmental processes, including tube morphogenesis and neuron projection morphogenesis (Fig. 3c).

Fig. 3 — a Volcano plot of RNA-seq DEGs in *Rps19bp1* KD mESCs (|log₂FC | > 0.58, p < 0.05) with up- and down-regulated genes highlighted in red and blue, respectively (n = 3 samples). b Clustered heatmap of DEGs in *Rps19bp1* KD mESCs, with up- and down-regulation indicated in red and green. c GO and KEGG pathway enrichment analysis of DEGs in *Rps19bp1* KD mESCs. Color intensity represents the enrichment ratio. d *Rps19bp1* KD induces the transcription of lineage-specific markers in undifferentiated mESCs. p = 0.0115; <0.0001; <0.0001; <0.0001; 0.0010; 0.0357; 0.0003; 0.0367; <0.0001. e *Rps19bp1* KD enhances the transcription of lineage-specific genes during EB differentiation. p = 0.0012; <0.0001; <0.0001. Expression levels of *Pax6* (red), *Gata4* (green), and *Fgfbp1* (blue) were assessed by qPCR at D0 and D4. The control group was shown as solid lines with filled circles, whereas the *Rps19bp1* KD group was shown as dashed lines with filled squares. f Venn diagram illustrating the overlapping DEGs between *Rps19bp1* KD and H2B-K108AcK samples. g GO and KEGG analysis of the overlapping DEGs from *Rps19bp1* KD and H2B-K108AcK samples. For GO-MF, enrichment analysis was conducted separately using the intersection genes of up-regulated (shown in red) and down-regulated (shown in blue) groups. GO-BP and KEGG pathway enrichment analyses were performed using all DEGs, with results displayed in red. Color intensity represents the enrichment ratio. h *Rps19bp1* KO promotes H2B occupancy at the promoters of ectoderm-specific genes. p = 0.0315; 0.0026; 0.0027. i *Rps19bp1* KO increases H2B occupancy at the promoters of endoderm- and mesoderm-specific genes. p = 0.0021; 0.0019; 0.0016. j H2B-K108 acetylation promotes H2B occupancy at the promoters of ectoderm-specific genes. p = <0.0001; 0.0004; 0.0003. k H2B-K108 acetylation increases H2B occupancy at the promoters of endoderm- and mesoderm-specific genes. p = 0.0006; 0.0071; 0.0005. In (d, h, i), the *Rps19bp1* KD or KO group was shown in greige, and the control group in gray. In j, k, the H2B-WT group was shown in light blue, and the H2B-K108AcK group in mauve. RNA-seq differential expression (a) was analyzed by two-sided Wald test with BH correction, and GO/KEGG enrichment (c, g) by one-sided Fisher’s exact test with BH correction (FDR < 0.01, overlap ≥ 3). For qPCR (d, e, h–k), data were presented as mean ± SEM from three independent experiments. P-values were determined by two-tailed Student’s ttests. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001. Source data are provided as a Source Data file.

To validate these findings, we selected a panel of differentiation- and pluripotency-related genes from the DEGs (Supplementary Fig. 4a). RT-qPCR confirmed significant upregulation of ectoderm, endoderm, and mesoderm markers in Rps19bp1 KD cells (Fig. 3d). In contrast, expression of core pluripotency factors remained largely unchanged (Supplementary Fig. 4b). To further assess the role of Rps19bp1 in early differentiation, we performed embryoid body (EB) assays and verified that Rps19bp1 expression remained decreased in the Rps19bp1 KD cells during differentiation (Supplementary Fig. 4c). Rps19bp1 KD EB displayed enhanced upregulation of lineage-specific markers compared to WT cells (Fig. 3e and Supplementary Fig. 4d-f), while pluripotency markers declined with time in both groups as expected (Supplementary Fig. 4g). These results together suggested that Rps19bp1 plays an important role in regulating mESC differentiation.

We next asked whether Rps19bp1-mediated changes are linked to H2B-K108 acetylation. Notably, GO terms enriched in Rps19bp1 KD cells showed considerable overlap with those identified in mESCs expressing H2B-K108AcK (Fig. 2f, 3c), supporting the potential role for Rps19bp1 in regulating H2B-K108 acetylation. Given the relatively broad substrate specificity of deacetylases, the regulatory function of Rps19bp1 in supporting Sirt1 activity is unlikely to be confined solely to H2B-K108 deacetylation. In fact, the transcriptional changes in H2B-K108AcK samples were less extensive than those in Rps19bp1 KD cells (Fig. 2d, 3a). To assess the relationship between Rps19bp1 and H2B-K108 acetylation, we performed an intersectional analysis of their respective DEGs. This comparison identified 163 overlapping DEGs (Fig. 3f). GO analysis of these shared genes revealed enrichment in key developmental processes, such as placenta development, tube morphogenesis, and sensory organ development (Fig. 3g). Further functional annotation revealed that the up-regulated genes were predominantly associated with transcriptional functions, such as DNA-binding transcription activator activity, while the down-regulated were enriched for DNA-binding transcription repressor activity (Fig. 3g).

Chromatin immunoprecipitation followed by qPCR (ChIP-qPCR) revealed increased H2B occupancy at the promoters—but not coding sequences—of several lineage-specific genes in Rps19bp1 KO cells (Fig. 3h, i and Supplementary Fig. 4h). Promoters of pluripotency genes showed no significant change in H2B occupancy (Supplementary Fig. 4i). Consistent with Rps19bp1 deficiency, H2B-K108AcK showed enhanced binding ability to promoters of these lineage-specific genes, but not their CDS (Fig. 3j, k and Supplementary Fig. 4j).

Together, these results suggest that the Rps19bp1-H2B-K108ac cascade modulates transcriptional programs critical for mESC differentiation.

H2B-K108 acetylation drives puncta formation and rewrites molecular interactions uncovered by the Site-specific-AcK tool

Interestingly, immunofluorescence staining showed that H2B-K108AcK, but not WT H2B, forms puncta in mESCs (Fig. 4a, b). These droplets are sensitive to 1,6-hexanediol (1,6-HD) treatment, suggesting that multivalent interactions contributed to the puncta formation (Supplementary Fig. 5a, b). Given our prior finding that Rps19bp1 regulates H2B-K108 deacetylation, we further assessed puncta formation in Rps19bp1 KD mESCs. Intriguingly, Rps19bp1 depletion recapitulated the puncta phenotype observed in H2B-K108AcK-expressing cells (Fig. 4c, d). To evaluate the biological relevance of this phenomenon, we monitored their dynamics during differentiation. H2B puncta appeared at 72 h post-differentiation, a time point that coincided with upregulated levels of Pax6—a canonical ectodermal marker (Supplementary Fig. 5c, d). Collectively, these findings indicate that H2B-K108 acetylation facilitates H2B puncta formation, which may in turn promote cellular differentiation.

In vitro reconstitution assays revealed that genomic DNA promotes the formation of H2B-K108ac droplets, while RNA exerts an inhibitory effect on this process (Supplementary Fig. 5e). Actinomycin D-mediated transcriptional inhibition substantially abolished nuclear H2B-K108ac puncta (Fig. 4e, f), establishing a functional link between H2B-K108ac puncta and transcription. To further verify this association, we performed immunofluorescence colocalization analysis using RNA polymerase II (Pol II), a canonical marker of active transcription^65–67. Spatial correlation analysis revealed partial colocalization of H2B-K108AcK puncta with transcriptional hubs marked by RNA polymerase II (Pol II) (Fig. 4g, h)^68,69. These observations indicate that H2B-K108ac puncta may facilitate transcriptional activation.

Interestingly, the evolutionary conserved H2B K108 residue localizes near the acidic patch (Supplementary Fig. 5f), a negatively charged cleft formed by clusters of acidic residues (aspartate and glutamate) on H2A/B dimers. The so-called “acidic patch” plays a critical role in higher-order chromatin organization^70–74. We thus hypothesized that the K108ac may modulate the property of the acidic patch.

To characterize interactome alterations induced by H2B-K108ac, we performed immunoprecipitation of H2B-K108AcK from Site-specific-AcK-engineered engineered mESCs, followed by MS analysis (Fig. 4i). GO analysis of significantly enriched interactors revealed predominant association with gene expression, RNA metabolic process, chromatin remodeling, and so on (Fig. 4j). Co-IP assays verified that H2BK108 acetylation strengthens its interaction with serine/arginine-rich splicing factors (Srsf1, Srsf3), chromatin modifiers (Psip1, Rybp), and RNA metabolism factors (Ewsr1, Cggbp1) (Fig. 4k–m and Supplementary Fig. 5g). Notably, positively charged surface were found in all identified interactors (Supplementary Fig. 5h). We thus generated mutants lacking these putative positively charged regions and demonstrated that loss of these regions significantly impaired their binding to H2B (Supplementary Fig. 5i–l). Together, these findings indicate that H2B-K108 acetylation may modulate chromatin accessibility.

H2B-K108 acetylation modulates chromatin accessibility

To characterize the genomic distribution of H2B-K108AcK, we applied Cleavage Under Targets and Tagmentation‌ sequencing (CUT&Tag-seq)^75,76 on mESCs expressing H2B-K108AcK-EGFP-3×Flag, with H2B-EGFP-3×Flag-expressing mESCs serving as the control. Cells were sorted based on GFP-positive using a Beckman Colter flow cytometry (Supplementary Fig. 6a). Given the lack of a validated specific antibody for H2B-K108ac, we enriched AcK-incorporated positive cells via EGFP sorting and utilized an anti-Flag antibody for subsequent CUT&Tag-seq profiling. A total of 1130 significantly enriched H2B-K108AcK peaks were identified (Fig. 5a), with the majority located in introns (48.3%) and intergenic regions (39.2%) (Fig. 5b). Notably, H2B-K108AcK signals were predominantly found in the distal (> 10 kb) downstream regions of actively transcribed genes (Fig. 5c). In parallel, Assay for Transposase-Accessible Chromatin using sequencing‌ (ATAC-seq)^77,78 analysis indicated a significant increase in 1,937 peaks and a repression of 35 peaks following the introduction of H2B-K108AcK into mESCs (Fig. 5d, e), implying that H2B-K108ac plays a role in modulating chromatin accessibility.

Fig. 5 — a Heatmap showing signals of H2B-K108AcK-enriched CUT&Tag-seq peaks (|log₂FC | > 1 and p < 0.05) in mESCs (n = 3 samples). Color scale represents the signal density. b Distributions of H2B-K108AcK-enriched CUT&Tag-seq peaks (|log₂FC | > 1 and p < 0.05) across different gene features in mESCs. c Scatter plot showing the distributions of expression levels of H2B-K108AcK-enriched CUT&Tag-seq peak-associated genes versus the distance of these peaks relative to nearest TSS. The distance is in bp. d Volcano plot showing differential ATAC-seq peaks in the H2B-K108AcK compared to H2B-WT overexpressing mESCs (n = 3 samples). Significantly up-regulated and down-regulated peaks are labeled with red and cyan dots, respectively. Statistical significance was assessed using a two-sided t-test. No adjustment for multiple comparisons was applied. e Heatmap of differentially accessible peaks. Each row is a peak; each column is a sample. Color gradient represents normalized ATAC-seq signal intensity. f GO enrichment analysis of overlapping target genes between ATAC-seq-upregulated and CUT&Tag-seq-enriched peaks in H2B-K108AcK overexpressing mESCs. g GO enrichment analysis of overlapping target genes between ATAC-seq-upregulated peaks and RNA-seq-upregulated DEGs in H2B-K108AcK overexpressing mESCs (|log₂FC | > 0.58, p < 0.05). GO/KEGG enrichment (f, g) was analyzed by one-sided Fisher’s exact test with BH correction (FDR < 0.01, overlap ≥ 3). Color intensity represents the enrichment ratio. Source data are provided as a Source Data file.

To elucidate the functional consequences of these chromatin changes, we conducted GO_BP analysis of the integrated ATAC-seq and CUT&Tag datasets. The analysis revealed significant enrichment in pathways related to developmental and differentiation processes, such as digestive system development, tube morphogenesis, and glandular epithelial cell differentiation (Fig. 5f). Gene Ontology Molecular Function (GO_MF) analysis further demonstrated marked enrichment in DNA-binding transcription activator activity and genes associated with growth factor binding (Fig. 5f).

In parallel, a combination of ATAC-seq and RNA-seq profiles of H2B-K108AcK-upregulated genes were enriched in GO_BP categories, including kidney epithelium development, regulation of cellular response to growth factor stimulus, regulation of smoothened signaling pathway, ear development, and regulation of postsynaptic membrane potential (Fig. 5g).

These observations demonstrate that H2B-K108 acetylation serves as a functional regulator of chromatin accessibility for development-associated genes.

Rps19bp1 deficiency promotes neural differentiation in mESC-derived teratoma

To understand the cell populations in Rps19bp1-deficient mESCs, we performed single-cell RNA sequencing analysis (scRNA-seq) on Rps19bp1 KD and control mESCs, subsequently integrating them using Harmony⁷⁹. Both samples exhibited comparable qualified QC metrics (Supplementary Fig. 7a) and Harmony embedding values distributions (Supplementary Fig. 7b), indicating effective batch effect correction. UMAP analysis^80,81 identified four distinct subpopulations (Fig. 6a) with signatures of pluripotent mESCs (Nanog, Oct4, Sox2) in clusters 1 and 3, neural lineage (Nrxn1, Negr1, Nes, Nrcam, Ctnna2, Wnt3, Fgf13, Sox2) in cluster 2, and endoderm cells (Gata4, Sox17) in cluster 4 (Supplementary Fig. 7c). Notably, cells in cluster 2 were significantly induced in Rps19bp1-deficient mESCs (Fig. 6a, b) and displayed markedly elevated levels of neural lineage marker genes (Fig. 6c). Additionally, developmental trajectory analysis suggested that cluster 2 represents a further differentiated lineage compared to pluripotent mESCs (Fig. 6d). GO analysis of genes enriched in cluster 2 highlighted significant neural lineage-specific processes, such as synapse organization and postsynaptic neurotransmitter receptor activity (Fig. 6e). Collectively, these findings indicate that the loss of Rps19bp1 promotes the differentiation of mESCs into neural lineages, even under standard mESCs culture conditions. Consistently, upregulation of neural-related genes (Wnt3, Pax6, and Pou4f3) in mESCs following H2B-K108AcK expression (Supplementary Fig. 8a) provides preliminary evidence for its potential involvement in neurogenesis, which requires future validation.

Fig. 6 — a scRNA-seq gene expression projected onto a UMAP space for control and *Rps19bp1* KD mESCs (n = 1 sample). b Proportions of cell clusters in indicated samples. c Gene expression feature plots of indicated genes within scRNA-seq UMAP clusters comparing control and *Rps19bp1* KD mESCs. d Scatter plots of developmental trajectory analysis using Monocle2 for selected clusters. Color scales represent the pseudotime. e GO enrichment analyses of DEGs in cluster 2. Statistical significance was analyzed by one-sided Fisher’s exact test with BH correction (FDR < 0.01, overlap ≥ 3). Color intensity represents the enrichment ratio. f The mRNA level of *Map2* and *NeuN* were dramatically elevated in *Rps19bp1* KO teratomas (p = 0.0076; 0.0004). g Representative IHC images showing NeuN immunoreactivity in WT and *Rps19bp1* KO teratomas. Scale bars, 2.5 mm. h Quantification of NeuN-positive area based on IHC staining (p = 0.0055). i *Rps19bp1* KO promotes neural-directed differentiation from mESCs, as evidenced by increased neurite outgrowth, filopodia formation, and Map2 expression. Scale bars, 10 μm. In (f, h), the *Rps19bp1* KO teratomas group was shown in greige, and the control group in gray. Quantitative experiments represented the mean ± SEM of three independent experiments. P-values were determined by two-tailed Student’s t-tests. ** p < 0.01, *** p < 0.001. Source data are provided as a Source Data file.

To validate the in vivo relevance of these findings, we conducted a teratoma formation assay in immunodeficient mice. While both WT and Rps19bp1 KO mESCs successfully generated teratomas (Supplementary Fig. 9a), KO-derived tumors displayed significantly smaller sizes and lower weights (Supplementary Fig. 9a–c). RT-qPCR confirmed reduced Rps19bp1 expression in the KO group (Supplementary Fig. 9d). Histological examination revealed fewer undifferentiated regions in KO teratomas (Supplementary Fig. 9e, f). Consistently, the neural lineage marker NeuN was significantly upregulated at both mRNA (Fig. 6f) and protein levels (Fig. 6g, h), suggesting enhanced neurogenesis in the Rps19bp1 KO group. RNA-seq of KO-derived teratomas identified 610 upregulated and 839 downregulated genes (p < 0.05, |log₂FC | > 2) (Supplementary Fig. 9g, h). Notably, genes involved in neural development were significantly elevated (Supplementary Fig. 9i). Moreover, neural-directed differentiation assays demonstrated increased neurite outgrowth and filopodia formation in Rps19bp1-deficient neural progenitor cells, along with elevated Map2 expression (Fig. 6i).

These results indicated that Rps19bp1 represses early neural differentiation, and loss of this repression enhances neurogenesis both in vitro and in vivo.

Discussion

Dynamic and site-specific histone modifications play a central role in regulating chromatin. We developed a generalized strategy that site-specifically programs histones to enable residue-centered interactome profiling and PTM functional analysis in living mESCs, with a special analysis of H2B-K108 acetylation. This platform allows precise dissection of causal relations between individual PTMs and cell fate decisions, providing mechanistic insights into the regulatory processes governing mESC differentiation.

Histone PTMs play critical roles in cell fate determination, yet conventional methods for their study are often limited by poor specificity, low accessibility, and disruption of native chromatin architecture. The GCE-based toolkits address these drawbacks by enabling precise, site-specific modification within native chromatin contexts. The chemically incorporated AcK structurally recapitulates endogenous acetylated lysine residues, ensuring faithful profiling of physiological protein-protein interactions. Importantly, the GCE system recapitulates the dynamic epigenetic regulation during mESC pluripotency maintenance and differentiation—an advantage that permanent K108Q mutation cannot achieve. In this study, the K108Q mutant is only used for auxiliary verification, while all core quantitative interaction and epigenetic profiling results rely on the high-precision GCE chemical platform.

Notably, previous GCE strategies have been largely restricted to immortalized cell lines, whereas our system achieves efficient site-specific incorporation of ncAAs in mESCs while preserving pluripotency homeostasis and resisting epigenetic silencing. Using this approach, we successfully revealed the functional significance of H2B-K108 acetylation during mESC differentiation. Although the current study focuses on acetylation as a proof-of-concept demonstration, which serves to validate the feasibility and robustness of our technical framework, the core GCE strategy is not limited to a single PTM type. Instead, it can be readily adapted to diverse histone modifications, such as acylation and sulfation, and further extended to functional characterization of non-histone proteins. Collectively, this work establishes a generalizable framework for decoding site-specific PTM functions in developmental systems, expands our mechanistic understanding of core-domain histone modifications, and provides a versatile tool for dissecting the regulatory mechanisms of discrete protein modifications under both physiological and pathological conditions.

Despite its clear functional relevance, Sirt1 was not detected in our H2B-K108DiZPK-based interactome. As the catalytic deacetylase responsible for H2B-K108ac modification, Sirt1 presumably interacts with H2B via highly transient and catalysis-dependent binding. Proximity crosslinking mediated by genetically encoded ncAAs is highly sensitive to spatial orientation, interacting-residue distance, and interaction duration within the UV activation window^82,83. Accordingly, it is reasonable that Sirt1 evades capture by the H2B-K108DiZPK probe.

While much attention has been given to N-terminal tail modifications, PTMs within the core domains are increasingly recognized as critical regulatory layers in nucleosome architecture and DNA-histone interactions^1,74,84–88. Applying our GCE platform, we elucidated the interactome of H2B-K108, a residue located on the outermost surface of the nucleosome core that participates in hydrogen bonding networks, which contribute to nucleosome stability^50,51. Intriguingly, reduced methylation at H2B-K108 has been observed in the frontal cortex of Alzheimer’s disease patients⁵¹. Modification at this residue is likely to disrupt inter-chain interactions, thereby promoting nucleosome disassembly. Furthermore, H2B-K108 was identified as the most reactive lysine residue near the acidic patch, and its acetylation has been suggested to enhance Dot1L activity in vitro⁵². Despite these clues, the specific cellular functions of H2B-K108 acetylation remain unclear.

Our findings reveal that H2B-K108 acetylation modulates chromatin accessibility in mESCs. We further identified its erasers and readers and elucidated their functional contributions to mESC differentiation. These results provide insights into the interplay between histone modifications and the regulation of stem cell fate. Notably, however, enhanced chromatin accessibility and transcriptional changes revealed by H2B-K108AcK are limited, which may be due to the relatively low expression of H2B-K108AcK compared to endogenous multi-copy H2B expression, as well as the stochastic genomic distribution of H2B-K108AcK.

In summary, we establish a GCE-based strategy to probe site-specific modifications in mESCs, advancing the molecular understanding of stem cell differentiation. This approach not only facilitates the functional characterization of individual PTMs but also holds broad potential for studying diverse protein modifications across biological contexts. Future applications include employing GCE technology to investigate the crosstalk and dynamics of multiple PTMs simultaneously, promising to reveal epigenetic regulation.

Methods

Reagent and resources

The reagents and tools used in this study are listed in Supplementary Data 1.

Cell culture and transfection

HEK293T cells were obtained from the National Collection of Authenticated Cell Cultures (NCACC) and cultured in Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS) at 37 °C under 5% CO₂.

ES-E14TG2a mESCs were obtained from the American Type Culture Collection (ATCC) and cultured in DMEM containing 15% FBS, 1× Glutamax, 1× penicillin-streptomycin, 1× non-essential amino acids (NEAA), 1× Embryomax Nucleosides, 0.1 mM β-mercaptoethanol (β-ME), and 1 × 10⁵ units/50 mL leukemia inhibitory factor (LIF) at 37 °C under 5% CO₂.

HEK293T cells were transfected with polyethylenimine (PEI) and suspended E14 were transfected with Lipoplus or Xtreme HP transfection reagent⁸⁹.

All cell lines were routinely confirmed to be free of mycoplasma contamination.

Plasmids

All genes were cloned using the ClonExpress II One-Step Cloning Kit. The mouse H2ax, H2b, H3.3, and H4 genes were synthesized by General Biol. The Sirt1 gene was synthesized by Tsingke Biotechnology. These genes were inserted into pEF1α-HA/Flag for mammalian expression. The EF1α promoter was amplified from the PaA036_pEGIP plasmid and cloned into the pCMV-HA/Flag plasmids generated in our previous work⁹⁰. The sequences of mouse Rps19bp1, Psip1, Rybp, Cggbp1, Ewsr1, Gar, Nhp2l1, Snrpd3, Srsf1, and Srsf3 genes were amplified from a mouse cDNA library and cloned into the pcDNA3.1-StrepII vector or the pcDNA3.1-EGFP vector. The pEF1α-H2B-XTAG-Flag plasmids and Rybp, Ewsr1, Srsf1, Srsf3, Cggbp1, and Psip1 truncated bodies were constructed through site-directed mutagenesis using KOD DNA polymerase. For prokaryotic expression, the H2B, H2B-XTAG, and Sirt1 genes were cloned into the pET28b (+) vector with a His, StrepII, StrepII-Flag or EGFP-His tag, and Rps19bp1 was cloned into the pGEX-4T-1 vector with a GST tag.

For gene knockdown and knockout, the oligos were resuspended to a final concentration of 20 μM in annealing buffer (5 μL each oligo, 5 μL 10x NEB buffer, 35 μL nuclease-free water). The mixture was subjected to an annealing protocol in a PCR machine as follows: incubation at 70 °C for 10 min, followed by gradual cooling to room temperature. The annealed oligonucleotides and the digested vector (pLKO.1-TRC vector for gene knockdown and plentiCRISPR V2 vector for knockout) were ligated at a molar ratio of 3:1 using a standard ligation reaction system (2 μL annealed oligos, 20 ng digested pLKO.1-TRC-cloning vector, 2 μL 10× NEB T4 DNA ligase buffer, 1 μL NEB T4 DNA ligase, nuclease-free water up to 20 μL total volume). The ligation mixture was incubated at 25 °C for 3 h.

The plasmids pRosa26-EF1α-DiZPKRS-4×PylT^M15_CCA and pRosa26-EF1α-AcKRS-4×PylT^M15_CCA were constructed for targeted integration of the GCE system into the Rosa26 locus in mESCs. Four tandem copies of the suppressor tRNA_CUA (4×PylT^M15_CCA) were commercially synthesized (Genewiz) and cloned into the pCMV-DiZPKRS or pCMV-AcKRS vector to generate the intermediate plasmid pCMV-DiZPKRS/AcKRS-4×PylT^M15_CCA. The EF1α promoter was then amplified using primers containing BsrGI, NotI, PacI, and HindIII restriction sites and used to replace the CMV promoter of the pCMV-DiZPKRS/AcKRS-4×PylT^M15_CCA plasmid via digestion with BsrGI and HindIII, followed by ligation. The EF1α-DiZPKRS/AcKRS fragment was subsequently prepared by PCR amplification with primers containing NheI and SalI restriction sites, and the 4×PylT^M15_CCA fragment was obtained by PacI-NheI digestion of the pEF1α-DiZPKRS/AcKRS-4×PylT^M15_CCA plasmid. Finally, the PaA036_Ai65 plasmid (obtained from ZhiHong Xue’s lab, which contains the left (1083 bp) and right (4266 bp) homology arms of the Rosa26 locus) was digested with PacI-SalI and ligated with the EF1α-DiZPKRS/AcKRS and 4×PylT^M15_CCA fragments to yield the pRosa26-EF1α-DiZPKRS/AcKRS-4×PylT^M15_CCA. Finally, a hygromycin resistance cassette was amplified from PaD005 and introduced via homologous recombination, replacing the original G418 resistance marker, resulting in the plasmid pRosa26-EF1α-DiZPKRS/AcKRS-4×PylT^M15_CCA.

For piggyBac-based donor plasmids, the Rosa26 homology arms of pRosa26-EF1α-DiZPKRS/AcKRS-4×PylT^M15_CCA were substituted with piggyBac inverted terminal repeats (ITRs) via restriction digestion and ligation, generating the pPiggy-EF1α-DiZPKRS/AcKRS-4×PylT^M15_CCA plasmids.

All plasmids were verified by restriction digestion and Sanger sequencing to confirm correct sequence assembly.

Recombinant protein expression and purification

The pGEX-2T-mLIF plasmid was transformed into E. coli BL21 (DE3) and cultured in LB medium supplemented with ampicillin at 37 °C until OD₆₀₀ reached 0.6. Protein expression was induced by 1 mM IPTG for 3 h at 37 °C. Cells were harvested, washed with PBS, and lysed in Tris-EDTA buffer containing lysozyme (0.5 mg/mL), DNase I, RNase H, and protease inhibitors, followed by three freeze-thaw cycles. The lysate was centrifuged, and the supernatant was incubated with Glutathione Sepharose® 4B beads overnight at 4 °C. After extensive washing, GST-mLIF fusion protein on beads was cleaved using thrombin to release mLIF. The eluted protein was concentrated, adjusted to a final concentration of 1 mg/mL in PBS containing 0.5% BSA, aliquoted, and stored at –80 °C.

For H2B-StrepII-Flag protein expression, BL21(DE3) cells with the pET28-H2B-StrepII-Flag plasmid were induced with 0.5 mM IPTG at 37 °C for 4 h. For H2B-K108AcK-StrepII-Flag expression, cells with the pET28-H2B-K108TAG-StrepII-Flag and pSupAR-AcK3RS-mbF (IPYE) plasmids were incubated with 5 mM Nε-acetyl-L-lysine and induced with 0.5 mM IPTG at 30 °C overnight. Samples were harvested, resuspended in StrepII lysis/binding buffer (100 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1% TritonX-100, pH 8.0) with 25 mM nicotinamide, and treated with lysozyme at 37 °C for 20 min, followed by Benzonase^® Nuclease treatment on ice for 20 min. Lysates were sonicated and centrifuged at 16,000 g for 20 min at 4 °C. The supernatant was incubated with Strep-Tactin beads for 3 h and washed three times with StrepII wash buffer (100 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1% TritonX-100, pH 8.0). Samples were eluted using StrepII elution buffer (100 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1% TritonX-100, 0.5% SDS, pH 8.0) containing 25 mM d-desthiobiotin. Purified protein was concentrated using a Millipore ultrafiltration device and stored at −80 °C.

For GST-Rps19bp1 protein expression, BL21(DE3) cells with the pGEX-4T-1GST-Rps19bp1 plasmid were induced with 0.5 mM IPTG at 18 °C overnight. After sonication and centrifugation, the supernatant was incubated with Glutathione Sepharose® 4B beads. The protein was eluted with GST lysis/binding buffer (50 mM Tris, 150 mM NaCl, 0.3 mM DTT, 1 g/L NP-40, pH 7.5) with 25 mM reduced glutathione. All other steps are the same as the H2B-StrepII-Flag purification protocol.

For His-tagged protein purification, the lysed supernatant was incubated with Ni-NTA beads and eluted with 500 mM imidazole in His wash buffer (50 mM Tris-HCl, 500 mM NaCl, 10% glycerol, 5 mM imidazole, pH 8.0). All other procedures remained consistent with the H2B-StrepII-Flag purification.

Site-specific-Link

For the DiZPK-based Site-specific-Link, mESC^R26_DiZPK or HEK293T cells were transiently transfected with plasmids encoding either wild-type or mutant histone constructs, as well as the pEF1α-DiZPKRS-4×PylT^M15_CCA plasmid, using LipoPlus. Six hours after transfection, the medium was replaced with medium containing 1 mM DiZPK. After 48 h of culture, cells were subjected to UV irradiation at 365 nm for 20 min on ice using a UV crosslinker. Cells were then collected, and proteins were extracted for further analysis.

Site-specific-AcK

For Site-specific-AcK, mESC^R26_AcK was transiently transfected in suspension with plasmids encoding either wild-type or mutant histone constructs, together with the pEF1α-AcKRS-4×PylT^M15_CCA plasmid using LipoPlus. Six hours after transfection, the medium was replaced with medium supplemented with 5 mM AcK. Following 48 h of culture, cells were harvested, and proteins were isolated for further analysis.

Acid extraction of histones from mESCs

For acid extraction of histones, cells were lysed in NETN buffer (150 mM NaCl, 5 mM EDTA, 20 mM Tris-HCl pH 8.0, 0.5% NP-40, and 1% protease inhibitor cocktail) for 10 min on ice, and centrifuged twice at 2,000 g for 10 min at 4 °C. The pellets were resuspended in 0.2 M H₂SO₄ buffer and extracted with gentle rotation for 4 h at 4 °C. After centrifugation at 2000 g for 15 min at 4 °C, proteins in the supernatant were precipitated with trichloroacetic acid (TCA) at 4 °C overnight. The pellets obtained after centrifugation at 20,000 g for 15 min at 4 °C were washed twice with cold acetone, air-dried, and resuspended in distilled water. For assays involving the detection of H2B acetylation, the NETN buffer was supplemented with 25 mM nicotinamide to inhibit deacetylase activity.

For MS analysis of co-immunoprecipitated covalent complexes, acid-purified histones from mESCs expressing H2B-K108DiZPK-Flag-StrepII were incubated with anti-Flag beads in Flag buffer (150 mM NaCl, 50 mM Tris-HCl, pH 7.4, and 1% protease inhibitor cocktail) for 4 h at 4 °C on a rotator. Beads were washed twice with Flag buffer, and bound proteins were eluted twice with 3–5 bead volumes of acidic elution buffer (0.1 M glycine-HCl, pH 3.0) for 5–10 min at room temperature with gentle rotation. The eluates were immediately neutralized by adding one-tenth volume of 1 M Tris-HCl (pH 8.0) to adjust the pH to 7.0–8.0. For the second purification step, the neutralized Flag eluates were incubated with Strep beads in Strep buffer (150 mM NaCl, 50 mM Tris-HCl, 1 mM EDTA, pH 8.0, and 1% protease inhibitor cocktail) for 2 h at 4 °C on a rotator. Beads were washed twice with Strep buffer, and bound proteins were boiled in loading buffer at 95 °C for 10 min.

For H2B-K108AcK-Flag-StrepII purification, acid-extracted histones were further purified using Strep-Tactin beads as described above and eluted with 50 mM desthiobiotin in Strep buffer.

For purification of H3.3-K56AcK-Flag, acid-extracted histones were further purified using anti-Flag beads. Beads were washed, and bound proteins were boiled with protein loading buffer at 95 °C for 10 min.

Fraction of chromatin

mESCs expressing H2B-K108AcK or H2B-K108DiZPK were lysed on ice using ice-cold NP-40 lysis buffer (150 mM NaCl, 0.05% NP-40, and 10 mM Tris-HCl, pH 7.5) with 1% protease inhibitor cocktail for 7 min. Transferring the lysate onto a sucrose pad (24% sucrose (w/v) in NP-40 lysis buffer) and centrifuging at 20,000 g and at 4°C for 10 min to obtain the supernatant as the cytoplasm. The pellets were washed once with 1 mL cold wash buffer (PBS with 1 mM EDTA) and centrifuged at 1000 g for 2 min at 4 °C. The pellets were gently resuspended with 200 µL glycerol buffer (20 mM Tris-HCl pH 8.0, 75 mM NaCl, 0.5 mM EDTA, 0.85 mM DTT, 50% glycerol, and 1% protease inhibitor cocktail) and mixed gently. Lysed the pellet within nuclear lysis buffer (1 mM MDT, 7.5 mM MgCl₂, 0.2 mM EDTA, 0.3 mM NaCl, 1 urea, 1% Nonidet P-40, and 10 mM EPES, pH 7.6) with 1% protease inhibitor cocktail for 2 min on ice and centrifuged at 14,000 g for 2 min at 4 °C. The supernatant is the soluble fraction of the nucleus. Wash the pellet twice with 1 mL PBS/1 mM EDTA and centrifuge at 14,000 g for 1 min at 4 °C to obtain the pellet, followed directly by solubilization in RIPA buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% Trion X-100, 1% sodium deoxycholate, 0.1% SDS, benzonase, and 1% protease inhibitor cocktail).

In vitro deacetylation assay

Purified H2B and H2B-K108AcK proteins were incubated either with purified Sirt1 or with Sirt1 and Rps19bp1 at 37 °C for 1 h in preheated deacetylation buffer (50 mM Tris HCl, 1 mM MgCl₂, 150 mM NaCl, pH 8.0) containing 10 mM NAD⁺.

Immunoprecipitation and immunoblotting

Cells were lysed in NETN buffer (250 mM NaCl, 5 mM EDTA, 20 mM Tris-HCl, 0.5% NP-40, pH 8.0) supplemented with protease inhibitor cocktail. Cell lysates were disrupted by passing through a 1 mL syringe more than ten times. The clarified lysates were then incubated with anti-Flag agarose beads, Strep agarose beads, or Ni-NTA beads overnight at 4 °C with gentle rotation. The beads were washed three times with lysis buffer, and bound proteins were eluted by boiling in SDS loading buffer for 5 min.

Samples were separated by electrophoresis on SDS-PAGE gels and transferred onto PVDF membranes. Membranes were blocked with 5% (w/v) skim milk in TBST and incubated with primary antibodies overnight at 4 °C. After washing, membranes were incubated with secondary antibodies (1:10,000) for 1 h at room temperature. Signals were detected using a SageCapture imaging system (Sinsage, Beijing, China). Uncropped and unprocessed blot images are provided in the Source Data file.

Pulldown assays

For the His pulldown assay, His-tagged proteins were incubated overnight with Ni-NTA beads at 4 °C in His binding buffer (50 mM Tris, 500 mM NaCl, 20 mM Imidazole, 1% Tween20, 10% glycerol, 1.4 g/L β-Me, pH 8.0) under gentle rotation. Prey proteins were then added and incubated for 4 h. Washed the beads three times with His wash buffer and denatured the samples using SDS loading buffer for subsequent immunoblotting analysis.

For the StrepII pulldown assay, StrepII-tagged proteins were incubated with Strep-Tactin beads in the presence of prey proteins in Strep lysis/binding buffer for 3 h at 4 °C with gentle mixing. The beads were washed three times with Strep wash buffer, and co-immunoprecipitated complexes were denatured using SDS loading buffer for immunoblotting. Uncropped and unprocessed blot images are provided in the Source Data file.

Microscale thermophoresis (MST) assay

MST binding assays were performed on a Monolith NT.115 instrument (NanoTemper Technologies, Germany) at 25 °C. All assays were conducted in MST buffer containing 20 mM HEPES (pH 7.5), 500 mM NaCl, 0.05% (v/v) Tween-20, and 1 mM DTT. The unlabeled H2B-His and H2B-K108Q-His ligands were prepared in a 1:2 serial dilution series, with concentration ranges from 50 μM to 1.53 nM and 6 μM to 0.18 nM, respectively. After 15 min incubation at room temperature, samples were loaded into standard Monolith NT.115 capillaries. Measurements were performed using 60% or 80% MST power with the blue channel.

Raw data were analyzed with MO. Affinity Analysis software (v2.3) using the built-in one-site binding model to calculate the K_d. For statistical comparison, K_d values from three independent biological replicates were compared via an unpaired t-test. All data are represented as mean ± SD (n = 3). All capillary scanning profiles were inspected to exclude samples showing aggregation or nonspecific surface adsorption artifacts.

Construction of monoclonal mESCs

To generate monoclonal mESC^PB_DiZPK and mESC^PB_AcK lines, the pPiggy-EF1α-DiZPKRS-mbp-4×PylT^M15_CCA or pPiggy-EF1α-AcKRS-mbp-4×PylT^M15_CCA plasmid and the helper plasmid encoding the piggyBac transposase (pCMV-hyPBase) were co-transfected into mESCs using the Xtreme HP transfection reagent. Forty-eight hours post-transfection, the cells were subjected to hygromycin selection for 4–7 days. Resistant single colonies were manually picked and cultured in 96-well plates. Positive clones were identified using PCR genotyping and further validated by Sanger sequencing.

To generate monoclonal mESC^R26_DiZPK and mESC^R26_AcK lines, mESCs were co-transfected with the pX330-U6-Chimeric_BB-CBh-hSpCas9 plasmid containing sgRNA targeting the Rosa26 locus and pRosa26-EF1α-DiZPKRS-mbp-4×PylT^M15_CCA or pRosa26-EF1α-AcKRS-mbp-4×PylT^M15_CCA. Seventy-two hours after transfection, the cells were subjected to hygromycin selection for 4-7 days. Single colonies were manually picked and cultured. Positive clones were identified using PCR genotyping and further validated through Sanger sequencing of the edited genomic region.

Construction of knockdown cells

The lentivirus was produced in HEK293T cells by co-transfecting pLKO.1-shRNA plasmid with the packaging plasmids psPAX2 and pMD2.G. Viral supernatant was collected 48 h after transfection. The mESCs were incubated with lentiviral particles in the presence of 8 µg/mL polybrene. After 24 h, the medium was replaced with fresh mESC culture medium. Stable knockdown cells were selected using puromycin (2 µg/mL) or G418 (1 mg/mL) for 3-5 days. Knockdown efficiency was confirmed by quantitative RT-PCR and immunoblotting analysis.

The shRNA sequences targeting Rps19bp1 were as follows:

5′-CCGGCACTGAAGAAGACTTCCAGAACTCGAGTTCTGGAAGTCTTCTTCAGTGTTTTTG-3′(shRNA1); 5′-CCGGAGGCAATTCAGGCCCAGAAACCTCGAGGTTTCTGGGCCTGAATTGCCTTTTTTG-3′(shRNA2);

5′-CCGGCCGGAAAGGAACGTATCTGTTCTCGAGAACAGATACGTTCCTTTCCGGTTTTTG-3′(shRNA3).

Construction of knockout cells

For Lenti-CRISPR/Cas9-mediated gene knockout, sgRNA targeting Rps19bp1 (5′-GGTCTCGACACTCTCGCTTC-3′) was cloned into the plentiCRISPR v2 plasmid. The resulting plentiCRISPR v2 plasmid was co-transfected with the packaging plasmids psPAX2 and pMD2.G into HEK293T cells to produce lentiviral particles. The supernatant was collected at 48 h post-transfection, filtered through a 0.45 μm filter, and used to infect mESCs. After 48 h post-infection, cells were selected with 2 μg/mL puromycin. Single colonies were picked and expanded. Genomic DNA was extracted from each clone and assessed using PCR amplification followed by Sanger sequencing of the target locus. Loss of target protein expression was further confirmed by immunoblotting.

Phase separation assays

The purified H2B-K108AcK-His₆ proteins were dialyzed in no salt buffer (50 mM Tris-HCl, pH 7.4) to remove redundant imidazole and concentrated. For the H2B-K108AcK-His₆ phase assay, the H2B-K108AcK-His₆ proteins were centrifuged at 16,000 g for 10 min to remove potential insoluble aggregates and added into Tris buffer (50 mM Tris-HCl, 125 mM NaCl, 10% glycerol, pH 8.0) with 10% Dextran-70 to a final concentration of 3 µM. To test if DNA or RNA affects puncta formation of H2B-K108AcK-His₆ proteins, an appropriate concentration of genomic DNA or RNA was added into the above solution^91,92. The double-sided tape was folded, punched with holes, and taped on a glass slide to create a chamber. The mixed solution was added into the chamber, covered with a cover glass, and observed by an Olympus upright microscope (Bx53) DIC.

Mass spectrum analysis

For MS analysis of H2B-K108DiZPK-Flag-StrepII/H2B-K108AcK-Flag-StrepII/H3.3-K56AcK-Flag, proteins were separated by SDS-PAGE and collected for in-gel digestion as descript before⁹³. In brief, gel pieces were destained and reduced with 10 mM dithiothreitol (DTT) at 56 °C for 1 h, then alkylated with 55 mM iodoacetamide (IAA) under dark at room temperature for 30 min. Digestion was performed using sequencing-grade modified trypsin overnight at 37 °C. The digested peptides were desalted using homemade C18 StageTips, resuspended in 0.1% formic acid, separated on a 25-cm C18 column, and analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) on an Orbitrap Exploris 480 mass spectrometer (Thermo Fisher Scientific) coupled to an EASY-nLC 1200 system.

Endogenous histones were extracted under acidic conditions, reduced with TCEP, and alkylated as described above. Samples were alkylated, and four volumes of methanol, three volumes of water, and one volume of chloroform were sequentially added, followed by centrifugation at 10,000 g for 10 min at room temperature. Four volumes of methanol were added to the pellet. The precipitate was washed and air-dried. Proteins were then digested overnight with trypsin, followed by an additional 4 h digestion using a fresh aliquot of trypsin the next day. For acetylation analyses of endogenous histone, digested peptides were incubated with anti-acetyl-lysine agarose beads at 4 °C for 4 h with gentle rotation. Beads were washed with NETN buffer and eluted with 0.1% formic acid. The peptides were desalted as described above.

Full MS spectra were acquired in the Orbitrap analyzer over an m/z range of 350-1800 with a resolution of 60,000, an automatic gain control (AGC) target of 300%, and a maximum injection time of 50 ms. MS/MS spectra were generated using higher-energy collisional dissociation (HCD) with a normalized collision energy (NCE) of 30% and acquired at a resolution of 15,000, with an AGC target of 75%, the maximum injection time was 22 ms, and the isolation window was 1.6 m/z.

The raw MS data were analyzed using MaxQuant (version 1.6.1.0) with the UniProt mouse reference database (downloaded on 16 July 2019). Trypsin was specified as the digestion enzyme with up to 2 missed cleavages. Carbamidomethylation was set as a fixed modification, and oxidation (M) and acetylation (protein N-term) as variable modifications. FDR was set to 1% at the protein and peptide level. LFQ was used for quantification. Acetylation sites from reverse sequences and potential contaminants were removed. For interactome analysis, proteins identified in at least two of the three biological replicates and exhibiting > 1.5-fold enrichment relative to wild-type H2B controls were retained for downstream analysis. Statistical significance values served as complementary information but were not used as the primary filtering criterion. Missing LFQ intensity values (reported as zero by MaxQuant) were replaced with a pseudovalue corresponding to one-half of the minimum non-zero LFQ intensity detected in the dataset. This imputation was applied to enable quantitative comparison and fold-change calculation for proteins undetected in one experimental group. Non-zero LFQ intensity values were left unchanged. Fold change was calculated as the ratio of the mean protein abundance between groups. P values were calculated using a two-sided Student’s t-test. The same statistical results were further used for Gene Ontology (GO) enrichment analysis.

mESC differentiation

For EB differentiation, mESCs were dissociated into single cells and resuspended in ES differentiation medium (mESC culture medium without LIF and Embryomax Nucleosides) at a density of 1 × 10⁵ cells/mL. A total of 2 × 10³ cells were seeded per well into ultra-low attachment 6-well dishes containing 2 mL of ES differentiation medium. Cultures were maintained at 37 °C under 5% CO₂, with medium replaced every other day. EBs were collected at designated time points for subsequent analyses.

Neural differentiation was performed according to a published protocol⁹⁴ with minor modifications. Briefly, mESCs were seeded at 1 × 10³ cells per well in Matrigel-coated 12-well plates and cultured in mESC medium containing LIF for 3–5 days to allow the formation of compact colonies. Neural induction was initiated by switching to Medium I (DMEM/F12 supplemented with 1× B27, 1× N2, 1× NEAA, 0.1 mM β-mercaptoethanol, 100 ng/mL Noggin, 20 μM SB431542, 2 μM dorsomorphin, and 3 μM CHIR99021) and maintained for 4 days with daily medium changes. At day 5, rosettes were detached using 1.5 mg/mL collagenase IV and transferred to uncoated plates containing Medium II (DMEM/F12 supplemented with 1× N2, 20 ng/mL FGF2, and 1.6 g/L glucose) for neurosphere formation over the next 4 days. The neurospheres were collected, dissociated into single cells using Accutase, and replated onto Matrigel-coated plates in Medium III (DMEM/F12 supplemented with 1× N2, 0.1× B27, 20 μg/mL insulin, 1.6 g/L glucose, 20 ng/mL EGF, and 20 ng/mL FGF2). After 24 h in Medium III, cells were fixed for subsequent analysis.

Immunofluorescence staining

Cells were rinsed with cold PBS and fixed with 4% paraformaldehyde at room temperature for 40 min. Permeabilization and blocking were performed using PBS containing 10% goat serum and 0.5% Triton X-100 for 1 h at room temperature. Primary antibodies were diluted in PBS with 2% goat serum and incubated overnight at 4 °C. After incubation, samples were washed 3–5 times with PBS and then incubated with appropriate species-specific fluorescent secondary antibodies for 1 h at room temperature in the dark. Samples were subsequently washed 3–5 times with PBS and rinsed three times with distilled water. Coverslips were mounted using mounting medium containing DAPI. Fluorescent images were acquired using a Zeiss LSM900 confocal microscope. To quantify the colocalization between Pol II and H2B-WT versus H2B-K108AcK, Pearson’s correlation coefficient was computed. Statistical comparisons between the H2B-WT and H2B-K108AcK groups were then performed using Student’s t-test on the derived correlation coefficients.

For the Act D treatment experiment, cells were treated with 2.5 µg/mL Act D for 8 h before immunofluorescence staining.

For the 1,6-HD experiment, cells were treated with 1.5% 1,6-HD for 5 min. For the recovery experiment, 1,6-HD-treated cells were washed and cultured in fresh medium for an additional 8 h before staining.

Teratoma formation and IHC, HE staining

Seven-week-old female NCG mice (NOD/ShiLtJGpt-Prkdc^em26Cd52Il2rge^m26Cd22/Gpt; GemPharmatech, Cat. No. T001475) were used in this study. All animal procedures were approved by the Institutional Animal Care and Use Committee of West China Hospital, Sichuan University (Approval No. 20230111007), and performed in accordance with institutional guidelines and national regulations for laboratory animal care and use.

Mice were group-housed (up to four per cage) in autoclaved individually ventilated cages under specific pathogen-free (SPF) conditions with a 12 h light/12 h dark cycle and ad libitum access to food and water. All animals were confirmed healthy and had not undergone any previous experimental procedures or drug treatments.

Approximately 3 × 10⁶ mESCs suspended in DMEM were subcutaneously injected into the groin region of 8-week-old female NCG mice. Tumor formation and progression were monitored weekly. Four weeks post-injection, teratomas were surgically excised, fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned for hematoxylin and eosin (H&E) staining and immunohistochemistry (IHC) analysis. The teratoma tissues were stored at −80 °C for further analysis. H&E staining was performed at Sichuan Youngster Technology Co., Ltd., and IHC was conducted at Bio-Longyan Pathology Service Co., Ltd.

Single-cell RNA sequencing and data analysis

Single-cell RNA sequencing

Cells were dissociated into a single-cell suspension using Accutase and then filtered through a 40 μm cell strainer. Cell viability and total cell counts were assessed using a cell counter (LoGoS). Dead cells were removed using a Dead Cell Removal Kit. Single-cell RNA libraries were prepared using the 10×Genomics 3’ Single Cell RNA-seq Kit according to the manufacturer’s instructions. The libraries were amplified and subjected to paired-end sequencing on an Illumina platform (llumina HiSeq PE150 or NovaSeq PE150) at Novogene Technologies.

Data processing and alignment

Single-cell RNA sequencing data were processed using the 10x Genomics pipeline. Raw sequencing reads were aligned to the human reference genome (mm10) and quantified using Cell Ranger (v8.0.0) with default parameters. The alignment included removal of polymerase chain reaction duplicates and quality filtering to generate a gene-cell count matrix for subsequent analysis.

Quality control, dimensionality reduction, clustering, and cell type annotation

The Seurat package (v5.3.0) was used for downstream analyses. Cells with unique feature counts less than 2500 or greater than 9000 were filtered out to remove potential doublets and low-quality cells. Additionally, cells with mitochondrial gene content exceeding 5% were excluded to eliminate compromised cells. Gene expression counts were normalized using the ‘LogNormalize’ method with a scale factor of 10,000. The top 2000 highly variable genes were identified using the ‘vst’ method for downstream dimensionality reduction analysis. Principal component analysis (PCA) was performed on scaled data using the highly variable genes. The top 20 principal components were selected based on the elbow plot for downstream analysis. Cells were embedded in two dimensions using uniform manifold approximation and projection (UMAP). Cell clustering was performed using the Louvain algorithm with a resolution parameter of 1 to identify distinct cell populations. Cell clusters were annotated based on the expression of well-established marker genes.

Differential Expression Analysis. Differentially expressed genes between conditions or cell types were identified using the Wilcoxon rank sum test implemented in Seurat. Genes with an adjusted p < 0.05 and an absolute log₂FC > 0.58 were considered statistically significant.

Pseudotemporal analysis

Pseudotemporal ordering of cells was performed using Monocle2 (v2.34.0) to reconstruct developmental trajectories. Data were normalized with estimateSizeFactors and dispersion with estimateDispersions. Dimensionality reduction was performed using DDRTree (max_components = 2). Cells were ordered along pseudotime with orderCells using differentially expressed genes across clusters.

Flow cytometry analysis and cell sorting

Cells were dissociated into single-cell suspensions using complete culture medium to terminate trypsinization, followed by removal of residual trypsin. The cell suspension was adjusted to a concentration of approximately 2–3 × 10 ⁶ cells/mL and maintained on ice before flow cytometric sorting. Prior to sorting, cell suspensions were passed through a 40 μm cell strainer to remove cell debris and aggregates.

Flow cytometric sorting was performed at MoFlo Astrios EQ flow cytometer equipped for fluorescence-activated cell sorting (FACS). Cells were collected into 1.5 mL low-binding microcentrifuge tubes. The sequential gating strategy was applied as follows: dead cells and debris were excluded based on forward scatter (FSC)/side scatter (SSC) characteristics, followed by removal of cell doublets or aggregates using singlet gating. GFP-positive cells were subsequently identified and sorted for downstream analyses. The complete sequential gating and sorting strategies were shown in Supplementary Fig. 6.

CUT&Tag-seq

For Cut&Tag analysis, cells expressing either H2B-EGFP-3×Flag or the H2B-K108AcK-EGFP-3×Flag were sorted for EGFP-positive populations using a Beckman Colter flow cytometer. CUT&Tag libraries were prepared using the Novoprotein CUT&Tag Kit according to the manufacturer’s instructions. Briefly, live sorted cells (1×10⁵) were incubated with an anti-Flag antibody (Abmart, Cat# 3M20008, diluted 1:200) at 4 °C for 12 h. A pAG-Tn5 transposase complex was assembled in situ.

The quality of libraries was evaluated using a Qsep100 Bio-Fragment Analyzer (BiOptic Inc.), which revealed a fragment size distribution predominantly between 200 and 700 bp, indicating successful library preparation. Paired-end sequencing (2 × 150 bp) was performed on the Illumina NovaSeq 6000 platform (Haplox Genomics).

ATAC-seq

Cells expressing either H2B-EGFP-3×Flag or the mutant H2B-K108AcK-EGFP-3×Flag were sorted based on EGFP-positive using a Beckman Colter flow cytometry. ATAC-seq libraries were prepared using the Novoprotein Chromatin Profile Kit for Illumina (N248) following the manufacturer’s instructions. Briefly, nuclei were isolated from 5 × 10⁴ cells and subjected to Tn5 transposase-mediated fragmentation and adapter incorporation. The fragmented DNA was then purified and amplified by PCR to generate sequencing-ready libraries. Amplified libraries were purified to remove primer dimers and other contaminants, and quality was evaluated as described for the CUT&Tag libraries. Sequencing was performed on the Illumina NovaSeq 6000 platform (Haplox Genomics) using paired-end reads (2×150 bp).

CUT&Tag-seq and ATAC-seq data analysis

Pair-end (PE) sequencing reads were processed through a standardized workflow for quality control, filtering, and adapter trimming to yield clean reads, utilizing fastp and Cutadapt. These reads were aligned to the mm10 mouse reference genome with Bowtie2 under the following parameters (--end-to-end --very-sensitive --no-mixed --no-discordant --phred33 -I 10 -X 500) to produce BAM files. High-quality mapped reads were extracted using the samtools with a mapping quality threshold of 40 (-q 40). Duplicate reads were eliminated using the Picard toolkit. Only uniquely mapped and deduplicated reads with a quality score exceeding 40 were retained for subsequent analysis.

Scale factors were calculated using counts of uniquely mapping reads. Coverage plots and heatmaps were generated by normalizing read coverage with the scale factor and converting BAM files to bigWig format using DeepTools. BAM files were transformed into normalized bedGraph files via the BEDTools genomecov command, utilizing the previously calculated scale factor. Peaks were identified from normalized bedGraph files with SEACR, employing a non-stringent setting of 0.05. High-confidence peaks were determined as consensus peaks between two replicates using the BEDTools intersect command. Differential peaks were assessed by merging peaks from distinct groups based on read coverage using bedtools. Coverage for each peak was normalized with the previously calculated scale factor. Genomic localization of peaks was annotated via HOMER annotatePeaks.pl. Peak-associated genes were identified as the nearest genes annotated by HOMER to the respective peaks.

RNA-seq

Total RNA was isolated using Trizol⁹². Briefly, 1 mL of Trizol was added directly to cell pellet, and the lysate was homogenized by vortexing and pipetting until no visible clumps remained. The mixture was incubated at room temperature for 5 min to ensure complete dissociation of nucleoprotein complexes. Chloroform (200 μL per sample) was added, and the tubes were shaken vigorously for 15 s until the solution became homogeneous, followed by a 5 min incubation at room temperature. Phase separation was achieved by centrifugation at 12,000 g for 15 min at 4 °C. The aqueous phase was carefully transferred to a new RNase-free microcentrifuge tube, and RNA was precipitated by adding an equal volume of isopropanol. Samples were mixed gently by inversion and incubated on ice for 10 min, then centrifuged at 12,000 g for 10 min at 4 °C. The RNA pellet was washed twice with 1 mL of 75% ethanol (prepared with DEPC-treated water) and centrifuged at 12,000 g for 5 min at 4 °C. The supernatant was discarded, and the pellet was air-dried at room temperature for 2–5 min. Finally, the RNA was dissolved in an appropriate volume of DEPC-treated water. For mRNA-seq, poly(A)⁺ RNA was enriched using an oligo(dT) primer and used to construct sequencing libraries. Library preparation and high-throughput sequencing were carried out by Majorbio, Novogene, and Personalbio.

Quantitative real-time PCR

Total RNA was extracted from cells using the FOREGENE Cell Total RNA Isolation Kit. cDNA was synthesized from extracted RNA using HiScript® II Q RT SuperMix. qPCR was performed using SYBR qPCR Master Mix on a Bio-Rad CFX96 PCR system. Mouse Gapdh was used as a control. Quantification was calculated using the 2^-∆∆CT method and displayed as fold change. The following primer sequences were used for RT-qPCR. The primers used for qRT-PCR are listed in Supplementary Data 2.

ChIP-qPCR

Cells were collected and crosslinked with 1% formaldehyde in PBS at 25 °C for 8 min. The reaction was quenched with 125 mM glycine on ice. Samples were centrifuged and resuspended in lysis buffer I (50 mM HEPES-KOH, 140 mM NaCl, 1 mM EDTA, 10% glycerol, 0.25% TritonX-100, 0.5% NP-40, pH 7.5), buffer II (10 mM Tris-HCl, 200 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, pH 8.0), and sonication buffer (20 mM Tris-HCl, 150 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% TritonX-100, pH 8.0), and then subjected to ultrasonic fragmentation at 40 W. After centrifugation at 16,000 g for 10 min, take a 10% aliquot of the sample as input. The left samples were divided into two parts and incubated with anti-Flag or anti-IgG antibodies in the presence of protein A/G magnetic beads overnight with rotation at 4 °C. Beads were washed with low-salt ChIP buffer (20 mM Tris-HCl, 150 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% TritonX-100, pH 8.0), high-salt ChIP buffer (20 mM Tris-HCl, 500 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% TritonX-100, pH 8.0), LiCl wash buffer (10 mM HEPES-KOH, 500 mM LiCl, 1 mM EDTA, 0.7% sodium deoxycholate, 1% NP40, pH 8.0), and TE buffer (10 mM Tris-HCl, 1 mM EDTA, 50 mM NaCl, pH 8.0) twice. Samples were eluted with elution buffer (50 mM Tris-HCl, 10 mM EDTA, 1% SDS, pH 8.0) and treated with 0.2 mM NaCl, 10 µg RNase A, and 30 µg proteinase K at 65 °C for 4-6 h. DNA was extracted with Tris-saturated phenol, 1-Br-3-Chloropropane, and isoamyl alcohol at a volume ratio of 25:24:1 and analyzed by qPCR using the following primers. Quantification was calculated using the 2^-∆∆CT method and displayed as fold change⁹². The primers used for ChIP-qPCR are listed in Supplementary Data 2.

Statistics and reproducibility

All experiments were independently repeated at least three times unless otherwise stated. Statistical analyses were performed using GraphPad Prism software. Comparisons between two groups were conducted using the unpaired Student’s t-test. For comparisons among four groups, one-way analysis of variance (ANOVA) was applied, followed by Tukey’s post hoc test for multiple comparisons. A p-value of less than 0.05 was considered statistically significant. For RNA-seq data, differential expression was assessed by DESeq2 using a two-sided Wald test with Benjamini-Hochberg (BH) correction for multiple testing. For GO and KEGG enrichment analyses, statistical significance was assessed using a one‑sided hypergeometric test (Fisher’s exact test) with Benjamini–Hochberg correction; the figure displays -log10 (raw P) and enrichment score, with FDR < 0.01 and overlap ≥ 3 as thresholds. Adjusted q-values or FDR are listed in Source Data. For proteomics data, differentially expressed proteins were identified based on fold change and P values calculated using a two-sided Student’s t-test. No adjustment for multiple comparisons was applied. Representative images, including immunoblots and microscopy images, were obtained from three independent experiments with similar results.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Supplementary information

Supplementary Information^{(11.9MB, pdf)}

41467_2026_77439_MOESM2_ESM.pdf^{(68.5KB, pdf)}

Description of Additional Supplementary Files

Supplementary Data 1-2^{(27.1KB, xlsx)}

Reporting Summary^{(127.8KB, pdf)}

Transparent Peer Review file^{(828.1KB, pdf)}

Source data

Source data^{(25.7MB, xlsx)}

Acknowledgements

We thank the members of the Mass Spectrometry Center of West China Hospital (Rui Wang and Xiyu Wu) for proteomic techniques and data interpretation. We thank Tingting Guo from the Respiratory Health Research Institute of West China Hospital for flow analysis and sorting techniques.

Author contributions

F.N.C., Q.H., and L.Z. designed and performed the experiments. F.N.C., Q.H., L.Z., Y.J.R., Y.P.Z., W.C., Z.H.X., and H.Y.R. designed the experiments and analyzed the results. H.Y.R. and Z.H.X. wrote the manuscript. H.Y.R. performed all project coordination, wrote the manuscript, and secured funding.

Peer review

Peer review information

Nature Communications thanks Han Xiao and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Funding

H.Y.R. discloses support for the research of this work from National Natural Science Foundation of China [grant number 32271315]. W.C. discloses support for the research of this work from National Science Fund for Distinguished Young Scholars [grant number 32225001]. H.Y.R. and W.C. disclose support for the research of this work from the 1.3.5 project for disciplines of excellence, West China Hospital, Sichuan University [grant numbers ZYYC23021 and ZYGD23018]. F.N.C. discloses support for the research of this work from China Postdoctoral Science Foundation [grant number GZC20231839]. All other authors declare no relevant funding.

Data availability

The scRNA-seq data generated in this study have been deposited in the Genome Sequence Archive (GSA) database under accession code CRA030465. The CUT&Tag-seq data generated in this study have been deposited in the GSA database under accession code CRA030446. The H2B-K108AcK bulk RNA-seq data generated in this study have been deposited in the GSA database under accession code CRA030444. The Rps19bp1-KO teratoma bulk RNA-seq data generated in this study have been deposited in the GSA database under accession code CRA030359. The Rps19bp1-KD bulk RNA-seq data generated in this study have been deposited in the GSA database under accession code CRA030443. The ATAC-seq data generated in this study have been deposited in the GSA database under accession code CRA030445. The Mass spectrometry data generated in this study have been deposited in the integrated proteome resources (iProX) under accession code PXD068269. The remaining data are available within the paper and the supplementary information or Source Data file. Source data are provided with this paper. Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request. Source data are provided with this paper.

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

These authors contributed equally: Fangni Chai, Qin Huang, Li Zhou.

Contributor Information

Zhihong Xue, Email: xuez@scu.edu.cn.

Haiyan Ren, Email: hyren@scu.edu.cn.

Supplementary information

The online version contains supplementary material available at https://doi.org/10.1038/s41467-026-77439-7.

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PERMALINK

Site-specific programming characterizes dynamic post-translational acetylation of histone H2B lysine 108 in mouse embryonic stem cells

Fangni Chai

Qin Huang

Li Zhou

Yijia Ren

Yanping Zhong

Wei Cheng

Zhihong Xue

Haiyan Ren

Abstract

Introduction

Results

Develop the site-specific toolkits in mESCs

Fig. 1. Development of Site-specific toolkits in mESCs.

Site-specific system mediate ncAAs incorporation into histones in living mESCs

Site-specific-Link revealed Rps19bp1 as a direct interacting partner of H2B-K108DiZPK in mESCs

Fig. 2. Proteomic analysis of H2B-K108DiZPK interactome and revealed Rps19bp1 regulating its deacetylation in mESCs.

Rps19bp1 promotes deacetylation of H2B K108 in a Sirt1-dependent manner

The Rps19bp1-H2B-K108ac cascade plays important roles in mESC differentiation

Fig. 3. The Rps19bp1-H2B-K108ac cascade plays important roles in mESC differentiation.

H2B-K108 acetylation drives puncta formation and rewrites molecular interactions uncovered by the Site-specific-AcK tool

Fig. 4. The Site-specific-AcK tool revealed H2B-K108 acetylation induces puncta formation and alters the interactome.

H2B-K108 acetylation modulates chromatin accessibility

Fig. 5. H2B-K108 acetylation modulates chromatin accessibility in mESCs.

Rps19bp1 deficiency promotes neural differentiation in mESC-derived teratoma

Fig. 6. Rps19bp1 deficiency promotes neural differentiation in mESC-derived teratoma.

Discussion

Methods

Reagent and resources

Cell culture and transfection

Plasmids

Recombinant protein expression and purification

Site-specific-Link

Site-specific-AcK

Acid extraction of histones from mESCs

Fraction of chromatin

In vitro deacetylation assay

Immunoprecipitation and immunoblotting

Pulldown assays

Microscale thermophoresis (MST) assay

Construction of monoclonal mESCs

Construction of knockdown cells

Construction of knockout cells

Phase separation assays

Mass spectrum analysis

mESC differentiation

Immunofluorescence staining

Teratoma formation and IHC, HE staining

Single-cell RNA sequencing and data analysis

Single-cell RNA sequencing

Data processing and alignment

Quality control, dimensionality reduction, clustering, and cell type annotation

Pseudotemporal analysis

Flow cytometry analysis and cell sorting

CUT&Tag-seq

ATAC-seq

CUT&Tag-seq and ATAC-seq data analysis

RNA-seq

Quantitative real-time PCR

ChIP-qPCR

Statistics and reproducibility

Reporting summary

Supplementary information

Source data

Acknowledgements

Author contributions

Peer review

Peer review information

Funding

Data availability

Competing interests

Footnotes

Contributor Information

Supplementary information

References

Associated Data

Supplementary Materials

Data Availability Statement

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