Inhibiting the Hif-1α-Drp1 axis alleviates mitochondrial dysfunction and reduces senescence-like changes in myocardial tissue after acute myocardial infarction

Abstract

Background

Cellular senescence is a significant pathological process in acute myocardial infarction (AMI), yet its upstream regulatory mechanisms remain unclear. This study reveals that the HIF-1α /Drp1 signaling axis serves as a critical link between hypoxia and senescence-like changes in myocardial tissue by disrupting mitochondrial homeostasis.

Methods

Using rat AMI model (in vivo) and hypoxia‑exposed H9c2 cardiomyocytes (in vitro), we assessed mitochondrial morphology, mtROS, and senescence markers. Genetic gain‑ and loss‑of‑function approaches were applied to modulate Drp1 and HIF‑1α. We further evaluated the effects of the mitochondrial fission inhibitor Mdivi‑1 and the SASP inhibitor Ruxolitinib on mitochondrial function, senescence, and apoptosis.

Results

AMI/hypoxia activated the ERK1/2-Akt pathway, promoting Drp1 Ser616 phosphorylation and mitochondrial translocation, leading to excessive fission, mtROS burst, and senescence-like changes in myocardial tissue (in vivo) / cardiomyocyte senescence (in H9c2 cells). HIF-1α transcriptionally regulated Drp1 expression. Mdivi-1 restored mitochondrial dynamics, reduced SASP-related inflammation, and improved cardiac function, indicating a myocardial protective effect. Ruxolitinib suppressed Drp1 Ser616 phosphorylation and alleviated senescence, apoptosis, and pyroptosis.

Conclusion

HIF-1α-mediated upregulation of Drp1 and ERK-dependent phosphorylation of Drp1 collectively drive mitochondrial dysfunction and senescence-like changes in myocardial tissue in AMI. Targeting these pathways or the SASP pathway represents a promising therapeutic strategy.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13062-026-00847-8.

Introduction

Acute Myocardial Infarction (AMI) is one of the leading cardiovascular diseases globally, resulting in elevated mortality and morbidity rates [1, 2]. AMI is typically caused by blockage in the coronary arteries, leading to insufficient blood supply to the myocardium, which triggers myocardial cell necrosis and causes irreversible damage to the heart muscle [3]. While clinical treatments primarily focus on restoring blood flow and minimizing acute damage, long-term myocardial functional impairment and fibrosis remain significant challenges [4]. Therefore, a deeper understanding of the mechanisms underlying AMI-induced myocardial cell injury is crucial for the development of new therapies.

Mitochondria are central to cellular energy metabolism, and their dynamic balance-including fission and fusion-is essential for maintaining mitochondrial function and cellular health [5, 6]. Imbalance in mitochondrial dynamics, particularly an increase in fission, has been closely linked to the pathogenesis and progression of various cardiovascular diseases [7, 8]. Drp1 (dynamin-related protein 1) is the main regulatory factor for mitochondrial fission, playing a critical role in facilitating mitochondrial division and influencing processes such as apoptosis, stress responses, cellular senescence, and energy metabolism [9, 10]. Mitochondrial dysfunction in cardiovascular diseases is often associated with abnormal activation of Drp1; excessive mitochondrial fission leads to decreased mitochondrial membrane potential and increased levels of reactive oxygen species (ROS), which in turn triggers myocardial cell death and worsens cardiac function [11]. Previous studies have demonstrated that Drp1 is upregulated in myocardial ischemia-reperfusion injury, promoting mitochondrial fission and resulting in mitochondrial dysfunction and cell death [12, 13].

Cellular senescence is an irreversible state of cell cycle arrest, accompanied by mitochondrial dysfunction, metabolic dysregulation, and the development of a senescence-associated secretory phenotype (SASP) [14–16]. Recent studies have revealed that cardiomyocyte senescence not only characterizes natural cardiac aging but also plays a significant role in acute cardiac injuries such as AMI [17, 18]. Ischemic stress can directly induce stress-induced premature senescence (SIPS) in cardiomyocytes, characterized by upregulation of senescence markers such as p16, p21, and p53, telomere shortening, and disruption of mitochondrial cristae structure [19–22]. More importantly, senescent cardiomyocytes secrete SASP-related inflammatory factors (e.g., IL-1α, IL-1β, IL-6, IL-8, MCP-1, etc.), which exacerbate local inflammatory responses, promote fibroblast activation and collagen deposition, thereby driving myocardial fibrosis and adverse ventricular remodeling [23, 24]. Of note, mitochondrial dynamics imbalance, particularly Drp1-mediated excessive mitochondrial fission, has been identified as a key upstream event triggering cardiomyocyte senescence. MITOL (mitochondrial E3 ubiquitin ligase) inhibits excessive mitochondrial fission by ubiquitinating and degrading Drp1; its deficiency leads to Drp1 accumulation, mitochondrial fragmentation, and ROS burst, ultimately accelerating cardiomyocyte senescence and cardiac dysfunction [25]. Furthermore, the interaction between Drp1 and the cytoskeletal protein filamin A has also been shown to participate in the regulation of post-AMI cardiomyocyte senescence [26]. However, the specific regulatory mechanism of Drp1 in AMI-induced cardiomyocyte senescence remains unclear, and whether targeting Drp1 can improve post-AMI cardiac prognosis by inhibiting cellular senescence awaits further elucidation.

This study aims to systematically assess the role of Drp1 in AMI-induced myocardial tissue senescence-like changes and mitochondrial dysfunction by establishing rat AMI models and H9c2 cardiomyocyte hypoxia models. Through experiments involving the overexpression and knockdown of Drp1, we will further investigate its specific mechanisms in regulating mitochondrial dynamics, ROS generation, and cellular senescence. Additionally, we will assess the protective effects of targeted inhibition of Drp1 on myocardial injury in AMI.

Materials and methods

Animal models and treatment

In this study, eight-week-old male SD rats were obtained from Beijing Vital River Laboratory Animal Technology Co., Ltd. All experimental procedures strictly adhered to ethical guidelines for animal experimentation. This study was approved by the Institutional Animal Care and Use Committee of The First Affiliated Hospital, Jiangxi Medical College, Nanchang University (IACUC Issue No: 202311QR015). The AMI model was induced by ligating the left anterior descending coronary artery (LAD) [27]. The permanent LAD ligation model was selected based on a previously published study [28], which successfully established an AMI rat model using this method and demonstrated reliable induction of myocardial injury. This well-validated modeling approach was adopted to ensure the reproducibility of our experiments. Post-surgery, the SD rats were randomly divided into the AMI + DMSO group and the AMI + Mdivi-1 (mitochondrial division inhibitor 1, mitochondrial fission inhibitor) group. As a prophylactic intervention in this experimental model, Mdivi-1 was administered as a single intraperitoneal injection at a dose of 1.2 mg/kg, 15 min prior to surgery [29]. The control group received the same dose of DMSO. Day 7 post-surgery was selected as the key time point for tissue collection and analysis, as at this time point the acute inflammatory phase has largely subsided while ventricular remodeling has just begun, allowing optimal detection of senescence-like changes. All specimens were selected from the papillary muscle position for image acquisition. As an important part of the heart, the papillary muscle can reflect the overall pathological changes of the heart, ensuring that the data is representative and consistent.

Cell culture and treatment

The rat cardiomyocyte cell line H9c2 (purchased from Procell, Cat. No. H9c2(2 − 1)) was cultured in DMEM (Sevier Biotech) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin in a 37 °C, 5% CO₂ environment. Cells were passaged once they reached 80% confluence.

A hypoxic cell model simulating AMI was established using H9c2 cells [30]. When cardiomyocytes at the logarithmic growth phase reached 70%–80% confluence, the culture flask caps were loosened to allow gas exchange, and the flasks were placed in a sealed plastic container. The container was then filled with a hypoxic gas mixture (94% N₂ + 5% CO₂ + 1% O₂) by continuous insufflation for 10 min. After sealing the container (with the flask caps kept loosened), it was transferred to a 37 °C incubator for culture. Cells were treated with 25 nM mito-TEMPO (MT; MCE) for 24 h [31] and 300 nM ruxolitinib (CAS No.: 941678-49-5; MCE) for 48 h.

Cell transfection

To achieve gene knockdown or overexpression, H9c2 cells were transfected using Lipofectamine™ 3000 transfection reagent (Invitrogen, L3000015). The transfection vectors included Drp1 siRNA, HIF-1α siRNA, with si-NC as the negative control (GenePharma), as well as the Drp1 overexpression plasmid pcDNA3.1-Drp1, with the empty vector pcDNA3.1 serving as the control (GenePharma). The specific procedure was as follows: Cells were seeded in 6-well plates and cultured until 70%–80% confluence. The siRNAs or overexpression plasmids were mixed with Lipofectamine™ 3000 in OPTI-MEM low-serum medium (Invitrogen, 31985070) following the standardized instructions provided by the manufacturer and incubated at room temperature for 20 min to form complexes. The complexes were then added to the cells, and after 4–6 h of incubation, the medium was replaced with complete medium containing 10% FBS. Experiments were conducted 48 h after cell transfection, with transfection efficiency confirmed by Western blot analysis.

EdU (5-Ethynyl-2’-deoxyuridine) detection of cell proliferation

EdU was utilized to assess the proliferation of H9c2 cells under hypoxic conditions. Following treatment, the cells were incubated with 20 µM EdU at 37 °C for 2 h. They were then fixed with 4% paraformaldehyde and permeabilized using PBS containing 0.1% Triton X-100 (Sigma). The Click reaction was performed using the BeyoClick™ EdU-488 Cell Proliferation Detection Kit (Beyotime), and DAPI was employed for nuclear staining. After staining, EDU-positive cells were observed using a fluorescence microscope (Keyence). Five random fields were selected for imaging, and ImageJ software was used for cell counting. The percentage of EDU-positive cells was calculated to assess the cell proliferation level.

Transmission electron microscopy (TEM) for mitochondria detection

H9c2 cell samples were fixed in 2.5% glutaraldehyde at 4 °C for 2 h, followed by fixation with 1% osmium tetroxide for 1 h. Samples were dehydrated through gradient ethanol and washed with acetone before being embedded in epoxy resin, with polymerization at 60 °C for 48 h. Using an ultramicrotome, samples were sliced into 70 nm thick sections, followed by contrast staining with uranyl acetate and lead citrate. Finally, the samples were observed using a transmission electron microscope to capture high-resolution images of mitochondrial ultrastructure.

Senescence-associated β-galactosidase (SA-β-gal) detection

To evaluate the degree of cellular senescence, a cell SA-β-gal staining kit (Beyotime) was utilized. For cells in different treatment groups, 1 mL of β-galactosidase staining fixative was added to each well for fixation. Following fixation, the fixative was removed, and cells were washed with PBS. Subsequently, 1 mL of staining working solution was added to each well, and the cells were observed under a standard optical microscope according to the product instructions. Five random fields were selected for imaging, and ImageJ software was used for cell counting to calculate the percentage of SA-β-gal positive cells.

Enzyme-linked immunosorbent assay (ELISA)

The levels of CK-MB, LDH, and CPK in rat serum were measured using ELISA kits (JL12296, JL13677, JL21154; Jianglai Bio). Following the manufacturer’s instructions, serum samples and standards were added to a 96-well plate, incubated, and detection antibodies were subsequently added, followed by a color substrate. The reaction was terminated, and absorbance was measured at 450 nm. All results were quantitatively analyzed based on a standard curve.

Tissue sectioning and staining

Heart tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned into 4–5 μm slices. The sections were then subjected to histological staining, including hematoxylin and eosin (H&E) staining and Masson’s trichrome staining, to evaluate cardiac pathological changes.

H&E staining

Heart tissue sections were first deparaffinized and rehydrated, then stained with hematoxylin (Sevier Biotech) for 5 min and rinsed. Differentiation was performed using 0.5% hydrochloric acid alcohol, followed by another rinse. The sections were then stained with eosin (Sevier Biotech) for 2 min, rinsed again, dehydrated, cleared, and finally mounted with coverslips using mounting medium. Pathological changes and severity of cardiac tissue were observed under a microscope (DM500, Leica).

Masson’s trichrome staining

Cardiac fibrosis was assessed using a Masson’s trichrome staining kit (Solarbio). After deparaffinization and rehydration, heart tissue sections were placed in distilled water for standby. The sections were then sequentially stained using Masson reagents: nuclei were stained with Weigert’s iron hematoxylin for 8 min, followed by rinsing and bluing with tap water. Acid fuchsin solution was applied for 8 min, briefly rinsed, then treated with phosphomolybdic acid for 1 min, and subsequently stained with aniline blue solution for 2 min. After staining, the sections were dehydrated with graded ethanol, cleared with xylene, and mounted using neutral balsam. The slides were observed under a light microscope (Leica DM500). Collagen fibers appeared blue, muscle fibers red, and nuclei blue-black. The fibrotic area was quantitatively analyzed using ImageJ software.

2,3,5-triphenyl tetrazolium chloride (TTC) staining

TTC staining was employed to evaluate the area of myocardial infarction. Fresh heart tissues were embedded in OCT compound without fixation and rapidly frozen at − 20 °C for 30 min. The hearts were then sliced transversely into 2–3 mm thick sections. The slices were incubated in TTC solution (Servicebio) at 37 °C in the dark for 30 min. After staining, the sections were fixed in 4% paraformaldehyde. Viable myocardium appeared red, while infarcted areas appeared white. The infarct size ratio was analyzed using ImageJ software.

Immunohistochemistry (IHC)

IHC was utilized to detect the expression of specific proteins in cardiac tissue. Cardiac tissue Sections.  (4 μm thick) were dewaxed and rehydrated, followed by treatment with 100 µL of endogenous peroxidase blocker. The sections were then blocked with 10% goat serum to prevent nonspecific binding. Subsequently, the sections were incubated overnight at 4 °C with primary antibody Drp1 (ab184247, 1:1000, Abcam). The following day, an HRP-conjugated secondary antibody (ab6721, 1:5000, Abcam) was applied for incubation. DAB (Sevier Biotech) was used for coloration, and nuclei were counterstained with hematoxylin (Beyotime). After dehydration through alcohol gradient, the sections were mounted and observed with an optical microscope (Leica).

MitoTracker red staining

To evaluate mitochondrial morphology and function, MitoTracker Red CMXRos (Beyotime) staining was employed. Cells were incubated with MitoTracker Red dye at concentrations ranging from 20 nM at 37 °C for 30 min to label active mitochondria. After staining, cells were washed three times with PBS and observed under a fluorescence microscope (Leica) for mitochondrial morphology. Changes in mitochondrial morphology were quantitatively analyzed using ImageJ software.

Chromatin immunoprecipitation (ChIP)

A ChIP kit (Beyotime, P2078) was used to detect the binding of HIF-1α to the promoter region of target genes under normoxic and hypoxic conditions. We first fix the cells to cross-link DNA and proteins. Subsequently, the cells were lysed and the chromatin was sheared using an ultrasonic cell crusher (Leipu Instruments, model LP-XP150). Next, immunoprecipitation was performed using HIF-1α-specific antibody (abcam#ab1) and IgG (negative control). Finally, the immunoprecipitated DNA was assayed by qPCR to analyze the binding enrichment of HIF-1α.

Dual-luciferase reporter gene assay

Experiments were performed using a dual-luciferase reporter gene assay kit (Beyotime, RG027). First, construct the PGL3-DRP1 vector (inserting the target gene promoter sequence) and the empty vector PGL3-Vector, and then use Lipo8000™ transfection reagent (Beyotime) to transfect them into cells respectively. Luciferase activity was measured under control and hypoxic conditions. The degree of activation of the DRP1 promoter under hypoxic conditions was evaluated by relative luciferase activity, using the empty vector as a control.

Immunofluorescence (IF) staining

DNA damage detection

DNA damage was detected through γ-H2AX IF staining. Fixed cardiac tissue sections were permeabilized and then blocked with a blocking solution for 2 min at room temperature to prevent nonspecific binding. The sections were incubated overnight at 4 °C with a rabbit anti-γ-H2AX primary antibody, followed by incubation the next day with an Alexa Fluor^® 488-conjugated anti-rabbit secondary antibody for 1 h at room temperature in the dark. Nuclei were counterstained with DAPI, and the sections were mounted with mounting medium. Fluorescence imaging was performed using a fluorescence microscope (Keyence). The experimental procedure was strictly carried out according to the instructions of the DNA damage detection kit (Beyotime, C2035S). The proportion of γ-H2AX–positive cells was quantitatively analyzed using ImageJ software to assess the extent of DNA damage.

Co-localization analysis of Drp1 and mitochondria

IF staining was performed to detect the co-localization of Drp1/p-Drp1(Ser616) with mitochondria in both H9c2 cells and myocardial tissue.

For H9c2 cells, cells were collected and adjusted to a density of 5 × 10⁵ cells/mL. Mitochondria were labeled with MitoTracker Red (Beyotime), followed by fixation with 4% paraformaldehyde for 15 min and permeabilization with 0.1% Triton X-100. Cells were then incubated overnight at 4 °C with anti-Drp1 antibody (1:1000, Wuhan Sanying), followed by HRP-conjugated secondary antibody (1:5000, Abcam) for 2 h. DAPI was used for nuclear staining.

For myocardial tissue, paraffin-embedded sections from four experimental groups (Sham + DMSO, Sham + Mdivi-1, AMI + DMSO, AMI + Mdivi-1) were dewaxed, rehydrated, and subjected to antigen retrieval using a microwave oven. After permeabilization with 0.2% Triton X-100 and blocking with 3% BSA, sections were incubated with anti-p-Drp1(Ser616) antibody (Abcam, ab314755, 1:200) overnight at 4 °C, followed by FITC-conjugated secondary antibody (Beyotime, A0562) for 1 h. Mito-Tracker Red CMXROS (Beyotime, 1:1000) was then applied at 37 °C for 1 h to label mitochondria, and DAPI was used for nuclear staining.

All images were captured using a fluorescence microscope (Keyence, BZ-X800) and fluorescence intensity was analyzed using ImageJ software.

Protein extraction and western blot

Heart tissue or H9c2 cells were lysed with RIPA buffer to extract total protein. After separation by SDS-PAGE, proteins were transferred to PVDF membranes. The membranes were blocked with 5% nonfat milk and incubated with antibodies against Drp1 (abcam#ab184247), Fis-1 (proteintech#10956-1-AP), Mfn-2 (abcam#ab124773), p16 (Cell Singaling#80772), p21 (abcam#ab109520), p53 (abcam#ab26), Lamin B1 (proteintech#12987-1-AP), Bax (proteintech#60267-1-Ig), Bcl-2 (BOSTER#A00040-2), HIF-1α (abcam#ab1), Phospho-DRP1 (Ser616) (Cell Singaling#3455), ERK1/2 Monoclonal antibody (proteintech#66192-1-Ig), Phospho-ERK1/2 (Thr202/Tyr204) (proteintech#28733-1-AP), Phospho-DRP1 (Ser637) (Cell Singaling#20990), Phospho-AKT (Ser473) Monoclonal (proteintech#66444-1-Ig), AKT (proteintech#10176-2-AP), GSDMD (Full length + N terminal) (abclonal#A20197SP), caspase-1 (Biodragon#RM8397), cle-caspase-1 (Affinity Biosciences#AF4005). All the above antibodies were used at a dilution of 1:1000, followed by incubation with HRP-conjugated secondary antibody (ab6721, 1:5000, Abcam). ERK1/2, HIF-1α and Bax were incubated with a different secondary antibody (ab6789, 1:5000, Abcam). β-tubulin (AB0039, 1:1000, Abcam) served as an internal control. Signals were detected using chemiluminescence, and images were quantitatively analyzed with ImageJ software, and all values were normalized to the signal values of the control samples.

Real-time quantitative PCR (qRT-PCR)

Total RNA was extracted from cardiac tissue or H9c2 cells, and cDNA was synthesized using the PrimeScript™ RT Reagent Kit (Takara). Subsequently, qRT-PCR was performed using TB Green^® Premix Ex Taq™ II (Takara) to monitor DNA amplification in real-time and analyze gene expression levels. The target genes included the mRNA expression of Drp1, Fis-1, Mfn-2, p16, p21, p53, Lamin B1, IL-1α, IL-1β, IL-8, MCP-1, ICAM-1, Bax, Bcl-2, and HIF-1α. Relative expression levels were calculated using the 2^^−ΔΔCt method, with β-actin as the internal control. The sequences of the target genes are shown in Table 1.

Table 1.

Primers used in qRT-PCR

Gene	Forward sequence (5’-3’)	Reverse sequence (5’-3’)
Drp1	CGCTGATCCCGGTCATCAAT	ACTCCATTTTCTTCTCCTGTTGT
p16	GCTCTCCTGCTCTCCTATGG	AGATACCGCAAATACCGCAC
p21	TCTGTGAGACAACAAGGGCAA	GTGGAACAGGTCGGACATCA
p53	CTCCTCTCCCCAGCAAAAGA	GTAGACTGGCCCTTCTTGGT
Lamin B1	AACAGCTGCTTGACGTGAAG	ACTGCTCGCCTCTGATTCTT
IL-1α	AGGATCGTCAAGCAGGAGTT	GGAGACTGCCCATTCTCGAC
IL-1β	GACTTCACCATGGAACCCGT	TGTCGTTGCTTGTCTCTCCT
IL-8	ACCGATGTCTACGTGCTGAA	GATGGCCAGGTATCGATCCA
MCP-1	GCTGCTACTCATTCACTGGC	GGTGCTGAAGTCCTTAGGGT
ICAM-1	TCGGTGCTCAGGTATCCATC	GCCACAGTTCTCAAAGCACA
Bax	GAGACACCTGAGCTGACCTT	CGTCTGCAAACATGTCAGCT
Bcl-2	CGGGAGAACAGGGTATGA	CAGGCTGGAAGGAGAAGA
Fis-1	AAAGAGGAGCAGCGGGATTA	TGGGGCTCAGTCTGTAACAG
Mfn-2	TCAGTAGCCAATCTGGACCT	TCTCTTGGATGTAGGCCCCC
HIF-1α	CCAGCAGACCCAGTTACAGA	TTCCTGCTCTGTCTGGTGAG
β-actin	CATTGCTGACAGGATGCAGAAGG	TGCTGGAAGGTGGACAGTGAGG

Flow cytometry detection of mitochondrial reactive oxygen species (mtROS) and cell apoptosis

mtROS detection

MitoSOX™ Red probe (Thermo Fisher) was used to detect mtROS. A total of 5 × 10⁵ logarithmic-phase cells were collected, washed twice with PBS, and resuspended in 200 µL of 1×Binding Buffer. MitoSOX dye was added in the dark and incubated at room temperature for 30 min. Flow cytometry (Agilent) was then performed, and FlowJo software was used to analyze the percentage of MitoSOX-positive cells to assess mitochondrial ROS levels.

Cell apoptosis detection

Apoptosis of cardiac cells was detected using the Annexin V/PI double staining method (Annexin V-FITC Apoptosis Detection Kit, Invitrogen). Cells in the logarithmic growth phase were digested with 0.25% trypsin without EDTA and counted. A total of 5 × 10⁵ cells were collected, washed with PBS, and resuspended in 1×Binding Buffer. Under light-protected conditions, 5 µL of Annexin V-FITC was added and incubated for 10 min, followed by the addition of 10 µL of 20 µg/mL PI for staining. The proportions of early and late apoptotic cells were then analyzed using a flow cytometer (Agilent).

Echocardiography to assess cardiac function

Cardiac function was assessed using echocardiography (MyLab™ SigmaPVET, Esaote), focusing particularly on ejection fraction (EF) and fractional shortening (FS). Prior to the procedure, the animals were prepared for image acquisition by removing chest hair and maintaining a stable body temperature. All animals were subjected to inhalation anesthesia using isoflurane, and echocardiographic measurements were performed within the same post-anesthesia time window across all animals. During the anesthesia period, heart rate was maintained at 400–600 bpm, and body temperature was maintained at 39 °C (range: 38.5–39.5 °C), both of which were continuously monitored and kept stable. Coupling gel was applied to the chest, and a high-frequency probe was used to acquire long-axis and short-axis M-mode echocardiographic images. Left ventricular internal diameter in diastole (LVIDd) and left ventricular internal diameter in systole (LVIDs) were recorded. EF and FS were automatically calculated by the built-in software to comprehensively evaluate cardiac pumping function.

Statistical analysis

Data were analyzed using GraphPad Prism 9.5 (GraphPad Software, USA) statistical software. All data are presented as means ± standard deviation (SD). Intergroup differences were analyzed using two-tailed t-test for comparisons between two groups, one-way ANOVA for comparisons involving a single variable, and two-way ANOVA for experiments involving two or more variables (e.g., Mdivi-1 and Ruxolitinib treatments), followed by Bonferroni correction for post hoc analysis. P < 0.05 was considered statistically significant.

Results

AMI induces increased Drp1 levels in rat cardiac tissue and promotes senescence-like changes in myocardial tissue, while in H9c2 cells it correlates with cardiomyocyte senescence

In the AMI model, the role of Drp1 in senescence-like changes in myocardial tissue and cardiac injury was validated (Fig. 1). Long-axis echocardiography results showed that compared to the Sham group, the EF, and FS in the AMI group were significantly decreased, while the LVIDd and LVIDs were significantly increased (Fig. 1A-B). Short-axis echocardiography further validated these results (Fig. S1A-B). The infarct area of the heart was marked using TTC staining, with 1–5 representing different regions of the heart. The results showed that the white infarct area in the AMI group was significantly larger, indicating substantial heart damage (Fig. 1C-D). H&E staining results showed that the myocardial cells in the Sham group rats were arranged regularly, and the muscle fibers were intact. In the AMI group rats, some myocardial cell nuclei were lost, accompanied by local hemorrhage and neutrophil infiltration (Fig. S1C). Masson staining further demonstrated a significant increase in collagen fibers and enhanced fibrosis in the myocardial tissue of the AMI group (Fig. S1D), indicating that AMI induced marked myocardial fibrosis. Compared with the Sham group, the AMI group showed significantly increased mRNA and protein levels of Drp1 (Fig. 1E-G), along with elevated myocardial injury markers (CK-MB, LDH, CPK), DNA damage (γ-H2AX), senescence-associated factors (p16, p21, p53), and inflammatory factors (IL-1α, IL-1β, IL-8, MCP-1, ICAM-1), while Lamin B1 was significantly decreased (Fig. 1H-L). Collectively, these in vivo findings indicate that AMI upregulates Drp1 in rat cardiac tissue, which is associated with cardiac dysfunction, myocardial injury, fibrosis, and senescence-like changes in myocardial tissue.

Fig. 1.

AMI induces an increase in Drp1 levels in rat heart tissue, leading to senescence-like changes in myocardial tissue. (A-B) Echocardiography and quantitative analysis of four key cardiac function parameters (n =6). (C-D) TTC staining to assess the myocardial infarction area and quantify infarct size (n =6). (E) qRT-PCR to detect Drp1 levels in tissue (n =3). (F) Western blot to detect Drp1 levels in tissue (n =3). (G) IHC to detect Drp1 levels in cardiac tissue (n =6). (H) ELISA to measure the serum levels of CK-MB, LDH, and CPK (n =3). (I) γ-H2AX IF assay to detect DNA damage (n =3). (J-K) qRT-PCR and Western blot to detect the mRNA levels and protein expression of senescence-related markers p16, p21, p53, and Lamin B1 in cardiac tissue (n =3). (L) qRT-PCR to detect the mRNA levels of SASP in cardiac tissue (n =3). All of the above experiments were biological replicates. *P < 0.05, ** P < 0.01, ***P < 0.001, vs Sham group

To further verify the effects of Drp1 on H9c2 cell proliferation and apoptosis, hypoxic treatment was performed to simulate AMI in vitro (Fig. S2). Compared with the control group, the hypoxia group showed significantly increased H9c2 cardiomyocyte senescence (increased SA-β-gal positivity, Fig. S2A; upregulation of p16, p21, p53, and downregulation of Lamin B1, Fig. S2B-C), elevated mRNA levels of inflammatory factors IL-1α, IL-1β, IL-8, MCP-1, and ICAM-1 (Fig. S2D), inhibited proliferation (EdU, Fig. S2E-F), increased apoptosis (Annexin V/PI, Fig. S2G-H), and altered Bax/Bcl-2 expression (upregulated Bax and downregulated Bcl-2, Fig. S2I-J). In summary, these in vitro experiments confirm that Drp1 regulates H9c2 cell proliferation, apoptosis, and cardiomyocyte senescence under hypoxic conditions.

ERK1/2-Akt kinases regulate Drp1 phosphorylation and mitochondrial fission in cardiomyocyte senescence induced by AMI

This study aimed to investigate the role of Drp1 in in senescence-like changes in myocardial tissue (in vivo) and cardiomyocyte senescence (in vitro) induced by AMI, as well as in associated mitochondrial dysfunction, and to evaluate the cardioprotective effects of targeted Drp1 inhibition. Through in vivo and in vitro experiments, we demonstrated that AMI and the hypoxic microenvironment lead to mitochondrial dynamics imbalance. In the AMI rat model, compared with the Sham group, the mitochondrial fission protein Fis-1 was significantly upregulated, while the fusion protein Mfn-2 was significantly downregulated, indicating a shift of mitochondrial dynamic balance toward excessive fission (Fig. 2A, Fig. S3A). In the H9c2 hypoxic cell model, we further observed significant mitochondrial structural and functional damage. Transmission electron microscopy revealed mitochondrial swelling and disruption of cristae structure (Fig. 2B); Mito-tracker staining showed punctate mitochondrial fragmentation (Fig. 2C, Fig. S3B); MitoSOX detection indicated that the proportion of mtROS-positive cells dramatically increased from 4.99% to 71.34% (Fig. 2D, Fig. S3C); Western blot and qRT-PCR results similarly confirmed the increase of Fis-1 and the decrease of Mfn-2 (Fig. 2E, Fig. S3D). From a mechanistic perspective, hypoxic conditions effectively activated Drp1. IF results demonstrated that hypoxia promoted Drp1 translocation to mitochondria (Fig. 2F); Western blot further revealed that hypoxia upregulated the pro-fission phosphorylation level of Drp1 at the Ser616 site while downregulating the inhibitory phosphorylation level at the Ser637 site (Fig. 2G-H). Upstream signaling pathway analysis showed that the levels of p-ERK1/2 and p-Akt were significantly increased in the hypoxic group (Fig. 2I). In summary, the AMI-induced hypoxic microenvironment (in vivo) activates the ERK1/2-Akt signaling axis, promotes Drp1 phosphorylation and mitochondrial translocation, induces mitochondrial fission imbalance, structural damage, and mtROS accumulation, ultimately leading to senescence-like changes in myocardial tissue; consistent findings were observed in hypoxic H9c2 cells (in vitro) regarding cardiomyocyte mitochondrial dysfunction.

Fig. 2.

Alterations in mitochondrial dynamics, mitochondrial damage, and related molecular pathways in rat myocardial tissue (in vivo) and hypoxic H9c2 cells (in vitro) models of AMI. Experimental groups (in vivo): Sham, AMI. (A) Western Blot analysis of protein bands for Fis-1 and Mfn-2 in cardiac tissue and visual analysis (n =3). Experimental groups (in vitro): Control, Hypoxia. (B) TEM was used to examine mitochondrial morphology (n =3). (C) Mitochondrial morphology was evaluated using MitoTracker Red CMXRos staining (n =3). (D) Flow cytometry was employed to measure mtROS production in H9c2 cells (n =3). (E) Western Blot analysis of the expression levels of Fis-1 and Mfn-2 in H9c2 cells (n =3). (F) IF was used to detect the co-localization of Drp1 and mitochondria, with quantitative analysis shown on the right (n =3). (G-I) Western blot analysis was performed to detect the protein levels of p-Drp1 (Ser616), p-Drp1 (Ser637), Drp1, p-ERK1/2, ERK1/2, p-Akt, and Akt (n =3). All of the above experiments were biological replicates. ***P < 0.001, vs Sham/Control group

ERK pathway activation promotes Drp1 Ser616 phosphorylation and mitochondrial translocation inducing excessive mitochondrial fission and cardiomyocyte injury

To clarify the regulatory role of the ERK pathway in Drp1-mediated mitochondrial damage, H9c2 cells were treated with the ERK inhibitor U0126 under hypoxic conditions. Western blot analysis confirmed that p-ERK1/2 and p-Drp1 (Ser616) levels were significantly elevated in the Hypoxia group compared with the Control group, whereas U0126 treatment effectively inhibited these changes, suggesting that hypoxia activates Drp1 through the ERK pathway (Fig. 3A-B). At the functional level, U0126 intervention significantly ameliorated hypoxia-induced mitochondrial damage: the mtROS-positive rate decreased from over 70% to 26.29% (Fig. 3C), mitochondrial fragmentation was alleviated with restoration of reticular structure (Fig. 3D), and the proportion of senescent cells was also markedly reduced (Fig. 3E).

Fig. 3.

Regulatory effects of the ERK/Drp1 signaling pathway on mitochondrial function and cellular senescence in hypoxic H9c2 cardiomyocytes. Experimental groups: Control, Hypoxia, Hypoxia + DMSO, and Hypoxia + U0126 (ERK inhibitor). (A-B) Western blot was performed to detect the expression of p-Drp1 (Ser616), Drp1, p-ERK1/2, and ERK1/2 (n =3). (C) Flow cytometry was employed to assess mtROS production (n = 3). (D) Mitochondrial morphology was assessed using MitoTracker Red CMXRos staining (n =3). (E) The SA-β-gal positive rate in H9c2 cells was measured using the SA-β-gal staining kit (n =3). Experimental groups: Hypoxia + Vector, Hypoxia + Drp1, Hypoxia + si-NC, Hypoxia + si-Drp1. (F) TEM was used to examine mitochondrial morphology (n =3). (G) Mitochondrial morphology was determined by MitoTracker Red CMXRos staining (n =3). (H) Flow cytometry was used to measure mtROS production (n =3). (I) Western blot detected the protein expression levels of Fis-1 and Mfn-2, followed by data visualization (n =3). All of the above experiments were biological replicates. **P < 0.01, ***P < 0.001, vs Control/Hypoxia + DMSO/Hypoxia + Vector/Hypoxia + si-NC

On this basis, to further validate the core role of Drp1 as a downstream effector molecule, Drp1 overexpression and knockdown interventions were performed in the H9c2 cell model of hypoxia-simulated AMI (efficiency shown in Fig. S3E-H). Transmission electron microscopy revealed that Drp1 overexpression exacerbated hypoxia-induced mitochondrial swelling, cristae loss, and rupture, whereas Drp1 knockdown reversed these changes (Fig. 3F). Correspondingly, MitoTracker staining confirmed that overexpression promoted fragmentation while knockdown restored reticular structure (Fig. 3G, Fig. S3I). MitoSOX quantification showed that Drp1 overexpression increased mtROS from 70.18% to 84.78%, whereas knockdown decreased it to 25.13% (Fig. 3H, Fig. S3J). Furthermore, Western blot analysis demonstrated that Drp1 overexpression upregulated the fission protein Fis-1 and downregulated the fusion protein Mfn-2, while Drp1 knockdown reversed these trends (Fig. 3I, Fig. S3K). IF further confirmed that Drp1 overexpression enhanced its mitochondrial translocation, whereas knockdown attenuated this process (Fig. S3L-M). Collectively, these findings indicate that Drp1 is a key effector molecule mediating excessive mitochondrial fission and injury under hypoxic conditions.

MT intervention rescues Drp1 overexpression-induced mitochondrial morphological damage and structural disruption

To investigate the rescue effect of MT on Drp1-mediated mitochondrial damage and cellular senescence, a Drp1 overexpression model was established in hypoxic H9c2 cardiomyocytes followed by MT intervention. Transmission electron microscopy revealed that the Hypoxia + Vector group exhibited mitochondrial swelling and morphological abnormalities; Drp1 overexpression (Hypoxia + Drp1 group) further exacerbated mitochondrial fragmentation, with cristae destruction or even loss; whereas combined MT treatment (Hypoxia + Drp1 + MT group) significantly improved mitochondrial morphology, with cristae structure largely restored (Fig. 4A). Notably, an additional MT-alone group was included in Fig. S4A-B to evaluate the basal protective effect of MT. MitoSOX detection showed that the mtROS-positive rate was 70.74% in the Hypoxia + Vector group; Drp1 overexpression increased it to 85.16%; MT alone reduced it to 47.51%; and combined treatment (Drp1 overexpression + MT) lowered it to 76.35%, which remained higher than that in the MT-alone group (Fig. S4A). MitoTracker staining demonstrated that Drp1 overexpression exacerbated mitochondrial fragmentation and length shortening; MT alone restored reticular structure and reduced fragmentation; whereas combined treatment only partially corrected excessive fission, with less improvement than the MT-alone group (Fig. S4B).

Fig. 4.

mtROS scavenging reverses Drp1 overexpression-induced mitochondrial damage and cellular senescence in hypoxic H9c2 cardiomyocytes. Experimental groups: Hypoxia + Vector, Hypoxia + Drp1, Hypoxia + Drp1 + MT. (A) TEM was used to examine mitochondrial morphology. (B) Western blot analysis was carried out to evaluate the protein expression levels of Fis-1 and Mfn-2. (C-D) IF was performed to detect Drp1 co-localization with mitochondria. (E-F) SA-β-gal activity was detected using a SA-β-gal staining kit. (G) Western blot analysis was performed to detect the expression of senescence-related markers p16, p21, p53, and Lamin B1 in cells. (H) The mRNA levels of SASP factors, including IL-1α, IL-1β, IL-8, MCP-1, and ICAM-1, were detected by qRT-PCR. All of the above experiments were biological replicates (n =3). *P < 0.05, **P < 0.01, ***P < 0.001, vs Hypoxia + Vector/Hypoxia + Drp1 group

All other experiments included only the combined MT treatment group (without an MT-alone group). Western blot and qRT-PCR analyses showed that Drp1 overexpression upregulated Fis-1 and downregulated Mfn-2, and combined MT treatment partially reversed these trends (Fig. 4B, Fig. S5A). IF co-localization analysis revealed that Drp1 overexpression significantly enhanced its fluorescence signal in the mitochondrial region, indicating increased mitochondrial localization and fission activity of Drp1; combined MT treatment effectively reversed this phenomenon, markedly reducing the Drp1 signal on mitochondria (Fig. 4C-D). EdU staining showed that Drp1 overexpression inhibited cell proliferation, whereas combined MT treatment restored this capacity (Fig. S5C-D). SA-β-gal staining confirmed that Drp1 overexpression promoted cellular senescence, while combined MT treatment reduced the proportion of senescent cells (Fig. 4E-F). Furthermore, combined MT treatment also suppressed the upregulation of senescence-associated genes (p16, p21, p53) and inflammatory factors (IL-1α, IL-1β, IL-8, MCP-1, ICAM-1), as well as reversed the downregulation of Lamin B1 induced by Drp1 overexpression (Fig. 4G-H, Fig. S5B). In summary, Drp1 overexpression exacerbates hypoxia-induced mitochondrial damage, oxidative stress, and cellular senescence, whereas MT intervention effectively reverses these changes and restores mitochondrial functional homeostasis.

HIF-1α regulates Drp1-mediated mitochondrial dysfunction

In the H9c2 cell hypoxia model, the knockdown of HIF-1α had a significant impact on Drp1-regulated mitochondrial function. Western blot results showed that HIF-1α protein levels were markedly increased under hypoxic conditions (Fig. 5A; Fig. S6A-B). ChIP assays revealed that under hypoxic conditions, HIF-1α binding to the target gene promoter was significantly enhanced (Fig. 5B). Dual-luciferase reporter assay demonstrated that DRP1 promoter activity was markedly increased, whereas empty vector activity showed no significant change (Fig. 5C). These results indicate that hypoxia promotes HIF-1α binding and activation of the DRP1 promoter, reflecting its hypoxia-responsive mechanism. Functionally, by transfecting si-NC or si-HIF-1α (Fig. S6C-D), relevant parameters were examined under normoxic and hypoxic conditions. MitoSOX staining showed that under normoxia, HIF-1α knockdown alone did not affect baseline mtROS levels; hypoxic treatment significantly increased the mtROS-positive rate to 70.82%, whereas HIF-1α knockdown dramatically reduced it to 20.61%, indicating that hypoxia is a key trigger of mitochondrial oxidative damage and that HIF-1α knockdown significantly alleviates excessive mtROS production (Fig. 5D-E). IF co-localization further revealed that under normoxia, HIF-1α knockdown led to reduced Drp1 expression and mitochondrial co-localization; under hypoxia, Drp1 fluorescence intensity was significantly upregulated with substantial translocation to mitochondria and obvious mitochondrial fragmentation; whereas HIF-1α knockdown significantly decreased Drp1 fluorescence intensity and mitochondrial co-localization, ameliorated punctate fragmentation, and restored reticular fusion structure (Fig. 5F-G). To validate the causal relationship of the HIF-1α/Drp1 axis, Drp1 was overexpressed as a rescue experiment in HIF-1α-knockdown hypoxic cells, with appropriate controls (normoxia, hypoxia alone, negative control, and empty vector control). MitoSOX flow cytometry showed that the mtROS-positive rate was significantly elevated in the hypoxia alone group, was dramatically reduced to 18.39% by HIF-1α knockdown, and rebounded to 46.78% upon Drp1 rescue, reversing the protective effect of HIF-1α knockdown (Fig. 5H). Western blot demonstrated that hypoxia upregulated HIF-1α expression and promoted Drp1 Ser616 phosphorylation; HIF-1α knockdown reduced the p-Drp1 (Ser616)/Drp1 ratio; whereas Drp1 rescue directly upregulated the p-Drp1 (Ser616)/Drp1 ratio without restoring HIF-1α, reactivating the mitochondrial fission program (Fig. 5I). Mitochondrial dynamics protein analysis showed that under hypoxic conditions, HIF-1α knockdown downregulated the fission proteins Drp1 and Fis-1 and upregulated the fusion protein Mfn-2; Drp1 rescue subsequently increased fission proteins and decreased fusion proteins, recapitulating the dysregulated dynamics phenotype (Fig. 5J). IF co-localization confirmed that hypoxia promoted Drp1 translocation to mitochondria and fragmentation; HIF-1α knockdown reduced Drp1 recruitment and restored reticular structure; Drp1 rescue increased co-localization intensity again, fully reversing the morphological protection (Fig. 5K). In summary, HIF-1α is a key upstream regulator of Drp1 activation and mitochondrial injury in hypoxic cardiomyocytes; inhibition of HIF-1α alleviates mitochondrial dysfunction in a Drp1-dependent manner, whereas Drp1 overexpression reverses this protective effect, establishing the central role of the HIF-1α/Drp1 axis in hypoxic cardiomyocyte mitochondrial injury.

Fig. 5.

HIF-1α regulates Drp1 Ser616 phosphorylation-mediated mitochondrial dysfunction in hypoxic H9c2 cardiomyocytes. Experimental groups: Control, Hypoxia. (A) Western blot detection of HIF-1α protein expression bands in H9c2 cells. (B) CHIP assay was conducted to detect the interaction between HIF-α and Drp1. (C) Dual-luciferase assay was performed to detect the interaction between HIF-α and Drp1. Experimental groups: Control + si-NC, Control + si-HIF-1α, Hypoxia + si-NC, Hypoxia + si-HIF-1α. (D-E) Flow cytometry was used to measure mtROS production in H9c2 cells. (F-G) IF was used to detect the colocalization of Drp1 and mitochondria. (H) Flow cytometry was used to detect mtROS production in H9c2 cells after four different transfection treatments, along with visual analysis. Experimental groups: Control, Hypoxia, Hypoxia + si-NC, Hypoxia + si-HIF-1α, Hypoxia + si-HIF-1α + Vector, Hypoxia + si-HIF-1α + Drp1. (H) mtROS levels were detected by flow cytometry using MitoSOX dye. (I) Western blot detected the protein expression of Drp1, p-Drp1 (Ser616), and HIF-1α. (J) Western blot detected the protein expression of Fis-1 and Mfn-2. (K) IF was used to detect the colocalization of Drp1 and mitochondria. All of the above experiments were biological replicates (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, vs Control/IgG/Control + si-NC/Hypoxia + si-NC/Hypoxia + si-HIF-1α + Vector group; ns means no significant difference vs Control/Hypoxia + si-HIF-1α + Vector group

Ruxolitinib inhibits the SASP pathway to alleviate hypoxic cardiomyocyte injury and Drp1 is a downstream key molecule

To investigate the protective effect of the SASP pathway inhibitor Ruxolitinib on hypoxia-induced cardiomyocyte injury, H9c2 cells were treated with Ruxolitinib in a hypoxia model, and its effects on cellular senescence and apoptosis were observed. The experiment included four groups: Control, Hypoxia, Hypoxia + DMSO, and Hypoxia + Ruxolitinib. The results showed that Ruxolitinib significantly inhibited the hypoxia-induced increase in the p-Drp1(Ser616)/Drp1 ratio (Fig. S7A), restored cell proliferation capacity (Fig. S7B-C), reduced the percentage of SA-β-gal-positive cells (Fig. 6A), reversed the abnormal expression of p16, p21, p53, and Lamin B1 (Fig. 6B-C), inhibited apoptosis (Fig. 6D-E), downregulated Bax and upregulated Bcl-2 expression (Fig. 6F-G).

Fig. 6.

Effects of targeted inhibition of the SASP pathway or modulation of Drp1 expression on senescence, apoptosis, and inflammatory response in hypoxic H9c2 cardiomyocytes. Experimental groups: Control, Hypoxia, Hypoxia + DMSO, Hypoxia + Ruxolitinib. (A) SA-β-gal staining was performed using a SA-β-gal staining kit. (B-C) qRT-PCR and Western blot were used to detect the mRNA and protein expression levels of senescence-related markers p16, p21, p53, and Lamin B1 in cells. (D-E) Annexin V/PI double staining was performed to detect cell apoptosis. (F-G) qRT-PCR and Western blot were used to detect the mRNA and protein expression levels of Bax and Bcl-2. Experimental groups: Hypoxia + Vector, Hypoxia + Drp1, Hypoxia + si-NC, Hypoxia + si-Drp1. (H-I) SA-β-gal staining was performed using a SA-β-gal staining kit. (J-L) qRT-PCR and western blot were used to detect the mRNA and protein expression levels of p16, p21, p53, and Lamin B1. (M) qRT-PCR detected the mRNA levels of SASP markers, including IL-1α, IL-1β, IL-8, MCP-1, and ICAM-1, in H9c2 cells. All of the above experiments were biological replicates (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, vs Hypoxia/Hypoxia + Vector/Hypoxia + si-NC group

To clarify the key role of Drp1 in H9c2 cell senescence and apoptosis, Drp1 overexpression and knockdown experiments were further performed. EdU assay showed that Drp1 overexpression significantly inhibited cell proliferation, while Drp1 knockdown enhanced proliferative capacity (Fig. S7D-E), suggesting that Drp1 negatively regulates cardiomyocyte proliferation. SA-β-gal staining indicated that Drp1 overexpression significantly increased the proportion of senescent cells, whereas Drp1 knockdown reduced the number of senescent cells (Fig. 6H-I), indicating that Drp1 promotes hypoxia-induced cardiomyocyte senescence. qRT-PCR and Western blot further confirmed that Drp1 overexpression upregulated the senescence-promoting factors p16, p21, and p53 and downregulated the anti-aging factor Lamin B1, while Drp1 knockdown reversed these changes (Fig. 6J-L), verifying the pro-senescence role of Drp1 at the molecular level. qRT-PCR analysis showed that Drp1 overexpression promoted the transcriptional activation of SASP-related inflammatory factors IL-1α, IL-1β, IL-8, MCP-1, and ICAM-1, whereas knockdown inhibited their expression (Fig. 6M), indicating that Drp1 induces a senescence-associated inflammatory microenvironment. Furthermore, Drp1 overexpression upregulated the pro-apoptotic protein Bax and downregulated the anti-apoptotic protein Bcl-2, while Drp1 knockdown had the opposite effect (Fig. S7F), suggesting that Drp1 also promotes apoptosis. Western blot results showed that the cle-Caspase-1/Caspase-1 and GSDMD-N/GSDMD ratios were significantly increased in the Drp1 overexpression group, while these ratios were markedly decreased in the Drp1 knockdown group, indicating that Drp1 positively regulates the hypoxia-induced activation of the Caspase-1/GSDMD pyroptosis pathway (Fig. S7G). In conclusion, Ruxolitinib alleviates hypoxia-induced senescence and apoptosis in H9c2 cells and restores cell proliferation capacity by inhibiting the SASP pathway; Drp1 positively regulates cardiomyocyte senescence, apoptosis, pyroptosis, and SASP-related inflammation, serving as a key downstream effector molecule of the SASP pathway.

Inhibition of Drp1 with Mdivi-1 attenuates mitochondrial dysfunction, senescence-like changes in myocardial tissue, and SASP-related inflammation, conferring cardioprotection in AMI rats

To investigate the protective effect of targeted inhibition of Drp1 on myocardial injury induced by AMI, an in vivo AMI model was established in SD rats by left anterior descending coronary artery ligation, with sham-operated (Sham) rats as controls, and prophylactic administration of either DMSO vehicle or the Drp1-specific inhibitor Mdivi-1. The experiment consisted of six groups: Sham, Sham + DMSO, Sham + Mdivi-1, AMI, AMI + DMSO, and AMI + Mdivi-1. qRT‑PCR and Western blot results showed that, compared with the Sham group, the AMI group exhibited a significant upregulation of the mitochondrial fission marker Fis-1 and a marked downregulation of the fusion marker Mfn-2 in myocardial tissue; Mdivi-1 intervention effectively reversed this imbalance and corrected mitochondrial dynamics disorder. In the Sham groups, Mdivi-1 had no significant effect on the expression of Fis-1 and Mfn-2, and the DMSO vehicle showed no interference (Fig. 7A‑B).

Fig. 7.

Prophylactic administration of Mdivi-1 alleviates senescence-like changes in myocardial tissue, inflammation, and cardiac dysfunction in AMI rats by restoring mitochondrial dynamics. Experimental groups: Sham, Sham + DMSO, Sham + Mdivi-1, AMI, AMI + DMSO, AMI + Mdivi-1. (A) The mRNA levels of Fis-1 and Mfn-2 in cardiac tissue were quantified using qRT-PCR (n = 3). (B) Western blot was applied to assess the protein expression of Fis-1 and Mfn-2 in cardiac tissue (n =3). Experimental groups: Sham + DMSO, Sham + Mdivi-1, AMI + DMSO, AMI + Mdivi-1. (C) γ-H2AX IF was utilized to determine the positive rate of the DNA damage marker γ-H2AX (n =3). (D-E) qRT-PCR and Western blot were performed to determine the mRNA and protein expression of p16, p21, p53, and Lamin B1, respectively (n=3). (F) qRT-PCR was employed to detect the mRNA levels of SASP (n =3). (G-H) TTC staining was used to evaluate the myocardial infarction region and the relative infarct size (n =6). (I-J) Echocardiography and quantitative analysis of four key cardiac function parameters were performed (n =6). All of the above experiments were biological replicates. *P < 0.05, **P < 0.01, ***P < 0.001, vs Sham/Sham + DMSO/AMI + DMSO group; ns means no significant difference vs Sham/Sham + DMSO/AMI group

Further analysis within the four groups (Sham + DMSO, Sham + Mdivi-1, AMI + DMSO, AMI + Mdivi-1) revealed that: in the Sham + Mdivi-1 group, Drp1 mRNA and the p‑Drp1 (Ser616)/Drp1 ratio were significantly downregulated; in the AMI + DMSO group, both were extremely significantly elevated; and the AMI + Mdivi-1 group significantly inhibited these elevations (Fig. S8A‑B). ERK pathway detection showed that the p‑Drp1/Drp1 ratio was significantly reduced in the Sham + Mdivi-1 group, while the p‑ERK/Drp1 ratio was extremely significantly upregulated in the AMI + DMSO group, and Mdivi-1 blocked this phosphorylation (Fig. S8C‑D). IF co-localization confirmed that p‑Drp1 translocated extensively to mitochondria in the AMI + DMSO group, whereas this translocation was significantly reduced in the AMI + Mdivi-1 group (Fig. S8E).

Based on these findings, the effects of Drp1 inhibition on post-AMI myocardial injury, inflammation, fibrosis, and cardiac function were further evaluated. First, under the Sham background, Mdivi-1 had no significant effect on any of the following measured parameters (Fig. 7C‑J, Fig. S8F‑J). γ‑H2AX IF showed severe DNA damage in the AMI + DMSO group, and the fluorescence signal was significantly decreased after Mdivi-1 treatment (Fig. 7C). Detection of senescence-like changes in myocardial tissue indicated that the mRNA and protein levels of p16, p21, and p53 were significantly elevated in the AMI + DMSO group, while Lamin B1 was significantly decreased; Mdivi-1 reversed these changes (Fig. 7D‑E). qRT‑PCR results for SASP-related inflammatory factors (IL‑1α, IL‑1β, IL‑8, MCP‑1, ICAM‑1) showed that all these factors were extremely significantly upregulated in the AMI + DMSO group, and Mdivi-1 intervention significantly suppressed them (Fig. 7F). TTC staining showed a significant increase in myocardial infarct size in the AMI + DMSO group, and Mdivi-1 treatment significantly reduced the infarct size (Fig. 7G‑H). H&E staining showed severe disorganization of myocardial structure, accompanied by edema, hemorrhage, and inflammatory infiltration in the AMI + DMSO group, which were markedly improved after Mdivi-1 treatment (Fig. S8F). Masson staining showed a very significant increase in myocardial fibrosis area in the AMI + DMSO group, and Mdivi-1 significantly reduced collagen deposition (Fig. S8G‑H). Echocardiography revealed that in the AMI + DMSO group, LVIDd and LVIDs were significantly increased, while EF% and left ventricular FS% were greatly decreased; after Mdivi-1 treatment, ventricular dilation was alleviated, and cardiac function parameters significantly recovered (Fig. 7I‑J). Short-axis echocardiography further validated these results (Fig. S8I‑J). In summary, in this experimental model, prophylactic administration of Mdivi-1, by inhibiting Drp1, comprehensively reversed AMI-induced mitochondrial fission-fusion imbalance, DNA damage, senescence-like changes in myocardial tissue, and SASP-related inflammatory factor release, leading to reduced infarct size and improved cardiac function. These findings demonstrate that Mdivi-1 confers myocardial protection in this preventive intervention setting.

Discussion

This study systematically explored the role of Drp1 in AMI and its potential mechanisms for myocardial protection through its inhibition. By establishing rat AMI models and H9c2 cardiomyocyte hypoxia models, we observed that Drp1 was significantly upregulated during AMI, which promoted mitochondrial fission and subsequently led to mitochondrial dysfunction, senescence-like changes in myocardial tissue (in vivo) / cardiomyocyte senescence (in vitro), and myocardial injury. Prophylactic administration of Mdivi-1 not only improved mitochondrial morphology and function but also significantly reduced myocardial cell senescence-like changes in myocardial tissue and apoptosis, ultimately enhancing cardiac function. The findings suggested that Drp1 could be a potential therapeutic target for AMI treatment and warranted further exploration.

In the AMI model, we observed a significant upregulation of Drp1 expression in myocardial tissue, which is consistent with previous studies [32–34]. Echocardiography results indicated that, compared to the Sham group, the AMI group exhibited significantly reduced EF and FS, along with significantly increased LVIDd and LVIDs, indicating impaired cardiac contractile function. Additionally, levels of CK-MB, LDH, and CPK were significantly elevated in the AMI model, reflecting myocardialinjury and death. Further H&E staining, Masson staining, and TTC staining results indicated that the pathological response, fibrosis, and infarct area in the AMI group were significantly increased, confirming irreversible myocardial structural damage.

Studies have shown that Fis-1, as a key receptor for mitochondrial fission, binds to activated Drp1, facilitating its localization to the outer mitochondrial membrane, thereby inducing excessive fission and disrupting mitochondrial dynamic balance [35]. Additionally, overactivation of Drp1 inhibits Mfn-2-mediated mitochondrial fusion, further compromising the integrity of the mitochondrial network [36]. In the DNA damage response, increased deposition of γH2Ax and p53-binding protein 1 (p53BP1) on chromatin, as well as the activation of ataxia telangiectasia mutated (ATM) kinase, which phosphorylates and activates p53, have been observed [37]. In our study, Drp1 significantly promoted mitochondrial fission in both the AMI model and hypoxia-treated H9c2 cells, leading to mitochondrial fragmentation and swelling, consistent with Drp1’s critical role in regulating mitochondrial dynamics [38, 39]. Furthermore, elevated levels of Fis-1 and reduced levels of Mfn-2 were detected in the AMI model, indicating an imbalance between mitochondrial fusion and fission. TEM, MitoTracker Red staining, and mtROS flow cytometry demonstrated that Drp1 activity directly influenced mitochondrial oxidative stress, with mtROS levels significantly increasing under hypoxic conditions. γ-H2AX IF revealed aggravated DNA damage in the AMI model. Flow cytometry indicated that apoptosis levels in H9c2 cells were significantly increased after hypoxic treatment, which was further confirmed by the detection of Bax and Bcl-2 protein and mRNA levels. These results suggested that Drp1 promoted mitochondrial fission and ROS production, which may have contributed to enhanced oxidative stress, DNA damage, and apoptosis in myocardial cells [40, 41].

Regarding the upstream regulatory mechanism of Drp1 activation, this study found that both ERK1/2 and Akt signaling pathways were significantly activated under hypoxic conditions, and the ERK inhibitor U0126 effectively inhibited hypoxia-induced elevation of p-Drp1(Ser616), confirming that ERK is a key upstream kinase regulating Drp1 Ser616 phosphorylation, which is consistent with previous reports [42, 43]. Furthermore, HIF-1α was also confirmed to regulate Drp1 Ser616 phosphorylation [44, 45]. These findings provide new experimental evidence for understanding the upstream regulatory mechanism of Drp1 activation under myocardial ischemic conditions.

p21, a member of the Cip/Kip family of cyclin-dependent kinase (CDK) inhibitors, plays a key role in cell cycle inhibition, senescence induction, tumor suppression, and regulation, with broad biological activity [46]. Upon binding to CDK, it forms a complex with PCNA, inhibiting PCNA-dependent DNA synthesis and causing cell cycle arrest. Similarly, p16, another CDK inhibitor, induces cellular senescence and growth arrest [47]. p53, an upstream effector of p21, mediates apoptosis by promoting BAX expression and inhibiting BCL-2 expression [48]. In studies on vascular aging and atherosclerosis, Drp1-mediated mitochondrial fission has been shown to induce oxidative stress and mitochondrial dysfunction, activating the p53 signaling pathway. In turn, p53 activation upregulates p16 and p21 expression, leading to cell cycle arrest and accelerating endothelial cell senescence [49]. In our study, Drp1 overexpression in the hypoxia-treated H9c2 cell model significantly exacerbated mitochondrial dysfunction, resulting in structural damage and increased ROS levels. IF further confirmed increased co-localization of Drp1 with mitochondria, indicating elevated cellular stress and Drp1’s involvement in mitochondrial fission. Conversely, Drp1 knockdown alleviated ROS generation and structural damage in mitochondria. Western blot and qRT-PCR analyses revealed that Drp1 regulates the expression of Fis-1 and Mfn-2, affecting mitochondrial fission and fusion processes. Additionally, Drp1 overexpression inhibited cell proliferation, upregulated senescence-associated genes (p16, p21, p53), and increased the expression of SA-β-gal, while Lamin B1 expression was downregulated. Drp1 knockdown reversed these effects. Furthermore, treatment with MT [50] effectively mitigated Drp1-induced cellular damage. The results of this study suggested that Drp1 may have been involved in the process of cardiomyocyte senescence by regulating mitochondrial function.

Cellular senescence and inflammation frequently co-occur and mutually reinforce one another [51, 52]. During senescence, senescent cells secrete significant quantities of pro-inflammatory factors, such as IL-1α, IL-1β, IL-6, and MCP-1, through the SASP, further exacerbating local nervous system inflammation and acute lung injury. Concurrently, inflammation accelerates mitochondrial dysfunction and cellular senescence by activating the Drp1-mediated mitochondrial fission pathway. DRP1 is a key regulator of mitochondrial fission, and its dysregulation can disrupt mitochondrial homeostasis and promote neuroinflammation, thereby exacerbating disease severity [53, 54]. In our study, hypoxia and Drp1 overexpression together led to increased expression of inflammatory factors, including IL-1α, IL-1β, IL-8, MCP-1, and ICAM-1. This inflammatory response was reversed by treatment with MT, suggesting that MT can suppress Drp1-induced cellular inflammation. Notably, this study also found that Drp1 overexpression significantly activated the Caspase-1/GSDMD pyroptosis pathway. Pyroptosis, as a pro-inflammatory form of programmed cell death, whose activation further exacerbates inflammatory responses [55]. Studies have shown that Drp1-mediated mitochondrial fission promotes inflammasome activation and Caspase-1 cleavage, thereby driving inflammatory responses [56]. Therefore, pyroptosis activation induced by Drp1 overexpression not only directly leads to cell death but may also amplify the local inflammatory microenvironment through the release of inflammatory factors, suggesting that Drp1 participates in cardiomyocyte injury and inflammatory regulation through both apoptotic and pyroptotic pathways.

HIF-1α is a transcription factor and a core component of the oxygen-sensing mechanism in mammalian cells. It is a crucial regulator in hypoxic and ischemic responses [57]. Under normoxic conditions, HIF-1α is degraded by prolyl hydroxylase enzymes, but during hypoxia, its degradation is inhibited, leading to its accumulation. HIF-1α translocates to the nucleus and forms a dimeric complex with HIF-1β, which binds to hypoxia-responsive genes and regulates downstream targets. HIF-1α plays a critical role in metabolic processes and has been shown to provide cardioprotection during ischemia/reperfusion injury (I/RI) by maintaining mitochondrial dynamic balance [58]. Under hypoxic conditions, HIF-1α is elevated in the portal hypertensive gastric mucosa and gastric cancer tissues of both humans and mice. It mediates mitochondrial dysfunction by upregulating METTL3/IGF2BP3-dependent Drp1 N6-methyladenosine modification, leading to increased mtROS production [59]. In addition, studies have shown that HIF-1α regulates the expression of various genes, such as VEGF, EPO, etc., by binding to the hypoxia response element (HRE) [60, 61]. On the other hand, Drp1, as an important regulator of mitochondrial dynamics, has been found to have significant changes in its expression and activity under various stress conditions [62–64]. However, there is currently no direct evidence indicating that HIF-1α can bind directly to the promoter region of Drp1 and regulate its expression. Nevertheless, based on existing literature, it is speculated that the Drp1 promoter region may contain potential HRE sites, allowing HIF-1α to bind and promote the transcriptional expression of Drp1 [64–66]. This study reveals the role of HIF-1α in Drp1-regulated mitochondrial function. We found that under hypoxic conditions, HIF-1α may promote the expression of Drp1 to some extent by enhancing its binding to the DRP1 promoter and activating its promoter activity. In addition, knockdown of HIF-1α significantly reduced Drp1 and Fis-1 expression, increased Mfn-2 levels, decreased mtROS production, and reduced Drp1 co-localization with mitochondria.

Although our study did not directly assess the impact of changes in ROS levels on the binding of HIF-1α to the Drp1 promoter, under hypoxic or stressful conditions, elevated ROS levels may be a key factor in promoting the stability and activity of HIF-1α. Previous studies have shown that ROS can significantly enhance the stability of HIF-1α [67]. In conjunction with our experimental results, we speculate that ROS may indirectly enhance the binding of HIF-1α to the Drp1 promoter and its regulatory function by upregulating HIF-1α activity. Specifically, under hypoxic conditions, the binding of HIF-1α to the Drp1 promoter was significantly higher than that in the normoxic control group, further suggesting that ROS may play an important role in this process. However, there is currently a lack of direct evidence to verify the specific role of ROS in this process, and future in-depth and comprehensive studies are needed to validate this hypothesis. In our study, HIF-1α regulated Drp1, which in turn affected mitochondrial dynamics, including the generation and clearance of mtROS, thereby promoting the survival and function of cardiomyocytes under hypoxic conditions. It is noteworthy that the overexpression of Drp1 partially reversed the protective effect caused by HIF-1α knockdown, indicating that Drp1 played a key regulatory role in HIF-1α-mediated cardioprotection. Therefore, HIF-1α and Drp1 jointly regulated mitochondrial dynamics to enhance cardiomyocyte adaptation to hypoxic stress.

SASP is a key marker of cellular senescence [68]. Ruxolitinib is a classical JAK/STAT signaling pathway inhibitor that alleviates cellular senescence by inhibiting SASP secretion [69, 70]. In this study, we found that Ruxolitinib treatment significantly reduced senescence and apoptosis of H9c2 cells, restored cell proliferation capacity, and inhibited hypoxia-induced p-Drp1(Ser616) phosphorylation, suggesting that the SASP pathway may act upstream of Drp1 activation and that the cardioprotective effect of Ruxolitinib is partially mediated through inhibiting Drp1 activation. Furthermore, Drp1 overexpression exacerbated cellular senescence and apoptosis, whereas Drp1 knockdown showed opposite effects, further supporting the key role of Drp1 in cardiomyocyte senescence. In summary, Ruxolitinib, as a SASP inhibitor, effectively alleviates cardiac cellular senescence and exhibits potential cardioprotective effects.

Mdivi-1, a derivative of quinazolinone that inhibits Drp1-dependent fission, promotes mitochondria, and reduces damage [71, 72]. In the AMI model of this study, prophylactic administration of Mdivi-1 (given 15 min prior to AMI induction) not only restored the balance of mitochondrial fission-fusion (downregulating Fis-1, upregulating Mfn-2), but also significantly improved mitochondrial function and alleviated oxidative stress-related DNA damage. These results are consistent with previous reports [73]. More importantly, this study found that Mdivi-1 effectively inhibited senescence-like changes in myocardial tissue (downregulating p16/p21/p53, upregulating Lamin B1) and the expression of SASP-related inflammatory factors. This finding provides new experimental evidence that Mdivi-1 exerts cardioprotective effects in this preventive intervention setting by inhibiting senescence-like changes in myocardial tissue, complementing the mechanistic understanding of its role in post-AMI ventricular remodeling. At the functional level, Mdivi-1 treatment significantly reduced myocardial fibrosis, infarct size, and improved cardiac function, indicating that inhibiting the Drp1-mediated senescence pathway confers long-term prognostic benefits in this model. Mechanistically, we demonstrated that Mdivi-1 blocked AMI-induced p-Drp1 (Ser616) phosphorylation and upstream p-ERK activation, further clarifying the key role of the ERK/Drp1 signaling axis in mediating myocardial injury and senescence-like changes in myocardial tissue. In summary, this study demonstrates that prophylactic Mdivi-1 administration provides myocardial protection in the AMI rat model, likely through inhibition of the ERK/Drp1 pathway-mediated mitochondrial fission and senescence-like changes in myocardial tissue. These findings offer experimental support for the potential cardioprotective application of Mdivi-1 in a preventive or very early intervention context, while further studies are needed to assess its therapeutic potential in a post-AMI setting.

Despite the important progress made in this study, several limitations should be acknowledged. First, the in vivo experiments were based on whole myocardial tissue analysis, which cannot precisely distinguish the cellular origin of senescence signals. We acknowledge that fibroblasts and infiltrating immune cells may also contribute to the observed senescence phenotype. Future studies utilizing cell-specific transgenic animals or single-cell technologies are warranted to further validate the primary cellular sources of senescence. Second, although Mdivi-1 is a widely used Drp1 inhibitor and the in vitro knockdown and overexpression experiments support its role in Drp1 regulation, Mdivi-1 may still have off-target effects to some extent. Moreover, the lack of in vivo genetic loss-of-function or gain-of-function studies (e.g., cardiomyocyte-specific Drp1 knockout or overexpression) limits our ability to establish definitive causal relationships. Future studies should incorporate genetic approaches to further validate the specific role of Drp1. Third, this study is based on a rat model and the H9c2 cell line; clinical translation remains to be validated. Fourth, this study lacks direct in vivo evidence of mitochondrial structural remodeling (e.g., three-dimensional reconstruction or high-resolution imaging by electron microscopy), and currently relies mainly on functional assays to indirectly infer mitochondrial dynamics. Although the functional data provide important clues, the introduction of orthogonal structural validation approaches (e.g., in vivo mitochondrial ultrastructural analysis) would help to more comprehensively support the causal relationship between mitochondrial remodeling and functional changes. Future studies are also needed to explore additional molecular mechanisms, develop safer Drp1 inhibitors, and investigate their effects at different stages of AMI.

In summary, this study validated the role of Drp1-mediated excessive mitochondrial fission in myocardial injury in a permanent ligation AMI model and integrated multidimensional phenotypes including senescence-like changes in myocardial tissue (in vivo) / cardiomyocyte senescence (in vitro), SASP-related inflammation, and pyroptosis. These findings provide experimental evidence for understanding the molecular mechanisms of post-AMI ventricular remodeling from the novel perspective of cellular senescence and inflammatory regulation, and offer theoretical support for cardioprotective strategies targeting mitochondrial dynamics in preventive or early intervention settings.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

AMI

Acute Myocardial Infarction

ATM

Ataxia telangiectasia mutated

ChIP

Chromatin Immunoprecipitation

Drp1

Dynamin-related protein 1

EdU

5-Ethynyl-2’-deoxyuridine

ELISA

Enzyme-linked immunosorbent assay

FBS

Fetal bovine serum

HIF-1α

Hypoxia-Inducible Factor-1 Alpha

H&E

Hematoxylin and Eosin

HRE

Hypoxia response element

IHC

Immunohistochemistry

IF

Immunofluorescence

LAD

Left anterior descending coronary artery

LVEF

Left ventricular ejection fraction

LVFS

Left ventricular fractional shortening

LVIDd

Left ventricular internal diameter in diastole

LVIDs

Left ventricular internal diameter in systole

Mdivi-1

Mitochondrial Division Inhibitor 1

mtROS

Mitochondrial Reactive Oxygen Species

MT

Mito-TEMPO

p53BP1

p53-binding protein 1

qRT-PCR

Real-Time Quantitative PCR

ROS

Reactive oxygen species

SASP

Senescence-associated secretory phenotype

SA-β-gal

Senescence-Associated β-Galactosidase

SEM

Standard error of the mean

TTC

2,3,5-Triphenyl Tetrazolium Chloride

Author contributions

Peng Yang: Conceptualization, Data curation, Formal analysis, Investigation, Writing-original draft, Writing-review & editing. Yiheng Yang: Conceptualization, Data curation, Investigation, Writing-original draft, Writing-review & editing. Xiao Huang: Conceptualization, Investigation, Formal analysis, Methodology, Writing-original draft, Writing-review & editing. Hongmin Zhu: Formal analysis, Software, Methodology, Writing-review & editing. Yuliang Zhan: Methodology, Writing-review & editing. Xinyong Cai: Software, Writing-review & editing. Zhenzhong Zheng: Project administration, Supervision, Writing-review & editing. Qingshan Tian: Project administration, Resources, Funding, Supervision, Writing-review & editing.

Funding

This work was supported by grants from the National Natural Science Foundation of China (82260383), the Key Research and Development Program of Jiangxi Province (20232BBG70018), the 2022 Senior Science and Technology Innovation Talent Program of Nanchang City (2022-321-13), and the Science and Technology Program of Jiangxi Provincial Administration of Traditional Chinese Medicine (2024B0183).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethical approval

This study was approved by the Institutional Animal Care and Use Committee of The First Affiliated Hospital, Jiangxi Medical College, Nanchang University (IACUC Issue No: 202311QR015).

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.