Runx1 drives cardiac fibrosis and heart failure through epigenetic activation of myofibroblasts in pressure overload-induced cardiac remodeling

Abstract

Background

Heart failure (HF) is often accompanied by cardiac fibrosis, a pathological process driven by activated cardiac fibroblasts (CFs) transitioning to a myofibroblast phenotype. The Runt-related transcription factor 1 (Runx1) has been implicated in various fibrotic diseases, but its role in cardiac fibrosis and HF progression remains unclear. This study aimed to elucidate the role of Runx1 in CF activation, cardiac fibrosis, and HF development.

Methods

HF was induced in mice using transverse aortic constriction (TAC). Runx1 expression was assessed in failing hearts via Western blot and qPCR, with immunostaining to localize Runx1 in CFs. In vitro, CFs were treated with TGF-β, and Runx1 knockdown was achieved using siRNA or adenoviral-mediated deletion. Myofibroblast-specific Runx1 knockout mice (PostnCre, Runx1F/F) were used to investigate the in vivo effects of Runx1 on cardiac function, fibrosis, and hypertrophy post-TAC. Chromatin immunoprecipitation (ChIP) and luciferase assays were conducted to evaluate Runx1’s regulation of Postn transcription.

Results

TAC-induced HF was associated with significant upregulation of Runx1 protein and mRNA levels, particularly in CFs. In vitro, Runx1 knockdown suppressed TGF-β-induced markers of CF activation, including α-smooth muscle actin (α-SMA), periostin (Postn), and collagen type I (Col1a1), and reduced CF migration and proliferation. PostnCre-Runx1F/F mice exhibited improved cardiac function, reduced hypertrophy, and decreased fibrosis compared to control mice post-TAC. Mechanistically, Runx1 was found to bind the Postn promoter and recruit the transcriptional coactivator P300, enhancing histone acetylation and promoting Postn transcription.

Conclusions

Runx1 plays a pivotal role in cardiac fibroblast activation and fibrosis, likely through epigenetic regulation of Postn expression, thereby driving heart failure progression. Targeting Runx1 may represent a promising therapeutic strategy for heart failure.

Background

Heart failure is the leading cause of morbidity and mortality within the global cardiovascular disease population, characterized by the pathological hallmarks of progressive cardiomyocyte loss and widespread interstitial fibrosis[1, 2] As the ultimate common pathway in response to a range of pathological insults, heart failure involves complex interactions among multiple intracellular and extracellular signaling cascades[3–5]. A key pathological process in heart failure progression is ventricular remodeling, marked by the accumulation and deposition of extracellular matrix (ECM), particularly collagen types I and III. This process increases myocardial stiffness and worsens cardiac dysfunction[6, 7]. Cardiac myofibroblasts, which primarily originate from resident cardiac fibroblasts (CFs), are the main drivers of ECM deposition and scar formation in the injured heart[8]. Chronic injury triggers continuous activation of cardiac fibroblasts, causing them to transdifferentiate into highly proliferative and migratory myofibroblasts, leading to irreversible cardiac fibrosis and ultimately end-stage heart failure[9]. Consequently, there is growing interest in identifying new molecular targets for pathological cardiac remodeling and dysfunction.

Runt-related transcription factor 1 (Runx1) belongs to the Runx gene family (Runx1, Runx2, and Runx3), all of which encode DNA-binding α subunits that partner with core binding factor β to form heterodimeric transcription factors[10]. Runx proteins can either activate or repress gene transcription during development and in pathological conditions[11]. Previous studies have suggested that Runx1 is closely associated with tissue fibrosis. For instance, specific deletion of Runx1 in renal tubular epithelial cells (RTECs) was shown to protect against renal fibrosis induced by unilateral ureteral obstruction (UUO) or folic acid (FA) treatment[12]. Additionally, silencing Runx1 in human lung fibroblasts (HLFs) reduced fibroblast-to-myofibroblast differentiation, as evidenced by decreased expression of α-SMA and ECM proteins[13]. In the liver, Runx1 knockdown reduced the viability and migration of hepatic stellate cells and attenuated CCL4-induced liver fibrosis in mice[14]. These findings suggest that Runx1 plays a significant role in fibrosis across various organs.

However, the role of Runx1 in cardiac fibrosis remains poorly understood. While a few studies have shown that Runx1 inhibition can alleviate cardiac fibrosis and dysfunction following myocardial infarction, these investigations have primarily focused on the role of Runx1 in cardiomyocytes rather than in cardiac myofibroblasts or other cell types[15–17]. Interestingly, bioinformatics analysis of systemic sclerosis fibroblast transcriptomes has identified upstream transcription factors, including Fosl2, Runx1, and Stat1, as key drivers of myofibroblast activation in this condition[18]. Therefore, it is necessary to evaluate the roles and mechanisms of Runx1 in cardiac myofibroblasts.

In our study, we observed a significant upregulation of Runx1 in the hearts of mice with heart failure. Using a tamoxifen-inducible, myofibroblast-specific Runx1 deletion model, we found that deletion of Runx1 significantly reduced fibrotic response and heart failure induced by transverse aortic constriction (TAC). These findings suggest that myofibroblast-specific Runx1 plays a critical role in cardiac fibrosis pathogenesis in mice and may represent a promising therapeutic target for treating heart failure.

Methods

Animals, surgery, and histology

All animal experiments were conducted in strict accordance with the National Research Council’s Guide for the Care and Use of Laboratory Animals and were approved by the Ethical Committee of Jiangxi Medical College. To generate myofibroblast-specific Runx1 knockout (KO) mice, Runx1 flox/flox mice (JAX Strain #: 008772), in which exon 4 of the Runx1 gene is flanked by loxP sites, were crossed with Postn-Cre mice (JAX Stock #: 029645), which express Cre recombinase under control of the Postn promoter. The resulting Runx1 flox/flox; Postn-Cre mice (referred to as Runx1 cKO) enable fibroblast-specific deletion of Runx1. Genotyping was performed by PCR using the following primers: for Runx1, forward 5’-GCGTTCCAAGTCAGTTGTAAGCC-3’ and reverse 5’-CTGCATTTGTCCCTTGGTTGACG-3’; the PCR product is 550 bp for the floxed allele and 362 bp for the wild-type allele. An alternative primer set was also used for confirming Cre-mediated recombination: wild-type forward, 5’-GGTGCTTCTGTAAGGCCATC-3’; mutant forward, 5’-GGTGGGACATTTGAGTTGCT-3’; and common reverse, 5’-CCTTGCAATAAGTAAAACAGCTC-3’. The expected PCR products were ~ 270 bp for the mutant allele and 380 bp for the wild-type allele. Male KO and littermate control mice, aged eight weeks, were subjected to transverse aortic constriction (TAC) surgery to induce pressure overload, following previously established protocols[19]. Sham-operated animals underwent the same procedures except for the banding of the transverse aorta. Mice were fed a tamoxifen diet (TAM, Envigo, TD.130860) starting 48 h before surgery and continued the TAM diet until tissue harvesting.

Mice were anesthetized, and the left common carotid artery was severed for exsanguination. Hearts were collected, rinsed in PBS, fixed in 4% paraformaldehyde for 48 h, and subsequently paraffin embedded. Cross-sections of the heart, 5-µm thick, were cut and deparaffinized in xylene twice, followed by rehydration in 100%, 95%, 80%, and 70% ethanol. Masson’s trichrome, Sirius red, and hematoxylin-eosin (H&E) staining were performed according to the manufacturer’s instructions (Masson’s Trichrome Stain Kit [Solarbio, G1340], Sirius Red Stain Kit [Solarbio, S8060], and H&E buffer [Solarbio, G1120], respectively). Digital images of stained sections were captured using a high-resolution whole-slide scanner (Leica Biosystems).

Echocardiography

Transthoracic echocardiography was conducted under anesthesia with sodium pentobarbital, both prior to surgery and six weeks post-TAC, using a dynamically focused 15 MHz linear-array transducer (EnVisor M2540A; Philips Medical System, Best, The Netherlands) with a depth setting of 1.5 cm. M-mode tracings were obtained from the short-axis view at the level of the papillary muscle of the left ventricle (LV).

Isolation and culture of cardiac fibroblasts (CFs)

cardiac fibroblasts were isolated as previously described[20] After euthanizing mice by cervical dislocation, hearts were rapidly excised via sternotomy and transferred into a 15 mL sterile Falcon tube containing pre-warmed (37 °C) phosphate-buffered saline (PBS), then moved to a laminar flow hood where all subsequent steps were performed under sterile conditions. Residual aortic tissue was removed, and hearts were washed twice with 10 mL of 37 °C PBS. The right and left atria were separated from the ventricles and placed in Petri dishes containing PBS. Tissues were minced into ~ 1 mm² pieces, transferred into T-25 flasks, washed three times with 10 mL of PBS, and subjected to serial 10-minute enzymatic digestions with 10 mL of pre-warmed dissociation medium at 37 °C in a 5% CO₂ incubator with gentle agitation (75–80 rpm) on an orbital shaker. After each digestion, supernatants were collected, centrifuged at 1500 rpm (405 × g) for 5 min at room temperature, and the cell pellets resuspended in 1 mL of complete M199 medium (M199 supplemented with 10% FBS and 2% penicillin-streptomycin, with optional Amphotericin B). Digestions were repeated until tissues were fully dissociated; all suspensions were pooled, centrifuged again, and the final cell pellet was resuspended in 4 mL of complete medium. Atrial or ventricular cell suspensions were plated onto either 35 mm collagen-coated soft hydrogel-bound dishes to obtain quiescent fibroblasts or uncoated standard 35 mm plastic dishes to promote myofibroblast activation. Cells were incubated at 37 °C and 5% CO₂ for 150 min (pre-plating), then washed 3–4 times with warm medium to remove non-adherent cells, followed by addition of 2 mL fresh medium. After 20 h, adherent cells were washed twice, replenished with fresh medium, and cultured until reaching 80–90% confluence; these were defined as passage 0 (P0) cardiac fibroblasts.

Cardiomyocyte isolation

Adult cardiomyocytes were isolated using a Langendorff-free enzymatic digestion protocol as previously described[21]. Briefly, mice were anesthetized, and the chest cavity was opened to expose the heart. The descending aorta and inferior vena cava were transected, and 7 mL of ice-cold EDTA buffer was injected into the right ventricle to flush blood. The ascending aorta was clamped using Reynolds or Lahey forceps, and the heart was excised and transferred into a 60-mm dish containing EDTA buffer. Sequential enzymatic digestion was performed by injecting 10 mL of EDTA buffer, followed by 3 mL of perfusion buffer, and then 30–50 mL of collagenase buffer into the apex of the left ventricle (LV) until tissue visibly softened. The heart was dissected into its chambers, and tissues were minced into ~ 1 mm³ pieces and gently triturated to dissociate cells. Enzymatic digestion was halted with 5 mL stop buffer. The resulting cell suspension was filtered through a 100-µm mesh, and cardiomyocytes were enriched via four rounds of gravity sedimentation with gradual calcium reintroduction using stepwise increasing calcium concentrations in reintroduction buffers. Rod-shaped, calcium-tolerant cardiomyocytes were collected, quantified using a hemocytometer, and plated on laminin-coated plates or coverslips in prewarmed plating medium (37 °C, 5% CO₂). After one-hour, unattached cells were removed and fresh culture medium was added, followed by media replacement every 48 h. The supernatant fractions containing nonmyocytes were pooled, centrifuged (300 × g, 5 min), resuspended in fibroblast growth medium, and plated onto tissue-culture treated plastic surfaces (approx. 23 cm² per LV) for downstream analysis or culture.

Isolation and culture of cardiac microvascular endothelial cells (CMECs)

Adult murine hearts were harvested and perfused through the aorta with ice-cold phosphate-buffered saline (PBS) to remove blood, followed by excision of the atria[22]. The ventricles were finely minced and transferred into 1 mL of pre-warmed enzymatic digestion solution containing 0.25 mg/mL Liberase TL (Roche, Mannheim, Germany) and 20 U/mL DNase I (Sigma-Aldrich, Steinheim, Germany) in sterile-filtered Hank’s Balanced Salt Solution (HBSS; with Ca²⁺/Mg²⁺, Gibco/Invitrogen), supplemented with 10 mM HEPES. The tissue suspension was incubated at 37 °C for 20 min and centrifuged at 900 rpm. The supernatant containing dissociated cells was then transferred to 8 mL of ice-cold Iscove’s Modified Dulbecco’s Medium (IMDM) supplemented with 20% fetal calf serum (FCS; Gibco/Invitrogen). The remaining tissue was subjected to a second round of enzymatic digestion using fresh enzyme solution. Cell suspensions from both digestion steps were pooled, filtered twice through 40 μm nylon cell strainers (BD Biosciences, Heidelberg, Germany) to remove undigested debris, and centrifuged at 300 × g for 10 min at 4 °C. The resulting cell pellet was resuspended in MACS buffer (PBS containing 2 mM EDTA and 0.5% bovine serum albumin; all from Gibco/Invitrogen). For magnetic separation, cells (approximately 400,000–800,000 per adult heart) were incubated with anti-CD31 magnetic microbeads (1:10 dilution; Miltenyi Biotec, Bergisch Gladbach, Germany) for 15 min at 8 °C. After washing twice with MACS buffer, labeled cells were passed through MACS LS-columns (Miltenyi Biotec) to isolate CD31⁺ cardiac endothelial cells. Purified CMECs were then collected for downstream applications.

Cell culture and transwell assay

Primary CFs were treated with TGF-β (10 ng/mL) for 24–48 h. HEK293T cells (ATCC) were cultured in high-glucose DMEM (Sigma) supplemented with 10% fetal bovine serum. Cells were transfected with the Postn promoter plasmid, Runx1 overexpression plasmid, or siRNAs (Santa Cruz, sc-37678) targeting Runx1 using Lipofectamine 3000 (Sigma, L3000008). Cells were infected with GFP or Cre adenovirus (Biowit, China) for 24 h. Promoter activity was measured using a luciferase reporter assay system (Promega), and cell migration was assessed using a Transwell assay in a 24-well Boyden chamber with an 8-µm pore insert. Cardiac fibroblasts (CFs) were harvested, resuspended in serum-free medium, and adjusted to a concentration of 5 × 10⁵ cells/mL. A 100–200 µL volume of the cell suspension was added to the upper chamber of each Transwell insert, while the lower chamber was filled with 600 µL of medium containing a chemoattractant (typically with serum). The plate was incubated at 37 °C for 12 h. After incubation, non-migrated cells on the top side of the membrane were gently removed with a cotton swab. Migrated cells on the lower membrane surface were fixed with 4% paraformaldehyde and stained with crystal violet. The number of migrated cells was quantified under a microscope.

5-ethynyl-2’-deoxyuridine (EdU) fluorescence staining

The proliferation of primary CFs was assessed using the Cell-Light Apollo 567 EdU Stain Kit (Ribobio, C10310-1), following the manufacturer’s instructions. Primary CFs were incubated with EdU fluorescent dye for 24 h with simultaneous treatment with TGF-β. EdU fluorescence was visualized using confocal scanning microscopy (LEXT, OLS4500).

RNA isolation and real-time PCR

Total RNA was extracted from cells and tissues using TRIzol Reagent (Sigma, 10296010). Reverse transcription was conducted with Hiscript III Reverse Transcriptase (Vazyme, R30), following the manufacturer’s protocol. Real-time PCR was performed on the LightCycler^® 96 system. Primer sequences are listed in Supplementary Table S1.

Protein extraction, immunoprecipitation, and western blotting

Proteins were extracted from cells and tissues using RIPA buffer (50 mM Tris [pH 7.4], 150 mM NaCl, 1% Triton X-100, 1% sodium deoxycholate, 0.1% SDS) supplemented with a protease inhibitor (Beyotime, P1005). For immunoprecipitation, cell lysates were prepared with IP buffer (Beyotime, P0013) with added protease inhibitor, incubated with antibodies overnight, and bound to Protein A/G-plus magnetic beads. Protein complexes were eluted by boiling in IP buffer with 1X sample loading buffer. Western blots were performed with the following antibodies: anti-β-actin (CST, 3700 S), anti-Runx1 (Abcam, ab23980), anti-P300 (CST, 57625 S), anti-α-SMA (Abcam, ab5694), and anti-collagen type I (Rockland, I600-403-103).

Chromatin immunoprecipitation

Chromatin immunoprecipitation (ChIP) and re-ChIP were performed using a ChIP kit (CST, 9003 S) according to the manufacturer’s instructions. Briefly, chromatin was cross-linked with 1% paraformaldehyde, sonicated, and immunoprecipitated with antibodies against Runx1 (CST, 8529 S), P300 (CST, 57625 S), IgG (CST, 3900), acetylated H3 (CST, 8173), and acetylated H4 (CST, 13534) overnight. Protein A/G-plus magnetic beads were added to enrich the chromatin. DNA was isolated from the enriched chromatin for real-time PCR. For re-ChIP, eluted chromatin was stripped from magnetic beads using elution buffer (100 mM NaCO3, 1% SDS), diluted with ChIP buffer, and subjected to a second immunoprecipitation with a second antibody of interest. ChIP primer sequences are listed in Supplementary Table S1.

Statistical analysis

Data are presented as mean ± SD. Differences between two groups were analyzed by unpaired t-test with Welch’s correction for normally distributed data. For experiments with more than two groups and one independent variable, one-way ANOVA with Tukey’s multiple comparisons test was used. For experiments with more than one independent variable, two-way ANOVA with Tukey’s multiple comparisons test was applied. Statistical significance was defined as P < 0.05.

Results

Increased runx1 expression in failing mouse hearts

To investigate the potential role of Runx1 in heart failure, we evaluated its expression levels in failing hearts from transverse aortic constriction (TAC)-induced wild-type male mice at 6 weeks post-surgery. Compared to sham controls, Runx1 protein (Fig. 1A and B) and mRNA (Fig. 1C) levels were significantly upregulated in failing hearts. In parallel, we observed a marked increase in the expression of genes associated with cardiac hypertrophy and fibrosis (Fig. 1C), indicating a pathological remodeling response. Further analysis of isolated cell types—cardiac endothelial cells (ECs), cardiomyocytes (CMs), and cardiac fibroblasts (CFs)—from failing hearts revealed that Runx1 expression was predominantly induced in CFs, with minimal changes observed in CMs or ECs (Fig. 1D). Additionally, consistent with these in vivo findings, stimulation of CFs isolated from adult mouse hearts with Transforming Growth Factor-β (TGF-β) led to increased Runx1 protein (Fig. 1E) and mRNA levels (Fig. 1F). These results indicate that Runx1 expression is upregulated in the failing heart, particularly in CFs, suggesting a key role for Runx1 in CF-mediated processes that may contribute to the progression of heart failure.

Fig. 1.

Upregulation of Runx1 in Failing Mouse Hearts and TGF-β-Stimulated Cardiac Fibroblasts (CFs). A-B Western blot analysis and quantification of Runx1, Atrial natriuretic peptide (ANP) and B-type natriuretic peptide (BNP) protein levels in hearts from wild-type (WT) mice subjected to 6 weeks of transverse aortic constriction (TAC) or sham surgery (n = 5). Protein levels were normalized to β-actin. C mRNA levels of ANP, BNP, Col1a1, Col3a1, α-SMA, and Runx1 in hearts from WT mice with 6 weeks of TAC or sham surgery, normalized to 18 S (n = 5). D Comparison of Runx1 mRNA levels in endothelial cells (ECs), cardiac fibroblasts (CFs), and cardiomyocytes (CMs) isolated from WT mice subjected to 6 weeks of TAC or sham surgery (n = 5), normalized to 18 S. E-F Runx1 expression measured by Western blot and qPCR in primary CFs isolated from adult WT mice and stimulated with TGF-β (10 ng/ml) for 24 h. All statistical analyses were performed using an unpaired t-test with Welch’s correction. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. “n” refers to independent biological replicates

Runx1 knockdown alleviates cardiac fibroblast activation in vitro

To investigate the role of Runx1 in cardiac fibroblast (CF) activation, we examined how Runx1 knockdown affects CF function in response to pro-fibrotic stimulation. Based on previous findings indicating that Runx1 expression is upregulated by pro-fibrotic signals in CFs, we hypothesized that reducing Runx1 expression would mitigate CF activation. We silenced Runx1 in CFs using a combination of three siRNAs, which effectively suppressed TGF-β-induced increases in mRNA levels of pro-fibrotic genes such as α-smooth muscle actin (α-SMA), periostin (Postn), and collagen type I (Col1a1) (Fig. 2A). Western blot analysis confirmed that Runx1 knockdown markedly reduced the TGF-β-induced expression of α-SMA and Col1a1 at the protein level (Fig. 2B). Since activated CFs exhibit increased migration and proliferation, we next assessed these functions. Runx1 knockdown significantly inhibited TGF-β-induced migration and proliferation of CFs (Fig. 2C–D). To further validate these findings, we employed an alternative approach using adenoviral Runx1 deletion. Primary CFs were isolated from Runx1^f/f mice and infected with adenovirus expressing Cre recombinase to specifically delete Runx1. As expected, CFs infected with Cre adenovirus showed a significant reduction in Runx1 protein levels compared to CFs infected with GFP adenovirus (Fig. 2E). Consistently, Runx1 deletion significantly attenuated TGF-β-induced CF activation (Fig. 2E–F), proliferation (Fig. 2G), and migration (Fig. 2H). These results indicate that Runx1 knockdown or deletion can effectively reduce CF activation and limit their pro-fibrotic responses to TGF-β stimulation, suggesting that Runx1 plays a critical role in CF activation and fibrotic processes.

Fig. 2.

Inhibition of Runx1 Expression Impairs Activation and Function of Primary CFs In Vitro. Primary CFs were isolated from adult WT mice, transfected with siRNA targeting Runx1 or scrambled RNA (SCR), and treated with TGF-β (10 ng/ml) or vehicle for 24 h. A-B Expression levels of Runx1 and pro-fibrogenic genes were assessed by qPCR and Western blot analysis. C-D The migratory and proliferative capacities of primary CFs were evaluated using transwell assays and EdU fluorescence staining, respectively. Additionally, primary CFs were isolated from adult Runx1^F/F mice, infected with adenovirus carrying GFP or Cre, and treated with TGF-β (10 ng/ml) or vehicle for 24 h. E-F Runx1 and pro-fibrogenic gene expression was measured by qPCR and Western blot. G-H The migration and proliferation of primary CFs were assessed as previously described. Statistical analyses were conducted using a Two-way ANOVA with Tukey’s multiple comparisons test. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

Runx1 loss in cardiac myofibroblasts reduces the fibrotic response to TAC

To investigate whether Runx1 deletion in cardiac myofibroblasts protects the heart from fibrosis in vivo, we generated a myofibroblast-specific Runx1 knockout (KO) mouse model using a Cre/Loxp system. Runx1^f/f mice were crossed with Postn-Cre mice to produce Runx1 KO mice (Postn-Cre, Runx1^f/f). KO and control mice, which lacked Cre recombinase activity, were subjected to transverse aortic constriction (TAC) surgery at 8 weeks of age and fed tamoxifen-containing food to induce Runx1 deletion until tissue collection post-TAC (Fig. 3A). We confirmed the significant down regulation of RUNX1 expression in isolated fibroblasts (Fig. 3B). Echocardiographic analysis showed that KO mice exhibited significantly improved cardiac function compared to control mice following TAC. Specifically, KO mice had a reduced interventricular septum diameter in diastole (IVS-d) and higher ejection fraction (EF) and fractional shortening (FS) values, indicating improved cardiac performance (Fig. 3C). Additionally, Runx1 deletion attenuated TAC-induced cardiac remodeling, as evidenced by a smaller increase in heart weight to body weight (HW/BW) and heart weight to hindlimb length (HW/HL) ratios in KO mice compared to controls (Fig. 3D-E). Histological analyses, including Sirius Red and Masson’s Trichrome staining, revealed a marked reduction in collagen deposition and fibrosis in KO mice (Fig. 3F-G). The expression levels of pro-fibrotic genes, including α-SMA, Postn, Col1a1, and Col3a1, were significantly lower in KO mice compared to control mice after TAC, indicating reduced fibrotic signaling (Fig. 3H). Importantly, KO mice showed no significant differences from control mice under baseline conditions, suggesting that Runx1 deletion does not affect heart structure or function in the absence of stress. Overall, these findings suggest that myofibroblast-specific Runx1 knockout mitigates the fibrotic response, helping to prevent cardiac remodeling and dysfunction in response to pressure overload-induced heart failure.

Fig. 3.

Myofibroblast-Specific Deletion of Runx1 Attenuates TAC-Induced Cardiac Remodeling and Dysfunction. Male Postn^Cre, Runx1^F/F mice (KO mice) and Runx1^F/F control mice (WT mice) underwent TAC surgery or sham procedure at 8 weeks of age, with tamoxifen (TAM) diet initiated 48 h prior to surgery and continued until tissue collection (n = 5–8 per group). A Schematic overview of the experimental design. B Runx1 gene expression was measured by qPCR. C Echocardiographic images comparing KO and WT mice following TAC surgery or sham procedure. Assessment of cardiac function through measurement of interventricular septum diameter in diastole, ejection fraction, and fractional shortening. D Photographs of global heart morphology. E Heart weight-to-body weight ratio (HW/BW) and heart weight-to-tibial length ratio (HW/TL) were calculated. F, G Fibrotic areas in heart sections visualized by Sirius Red and Masson’s Trichrome staining. H Expression of pro-fibrogenic genes (α-SMA, Postn, Col1a1, Col3a1) was quantified by qPCR. Statistical analyses were performed using a Two-way ANOVA with Tukey’s multiple comparisons test. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. “n” refers to independent biological replicates

Runx1 regulates postn transcription during cardiac fibroblast activation

Previous studies have suggested that Runx1 acts as an upstream transcription factor driving myofibroblast activation in conditions such as systemic sclerosis by influencing fibroblast transcriptome[18], we observed that among the pro-fibrogenic genes affected by Runx1 knockdown, Postn was particularly responsive, suggesting that Runx1 might directly regulate Postn expression in cardiac fibroblasts (CFs). To explore this hypothesis, we analyzed the potential of Runx1 as a transcription factor for Postn using the JASPAR database. Our results revealed that the Postn promoter region (−2000/+54) contains a Runx1-binding motif (−622/−612), suggesting a possible regulatory interaction (Fig. 4A). To confirm whether Runx1 could directly activate Postn transcription, we constructed a series of truncated Postn promoter-luciferase reporter plasmids. Our findings demonstrated that the full-length promoter (−2000/+54) showed strong activation by Runx1, whereas the shortest promoter fragment (−500/+54) lost responsiveness to Runx1 overexpression, supporting the importance of the Runx1-binding motif within this region (Fig. 4B). Chromatin immunoprecipitation (ChIP) assays further validated this regulatory mechanism. TGF-β stimulation enhanced Runx1 binding to the Postn promoter, with maximal enrichment observed at 24 h post-treatment, while no such binding was detected at the Gapdh promoter (Fig. 4C). Runx1 silencing significantly reduced this binding (Fig. 4D). We repeated the ChIP assay using adenoviral-mediated Runx1 depletion, which similarly demonstrated a marked decrease in Runx1 binding to the Postn promoter (Fig. 4E). Collectively, these findings indicate that Runx1 directly activates Postn transcription during cardiac fibroblast activation, positioning Runx1 as a critical regulator of pro-fibrotic gene expression in this context.

Fig. 4.

Runx1 Directly Regulates Postn Transactivation in Activated CFs. A Prediction of Runx1 binding motifs on the Postn promoter using JASPAR database. B Luciferase reporter assays in 293 T cells transfected with varying lengths of the Postn promoter-luciferase plasmid alongside Runx1 plasmid. Luciferase activity was normalized to protein concentration and GFP fluorescence. C Primary CFs treated with or without TGF-β (10 ng/ml) for 24–48 h underwent ChIP analysis with anti-Runx1 or anti-IgG. D Primary CFs transfected with siRNA targeting Runx1 or SCR were treated with TGF-β (10 ng/ml) or vehicle for 24 h, followed by ChIP analysis. E Primary CFs isolated from adult Runx1^F/F mice were infected with adenovirus carrying GFP or Cre, followed by TGF-β (10 ng/ml) or vehicle treatment for 24 h. ChIP analysis was performed with anti-Runx1 or anti-IgG. Statistical analyses were conducted using one-way ANOVA with Tukey’s multiple comparisons test for B-C and Two-way ANOVA for D-E. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

Runx1 interacts with P300 to epigenetically regulate postn transcription

To investigate the mechanism by which Runx1 regulates Postn transcriptional activation, we utilized the STRING database (Search Tool for the Retrieval of Interacting Genes/Proteins) to identify potential Runx1-interacting proteins. By inputting ‘Runx1’ into the STRING database (https://string-db.org/), we obtained a protein–protein interaction (PPI) network based on evidence from experimental data, curated databases, text mining, co-expression, and predictive computational methods. The interaction panel provided a visual network of known and predicted Runx1-associated proteins, highlighting key candidates potentially involved in the transcriptional regulation of Postn (Fig. 5A). Since we aimed to explore a mechanism involving positive epigenetic regulation, we selected P300, a known transcriptional coactivator. Previous studies have shown that Runx1 can interact with coactivators like P300 to enhance gene transcription[23, 24]. We hypothesized that Runx1 recruits P300 to facilitate Postn transcription during cardiac fibroblast (CF) activation. Co-immunoprecipitation assays confirmed a strong interaction between Runx1, and P300 in CFs stimulated with TGF-β (Fig. 5B). To examine whether this interaction occurs at the Postn promoter, we performed a re-ChIP assay, which revealed that TGF-β treatment significantly increased the recruitment of the Runx1-P300 complex to the Postn promoter, but not to the Gapdh promoter, in CFs (Fig. 5C). Further analysis showed that Runx1 deletion via adenoviral infection significantly reduced P300 occupancy on the Postn promoter, as detected by ChIP (Fig. 5D). Similarly, P300 knockdown by siRNA notably decreased Runx1 recruitment to the Postn promoter (Fig. 5E). Given P300’s role as a histone acetyltransferase (HAT), we examined whether it affects histone modifications on the Postn promoter. ChIP assays demonstrated that TGF-β stimulation enhanced the enrichment of active histone marks, specifically acetylated H3 and H4, on the Postn promoter. Silencing P300 abolished these histone modifications, indicating P300’s essential role in epigenetic activation of Postn (Fig. 5F). In summary, these findings suggest that Runx1 recruits P300 to the Postn promoter, where it enhances Postn transcription through epigenetic modifications, highlighting a collaborative mechanism in CF activation.

Fig. 5.

Runx1 Recruits P300 to Regulate Postn Transcription. A The protein-protein interaction (PPI) network constructed using the STRING database. B Immunoprecipitation experiments performed with anti-Runx1, anti-P300, or anti-IgG in primary CFs treated with TGF-β (10 ng/ml) for 24 h. C ChIP assays conducted on primary CFs treated with or without TGF-β (10 ng/ml) for 24 h, using anti-Runx1, anti-P300, or anti-IgG. D Primary CFs isolated from adult Runx1^F/F mice were infected with adenovirus carrying GFP or Cre, followed by treatment with TGF-β (10 ng/ml) or vehicle for 24 h, with ChIP analysis performed using anti-P300 or anti-IgG. E Primary CFs were transfected with siRNA targeting P300 or SCR, followed by TGF-β (10 ng/ml) or vehicle treatment for 24 h, with ChIP analysis using anti-Runx1 or anti-IgG. F Primary CFs treated with or without TGF-β (10 ng/ml) for 24 h underwent ChIP analysis with anti-AcH3, anti-AcH4, or anti-IgG. Statistical analyses were performed using unpaired t-test with Welch’s correction for B, and Two-way ANOVA with Tukey’s multiple comparisons test for C-E. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

Discussion

Cardiac fibrosis is a critical process linking cardiac remodeling to heart failure. The activation of cardiac fibroblasts is recognized as the primary contributor to heart fibrosis[25]. While effectively targeting the damaging activation of cardiac fibroblasts may limit fibrosis, specific clinical treatments for cardiac fibrosis remain inadequate[26]. In the current study, we present the first report on the expression and function of Runx1 in cardiac fibroblasts. Importantly, our findings reveal that myofibroblast-specific deletion of Runx1 regulates cardiac remodeling by limiting cardiac fibroblast activation. Notably, the deletion of Runx1 in cardiac fibroblasts mitigates remodeling and heart failure induced by transverse aortic constriction (TAC). Mechanistically, we uncover a novel interaction between Runx1 and P300, which appears to contribute to the transactivation of the Postn gene in cardiac fibroblasts (CFs). This research highlights the potential therapeutic implications of targeting Runx1 in cardiac remodeling and heart failure.

Previous research has primarily focused on the effects of Runx1 in acute leukemia and cancer[27–29]. However, the role of Runx1 in the pathological processes of non-cancerous conditions, such as tissue fibrosis, has recently gained attention. For instance, Woosook Kim et al. demonstrated that Runx1 is a key transcription factor in the proliferation and differentiation of mesenchymal stem cells into myofibroblasts[30]. Increased expression of Runx1 has been shown to promote processes involved in lung fibrosis, including fibroblast activation and transdifferentiation to the myofibroblast phenotype, while Runx1 deletion significantly impairs lung fibrosis progression[13]. Other studies have indicated that master regulators, including Runx1 and CREB3L1, activate hepatic stellate cells (HSCs) during fibrogenesis, as evidenced by public single-cell data from fibrotic and cirrhotic human livers[31], which corroborates Tracy et al.‘s findings[18]. In contrast, little is known about the role of Runx1 in the myofibroblasts of cardiac fibrosis, despite Runx1 being identified as a key regulator of adverse cardiac remodeling [15, 32–34]. In our study, we observed that Runx1 expression is increased in TAC-induced failing mouse hearts, suggesting its involvement in heart failure and the advanced stages of cardiac fibrosis. In vitro, Runx1 knockdown alleviated cardiac fibroblast activation across various biochemical experiments. In vivo, loss of Runx1 in cardiac myofibroblasts reduced the fibrotic response to TAC. These findings strongly indicate that Runx1 plays a critical role in regulating the trans differentiation of cardiac fibroblasts to myofibroblasts. Integrating our findings with previous studies, we propose that Runx1 may serve as a core transcription factor in fibroblast activation. Its regulatory role in the progression of cardiac fibroblast trans differentiation to myofibroblasts may extend beyond the heart, potentially influencing tissue fibrosis across various biological contexts. Excitingly, Runx1 may also emerge as a novel fibrotic marker and a promising target for new therapeutic strategies aimed at mitigating tissue fibrosis.

As a transcriptional coactivator with intrinsic acetyltransferase activity, P300 regulates the expression of numerous genes. Recent evidence links P300 closely to fibrotic responses. Ghosh et al. found that ectopic P300 expression accelerates Smad-mediated collagen gene expression in normal skin and lung fibroblasts[35]. Conversely, depletion of P300 using ribozymes abrogates TGF-β-induced collagen synthesis in normal dermal fibroblasts[36]. Furthermore, disrupting P300 levels or activity has been shown to attenuate the differentiation of activated HSCs into myofibroblasts in the liver[37]. Importantly, treatment with the P300 inhibitor L002 has been demonstrated to prevent Ang II-induced cardiac hypertrophy and reverse myofibroblast activation in murine ventricles[38]. These data indicate that, akin to Runx1, P300 plays a pivotal role in the activation of fibroblasts. Given that previous research suggests an interaction between P300 and Runx1 through immunoprecipitation and immunofluorescence assays[23, 24, 39], we aimed to investigate whether Runx1 could interact with P300 in TGF-β-stimulated cardiac fibroblasts.

In this study, we uncover a novel epigenetic mechanism by which Runx1 regulates cardiac fibrosis through its interaction with the histone acetyltransferase P300 to drive Postn (periostin) transcription in activated cardiac fibroblasts (CFs). This finding not only identifies Runx1 as a central transcriptional regulator of fibrogenesis but also highlights a previously unrecognized chromatin-based regulatory axis involving P300-mediated histone acetylation at the Postn promoter. P300 is a well-characterized histone acetyltransferase that promotes transcription by loosening chromatin structure through acetylation of histones H3 and H4, thus facilitating gene expression[40]. Our data demonstrate that Runx1 recruits P300 to the Postn promoter, enhancing local histone acetylation and chromatin accessibility, thereby initiating periostin transcription. This mechanism aligns with epigenetic regulatory patterns observed in fibrosis of other organs such as the liver and lungs, where P300 similarly drives pro-fibrotic gene programs[37, 41–43]. The Runx1-P300 interaction presents a compelling therapeutic target: small-molecule P300 inhibitors such as A-485, L002, and C646 have been shown to suppress fibrosis-related gene expression and attenuate pathological cardiac remodeling in preclinical studies[38, 44–47]. Given that Postn encodes periostin—a matricellular protein integral to ECM remodeling, collagen crosslinking, fibroblast recruitment, and myofibroblast persistence—its upregulation is a hallmark of fibrotic and failing hearts[48, 49]. Genetic ablation of Postn in mice confers resistance to pressure overload-induced fibrosis and preserves cardiac function[50, 51]. Therefore, targeting the Runx1-P300-Postn axis may offer a more specific anti-fibrotic strategy than broadly inhibiting upstream pathways like TGF-β, which are pleiotropic and essential for tissue homeostasis. Future approaches could include development of molecules or peptides to disrupt the Runx1-P300 interaction, neutralizing antibodies against periostin, or gene therapy using fibroblast-specific promoters (e.g., Postn promoter) to express dominant-negative Runx1. These strategies hold promise to mitigate fibrotic remodeling and progression to heart failure. Furthermore, the role of Runx1 may be conserved across organs affected by fibrotic diseases such as systemic sclerosis, liver cirrhosis, and pulmonary fibrosis, emphasizing its potential as a universal fibrotic regulator. Integrating genome-wide chromatin profiling techniques such as ATAC-seq or ChIP-seq will be instrumental in mapping the broader epigenomic landscape controlled by Runx1-P300 complexes and may reveal additional fibrogenic targets for intervention.

Several limitations of this study warrant consideration. First, our investigation focused exclusively on the ablation of Runx1 in myofibroblasts and its effect on attenuating TAC-induced cardiac fibrosis, without assessing the potential impact of Runx1 deletion in cardiomyocytes. Second, although we demonstrated that Postn is a downstream target of RUNX1 and provided in vitro evidence that exogenous POSTN partially rescues fibroblast activation following RUNX1 knockdown, these results remain limited and should be interpreted with caution. More comprehensive in vitro and in vivo studies are required to firmly establish the extent to which RUNX1-mediated regulation of POSTN controls fibroblast activation. Third, it is likely that RUNX1 regulates a broader pro-fibrotic transcriptional program beyond Postn. We did not explore other potential RUNX1 target genes, and future transcriptomic or epigenomic profiling would help to comprehensively define RUNX1-dependent gene networks in cardiac fibroblasts. Fourth, while we observed increased Runx1 expression in fibroblasts following pressure overload, the upstream signaling mechanisms responsible for this induction remain unclear. Pathways such as TGF-β, NF-κB, or MAPK signaling may be involved and warrant further investigation. Fifth, we did not investigate the interaction between RUNX1 and P300 at the physiological level or consider whether P300 silencing could modulate myofibroblast function. Sixth, although our study emphasized the RUNX1-P300 complex, other epigenetic cofactors may participate in regulating Postn expression and the fibrotic response. Lastly, the clinical relevance of our findings in the context of human heart failure remains to be determined. Addressing these limitations in future studies will be essential for advancing our understanding of RUNX1-mediated fibrotic remodeling.

Conclusions

This study was designed to investigate the functional role of Runx1 in cardiac fibroblast activation and its contribution to cardiac fibrosis and heart failure. Using both in vitro and in vivo models, we demonstrated that Runx1 is significantly upregulated in activated cardiac fibroblasts and that its deletion in myofibroblasts markedly reduces fibrosis and improves cardiac function in a pressure overload model. Mechanistically, we uncovered that Runx1 may drives fibroblast activation by epigenetically promoting Postn transcription via recruitment of the transcriptional coactivator P300. Our findings reveal a novel regulatory axis—Runx1/P300/Postn—that contributes to pathological cardiac remodeling. This suggests that targeting Runx1 in cardiac fibroblasts may represent a promising therapeutic strategy for mitigating cardiac fibrosis and heart failure. Further research should investigate the translational potential of Runx1 inhibition in clinical settings, explore the involvement of additional epigenetic co-regulators in the Runx1 complex, and assess whether similar mechanisms operate in human fibrotic heart disease. Furthermore, determining the cell-type specificity and safety of targeting Runx1 systemically will be crucial for therapeutic development.

Authors’ contributions

JA, WZ, LX, and QW contributed equally to the conception and design of the study, performed experiments, collected data, and drafted the initial manuscript. XP and ZZ participated in methodology development, data acquisition, and analysis. XW and YZ contributed to data interpretation and visualization. JH and LG were responsible for validation and assisted in data analysis. MW contributed to software and statistical analysis. JZ supervised the project, provided critical revisions to the manuscript for important intellectual content, and secured funding.

Funding

This study was supported by the Natural Science Foundation of Jiangxi Province (GrantNo.20204BCJ23017), National Natural Science Foundation of China (Grant No.81960061), Science and technology innovation base plan of Jiangxi Province (20212BCD42006) and Science Foundation of Traditional Chinese Medicine of Jiangxi Provincial Health Commission (Grant No.2022B981).

Data availability

The datasets used and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Declarations

Ethics approval and consent to participate

All methods are reported in accordance with ARRIVE guidelines. The animal study protocol (2021-12-002) was approved by the Institutional Review Board of the Animal Care and Use Committee of First Affiliated Hospital of Nanchang University, Jiangxi, China.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.

All methods are reported in accordance with ARRIVE guidelines. The animal study protocol (2021-12-002) was approved by the Institutional Review Board of the Animal Care and Use Committee of First Affiliated Hospital of Nanchang University, Jiangxi, China. Clinical trial number: not applicable.