ARID3A coordinates the proliferation-differentiation switch of transit-amplifying cells in the intestine

Intestinal stem cells (ISCs) at the crypt base divide and give rise to progenitor cells that have the capacity to proliferate and differentiate into various mature epithelial cell types in the transit-amplifying (TA) zone. Here, we identified the transcription factor ARID3A as a novel regulator of intestinal epithelial cell proliferation and differentiation at the TA compartment. We show that ARID3A forms an expression gradient from villus tip to the early progenitors at the crypts mediated by TGF-β and WNT signalling. Intestinal epithelial-specific deletion of Arid3a reduces proliferation of TA cells. Bulk and single cell transcriptomic analysis shows increased enterocyte differentiation and reduced secretory cells in the Arid3a cKO intestine. Interestingly, upper-villus gene signatures of both enterocytes and secretory cells are enriched in the mutant intestine. We find that the enhanced enterocyte differentiation in the Arid3a cKO intestine is caused by increased binding of HNF1 and HNF4. Finally, we show that loss of Arid3a impairs irradiation-induced regenerative process by altering the dynamics of proliferation and apoptosis. Our findings imply that ARID3A may play a gatekeeping role in the TA compartment to maintain the “just-right” proliferation-to-differentiation ratio for tissue homeostasis and plasticity.


Introduction
The intestinal epithelium is one of the fastest renewing and regenerating tissues and its high turnover in cell composition is facilitated by Lgr5+ intestinal stem cells (ISCs) residing at the intestinal crypts. ISCs divide to generate a daughter cell that will either self-renew to generate another stem cell or will enter the transit-amplifying (TA) zone for subsequent lineage specification (1)(2)(3). Intestinal lineage decision takes place at cell positions +4/+5, where progenitor cells are located (4). These progenitors can be broadly divided into two main subtypes, absorptive and secretory, which are highly plastic and are able to re-acquire stemness for tissue regeneration upon injury (5)(6)(7). The stem cell-to-daughter cell transition in the intestinal epithelium is a highly dynamic and plastic process. Maintenance and regulation of the stem cell pool is controlled both by an epithelial cellular niche as well as by the mucosal stromal microenvironment (8).
A variety of signalling cascades has been well-described in regulating ISC maintenance, fate decision and terminal maturation. WNT and NOTCH signalling are the key drivers to maintain stem cell identity at the bottom of the crypt. At the +4/+5 progenitors, NOTCH dictates the initial absorptive versus secretory fate decision: "NOTCH-ON" promotes enterocyte differentiation, whilst "NOTCH-OFF" de-represses the master regulator ATOH1 to drive secretory lineage specification. We have recently identified that the transcription corepressors MTG8 and MTG16 are also key regulators expressed at the progenitors to facilitate this binary fate decision process by repressing ATOH1 transcription (9). After the initial lineage commitment process at +4/5 cells, these progenitors will continue to proliferate and differentiate at the TA cells in the upper crypts and will eventually exit the cell cycle for terminal differentiation in the villi. Emerging evidence reveals that enterocytes, goblet cells and tuft cells exhibit a broad zonation of their gene expression programme along the cryptvillus axis to facilitate different functions, highlighting the complexity of the cellular differentiation process (10)(11)(12). Whilst the initial lineage specification at +4/5 cells has been extensively studied, the molecular control of proliferation and differentiation states at the TA cells has been largely overlooked in the past. Characterising the TA cell regulation will be important for the understanding of intestinal epithelial cell type composition under homeostasis and injury-induced regeneration.
Here, we report the transcription factor ARID3A as a novel regulator of intestinal homeostasis that plays a central role in controlling proliferation and differentiation dynamics of the TA cells. ARID3A forms an expression gradient from villus tip to crypt progenitor cells driven by TGF-b and WNT signalling. Loss of Arid3a inhibits TA cell proliferation, perturbs the absorptive versus secretory cell differentiation and increases the expression of villus-tip gene signatures, leading to reduced regenerative capacity upon irradiation-induced injury. Our findings reveal the hitherto unrecognised role of ARID3A in coordinating TA cell proliferation and differentiation in the intestine.

Arid3a forms an expression gradient from villus tip to crypt progenitor cells
We have previously identified a set of genes that are enriched at the +4/+5 progenitor cells compared to the Lgr5+ ISCs (9). Screening for transcription factors that are enriched at the +4/+5 cells identified the A+T Rich Interaction Domain 3a (Arid3a) as a putative modulator of intestinal epithelial homeostasis ( Figure 1A). Quantitative reverse transcription (qRT-PCR) analysis of Lgr5-GFP sorted cells isolated from a Lgr5-EGFP-ires-CreERT2 intestinal crypts confirmed the enrichment at the GFP-low progenitors ( Figures 1B and S1A). To further probe the localisation of Arid3a at the crypt, we performed double RNAscope co-staining of Arid3a with Lgr5 (ISC marker) and Atoh1 (secretory progenitor and Paneth cell marker). While Arid3a showed both overlapping and exclusive staining with Lgr5 and Atoh1, the co-localisation was minimal ( Figure 1C). Quantification of the RNAscope data revealed that Arid3a not only was enriched at the +4/+5 cells as compared to the ISCs, but its expression was maintained throughout the upper crypt ( Figure 1D). Interestingly, expression analysis of ARID3A at both mRNA and protein levels showed a strong expression gradient from the villus tip to early progenitor cells at the crypt ( Figures 1E and 1F). To confirm the enrichment of Arid3a at the villus compartment, we performed crypt-villus fractionation of mouse proximal small intestinal tissue followed by qRT-PCR analysis of the two compartments. As expected, the stem cell-specific marker Olfm4 was enriched at the crypt fraction, while the enterocyte marker Alkaline phosphatase (Alpi) was enriched at the villus. In accordance with our RNAscope data, Arid3a showed a 20-fold upregulation in the villus compared to the crypt ( Figure S1B). Stromal expression of Arid3a was also detected ( Figures 1E and 1F) since Arid3a is also expressed in B cells (13,14).
Enterocytes are the most prominent type of intestinal epithelial cells exhibiting characteristic microvilli structures (15). Careful examination of ARID3A protein staining confirmed the expression of ARID3A in brush border-bearing enterocytes ( Figure 1F). We further tested if Arid3a is also expressed at secretory cells. By combing RNAscope staining of Arid3a and immunofluorescent staining to detect the protein levels of Mucin 2 (MUC2), Chromogranin A (CHGA) and Lysozyme (LYZ), we confirmed that Arid3a is also expressed in goblet, enteroendocrine and Paneth cells, respectively ( Figure S1C).

WNT and TGF-β regulate expression of Arid3a in epithelial cells
Next, we sought to characterise the driver of Arid3a expression gradient. Arid3a forms an expression gradient from the upper crypt in a pattern opposite to the WNT gradient, implying that WNT signalling may regulate its expression. To test the regulatory role of WNT, ex vivo wild-type (WT) mouse intestinal organoids were treated with two different WNT inhibitors, LF3 and LGK974, for 48h. Successful WNT inhibition was confirmed by downregulation of known WNT target genes, such as Axin2, Sox9 and Cyclin D1 (Figure 2A and S2A). In contrast, Arid3a expression was upregulated upon treatment with either of the two inhibitors, suggesting an repressive role of WNT in Arid3a expression (Figures 2A and S2A). To validate the negative role of WNT in Arid3a expression, we further examined an independent WNT overactivation model using our previously published WNT-high mouse organoids carrying a truncated APC (APC5) (16). Consistent with the observations earlier, Arid3a expression was downregulated, while the WNT targets Axin2 and Cyclin D1 were significantly increased in the WNT-high Apc5 organoids compared to WT control ( Figure 2B). Since WNT signalling regulates ISCs self-renewal and differentiation, the expression changes of Arid3a upon modulation of WNT in organoids might be caused indirectly by cell fate changes. To validate if WNT directly regulates Arid3a without change of cell fate, we switched to the human colorectal cancer cell line LS174T -with activated WNT signalling driven by β-catenin mutation -by treating them with two different doses of LF3 inhibitor (10 and 30μΜ) for 24 hours. qRT-PCR analysis showed a robust upregulated expression of ARID3A and downregulation of WNT target AXIN2 in a dose-dependent manner ( Figure 2C). Immunofluorescent staining further confirmed upregulated expression of ARID3A at the protein level in LF3-treated LS174T cells ( Figure S2B).
Since Arid3a is enriched at the early progenitor cells, we asked if its expression might be regulated by NOTCH signalling, the master regulator of early fate decision at the progenitors (7). Intestinal organoids were treated with the γ-secretase inhibitor DAPT (10μM) for 48h followed by qRT-PCR analysis. While DAPT treatment inhibited NOTCH target Hes1 and de-repressed Atoh1 expression, we did not observe any significant changes in Arid3a expression, indicating that its expression is independent of Notch signalling ( Figure S2C).
Apart from WNT and NOTCH, TGF-β and BMP signalling are also involved in intestinal homeostasis, while their dysregulation has been linked to cancer and other gastrointestinal diseases (17)(18)(19)(20). Interestingly, both TGF-β and BMP signalling form an expression gradient similar to that of Arid3a. To test whether the expression of Arid3a is regulated by the TGF-β superfamily, we treated mouse intestinal organoids with recombinant TGF-β1 and BMP4 ligands. As expected, BMP treatment activated its target gene Id1 and inhibited ISC marker Lgr5 expression (21,22). However, the expression of Arid3a was unaffected by BMP ( Figure   S2D). In contrast, treatment of organoids with recombinant TGF-β1 for 4h, 12h and 24h revealed that Arid3a expression was increased in a dose-dependent manner, similar to the expression of TGF-β target genes Smad7 and Id1 ( Figure 2D). It is important to note that the intestinal differentiation markers were not upregulated until 24h after TGF-β induction, indicating that Arid3a expression was driven by TGF-β signalling directly rather than because of increased differentiation ( Figure 2D).

Arid3a regulates transit-amplifying cell proliferation
It has been previously reported that Arid3a-null mice exhibit embryonic lethality due to defects in haematopoiesis (23). In order to characterise the functional role of ARID3A in intestinal epithelial cells, we generated an in vivo conditional knockout mouse model, VillinCre-ERT2 +/-; Arid3a fl/fl (Arid3a cKO), to delete Arid3a in Villin+ intestinal epithelial cells upon tamoxifen induction (24,25). Analysis of intestinal tissues 1 month post-tamoxifen administration confirmed complete abolishment of ARID3A expression at the epithelial cells, while the expression remained unchanged at stromal cells ( Figure 3A). Haematoxylin and Eosin (H&E) staining did not show any noticeable changes in the gross morphology of the Arid3a-depleted intestine ( Figure S3A). However, semi-quantitative pathologist scoring revealed that Arid3a cKO animals exhibited a minimal to mild villus atrophy ( Figures S3A and   S3B). Moreover, Arid3a cKO mice were also found to have shorter small intestine (mean= 35.35cm) when compared to WT (mean= 37.60cm) ( Figure 3B).
To characterise the molecular changes caused by Arid3a deletion, unbiased RNA sequencing (RNA-seq) analysis was performed on the Arid3a cKO and WT intestine.
Hierarchical clustering analysis showed that samples with the same genotypes readily clustered together ( Figure S3C). Differential gene expression analysis revealed a total of 4387 genes differentially expressed between the two groups (FDR cut-off <0.05), with 2413 genes upregulated and 1974 downregulated in the Arid3a cKO intestine (Supplemental information 1). In particular, we observed a mild to moderate decrease of various cell cycle and cell proliferation markers in the Arid3a-depleted intestinal crypts ( Figure 3C). Indeed, Gene Set Enrichment Analysis (GSEA) of mitotic cell cycle regulators indicated a strong downregulation in cKO intestine ( Figure 3D), while Metacore analysis identified the regulation of cell population proliferation as the most significantly affected Gene Ontology (GO) biological process ( Figure S3D). To assess the number of mitotically active cells in the crypts, animals were injected with a short pulse of 5-ehtynyl-2'deoxyuridine (EdU) to label cells undergoing de novo DNA synthesis or S-phase synthesis of the cell cycle. In accordance with our previous findings, Arid3a cKO intestine showed a reduction in total numbers of proliferative cells per crypt compared to WT animals (WT mean=9.73 cells; cKO mean=7.2 cells) ( Figure 3E).
Interestingly, quantitation of EdU-positive cells revealed that reduced proliferation was mostly observed at cell positions 9-15 (counting from the crypt bottom), whereas EdUlabelling was largely unchanged at cell positions 1-5 ( Figure 3F). This result suggests that Arid3a depletion inhibits proliferation at TA cells in the upper crypt whilst ISCs at the crypt base are mostly unaffected. Indeed, GSEA revealed a significant transcriptional downregulation of and TA cell gene signature (26) as well as in WNT signalling (27) at Arid3a cKO crypts ( Figures 3G and S3E). Of note, RNAscope analysis confirmed no difference in the expression of the ISC-specific marker Olfm4 between WT and Arid3a cKO tissues, indicating that the number of stem cells are not affected upon Arid3a deletion ( Figure S3F).
To validate the in vivo data, we generated ex vivo organoid cultures for functional analysis. Organoid formation analysis showed a significant decrease in organoid formation capacity in Arid3a cKO derived crypts (WT mean=64.1 organoids, cKO mean=48.4 organoids) ( Figure 3H). We further challenged the organoids by depleting one of the essential growth factors and WNT agonist, RSPONDIN (RSPO), in the culture medium to test the organoid dependency on exogenous WNT signal. Murine organoid cultures rely on supplementation of exogenous growth factors RSPO, EGF and NOGGIN to survive (28). Under normal condition, organoids were cultured in medium containing 5% RSPO conditioned media (CM). Organoids derived from WT and cKO animals were assessed and quantified based on their morphologies: organoids with >3 buds were considered healthy, organoids with 1-3 buds were considered unhealthy and organoids that failed to bud or form cysts were considered collapsed. WT and cKO organoids did not exhibit any major morphological differences in the presence of 5% RSPO CM ( Figure S3G). However, when organoids were challenged with a lower RSPO concentration (1%), a higher percentage of collapsed and a lower percentage of healthy organoids were observed ( Figure S3G). The increased dependence on exogenous RSPO in the cKO organoids suggests a reduction of endogenous WNT signalling in the Arid3a-depleted crypt cells, which is consistent with the GSEA data observed in Figure S3E.

Loss of Arid3a perturbs absorptive and secretory cell differentiation
Next, we asked if functional differentiation is affected in the cKO intestine. We first looked into differentiation of Paneth cells, the only specialised epithelial cells that reside at the crypt base adjacent to ISCs (7). RNA-seq analysis revealed downregulation of various Paneth cell markers in Arid3a cKO intestine, including Lyz1, the newly discovered marker Mptx2 (29), as well as a large number of anti-microbial peptides (α-defensins, Defa) secreted specifically from Paneth cells (30) (Figure 4A). Immunostaining of LYZ further confirmed the reduction of Paneth cells numbers in the cKO intestine (WT mean=14.53 cells/5 crypts, cKO mean=11.82 cells/5 crypts) ( Figure 4B). Paneth cells function as ISC niche by secretion of essential niche factors such as WNT ligands (31). Reduced Paneth cell numbers in Arid3adeficient intestine may shorten the WNT signal gradient in the crypt, leading to the observation of reduced WNT signature and TA cell proliferation.
In adult intestinal crypts, ISCs give rise to progenitor cells at the TA zone that will adopt either absorptive or secretory fate. Apart from Paneth cells, all other differentiated epithelial cells migrate up towards the villus after lineage commitment. GSEA showed significant enrichment of absorptive gene signature and an overall reduced secretion signature in the mutant intestine ( Figure 4C), suggesting that Arid3a regulates the differentiation ratio between absorptive and secretory cells. Recent studies further revealed that the differentiated cells do not acquire their terminal identity at the TA zone. Rather, the committed cells are further zonated along the villi to carry out different functions (10,11). In particular, enterocyte zonation can be grouped into five distinct functional clusters across the crypt/villus axis (11) ( Figure 4D). Interestingly, differential gene expression analysis and GSEA showed that Cluster 1 (early crypt enterocytes) was strongly enriched at WT animals, while the gene signatures of Clusters 2-5 were upregulated in Arid3a cKO intestines ( Figures 4E and   4F). Immunostaining of the villus tip-enriched markers APOA4 and ASPA confirmed their upregulation and perturbed zonated expression in the cKO intestines ( Figures S4A and S4B).
It has been previously shown that genes in Cluster 2 are associated with mitochondrial activity and clusters 3-5 are associated with absorption, nutrient transport, brush border function and cell adhesion (11). Interestingly, Metacore analysis of the significantly upregulated genes of the RNA-seq dataset (FDR<0.05, Fold change>1.5) showed that most of the top upregulated GO biological processes were related to an increase of different metabolic processes ( Figure   S4C), which are likely facilitated by the increased enterocyte absorptive gene signatures in Clusters 2-5. Furthermore, disaccharidase functional assay confirmed an increased sucrose breakdown to glucose in the Arid3a cKO organoids, indicating higher enterocyte digestion activity in the Arid3a-depleted intestinal epithelium compared to WT control ( Figure 4G). distinct enterocytes clusters that exhibit similarities with the previously described zonated clusters along the villus axis (11) ( Figure 5A). In accordance with the results observed earlier, we found that loss of Arid3a resulted in an increase in enterocyte populations (WT=51.3%, cKO=56%) and a reduction of secretory cells (WT=15.8%, cKO=12.4%) and TA cells (WT=13.2%, cKO=12.3%), whilst the number of Lgr5+ ISCs remains largely unaffected ( Figure   5B).
To better understand the changes in differentiation and cell fate dynamics, we performed pseudotime trajectory analysis which revealed 3 distinct major trajectories: differentiation trajectory towards 1) enterocyte lineage, 2) Goblet, EE and Paneth cells and 3) Tuft cells ( Figure 5C). This indicates that Tuft cells is a distinct cell lineage from other secretory cell types. Loss of Arid3a did not cause major alteration to the overall differentiation dynamics. Since we observed a significant increase in enterocyte signatures in Arid3a cKO intestine from bulk RNA-seq data, we ask if loss of Arid3a would affect enterocyte differentiation dynamics. Indeed, spatial mapping of the most differentially expressed enterocyte genes (based on the bulk RNA-seq) against pseudotime showed enriched expression of these genes towards earlier pseudo-timeline in the absence of Arid3a ( Figure   5D). This was confirmed by mapping the expression of selected enterocyte markers (S100g, Muc3, Ms4a18 and Ace2) in UMAP, which showed increased expression of these genes in the Arid3a cKO cells aligned at earlier pseudo-temporal trajectory compared to WT ( Figure 5E and  Figure 5H and Figure S5C).
Together, our data shows that Arid3a deletion resulted in reduced proliferation in the TA cells and enhanced enterocyte differentiation. Transcriptomic analysis using both bulk and scRNA-seq indicates that ARID3A modulates intestinal differentiation dynamics in the villus by fine-tuning the expression level of the spatial differentiation markers at the corresponding zones.

Increased binding and transcription of HNF1 and HNF4 in Arid3a-depleted intestine
In order to understand how loss of Arid3a perturbs intestinal spatial differentiation and causes changes in cell composition of the small intestinal epithelium, we performed ATAC-seq to assess the differences in chromatin accessibility between WT and Arid3a cKO intestine. Analysis of the ATAC-seq data showed that the Arid3a cKO intestine exhibits a more global open chromatin pattern when compared to WT ( Figure S6A). Although these changes were mild, the result was unexpected considering that Arid3a deletion caused increased differentiation that is often associated with less open chromatin (34). This suggests that the overall enhanced chromatin accessibility may not fully explain the perturbation of proliferation and differentiation caused by Arid3a loss. To gain more insight into the mechanism of ARID3A function, we performed footprinting analysis of our ATAC-seq using a recently published methodology -Transcription factor Occupancy prediction By Investigation of ATAC-seq Signal (TOBIAS) (35), which enables genome-wide analysis of transcription factor (TF) dynamics and calculates enriched motif binding using publicly available binding motifs of hundreds of transcription factors. Interestingly, TOBIAS analysis of the ATAC-seq data showed an enrichment of transcription factor binding sites in AT-rich genomic regions of the Arid3a cKO intestine ( Figure 6A). These included members of the ARID family (ARID3B and ARID5A) as well as members of the HNF family (Hnf1 and Hnf4). On the other side, WT intestine showed enrichment of binding motifs of FOS/JUN dimers (AP-1 pathway) ( Figure 6A), which has been previously linked with proliferation, apoptosis and carcinogenesis (36). To confirm this result, we utilised ChromVAR to assess the TF-associated chromatin accessibility. TFs with variability score >5 are associated with dynamic chromatin contributing to phenotypic changes, while TFs with variability score <5 are associated with permissive chromatin (37). In accordance with the TOBIAS analysis, FOS/JUN dimers and HNF4A-associated motifs were at the top two highest variability scores ( Figure 6B), indicating that changes of these TF dynamics contribute to the altered proliferation and differentiation in Arid3a cKO intestine.
Interestingly, HNF proteins have been previously associated with terminal maturation of enterocytes (38)(39)(40). It is conceivable that enrichment of HNF activity in the Arid3a cKO intestine results in increased spatial differentiation of enterocytes along the villus. Indeed, GSEA of the RNA-seq data showed a strong enrichment of the previously published Hnf4a/g target genes in the Arid3a cKO intestine ( Figure 6C), confirming the increased HNF4 transcription in the absence of Arid3a. In accordance with this finding, previously described intestinal epithelial-specific HNF4 targets (Clec2e and Slc30a10), showed higher chromatin accessibility in the promoter regions of the Arid3a-deficient intestine compared to WT ( Figure   6D). We further performed GO analysis of the 5000 most variable peaks between WT and Arid3a cKO intestine to evaluate the biological processes behind. Interestingly, we observed significant upregulation of many processes related to enterocyte functions in the cKO intestine including carbohydrate/monosaccharide metabolic processes and intestinal absorption ( Figure 6E), supporting the notion that HNF-mediated terminal differentiation of enterocytes is enriched in the mutant. Together with the transcriptomic data, we propose that ARID3A functions to maintain cell proliferation and prevent pre-mature HNF-mediated terminal differentiation at the TA progenitor cells.

Loss of Arid3a impairs irradiation-induced regeneration
Since deletion of Arid3a inhibits proliferation of the TA cells where progenitors reside, we asked whether this would affect the regenerative capacity of the intestine upon irradiation. Intestinal epithelium regenerates rapidly within days after irradiation: (1) apoptotic phase (1-2 days post-irradiation, dpi), (2) hyperproliferating/regenerating phase (3-4 dpi) and (3) normalisation phase (5 dpi onwards) (41). We have shown earlier that Arid3a forms an expression gradient from villus tip to the early progenitor cells at the crypt ( Figures   1C and 1D). We first asked if ARID3A expression is changed upon irradiation. To address that, we irradiated WT mice and collected the irradiated intestinal tissues at 1, 2, 3 and 4 dpi as well as the non-irradiated controls. Immunohistochemistry analysis showed collapsed crypts and transient loss of Arid3a+ cells on 1dpi, followed by crypt elongation and increased numbers of Arid3a+ cells at the upper crypt during the regenerating phase (3dpi).
Approaching the normalisation phase on day 4, the number of ARID3A+ cells returned to homeostatic levels ( Figure S7A). This suggests that Arid3a may play a role in intestinal regeneration and restoration of tissue homeostasis after injury.
Next, we investigated whether deletion of Arid3a perturbs the regenerative response.
Expert pathological analysis was performed on the irradiated tissues to assess the damage of the lamina propria, mucosa and gut-associated lymphoid tissue, and confirmed a more extensive tissue damage of cKO animals from day 2 to day 4 when compared to WT ( Figure   S7B and S7C). We then performed immunostaining of KI67 and Cleaved caspase-3 (c-CASP3) to assess tissue proliferation and apoptosis respectively ( Figure 7A and 7B). No major differences were observed between WT and cKO animals on 1 dpi, where extended cell death led to a dramatic reduction of proliferation. On 2 and 3 dpi, WT intestine started regenerating as evident by crypt expansion and increased proliferation, whilst Arid3a cKO intestine showed much lower numbers of KI67+ cells with minimal crypt expansion. By 5 dpi, WT intestine had returned to the normalisation phase, whereas the majority of Arid3a cKO crypts were still in the hyperproliferation phase with elongated crypts ( Figure 7A). To confirm the reduced proliferative capacity of crypts at early timepoints, we isolated crypts from WT and Arid3a cKO intestine collected on 1 and 3 dpi and performed organoid formation assay. No differences were observed on day 1 apoptotic phase where organoid formation efficiency was less than 10% ( Figure S7D). However, on 3 dpi, WT crypts had restored their capacity to form organoids while the colony formation efficiency remained low for the Arid3a depleted crypts (p-value=0.099) ( Figure S7D). This is consistent with the earlier observation that the regeneration phase of Arid3a cKO intestine was impaired on 3 dpi.
Interestingly, c-CASP3 staining showed no differences in the number of apoptotic cells between WT and mutant at the initial apoptotic and regenerating phase (1-3 dpi) (Figures 7B and 7C), suggesting that perturbation of regeneration was not directly caused by increased apoptosis. However, approaching the normalisation phase (4-5 dpi), Arid3a cKO intestine showed a significantly higher percentage of apoptotic crypts compared to control ( Figures 7B   and 7C). It is interesting to note that the number of apoptotic crypts in both WT and mutant intestine appeared to decrease gradually over time in a wave rather than linear pattern ( Figure 7D), suggesting that crypts unable to regenerate in the first wave after irradiation may collapse again. On the other hand, the apoptotic wave in the Arid3a-depleted intestine failed to subside and remained high over time ( Figure 7D). Of note, increased expression of apoptotic markers was also observed in the cKO intestine during homeostasis, suggesting that ARID3A may also play a role in regulating apoptosis ( Figure S7E).

Discussion
ARID3A is a transcription factor with DNA binding domain that interacts with A+T rich genomic regions (42,43). Functionally, ARID3A has been shown to drive normal development of both myeloid and B cell lineage specification, whereas deletion of the mouse Arid3a, leads to embryonic lethality due to defective haematopoiesis (44). Interestingly, Arid3a also regulates B cell response to antigen via post-translational palmitoylation of cytoplasmic ARID3A, leading to lipid rafts accumulation and B-cell antigen receptor (BCR) signalling (45).
Moreover, ARID3A is enriched in megakaryocytes compared to haematopoietic progenitor cells and has been shown to promote terminal megakaryocytic differentiation (46). However, the role of ARID3A in intestinal epithelium has not yet been explored. A recent single-cell analysis of the developing gut has identified Arid3a as one of the key regulators of intestinal epithelial development through transcription factor regulatory network mapping, whilst the mechanism remains unknown (47).
Here, we report for a previously unrecognised role of ARID3A in adult intestinal epithelial differentiation. We showed that ARID3A is regulated by WNT and TGF-β signalling Besides the binary cell fate decision at the early progenitors, it has been recently shown that both absorptive and secretory cells undergo spatial differentiation along the villus axis, resulting in regional and functional heterogeneity of all cell types (10,11). This implies that intestinal differentiation is a continuous process throughout the crypt-villus axis. More recent studies have shown that the zonation patterning of enterocytes is regulated by a BMP signalling gradient or villus tip telocytes (12,50), while BMP's downstream target c-MAF has also been reported to act as a regulator of the intestinal villus zonation programme (51). Our data suggests that TGF-β may also play a role in the spatial gene expression programme via ARID3A. The overall increased expression of villus gene signatures across all intestinal epithelial cell types in the mutant animals suggests that ARID3A may have an additional role of spatial differentiation in the villus by fine-tuning the expression levels of zonated genes.
Our data further shows a role of ARID3A in intestinal regeneration. It is believed that progenitor cells in the crypt are highly plastic to allow dedifferentiation into regenerative ISCs upon injury. Deletion of Arid3a switches the TA cells to higher differentiation-to-proliferation ratio, which may explain the impaired regenerative capacity by reduced plasticity. This suggests that ARID3A plays a gatekeeping role in the TA compartment to maintain the "justright" proliferation to differentiation ratio for tissue homeostasis and plasticity. Further investigation of the transcription factor network of ARID3A in the intestinal crypt-villus axis would help understand its unique role in fine-tuning the proliferation-differentiation switch.

Animals, drug administration and treatments
All animals were maintained with appropriate care according to the United Kingdom Animal Scientific Procedures Act 1986 and the ethics guidelines of the Francis Crick Institute.
The full list of transgenic mice used for this study is shown below in Table1. To induce conditional deletion, wild-type (WT) and knockout (cKO) animals were injected intraperitoneally with tamoxifen at 1.5mg/10g of mouse weight (from a 20mg/ml stock solution). Mice were culled by schedule 1 procedure (S1K) at the desired timepoint.
For irradiation induced-injury experiments mice were exposed to controlled 12 Gray (12Gy) total body ionising irradiation to induce damage using a Caesium (γ) irradiator. The dosage rate was 0.779Gy/min. Mice were culled by S1K at the desired timepoint.

Cell line immunofluorescence
For immunofluorescence (IF) experiments, LS174T cells were grown on sterilised glass coverslips in 24-well plates. Coverslips were not coated with poly-L-lysine before seeding the cells, since LS174T cells are very adherent. Cells were fixed with 4% paraformaldehyde (PFA) for 15min and permeabilised using 0.5% Triton X-100 in PBS for 15min. Cells were blocked with 1% Bovine Serum Albumin (BSA) for 1hr at room temperature before overnight incubation with primary antibodies at 4 o C (see table 3  Images were acquired as z-stacks using a Leica SPE confocal microscope and processed using Fiji.

Establishment and maintenance of mouse organoid cultures
Organoids were established from freshly isolated adult small intestine, as previously described (52). In brief, 2cm of jejunal small intestinal tissue was opened longitudinally and villi was scrapped using a glass cover slip. The remaining tissue was incubated in 15mM EDTA and 1.5mM DTT at 4 o C for 10min and moved to 15mM EDTA solution at 37 o C for an extra 10min. Subsequently, the tissue was shaken vigorously for 30sec to release epithelial cells from basement membrane and the remaining remnant intestinal tissue was removed. Cells were washed once, filtered through a 70μm cell strainer and resuspended in Cultrex BME Type 2 RGF Pathclear (Amsbio, 3533-01002). All freshly isolated organoids were maintained in either Intesticult medium (Stem Cell technologies, #06005) or in-house made basal medium containing EGF (Invitrogen PMG8043), NOGGIN, RSPONDIN and WNT3A (WENR medium), as previously described (28). The Rho kinase inhibitor Y-27632 (Sigma, Y0503) was added to the culture during first week of crypt isolation and single cell dissociation. For WNT, Notch, TGFβ and BMP pathway manipulation, organoids were passaged and allowed to recover for 72 hours before treatment. For WNT signalling inhibition, organoids were treated for 48hr with either 5μM of LGK974 inhibitor or 30μM of LF3 inhibitor (Sigma, SML1752); for Notch signalling inhibition, organoids were treated with 10μM of DAPT inhibitor (Sigma, D5942) for the indicated timepoints; for TGF-β signalling manipulation, organoids were treated with 0.1ng/ml of recombinant TGF-β1 (Sigma, 11412272001) for the indicated timepoints; for BMP signalling manipulation, organoids were treated with 20ng/mL of recombinant BMP4 (Peprotech, 120-05ET) for the indicated timepoints.
NOGGIN and RSPONDIN conditioned media were generated by HEK293T cells. WNT3A conditioned medium was generated from L cells. All images were acquired using an EVOS FL Cell Imaging System (Life technologies) and image brightness was adjusted using Adobe Photoshop (exactly same parameters were applied to all samples of the same experiment).

Mouse organoids assays
Organoids were established as described in the previous section. For organoid formation assay, crypts were counted using a brightfield microscope and 200 crypts were seeded in 20μl of Cultrex BME Type 2 RGF Pathclear in individual wells of a 48-well plate and cultured in WENR medium for 5 days until counted. Three technical replicates were performed per animal.
For RSPONDIN withdrawal assay, organoids were passaged and seeded in 3 10μl droplet per well of a 24-well plate. Organoids were allowed to re-establish in normal ENR medium (5% RSPONDIN) for 48hr. Subsequently, organoid medium was replaced and have organoids were cultured in ENR medium containing either 5% or 1% of RSPONDIN.
To detect disaccharide levels in organoids supernatants, organoids were washed twice with PBS and incubated with a 56mM solution of sucrose during 1h. Supernatants were collected and frozen until the assay was performed. To detect glucose content, Amplex® Red Glucose/Glucose Oxidase Assay Kit (Invitrogen, A22189) was used. Samples were diluted when necessary and incubated with the reaction buffer containing Amplex Red®, horseradish peroxidase and glucose oxidase. Fluorescence was measured in a Tecan microplate reader with an excitation wavelength of 540nm and fluorescence emission detection at 590nm.
Glucose concentration was assessed using a glucose standard curve from 0 to 200µM.

Crypt-villus fractionation
4cm of jejunal small intestinal tissue was opened longitudinally and villi was scrapped using a glass cover slip. Villi and the remaining intestinal tissue were transferred into two separate tubes and washed once. Both parts of the small intestine were incubated in 15mM EDTA and 1.5mM DTT at 4 o C for 10min and moved to 15mM EDTA solution at 37 o C for an extra 10min (as described in section 2.3.1) for isolation of epithelial cells of villi and crypt fragments. Pelleted cells were re-suspended in RLT buffer and stored at -80 0 C before proceeding to RNA extraction.

Fluorescent-activated Cell Sorting (FACS) of GFP-positive cells from Lgr5-EGFP-ires-CreERT2 mice
Crypts were harvested from the proximal jejunum (~10cm) as described in section Slides were scanned using an Olympus VS120 slide scanner and images were processed using QuPath (53).
For immunofluorescence, slides were incubated with Alexa-Fluor 488 or Alexa-Fluor 568 antibody for 1h, washed three times with PBS, incubated with 4',6'-diamidino-2phenylindole (DAPI) for 15 min to visualize nuclear DNA and mounted with ProLong Gold Antifade Mountant. Images were acquired as z-stacks using a Leica SPE or a Leica SP8 confocal microscope and processed using Fiji. For whole slide imaging, slides were scanned using an Olympus VS120 slide scanner and images were processed using QuPath (53). Mayer and for immunofluorescence slides were incubated with DAPI for 10min for DNA visualisation. Images acquired with an Olympus VS120 slide scanner and images were processed using QuPath (53).
For combined RNAscope and immunofluorescence of Arid3a with MUC2, CHGA or LYZ, samples were first stained for Arid3a using the red channel of duplex RNAscope kit, followed by antibody immunostaining as described above.

Bulk RNA-seq sample preparation
Crypts or villi were isolated from 10cm of mouse jejunal small intestinal tissue as described in "Crypt-villus fractionation" section and RNA was isolated as described in "RNA isolation" section. RNA integrity (RIN) was examined using Bioanalyzer 2100 RNA 6000 Nano kit from Agilent and RIN cut-off was set to 7. For crypt samples, libraries were prepared using The quality of the final DNA library was confirmed on the Agilent Tapestation before the samples were submitted to sequencing.

ATAC-seq data analysis
The nf-core/atacseq pipeline (version 1.  (66)). All data was processed relative to the mouse Ensembl GRCm38 release 95. A set of consensus peaks was created by selecting peaks that appear in at least one sample. Counts per peak per sample was then imported on DESeq2 within R environment for differential expression analysis. Pairwise comparisons between genotypes in each condition, and between conditions per genotype were carried out and differential accessible peaks were selected with an FDR < 0.05.
For footprinting analysis TOBIAS (v 0.12.10) (35) was used by running the following pipeline (https://github.com/luslab/briscoe-nf-tobias). The pipeline runs TOBIAS' ATACorrect, ScoreBigwig, BINDetect and generates PlotAggregate metaplots on merged replicate bam files. TOBIAS was run on set of consensus peaks used for the differential analysis (see above). As described before (35), all TFs with -log10(p-value) above the 95% quantile or differential binding scores smaller/larger than the 5% and 95% quantile are coloured. Selected TFs are also shown with labels.
For ChromVar analysis, the previously published R package was used: (http://www.github.com/GreenleafLab/chromVAR), to analyse sparse chromatinaccessibility data by estimating gain or loss of accessibility within peaks sharing the same motif or annotation while controlling for technical biases. Identified TFs were sorted based on their variability score.
For GO analysis, the 5000 peaks with the highest variability between WT and Arid3a cKO samples were chosen, and peaks were annotated with the closest gene prior to the analysis.

scRNA-seq sample preparation
Isolated mouse crypts were dissociated to single cells as

Data and code availability
The accession numbers for the raw and processed data used for this study was deposited on GEO and is publicly available (Bulk RNA-seq GEO: GSE242983, scRNA-seq GEO: GSE243155 and ATAC-seq GEO: GSE212560).