Increased matrix stiffness promotes fibrogenesis of hepatic stellate cells through AP-1-induced chromatin priming

doi:10.21203/rs.3.rs-4893653/v1

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Increased matrix stiffness promotes fibrogenesis of hepatic stellate cells through AP-1-induced chromatin priming

https://doi.org/10.21203/rs.3.rs-4893653/v1

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Matrix stiffness can have significant effects on cell behavior, regulating processes such as proliferation, differentiation, migration, and extracellular matrix production; however, less is known regarding the epigenomic and transcriptional regulation underling the effect of matrix stiffness on cell phenotypic shifts. In the present study, we utilized an in vitro system to assess the phenotypic shifts of hepatic stellate cells (HSCs) following changes in matrix stiffness, in addition to integrating multi-omics with imaging and biochemical assays to investigate the mechanism underlying the effect of mechanical stimuli on fibrosis. We show that cells cultured on a stiff matrix display more accessible chromatin sites, which consist of primed chromatin regions that become more accessible prior to the upregulation of nearby genes. These regions are enriched in fibrosis-associated genes that function in cytoskeletal organization and response to mechanical stimuli. Mechanistically, we demonstrate that activation of the AP-1 transcription factor family is responsible for chromatin priming, among which activated p-JUN is critical for the promotion of fibrogenic phenotypic shifts. The identified chromatin accessibility-dependent effect of matrix stiffness on cellular phenotypic shifts may be responsible for various fibrotic diseases and provide insight into intervening approaches.

Biological sciences/Genetics/Epigenomics

Biological sciences/Developmental biology/Transdifferentiation

Biological sciences/Genetics/Gene regulation

Matrix stiffness

phenotypic shifts

primed chromatin

pioneer transcription factors

p-JUN

Cells are subjected to a wide variety of mechanical stimuli, acting in multiple biological processes or diseases. For example, matrix stiffness, which refers to the rigidity of the extracellular matrix (ECM) surrounding cells, exerts various effects on cellular behavior such as proliferation, differentiation, migration, and ECM production (Wells, 2008). During development, mechanical cues can guide the differentiation of stem cells (Engler et al., 2006). Additionally, in most solid tumors, matrix stiffness alters the force-generation capability and metastatic potential of the cancer cells (Kraning-Rush et al., 2012). While it is known that matrix stiffness can induce changes in cytoskeletal organization and the expression of adhesion molecules, the manner by which matrix stiffness influences the expression and activity of epigenetic regulators and transcription factors, which play critical roles in the regulation of gene expression and cell fate, remains unclear. Moreover, fibrotic diseases, ECM dysregulation, wound healing, and tumorigenesis, are intimately associated with alterations in the mechanical microenvironment (Allen et al., 2020; Fattet et al., 2020; Jang et al., 2020; Stowers et al., 2019). Matrix stiffness has been implicated in fibrotic progression; however, the exact mechanisms underlying its contribution to disease pathogenesis remain elusive.

Recent advances have shown that mechanical force can regulate chromatin organization and that mechanosensitive transcription factors participate in the regulation of gene expression and cell state (Tajik et al., 2016; Uhler and Shivashankar, 2017). Considering the diverse phenotypic shifts that occur in hepatic stellate cells (HSCs) during the development and regression of liver fibrosis, the epigenetic changes induced by different matrix stiffness may be important contributors to shifts in cellular state (Mallat and Lotersztajn, 2013). The epigenome can act as a bridge connecting genotype and phenotype, and many studies have revealed that certain epigenomic changes can precede or foreshadow changes in gene expression (Ma et al., 2020; Zhang et al., 2021). For example, transient nuclear deformation and cyclic tensile stretch alter the levels and distribution of histone H3 trimethylation at lysine 9 (H3K9me3) and histone H3 trimethylation at lysine 27 (H3K27me3) to promote cell reprogramming or maintain genome integrity (Nava et al., 2020; Song et al., 2022). In addition, many transcription factors, such as YAP/TAZ and MRTF, are regulated by mechanical stimuli (Dupont et al., 2011; Miralles et al., 2003); however, the relationship between chromatin state and mechanotransduction during these processes is unclear, and the manner by which gene expression is regulated by chromatin remodeling needs to be explored further. Moreover, a recent advancement in multi-omics sequencing technologies, which enables joint profiling of the transcriptome and chromatin conformation, uncovered that widespread chromatin interactions are rewired before transcriptional activation (Liu et al., 2023).

Here, we established an in vitro system to explore the function of matrix stiffness on cell phenotypic shifts. LX-2, a human HSC line, was seeding on hydrogel, the stiffness of which was adjusted to physiologically relevant conditions of cirrhotic and normal liver tissues. The myofibroblastic differentiation of HSC is a critical event in liver fibrosis and is part of the final common pathway to cirrhosis in chronic liver disease from all causes, and the matrix is reported a determinant of its differentiation (Olsen et al., 2011). To investigate the mechanism underlying the regulation of HSC phenotypic shifts by matrix stiffness, we performed RNA-seq and ATAC-seq at two timepoints using cells exposed to an artificial matrix with two different stiffnesses, which allowed us to study the sequence of chromatin state changes and transcriptional regulation. Firstly, we found that cells cultured on a stiff matrix display a myofibroblastic phenotype. Meanwhile, chromatin accessibility and gene expression undergo reprogramming according to different matrix stiffnesses. Integrated analysis of the sequencing data identified primed chromatin sites that become accessible prior to the onset of gene expression, where the genes adjacent to these sites are mechanoresponsive and involved in fibrosis. Furthermore, we reveal that AP-1 family transcription factors play a key role in the reconstruction of chromatin accessibility and gene regulation in response to a stiff matrix. In summary, our data provide key insights into the molecular mechanism underlying the activation of HSCs by a stiff matrix and further explore the manner by which gene expression is reprogrammed through chromatin priming. These results also provide an explanation of the function of mechanical signaling in cell behavior, epigenomic landscaping, and transcription, which has important implications for better understanding biomechanics and disease pathogenesis.

Matrix stiffness induces shifts in the fibrotic phenotype and transcriptome

To decode the mechanism underlying the induction of cell phenotypic shifts by mechanical signaling, we utilized a well-studied in vitro system in which matrix stiffness can be tuned to mimic that of physiological tissue (Stowers et al., 2019), allowing us to study the effect of mechanical stimuli on cell behavior. Human LX-2 hepatic stellate cells, a commonly used cell line for the study of liver disease (Castilho-Fernandes et al., 2011; Wang et al., 2021b; Xu et al., 2005), were seeding on hydrogels possessing a Young’s moduli similar to that of normal (~ 2 kPa, soft) or cirrhotic (~ 40 kPa, stiff) liver tissue (Berzigotti, 2017; Yeh et al., 2002) for four days. When the HSCs cultured on tissue culture plastic for 7 days (similar to LX-2 cell in our work) were then transferred to either a soft or stiff matrix for further cultivation, the cells take a few days transiting from a rounded shape to a more spread, myofibroblast-like phenotype (Caliari et al., 2016). These results indicated that quickly after being transferred from TCP to matrices, cells start to set on activation and eventually adapt to the new mechanical environment. Based on this evidence, we collected the cells separately at two timepoints (two or four days) and conducted multi-omics analysis to explore the mechanism of cell phenotype transition induced by stiffness (Fig. 1a).

Firstly, we characterized the phenotype of cells on different hydrogels at both timepoints. Immunofluorescence of α-smooth muscle actin (α-SMA) stress fibers was used to identify activated HSCs since it is a common marker of fibrotic HSCs. The cells cultured on the stiff matrix displayed well-organized actin stress fibers than those on the soft matrix (Fig. 1b, Supplementary Fig. 1a). To quantitatively detect the cytoskeletal gene, the α-SMA expression levels were compared, and the results showed that α-SMA was slightly up-regulated after 2 days of culture on a stiff matrix, and significantly up-regulated after 4 days of culture (Supplementary Fig. 1b). These results demonstrate that LX-2 cells cultured on a stiff matrix display a fibrotic phenotype consistent with that of activated primary HSCs during fibrosis (Olsen et al., 2011).

Next, we asked the question whether the shift toward a myofibroblastic phenotype is accompanied by global transcriptomic changes. RNA-seq uncovered genome-wide changes in the transcriptional landscape in response to mechanical stimuli after two and four days. The expression of VCAN (Versican) and ITGB2 (Integrin Subunit Beta 2), markers of fibrotic cells (Berzigotti, 2017), was increased in cells cultured on the stiff matrix. The expression levels of MMP2 and MMP9, encoding matrix metalloproteinases that degrade the extracellular matrix, were decreased during fibrosis (Fig. 1c, 1d). A greater number of genes were differentially expressed at four days than at two days, and this alteration in gene expression is consistent with those reported in previous studies on active HSCs in liver fibrosis (Marquardt et al., 2012; Tsuchida and Friedman, 2017; Zhang et al., 2020). Gene Ontology (GO) enrichment analysis shows that the upregulated genes are enriched in fibrotic pathways and biological processes associated with activated HSCs, such as cell-cell adhesion, extracellular matrix organization, and wound healing processes (Fig. 1e, 1f). On the contrary, downregulated genes are enriched in immune response pathways (Supplementary Fig. 1c, 1d). And the previously reported gene signatures defined in the HSC activation mouse model were upregulated in cells cultured on a stiff matrix for two and four days (Supplementary Fig. 1e, 1f) (De Smet et al., 2021). Taken together, these results show that a stiff matrix induces the transition toward a fibrotic phenotype, in addition to the upregulation of fibrosis-related genes in HSCs. The stiff matrix-induced fibrotic phenotype became increasingly pronounced during continuous stimulation.

Increased matrix stiffness is accompanied by dynamic chromatin accessibility

To explore the mechanisms underlying transcriptomic changes in response to a stiff matrix, we assessed the chromatin accessibility dynamics using transposase-accessible chromatin with sequencing (ATAC-seq). Analysis of differentially accessible peaks (DAPs) on day 4 revealed 3,786 significantly more accessible peaks in cells cultured on the stiff matrix, with only 256 peaks found to be significantly less accessible (Fig. 2a). The genes adjacent to the more accessible regions in cells cultured on the stiff matrix for four days were enriched in cell proliferation and adhesion pathways (Supplementary Fig. 2a). To reveal the changes that occur during the HSC activation process and further study the function of alterations in chromatin state in gene regulation, ATAC-seq was performed using cells cultured on the stiff matrix for two days. A total of 750 more accessible peaks and 147 less accessible peaks were found (Fig. 2a). The genes adjacent to the more accessible regions in cells cultured on the stiff matrix for two days were enriched in protein phosphorylation and signal transduction pathways as well as wound healing, which is known to be associated with the activated HSC phenotype (Supplementary Fig. 2b). These data demonstrated that culturing HSCs in a stiff matrix induced a greater number of more accessible chromatin sites, which may be responsible for the upregulation of fibrosis-associated genes. To explore the manner by which DAPs induced by matrix stiffness affect the transcriptome, we characterized DAPs according to their relative locations to genes. DAPs were mostly located in intergenic and intronic regions, where enhancers were abundant and may play a key role in gene regulation (Fig. 2b) (Borsari et al., 2021).

Stiffness-induced dynamic chromatin accessibility precedes changes in gene expression

To better understand the relationship between chromatin accessibility and the regulation of fibrosis-associated genes in response to matrix stiffness, we compared the differentially accessible regions with gene expression patterns. We first identified the open chromatin regions unique to each sample and then defined regions either shared between cells cultured for two and four days or between soft and stiff matrices, classifying them into eight clusters (Fig. 2c). Subsequently, we evaluated the RNA expression patterns of genes adjacent to the chromatin regions in each cluster. As expected, the overall gene expression levels were positively correlated with promoter accessibility, for example IGFBP3 (Insulin-like Growth Factor Binding Protein 3) (Fig. 2c, Supplementary Fig. 2c), which is consistent with the notion that chromatin accessibility governs the expression of nearby genes (Shema et al., 2019). Notably, a larger number of chromatin regions became more accessible in cells cultured for two days on stiff matrix relative to soft matrix; however, the genes adjacent to these regions were not upregulated until day 4 (Fig. 2d; the “D2 stiff + D4 stiff” cluster). Such regions contain several fibrosis marker genes such as ITGB2 and COL8A1 (Collagen Type VIII Alpha 1 Chain) (Fig. 2d). We defined these regions as mechanically primed chromatin regions, where dynamic chromatin accessibility precedes and foreshadows changes in gene expression in response to mechanical stimuli (Fig. 2e). Previous studies have reported that chromatin priming plays an important role in facilitating gene activation during cell differentiation and lineage specification, since it allows for a more rapid response to external signals (Cruz-Molina et al., 2017; Spitz and Furlong, 2012). Based on this evidence, it is reasonable to suggest that primed chromatin may regulate fibrosis-associated gene expression in response to mechanical signaling.

We next examined the enrichment of GO terms for genes adjacent to each cluster of primed chromatin regions. Although individual clusters were enriched in GO terms for various functions, most clusters were enriched in GO functions related to HSC activation in liver disease (Fig. 2c) (Ramachandran et al., 2019; Seet et al., 2017). Notably, the genes associated with primed chromatin regions were enriched in functions related to actin cytoskeleton organization, response to mechanical stimuli, and mesenchymal-to-epithelial transition (Fig. 2c), which have been reported in activated HSCs (Long et al., 2022; Wagh et al., 2021). In addition, the genes that became more accessible in cells cultured on a stiff matrix for two days (the “D2 stiff” cluster) tended to participate in housekeeping kinase signaling pathways essential to the early stages of cell proliferation (Fig. 2c). However, the genes that became more accessible in cells cultured on a stiff matrix for four days (the “D4 stiff” cluster) were related to liver development and membrane protein proteolysis (Fig. 2c) (Principi et al., 2023), which have been reported to be activated during fibrotic progression (Brusatin et al., 2018; Hernandez-Guerra et al., 2019), indicating that the activation of mechanical signaling transduction pathways affects cell behavior. The accessible peaks shared by cells cultured on the soft and stiff matrix for two days were enriched in cell migration and cytoskeletal organization pathways that contribute to cell spread and colonization on hydrogels (Fig. 2c). By contrast, the peaks shared by cells on day 4 were enriched in adherens junction organization and epithelial transition pathways related to increased cell density (Fig. 2c). Collectively, dynamic chromatin accessibility induced by the stiff matrix is associated with diverse functional pathways converging to mechanosensing genes in response to mechanical stimuli.

Primed chromatin accessibility remodeling is positively associated with activation of the H3K27ac modification

To explore the mechanism underlying the establishment of primed chromatin, we characterized the genomic features of the primed chromatin regions and subsequently employed chromHMM to train 10 chromatin states of HSCs using five epigenetic markers (Ernst and Kellis, 2012). The chromatin states were trained using ENCODE ChIP-seq data generated from primary HSCs. Previous studies have reported that changes in histone modifications may precede changes in gene expression, resulting in the establishment of primed chromatin (Bernstein et al., 2006). We found that primed chromatin regions were enriched in bivalent promoters (Fig. 3a) containing both H3K27me3 and H3K4me3, representing repressive and activating histone modifications that maintain these chromatin regions poised for activation (Bernstein et al., 2006). Moreover, primed chromatin regions were also enriched in active enhancers (Fig. 3a) containing H3K27ac which are responsible for enhancer activation (Creyghton et al., 2010). These results are in accordance with those previously reported that H3K27ac modifications may mark genomic regions primed for activation (Wang et al., 2022).

To further explore the association between histone modifications and the establishment of primed chromatin in response to matrix stiffness, we performed Cut&Tag experiments targeting H3K4me3, H3K27me3, and H3K27ac in LX-2 cells cultured for two days on soft and stiff matrices, and compared the changes in histone modifications among the 8 chromatin accessibility clusters (Kaya-Okur et al., 2019). H3K4me3 and H3K27me3 levels remained stable in all the clusters in response to the stiff matrix (Fig. 3b). Correlation results further validated that histone methylation is only weakly correlated with primed chromatin accessibility (Supplementary Fig. 3a, b). Consistently, immunofluorescence imaging shows a globally similar distribution of histone methylation modifications (Fig. 3d). By contrast, the H3K27ac modification level was specifically increased in the “primed chromatin” and “D2 stiff” clusters but not in the other clusters, indicating specifically increased chromatin accessibility in these regions in response to the stiff matrix (Fig. 3b). As an example, the fibrosis marker gene ITGB2 displayed increased chromatin accessibility and H3K27ac modification levels; however, there was no change in the histone methylation levels within its promoter region (Fig. 3c). This finding is consistent with previous reports that H3K27ac contributes to chromatin accessibility and distinguishes active from inactive regulatory elements (Creyghton et al., 2010). Taken together, these results suggest that H3K27ac is involved in the process by which stiffness-induced primed chromatin is prepared for transcriptional activation.

AP-1 factor motifs are enriched in primed chromatin regions in response to matrix stiffness

The universality of priming mechanisms that include both selective histone modifications and transcription factor (TF) binding has been confirmed by genome-wide studies (Gifford et al., 2013; Wamstad et al., 2012). Moreover, one study focusing on chromatin reprogramming and genome activation revealed that pioneer factors can bind nucleosomal DNA to regulate histone modifications and chromatin accessibility during zygotic genome activation (ZGA), suggesting that H3K27ac is not absolutely required for chromatin opening (Schulz and Harrison, 2019). Furthermore, TFs are regarded as master regulators of the epigenome and global transcriptional activity (Balsalobre and Drouin, 2022; Barrero et al., 2010). To explore potential TFs responsible for the primed chromatin regions induced by a stiff matrix, we performed HOMER enrichment analysis to identify enriched TF binding motifs (Arpa and Durbeej, 2023). The activator protein-1 (AP-1) and TEAD families were the most enriched motifs in primed chromatin regions (Fig. 4a). TFs in the AP-1 family bind DNA and contribute to genome-wide remodeling of the epigenetic landscape, and thus may function as pioneer factors (Shaulian and Karin, 2001; Soufi et al., 2015; Vierbuchen et al., 2017). In support of this role, the AP-1 motif had stronger footprint signals in primed chromatin regions of cells cultured on the stiff matrix (Fig. 4b). These results suggest that AP-1 family TFs may be responsible for primed chromatin regions and downstream changes in gene expression.

Next, we asked the question whether the increased mRNA expression or enhanced binding of activated AP-1 family TFs contributes to primed chromatin remodeling in response to matrix stiffness. As the previous studies reported, AP-1 is a menagerie of dimeric basic region-leucine zipper (bZIP) proteins that belong to the Jun (c-Jun, JunB, JunD), Fos (c-Fos, FosB, Fra-1 and Fra2), Maf (c-Maf, MafB, MafA, MafG/F/K and Nrl) and ATF (ATF2, LRF1/ATF3, B-ATF, JDP1, JDP2) sub-families, each sub-family of TFs share some similarities in structure and function (Eferl and Wagner, 2003; Shaulian and Karin, 2002). RNA-seq shows that the expression of most AP-1 TFs remained stable in cells cultured on the soft and stiff matrices (Fig. 4c). The scRNA-seq using primary HSCs from patients with liver cirrhosis and healthy donors (Ramachandran et al., 2019) showed that the transcription level of JUN was moderately decreased in cirrhotic livers (Supplementary Fig. 4a-d). Taken together, these results suggested that JUN expression remained stable and the mRNA expression level was not the main reason for its enhanced enrichment on primed chromatin in response to the stiff matrix.

Although TF expression provides the first line of evidence for the locations in which they may function, their activity often depends on post-transcriptional events and the mRNA levels do not necessarily indicate regulatory activity (Vaquerizas et al., 2009). As TFs can move around and bind primed chromatin without necessarily increased expression, we next focused on exploring whether the binding of activated AP-1 TFs was enhanced resulting in primed chromatin state. Since some of the AP-1 TFs (JUN, FOS) can be phosphorylated at distinct residues by different kinases and thus possesses enhanced transcriptional activity (Karin, 1996), we compared the change of protein levels of JUN, p-JUN, FOS, p-FOS and ATF3 by western blot. We found that p-JUN and ATF3 expressed at higher protein levels in cells cultured on a stiff matrix for two days, while other factors in the AP-1 family remained stable at the protein level (Fig. 4d). Next, we used the Cut&Tag technique targeting p-JUN and ATF3 in the same experimental batch to capture its binding affinity in the different clusters of chromatin regions. An increased binding level of p-JUN was found specifically in primed chromatin regions (Fig. 4e), and the fraction of p-JUN on accessible peaks also showed a preference for primed chromatin regions (Fig. 4f). However, the enrichment of ATF3 in primed chromatin regions remains unchanged in response to matrix stiffness (Supplementary Fig. 5a). Additionally, the fraction of ATF3 on accessible peaks does not show a preference for primed chromatin regions like p-JUN (Supplementary Fig. 5b). JUN is the most efficacious transcriptional activator in the AP-1 family (Shaulian and Karin, 2002) and is considered an important regulator of hepatic stress responses (Schulien et al., 2019). Taken together, these results indicate that AP-1 factors, in particular p-JUN, are enriched in primed chromatin regions and may mediate their establishment in response to mechanical stimuli.

Phosphorylation of JUN promotes chromatin accessibility in response to matrix stiffness

To further explore the mechanism underlying the regulation of chromatin accessibility by p-JUN, we designed plasmids containing mutated phosphorylation sites at Ser⁶³ and Ser⁷³ within JUN to achieve enhancement (the JUN-EE mutant) of its transcriptional activation. Moreover, we also designed the JUN-EE-Δbasic and JUN-EE-Δleucine mutants to interfere with the DNA-binding activity of JUN and its dimerization with other TFs, respectively (Fig. 5a) (Lynn et al., 2019). To quantify the phosphorylation level of JUN mutants (JUN-EE, JUN-AA), we utilized the normalized fluorescent intensity to compare the relative expression levels of p-JUN in cells. The results showed that the phosphorylation level of cells with JUN-EE mutant was approximately 3-fold higher than those in control cells, while the phosphorylation level of JUN-AA mutant was about 3-fold lower than control cells (Fig. 5b, 5c), which is consistent with the expected functions of JUN mutants (Lynn et al., 2019).

After that, ATAC-seq was performed to identify changes in chromatin accessibility. Enhanced phosphorylation of JUN (the JUN-EE mutant) resulted in increased chromatin accessibility in the primed chromatin regions of cells cultured on a soft matrix (Fig. 5d). By contrast, the JUN-EE-Δbasic and JUN-EE-Δleucine mutants resulted in reduced chromatin accessibility as compared with JUN-EE and causing the chromatin accessibility similar to that of control cells (Fig. 5d), which was in line with the mechanism that the dimerization and DNA binding activity were required for p-JUN to perform its transcription function (Shaulian and Karin, 2002). Accordingly, we also compared the ATAC-seq signal in the other chromatin regions excluding primed chromatin and found that the chromatin accessibility increased moderately, suggesting a specific function of p-JUN in the establishment of primed chromatin (Supplementary Fig. 4e). We then investigated whether p-JUN is indispensable for chromatin accessibility in response to matrix stiffness. The deficient phosphorylation of JUN (the JUN-AA mutant) resulted in decreased chromatin accessibility in the primed chromatin regions of cells cultured on a stiff matrix (Fig. 5e). Accordingly, when comparing the ATAC-seq signal in the other chromatin regions excluding primed chromatin, we also found decreased chromatin accessibility (Supplementary Fig. 4f). These results indicate that p-JUN is required for the establishment of accessible chromatin globally.

Furthermore, we examined the transcriptional changes of fibrotic genes upon the gain and loss of function of p-JUN by day 4, the results showed that the expression of the fibrosis marker α-SMA was increased in cells with enhanced p-JUN, while its expression was decreased in cells with deficient p-JUN (Fig. 5f). Immunofluorescence imaging shows that the cells containing the JUN-EE mutant had well-organized actin stress fibers compared with the control cells expressing wild-type JUN, conversely, the JUN-AA resulted in deficient stress fiber organization (Fig. 5g). These results indicated that the phosphorylation of JUN could promote the accessibility of primed chromatin and the phenotypic shift towards fibrosis.

Given that the phosphorylation of JUN regulates chromatin dynamics under mechanical stimulation, we further sought to elucidate the upstream signaling pathways responsible for the changes in extracellular matrix stiffness and regulation of TF phosphorylation levels. It has been suggested that extracellular matrix stiffness may regulate the activities of cytoplasmic kinases and subsequently influence cell behavior (Wang et al., 2021a). The conserved extracellular signal-regulated kinase (ERK) signaling pathway is activated by a variety of extracellular stimuli (Lavoie et al., 2020). To address the participation of upstream signaling pathways after the application of matrix stiffness, we performed immunofluorescence assays targeting p-ERK. We found that p-ERK had a higher nuclear-to-cytoplasmic fluorescence ratio in response to a stiff matrix (Fig. 5h, i). Next, we used specific inhibitors of ERK, JNK kinases, and p38, which have been reported to be important in the response to stimuli and tumorigenesis (Fig. 5j, k). Lower p-JUN levels were observed following inhibition of p-ERK, while the inhibition of other kinases or pathways had no effect, suggesting that p-ERK is critical for the activation of JUN. These results indicate that the ERK pathway acts as a molecular link between matrix stiffness and chromatin accessibility, which in turn enables cells to dynamically regulate gene expression and adapt to altered mechanical environments. Moreover, we continuously observed the nuclear-cytoplasmic fluorescence ratio of p-ERK on the stiff matrix to explore the timing and duration of p-ERK activity. The results showed that the activation of ERK responded to stiffness in a relatively short period of time (within 2 hours) and maintained the activated state for a long time (Supplementary Fig. 6a, 6b). Previous publications reported that the translocation of ERK can be either transient, with a peak at around 10–15 min after stimulation, or prolonged, where most ERK molecules enter the nucleus within 10 min and can be retained there by various nuclear anchoring moieties for up to 3 h or even longer (> 10 h) (Ram et al., 2023; Shankaran et al., 2009). Based on these, we proposed that the matrix stiffness can be regarded as continuous and long-term extracellular stimulus to the cell, thereby leading to a sustained ERK activity responding dynamics.

Here, we demonstrate that extracellular matrix stiffness induces cell phenotypic transition through the remodeling of chromatin accessibility. Moreover, we show that the establishment of primed chromatin in response to matrix mechanical stimuli precedes the onset of gene transcription. Overall, based on our work and prior knowledge, we propose a model underlying the regulation of gene expression in response to mechanical stimuli. At the first stage, the cells are subjected to the increased matrix stiffness on a shorter time scale, the cytoplasmic kinases ERK would be activated and translocate to the nucleus and then phosphorylate the pioneer factor (JUN), leading to its transcriptional activation state (p-JUN) and promoting its binding to chromatin. Subsequently, the pioneer factor can initiate chromatin opening to establish a primed chromatin state. During this process, we find that the H3K27ac modification is increased, which is consistent with the previous report that the pioneer factor can facilitate the recruitment of the general transcription co-activator p300–CREB-binding protein (CBP) and acetylated H3K27 (Balsalobre and Drouin, 2022). However, the nearby genes are not yet transcriptionally activated. At the second stage, when the cells are subjected to stiff matrix stimuli on a longer time scale, the genes near the pioneer factor start to be activated and expressed, and the cells exhibit a pronounced fibrotic phenotype (Fig. 6). In summary, our findings emphasize a major role for chromatin priming in the response to extracellular matrix mechanical stimuli, which sheds light on the gene regulation mechanism in cells cultured on a stiff matrix. Given that both histone acetylation and pioneer factors are involved in dynamic chromatin accessibility, the detailed functions of AP-1 factors and histone acetyltransferases (HATs) in the regulation of the chromatin landscape remain unclear. Although previous studies have revealed that pioneer factors cooperate with HATs (such as p300/CBP) to modulate H3K27ac modification (Choi et al., 2016), it remains to be established whether the AP-1 family could promote HAT recruitment.

Several significant works have discovered the contribution of extracellular matrix stiffness to cell phenotypic transition through transcriptome alterations (Engler et al., 2006; Ondeck et al., 2019; Panciera et al., 2020). Moreover, the chromatin landscape can also be affected by matrix stiffness; for example, changes in matrix stiffness can drive changes to the nucleus and chromatin state through Sp1-HDAC3/8-mediated pathways, in which Sp1 can recruit HDACs (histone deacetylases) to further modulate chromatin accessibility (Stowers et al., 2019). Although chromatin epigenetics is considered a common mechanism for gene expression regulation, it has not been systematically studied with respect to mechanosignaling-induced cell state changes. Here, by collecting cells cultured on a stiff matrix for different periods and detecting their chromatin accessibility and gene regulation, we observed that changes in chromatin accessibility precede changes in gene expression, creating primed chromatin states that prepare genes for activation to alter the hepatic stellate cell phenotype. These observations are consistent with the previous definition about primed chromatin state (Calo and Wysocka, 2013) and indicate that mechanical stimuli initially affect chromatin epigenetics, which subsequently causes changes in cell phenotype, suggesting that the regulation of chromatin accessibility may potentially aid in the diagnosis and treatment of fibrotic diseases at an earlier stage.

The transcriptional regulation controlled by epigenetics, such as DNA methylation, histone modifications, has been widely explored in the context of hepatic stellate cells (HSCs) activation and the mechanical activation of other mesenchymal cells. Epigenetic events that control HSC activation and function are highly dynamic processes (Marcher et al., 2019), and several relevant epigenetic mechanisms has been reported. For example, the DNA methyltransferases (DNMTs), which could regulate the CpG methylation, mediated hypermethylation of phosphatase and tensin homolog (PTEN) activates HSCs (Bian et al., 2012). Tian et al. reported that Myocardin-related transcription factor A (MRTFA, also known as MKL/myocardin-like protein 1) orchestrates profibrogenic transcription by recruiting a histone methyltransferase complex to the promoters of fibrogenic genes by regulating the H3K4 methylation (Tian et al., 2016). In our work, we defined the primed chromatin regions, where the chromatin accessibility precedes and foreshadows changes in gene expression. Moreover, we characterized the genomic features of these regions and revealed that the primed chromatin was enriched in bivalent promoters and active enhancers. And we further uncovered that their accessibility is associated with H3K27ac modifications, indicating that the histone modification plays a role in chromatin dynamic in response to the mechanical stimuli.

Epigenetic regulators and transcription factors can also organize the genome into accessible or closed regions orchestrating the appropriate transcriptional program in the cell (Tee and Reinberg, 2014). Many previous studies have reported that YAP1 is a critical driver of HSC activation, as its nuclear translocation is followed by the induction of YAP1 target genes during activation (Machado et al., 2015; Mannaerts et al., 2015; Swiderska-Syn et al., 2016), and pharmacological inhibition of YAP1 improves liver fibrosis in mice (Martin et al., 2016). Jang et al. using the human gastric cancer cell line revealed that the DNA methylation of the promoter region of the oncogenic YAP is reversibly regulated by the stiffness of the extracellular matrix (Jang et al., 2020). Additionally, TEAD proteins have been shown to interact with YAP/TAZ, a key component of the Hippo signaling pathway, could sense the physical nature and drive tumorigenesis (Panciera et al., 2017; Piccolo et al., 2014). In our work, we found that AP-1 family, including the JUN, FOS, and ATF sub-families, which are known as the pioneer transcription factors can initiating chromatin remodeling and involved in the regulation of the developmental process (Shaulian and Karin, 2002; Thompson, 2020), are involved in the stiffness-induced cell phenotypic transition. Our findings are consistent with previously studies that AP-1 act as “activating” transcription factors in the activated HSCs by controlling transcription of potent pro-fibrogenic genes (Park et al., 2005; Smart et al., 2001). Moreover, YAP/TAZ/TEAD and AP-1 form a complex that synergistically activates target genes, and the YAP/TAZ-induced oncogenic growth is strongly enhanced by gain of AP-1 and severely blunted by its loss (Zanconato et al., 2015). Taken together, although there have been several indications that chromatin epigenomic remodeling and transcription factor play a key role in mechanical transduction and cell phenotype shifts, our work reveals a complex interplay by which mechanical signals can be transduced to the nucleus to alter chromatin accessibility through the activation of the regulatory factor JUN, which initiates chromatin priming and drives cell phenotypic shifts. The detailed mechanism between the epigenetic regulators and TFs are still needed further study.

In addition, the in vitro system, in which the hydrogel stiffness was adjusted to physiologically relevant conditions of cirrhotic and normal liver tissues, enables us to study the effects of mechanical stimuli on cell behavior during the fibrotic disease process. Liver fibrosis is directly caused by chronic liver inflammation and leads to the development of cirrhosis and hepatocellular carcinoma (Kisseleva and Brenner, 2021). HSCs are the main contributors of the extracellular matrix to the fibrotic liver, increasing the stiffness of the microenvironment. Simultaneously, this stiffness can be detected by HSC surface receptors called integrins, allowing HSC activation and differentiation to myofibroblasts (Hynes, 2002; Zhang and Friedman, 2012). Current models regard the relationship between extracellular matrix stiffness and HSC phenotypic transition as a chain of circumstances that results in end-stage fibrotic disease (Mueller and Sandrin, 2010). Although liver cirrhosis is a relatively complex process in vivo, our experiment built on foundational studies was able to partially delineate the differentiation process from HSCs to myofibroblasts in response to matrix stiffness. Despite many signaling pathways, molecules, and mechanisms reported to be involved in phenotypic shifts of HSCs (Kisseleva and Brenner, 2021), it remains unclear to what extent matrix stiffness can drive phenotypic shifts of HSCs and whether these shifts are accompanied by epigenetic changes. Here, we explored the mechanism underlying the effect of mechanical cues from the microenvironment on cell phenotype using matrix stiffness as the sole variable. More broadly, fibrotic diseases encompass pathological angiogenesis in multiple organs, among which the common feature of fibrosis is conserved.

We reveal that the activated AP-1 could response to the stiffen matrix and established the primed chromatin state that prepare genes for cell phenotypic shifts. Although we emphasized that the phosphorylated and activated of p-JUN play a key role in transducing the mechanical stimuli to the nucleus, further studies are needed to cexplore whether p-ERK nuclear activity sufficient to overcome soft matrix effects, and whether this explicitly requires Jun phosphorylation. And acknowledging the complexity of cellular mechanosensing pathways is also crucial.

In summary, this work integrates time-resolved, genome-wide profiles to explore fibrotic cell fate and provides insights into the involvement of transcription factors and chromatin dynamics in mechanical-related fibrotic diseases and liver homeostasis. Understanding the molecular events involved in the establishment and maintenance of chromatin accessibility and gene regulation in response to mechanical stimuli will benefit the design of strategies for cellular reprogramming, differentiation, and cancer treatment.

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REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
rabbit anti-SMA	Abways	CY5295
rabbit anti-P-FOS	Cell Signaling Technology	5348T
rabbit anti-P-JUN	Cell Signaling Technology	3270T
rabbit anti-ATF3	Abcam	ab254268
mouse anti-c-JUN	Santa Cruz Biotechnology	sc-74543
mouse anti-c-FOS	Santa Cruz Biotechnology	sc-166940
mouse anti-GAPDH	Abcam	Ab8245
rabbit anti-H3K27ac	Abcam	ab4729
rabbit anti-H3K4me3	Abcam	ab8580
mouse anti-H3K27me3	Abcam	ab6002
Biological samples
Goat anti-Rabbit IgG HRP	EASYBIO	BE0101
Goat anti-Mouse IgG HRP	EASYBIO	BE0102
Alexa Fluor 594 donkey anti-mouse	Thermo Fisher Scientific	A21208
Alexa Fluor 594 donkey anti-rabbit	Thermo Fisher Scientific	A21207
Alexa Fluor 488 donkey anti-rabbit	Thermo Fisher Scientific	A21206
Alexa Fluor 647 goat anti-mouse	Thermo Fisher Scientific	A21235
Alexa Fluor 488 donkey anti-mouse	Thermo Fisher Scientific	A21202
Chemicals, peptides, and recombinant proteins
TruePrep DNA Library Prep Kit V2	Vazyme	TD502
HiScript® II 1st Strand cDNA Synthesis Kit	Vazyme	R211-01
NovoNGS® CUT&Tag 3.0 High-Sensitivity Kit	NovoProtein	N259-YH01
Nuclear and Cytoplasmic Protein Extraction Kit	YEASEN	20126ES50
MolPure® Cell RNA Kit	YEASEN	19231ES50
ERK1/2 inhibitor	MedChemExpress	PD98059
JNK1/2/3 inhibitor	MedChemExpress	SP600125
P38 inhibitor	MedChemExpress	SB203580
Deposited data
RNA-seq, ATAC-seq, Cut&Tag	This paper
ChIP-seq	ENCODE	ENCFF781SNK, ENCFF050OMW, ENCFF760ABV, ENCFF064AZN, ENCFF055ACA, ENCFF539DCQ, ENCFF414RVB
Experimental models: Cell lines
Human LX-2 cells	Procell	CL-0560
Oligonucleotides
The primers used for qPCR can be found in Table S1.	This paper
Software and algorithms
Bowtie2	Langmead and Salzberg (2012)	https://bowtie-bio.sourceforge.net/bowtie2/index.shtml
MACS2	Zhang et al. (2008)	https://github.com/taoliu/MACS
Homer	Heinz et al. (2010)	http://homer.ucsd.edu/homer/motif/
STAR	Dobin et al. (2013)	https://github.com/alexdobin/STAR
DESeq2	Love et al. (2014)	http://www.bioconductor.org/packages/release/bioc/html/DESeq2.html
DiffBind	Stark and Brown (2011)	https://bioconductor.org/packages/release/bioc/html/DiffBind.html
Samtools	Li et al. (2009)	http://samtools.sourceforge.net
Trimgalore	N/A	http://www.bioinformatics.babraham.ac.uk/projects/trim_galore
R	N/A	https://www.rproject.org/
ImageJ	N/A	https://imagej.net/ij/index.html
IGV	Robinson et al. (2011)	https://www.igv.org/
chromHMM	Ernst and Kellis (2012)	https://compbio.mit.edu/ChromHMM/
PrimerBank	Wang et al. (2012)	https://pga.mgh.harvard.edu/primerbank/

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Yujie Sun ([email protected]).

Materials availability

This study did not generate new unique reagents and biological materials

Data and code availability

The raw data of ATAC-seq, RNA-seq and Cut&Tag have been deposited in Gene Expression Omnibus (GEO) under the accession number GSE220703.

This paper does not report original code. The software used in this study is described in the above section in details. Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

Method details

Hydrogel formation

The polyacrylamide (PA) hydrogels were prepared following previous protocols (Kaylan et al., 2018; Tse and Engler, 2010; Wen et al., 2014). Briefly, clean glass coverslips were functionalized using 3-(trimethoxysilyl) propyl methacrylate to facilitate covalent attachment of polyacrylamide gel. The polymer solution contained acrylamide monomers, the crosslinker N, N methylene-bis-acrylamide, ammonium persulphate, and N, N, N’, N’-tetramethyl ethylenediamine (TEMED, T22500, Sigma-Aldrich). Polyacrylamide gel was sandwiched between a functionalized coverslip and a dichlorodimethylsilane-treated slide (40140, Sigma-Aldrich). The ratio of % acrylamide/% bis-acrylamide was 7.8/0.8 and 10.4/0.5 for soft and stiff gel, respectively. Subsequently, substrates were incubated in 0.1 mg/mL N-sulphosuccinimidyl-6-(40-azido-20-nitrophenylamino) hexanoate (sulpho-SANPAH, 22589, Sigma-Aldrich) and activated with ultraviolet light. Finally, a thin layer of plasma fibronectin (0.1 mg/mL, F2006, Sigma-Aldrich) was crosslinked to the gel surface by overnight incubation at 4°C.

Cell culture

Immortalized human HSC cell line LX-2 (Procell, CL-0560) were cultured in DMEM supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin at 37°C with 5% CO2 as described previously (Xu et al., 2005). Cells were seeding on the hydrogels at 50,000 cells ml−1 final concentration and cultured for 2 or 4 days and then were collected for downstream experiments.

Knockdown and overexpression

Mutants and truncations of human JUN (Gene ID: 3725) were cloned into the pEGFP-C3 vector for transient expression in LX-2 cells. The JUN-AA mutant featured point mutations where Ser⁶³ and Ser⁷³ were changed to Ala, simulating the inhibition of phosphorylation. The JUN-EE mutant contained point mutations where Ser⁶³ and Ser⁷³ were altered to Glu, simulating phosphorylation. JUN-EE-Δbasic contained a truncated DNA-binding domain through deletion of the 254–280 gene segment based on JUN-EE. JUN-EE-Δleucine contained a truncated dimerization domain through deletion of the 280–314 gene segment based on JUN-EE. Each of plasmid contains a GFP tag. Cells were transfected with the plasmids using Neofect™ DNA transfection reagent.

Immunofluorescence

Cells were fixed with 4% paraformaldehyde for 15 min at room temperature, washed twice with PBS, and blocked with 5% fetal bovine serum for 1 h at room temperature. Subsequently, cells were incubated with primary antibodies at room temperature for 1 h, washed three times with PBS, and incubated with the corresponding donkey anti-rat/rabbit Alexa Fluor^® 488/594-conjugated secondary antibodies (at final concentration 2 µg/mL) for 1 h at room temperature. The following primary antibodies were used in this study: rabbit anti-α-SMA (Cat. no. CY5295, 1:100, Abways), rabbit anti-P-JUN (Cat. 3270T, 1:100, Cell Signaling Technology). Finally, samples were observed using confocal fluorescence microscopy. The data was processed using ImageJ software. As for the normalized mean fluorescence intensity of immunostaining signals of p-JUN in cells, the fluorescence background in the cytoplasm was subtracted. The total fluorescence intensity was then normalized by dividing the maximum value of each panel to adjust shown range in y-axis. Each point on the scatter plot represents a cell. About 15 pairs of cells within the same imaging view were analyzed for each independent experiment.

Western blotting

Total protein was extracted from the cells using SDS-PAGE protein loading buffer (YEASEN, 20315ES05) containing protease inhibitors (PMSF) and phosphatase inhibitors (Roche, PhosSTOP). Cytoplasmic and nuclear proteins were extracted separately using the Nuclear and Cytoplasmic Protein Extraction Kit (YEASEN, 20126ES50) according to the manufacturer’s protocol. Samples were subjected to SDS-PAGE and transferred to nitrocellulose membrane, which was then blocked with 5% skimmed milk for 30 min at room temperature. The membranes were incubated overnight at 4°C with specific primary antibodies. After washing three times with TBST, the membranes were incubated with secondary antibodies for 2 h at room temperature. Protein signals were acquired using a membrane imaging system (ChemiDoc™ XRS+, Bio-Rad). The following primary antibodies were used in this study: rabbit anti-α-SMA (Cat. no. CY5295, 1:1,000, Abways), rabbit anti-p-FOS (Cat. 5348T, 1:1,000, Cell Signaling Technology), rabbit anti-P-JUN (Cat. 3270T, 1:1,000, Cell Signaling Technology), rabbit anti-ATF3 (Cat. ab254268, 1:1,000, Abcam), mouse anti-c-FOS (Cat. sc-166940, 1:1,000, Santa Cruz Biotechnology), mouse anti-c-JUN (Cat. sc-74543, 1:1,000, Santa Cruz Biotechnology), mouse-anti-GAPDH (Cat. Ab8245, 1:1,000, Abcam). And the secondary antibodies were Goat anti-Rabbit IgG HRP (Cat. BE0101, 1:10,000, EASYBIO), Goat anti-Mouse IgG HRP (Cat. BE0102, 1:10,000, EASYBIO).

RT-qPCR

TRIzol™ reagent was used to extract total RNA from cultured cells according to the manufacturer’s instructions. Next, RNA was resuspended in DEPC-treated water and quantitative RT-PCR was performed using the HiScript® II One Step qRT-PCR SYBR Green Kit according to the product instructions. The Quantagene q225 qPCR System was used to analyze cDNA levels with specific primers and normalize the results to GAPDH. The primers used for RT-qPCR are designed by PrimerBank (https://pga.mgh.harvard.edu/primerbank/). The primers were validated by examining the primer efficiency and melt curve parameters. The primer efficiency assesses whether the detection of the fluorescent dye reflects this dilution and that the template DNA is truly doubling every cycle. And the primer efficiency falls in the range of 90-110%. Additionally, the melting curve is used to check whether primers are specific to a single gene of interest, and are dissociate at a single temperature. The data showed in this word was passed validation. All qPCR-related primers are shown in Table S1.

Usage of inhibitors

The inhibitors of ERK, JNK and p38 (listed in Key resources table) were diluted to 10μM as instructions and added to the cells on stiff matrix, after culturing for two days, the cells were collected for downstream analysis.

RNA-seq library preparation

Total RNA was extracted using the MolPure^® Cell RNA Kit (YEASEN, 19231ES50). RNA sequencing libraries were constructed by GENEWIZ, Inc. (Suzhou, China). RNA-seq paired-end reads were sequenced on the Illumina™ HiSeq XTen platform.

ATAC-seq library preparation

Samples were centrifuged for 5 min at 500 × g, 4°C and the cell pellet was resuspended in moderately cold lysis buffer. The TruePrep DNA Library Prep Kit V2 for Illumina^® (TD502) was used for library construction (Vazyme Biotech Co., Ltd) after 12 cycles of PCR amplification. After purification, paired-end sequencing was performed on the Illumina^® Novaseq 6000 platform.

Cut&Tag library preparation

The CUT&Tag assay was performed using the NovoNGS^® CUT&Tag 3.0 High-Sensitivity Kit (NovoProtein, N259-YH01). A total of 1 × 10⁵ cells were washed twice with 1.5 mL wash buffer and then mixed with activated concanavalin A beads. After successive incubation with the primary (room temperature, 2 h) and secondary (room temperature, 1.5 h) antibodies, cells were washed and incubated with pAG-Tn5 for 1.5 h. The following primary antibodies were used in this study: rabbit anti-H3K27ac (Cat. Ab4729, 1:50, Abcam), rabbit anti-H3K4me3 (Cat. Ab8580, 1:50, Abcam), mouse anti-H3K27me3 (Cat. Ab6002, 1:50, Abcam), rabbit anti-P-FOS (Cat. 5348T, 1:50, Cell Signaling Technology), rabbit anti-P-JUN (Cat. 3270T, 1:50, Cell Signaling Technology), rabbit anti-ATF3 (Cat. ab254268, 1:50, Abcam). And the secondary antibodies were Goat anti-Rabbit IgG HRP (Cat. BE0101, 1:100, EASYBIO), Goat anti-Mouse IgG HRP (Cat. BE0102, 1:100, EASYBIO). Subsequently, tagmentation was performed for 1 h in the provided buffer, and the target DNA fragments were purified using tagmentation DNA extraction beads. After washing the beads in 80% ethanol, the libraries were sequenced using the Illumina^®NovaSeq 6000 platform.

RNA-seq data analysis

The raw sequences were cleaned using TrimGalore (https://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) and mapped to the human reference genome (hg19) by STAR (v2.7.1a) with default parameters. All mapped BAM files were converted to bigwig using bedtools (v2.24.0) (Quinlan, 2014) for visualization in IGV. High-quality mapped reads were quantitated using htseq-count (v0.11.3) (Anders et al., 2015). Differentially expressed genes were analyzed by DEseq2 (Love et al., 2014). Functional enrichment of previously reported gene sets in the transcriptomes was determined using the GSEA software package (Mootha et al., 2003; Subramanian et al., 2005), and GO enrichment analysis was performed using DAVID (Sherman et al., 2022). The gene signatures used in RNA-seq analysis have been reported previously (De Smet et al., 2021). The active fibrogenic HSC signature was based on the CCL₄ model of liver fibrosis, in which differentially expressed genes in HSCs isolated from CCL₄-treated mice were compared with HSCs isolated from both healthy and recovered mice. This signature constitutes genes directly associated with ECM deposition and fibrotic HSCs. The conserved initiatory HSC signature was generated by intersecting “in vivo” and “in vitro” genes. The in vivo liver injury signature was defined as up- and downregulated genes in HSCs at 24 h after a single injection of CCL₄ as compared with HSCs isolated from healthy mice.

ATAC-seq data analysis

Sequencing adaptors were removed from the raw ATAC-seq reads using TrimGalore (https://www.bioinformatics.babraham.ac.uk/projects/trim_galore/), and clean data were mapped to the human reference genome (hg19) using Bowtie2 (v.2.4.1) (Langmead and Salzberg, 2012). PCR duplicates were removed using Picard (https://broadinstitute.github.io/picard/). Peaks were called using Macs2 (Zhang et al., 2008) with a relaxed q-value threshold of 0.05. The HOMER motif discovery algorithm findMotifsGenome.pl was used to elucidate the enriched motifs of specific accessible regions (Heinz et al., 2010).

Cut&Tag data analysis

TrimGalore (https://www.bioinformatics.babraham.ac.uk/projects/trim_galore/) was used to cut adapters, and trimmed reads were then aligned to the human genome (hg19) using Bowtie2 as described in the ATAC-seq data analysis section. Reads were sorted and converted to BAM format, and duplicates were marked using Picard (https://broadinstitute.github.io/picard/). Chromatin markers were used to train the chromatin states in mouse HSCs by chromHMM (Ernst and Kellis, 2012). For comparison of the ChIP-seq and ENCODE data, H3K4me1, H3K27ac, and CTCF ChIP-seq data generated in MEFs were downloaded from https://www.encodeproject.org/. After normalization of the sequencing depth, the deepTools2 (v 3.3.1) software with default parameters (Ramirez et al., 2016) was used to plot the heatmaps showing signals around peak regions.

Identification of unique and shared differentially accessible regions

The read pair numbers inside each ATAC-seq peak were calculated using DiffBind (bioconductor.org/packages/release/bioc/html/DiffBind.html). The differentially accessible regions were identified using DEseq2 (Love et al., 2014) with a cutoff of Fold Change ≥ 2 and p-value < 0.05 through pairwise spatiotemporal comparisons (D2 soft vs. D2 stiff, D4 soft vs. D4 stiff, D2 soft vs. D4 soft, D4 stiff vs. D4 stiff). Four unique clusters and four shared clusters were selected from the unique and shared peaks; for example, D4 stiff unique peaks were selected from the differentially accessible regions with higher signals by intersecting D2 stiff vs. D4 stiff and D4 soft vs. D4 stiff.

Quantification and statistical analysis

The high-through sequencing experiments consisted of at least two biological replicates. And the biochemical assays consisted of at least three biological replicates. For comparison between conditions in this study, Wilcoxon tests were performed using the function in R. The statistical significance of qPCR was determined by One-way ANOVA. The data expressed as the mean ± standard deviation (SD) was representative of all independent experiments. P values of < 0.05 were considered significance.

Acknowledgments

W.Y., T.D. and Y.S. were supported by the National Key R&D Program of China (No. 2022YFA1303103) and the National Science Foundation of China (21825401). W.Z. and C.L. were supported by the National Natural Science Foundation of China (32288102, 32025006) and the National Key Research and Development Program of China (2021YFA1100300). Z.F. was supported by the S&T Program of Hebei No.22377703D. W.Q. was supported by Hebei Natural Science Foundation (Project H2021206314). Part of the data analysis was performed on the High-Performance Computing Platform of the Center for Life Sciences, Peking University.

Author contributions

Y.S. and C.L. supervised the study. Z.F. and W.Q. gave some of the experimental guidance and provided part of samples. W.Z. and W.Y. designed the experiments and completed the sequencing library construction. W.Y. and T.D. conducted all the imaging experiments, chemical assays and data analysis. W.Z. performed the bioinformatics analysis. W.Z, W.Y., and T.D. wrote the manuscript with input from all authors.

Declaration of interest

The authors declare no competing financial interests.

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Increased matrix stiffness promotes fibrogenesis of hepatic stellate cells through AP-1-induced chromatin priming

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Abstract

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Introduction

Results

Matrix stiffness induces shifts in the fibrotic phenotype and transcriptome

Increased matrix stiffness is accompanied by dynamic chromatin accessibility

Stiffness-induced dynamic chromatin accessibility precedes changes in gene expression

Primed chromatin accessibility remodeling is positively associated with activation of the H3K27ac modification

AP-1 factor motifs are enriched in primed chromatin regions in response to matrix stiffness

Phosphorylation of JUN promotes chromatin accessibility in response to matrix stiffness

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