Canonical BAF chromatin remodeling complex specifies stem cell fate via cell-type-specific co-factor recruitment

Domain: chemistry
Published: 2026-03-03
Editorial outcome: minor_revision

Abstract
Individual subunits within the canonical BAF (cBAF) chromatin remodeling complex are known to regulate stem cell behavior, with some functional redundancy across subunits. Yet, how the cBAF complex directs adult stem cell fate specification and maintains stem cell niches remains unclear. Using the adult mouse incisor, we show that cBAF specifies mesenchymal stem cell (MSC) fate by recruiting distinct transcriptional co-factors to shape cell-type-specific chromatin regulation. Through single-cell multi-omics and in vivo functional analyses, we identify ARID1-containing cBAF as an essential gatekeeper that maintains the dynamic balance between MSC self-renewal and differentiation. Specifically, cBAF–DLX2 interactions preserve niche identity by remodeling intronic chromatin accessibility of niche-defining marker Runx2, while cBAF–FOXO1 directly modulates promoter accessibility of lineage-regulating transcription factors, including STAT3 and TRP53, among others, to balance progenitor proliferation and differentiation. Functional perturbation of RUNX2 and TRP53 confirms their roles downstream of cBAF in niche maintenance and fate specification. Our findings establish cBAF as a central regulator of adult stem cell niches and lineage commitment, and highlight cofactor-dependent mechanisms that may have broader implications for tissue regeneration and BAF-associated disorders.

Introduction
Epigenetic regulation orchestrates gene expression programs essential for cellular identity, lineage specification, and tissue maintenance without altering the DNA sequence. The mammalian SWI/SNF complex, a critical epigenetic regulator also known as the BAF (BRG1/BRM-associated factor) complex, dynamically modulates nucleosome positioning and chromatin accessibility in an ATP-dependent manner, enabling transcription factors and RNA polymerase II to access DNA sequences and regulate gene expression1-5. Dysregulation of the canonical BAF (cBAF) complex is prevalent across diverse pathological contexts in humans. Notably, mutations in cBAF subunits are involved in approximately 20% of human cancers and are linked to developmental disorders such as Coffin-Siris syndrome and autism spectrum disorders, highlighting their pivotal roles in cancer suppression, embryonic development, and tissue specification4,6-14. Recent findings further demonstrate that cBAF acts as a negative regulator of T cell fate by governing enhancer landscapes in activated CD8+ T cells, thereby influencing transcription factor recruitment and suggesting a potential therapeutic target in immunotherapy15,16. Although individual roles of cBAF subunits have been studied across various biological contexts17-24, significant gaps remain in our understanding of the precise mechanisms by which cBAF modulates chromatin accessibility profiles and transcriptional programs in a cell-type-specific manner, particularly in adult tissues. Clarifying these mechanisms is vital not only to comprehend fundamental stem cell biology but also to identify novel therapeutic targets for diseases arising from aberrant chromatin remodeling or stem cell dysregulation.

The BAF chromatin remodeling complex exerts diverse, context-dependent functions across tissues, shaped by its interactions with lineage-specific transcription factors (TFs) and chromatin modifiers. In embryonic stem cells, the cBAF subunit BRG1 is recruited by core pluripotency factors (OCT4, SOX2, NANOG) and plays an important role in this transcriptional network essential for maintaining stemness25,26. In cancer, BAF acts as a tumor suppressor by partnering with key regulators, such as MYC and p53, to control gene expression and cell behavior27,28. In bone marrow, BAF is engaged by CUX1 to regulate hematopoietic lineage commitment10. These interactions confer transcriptional specificity, enabling BAF to direct tissue-specific gene regulation. Beyond these contexts, the TFs that interact with the BAF complex to control stem cell fate commitment warrant further investigation. FOXO transcription factors have been identified as BAF partners that facilitate chromatin remodeling during cellular stress responses and differentiation29. DLX family homeobox proteins are implicated in craniofacial and skeletal development, and their mutations are linked to neurodevelopmental disorders such as autism30. Whether FOXO and DLX proteins can serve as context-specific co-factors of cBAF in regulating stem cell fate and the molecular mechanisms underlying their cooperation remain largely uncharacterized.

Adult tissue homeostasis relies on the precise coordination of stem cell behavior within specialized microenvironments known as stem cell niches, where important interactions occur between niche cells, nerves, circulatory vessels, and other cellular components31-35. These niche interactions regulate critical processes, such as stem cell quiescence, proliferation, differentiation, and lineage commitment, to maintain continuous tissue renewal throughout an organism's lifespan36-39. Regenerative tissues, such as the hair follicle, intestine, and mouse incisor, exhibit conserved and spatially organized hierarchies of stem cells, transit-amplifying cells (TACs), niche cells, and differentiated cells. These tissues thus serve as excellent models for investigating mechanisms governing adult stem cell regulation and fate determination36,40-42. The mouse incisor model provides a uniquely accessible system with specific cellular markers, precise spatial organization, and well-defined differentiation trajectories, making it ideal for studying the regulation of stem cell niches and lineage commitment (Figure 1a). Previous studies established that ARID1A and ARID1B, mutually exclusive subunits of the cBAF complex, display distinct expression patterns within the mouse incisor. ARID1A promotes cell cycle exit in TACs, whereas ARID1B maintains stem cell quiescence17,19. These initial insights inspired us to ask questions about the crucial roles of cBAF within the stem cell tissue, such as how cBAF differentially regulates chromatin accessibility in stem cells, TACs, and niche cells; whether it recruits distinct transcriptional co-factors in distinct cell types; and how these interactions coordinate lineage-specific transcriptional programs. Addressing this critical knowledge gap is essential for understanding basic stem cell biology and tissue homeostasis, as well as for potential clinical translation targeting diseases associated with aberrant stem cell behavior or chromatin dysregulation.

Thus, our study systematically investigates the cell-type-specific regulatory mechanisms by which cBAF recruits co-factors to orchestrate adult mesenchymal stem cell fate commitment. By integrating single-cell multi-omic profiling and rigorous in vivo functional assays in the adult mouse incisor model, we reveal that cBAF remodels chromatin accessibility differently across various cell types. Our integrative analysis has uncovered co-factor-guided regulatory mechanisms of cBAF with cell-type-specific functions. DLX2 and FOXO1, as co-factors of cBAF, help remodel chromatin accessibility profiles to regulate precise transcriptional programs, thereby maintaining niche cell identity and balancing TAC proliferation and differentiation. This co-factor-guided regulatory framework is crucial to the roles of the cBAF complex in specifying stem cell fate, preserving stem cell niches, and maintaining tissue homeostasis and regeneration. Taken together, our findings improve understanding of how epigenetic regulators differentially determine stem cell fate, and the identified downstream targets provide a potential foundation for future therapies targeting chromatin remodeling complexes in areas such as tissue regeneration, cancer, and developmental disorders.

Results
Loss of cBAF significantly impairs adult tissue growth, homeostasis, and regeneration

Dual loss of ARID1A and ARID1B causes dissociation of the cBAF complex43, making the Arid1a and Arid1b double-knockout mouse model valuable for studying the fundamental role of cBAF. We first confirmed the expression patterns of ARID1A and ARID1B, which are widely expressed in the proximal region of the incisor tissue, with ARID1A specifically expressed in the TAC region and ARID1B expressed in the MSC region (Figure S1a-d). To investigate how cBAF complex plays its functional roles in the stem cell microenvironment and regulates lineage commitment in adult tissue homeostasis and regeneration, we deleted both Arid1a and Arid1b specifically in the GLI1+ MSC lineage in the continuously growing mouse incisor, by generating tamoxifen-inducible Gli1-CreER;Arid1afl/fl;Arid1bfl/fl/fl (DKO) mice. To study adult stem cell homeostasis, tamoxifen was administered at postnatal 4-5 weeks in mice, when mouse incisors are fully mature (Figure 1b). We began this study by confirming that Arid1a and Arid1b were efficiently deleted in the dental mesenchyme and epithelium of the mouse incisor 5 days post-tamoxifen induction (dpt) (Figure S1e-h).

To investigate the role of the cBAF complex in mouse incisor growth and post-injury regeneration, we first performed a notch movement assay to evaluate the growth capacity of DKO incisors. We found the growth rate of the incisor was markedly reduced in DKO mice compared to controls when measured 10 days after notch creation (Figure 1c,d). We next conducted an incisor clipping assay to assess the tissue injury repair ability following the loss of the cBAF complex. The control group fully restored the clipped incisor length within two days, whereas DKO mice achieved less than 50% restoration at the same time frame (Figures 1e,f). These findings indicate the essential roles of the cBAF complex in supporting tissue growth and regenerative capacity of adult mouse incisors.

Next, we examined changes in tissue morphology one week after loss of cBAF. We observed that the well-aligned odontoblasts and ameloblasts in the control were severely disrupted and disorganized in the DKO mouse incisors, and nuclei in the TAC region displayed a round morphology in the control, whereas they exhibited an elongated, oval-shaped morphology in the DKO mouse incisors (Figure 1g). Moreover, significantly less dentin and enamel, formed by odontoblasts and ameloblasts, respectively, were deposited in the DKO incisors compared to the controls. Together, these severe phenotypic abnormalities support the essential role of the cBAF complex in maintaining adult tissue homeostasis.

Given that GLI1+ cells contribute to both mesenchymal and epithelial lineages in the mouse incisor, we further explored whether the observed dental mesenchymal defects in DKO mice could be attributed to changes in the dental epithelial cells. To specifically delete Arid1a and Arid1b in the epithelial lineage, we generate Sox2-CreER;Arid1afl/fl;Arid1bfl/fl mice. While we noted slight misalignment in the ameloblast layer of Sox2-CreER;Arid1afl/fl;Arid1bfl/fl mice, no morphological defects were observed in mesenchymal tissue in these mice when compared to controls one week post-induction (Figure S1i-j,l-m). Also, odontoblast marker Dspp expression showed no obvious difference between the Sox2-CreER;Arid1afl/fl;Arid1bfl/fl mice and controls (Figure S1k,n). These findings suggest that the loss of the cBAF complex in dental epithelial cells does not affect the mouse incisor mesenchyme, supporting the essential role of the cBAF complex specifically in dental mesenchymal cells for maintaining incisor growth, injury repair, and adult tissue homeostasis.

cBAF preserves the MSC-TAC-niche cell domains and progenitor cell fate commitment

The MSCs give rise to TACs in the proximal region of the mouse incisor, supporting tissue growth and regeneration. In this region, a specialized niche cell population, marked by co-expression of RUNX2 and GLI1 (RUNX2+GLI1+), supports the transition from MSCs to TACs41. To reveal the cellular changes associated with the observed phenotypes after the loss of cBAF complex in the GLI1+ lineage, we performed single-cell RNA sequencing (scRNA-seq) on cells collected from the proximal ends of DKO mouse incisors at 4 days post-induction, when the MSC, TAC, and niche cell domains have been disturbed (Figure S2). We integrated the DKO (8182 cells) scRNA-seq data with our previously published control (9625 cells) scRNA-seq dataset and analyzed the cell clusters using Seurat software19,44,45. The cell clusters were annotated using published markers, and a large cell cluster was defined as dental mesenchymal cells (Figure S3a,b)19,46. We further subclustered the dental mesenchymal cells into five cell clusters: PDF (proximal dental follicle cells), MSC (mesenchymal stem cells), DP (dental pulp cells), TAC (transit-amplifying cells), and Pre-od (pre-odontoblasts) (Figure 2a, Figure S3c,d), corresponding to their distinct in vivo locations (Figure 2b). Because RUNX2+GLI1+ niche cells are a small subset of GLI1+ cells, it is difficult to precisely isolate the niche cell population in scRNA-seq data. Therefore, we did not include niche cells in the clustering; instead, we compared them using in vivo analysis. We compared the proportions of each cell type between control and DKO samples. TACs were found to be twice as numerous in the DKO samples compared to controls, whereas MSCs and the differentiated pre-odontoblasts and dental pulp cells showed slight reductions (Figure 2c). To determine whether stem cell lineage differentiation is affected by the loss of the cBAF complex, we performed pseudotime analysis on mesenchymal cell clusters from scRNA-seq data using the Monocle2 algorithm in an unbiased manner47,48. This analysis revealed the single-cell trajectories over time as MSCs transition into TACs and subsequently differentiate into pre-odontoblasts and dental pulp cells (Figure 2d,e). Throughout this differentiation process, the expression of the MSC marker Gli1 was initially high and gradually decreased, while markers for pre-odontoblasts (Notum) and dental pulp cells (Fmod) increased in later stages. The proliferation marker Mki67 peaked during the middle stages, preceding the rise in pre-odontoblast marker levels (Figure S3e). By comparing differentiation trajectories between control and DKO samples along the pseudotime, we observed an early accumulation of TACs in the DKO sample at a stage typically occupied by MSCs in the control samples. Additionally, there was a loss of differentiated pre-odontoblasts after the branching point, along with a reduction in differentiated dental pulp cells in the DKO samples compared to controls (Figure 2e). We compared gene expression dynamics along pseudotime trajectories between control and DKO samples to elucidate the cause of the changes in lineage commitment. The pseudotime heatmap compared the expression enrichment of 772 genes linked to differentiation trajectories between the control and DKO (Figure 2f). Notably, genes within the green module, which are usually activated at early pseudotime points in control samples, showed later activation in DKO samples, shifting their expression toward more differentiated cell populations following cBAF loss. Functional annotation revealed that these temporally dysregulated genes are predominantly involved in critical biological processes, including cell proliferation, differentiation, and apoptosis (Figure S3f). We therefore sought to verify whether these cellular processes were detected in the DKO mice. We observed a progressive reduction in GLI1+ MSCs and, unexpectedly, a decrease in TACs in the DKO mice at 5 dpt compared to controls (Figure S4a,b). Because our transcriptomic analysis revealed altered expression of apoptosis-related genes, we performed a time-course TUNEL assay to determine whether hyperproliferation at 3 dpt ultimately led to cell death. Apoptosis was first detected at 4 dpt in the TAC region and became pronounced by 5 dpt (Figure S4c,d), suggesting that hyperproliferation may trigger apoptosis and consequently reduce the number of proliferating cells from 3 dpt to 5 dpt. These findings highlight the dynamic cellular responses following loss of cBAF.

Three distinct spatial cellular domains, including MSCs, TACs, and niche cells, are well-defined under normal conditions. To simultaneously visualize these spatial domains in vivo, we performed co-staining for β-Galactosidase (β-GAL), RUNX2, and EdU (a 2-hour EdU pulse to label proliferating cells in S-phase) in Gli1-LacZ and DKO;Gli1-LacZ mouse incisors. We observed that the well-defined cellular domains are disrupted following the loss of the cBAF complex, with a reduced GLI1+ MSC population, widespread proliferating cells, and increased RUNX2+ cells (Figure 2g). This data indicates that the cBAF complex is indispensable for maintaining the proper spatial organization of cell domains in the proximal end of the mouse incisor.

To further investigate how the cBAF complex regulates MSC fate commitment, we performed lineage tracing of GLI1+ MSCs using tdTomato reporter mice. We generated DKO;tdTomato mice and compared their MSC differentiation capabilities to those of Gli1-CreER;tdTomato control mice. Analysis of tdTomato fluorescence revealed significantly fewer labeled dental pulp cells in DKO;tdTomato incisors compared to controls (Figure 2h), indicating compromised lineage differentiation of MSCs following loss of cBAF. Additionally, we specifically monitored the capacity of TACs to differentiate into odontoblasts by injecting EdU to label proliferating TACs during S-phase and analyzed them 48 hours later. We performed co-staining for the odontoblast differentiation marker Dspp and EdU, and overlapped cells indicated that the labeled TACs differentiated into odontoblasts. In control incisors, we clearly observed Dspp+EdU+ odontoblasts differentiated from labeled TACs. However, in DKO incisors, this differentiation was disturbed, with minimal overlap between EdU and Dspp staining (Figure 2i), indicating that the ability of TACs to differentiate into odontoblasts was completely impaired after the loss of cBAF. These findings suggest that the loss of the cBAF complex causes MSCs to fail in properly committing to differentiated lineages. These data collectively demonstrate that the cBAF complex is essential for organizing the stem cell microenvironment, including MSC, TAC, and niche cells, and for ensuring appropriate stem cell lineage progression, thereby supporting adult tissue homeostasis and regeneration.

cBAF remodels chromatin accessibility in a cell-type-dependent manner

The cBAF complex is an essential remodeler of chromatin structure, enabling RNA Polymerase II (RNAPII) and transcription factors (TFs) to access promoter and enhancer regions, thereby regulating downstream gene expression. However, how the cBAF complex precisely regulates chromatin accessibility across various cell types and whether it exerts cell-type-specific effects on maintaining homeostasis and facilitating injury repair in adult tissues remain unexplored in vivo. To investigate how cBAF regulates chromatin accessibility in adult stem cells and their progeny at the single-cell level, we performed single-cell ATAC-sequencing (scATAC-seq) on the proximal regions of DKO mouse incisors 4 days post-tamoxifen induction. The scATAC-seq data from the DKO samples were compared with data from control mouse incisors in our previously published datasets19. We integrated and analyzed 5,522 control and 5,663 DKO cells using Signac49. Strikingly, the integrated UMAP plot showed a distinct separation of cell clusters between control and DKO samples, highlighting a significant reshaping of the chromatin accessibility upon loss of the cBAF complex (Figure S5a). Pseudobulk analysis revealed 56,119 peaks open in control samples but closed or reduced in the DKO (closed peaks), alongside 17,194 peaks opened or enhanced in the DKO (opened peaks) (Figure 3a). Using ChIPseeker, we annotated these peaks and compared the genomic distributions of unchanged, closed, and opened peaks50,51. Notably, we found that changed peaks (including both opened and closed peaks) were more frequently located in intronic and distal intergenic regions (enhancers) than the unchanged peaks, suggesting that enhancers were preferentially affected by cBAF disruption (Figure S5b). Heatmap analysis further confirmed that both closed and opened chromatin regions were enriched at enhancers rather than promoters (Figure 3b). Together, these findings provide robust in vivo evidence that cBAF predominantly regulates chromatin accessibility at enhancer regions in the adult mouse incisor.

To characterize cell-type-specific chromatin accessibility patterns, we annotated cell clusters in control and DKO mesenchymal populations based on putative gene activities and established cell-type markers (Figure 3c, Figure S5c). Comparative analysis revealed that the loss of cBAF led to a large number of peaks closed in the TAC region, including TACs and pre-odontoblasts. In contrast, MSC, PDF, and DP populations displayed comparable numbers of closed and opened peaks following cBAF loss (Figure 3d). Integration of chromatin accessibility data with transcriptional profiles showed that genes associated with closed peaks were generally downregulated in DKO samples, while those linked to opened peaks were upregulated (Figure S5d,e). Further analysis of genomic distribution revealed that DKO samples exhibited stronger peak signals in promoter regions, while in the TAC populations, promoter regions displayed a mixed pattern with both enhanced and reduced peak signals (Figure S5f). Notably, enhancer regions across all populations showed a global reduction in accessibility upon loss of cBAF (Figure S5f). As chromatin accessibility at promoter regions reflects gene expression levels, we specifically compared gene expression associated with closed and opened promoter peaks. Promoter regions with reduced or closed peaks exhibited decreased gene expression, and vice versa (Figure S5g,h). These results suggest that the cBAF complex exerts differential regulatory effects on promoter and enhancer elements to fine-tune gene expression across distinct mesenchymal cell types.

To assess how the cBAF complex modulates chromatin accessibility in a cell-type-specific manner, we annotated closed and opened peaks across distinct mesenchymal populations. Using UpSet plots, we compared genes linked to differentially accessible regions among these cell types. For genes associated with closed peaks, 1,077 genes were commonly affected across all five populations, with the TAC and pre-odontoblast clusters exhibiting the highest numbers of uniquely changed genes, highlighting a pronounced role for cBAF in regulating gene expression within the TAC region (Figure S6a). In contrast, 814 genes were commonly associated with opened peaks, with the differentiated dental pulp cell population showing the most distinct gene expression. MSCs and adjacent PDF cells also showed substantial transcriptional changes associated with increased chromatin accessibility (Figure S6b). These findings indicate that cBAF exerts distinct regulatory influences on chromatin accessibility and transcriptional output in a cell-type-dependent manner within the adult incisor mesenchyme.

We next investigated how cBAF differentially regulates distinct domains, focusing on the altered populations in the proximal region, including MSCs, TACs, and niche cells. To specifically investigate how the cBAF differentially regulates MSCs and TACs, we compared closed and opened chromatin peaks between MSCs and TACs. We identified that 1,778 changed genes were associated with closed peaks and 1,125 genes were associated with opened peaks across these two cell types. Notably, closed peaks were predominantly enriched in TACs, while opened peaks were more prevalent in MSCs (Figure S6c,e). To determine the functional implications of these accessibility changes, we performed Gene Ontology (GO) analysis on the annotated genes. Among the genes in MSCs associated with closed peaks, which reflect genes controlled by cBAF, we found enrichment for pathways involved in Wnt signaling, connective tissue development, mesenchyme development, and sensory nerve system development. Meanwhile, genes associated with closed peaks in TACs were enriched for ossification, neurogenesis, cell junction assembly, and adhesion, indicating potential regulation of these processes by the cBAF complex (Figure S6d). These GO analyses indicated that the cBAF complex exhibits distinct functional roles within MSC and TAC populations. The opened peak analysis further highlighted altered pathways involved in chromatin remodeling changes in both MSCs and TACs. The changes were linked to disrupted cell cycle transitions and Wnt signaling in MSCs, while showing enrichment for double-strand break repair, apoptosis, and mitotic phase transition in TACs, consistent with their increased apoptosis observed in vivo (Figure S6f). To confirm that the identified signaling pathways are indeed influenced by cBAF, we selected Wnt signaling and DNA damage for in vivo validation. In situ hybridization for Axin2, a readout of the Wnt pathway, revealed ectopic activation in MSCs and DP cells, along with upregulation in TACs, in the DKO samples compared to controls (Figure S6g), suggesting the role of cBAF complex in restricting Wnt signaling activation in TACs. Immunostaining for γH2AX, a marker of DNA double-strand breaks, showed significantly increased DNA damage in TACs of DKO samples compared to controls (Figure S6h), indicating that loss of cBAF triggers DNA double-strand breaks. These data confirm that cBAF plays distinct and critical roles in maintaining the chromatin accessibility of MSCs and TACs, as well as their associated cellular functions.

To investigate how the cBAF complex modulates chromatin accessibility and specifically regulates niche cells, we isolated RUNX2+GLI1+ cells from the MSC population and performed comparative chromatin accessibility analysis. Heatmap profiling revealed approximately twice as many regions with reduced accessibility (closed peaks) as with increased accessibility (opened peaks) following cBAF loss (Figure 3e). GO analysis of these differentially accessible regions showed substantial overlap with the signaling pathways affected in the broader MSC population (Figure 3f). Notably, morphogenesis and cell projection organization were enriched among the top signaling pathways identified in closed peaks in niche cells, suggesting unique chromatin remodeling events in these cells. Furthermore, pathways associated with extracellular stimulus response and cilium organization were selectively impaired in the niche cell population, underscoring the critical role of cBAF in maintaining the functional integrity of the stem cell niche.

Together, these findings indicate that cBAF exerts context-dependent regulatory control over chromatin accessibility across distinct cell populations. This cell-type-specific modulation may be driven by differential recruitment of co-factors, enabling cBAF to finely tune transcriptional programs within distinct cell types and to preserve tissue homeostasis in adult regenerative tissues.

Transcriptional co-factors, FOXO1 and DLX2, determine the regulatory specificity of cBAF

To identify potential co-factors interacting with cBAF in the cell populations of the proximal incisor, we conducted motif analysis of the control-specific (closed) peaks identified from our scATAC-seq data. Using HOMER motif analysis, we found that the top-enriched motifs were similar in promoter and enhancer peaks, indicating that the DNA-binding preferences of cBAF are consistent across different genomic contexts. The top 10 motifs belonged to the DLX, FOX, and RUNX families, which are AT-enriched, confirming that the cBAF complex recognizes AT-enriched DNA sequences irrespective of their binding regions (Figure 4a, Figure S7a). To further explore whether the identified TFs are potential co-factors of cBAF in proximal mesenchymal cells, we evaluated their expression patterns in scRNA-seq datasets (Figure S7b). In the DLX family, we found that Dlx1, Dlx2, Dlx3, and Dlx5 are widely expressed across different cell types, although fewer signals are detected in dental pulp cells for Dlx1, Dlx2, and Dlx5. In the FOX family, we found that Foxo1 and Foxo3 are expressed at low levels in MSCs and TACs but are more abundantly expressed in dental pulp cells, whereas Foxk1, as well as Isl1, are rarely detected in any of these populations. Regarding Runx1 and Runx2, we observed that Runx1 is barely detectable, while Runx2 is mainly expressed in the proximal dental follicle, consistent with previous studies that identify Runx2 as a niche cell marker in the mouse incisor41. The widely expressed and barely expressed genes will be excluded from further analysis.

To verify whether these TFs physically interact with ARID1A-containing and/or ARID1B-containing cBAF, we performed co-immunoprecipitation (Co-IP) to identify protein-protein interactions. We found that DLX2 and FOXO1 can both interact with ARID1A, whereas only DLX2 interacts with ARID1B (Figure 4b, Figure S7c). We then performed in situ hybridization to reveal their expression patterns in vivo. Foxo1 is expressed in dental pulp cells and less in TACs, while Dlx2 is expressed in MSCs and TACs, as well as in dental pulp (Figure 4c). To further confirm the functional binding activities of these TFs, we utilized TF footprinting analysis with scATAC-seq data. TF footprinting appears as induced transposase (Tn5) insertion rates at sites of direct protein-DNA interactions, providing precise single-cell-level evidence of TF occupancy52. Footprinting analysis revealed clear dips indicative of strong TF binding in both control and DKO samples. However, the control cell types (represented by yellow-green lines) displayed higher Tn5 insertion than the DKO cell types (represented by pink-blue lines), suggesting impaired binding activity of DLX2 and FOXO1 following cBAF loss (Figure 4d). To substantiate these observations at the molecular level, we performed Cleavage Under Targets & Release Using Nuclease sequencing (CUT&RUN-seq) for ARID1A, ARID1B, FOXO1, and DLX2 using control incisor tissue. Since ARID1A and ARID1B are DNA-binding proteins that lack a specific binding motif, we analyzed the CUT&RUN-seq results to identify their tissue-specific binding sites within the proximal end of the adult mouse incisor (Figure S7d). Genomic distribution analysis showed that ARID1A and FOXO1 exhibit more promoter regulatory activity than ARID1B and DLX2, and our analysis revealed a subset of target genes shared by these factors (Figure S7e-g). To verify whether these co-factors could interact with ARID1A and ARID1B in specific cell types, we focused on the intersections between TAC-specific closed peaks and ARID1A binding sites, as well as MSC-specific closed peaks and ARID1B binding sites. We confirmed that FOXO1 and DLX2 co-occupy ARID1A binding peaks, indicating their collaborative regulatory role in TACs. Similarly, DLX2 co-localizes with ARID1B binding peaks in MSCs, supporting its function as a co-factor for ARID1B-containing cBAF in the MSC population (Figure 4e,f). These findings suggest that cBAF recruits distinct co-factors to regulate chromatin accessibility profiles differently across various cell types.

cBAF complex with co-factor DLX2 safeguards stem cell niche composition

To further delineate how cBAF recruits distinct co-factors to define the niche cell-MSC-TAC domains, we revisited our previous findings that established individual roles for ARID1A and ARID1B. Specifically, ARID1A regulates TAC cell cycle exit, while ARID1B maintains MSC quiescence, as reported in prior work17,19.

Extending this work, we aimed to determine whether the cBAF complex specifically regulates the niche cell population. Given that we observed an increase in RUNX2+ cells in the DKO incisors compared with the controls, we first examined whether cBAF directly regulates Runx2 expression. Using CUT&RUN-seq data at the Runx2 gene locus, we observed that both ARID1A and ARID1B, with co-factor DLX2, show strong binding activity in the intronic region of Runx2 (Figure 5a). We also performed FOXO1 and DLX2 CUT&RUN sequencing using DKO mouse incisor tissue and found that DLX2 binding activity was lost upon cBAF loss, confirming DLX2 as a cBAF-dependent cofactor in regulating Runx2 expression (Figure 5a). To further validate the regulatory role of cBAF complex, we examined chromatin accessibility at the Runx2 locus using scATAC-seq data from control and DKO samples. Our data showed open chromatin peaks at the ARID1A/ARID1B/DLX2 binding site of Runx2 in control samples, while these peaks were significantly reduced in DKO cell clusters, indicating that the cBAF complex regulates the chromatin accessibility in the intronic region of Runx2 to suppress its expression (Figure 5a,b). Additionally, the chromatin accessibility of the Runx2 promoter region increased in DKO samples, confirming elevated Runx2 transcriptional activity following the loss of the cBAF complex. (Figure 5a). We then confirmed Runx2 upregulation in vivo by immunostaining. The result showed obvious ectopic activation of Runx2 in mesenchymal cells, especially in MSCs, in the DKO compared to the control (Figure 5c). To validate that the Runx2 intron is a functional regulatory element, we conducted a luciferase reporter assay. We detected significantly increased Gluc activity in the DKO incisor primary cell culture compared to controls (Figure S8a), indicating that cBAF suppresses Runx2 transcriptional activity by binding to the intronic region. To assess DLX2's role in regulating Runx2 expression, co-staining of RUNX2 and Dlx2 revealed mutually exclusive expression patterns (Figure 5d). Meanwhile, we knocked down Dlx2 with siRNA in control primary cells and observed an increase in Runx2 expression (Figure 5e). These data indicate that the presence of the cBAF complex recruits DLX2 to suppress Runx2 expression in mesenchymal cells by binding to its intron.

To test whether ARID1A and ARID1B are functionally redundant in regulating Runx2 expression, we performed bulk RNA sequencing of DKO samples and conducted GSA analysis comparing them to our published control samples19. We also checked Runx2 expression from the published bulk RNA sequencing results for control and Gli1-CreER;Arid1afl/fl (Arid1a KO) mice, as well as for control and Gli1-CreER;Arid1bfl/fl (Arid1b KO) mice17,19. Runx2 expression did not show significant changes between control and Arid1a KO or Arid1b KO, but it showed significant upregulation in the DKO (Figure S8b). We also performed immunostaining for RUNX2 in control, Arid1a KO, and Arid1b KO incisors, confirming that there was no obvious difference in RUNX2 expression, which indicates that ARID1A and ARID1B functionally compensate to regulate Runx2 expression and preserve the stem cell niche composition (Figure S8c). We also reanalyzed previously published scATAC-seq data from control and Arid1b KO samples19, showing no significant differences in chromatin accessibility peaks at the Runx2 locus, especially at the ARID1A/ARID1B/DLX2 binding site and the promoter region between the control mesenchymal cell (Control-MC) and Arid1b KO-MC samples (Figure S8d,e). This result suggests that loss of ARID1B alone does not affect Runx2 expression.

Given that both Runx2 and Arid1b are located on chromosome 17, it was not feasible to generate DKO mice with a Runx2 heterozygous deletion for functional validation. To circumvent this issue and assess the functional significance of RUNX2 as a downstream effector of the cBAF complex, we performed local knockdown experiments. Specifically, lentiviral particles encoding Runx2-targeting shRNA or control shRNA were injected into the proximal region of mouse incisors at 4-5 weeks of age, followed by tamoxifen induction, and samples were collected at 5 dpt (Figure 5f). Immunostaining confirmed efficient Runx2 knockdown in Runx2 shRNA-treated DKO;Gli1-LacZ incisors compared to those treated with control shRNA (Figure 5g). Additional analyses using β-GAL and Ki67 staining demonstrated significant restoration of disrupted MSC and TAC populations upon Runx2 knockdown (Figure 5g-i). These results indicate that RUNX2 is a critical downstream effector of the cBAF complex in maintaining niche cell function and regulating MSC and TAC populations.

Collectively, our findings demonstrate that the cBAF complex recruits DLX2 to regulate niche cell marker Runx2 expression. Within this regulatory complex, ARID1A and ARID1B exhibit functional redundancy, ensuring the appropriate expression levels of Runx2 to preserve stem cell niche integrity and sustain niche cell function.

cBAF-FOXO1 directly controls lineage-specific transcription factors for cell fate decisions

Since we observed severe phenotypes and cellular changes caused by disruption of the cBAF complex, we hypothesize that these changes arise not only from dysregulation of the stem cell niche but also from the disruption of other cellular processes. Therefore, we aimed to determine whether the loss of the cBAF complex directly causes the observed lineage commitment defects. Since our scRNA-seq analysis revealed significant changes in genes involved in proliferation, differentiation, and apoptosis along the cell differentiation trajectory (Figure 2f, Figure S3f), we sought to clarify how the cBAF complex controls TAC fate. To identify potential upstream regulators responsible for these gene expression changes, we performed TRRUST analysis, which pinpointed the top five TFs involved in this regulatory network (Figure 6a). We analyzed the expression patterns and differences in Stat3, Trp53, Yy1, Rela, and Sp1 between the control and DKO scRNA-seq datasets, showing that all were upregulated in the DKO samples, especially within the TACs and DP cells (Figure 6b). We then validated the expression of the top two TFs by in situ hybridization and confirmed increased expression of Stat3 and Trp53 in the DKO (Figure 6c). Considering the critical roles of these TFs in regulating cell differentiation and cell fate decisions during development and cancer progression53-59, we hypothesized that their elevated expression and activity may contribute to the proliferation and differentiation defects observed in TACs.

To test whether the cBAF complex regulates key TFs in vivo, we assessed chromatin accessibility at their gene loci using scATAC-seq data. We aggregated control and DKO cell populations into control mesenchymal cells (Control-MC) and DKO mesenchymal cells (DKO-MC), enabling a direct comparison of global chromatin accessibility between these two groups (Figure S9a). Our analysis revealed a significant increase in chromatin accessibility in the promoter regions of these five crucial TFs in the DKO-MC sample, reflecting enhanced transcriptional activation following cBAF loss (Figure 6d, Figure S9b). To further examine the direct regulatory interactions between cBAF complexes and these TF loci, as well as the co-factors FOXO1 and DLX2, we analyzed CUT&RUN-seq data between control and DKO samples. The data revealed predominant ARID1A and FOXO1 occupancy at the promoters of these TFs than ARID1B and DLX2 in the control sample, whereas FOXO1 and DLX2 binding activity showed varying levels of reduction in the DKO sample (Figure 6e, Figure S9c). To further validate cBAF-mediated transcriptional regulation of these TFs, we selected Stat3 and Trp53 as representative targets and cloned their promoter-binding regions upstream of Gluc for a luciferase reporter assay (Figure 6f). The DKO cells exhibited modest but significantly higher promoter-driven normalized Gluc activity than controls (Figure 6g), indicating that cBAF binds to promoter regions to repress TF expression. These transcriptional binding activities highlight the essential and predominant role of ARID1A-containing cBAF in interacting with FOXO1 to control the expression of key TFs.

To validate the functional roles of these TFs, we comprehensively investigated the cBAF complex in adult mesenchymal tissue by analyzing bulk RNA-seq data and identified over 3,000 genes that were significantly differentially expressed in the DKO compared to the control (Figure S10a). Using GO analysis, we further highlighted the top 10 altered signaling pathways (Figure S10b). Notably, several key pathways, including PDGF signaling, angiogenesis, the p53 pathway, and apoptosis signaling, are associated with TRP53. Given this connection, we chose TRP53 as a representative TF for rigorous testing of its functional role downstream of cBAF in regulating TAC fate by balancing proliferation, differentiation, or apoptosis. We generated a genetic rescue model by deleting one Trp53 allele on the DKO background (DKO;Trp53+/-). Following tamoxifen induction at 4-5 weeks of age, samples were collected at 5 dpt. At the cellular level, the disturbed TAC proliferation and impaired differentiation observed in the DKO incisors were partially rescued in the DKO;Trp53+/- model (Figure 6h). Additionally, the increased apoptosis in the DKO sample was partially ameliorated by the reduction of Trp53 (Figure S10c). These data collectively indicate that TRP53 acts as a key downstream TF of the cBAF complex, with critical roles in mediating TAC behavior and cell fate commitment.

Taken together, our findings demonstrate that the cBAF complex orchestrates the mesenchymal stem cell microenvironment and regulates lineage commitment through multiple, functionally distinct mechanisms. Specifically, cBAF maintains stem cell niche populations by modulating DLX2-mediated repression of Runx2, thereby preserving the stem cell microenvironment. Concurrently, cBAF regulates global chromatin accessibility and transcription factor occupancy, thereby guiding key biological processes, such as proliferation, differentiation, and apoptosis, in TACs. These results establish the cBAF complex as a central regulatory hub with numerous important cell-type-specific functions, achieved through distinct co-factors, including safeguarding stem cell niche composition, regulating progenitor fate commitment, and maintaining adult mesenchymal tissue homeostasis.

Methods
Ethics
This research was conducted in accordance with federal guidelines under a protocol approved by the Institutional Animal Care and Use Committee at the University of Southern California. Mice were housed in a pathogen-free environment, with comprehensive precautions taken to ensure their welfare. At specified time points, the mice were humanely euthanized using an overdose of carbon dioxide, followed by confirmation of death through cervical dislocation.

Animals
The following mouse lines were used and cross-bred in this study: Arid1afl/fl (JAX: 027717)93; Arid1bfl/fl (JAX: 032061)94; Trp53fl/fl (JAX: 008462)95; Gli1-CreER (JAX:007913)96; Sox2-CreER (JAX: 017593)97; Gli1-LacZ (JAX:008211)98; tdTomato (JAX: 007905)99. Male and female mice showed similar phenotypes and were both used in this study.

Tamoxifen administration
Tamoxifen (Sigma-Aldrich, T5648) was dissolved in corn oil (Sigma-Aldrich, C8267) at a concentration of 20 mg/mL. It was administered to mice at one month of age via intraperitoneal injection on three consecutive days, each at a dose of 1.5 mg per 10 g body weight, to induce Cre activity. Mouse mandibles were collected at multiple time points for molecular, cellular, and phenotypic analyses.

Notch movement and clipping assays
Control and DKO mice one week after tamoxifen induction were used for these assays. After administering anesthesia, a notch was made in the enamel of the left mandibular incisor above the gumline using a carbide bur (Brasseler USA, 018554U0). Ten days later, the distance of notch movement was measured as an indicator of incisor growth.

For the clipping assay, the left mandibular incisors of anesthetized control and DKO mice were clipped along the gingival papilla one week after tamoxifen induction. Two days after clipping, the injury repair ability was evaluated by measuring the amount of incisor growth.

Histological analysis
The mouse mandibles were sampled and dissected, then fixed in 10 % formalin solution at 4 °C overnight and decalcified in 10 % EDTA/PBS solution for 2-4 weeks. Following decalcification, the mandibles underwent a series of ethanol and xylene treatments for dehydration and were subsequently embedded in paraffin. The samples were sectioned at 5 µm thickness using a microtome (Leica, RM2235) and subjected to Hematoxylin and Eosin (HE) staining following the standard protocol for histological analysis.

Immunofluorescence and in situ hybridization
The mouse mandibles were collected, fixed, and decalcified as described for histological analysis. Prior to cryosectioning, the mandibles were dehydrated in sucrose/PBS solutions of increasing concentrations (15%, 30%, and sucrose/OCT) and then embedded in OCT compound. The samples were sliced into 8 µm sections with a cryostat (Leica, CM1850) and collected onto slides for immunofluorescence staining and in situ hybridization.

For immunofluorescence staining, the slides were incubated in a blocking solution (PerkinElmer, FP1012) for 1 hour at room temperature, then incubated with primary antibodies at 4 ˚C overnight. The slides were rinsed with PBST 3 times before incubation at room temperature for 2 hours with Alexa-conjugated secondary antibodies (1:200, Invitrogen). After that, the slides were counterstained with DAPI (Thermo Fisher Scientific, 62248) and then mounted. The primary antibodies used in this study were: ARID1A (1:100, Abcam, ab182561), ARID1B (1:100, Abcam, ab244351), Ki67 (1:100, Abcam, ab15580), β-galactosidase (β-GAL) (1:100, Abcam, ab9361), RUNX2 (1:100, Cell Signaling Technology, 12556), and Phospho-Histone H2A.X (Ser139) (Cell Signaling Technology, 2577).

RNAscope in situ hybridization was performed using the RNAscope Multiplex fluorescent v2 kit (Advanced Cell Diagnosis, 323100) according to the manufacturer’s instructions. The probes used in this study were synthesized by Advanced Cell Diagnostics: Probe-Mm-Dspp (448301), Probe-Mm-Trp53 (402331), Probe-Mm-Dlx2 (555951), Probe-Mm-Foxo1 (485761), Probe-Mm-Axin2 (400331), and Probe-Mm-Stat3 (425641).

Imaging
The histological HE-stained images were captured with a fluorescence microscope in brightfield mode (Keyence, BZ-X810). The immunofluorescent and RNAScope in situ hybridization fluorescent signals on histological sections were captured with a fluorescence microscope (Keyence, BZ-X810).

EdU incorporation, staining, and TUNEL assays
Both control and DKO mice were administered a single intraperitoneal injection of 5-ethynyl-2′-deoxyuridine (EdU) at a dosage of 25 µg/g body weight 2 hours and 48 hours before harvest. The mandibles were then dissected, fixed, decalcified, and sectioned as described above. EdU signal was detected using the Click-iT plus EdU cell proliferation kit (Thermo Fisher Scientific, C10637). Cell apoptosis was detected using the Click-iT plus TUNEL Assay kit (Thermo Fisher Scientific, C10617). Both procedures were performed according to the manufacturer's instructions. All the images were captured with a fluorescence microscope (Keyence, BZ-X810).

Bulk RNA-sequencing analysis
The proximal mesenchymal tissue of the DKO mouse incisor was carefully dissected four days after tamoxifen induction. RNA was extracted following the protocol provided by the RNeasy Micro Kit (Qiagen, 74004). Afterward, the cDNA library was constructed and sequenced at the Molecular Genomics Core at the University of Southern California (USC). The raw sequencing data were preprocessed, aligned to the mouse mm10 genome, and analyzed together with published mouse incisor control RNA-seq data using Partek Flow to identify differential gene expression19. Significance was assessed at a false discovery rate (FDR) of ≤ 0.1 and absolute fold change > 1.5.

Mesenchymal cell isolation
Mesenchymal tissue from the proximal end of the mouse incisor was collected and mechanically fragmented into tiny pieces to facilitate cell digestion. A solution of 2 mg/mL collagenase type I (Worthington, LS004196) and 4 mg/mL dispase II (Sigma-Aldrich, 54905400) was added to the chopped tissue in α-MEM medium (Gibco, 12571063). After fragmentation, the tissue was incubated at 37°C in a thermomixer (Eppendorf) at 600 rpm for 15 minutes to facilitate the digestion. The process was terminated by adding α-MEM medium with 10% FBS (Gibco, 16000044). The mixture was centrifuged at 600 × g for 5 minutes, and the supernatant was discarded. The pellet was resuspended in α-MEM medium with 10% FBS for further processing.

Single-cell RNA sequencing
Single cells were isolated from the proximal mesenchymal tissue of DKO mouse incisors at four days post-tamoxifen induction, following the protocol outlined in the 'Mesenchymal cell isolation' section. These cells were processed using the 10X Chromium system and tagged with barcodes from the Chromium Next GEM Single Cell 3ʹ v3.1 kit, preparing them for single-cell RNA sequencing. The library sample was constructed following the guidelines in the Chromium Next GEM Single Cell 3' Reagent Kits User Guide (v3.1 Chemistry) and then sequenced on an Illumina Novaseq System at the Technology Center for Genomics and Bioinformatics, University of California, Los Angeles (UCLA). Quality control, mapping, and assembly of the library count tables were executed using the CellRanger pipeline. A total of 8,182 cells from the DKO sample were sequenced. The raw read counts generated by the CellRanger count command were analyzed alongside published control scRNA-seq data using the Seurat v5 R package19,100.

Single-cell ATAC sequencing
Single cells were extracted from the proximal mesenchymal tissue of DKO mouse incisors at four days post-tamoxifen induction, following the protocol outlined in the 'Mesenchymal cell isolation' section. Following extraction, cells were lysed to release nuclei, and scATAC-seq libraries were then prepared using the Chromium Next GEM Single Cell ATAC Library & Gel Bead Kit (10x Genomics, PN-1000176). The suspension of nuclei was loaded onto the 10X Chromium system for barcoding. Library construction was performed following the v1.1 Chemistry protocol of the Chromium Single Cell ATAC Library & Gel Bead Kit. Sequencing of the scATAC-seq libraries was conducted on an Illumina NovaSeq System at the Technology Center for Genomics and Bioinformatics, UCLA. The raw data was processed and aligned to the reference genome mm10 post-sequencing using the ATAC CellRanger pipeline. Comparative analysis and peak calling were performed using Signac alongside published control scATAC-seq data19,52,101.

CUT&RUN sequencing and data analysis
Pulled mesenchymal tissue from the proximal end of the incisor was collected from control and DKO mice, respectively, and the tissue was digested as described in the ‘Mesenchymal cell isolation’ section. The CUT&RUN procedure was performed following the protocol for the Epicypher CUTANA™ ChIC/CUT&RUN kit (Epicypher, 14-1048), and library construction followed by the protocol of the Epicypher CUTANA™ library prep kit (Epicypher, 14-1001). The antibodies used for ARID1A were 1 μl of Abcam ab182561 and 2 μl of Cell Signaling Technology 12354. The antibodies used for ARID1B were 1 μl of Abcam ab57461 and 2 μl of Cell Signaling Technology 92964. The antibody used for FOXO1 was 2 μl of Proteintech 18592-1-AP. The antibody used for DLX2 was 2 μl of Abcam ab272902. The antibody used for IgG (control) was 1 μl of Epicypher 13-0042.

The CUT&RUN-seq libraries were sequenced on an Illumina NovaSeq System at the Technology Center for Genomics and Bioinformatics, UCLA. CUT&RUN-seq reads were trimmed for adapters and low-quality sequences using Galore and subsequently mapped to the mouse mm10 reference genome using Bowtie2 with default parameters102. PCR duplicates were marked using Picard. Coverage tracks were generated using the bamCoverage tool from deepTools with the RPKM normalization method103, and peak calling was conducted using MACS2104. Peak annotation was performed with ChIPseeker (RStudio). Visualization of CUT&RUN peaks was carried out using Integrative Genomics Viewer (IGV, Broad Institute). Motif analysis of enriched peaks was performed with HOMER (Hypergeometric Optimization of Motif EnRichment). Each control CUT&RUN sequencing sample has two replicates, and each DKO CUT&RUN sequencing sample has one replicate.

Co-immunoprecipitation
The proximal ends of control mouse incisors were collected and lysed in IP lysis buffer. The soluble fraction was centrifuged at 14,000 × g at 4°C for 10 min. The protein extracts were incubated overnight at 4°C with antibodies against ARID1A (Abcam, ab182560), ARID1B (Cell Signaling Technology, 92964), or control IgG (Cell Signaling Technology, 3900) under constant rotation. On the following day, pre-washed Protein G Sepharose beads were added to the lysate-antibody mixtures and incubated for an additional 2 hours at 4°C with rotation. The immune complexes were then washed three times. Input samples (total protein extracts before immunoprecipitation) and immunoprecipitated proteins (IgG control and specific antibody-target complexes) were separated using 4–15% SDS–PAGE gels, transferred onto PVDF membranes, and blocked with 5% non-fat dry milk. The membranes were incubated with primary antibodies against FOXO1 (Cell Signaling Technology, 2880), DLX2 (Abcam, ab272902), DLX1 (Cell Signaling Technology, 96585), DLX5 (Abcam, ab109737), and FOXO3A (Cell Signaling Technology, 2497), followed by detection using the Azure 300 imaging system (Azure Biosystems).

siRNA treatment of primary mesenchymal cells
Tissue from the proximal ends of control mouse incisors was dissected, minced, and plated on 6-well plates. Once cells had migrated out and reached confluence, they were harvested and replated at a density of 6 × 10⁴ cells per well into 24-well plates. The following day, cells were transfected with either control siRNA (Invitrogen, 4390844) or Dlx2 siRNA (Invitrogen, 4390771, ID s65010) using Lipofectamine RNAiMAX (Invitrogen, 13778150), according to the manufacturer’s protocol. RNA was extracted 3 days post-transfection using the RNeasy Plus Micro Kit (Qiagen, 74034), and quantitative PCR (qPCR) was performed to assess relative gene expression. The following primer sequences were obtained from PrimerBank (https://pga.mgh.harvard.edu/primerbank/) and used for qPCR reactions: Dlx2 (Forward: 5’-GTGGCTGATATGCACTCGACC-3’; Reverse: 5’-GCTGGTTGGTGTAGTAGCTGC-3’); Runx2 (Forward: 5’-GACTGTGGTTACCGTCATGGC-3’; Reverse: 5’-ACTTGGTTTTTCATAACAGCGGA-3’)

shRNA-lentivirus injection
Mice were anesthetized with isoflurane (2.5% induction, 1.5% maintenance). After clipping hair from the cheek, the skin was aseptically prepared using alternating Betadine scrub and alcohol. A sagittal incision was made through the skin and masseter muscle to expose the incisor region. Two full-thickness bone defects, each 0.3–0.4 mm in diameter and spaced 3.0–4.0 mm apart, were created using a microsurgical drill equipped with a trephine bit. One defect was positioned at the proximal end of the mouse incisor, while the other was located in the dental pulp region directly overlying the incisor mesenchyme. For lentiviral injection, a 10 μl Hamilton syringe (Hamilton Company, Reno, NV) connected to a glass capillary micropipette was inserted into the bone defect at the proximal end of the incisor. Subsequently, 3 μl of lentivirus carrying Runx2 shRNA (Santa Cruz, sc-37146-v) or control shRNA (Santa Cruz, sc-108080) was injected into the proximal end of the incisor mesenchyme at a flow rate of 20 nl/s. Mice were euthanized at 9 days post-injection for downstream analyses.

Luciferase reporter assay
Tissue from the proximal ends of control and DKO mouse incisors was dissected, minced, and plated on 6-well plates. Once cells had migrated out and reached confluence, they were harvested and replated at a density of 1 × 105 cells per well in 24-well plates. The Runx2 promoter, along with the cBAF binding sequence in the Runx2 intron (GeneCopoeia, CS-MPRM76664-LvPG04-01-50), the cBAF binding sequence in the Stat3 promoter (GeneCopoeia, CS-GS849T-LvPG04-01), and the cBAF binding sequence in the Trp53 promoter (GeneCopoeia, CS-GS850T-LvPG04-01) were cloned into the pEZX-LvPG04 vector, which contains Gaussia luciferase (Gluc) and secreted alkaline phosphatase (SeAP) in a dual-reporter system. These vectors were transfected using EndoFectin™ Lenti transfection reagent (GeneCopoeia, EF001) 48 hours after plating cells, following the manufacturer's protocol. After 24 hours of incubation, the transfected cells were changed to fresh medium. After another 48 hours, the cell culture medium was collected for analysis using the Secrete-Pair dual-luminescence assay kit (GeneCopoeia, FL033) according to the manufacturer's instructions. The SeAP signal was used as an internal standard control for the relative Gluc activity (Gluc/SeAP ratio) analysis of all samples. At least three replicates were performed, and each replicate sample was read three times to calculate the average.

Statistical analysis and reproducibility
Statistical analysis of the data was conducted using GraphPad Prism10 software. To determine the significance of the differences between paired groups, at least three samples from each comparison group were included, and unpaired two-tailed Student's t-tests were applied. The results are expressed as mean ± standard error of the mean (SEM). A p-value of less than 0.05 was considered to indicate statistical significance. Each experiment included at least three biological replicates with similar results.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript by Zhang et al reports how the canonical BAF (cBAF) chromatin remodeling complex regulates dental mesenchymal stem cell fate specification and maintains stem cell niches. They show that cBAF specifically recruits the transcriptional cofactors to activate different transcriptional programs. While cBAF recruits DLX2 to maintain stem cell-derived niche cells by remodeling intronic chromatin accessibility of niche-defining marker Runx2, cBAF-FOXO1 balances progenitor proliferation and differentiation through directly regulating promoter accessibility of lineage-specific transcription factors. These findings are interesting and novel. Another strength of this manuscript is that the authors used both genetic and epigenetic approaches combined with excellent mouse histology. Before this manuscript can be accepted, I suggest that there are two questions should be addressed.\n1) In addition to mesenchymal stem cells in mouse incisors, can these findings be applied to other mesenchymal tissues in mouse or human?\n2) It is very nice that the authors show that cBAF interacts with DLX2 and FOXO1. It will be more convincing if the authors perform re-ChIP assays to confirm these important findings, if possible.", "reviewer_2": "The study discovered that the cBAF complex functions as a central hub mechanism in regulating stem cell niches and fate determination. The focus of the research is on the specific roles played by cBAF in different cell populations. The results in the manuscript have some problems, so below points need to be addressed in order to substantiate the authors’ findings.\n1. Regarding DLX2 and FOXO1 as cofactors of cBAF, is there any relevant literature or research background supporting this premise? It is recommended to include such information in the Background section, while reducing the extent of discussion in the Results section.\n2. When describing the colocalization of fluorescence signals, it is advisable to add indicative arrows in the figures, along with quantitative colocalization analysis and schematic diagrams.\n3. The conclusion that the cBAF complex regulates cell fate requires deeper substantiation. Further exploration or discussion of the role of the cellular microenvironment may provide valuable insights.\n4. AXIN2 is used in this study as a marker of the Wnt pathway. However, it is known that AXIN2 itself plays an intrinsic role in craniofacial development. Has its specific role in tooth development been considered in the present work?\n5. Please clarify how the timing of tamoxifen injection in mice was determined.\n6. It is recommended to incorporate some of the results currently presented in the supplementary figures into the main figures of the published article, in order to maintain continuity in reading.", "reviewer_3": "Zhang’s work offers a comprehensive exploration of the role of the canonical BAF chromatin remodeling complex in regulating stem cell fate and maintaining tissue homeostasis, using the adult mouse incisor as a model. A key strength is the integrated use of single-cell multi-omics alongside in vivo perturbations to reveal how ARID1A/ARID1B and cell-type specific co-factors, particularly DLX2 and FOXO1, shape chromatin accessibility programs that coordinate niche maintenance and progenitor fate. The systems-level framework is compelling. Relative to the solid multi-omics analyses, several biological mechanistic links rely on correlative evidence. Strengthening these causal tests would considerably enhance the manuscript.\n1, The manuscript profiles dental mesenchymal cells at 4dpt by scRNA-seq and scATAC-seq, but does not articulate why this specific time point was chosen. Figures just emphasize 3 dpt and 5 dpt immunofluorescence, leaving a gap at day 4 despite single-cell sampling at that time. Authors may add day-4 immunofluorescence (e.g., β-GAL/Gli1, Ki67, or short-pulse EdU) to provide morphological context for the single-cell datasets, and justify the selection of 4 dpt (e.g., earliest inflection prior to overt MSC-TAC-niche cell domains disorganization). These additions would align modality, time point, and interpretation, strengthening causal inferences about early fate imbalance.\n2, The Runx2⁺Gli1⁺ niche cell is central to the study, yet its identity and dynamics are not shown explicitly in the single-cell analyses. In Fig. 2 and Suppl. Fig. 2, neither the clustering nor the pseudotime views delineate a niche subset or quantify its change after DKO. By contrast, the feature plots (Suppl. Fig. 6b: Runx2; Suppl. Fig. 2c: Gli1) imply that “niche” cells may align with a PDF subpopulation, whereas the schematics in Fig. 1a/Fig. 2b—and the text stating you “isolated Gli1⁺Runx2⁺ cells from the MSC population”—depict the niche as an MSC subset, creating a definition and mapping mismatch. To resolve this, authors may annotate the niche directly in Fig. 2 by co-expression gating (Gli1⁺ & Runx2⁺) on the scRNA-seq UMAP for control and DKO, and quantify niche proportions and nearest-neighbor relationships to MSC, PDF, and TAC clusters and also specify how Gli1⁺Runx2⁺ cells were defined computationally and used in downstream analysis in method.\n3, The motif and expression data suggest broader candidate co-factors than the two emphasized. In Fig4a and Suppl. Fig. 6a, DLX and FOX motifs rank among the top hits, with DLX1/DLX5 and FOXO3 notably prominent; Suppl. Fig. 6b shows similar family-level expression patterns across dental mesenchymal populations. Yet the manuscript focuses on DLX2 and FOXO1, supported by co-IP, footprinting, CUT&RUN, and co-occupancy with ARID1A/ARID1B (Fig. 4a–f), without excluding roles for DLX1/DLX5 or FOXO3. Please substantiate this prioritization (or to rule out alternatives).\n4, The current argument that cBAF co-regulates targets (e.g., Runx2, Trp53) via DLX2 and FOXO1 rests largely on control-side CUT&RUN (ARID1A/B, DLX2, FOXO1), DKO-side scATAC accessibility changes, and in vivo readouts. While strongly suggestive, authors could provide DKO-side CUT&RUN for DLX2 and FOXO1, focusing on the defined Runx2 intronic element and the promoters of Trp53/Stat3 and validate the function of the Runx2 intronic site and Trp53/Stat3 promoters.\nMinor points:\n1. Fig. 6 and Suppl. Fig. 8 show that promoters of the five nominated TFs (STAT3, TRP53, YY1, RELA, SP1) display increased accessibility in DKO, whether DLX2/FOXO1 are also enriched and occupied at the promoter-opened regions.\n2. The manuscript concludes that enhancers preferentially lose accessibility whereas promoters tend to open after DKO, but it is unclear whether these map to distinct or overlapping gene cohorts, what functions they serve in MSC, TAC, and Runx2⁺Gli1⁺ niche cells, and how they relate to ARID1A/ARID1B CUT&RUN binding. A compact summary would help.\n3. In Fig. 4c the Foxo1 in situ signal appears weak, which seems inconsistent with the manuscript’s claim that FOXO1 partners with ARID1A to regulate TAC-biased TF hubs and with the stated higher Foxo1 expression in TAC/DP relative to MSCs (Suppl. Fig. 6b). The authors could repeat immunofluorescence with FOXO1 antibody and co-stain for Ki67 or other TAC-domain markers to localize FOXO1 within TACs.\n4. Fig. 5g does not convincingly show the reduction of Gli1⁺ MSCs and Ki67⁺ TACs in DKO or their rescue after Runx2 knockdown. Please replace the representative images and add quantitative support."}, "author_response": "We are grateful to the reviewers for their insightful comments and suggestions, which have greatly improved this manuscript. We have evaluated each point raised by the reviewers and provided responses to address all questions and concerns.\n\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nThis manuscript by Zhang et al reports how the canonical BAF (cBAF) chromatin remodeling complex regulates dental mesenchymal stem cell fate specification and maintains stem cell niches. They show that cBAF specifically recruits the transcriptional cofactors to activate different transcriptional programs. While cBAF recruits DLX2 to maintain stem cell-derived niche cells by remodeling intronic chromatin accessibility of niche-defining marker Runx2, cBAF-FOXO1 balances progenitor proliferation and differentiation through directly regulating promoter accessibility of lineage-specific transcription factors. These findings are interesting and novel. Another strength of this manuscript is that the authors used both genetic and epigenetic approaches combined with excellent mouse histology. Before this manuscript can be accepted, I suggest that there are two questions should be addressed.\nWe appreciate the reviewer's positive evaluation and valuable comments for helping us improve our manuscript. Below, we provide detailed, point-by-point responses to address each comment.\n1) In addition to mesenchymal stem cells in mouse incisors, can these findings be applied to other mesenchymal tissues in mouse or human?\nWe thank the reviewer for raising this important point regarding the broader applicability of our findings to other mesenchymal tissues. Since the mouse incisor is derived from cranial neural crest cells (CNCCs), we examined additional mesenchymal tissues, including the coronal suture (also CNCC-derived), the sagittal suture (with its anterior portion derived from CNCCs and posterior portion from mesoderm), and the femur, a representative mesoderm-derived mesenchymal tissue.\nWe induced Cre activity at 1 month of age and harvested tissues 1 month after induction using Arid1afl/fl;Arid1bfl/fl as the control, comparing these mice to Gli1-CreER;Arid1afl/fl;Arid1bfl/fl (DKO) with loss of the cBAF complex. MicroCT analysis of control and DKO samples revealed distinct phenotypes across tissues. As expected, the mouse incisor displayed severe defects, with complete loss of dentin and enamel formation (Response Figure 1a). In contrast, no apparent morphological changes were observed in the coronal or sagittal sutures (Response Figure 1a).Although the femur in 1-month-old mice is approaching skeletal maturity, the growth plate remains active and supports continued bone growth and remodeling. In DKO femurs, microCT analysis showed reduced length and decreased trabecular bone formation, indicating that cBAF loss impairs postnatal bone growth and tissue differentiation (Response Figure 1a–1b).\nNext, we focused on the femur metaphysis, including the growth plate and subchondral bone, and assessed the expression patterns of ARID1A, ARID1B, FOXO1, and DLX2 at 1 month of age. Immunostaining showed that both ARID1A and ARID1B were broadly expressed in the growth plate, subchondral bone, and differentiated trabecular bone, with ARID1A showing higher expression at the growth plate. In situ hybridization further revealed stronger Foxo1 expression compared to Dlx2 in the subchondral bone (Response Figure 1c). Given that GLI1 marks osteogenic progenitor cells involved in bone formation1, we analyzed GLI1+ cells using Gli1-CreER;tdTomato and DKO;tdTomato mice. Cre activity was induced at 1 month of age, and tissues were collected 5 days later. We observed fewer tdTomato-positive cells, particularly at the subchondral bone, in DKO mice compared to controls, suggesting a disruption of the osteogenic progenitor population following cBAF loss (Response Figure 1d). Meanwhile, we also examined the expression of key cBAF downstream targets, RUNX2, Stat3, and Trp53, and found that all three were significantly upregulated in the subchondral bone of DKO femurs, similar to the changes observed in the incisor model (Response Figure 1d).\nThese results indicate that cBAF plays a critical role in regulating osteogenic progenitor behavior in the femur. While cBAF may interact with distinct co-factors in the metaphysis compared to the incisor, the regulation of downstream transcriptional targets appears to be at least partially conserved.\nResponse Figure 1. cBAF plays essential roles in regulating femur growth and tissue homeostasis.\n(a) MicroCT images of incisor, coronal suture, sagittal suture, and femur in control and DKO mice induced at 1 month and examined 1 month post-tamoxifen induction (1mpt). Yellow arrows indicate the normal differentiated tissue, and asterisks mark areas of tissue loss. (b) Quantification of femur length in control and DKO samples. Data are presented as mean ± SEM, n = 3, using an unpaired two-tailed Student’s t-test. p = 0.0163. (c) Immunostaining of ARID1A and ARID1B, and in situ hybridization of Foxo1 and Dlx2 in the femur at 1 month of age. The lower panel shows a higher magnification of the boxes in the upper panel. Yellow arrows indicate positive signals. (d) Comparative analysis of tdTomato-positive GLI1, RUNX2, Trp53, and Stat3 between control and DKO samples at 5 days post-tamoxifen induction (5dpt). Yellow arrows mark positive signals. n = 3. Scale bars: 100 μm.\n2) It is very nice that the authors show that cBAF interacts with DLX2 and FOXO1. It will be more convincing if the authors perform re-ChIP assays to confirm these important findings, if possible.\nWe thank the reviewer for this insightful suggestion. We agree that re-ChIP experiments would provide strong evidence to further validate the co-occupancy of cBAF with DLX2 and FOXO1. Unfortunately, because of the limited amount of chromatin obtainable from the proximal tissue of mouse incisors, performing re-ChIP assays from this very restricted tissue source turned out to be very challenging.\nAs an alternative strategy to support our conclusion, we performed CUT&RUN sequencing on DKO mouse incisor samples to investigate whether the loss of the cBAF complex affects DLX2 and FOXO1 binding. We observed a marked reduction of DLX2 binding at the intronic regulatory region of Runx2 upon loss of the cBAF complex (Revised Figure 5a), supporting the role of DLX2 as a cBAF-dependent co-factor in maintaining niche cells. Additionally, we examined FOXO1 and DLX2 binding profiles at key downstream target loci (Stat3, Trp53, Yy1, Rela, and Sp1) and observed varying degrees of peak signal reduction in DKO samples (Revised Figure 6e, Supplementary Figure 9c). These results collectively further supported the conclusion that DLX2 and FOXO1 function as cBAF co-factors in regulating lineage-specific transcriptional programs.\n\nReviewer #2 (Remarks to the Author)\nThe study discovered that the cBAF complex functions as a central hub mechanism in regulating stem cell niches and fate determination. The focus of the research is on the specific roles played by cBAF in different cell populations. The results in the manuscript have some problems, so below points need to be addressed in order to substantiate the authors’ findings.\nWe thank the reviewer for the feedback on our study and the valuable suggestions to improve our manuscript. Below, please find our point-by-point responses to all of the questions raised.\n1. Regarding DLX2 and FOXO1 as cofactors of cBAF, is there any relevant literature or research background supporting this premise? It is recommended to include such information in the Background section, while reducing the extent of discussion in the Results section.\nWe thank the reviewer for raising this point. We have now incorporated more background on the molecular functions of DLX2 and FOXO1 in cell regulation in the Introduction (lines 61-76) and reduced the discussion in the Results section accordingly.\n2. When describing the colocalization of fluorescence signals, it is advisable to add indicative arrows in the figures, along with quantitative colocalization analysis and schematic diagrams.\nWe thank the reviewer for the suggestion. In the revised Figure 2g, we have added arrows to highlight regions of colocalization in the DKO;Gli1-LacZ samples and included schematic diagrams for better visualization. Additionally, we quantified RUNX2+/GLI1+ double-positive cells in the revised Supplementary Figure 2f. For Figure 2i, where no colocalized fluorescence signals were observed in the DKO samples, we marked these areas with asterisks (*) to clearly indicate the absence of overlap.\n3. The conclusion that the cBAF complex regulates cell fate requires deeper substantiation. Further exploration or discussion of the role of the cellular microenvironment may provide valuable insights.\nWe thank the reviewer for the suggestion. To provide context and further substantiate this point, we revised the discussion related to cBAF’s regulation of the stem cell niche and cellular microenvironment to lines 500-518.\n4. AXIN2 is used in this study as a marker of the Wnt pathway. However, it is known that AXIN2 itself plays an intrinsic role in craniofacial development. Has its specific role in tooth development been considered in the present work?\nWe thank the reviewer for this comment. In our DKO mouse model, we observed that several direct downstream targets of cBAF, including Runx2, Stat3, and Trp53, showed altered expression as early as 4 days post-tamoxifen induction (dpt; see revised Figure 5c, Figure 6c). In contrast, Axin2 expression did not change significantly at this time point in our bulk RNA-seq data. Changes in Axin2 expression were detected at 5 dpt, suggesting that altered Wnt signaling is not one of the primary changes following loss of the cBAF complex.\nWe agree with the reviewer that Wnt signaling is crucial for craniofacial development, and that it would be interesting to investigate whether/how Wnt signaling plays a role in adult stem cell behavior in the mouse incisor in our future studies.\n5. Please clarify how the timing of tamoxifen injection in mice was determined.\nWe thank the reviewer for the opportunity to clarify this rationale. To study adult stem cell homeostasis, we administer tamoxifen injections at postnatal 4-5 weeks, when the incisors are already at a mature stage. This induction regime was established in our previous publications2-4. We have now included this rationale for the induction timing on lines 123-125.\n6. It is recommended to incorporate some of the results currently presented in the supplementary figures into the main figures of the published article, in order to maintain continuity in reading.\nWe thank the reviewer for the suggestion. To improve the flow for readers, we reorganized the figures and added former Supplementary Figure 7a to revised Figure 5d, former Supplementary Figure 8c to revised Figure 6c, and former Supplementary Figure 8b to revised Figure 6d.\n\nReviewer #3 (Remarks to the Author)\nZhang’s work offers a comprehensive exploration of the role of the canonical BAF chromatin remodeling complex in regulating stem cell fate and maintaining tissue homeostasis, using the adult mouse incisor as a model. A key strength is the integrated use of single-cell multi-omics alongside in vivo perturbations to reveal how ARID1A/ARID1B and cell-type specific co-factors, particularly DLX2 and FOXO1, shape chromatin accessibility programs that coordinate niche maintenance and progenitor fate. The systems-level framework is compelling. Relative to the solid multi-omics analyses, several biological mechanistic links rely on correlative evidence. Strengthening these causal tests would considerably enhance the manuscript.\nWe thank the reviewer for the supportive comments and valuable suggestions to improve our manuscript. Below, please find our point-by-point responses.\n1, The manuscript profiles dental mesenchymal cells at 4dpt by scRNA-seq and scATAC-seq, but does not articulate why this specific time point was chosen. Figures just emphasize 3 dpt and 5 dpt immunofluorescence, leaving a gap at day 4 despite single-cell sampling at that time. Authors may add day-4 immunofluorescence (e.g., β-GAL/Gli1, Ki67, or short-pulse EdU) to provide morphological context for the single-cell datasets, and justify the selection of 4 dpt (e.g., earliest inflection prior to overt MSC-TAC-niche cell domains disorganization). These additions would align modality, time point, and interpretation, strengthening causal inferences about early fate imbalance.\nWe thank the reviewer for the suggestion. We compared GLI1+ MSCs, Ki67+ TACs, and RUNX2+/GLI1+ niche cells at 3 dpt and 4 dpt using immunostaining and observed a significant increase in Ki67+ cells in the DKO samples compared to the control at 3 dpt. By 4 dpt, there was a notable reduction in GLI1+ MSCs and an expansion of RUNX2+/GLI1+ cells in the DKO samples. Based on these findings, we selected 4 dpt as the time point for scRNA-seq analysis, when the spatial organization of cell domains at the proximal end of the mouse incisor was disrupted.\nThese data have been added to the revised Supplementary Figure 2, with corresponding descriptions on lines 163-164.\n2, The Runx2⁺Gli1⁺ niche cell is central to the study, yet its identity and dynamics are not shown explicitly in the single-cell analyses. In Fig. 2 and Suppl. Fig. 2, neither the clustering nor the pseudotime views delineate a niche subset or quantify its change after DKO. By contrast, the feature plots (Suppl. Fig. 6b: Runx2; Suppl. Fig. 2c: Gli1) imply that “niche” cells may align with a PDF subpopulation, whereas the schematics in Fig. 1a/Fig. 2b—and the text stating you “isolated Gli1⁺Runx2⁺ cells from the MSC population”—depict the niche as an MSC subset, creating a definition and mapping mismatch. To resolve this, authors may annotate the niche directly in Fig. 2 by co-expression gating (Gli1⁺ & Runx2⁺) on the scRNA-seq UMAP for control and DKO, and quantify niche proportions and nearest-neighbor relationships to MSC, PDF, and TAC clusters and also specify how Gli1⁺Runx2⁺ cells were defined computationally and used in downstream analysis in method.\nWe thank the reviewer for the suggestion to explicitly annotate the RUNX2+GLI1+ niche cell population within the scRNA-seq data. Published studies indicate that these niche cells co-expressing RUNX2 represent a small subset of GLI1+ mesenchymal cells. This is a very small population that can be visualized by immunostaining but lacks additional unique transcriptional markers that would allow precise isolation as a discrete cluster in scRNA-seq. In light of this limitation, we validated niche cell dynamics using in vivo immunostaining, which provides more reliable detection of co-expressing cells. As shown in the revised Figure 2g and Supplementary Figure 2f, we quantified RUNX2+GLI1+ cells in control and DKO incisors and observed an increase in co-stained cells following loss of the cBAF complex. We added a note regarding this technical limitation on lines 171-173.\nThe reviewer also raises the important question of whether the RUNX2+GLI1+ cells in the DKO incisors retain niche-like molecular features. Prior work has shown that RUNX2+ niche cells secrete IGFBP3, a key mediator of MSC-derived IGF2 signaling that promotes TAC proliferation5. We therefore evaluated the expression of Igfbp3, along with Runx2, in our scRNA-seq datasets. Both genes were highly enriched in the proximal dental follicle and GLI1+ MSC clusters in the control, and showed further increased expression in the DKO sample (Response Figure 2a). Bulk RNA-seq analysis also confirmed significant upregulation of Igfbp3 in DKO tissue (Response Figure 2b). These data suggest that loss of cBAF expands the RUNX2+ niche-like population, although further experiments will be required to fully validate their functional identity. We also added more discussion about the niche cells on lines 509-518.\nResponse Figure 2. Igfbp3 is significantly upregulated after loss of the cBAF complex. (a) FeaturePlot of Runx2 and Igfbp3 expression in control and DKO incisor mesenchyme scRNA-seq datasets. (b) Quantification of Igfbp3 expression based on the incisor mesenchyme bulk RNA-seq data.\n3, The motif and expression data suggest broader candidate co-factors than the two emphasized. In Fig4a and Suppl. Fig. 6a, DLX and FOX motifs rank among the top hits, with DLX1/DLX5 and FOXO3 notably prominent; Suppl. Fig. 6b shows similar family-level expression patterns across dental mesenchymal populations. Yet the manuscript focuses on DLX2 and FOXO1, supported by co-IP, footprinting, CUT&RUN, and co-occupancy with ARID1A/ARID1B (Fig. 4a–f), without excluding roles for DLX1/DLX5 or FOXO3. Please substantiate this prioritization (or to rule out alternatives).\nWe thank the reviewer for raising this concern. We did not further investigate DLX1, DLX5, or FOXO3A because these proteins were not detected in our Co-IP assay. We have now included the Co-IP results to show that neither ARID1A nor ARID1B interacts with these proteins. This data has been added to the revised Supplementary Figure 6c.\n4, The current argument that cBAF co-regulates targets (e.g., Runx2, Trp53) via DLX2 and FOXO1 rests largely on control-side CUT&RUN (ARID1A/B, DLX2, FOXO1), DKO-side scATAC accessibility changes, and in vivo readouts. While strongly suggestive, authors could provide DKO-side CUT&RUN for DLX2 and FOXO1, focusing on the defined Runx2 intronic element and the promoters of Trp53/Stat3 and validate the function of the Runx2 intronic site and Trp53/Stat3 promoters.\nWe thank the reviewer for the comments. We conducted the CUT&RUN sequencing of DLX2 and FOXO1 in DKO mouse incisors as suggested. In the DKO samples, we found a marked reduction in DLX2 binding activity at the intron region of Runx2, where ARID1A, ARID1B, and DLX2 bind in the control (Revised Figure 5a), supporting the role of DLX2 as a cBAF-dependent co-factor in maintaining niche cells. Additionally, we examined FOXO1 and DLX2 binding profiles at key downstream target loci (Stat3, Trp53, Yy1, Rela, and Sp1) and observed varying degrees of peak signal reduction in DKO samples (Revised Figure 6c). These results collectively reinforce the conclusion that DLX2 and FOXO1 function as co-factors of cBAF in regulating lineage-specific transcriptional programs.\nTo validate the functional regulation of the cBAF binding region in the Runx2 intron and the promoters of Trp53 and Stat3, we performed a luciferase reporter assay. We observed significantly increased Gluc activities in the DKO cell culture compared to the control (Figure S8a, Figure 6f,g), indicating that cBAF suppresses Runx2 transcriptional activity by binding to the intron region and suppresses Stat3 and Trp53 expression by binding to their promoters. We added these results to lines 396-399 (Revised Figure S8a) and lines 465-471 (Revised Figure 6f,g).\nMinor points:\n1. Fig. 6 and Suppl. Fig. 8 show that promoters of the five nominated TFs (STAT3, TRP53, YY1, RELA, SP1) display increased accessibility in DKO, whether DLX2/FOXO1 are also enriched and occupied at the promoter-opened regions.\nWe thank the reviewer for the comment. In the revised Figure 6e and Supplementary Figure 9c, FOXO1 is more enriched and has higher occupancy at the promoter-opened regions than DLX2, as shown by the DLX2 and FOXO1 CUT&RUN-seq results in the control sample. However, both show varying levels of reduced binding activity after loss of cBAF.\n2. The manuscript concludes that enhancers preferentially lose accessibility whereas promoters tend to open after DKO, but it is unclear whether these map to distinct or overlapping gene cohorts, what functions they serve in MSC, TAC, and Runx2⁺Gli1⁺ niche cells, and how they relate to ARID1A/ARID1B CUT&RUN binding. A compact summary would help.\nWe thank the reviewer for the comment. The total number of changed peaks across all cells we collected was 73,131, which is much higher than the number of genes we could sequence. Therefore, it may involve overlapping gene cohorts, with genes that lose accessibility at the enhancer and gain accessibility at the promoter after cBAF loss. For example, our niche cell marker Runx2 showed reduced accessibility at the ARID1A/ARID1B binding site in the intron and higher accessibility at the promoter.\nAs we described, enhancers tend to lose accessibility, whereas promoters often gain accessibility, after cBAF loss. When we performed GO analysis of opened and closed peaks for annotated genes in MSCs, we identified overlapping signaling pathways (Revised Supplementary Figure 6d,f). The same applies to the TACs and niche cells. Thus, it is difficult to specify the individual roles of enhancers or promoters in MSC, TAC, and RUNX2+GLI1+ niche cells.\n3. In Fig. 4c the Foxo1 in situ signal appears weak, which seems inconsistent with the manuscript’s claim that FOXO1 partners with ARID1A to regulate TAC-biased TF hubs and with the stated higher Foxo1 expression in TAC/DP relative to MSCs (Suppl. Fig. 6b). The authors could repeat immunofluorescence with FOXO1 antibody and co-stain for Ki67 or other TAC-domain markers to localize FOXO1 within TACs.\nWe thank the reviewer for the comment. Unfortunately, we were unable to find antibodies of sufficient quality for FOXO1 staining. To address the reviewer’s concern, we co-stained Foxo1 with EdU injection for 2 hours, which labels cells in the S-phase, to show the co-localization of Foxo1 within TACs (Response Figure 3a-3b). We also co-localized Foxo1 with ARID1A (Response Figure 3c-3d). Based on the in situ hybridization, we revised the description to the more accurate “Foxo1 is expressed in dental pulp cells and less in TACs” on line 350.\nResponse Figure 3. Co-localization staining of Foxo1 with proliferating cells and ARID1A+ cells. (a-b) Co-staining of Foxo1 with EdU, which was injected 2 hours before harvesting the mouse incisor samples. b shows the high magnification of the box region in a. (c-d) Co-staining of Foxo1 with ARID1A in the mouse incisors. d shows the high magnification of the box region in c. Yellow arrows indicate the co-stained cells. Scale bars: 200 μm.\n4. Fig. 5g does not convincingly show the reduction of Gli1⁺ MSCs and Ki67⁺ TACs in DKO or their rescue after Runx2 knockdown. Please replace the representative images and add quantitative support.\nWe thank the reviewer for the comment. We have now included more representative staining images of GLI1+ and Ki67+ cells in Fig. 5g and added the quantification results to Fig. 5h and 5i."}, {"round": 2, "reviewer_comments": {"reviewer_1": "My concerns have been fully addressed.", "reviewer_2": "no other questions.", "reviewer_3": "I thank the authors for their comprehensive and point-by-point responses to my comments, and for the corresponding revisions made to the manuscript. All of my concerns have been adequately addressed, and the manuscript has been significantly improved. I find it now suitable for publication and therefore recommend its acceptance."}, "author_response": ""}]

Decision letter