(Pro)renin receptor promotes cardiomyocyte senescence via tripartite motif-containing 24-mediated stabilization of p53 in diabetic cardiomyopathy

Abstract

Diabetic cardiomyopathy (DCM) is a major complication of diabetes and a leading contributor to heart failure, in which cardiomyocyte senescence plays an increasingly recognized role. However, the underlying mechanisms driving this process remain poorly defined. Here, we identify the (pro)renin receptor (PRR) as a critical mediator of cardiomyocyte senescence in DCM. Using a high-fat diet and streptozotocin (STZ)-induced DCM mouse model, as well as primary cardiomyocytes exposed to high glucose and palmitic acid, we demonstrate that PRR expression is significantly upregulated in diabetic hearts and closely associated with key senescence markers, including SA-β-gal, γ-H2AX, p16, and p21. PRR overexpression exacerbates these senescence phenotypes and promotes the secretion of profibrotic senescence-associated secretory phenotype factors, contributing to increased myocardial fibrosis and cardiac dysfunction. Mechanistically, PRR stabilizes the p53 protein by inhibiting tripartite motif-containing 24 (TRIM24)-mediated ubiquitination and proteasomal degradation, thereby activating the p53-p21 axis. These findings reveal a novel role of the PRR in diabetic myocardial senescence and provide potential therapeutic targets for attenuating DCM progression.

Introduction

Diabetic cardiomyopathy (DCM) is a major cardiac complication of diabetes characterized by diastolic dysfunction and progressive systolic impairment, contributing to the high incidence of heart failure and adverse clinical outcomes in patients [1]. The pathogenesis of DCM is multifactorial and involves metabolic dysregulation, oxidative stress, and chronic inflammation [ 2, 3] . In recent years, cellular senescence has emerged as a pivotal contributor to the advancement of DCM pathology [ 4, 5] . This process is characterized by a spectrum of molecular changes, such as DNA damage accumulation, the induction of cell cycle inhibitors such as p16 and p21, and the secretion of a broad array of cytokines, chemokines, growth factors, and proteases, collectively known as the senescence-associated secretory phenotype (SASP) [ 6, 7] . These alterations can be initiated not only by physiological aging but also by pathological stimuli, including redox imbalance and metabolic stress [8]. Cardiomyocytes are particularly vulnerable to senescence, with senescent cells exhibiting widespread transcriptomic reprogramming [9]. Furthermore, increased senescent cell burden within cardiac tissue has been closely linked to progressive deterioration in heart function [ 4, 10] . The pharmacological elimination of senescent cells through senolytic compounds such as dasatinib and quercetin has demonstrated beneficial effects on cardiac structure and performance in experimental DCM models [11]. Although accumulating studies support the involvement of cellular senescence in the pathophysiology of DCM, the molecular mechanisms underlying cardiomyocyte senescence remain poorly understood.

The (pro)renin receptor (PRR) was initially identified as a component of the renin-angiotensin system (RAS), facilitating the generation of angiotensin I through its interaction with renin and prorenin, thereby contributing to blood pressure regulation and fluid balance [12]. In addition to this classical RAS-dependent role, PRR also activates angiotensin-independent pathways, including the MAPK and Wnt/β-catenin signaling pathways, which contribute to cardiovascular inflammation and fibrotic remodeling [13]. These angiotensin-independent roles of PRR are now recognized as critical contributors to the development of hypertension, adverse cardiac remodeling, and heart failure. Moreover, PRR serves as an accessory subunit of the vacuolar H⁺-ATPase (V-ATPase), playing a critical role in lysosomal acidification and intracellular pH regulation, processes essential for maintaining cellular homeostasis and cardiovascular integrity [ 14, 15] . Recent research has increasingly linked PRR to the modulation of cellular senescence. In skeletal muscle cells, PRR has been reported to promote senescence through activation of the Wnt/YAP axis  [16]. Upregulation of PRR expression has also been observed in the thoracic aorta and renal cortex of aged mice [ 17, 18] , and elevated levels of circulating soluble PRR (sPRR) have been detected in elderly individuals with heart failure [ 19, 20] . These findings collectively point to a possible regulatory role for PRR in senescence processes; however, its involvement in driving cardiomyocyte senescence under diabetic conditions remains insufficiently characterized.

p53 functions as a central transcriptional regulator of cellular senescence, responding to various stressors, such as redox imbalance, DNA damage, and metabolic disturbances [ 21, 22] . Upon activation, p53 induces the expression of downstream effector genes such as p21, thereby promoting cell senescence. Elevated p53 expression has been observed in diabetic myocardial tissues and is linked to increased expression of markers of cellular senescence [5]. The intracellular stability of p53 is tightly controlled by the ubiquitin-proteasome system, which governs its degradation through various E3 ubiquitin ligases [23]. Among these proteins, the tripartite motif-containing (TRIM) protein family has drawn increasing attention, with TRIM24 in particular being reported to facilitate p53 ubiquitination in cancer cells [ 24, 25] . However, the involvement of TRIM24 and p53 in DCM and their potential contributions to cardiomyocyte senescence have yet to be elucidated.

In this study, we sought to elucidate the contribution of PRR to cardiomyocyte senescence in DCM and to elucidate its underlying molecular mechanisms. Our results revealed a significant increase in PRR expression within the myocardium of DCM mice. Both in vivo and in vitro, PRR overexpression aggravated cardiomyocyte senescence and enhanced the secretion of profibrotic SASP components. Mechanistically, PRR promotes the stabilization of the p53 protein by inhibiting its TRIM24-mediated ubiquitination and proteasomal degradation, thereby facilitating p53-dependent transcriptional activation and upregulation of p21.

Materials and Methods

Animals and treatment

Male C57BL/6J mice (6‒8 weeks old, 20‒25 g) were obtained from Beijing Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China). Animals were maintained under specific pathogen-free conditions in a temperature- and light-controlled facility with unrestricted access to standard chow and water. Following a one-week acclimatization period, the mice were randomly assigned to four experimental groups ( n = 10): a normal diet (ND) control group, a DCM group induced with a high-fat diet combined with streptozotocin (HFD/STZ), a DCM group receiving control adenovirus expressing EGFP (Ad-EGFP), and a DCM group treated with adenovirus overexpressing PRR (Ad-PRR). The adenovirus was constructed by GeneChem Co., Ltd. (Shanghai, China). To establish the DCM model, the mice were given a high-fat diet providing 60% total caloric intake from fat for 8 weeks, followed by intraperitoneal administration of STZ (Sigma-Aldrich, St Louis, USA; 50 mg/kg/day, freshly prepared in 0.1 M citrate buffer) for five consecutive days. The control animals were maintained on a normal chow diet and received equivalent volumes of citrate buffer. Blood glucose levels were monitored one week after the final STZ injection, and the mice were classified as diabetic if their fasting glucose level exceeded 16.7 mM on three consecutive days. After 3 months of sustained hyperglycemia, cardiac function was assessed via transthoracic echocardiography to confirm the establishment of DCM. DCM mice were then injected with either Ad-PRR or Ad-EGFP at a dose of 1 × 10⁹ PFU per animal via the tail vein. Mice were sacrificed 4 weeks post-injection for further analyses. All procedures were approved by the Animal Ethics Committee of Shandong Provincial Hospital (No. 2022-104) and complied with the standards of the Institutional Animal Care and Use Committee.

Echocardiography

Cardiac performance was assessed via a Vevo® 2100 high-frequency ultrasound system (FUJIFILM VisualSonics, Toronto, Canada) under light isoflurane anesthesia. Key parameters of systolic function, including the left ventricular ejection fraction (LVEF), fractional shortening (LVFS), end-diastolic diameter (LVEDD), end-systolic diameter (LVESD), cardiac output (CO), and stroke volume (SV) were recorded via M-mode imaging. To evaluate diastolic function, pulsed-wave (PW) Doppler was employed to determine the mitral inflow E/A ratio, and tissue Doppler imaging was used to measure the E/e′ values.

Masson’s trichrome staining

Paraffin-embedded heart sections were processed through standard dewaxing and rehydration steps. Masson’s trichrome staining was performed via a commercial kit (Solarbio, Beijing, China) following the manufacturer’s protocol. In brief, the slides were treated with modified fixative at 56°C for 1 h, followed by nuclear staining with iron hematoxylin, muscle fiber staining with acid fuchsin, background differentiation with phosphomolybdic acid, and collagen fiber staining with aniline blue. After dehydration and mounting, the stained tissues were examined under a light microscope. Fibrosis was quantified by calculating the collagen area fraction in the myocardial interstitium and perivascular regions via ImageJ software.

Immunohistochemical staining

Paraffin-embedded heart sections were dewaxed, rehydrated, and subjected to antigen retrieval in citrate buffer (pH 6.0). After blocking endogenous peroxidase and nonspecific binding, the sections were incubated overnight at 4°C with primary antibodies diluted in 1% BSA: anti-p16 (sc-1661, 1:200; Santa Cruz, Dallas, USA), anti-collagen I (COL I, 14695-1-AP, 1:1000; Proteintech, Wuhan, China), anti-p21 (sc-6246, 1:200; Santa Cruz), anti-PRR (10926-1-AP, 1:400; Proteintech), and anti-collagen III (COL III; 22734-1-AP, 1:1000; Proteintech) antibodies. HRP-conjugated secondary antibodies and DAB were used for visualization, followed by hematoxylin counterstaining. Images were acquired via light microscopy, and positive staining areas or positive cell numbers were quantified via ImageJ. For immunohistochemical quantification, three randomly selected nonoverlapping fields per section and three sections per heart were analyzed.

Cell culture and treatment

Neonatal Wistar rats were obtained from Beijing Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China) and housed in a specific-pathogen-free facility under controlled conditions (22 ± 2 °C, 50–60% relative humidity, 12-h light/dark cycle) with ad libitum access to standard chow and water. All procedures were approved by the Animal Ethics Committee of Shandong Provincial Hospital (No. 2023-813) and complied with Institutional Animal Care and Use Committee guidelines. Ventricular cardiomyocytes were isolated from 1 to 3-day-old neonatal rats via enzymatic digestion with trypsin and collagenase II. After preplating to remove fibroblasts, the cells were cultured in DMEM supplemented with 10% fetal bovine serum (FBS; Gibco, Waltham, USA), 1% penicillin-streptomycin, and 5-bromo-2′-deoxyuridine (Solarbio) at 37°C and 5% CO₂.

Cardiomyocytes were assigned to the following treatment groups. The low glucose/high osmolarity (LG/HO) group received 5.5 mM glucose and 25 mM mannitol supplemented with 10% fatty acid-free bovine serum albumin (BSA; Sigma-Aldrich), where mannitol was used to match the osmolarity of the high-glucose condition. The high glucose/palmitic acid (HG/PA) group received 33.3 mM glucose and 500 μM palmitate (containing 10% fatty acid-free BSA) to mimic diabetic metabolic stress. The HG/PA+Ad-EGFP group received HG/PA treatment along with Ad-EGFP. The HG/PA+Ad-PRR group received HG/PA combined with Ad-PRR. Additionally, the HG/PA+Ad-PRR+PFT-α group was treated with HG/PA, Ad-PRR, and pifithrin-α (PFT-α, 20 μM; Sigma-Aldrich), an inhibitor of p53 transcriptional activity. A short hairpin RNA targeting PRR (shPRR: 5′-GCTCCGTAATCGCCTGTTTCA-3′) and a nontargeting control (shNC: 5′-TTCTCCGAACGTGTCACGT-3′) were cloned and inserted into adenoviral vectors. Recombinant adenoviruses (Ad-shPRR and Ad-shNC) were packaged and amplified in HEK293 cells and purified by GeneChem (Shanghai, China).

Immunofluorescence staining

Frozen myocardial sections or cultured cardiomyocytes were fixed with paraformaldehyde, permeabilized with 0.3% Triton X-100, and blocked with 10% goat serum. The samples were incubated overnight at 4°C with an anti-γ-H2AX antibody (ab81299, 1:200; Abcam, Cambridge, UK). After the samples were washed with PBS, an Alexa Fluor 647-conjugated secondary antibody (ab150083, 1:800; Abcam) was added, and the nuclei were counterstained with DAPI. Fluorescence images were acquired via a fluorescence microscope (Ti2-E; Nikon, Tokyo, Japan), and the signal intensity was quantified via ImageJ.

Senescence-associated β-galactosidase (SA-β-Gal) staining

SA-β-Gal activity was evaluated via a modified protocol based on established methodologies [26]. Briefly, frozen heart sections or cultured cardiomyocytes were fixed with fixative for 15 min and incubated with prewarmed staining solution (Beyotime, Shanghai, China) at 37°C in a CO₂-free incubator for 2 weeks or overnight. SA-β-Gal-positive areas (blue staining) were visualized under a light microscope (BX53; Olympus, Tokyo, Japan). The quantification of the positive staining area or cells was performed via ImageJ.

Conditioned medium experiment

Cardiac fibroblasts were obtained from neonatal rat ventricular tissues via the same digestion method used for cardiomyocytes. After digestion, the cell suspension was directly plated, and adherent fibroblasts were obtained via differential adhesion. Primary fibroblasts were cultured and used for subsequent experiments. For paracrine stimulation, conditioned medium from treated cardiomyocytes was collected, centrifuged, filtered, and diluted in fresh DMEM before incubating fibroblasts for 48 h. Fibroblast activation was evaluated by detecting α-smooth muscle actin (α-SMA) expression via western blotting, while cell proliferation was measured via the Cell Counting Kit-8 (CCK-8) assay. Fibroblasts were seeded into 96-well plates, treated with the conditioned medium as described, and then incubated with CCK-8 reagent (Beyotime) for 1 h at 37°C. The absorbance at 450 nm was recorded with a microplated reader (Molecular Devices, San Jose, USA) to assess viability.

Western blotting

Proteins from myocardial tissue or cardiomyocytes were extracted via RIPA buffer (Beyotime) supplemented with protease and phosphatase inhibitors (Beyotime) and quantified via a BCA assay kit (Beyotime). Equal amounts of protein were separated via 8%–12% SDS-PAGE, transferred to PVDF membranes (Merck Millipore, Darmstadt, Germany), blocked with 5% skim milk, and incubated overnight at 4°C with the following primary antibodies: anti-p16 (sc-1661, 1:500; Santa Cruz), anti-PRR (A6531, 1:800; ABclonal, Wuhan, China), anti-p53 (10442-1-AP, 1:3000; Proteintech), anti-GAPDH (10494-1-AP, 1:20000; Proteintech), anti-α-SMA (14395-1-AP, 1:6000; Proteintech), anti-p21 (sc-6246, 1:500; Santa Cruz), anti-α-Tubulin (14555-1-AP, 1:10,000; Proteintech), anti-MDM2 (66511-1-Ig, 1:3000; Proteintech), and anti-TRIM24 (14208-1-AP, 1:3000; Proteintech). After the membranes were washed with 1×TBST (Solarbio), HRP-conjugated secondary antibodies (Proteintech) were applied, and the bands were visualized via enhanced chemiluminescence (ECL) reagents (Thermo Fisher Scientific, Waltham, USA).

Enzyme-linked immunosorbent assay (ELISA)

The supernatants from the cultured cells were centrifuged for 10 min to eliminate debris. The concentrations of TGF-β1, IL-6, TGF-β2, and IL-1β were measured via commercial ELISA kits, incuding the TGF-β1 and TGF-β2 kits (Solarbio) and the IL-6 and IL-1β kits (ABclonal). Assays were performed following the manufacturer’s protocols. Standards and samples were loaded into 96-well plates precoated with capture antibodies, followed by incubation, washing, and sequential addition of biotin-labeled detection antibodies and HRP reagents. After substrate addition and color development, the reactions were stopped, and the absorbance at 450 nm was recorded via a microplate reader (Molecular Devices). The values were calculated from standard curves.

Pharmacological inhibition experiment

To investigate the degradation pathways regulating p53 stability, cardiomyocytes were treated with MG132 (10 μM; Selleck, Houston, USA), a proteasome inhibitor, or bafilomycin A1 (100 nM; Selleck), a lysosomal inhibitor. Both compounds were dissolved in DMSO, and control cells received equivalent volumes of DMSO as vehicle controls. To assess the protein half-life, cycloheximide (CHX, 50 μg/mL; Selleck) was used to inhibit new protein synthesis. Cultured cardiomyocytes were collected at 0, 1, 1.5, and 2 h after CHX treatment, after which protein extraction and western blotting were performed to evaluate p53 protein degradation over time.

Immunoprecipitation and ubiquitination assay

Cardiomyocyte lysates were prepared via IP lysis buffer with protease inhibitors (Beyotime), and immunoprecipitation was conducted via the Pierce™ Immunoprecipitation Kit (Thermo Fisher Scientific) following the manufacturer’s protocol. The lysates were incubated with an anti-p53 antibody (10442-1-AP; Proteintech) prebound to AminoLink™ Plus resin at 4°C overnight. The immune complexes were washed, eluted, and analyzed by western blotting with anti-TRIM24 (14208-1-AP, 1:1000; Proteintech). For the detection of p53 ubiquitination, cells were pretreated with MG132 (10 μM, 6 h; Selleck) to inhibit proteasomal degradation. After immunoprecipitation with an anti-p53 antibody as described, ubiquitinated p53 was identified by western blotting with an anti-ubiquitin antibody (10201-2-AP, 1:1000; Proteintech).

RNA sequencing and analysis

Cardiomyocyte RNA was isolated via TRIzol, and quality control was performed with a NanoDrop and Agilent 2100 Bioanalyzer. Samples meeting quality standards were sent to BGI Genomics (Shenzhen, China) for transcriptome sequencing. Library preparation, sequencing, and initial data processing were performed by BGI following standard procedures. Differential gene expression analysis was conducted via the DESeq2 package. Genes with an absolute log2-fold change (|log2FC|) ≥ 1 and an adjusted P value < 0.05 were considered differentially expressed genes (DEGs). The functional annotation of the DEGs was performed through Gene Ontology (GO) pathway enrichment analyses. Additionally, publicly available transcriptomic data from human cardiac tissue (GSE26887) were downloaded from the Gene Expression Omnibus (GEO) database. This dataset comprises left ventricular samples from 7 patients with type 2 diabetes mellitus and 5 nondiabetic controls. Differential gene expression in this dataset was analyzed via the GEO2R online tool. Gene set enrichment analysis (GSEA) was further used to evaluate the enrichment of senescence-related pathways in diabetic versus nondiabetic myocardial samples.

Statistical analysis

The data are expressed as the mean±SD. Normality was assessed via the Shapiro-Wilk test. For comparisons between two groups, unpaired two-tailed Student’s t tests were used, whereas one-way ANOVA with Tukey’s multiple comparisons test was used for analysis involving more than two groups. Correlations were analyzed via the Pearson method. Statistical analyses were conducted via GraphPad Prism version 9.5.1, with significance set at P  < 0.05.

Results

PRR upregulation is correlated with enhanced cellular senescence in diabetic hearts

To investigate senescence-related alterations in diabetic hearts, we first analyzed publicly available transcriptomic data (GSE26887) comparing cardiac tissues from diabetic and nondiabetic patients. GSEA of DEGs revealed significant enrichment of multiple pathways associated with cellular senescence in diabetic samples. The core cellular senescence signaling pathway was prominently upregulated ( Figure 1A). In addition, several prosenescent and SASP-associated pathways, including the MAPK, PI3K-Akt, NF-κB, TNF, IL-17, and JAK-STAT signaling pathways, were also enriched in diabetic hearts ( Figure 1B and Supplementary Figure S1), suggesting broad activation of senescence-associated signaling under diabetic conditions.

Figure 1 .

PRR expression is associated with senescence markers in diabetic hearts

(A) Gene set enrichment analysis (GSEA) of cardiac transcriptomic data (GSE26887) showing enrichment of the cellular senescence pathway in diabetic patients. (B) Bubble plot illustrating GSEA enrichment of cellular senescence-related and inflammatory pathways in diabetic hearts. (C,D) SA-β-Gal staining and quantification in myocardial tissues from normal diet (ND) controls and high-fat diet/streptozotocin (HFD/STZ)-induced DCM model mice, scale bar: 100 μm. (E,F) Immunofluorescence detection and quantification of γ-H2AX, scale bar: 10 μm. (G,H) Immunohistochemical staining and quantification of p16, scale bar: 20 μm. (I,J) Immunohistochemical staining and quantification of p21, scale bar: 20 μm. (K,L) Immunohistochemical staining and quantification of PRR expression, scale bar: 20 μm. (M‒P) Correlation analysis between PRR expression and the levels of SA-β-Gal, γ-H2AX, p16, and p21. Data are presented as the mean ± SD (n = 10). ** P < 0.01, *** P < 0.001, unpaired t-test.

To validate these findings, we established an HFD/STZ-induced DCM mouse model, as confirmed by hyperglycemia, cardiac hypertrophy, and impaired cardiac function ( Supplementary Tables S1–S3). Compared with ND control mice, DCM mice presented increased SA-β-Gal activity ( Figure 1C,D), elevated levels of γ-H2AX ( Figure 1E,F), and increased expression of p16 and p21 ( Figure 1G‒J), which was consistent with the occurrence of activated senescence in the hearts of diabetic patients. Importantly, the expression of PRR was also significantly elevated in DCM hearts ( Figure 1K,L). Western blotting further confirmed the upregulation of the senescence markers p16 and p21 and demonstrated a significant increase in PRR expression in the hearts of DCM mice ( Supplementary Figure S2). Correlation analyses revealed positive associations between PRR expression and multiple senescence markers, including SA-β-Gal activity, γ-H2AX fluorescence intensity, and p16 and p21 levels ( Figure 1M‒P). These findings indicate a potential link between increased PRR expression and enhanced cellular senescence in diabetic hearts.

PRR overexpression promotes cellular senescence in the myocardium of DCM mice

To explore the potential role of PRR in cellular senescence, we overexpressed PRR in DCM mice via adenoviral delivery. Western blotting confirmed a significant increase in PRR protein levels in Ad-PRR-treated hearts compared with Ad-EGFP controls ( Supplementary Figure S3). Compared with the Ad-EGFP control group, the PRR-overexpressing group presented significantly greater SA-β-Gal-positive staining in myocardial tissues ( Figure 2A,B). Additionally, immunohistochemical analysis revealed elevated expression of the senescence markers p16 and p21 in the PRR-overexpressing group ( Figure 2C‒E), along with increased fluorescence intensity of γ-H2AX ( Figure 2F,G). These changes suggest that PRR overexpression may aggravate cellular senescence in the diabetic myocardium.

Figure 2 .

PRR overexpression promotes cellular senescence in the diabetic myocardium

(A,B) SA-β-Gal staining and quantification in myocardial tissues from HFD/STZ-induced DCM mice treated with adenovirus encoding PRR (Ad-PRR) or control EGFP (Ad-EGFP), scale bar: 50 μm. (C–E) Immunohistochemical staining and quantification of p16 and p21, scale bar: 40 μm. (F,G) Immunofluorescence detection and quantification of γ-H2AX in cardiac tissue, scale bar: 10 μm. Data are presented as the mean ± SD (n = 10). *P < 0.05, **P < 0.01, one-way ANOVA.

PRR overexpression enhances cardiomyocyte senescence and profibrotic SASP secretion under HG/PA exposure

To further explore the involvement of PRR in cardiomyocyte senescence, we overexpressed PRR in primary cultured cardiomyocytes exposed to HG/PA treatment. Western blotting verified elevated PRR expression relative to Ad-EGFP-treated controls ( Supplementary Figure S4). HG/PA exposure alone resulted in increased SA-β-Gal-positive staining ( Figure 3A,B), upregulated the expression of p16 and p21 ( Figure 3C‒E), and increased γ-H2AX fluorescence intensity ( Figure 3F,G); and these senescence-associated markers were further augmented by PRR overexpression. Conversely, knockdown of PRR attenuated HG/PA-induced senescence phenotype, as evidenced by reduced SA-β-Gal activity, decreased γ-H2AX intensity, and downregulation of p16 and p21 ( Supplementary Figure S5). These data suggest that elevated PRR expression may intensify cellular senescence in cardiomyocytes under HG/PA exposure.

Figure 3 .

PRR overexpression promotes cardiomyocyte senescence and enhances fibroblast activation under HG/PA conditions

(A,B) SA-β-Gal staining and quantification in primary cardiomyocytes treated with low glucose/high osmolarity (LG/HO), high glucose and palmitic acid (HG/PA), HG/PA+Ad-EGFP, or HG/PA+Ad-PRR, scale bar: 20 μm. (C–E) Western blotting and quantification of p16 and p21 expression across treatment groups. (F,G) Immunofluorescence staining and quantification of γ-H2AX, scale bar: 20 μm. (H,I) ELISA measurement of secreted SASP factors (TGF-β1 and TGF-β2) in the supernatants of cultured cardiomyocytes. (J,K) Western blotting and quantification of α-SMA expression in fibroblasts after exposure to conditioned medium (CM) collected from cardiomyocytes. (L,M) Cell proliferation of fibroblasts following CM treatment was assessed via CCK-8 assay, scale bar: 100 μm. Data are presented as the mean ± SD (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, one-way ANOVA.

To evaluate the potential role of PRR in modulating SASP in cardiomyocytes, we measured the levels of representative SASP factors, including TGF-β1, TGF-β2, IL-1β, and IL-6, in the culture supernatants. Compared with the LG/HO control group, HG/PA treatment increased the secretion of these factors, indicating SASP activation under metabolic stress; and PRR overexpression in HG/PA-treated cardiomyocytes further elevated the TGF-β1 and TGF-β2 levels ( Figure 3H,I), whereas the IL-1β and IL-6 levels remained unchanged ( Supplementary Figure S6). This pattern suggests a preferential upregulation of profibrotic SASP components in response to PRR elevation.

To assess the potential paracrine impact of these secreted factors, we exposed primary cardiac fibroblasts to conditioned medium from PRR-overexpressing cardiomyocytes. This treatment increased α-SMA expression in fibroblasts ( Figure 3J,K), indicating myofibroblast activation, whereas no significant change in fibroblast proliferation was detected ( Figure 3L,M). Collectively, these results suggest that under HG/PA-induced stress, PRR may contribute to fibroblast activation by selectively enhancing profibrotic SASP factor secretion in cardiomyocytes.

PRR overexpression aggravates myocardial fibrosis and cardiac dysfunction in DCM mice

We next evaluated the impact of PRR on cardiac remodeling in DCM mice. Histological analysis revealed increased extracellular matrix deposition, particularly in the interstitial and perivascular regions of the myocardium, in PRR-overexpressing mice ( Figure 4A‒C), indicating an exacerbation of fibrotic remodeling. Collagen subtype analysis revealed elevated COL I level, whereas COL III expression remained largely unchanged, resulting in an increased COL I/III ratio ( Figure 4D‒F). This alteration points to reduced myocardial compliance and pathological stiffening.

Figure 4 .

PRR overexpression aggravates myocardial fibrosis and impairs cardiac function in DCM mice

(A) Masson’s trichrome staining showing fibrotic changes in heart sections, scale bar: 50 μm. (B,C) Quantification of interstitial and perivascular collagen deposition. (D) Immunohistochemical staining of collagen type I (COL I) and collagen type III (COL III) in myocardial tissue, scale bar: 100 μm. (E,F) Quantification of the COL I-positive area and the COL I/III ratio. (G) Representative echocardiographic images acquired by M-mode, PW Doppler, and tissue Doppler imaging. (H,I) Quantification of the left ventricular ejection fraction (LVEF) and fractional shortening (LVFS). (J,K) Assessment of diastolic function on the basis of the E/A and E/e′ ratios. Data are presented as the mean ± SD (n = 10). * P < 0.05, ** P < 0.01, *** P < 0.001, one-way ANOVA.

Functionally, PRR overexpression further impaired both systolic and diastolic function in DCM mice. LVEF and LVFS were reduced in the DCM group and decreased further with PRR overexpression ( Figure 4G‒I). A mild increase in LVESD was observed, whereas LVEDD remained unchanged ( Supplementary Figure S7A,B). Cardiac output (CO) was further reduced, whereas stroke volume (SV) did not differ significantly between the groups ( Supplementary Figure S7C,D). Diastolic dysfunction, as indicated by a decreased E/A ratio and an increased E/e′ ratio, was evident in diabetic mice and more pronounced in the PRR-overexpressing group ( Figure 4J,K). These data indicate that PRR overexpression aggravates myocardial fibrosis and contributes to further deterioration of cardiac function in DCM.

PRR promotes cardiomyocyte senescence via p53 stabilization and activation

Given that PRR overexpression upregulated p21 expression in both diabetic myocardial tissues and HG/PA-treated cardiomyocytes and that p21 is a downstream effector of p53, we hypothesized that PRR may regulate cellular senescence through p53 signaling. Our data showed that PRR overexpression increased p53 protein level in cardiomyocytes ( Figure 5A,B) without altering p53 mRNA expression ( Supplementary Figure S8), suggesting a posttranscriptional regulatory mechanism. Conversely, PRR silencing reduced p53 protein level without significantly affecting p53 mRNA level ( Supplementary Figure S9), further supporting the role of PRR in stabilizing p53 protein. CHX chase assay revealed that PRR increased p53 protein stability ( Figure 5C,D), indicating a reduction in p53 degradation. Proteasomal inhibition by MG132 restored p53 accumulation, whereas lysosomal inhibition by bafilomycin A1 had a minimal effect ( Figure 5E,F), indicating that PRR predominantly inhibits the proteasomal degradation of p53. Moreover, transcriptional inhibition of p53 via PFT-α diminished PRR-induced p21 expression ( Figure 5G,H), decreased the secretion of SASP components TGF-β1 and TGF-β2 ( Figure 5I,J), and reduced SA-β-Gal positivity ( Figure 5K,L). These findings indicate that PRR promotes cardiomyocyte senescence and SASP activation through the stabilization and activation of p53.

Figure 5 .

PRR promotes cardiomyocyte senescence through the stabilization and activation of p53

(A,B) Western blotting and quantification of p53 protein levels in HG/PA-treated cardiomyocytes overexpressing PRR. (C) CHX chase assay assessing p53 protein stability after PRR overexpression. (D) Quantification of p53 degradation rates. (E,F) Effects of the proteasome inhibitor MG132 and the lysosomal inhibitor bafilomycin A1 on p53 protein levels. (G,H) Western blotting and quantification of p21 expression following the inhibition of p53 transcriptional activity via pifithrin-α (PFT-α). (I,J) ELISA measurement of the concentrations of TGF-β1 and TGF-β2 in the culture supernatants of cardiomyocytes with PRR overexpression or PFT-α. (K,L) SA-β-Gal staining and quantification in treated cardiomyocytes, scale bar: 20 μm. Data are presented as the mean ± SD (n = 3). * P < 0.05, ** P < 0.01, one-way ANOVA.

PRR stabilizes p53 by suppressing TRIM24-mediated ubiquitination and degradation

Given the distinct subcellular localization of PRR and p53, we hypothesized that PRR might regulate p53 through intermediate factors. To explore this, transcriptomic profiling was conducted on HG/PA-treated cardiomyocytes overexpressing PRR, revealing 1277 DEGs ( Figure 6A). Among these ligases, the E3 ubiquitin ligases MDM2 and TRIM24 were identified as potential contributors to p53 regulation ( Figure 6B‒D). Further western blotting revealed that PRR overexpression resulted in decreased TRIM24 expression ( Figure 6E,F), whereas MDM2 level remained unchanged ( Supplementary Figure S10). Coimmunoprecipitation analysis confirmed a direct interaction between TRIM24 and p53, which was diminished following PRR overexpression ( Figure 6G). TRIM24 enhanced p53 ubiquitination, an effect that was attenuated by PRR coexpression ( Figure 6H). Moreover, TRIM24 overexpression reduced p53 protein levels, whereas PRR coexpression counteracted this reduction ( Figure 6I,J). Proteasome inhibition with MG132 restored p53 levels in TRIM24-overexpressing cells ( Figure 6K,L), indicating that TRIM24 mediates p53 degradation via the proteasomal pathway. Furthermore, TRIM24 coexpression partially mitigated the cellular senescence phenotype induced by PRR, as evidenced by reduced SA-β-Gal positivity ( Figure 6M,N). Additionally, TRIM24 attenuated the increased secretion of the profibrotic SASP mediators TGF-β1 and TGF-β2 observed in PRR-overexpressing cardiomyocytes ( Figure 6O,P). These findings suggest that PRR stabilizes p53 by suppressing TRIM24-mediated ubiquitination and proteasomal degradation, thereby promoting cardiomyocyte senescence under diabetic stress.

Figure 6 .

PRR stabilizes p53 by inhibiting TRIM24-mediated ubiquitination and degradation in HG/PA-treated cardiomyocytes

(A) Volcano plot of differentially expressed genes (DEGs) from transcriptomic analysis of HG/PA-treated cardiomyocytes with PRR overexpression. (B–D) Heatmaps of DEGs enriched in p53 binding (B) and ubiquitin ligase activity (C) and their overlap (D). (E,F) Western blotting of TRIM24 protein expression in HG/PA-treated cardiomyocytes overexpressing PRR. (G) Coimmunoprecipitation analysis of the interaction between TRIM24 and p53 in PRR-overexpressing cells. (H) Ubiquitination assay evaluating the effect of PRR on TRIM24-mediated p53 ubiquitination. (I,J) Western blotting and quantification of p53 protein levels. (K,L) Western blotting was used to assess the effect of MG132 treatment on TRIM24-mediated p53 degradation. (M,N) SA-β-Gal staining and quantification of positive cardiomyocytes. (O,P) ELISA was used to measure the concentrations of TGF-β1 and TGF-β2 in the culture supernatants. Data are presented as the mean ± SD (n = 3). *P < 0.05, **P < 0.01, one-way ANOVA.

Discussion

PRR has been implicated in diverse cellular processes beyond its classical roles, yet its involvement in cardiomyocyte senescence remains unclear. In this study, we reported a new function of PRR in promoting cardiomyocyte senescence in DCM. We demonstrated that PRR expression levels were significantly elevated in the myocardia of DCM mice and that, functionally, PRR overexpression exacerbated cardiomyocyte senescence and induced the secretion of profibrotic SASP factors. Mechanistically, PRR stabilized the p53 protein by suppressing TRIM24-dependent ubiquitination and proteasomal degradation, thereby increasing p53 transcriptional activity and inducing p21 expression.

Since its initial identification as a component of the RAS, PRR has been recognized as an important regulator of cardiovascular and renal diseases [ 12, 27] . Because global PRR knockout results in embryonic lethality, studies investigating its cardiac and renal functions have relied primarily on overexpression models [ 28, 29] . Recent evidence indicates that transgenic PRR overexpression promotes skeletal muscle senescence and accelerates sarcopenia, implicating PRR in age-related cellular decline [16]. In contrast, another study revealed that even marked PRR overexpression (up to 400-fold) in healthy myocardium does not trigger hypertension or fibrotic remodeling [29]. On the basis of these contrasting findings, PRR may function as amplifiers of pathological signaling rather than directly initiating disease processes [30]. Consistent with this notion, our study revealed significant upregulation of myocardial PRR expression under diabetic stress, which contributes to enhanced p53-dependent senescence signaling.

In this study, we found that PRR stabilizes the p53 protein through the inhibition of TRIM24-mediated ubiquitination and proteasomal degradation. These findings expand the current understanding of PRR beyond its conventional roles, positioning it as a modulator of protein homeostasis. Previous research has implicated PRR in regulating the degradation dynamics of several target proteins, such as CD36 and LDLR, with downstream effects on lipid metabolism and cellular uptake [ 31, 32] . Additionally, PRR has been shown to prevent Wnt3 degradation by inhibiting NEDD4L-mediated ubiquitination, thereby promoting oncogenic signaling in colorectal cancer [33]. Although p53 is well known to regulate apoptosis and autophagy, transcriptome analysis of PRR-overexpressing cardiomyocytes revealed no significant induction of classical downstream targets such as Bbc3 (PUMA), Fas, Dram1, or Prkab1 ( Supplementary Figure S11). These results suggest that PRR-induced senescence may occur independently of canonical p53-mediated apoptotic or autophagic pathways, although this hypothesis warrants further validation. TRIM24, a known E3 ligase for p53 in tumor models  [ 24, 25] , functions similarly in cardiomyocytes by influencing p53 turnover. This finding represents a novel extension of the regulatory role of TRIM24 in cardiovascular disease. In support of the importance of TRIM family proteins in heart disease, clinical data have linked the downregulation of TRIM32, another TRIM member, with maladaptive cardiac remodeling and decompensated function in patients with dilated cardiomyopathy [34]. Our study provides preliminary evidence that pharmacological inhibition of p53 or TRIM24 overexpression partially reversed the senescent phenotype, suggesting that the PRR-TRIM24-p53 axis represents a promising therapeutic target. Given the pleiotropic functions of p53, this axis may act as a broader molecular switch in cellular stress responses. However, owing to the essential tumor-suppressive role of p53, targeting TRIM24 or PRR may offer a more specific and safer therapeutic strategy, warranting further investigation.

SASP represents a key feature of senescent cells, mediating their impact on surrounding cells and the tissue microenvironment via paracrine signaling. In this study, we demonstrated that PRR overexpression led to increased expression of the profibrotic SASP factors TGF-β1 and TGF-β2, rather than inflammatory components such as IL-6 and IL-1β, in cardiomyocytes exposed to HG/PA. This selective enrichment aligns with previous reports showing that senescent cardiomyocytes preferentially secrete profibrotic and prohypertrophic factors such as TGF-β2 and GDF15 rather than classical inflammatory cytokines [35]. In line with this fibrotic shift, conditioned medium derived from PRR-overexpressing cardiomyocytes stimulated α-SMA expression in cardiac fibroblasts, indicating the activation of myofibroblast phenotypes. Interestingly, this activation was not accompanied by increased fibroblast proliferation, suggesting the possibility of a senescence-like phenotype in fibroblasts driven by cardiomyocyte-derived paracrine factors, a phenomenon also described in other models of paracrine senescence [35]. Mechanistically, the selective induction of profibrotic SASP components may involve p53 activation. Previous studies have shown that p53 can suppress NF-κB-driven inflammatory cytokines while promoting TGF-β/Smad-dependent fibrotic signaling pathways [ 36, 37] . Therefore, our findings suggest that PRR-mediated stabilization and activation of p53 not only accelerate cardiomyocyte senescence but also skew the SASP composition toward profibrotic factors, thereby contributing to myocardial fibrosis and functional deterioration in DCM.

Several limitations should be considered in this study. First, we were unable to directly assess the effects of PRR depletion, as the cardiomyocyte-specific PRR-knockout mice we generated exhibited early postnatal lethality, which is consistent with previous findings [14]. The phenotype observed in PRR-deficient models is largely attributable to the disruption of V-ATPase function, complicating the use of knockout models to dissect PRR-specific roles. Although the handle region peptide (HRP) has been applied as a PRR antagonist, its inhibitory effect is confined to blocking (pro)renin binding, with limited influence on RAS-independent pathways such as the MAPK and Wnt/β-catenin signaling pathways [ 38, 39] . Thus, HRP may not effectively target the PRR-TRIM24-p53 axis explored in this study. Future research utilizing targeted small molecule inhibitors or antibodies against the intracellular domains of PRR may provide more specific insights into their nonclassical functions. Second, although TRIM24 has been well established to promote K48-linked ubiquitination of p53 via its RING domain [ 25, 40] , this mechanism was not directly validated in our experiments. Considering the complex network of E3 ligases regulating p53 stability, further interactome studies are needed to clarify the precise contribution of TRIM24 to diabetic stress. Third, although PRR overexpression exacerbated cardiac fibrosis and dysfunction in DCM mice, whether these effects can be reversed by p53 inhibition or TRIM24 overexpression in vivo remains to be determined. Future studies are needed to evaluate the therapeutic importance of targeting this pathway in DCM. Finally, our evaluation of SASP components was limited to selected factors. Comprehensive profiling of the SASP secretome via transcriptomic or proteomic approaches will be necessary to better understand its role in myocardial remodeling during DCM progression.

In summary, this work identified PRR as a regulator of cardiomyocyte senescence in DCM through stabilization of p53 via TRIM24 suppression. These findings expand the role of PRR beyond its classical involvement in RAS signaling and suggest novel therapeutic targets for attenuating diabetic myocardial remodeling and dysfunction.

Supplementary Data

Supplementary data are available at Acta Biochimica et Biophysica Sinica online.

COMPETING INTERESTS

The authors declare that they have no conflict of interest.

Funding Statement

This work was supported by the grants from the National Natural Science Foundation of China (Nos. 82371574, 82070382, and 81570729) and the Taishan Scholars Programme (No. ts20190979).