MYC promotes myocardial fibrosis via METTL1-mediated m7G modification of HILPDA

Abstract

Background

Post-infarction myocardial fibrosis is a pivotal pathological process leading to heart failure; however, its epitranscriptional regulatory mechanisms remain poorly defined. The role of the MYC-METTL1-HILPDA axis in this process remains unexplored.

Methods

Myocardial infarction (MI) models were established in mice, and hypoxia-induced mouse cardiac fibroblasts were used. A range of molecular techniques, including qRT-PCR, Western blot, immunohistochemistry, RNA immunoprecipitation, chromatin immunoprecipitation, and dual-luciferase reporter assays, were employed to investigate the MYC-METTL1-HILPDA axis.

Results

Following MI, METTL1 and HILPDA were significantly upregulated in cardiac tissue. METTL1 stabilized HILPDA mRNA via m7G modification, thereby enhancing its protein expression. Functional studies demonstrated that HILPDA overexpression induced mitochondrial dysfunction and fibroblast activation, whereas HILPDA knockdown attenuated these effects. Furthermore, the transcription factor MYC was identified as an upstream regulator that directly binds the METTL1 promoter to activate its transcription. Crucially, HILPDA knockdown improved cardiac function, attenuated fibrosis, and reduced infarct size in mice.

Conclusion

This study identifies the MYC-METTL1-HILPDA axis as a novel driver of post-infarction myocardial fibrosis, which promotes mitochondrial dysfunction and fibroblast activation through m7G-mediated stabilization of HILPDA mRNA. These findings provide new mechanistic insights and reveal potential therapeutic targets for preventing heart failure.

Graphical Abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s13062-026-00828-x.

Introduction

Myocardial infarction (MI) is a leading cause of cardiovascular disability and death worldwide [1]. Its pathophysiological process encompasses multiple critical stages, including cardiomyocyte death, inflammatory responses, and myocardial fibrosis [2]. As a core pathological feature of cardiac remodeling following MI, myocardial fibrosis is characterized by abnormal activation, excessive proliferation of cardiac fibroblasts (CFs), and deposition of extracellular matrix, ultimately leading to severe impairment of cardiac structure and function [3]. Although classical signaling pathways such as transforming growth factor-β 1 (TGF-β1) have been implicated in fibrosis regulation [4], the molecular mechanisms driving persistent post-MI fibrosis, particularly at the post-transcriptional level, remain poorly defined. Therefore, elucidating the molecular mechanisms underlying myocardial fibrosis and identifying novel therapeutic targets hold significant scientific and clinical value.

In recent years, epigenetic modifications in cardiovascular diseases have garnered increasing attention [5]. Among these, N7-methylguanosine (m7G), a common methylation modification in messenger RNA, plays a crucial role in regulating RNA stability, localization, and translation efficiency [6]. METTL1, a key m7G methyltransferase, forms a complex with WDR4 to catalyze m7G modification of specific mRNAs, thereby influencing their translation [7]. Studies indicate that METTL1 expression is significantly upregulated in myocardial ischemia-reperfusion models, where it participates in regulating cardiomyocyte apoptosis through m7G modification of specific mRNAs [8]. Furthermore, METTL1 has been reported to regulate cardiac fibroblast proliferation and myofibroblast transdifferentiation, suggesting its potential role in myocardial fibrosis [9]. However, the specific downstream target genes through which METTL1 regulates fibroblast activation under MI conditions remain unidentified.

Metabolic dysfunction has been recognized as a hallmark feature of fibrotic diseases [10]. Hypoxia-inducible lipid droplet-associated protein (HILPDA), a key regulator of intracellular lipid metabolism, promotes lipid accumulation by inhibiting adipose triglyceride lipase (ATGL), thereby exacerbating lipid peroxidation and mitochondrial dysfunction [11, 12]. Mitochondrial dysfunction has been established as a key driver of post-infarction myocardial fibrosis [13]. Following MI, ischemic stress induces excessive mitochondrial reactive oxygen species production, loss of membrane potential, impaired ATP synthesis, and dysregulated mitochondrial dynamics [14]. This persistent mitochondrial dysfunction activates pro-fibrotic signaling pathways such as TGF-β and MAPK, directly promoting cardiac fibroblast-to-myofibroblast transdifferentiation, proliferation, and extracellular matrix deposition, thereby accelerating myocardial fibrosis and ventricular remodeling [15]. Increasing evidence indicates that HILPDA may promote fibrosis through multiple mechanisms, including inducing mitochondrial dysfunction and upregulating α-smooth muscle actin (α-SMA) and type I collagen expression [16, 17]. HILPDA is significantly upregulated in both kidney and liver diseases [18, 19]. However, its specific role and molecular mechanisms in myocardial fibrosis remain incompletely understood. Notably, bioinformatics analysis revealed potential m7G modification sites within the HILPDA mRNA sequence, and HILPDA expression in cardiac tissue showed a significant positive correlation with METTL1. Therefore, we hypothesize that HILPDA is a critical downstream effector of METTL1, mediating the pro-fibrotic effects of m7G modification on fibroblasts.

Regarding the upstream regulation of METTL1, the mechanism underlying the upregulation of METTL1 expression following MI remains unclear. Importantly, multiple studies indicate that the transcription factor MYC is elevated in fibrosis models across various organs and during fibroblast activation, playing a central regulatory role [20–22]. Our database mining revealed genetic interactions and co-expression relationships between MYC and METTL1. Further predictions suggest MYC can bind to specific sequences within the METTL1 promoter region, implying MYC may function as an upstream transcription factor regulating METTL1 transcriptional activation following MI. Therefore, we hypothesized that following MI, upregulation of MYC promotes METTL1 transcription. METTL1 mediates m7G modification of HILPDA mRNA, enhancing its stability and protein expression. Consequently, the upregulated HILPDA induces mitochondrial dysfunction and oxidative stress, ultimately driving the activation of cardiac fibroblasts and the progression of myocardial fibrosis.

This study focuses on the molecular regulatory mechanisms of myocardial fibrosis. By establishing MI mouse models and hypoxia-induced cellular models, it aims to systematically elucidate the pivotal role of the MYC/METTL1/HILPDA signaling axis in driving myocardial fibrosis. This research provides a novel perspective for deciphering the regulatory network governing ventricular pathological remodeling following MI and offers a theoretical foundation for developing therapeutic targets to intervene in fibrosis during heart failure.

Materials and methods

Animals

Male C57BL/6 mice (6–8 weeks old) were purchased from Beijing SPF Biotechnology Co., Ltd. All animals were housed under SPF conditions in a controlled environment (temperature: 21–26 °C, humidity: 40–70%) with a 12-h light/dark cycle. During the experiment, the mice had free access to food. After a one-week acclimatization period, experimental procedures were initiated. All experimental protocols were approved by.

The Institutional Animal Care and Use Committee of Nanchang Royo Biotech Co., Ltd (RYE2025070602).

Establishment of MI models

Mice were randomly divided into two groups (n = 9 per group): Sham and MI. The MI model was induced by permanent ligation of the left anterior descending coronary artery (LAD). Briefly, the anesthetized mice were intubated and mechanically ventilated. Following thoracotomy between the 3rd and 4th intercostal spaces, the heart was exposed, and the LAD was ligated with an 8 − 0 prolene suture. The chest wall and skin were then sutured closed. Sham-operated mice underwent the same surgical procedure except for LAD ligation. For the time course experiment, mice were euthanized at indicated time points (6 h, 24 h, 3 d, 7 d, and 14 d) post-MI or sham operation (n = 3 per time point). Cardiac tissues were collected, and primary cardiac fibroblasts were isolated from these tissues for qRT-PCR and Western blot (WB) analysis of HILPDA expression.

For the in vivo knockdown experiment, mice were randomly assigned to four groups (n = 9 per group): Sham, MI, MI + sh-NC, and MI + sh-HILPDA. The Sham and MI groups were treated as described above. MI + sh-NC group: Mice received an intravenous injection of a control AAV (carrying non-targeting shRNA) four weeks before LAD ligation. MI + sh-HILPDA group: Mice received an intravenous injection of AAV carrying HILPDA-targeting shRNA driven by the fibroblast-specific periostin (Postn) promoter at a titer of 1 × 10¹² vg/mL (total of 1 × 10¹¹ vg per mouse) four weeks before LAD ligation. This AAV-based strategy has been validated to achieve efficient and specific transgene expression in activated cardiac fibroblasts after MI [23]. The shRNA sequences are provided in Table 1.

Table 1.

Target sequences of shRNA and siRNA

Gene	Forward sequence (5’-3’)	Reverse sequence (5’-3’)
sh-HILPDA-1	CCGGAGTTCATGCTGAACCTCTATGCTCGAGCATAGAGGTTCAGCATGAACTTTTTTG	AATTCAAAAAAGTTCATGCTGAACCTCTATGCTCGAGCATAGAGGTTCAGCATGAACT
sh-HILPDA-2	CCGGGCTTTCCATCTTTGTTAGAGTCTCGAGACTCTAACAAAGATGGAAAGCTTTTTG	AATTCAAAAAGCTTTCCATCTTTGTTAGAGTCTCGAGACTCTAACAAAGATGGAAAGC
si-HILPDA-1	AAACCAAAAUCCAUUCUCCCA	GGAGAAUGGAUUUUGGUUUCG
si-HILPDA-2	UCCAAAAGAGUCCUUUAGGAC	CCUAAAGGACUCUUUUGGACC
si-MYC-1	AAAAGCUUCUUUUAUACUGCG	CAGUAUAAAAGAAGCUUUUCG
si-MYC-2	AUUUCUUCCAGAUAUCCUCAC	GAGGAUAUCUGGAAGAAAUUC

In each experimental group, a total of nine mice were used, with three mice assigned for histological analysis, three for molecular biology assays, and three for TTC staining. Echocardiography was performed 7 days after surgery, after which all mice were euthanized, and cardiac tissues were harvested for subsequent analyses, including histology, immunohistochemistry, WB, and TTC staining.

Isolation of mouse primary cardiac fibroblasts

Primary cardiac fibroblasts were isolated from mouse hearts. Briefly, hearts were excised, minced into 1 mm³ fragments, and digested with collagenase IV (2 mg/mL) and dispase II (1.2 U/mL) in DMEM containing 1% FBS at 37 °C for 10–15 min per cycle (3–5 cycles). The supernatants were collected, neutralized with DMEM containing 10% FBS, filtered, and centrifuged at 300–400×g for 10 min. The cell pellet was resuspended in complete medium (DMEM + 10% FBS + 1% penicillin-streptomycin) and seeded onto gelatin-coated dishes. After 1–2 h of incubation, non-adherent cells were removed, and adherent cardiac fibroblasts were cultured at 37 °C in 5% CO₂. Cells at passages 2–4 were used for subsequent experiments.

Hematoxylin and Eosin (H&E) staining

Cardiac tissues were fixed in 4% paraformaldehyde, paraffin-embedded, and sectioned into 4-µm-thick slices. Following deparaffinization in xylene and rehydration through a graded ethanol series, the sections were stained with hematoxylin solution (Servicebio, Wuhan, China) for 5 min to label nuclei, followed by differentiation and bluing. Subsequently, the cytoplasm was counterstained with eosin solution (Servicebio) for 2 min. Finally, the stained sections were dehydrated through an ethanol series, cleared in xylene, and mounted with a synthetic resin. The morphological changes, including myocardial cell arrangement, necrosis, and inflammatory cell infiltration, were examined and imaged under a light microscope (Keyence, Osaka, Japan).

Masson staining

To assess the extent of myocardial fibrosis, paraffin-embedded cardiac tissue sections.  (4 μm) were stained using the Masson’s trichrome staining kit (Solarbio, Beijing, China). Following deparaffinization and rehydration, sections were sequentially stained with Weigert’s hematoxylin for 5–10 min, Biebrich scarlet-acid fuchsin for 5–10 min, and aniline blue for 1–2 min, with differentiation using phosphomolybdic-phosphotungstic acid between red and blue staining. After rapid dehydration and clearing, sections were mounted for microscopic (Leica, Wetzlar, Germany) examination. Collagen deposition was quantified in interstitial and perivascular areas using ImageJ software (Version 1.53t, National Institutes of Health, Bethesda, MD, USA).

Immunohistochemistry (IHC)

Paraffin-embedded heart sections.  (4 μm thick) were deparaffinized in xylene and rehydrated through a graded ethanol series. After antigen retrieval in citrate buffer (pH 6.0) using a microwave heating method, endogenous peroxidase activity was quenched by incubation with 3% hydrogen peroxide. The sections were then blocked with 5% BSA to prevent non-specific binding. Subsequently, they were incubated for 30 min at 37 °C with primary antibodies against HILPDA (1:100, 14994-1-AP, Proteintech, Wuhan, China) and METTL1 (1:100, APR24316N, Biocompare, San Francisco, CA, USA). Following washes, the sections were incubated with universal secondary antibody (PV-6000, Zsbio, Beijing, China) at 37 °C for 20 min. The antigen-antibody complex was visualized using the DAB kit (Servicebio), which produced a brown precipitate. Finally, the sections were counterstained with hematoxylin (Servicebio) to label nuclei, dehydrated, cleared, and mounted. The stained sections were examined under a light microscope (Leica).

Immunofluorescence (IF)

Paraffin-embedded cardiac tissue Sect.  (4 μm) were deparaffinized, rehydrated, and subjected to antigen retrieval in citrate buffer. Cultured cells grown on coverslips were fixed with 4% paraformaldehyde for 15 min and permeabilized with 0.1% Triton X-100 for 10 min. All samples were blocked with 3% BSA for 30 min at room temperature.

For single staining, sections or cells were incubated with primary antibody against α-SMA (1:200, BM0002, Boster, Pleasanton, CA, USA), S100A4 (1:200, cy5799, ABways, Shanghai, China), and COL1A1 (1:200, BA0325, Boster) overnight at 4 °C. Following PBS washes, sections were incubated with CY3-labeled goat anti-mouse IgG secondary antibody (1:500, A0516, Beyotime, Shanghai, China) or FITC-labeled goat anti-rabbit IgG secondary antibody (1:500, A0562, Beyotime) for 1 h at room temperature protected from light. Nuclei were counterstained with DAPI.

For double staining in mouse cardiac tissues (HILPDA/Vimentin, HILPDA/cTnT, HILPDA/CD31, Ki67/CD31, Ki67/Vimentin), tyramide signal amplification (TSA) was employed. After incubation with the first primary antibody: anti-HILPDA (1:200, APR24316N, Biocompare) or anti-Ki67 (1:200, 28074-1-AP, Proteintech) overnight at 4 °C and HRP-conjugated goat anti-rabbit IgG (1:500, A0208, Beyotime) for 1 h, TSA working solution (iF488-Tyramide, Servicebio) was applied for 10 min, followed by microwave stripping. The second primary antibody: anti-Vimentin (1:200, PB9359, Boster), anti-cTnT (1:200, 15513-1-AP, Proteintech), or anti-CD31 (1:200, 28083-1-AP, Proteintech) were then incubated overnight at 4 °C, followed by CY3-conjugated goat anti-rabbit IgG (1:500, A0516, Beyotime) for 1 h. Nuclei were counterstained with DAPI.

For ROS detection, sections were incubated with DHE fluorescent probe (BB-470516, BestBio, Shanghai, China) at 37 °C for 30 min in the dark.

Finally, for all assays, fluorescence signals were visualized using a fluorescence microscope (Keyence).

Echocardiography

Mice were anesthetized with isoflurane (RWDLS, Shenzhen, China) inhalation, securely immobilized in a supine position, and the precordial region was depilated to expose the chest skin. Cardiac function was then evaluated using an ultrasound system (esaote, Genoa, Italy). Two-dimensional guided M-mode tracings were acquired from the parasternal short-axis view at the papillary muscle level. Left ventricular functional parameters, including left ventricular ejection fraction (LVEF) and left ventricular fractional shortening (LVFS), were measured from these recordings.

Sirius red staining

Paraffin-embedded cardiac tissue Sect.  (4 μm) were deparaffinized, rehydrated, and stained with Sirius Red solution (maokangbio, Shanghai, China) for 1 h at room temperature. After staining, sections were washed twice with acidified water, dehydrated through a graded ethanol series, cleared in xylene, and mounted with resinous medium. Collagen fibers were visualized under light microscopy.

TTC staining

The ventricles were cut into uniform 1 mm transverse slices and incubated in 2% TTC solution (Solarbio) at 37 °C for 30 min in the dark. Viable myocardium stained brick red due to formazan formation by dehydrogenase enzymes, while infarcted areas lacking enzyme activity remained pale. Sections were fixed in 4% paraformaldehyde to enhance contrast, and the infarct area and total ventricular area were quantified planimetrically using ImageJ software. Infarct size was expressed as the percentage of infarct area relative to the total left ventricular area.

ATP assay

ATP levels in cardiac tissues were quantified using a commercial ATP assay kit (Jiancheng Bioengineering Institute, Nanjing, China) according to the manufacturer’s protocol. ATP concentration was calculated based on the standard curve and normalized to the protein content of the homogenate.

Cellular ATP levels were quantified using an ATP Assay Kit (Solarbio). Cells from all groups were harvested, sonicated in distilled water, and centrifuged. The supernatant was deproteinized with chloroform, followed by centrifugation to collect the aqueous phase for immediate measurement. According to the kit protocol, the reaction mixture was prepared, and its absorbance at 340 nm was measured immediately (0 s) and after incubation at 37 °C (3 min) using a microplate reader (Thermo Fisher Scientific, Waltham, MA, USA). The ATP content was calculated based on the change in absorbance.

Cell culture and treatment

Mouse cardiac fibroblasts (MCFs) were purchased from iCell Bioscience Inc. Cells were maintained in a specialized fibroblast growth medium (iCell Bioscience Inc.) and cultured at 37 °C under a 5% CO₂ atmosphere. To establish an in vitro hypoxia model, cells at a stable growth state were placed in anaerobic culture bags and incubated for 24 h to induce hypoxic conditions.

MCFs were transfected with the following plasmid/siRNA constructs (see Table 1) using Lipofectamine 8000 (Beyotime) according to experimental groups: Hypoxia + OE-NC (empty vector), Hypoxia + OE-HILPDA (HILPDA overexpression plasmid), Hypoxia + si-NC (non-targeting siRNA), Hypoxia + si-HILPDA (HILPDA-targeting siRNA), Hypoxia + OE-METTL1 (METTL1 overexpression plasmid), Hypoxia + OE-METTL1 + si-NC (METTL1 plasmid + control siRNA), Hypoxia + OE-METTL1 + si-HILPDA (METTL1 plasmid + HILPDA siRNA), Hypoxia + OE-MYC (MYC overexpression plasmid), and Hypoxia + si-MYC (MYC-targeting siRNA). Following transfection, cells were cultured under standard conditions for 24–48 h before functional analysis.

For the TGF-β1-induced fibroblast activation model, MCFs were treated with TGF-β1 (10 ng/mL; 91701ES10, Yeasen, Shanghai, China) for 48 h to induce myofibroblast activation. For HILPDA knockdown experiments, cells were transfected with si-NC or si-HILPDA for 24 h before TGF-β1 treatment.

Lipid metabolism assays

To evaluate lipid accumulation, cells were subjected to Oil Red O staining. Briefly, cells were fixed with 4% paraformaldehyde for 10 min, washed with PBS, and then stained with Oil Red O working solution (C0157S, Beyotime) for 20 min at room temperature. After washing with PBS, images were captured under a light microscope (Olympus).

Cellular triglyceride (TG) and total cholesterol (TC) levels were measured using commercial assay kits (TG: A110-1-1, Nanjing Jiancheng; TC: 60723ES60, Yeasen) according to the manufacturer’s protocols. Cells were harvested and lysed, followed by the determination of TG and TC concentrations using a microplate reader at 500 nm. The results were normalized to total protein content and expressed as a percentage relative to the control group.

DCFH-DA assay

Intracellular ROS levels were measured using a ROS assay kit (Beyotime) with the fluorescent probe DCFH-DA. Cells from different groups were harvested and seeded at equal densities into 48-well plates, with three replicate wells per group, and cultured for 24 h at 37℃. Subsequently, the culture medium was removed, and 200 µL of DCFH-DA working solution, diluted 1:500 in serum-free medium to a final concentration of 20 µM, was added to each well. The plates were incubated at 37 °C in the dark for 30 min. After incubation, the cells were washed three times with serum-free culture medium to thoroughly remove any excess DCFH-DA probe. Finally, the cells were immediately observed under an inverted fluorescence microscope (Keyence), and images were captured for analysis.

JC-1 staining for mitochondrial membrane potential

Mitochondrial membrane potential was assessed using the JC-1 kit (Beyotime), followed by flow cytometry (Agilent Technologies, Santa Clara, CA, USA). Briefly, pretreated cells were harvested, washed with PBS, trypsinized, and centrifuged. The cell pellets were resuspended in 1 mL of JC-1 working solution, mixed thoroughly, and incubated at 37 °C for 20 min. Meanwhile, a sufficient amount of 1×JC-1 staining buffer was prepared and kept on ice. After incubation, the JC-1 staining solution was removed by centrifugation, and the cells were washed once with the pre-chilled 1×JC-1 staining buffer. The stained cells were then immediately analyzed using a flow cytometer. The mitochondrial membrane potential was expressed as the red/green fluorescence ratio.

Cell counting kit-8 (CCK-8) assay

Cells from all experimental groups in the logarithmic growth phase were harvested and adjusted to a density of 1 × 10⁶ cells/mL. Following cell counting and trypan blue staining to ensure cell viability, the single-cell suspensions were seeded into a 96-well plate and cultured in a 37 °C, 5% CO₂ incubator. After 24 h, 10 µL of CCK-8 solution (Yeasen) was added to each well, and the plate was incubated in the dark for 1 h under the same conditions. The absorbance (OD value) at 450 nm was then measured using a microplate reader (Thermo Fisher Scientific).

Transwell assay

Cell migration was assessed using Transwell chambers. Cells from all groups were resuspended in serum-free medium and seeded into the upper chambers at a density of 2 × 10⁵ cells/mL. The lower chambers were filled with complete medium containing 10% FBS. Following 24 h of incubation, the non-migrating cells on the upper membrane surface were removed with a cotton swab. The migrated cells on the lower surface were fixed with 4% paraformaldehyde, stained with 0.2% crystal violet, and enumerated in five random fields under an inverted microscope (Keyence). Quantitative analysis was performed using ImageJ software.

Quantitative real-time PCR (qRT-PCR)

Total RNA was extracted from cardiac tissues and MCFs using TRIzol reagent (Servicebio), and its concentration and purity were determined using a spectrophotometer. Subsequently, 1 µg of total RNA from each sample was reverse-transcribed into cDNA using the PrimeScript™ RT Kit (Takara, Japan) according to the manufacturer’s instructions. qRT-PCR was then performed in triplicate using TB Green^® Premix Ex Taq™ II Kit (Takara) on a real-time PCR detection system. The relative expression levels of target genes were analyzed using the 2^-ΔΔCt method, with GAPDH serving as the internal control. Table 2 provides detailed information on all primers used in this experiment.

Table 2.

Primers used in qRT-PCR

Gene	Forward sequence (5’-3’)	Reverse sequence (5’-3’)
GAPDH	CATCACTGCCACCCAGAAGACTG	ATGCCAGTGAGCTTCCCGTTCAG
HILPDA	GTTCCATTTCTCGGGAGGCA	GTGGCTGGAACTCCCATACC
Col1α1	CCTCAGGGTATTGCTGGACAAC	CAGAAGGACCTTGTTTGCCAGG
α-SMA	TGCTGACAGAGGCACCACTGAA	CAGTTGTACGTCCAGAGGCATAG
CTGF	TGCGAAGCTGACCTGGAGGAAA	CCGCAGAACTTAGCCCTGTATG
Collagen Ⅲ	TTCTGTGGGTCCTGCTGGGAAA	TTGTCACCTCGGATGCCTTGAG
METTL1	GTGGAGTTTGCAGACATAGGCTG	GCACATAGTCCGACACCTTCAC
MYC	TCGCTGCTGTCCTCCGAGTCC	GGTTTGCCTCTTCTCCACAGAC

WB

Total protein was extracted from mouse cardiac tissues, primary cardiac fibroblasts, and MCFs using RIPA lysis buffer (Biosharp, Hefei, China) supplemented with protease and phosphatase inhibitors. The protein concentration was determined with a BCA assay kit (Biosharp). SDS-PAGE was used to separate equal amounts of protein (40 µg) from each sample, and the proteins were subsequently transferred onto a PVDF membrane. After blocking with 5% non-fat milk for 2 h at room temperature, the membranes were incubated overnight at 4 °C with the following primary antibodies (all at 1:1000 dilution): anti-HILPDA (orb389656, Biorbyt, Cambridge, UK), anti-α-SMA (ab7817, Abcam, Cambridge, UK), anti-CTGF (25474-1-AP, Proteintech), anti-Collagen Ⅲ (ab184993, Abcam), anti-Collagen Ⅰ (ab270993, Abcam), anti-METTL1 (ab271063, Abcam), anti-MYC (ab32072, Abcam), PPARα (15540-1-AP, Proteintech), PPARβ (60193-1-Ig, Proteintech), and anti-GAPDH (ab8245, Abcam). Following incubation with HRP-labeled mouse IgG secondary antibody (1:5,000, ab6789, Abcam) and rabbit IgG secondary antibody (1:5,000, ab6721, Abcam) for 2 h at room temperature, the protein bands were visualized using the ECL detection system. GAPDH was used as a loading control for normalization, and band intensities were quantified using ImageJ.

Dot blot assay

Total RNA was extracted from the samples, and its concentration was measured. Equal amounts of RNA from each group were denatured and spotted directly onto a positively charged nylon membrane. The RNA was then cross-linked to the membrane using UV light. After blocking with 5% non-fat milk, the membrane was incubated overnight at 4 °C with an m7G primary antibody (DB-m7G, RayBiotech, Peachtree Corners, GA, USA). Subsequently, the membrane was incubated with HRP-conjugated secondary antibody (RayBiotech) for 2 h at room temperature. The m7G signals were finally visualized using an ECL detection system and quantified using ImageJ software. Methylene blue staining (Solarbio) of the membrane was performed to confirm equal RNA loading.

RNA immunoprecipitation (RIP)

The interaction between the METTL1 protein and HILPDA mRNA was assessed using the RIP Kit (GeneCreate, Wuhan, China). Cardiac tissues or cells were lysed in complete RIP lysis buffer (Invitrogen, Waltham, MA, USA). The lysates were then incubated with magnetic beads conjugated to 3–5 µg of anti-METTL1 antibody (ab309096, Abcam), with normal mouse IgG serving as a negative control. After extensive washing, the co-precipitated RNAs were isolated and purified. The enrichment of HILPDA mRNA in the immunoprecipitates was subsequently analyzed by qRT-PCR, with the results presented as fold-change relative to the IgG control group.

Actinomycin D chase assay

The stability of HILPDA mRNA was assessed using an actinomycin D chase assay. Cells from respective groups were treated with actinomycin D (100 µg/mL; MCE, Monmouth Junction, NJ, USA) to halt de novo RNA transcription. Total RNA was then isolated at predetermined time points (0, 4, 8, 12 h) post-treatment. The relative levels of HILPDA mRNA at each time point were quantified by qRT-PCR and normalized to the internal control GAPDH.

Chromatin immunoprecipitation (ChIP)

The binding of MYC to the METTL1 promoter region was analyzed by the ChIP kit (Beyotime). MCFs were cross-linked with 1% formaldehyde, and the chromatin was subsequently sheared by sonication to fragments of 200–1000 bp. The lysates were then immunoprecipitated overnight at 4 °C using 0.5 µg anti-MYC antibody, with normal IgG serving as a negative control. Following reversal of cross-links and proteinase K digestion, the purified DNA was analyzed by qPCR using primers specific for the METTL1 promoter region. The enrichment of the target DNA fragment in the MYC immunoprecipitate was calculated relative to the IgG control.

Dual-luciferase reporter assay

The METTL1 promoter fragment was cloned into a pGL3-based firefly luciferase vector (constructed by Tsingke Biotechnology). MCFs seeded in 6-well plates were co-transfected with the reporter plasmid and the Renilla luciferase internal control plasmid using Lipo8000™ transfection reagent (Beyotime). After 24–48 h, cells were lysed, and luciferase activities were measured sequentially with firefly and Renilla luciferase substrates using a luminometer. Firefly luciferase activity was normalized to Renilla luciferase activity for each sample, and relative promoter activity was expressed as the Firefly/Renilla luminescence ratio.

Bioinformatics analysis

The RMbase database (http://rna.sysu.edu.cn/rmbase/) was first employed to predict putative m7G modification sites on the HILPDA mRNA transcript, using default parameters for human transcriptome-wide analysis. To explore the potential interaction between METTL1 protein and HILPDA mRNA, we queried the ENCORI platform (https://rnasysu.com/encori/) with stringent filtering criteria (CLIP-seq data threshold: ≥ 2; pan-cancer analysis). For expression correlation analysis in cardiac tissues, the GEPIA database (http://gepia.cancer-pku.cn/) was utilized to calculate the Pearson correlation coefficients between METTL1 and HILPDA, as well as between METTL1 and MYC, using TCGA and GTEx normal heart tissue datasets. To investigate transcriptional regulation, the JASPAR database (https://jaspar.genereg.net/) was accessed with a relative profile score cutoff of 85% to predict the binding of MYC transcription factor to the “CCACGTGG” sequence identified in the METTL1 promoter region. Finally, the GeneMANIA database (https://genemania.org/) was used to construct a comprehensive interaction network between METTL1 and MYC, analyzing multiple interaction types, including genetic interactions and co-expression. The results were visualized using the respective platforms’ built-in tools.

Correlation analysis

Protein expression levels of MYC, METTL1, and HILPDA in cardiac tissues from Sham and MI mice were quantified by WB. Correlation coefficients (r) and p values were calculated using GraphPad Prism software. For bioinformatic correlation analysis, gene expression data from the GSE153494 dataset were downloaded and analyzed. Pearson correlation coefficients were calculated to assess the pairwise correlations among MYC, METTL1, and HILPDA.

Statistical analysis

All data are presented as mean ± standard deviation (SD) from at least three independent experiments. For animal studies, n represents the number of mice per group (n = 3). For in vitro experiments, n represents the number of independent biological replicates (n = 3). Statistical analyses were performed using GraphPad Prism software (version 8.0). Differences between two groups were analyzed using the unpaired Student’s t-test, while comparisons among multiple groups were evaluated by one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test. A value of p < 0.05 was considered statistically significant.

Results

HILPDA is upregulated in both cardiac tissue and primary fibroblasts after MI

We first analyzed differential gene expression following MI using the public dataset GSE153494. The volcano plot revealed that HILPDA expression was significantly upregulated in the MI 24 h group (M24 h) (Fig. 1A). A violin plot further visually demonstrated the elevated expression of HILPDA in the M24 h group (Fig. 1B). These bioinformatic findings suggest a potential role for HILPDA in MI pathogenesis. To investigate the temporal expression pattern of HILPDA after MI, the MI mouse model was established by LAD ligation. Primary cardiac fibroblasts were isolated from Sham and MI mouse hearts at the indicated time points (6 h, 24 h, 3 d, 7 d, and 14 d) post-MI. qRT-PCR and WB analysis revealed that HILPDA mRNA and protein expression level began to increase at 24 h, progressively increased, and peaked at 7 d after MI (Fig. S1A-B). These results are consistent with the bioinformatic findings at 24 h and further demonstrate that HILPDA upregulation persists into the fibrotic phase. Therefore, the 7-d time point was selected for subsequent in vivo experiments.

Fig. 1.

Expression of HILPDA in cardiac tissue and primary fibroblasts after MI. (A) Volcano plot depicting differential gene expression between the MI-control (Mcon) and MI-24 h (M24 h) groups in the GSE153494 dataset. (B) Violin plot depicting the expression of the target differential gene HILPDA in the Mcon and M24 h groups from the GSE153494 dataset. MI models were established, followed by the isolation of cardiac tissue. In vivo experimental groups: Sham and MI. (C) H&E staining was conducted to assess pathological damage in cardiac tissue. (D) Masson staining was applied to detect fibrosis in cardiac tissue. (E-G) The expression of HILPDA in cardiac tissue was evaluated by qRT-PCR, IHC, and WB. Primary cardiac fibroblasts were isolated from Sham and MI mouse hearts. Experimental groups: Sham-CF and MI-CF. (H-I) qRT-PCR and WB were utilized to assess the expression level of HILPDA. Data are presented as mean ± SD. n = 3. ^***p ˂ 0.001 vs. Sham/ Sham-CF

To validate this finding, we examined cardiac tissue at 7 d post-MI. H&E staining revealed marked pathological damage in cardiac tissue of the MI group, characterized by disorganized myocardial cell arrangement, necrosis, and inflammatory cell infiltration (Fig. 1C). Masson staining further revealed significantly increased myocardial fibrosis in the MI group (Fig. 1D). At the molecular level, qRT-PCR, IHC, and WB results consistently demonstrated that both mRNA and protein expression of HILPDA was significantly upregulated in the cardiac tissue of the MI group (Fig. 1E-G). To further confirm HILPDA upregulation specifically in cardiac fibroblasts at the fibrotic stage, primary cardiac fibroblasts were isolated from Sham and MI mouse hearts at 7 d post-MI. Consistent with the tissue-level findings, qRT-PCR and WB analysis demonstrated that HILPDA mRNA and protein expression level was significantly elevated in the cardiac fibroblasts isolated from MI mice (MI-CFs) (Fig. 1H-I). These results collectively suggest that HILPDA may contribute to the development and progression of post-infarction fibrosis.

HILPDA knockdown ameliorates cardiac impairment in MI

To investigate the cardiac function of HILPDA in vivo, an AAV lentiviral vector carrying sh-HILPDA was administered via the tail vein into the MI mouse model. Knockdown efficiency was verified by qRT-PCR and WB (Fig. S2A-B). Echocardiography results revealed significantly reduced LVEF and LVFS scores in the MI group, whereas sh-HILPDA treatment markedly restored these cardiac function metrics, indicating effective improvement in cardiac contractility (Fig. 2A-C). TTC staining revealed a significant increase in infarct size of MI mice, whereas sh-HILPDA treatment suppressed infarct expansion (Fig. 2D). H&E staining revealed structural disorganization, extensive necrosis, and inflammatory cell infiltration in cardiomyocytes of the MI group, whereas HILPDA knockdown markedly mitigated these pathological lesions (Fig. 2E). Additionally, Sirius red and Masson trichrome staining further confirmed that HILPDA knockdown effectively alleviated myocardial fibrosis (Fig. 2F-H).

Fig. 2.

The effect of HILPDA knockdown on improving cardiac function and myocardial fibrosis after MI. MI mice were administered with sh-NC/ sh-HILPDA via tail vein injection 4 weeks before MI surgery. Echocardiography was performed 7 days after MI, followed by cardiac tissue collection. Experimental groups: Sham, MI, MI + sh-NC, MI + sh-HILPDA. (A-C) Echocardiography was used to measure LVEF and LVFS to assess cardiac function. (D) Infarct size was visualized by TTC staining. (E) Pathological injury was observed by H&E staining. (F-H) Sirius Red and Masson staining were carried out to examine myocardial fibrosis. (I-J) Double IF staining was used to detect the co-localization of HILPDA with Vimentin, cTnT, or CD31. (K-L) Double IF staining was evaluated for Ki67 with CD31 or Vimentin. Data are presented as mean ± SD. n = 3. ^*p <0.05, ^**p <0.01, ^***p <0.001, ns means no significant difference vs. Sham/ MI + sh-NC

To investigate the cellular specificity of HILPDA and its role in post-MI myocardial remodeling, we performed immunofluorescence co-localization analysis. The results showed that HILPDA exhibited varying degrees of co-localization with Vimentin, cTnT, and CD31 in all groups. However, in the MI model, HILPDA knockdown only significantly reduced the co-localization fluorescence intensity of HILPDA with the fibroblast marker Vimentin, while its co-localization fluorescence intensity with the cardiomyocyte marker cTnT and endothelial cell marker CD31 remained unchanged (Fig. 2I-J), indicating that HILPDA primarily functions in cardiac fibroblasts post-MI. Ki67 double staining with CD31 or Vimentin further revealed that HILPDA knockdown significantly reduced both Ki67/CD31 co-localization fluorescence intensity (endothelial proliferation) and the area of Ki67⁺Vimentin⁺ cells (fibroblast proliferation) in MI mice (Fig. 2K-L), demonstrating that HILPDA knockdown suppresses critical proliferative events, including fibroblast proliferation as a core cellular event in myocardial fibrosis. These data collectively demonstrate that HILPDA knockdown effectively improves cardiac function post-MI, reduces myocardial fibrosis, diminishes infarct size, and suppresses endothelial proliferation, thereby providing clear cardioprotective effects.

HILPDA knockdown ameliorates fibroblast activation, mitochondrial dysfunction, and lipid metabolism dysregulation

To elucidate the role of HILPDA in myocardial fibrosis, MCFs were cultured under normoxic or hypoxic conditions and then transfected with si-HILPDA or OE-HILPDA. The overexpression and knockdown efficiency of HILPDA were successfully validated (Fig. S2C-F). qRT-PCR and WB analysis revealed that hypoxia significantly upregulated HILPDA expression, an effect that was further enhanced by OE-HILPDA. In contrast, si-HILPDA treatment effectively reduced HILPDA expression (Fig. 3A-B). The impact of HILPDA on hypoxia-induced mitochondrial dysfunction was then evaluated. Hypoxia stimulation caused elevated ROS levels (Fig. 3C-D), decreased mitochondrial membrane potential (red/green ratio; Fig. 3E-F), and reduced ATP synthesis in MCFs (Fig. 3G), indicating impaired mitochondrial function. HILPDA overexpression further exacerbated these abnormalities, whereas HILPDA knockdown significantly alleviated hypoxia-induced mitochondrial dysfunction. At the cellular functional level, CCK-8 and Transwell assays revealed that hypoxia and OE-HILPDA treatment significantly promoted the proliferation and migration of MCFs. In contrast, si-HILPDA treatment effectively reversed abnormal cellular proliferation and migration (Fig. 3H-J). At the molecular phenotype level, hypoxia exposure significantly upregulated mRNA and protein expression of myofibroblast transdifferentiation markers (Col1α1, α-SMA, and CTGF). This trend was further amplified upon HILPDA overexpression but was significantly reversed following HILPDA knockdown (Fig. 3K-L). IF staining further confirmed that HILPDA knockdown significantly suppressed hypoxia-induced Col1α1 and α-SMA expression (Fig. 3M, Fig. S3A).

Fig. 3.

The effects of HILPDA on hypoxia-induced mitochondrial dysfunction and fibroblast activation. Hypoxia-induced cell model was established using MCFs. Experiments groups: Normoxia, Hypoxia, Hypoxia + OE-NC, Hypoxia + OE-HILPDA, Hypoxia + si-NC, Hypoxia + si-HILPDA. (A-B) qRT-PCR and WB were utilized to quantify the expression of HILPDA. (C-D) ROS levels were assessed using DCFH-DA assay kit. (E-F) Flow cytometry was used to assess mitochondrial membrane potential using JC-1 staining, and the results are presented as the red/green fluorescence ratio. (G) ATP production rate was confirmed by enhanced ATP assay kit. (H) CCK-8 was utilized to evaluate cell viability. (I-J) Transwell assay was carried out to examine fibroblast migration ability. (K-L) The expression levels of myofibroblast transformation markers (Col1α1, α-SMA, and CTGF) were determined by qRT-PCR and WB. (M) IF was employed to detect the expression of Col1α1 and α-SMA. Data are presented as mean ± SD. n = 3. ^*p <0.05, ^**p <0.01, ^***p <0.001 vs. Normoxia/ Hypoxia + OE-NC/ Hypoxia + si-NC

To investigate whether HILPDA affects fibroblast activation through lipid metabolism dysregulation, we examined lipid-related parameters in hypoxia-induced MCFs. Oil Red O staining revealed that hypoxia significantly increased lipid droplet accumulation, an effect that was further enhanced by HILPDA overexpression, whereas HILPDA knockdown markedly reduced lipid droplet formation (Fig. S3B-C). Consistently, TG and TC levels were significantly elevated under hypoxic conditions, with OE-HILPDA further increasing and si-HILPDA decreasing their levels (Fig. S3D-E). Furthermore, WB analysis further showed that HILPDA knockdown reduced the protein expression levels of lipid oxidation markers (PPARα and PPARβ) (Fig. S3F).

To further validate our findings in a more physiologically relevant fibrosis model, we employed TGF-β1 to activate MCFs. Consistent with the hypoxia results, TGF-β1 induction significantly upregulated HILPDA expression level, which was effectively reversed by HILPDA knockdown (Fig. S4A-B). TGF-β1 stimulation also induced mitochondrial dysfunction (increased ROS, reduced ATP, decreased mitochondrial membrane potential (red/green ratio)), lipid metabolism abnormalities (elevated TG and TC levels, increased PPARα and PPARβ protein expression levels, enhanced lipid droplet accumulation), and enhanced cell proliferation and migration, all of which were ameliorated by HILPDA knockdown (Fig. S4C-K). Moreover, TGF-β1-induced upregulation of myofibroblast markers (Col1α1, α-SMA, CTGF) was significantly attenuated by si-HILPDA treatment (Fig. S4L-O). Collectively, these results demonstrate that HILPDA promotes fibroblast activation, induces mitochondrial dysfunction, and disrupts lipid metabolism under both hypoxic and TGF-β1 stimulation, supporting its pro-fibrotic role following MI.

METTL1 promotes fibroblast activation and myocardial fibrosis by mediating m7G modification of HILPDA

Bioinformatics analysis first predicted potential m7G modification sites on HILPDA mRNA using the RMbase database, and the ENCORI database suggested potential binding between METTL1 and HILPDA mRNA (Fig. S5A-B). Gepia analysis further revealed a significant positive correlation between METTL1 and HILPDA expression in cardiac tissue, suggesting a potential functional association (Fig. S5C). Therefore, we further explored the role of METTL1 in myocardial fibrosis and its regulatory mechanism governing HILPDA. In the MI model, both mRNA and protein expression of METTL1 were significantly upregulated in cardiac tissue, accompanied by markedly elevated m7G modification level (Fig. 4A-D, Fig. S5D). RIP assays in MCFs further demonstrated that METTL1 protein specifically binds to HILPDA mRNA under hypoxic conditions (Fig. 4E).

Fig. 4.

The effects of METTL1-mediated m7G modification of HILPDA on fibroblasts and myocardial fibrosis. In vivo experiments groups: Sham, MI. (A-C) qRT-PCR, WB, and IHC were utilized to quantify the expression of METTL1. (D) Dot Blot assay was used to detect the m7G level. In vitro experiments groups: Normoxia, Hypoxia. (E) RIP analyzed the interaction between the METTL1 protein and HILPDA mRNA. Hypoxia-induced MCFs were transfected with OE-NC/ OE-METTL1/ OE-METTL1 + si-NC/ OE-METTL1 + si-HILPDA. (F) Dot Blot assay was performed to assess the m7G level. (G-H) qRT-PCR and WB were conducted to quantify the expression of HILPDA. (I) Actinomycin D assay was used to evaluate HILPDA RNA stability. (J-K) ROS levels were determined using DCFH-DA assay kit. (L) ATP production rate was confirmed by enhanced ATP assay kit. (M) CCK-8 was conducted to measure cell viability. (N-O) Fibroblast migration capacity was examined by Transwell assay. (P-Q) qRT-PCR and WB were applied to analyze the expression of collagen deposition-related markers (Collagen I and Collagen III). (R) Cardiac fibroblast activation markers (S100A4 and α-SMA) were visualized by IF. Data are presented as mean ± SD. n = 3. ^*p <0.05, ^**p <0.01, ^***p <0.001, ns means no significant difference vs. Sham/ IgG/ Hypoxia + OE-NC/ Hypoxia + OE-METTL1 + si-NC

To validate the regulatory role of METTL1 on HILPDA, hypoxia-induced MCFs were transfected with OE-METTL1 or combined with si-HILPDA. OE-METTL1 transfection efficiency was validated (Fig. S2G-H). Dot blot assay confirmed that METTL1 overexpression significantly elevated the m7G modification level, whereas si-HILPDA treatment showed the opposite trend (Fig. 4F). qRT-PCR and WB results demonstrated that METTL1 overexpression significantly elevated HILPDA expression level, while HILPDA knockdown reversed this effect (Fig. 4G-H, Fig. S5E). Actinomycin D chase assay revealed that METTL1 overexpression stabilized HILPDA mRNA, the effect that was abrogated by HILPDA knockdown (Fig. 4I). At the mitochondrial level, OE-METTL1 treatment exacerbated hypoxia-induced dysfunction, leading to elevated ROS levels and reduced ATP production. Conversely, this phenotype was rescued by si-HILPDA treatment (Fig. 4J-L). At the cellular functional level, OE-METTL1 treatment promoted cell proliferation and migration, whereas HILPDA knockdown abolished these effects (Fig. 4M-O). Regarding the fibrotic phenotype, results further demonstrated that si-HILPDA treatment effectively suppressed the upregulation of collagen deposition markers (collagen I, collagen III) and fibrosis-related markers (S100A4, α-SMA) (Fig. 4P-R, Fig. S5F-G). These findings demonstrate that METTL1 enhances HILPDA stability by binding to its mRNA and mediating m7G modification, ultimately driving fibroblast activation and myocardial fibrosis progression.

MYC enhances myocardial fibrosis by promoting METTL1 transcription

To elucidate the upstream transcriptional regulation mechanism of METTL1, we conducted bioinformatics analysis. Gepia database analysis revealed a significant positive correlation between the expression of transcription factor MYC and METTL1 in cardiac tissue (Fig. S6A). Furthermore, GeneMania website analysis predicted genetic interactions and co-expression relationships between the two, suggesting that MYC may be a potential upstream regulator of METTL1 (Fig. S6B). In the MI model, qRT-PCR and WB results confirmed that MYC mRNA and protein expression were significantly upregulated in cardiac tissue (Fig. 5A-B). Analysis via the JASPRA website revealed that MYC binds to the “CCACGTGG” sequence in the METTL1 promoter region (Fig. 5C). Subsequent ChIP and dual luciferase reporter assays performed in hypoxia-induced MCFs validated that MYC directly binds the METTL1 promoter region and significantly enhances its transcriptional activity (Fig. 5D-E).

Fig. 5.

The role of MYC in regulating METTL1 transcription during myocardial fibrosis. In vivo experimental groups: Sham, MI. (A-B) The expression of MYC in cardiac tissues was quantified by qRT-PCR and WB. (C) Binding of MYC to the “CCACGTGG” sequence within the METTL1 transcriptional regulatory region was predicted using the JASPAR database. In vitro experimental groups: Normoxia, Hypoxia. (D) ChIP was utilized to analyze the interaction between MYC and the METTL1 promoter region. (E) METTL1 promoter activity was assessed using dual-luciferase reporter assay. In vitro experimental groups: Normoxia, Hypoxia, Hypoxia + OE-NC, Hypoxia + OE-MYC, Hypoxia + si-NC, Hypoxia + si-MYC. (F-H) qRT-PCR and WB were used to quantify the expression of METTL1 and HILPDA. (I) The m7G level was measured by Dot Blot assay. (J) CCK-8 was employed to evaluate cell viability. (K-L) Fibroblast migration capacity was examined by Transwell assay. (M-N) qRT-PCR and WB were conducted to examine the expression of Col1α1, α-SMA, and CTGF. (O-P) The expression levels of Col1α1 and α-SMA were visualized by IF. Data are presented as mean ± SD. n = 3. ^**p <0.01, ^***p <0.001, ns means no significant difference vs. Sham/ IgG/ Normoxia/ Hypoxia + OE-NC/ Hypoxia + si-NC

To investigate the biological function of MYC, hypoxia-induced MCFs were transfected with OE-NC/ OE-MYC/ si-NC/ si-MYC. The transfection efficiency of si-MYC and OE-MYC was verified (Fig. S2I-L). qRT-PCR and WB results showed that hypoxia induction or OE-MYC treatment synchronously upregulated METTL1 and HILPDA mRNA and protein expressions, increasing m7G modification level, while si-MYC treatment significantly reversed these molecular changes (Fig. 5F-I). In terms of cellular function and fibrotic phenotype, hypoxia stimulation significantly promoted MCF proliferation (Fig. 5J), migration (Fig. 5K-L), and expression of myofibroblastic transdifferentiation markers (Col1α1, α-SMA, CTGF) (Fig. 5M-P). These effects were further exacerbated by OE-MYC treatment. Conversely, MYC knockdown significantly suppressed these changes.

To further validate the regulatory relationships among MYC, METTL1, and HILPDA, we performed correlation analyses. Bioinformatic analysis of the GSE153494 dataset revealed significant positive correlations among METTL1 and MYC, METTL1 and HILPDA, and HILPDA and MYC (Fig. S6C). Consistently, Pearson correlation analysis of WB results from Sham and MI mice (Figs. 1G, 4B and 5B) revealed strong positive correlations among METTL1 and MYC, METTL1 and HILPDA, and HILPDA and MYC (Fig. S6D). These results support the existence of a coordinated regulatory axis among MYC, METTL1, and HILPDA after MI. Collectively, these findings demonstrate that MYC promotes myocardial fibrosis by activating METTL1 transcription, with MYC, METTL1, and HILPDA expression levels showing significant positive correlations.

HILPDA knockdown alleviates mitochondrial dysfunction and reduces myocardial fibroblast activation in vivo

To validate the in vivo function of HILPDA, an AAV vector carrying sh-HILPDA was constructed and administered via the tail vein into the MI mouse model. qRT-PCR and WB analysis showed that HILPDA, METTL1, and MYC expression levels were all upregulated in MI cardiac tissues. sh-HILPDA treatment significantly reduced HILPDA expression but did not affect MYC or METTL1 expression levels (Fig. 6A-B), indicating that HILPDA knockdown does not feedback to regulate its upstream regulators. IF and biochemical assays revealed significantly elevated ROS and impaired ATP synthesis in the MI group. In contrast, sh-HILPDA treatment effectively attenuated these abnormalities (Fig. 6C-D). Regarding the fibrotic phenotype, results further demonstrated that sh-HILPDA treatment significantly downregulated the expression of Collagen I, Collagen III, S100A4, and α-SMA (Fig. 6E-G). These findings indicate that HILPDA knockdown effectively improves mitochondrial dysfunction after MI, suppresses fibroblast activation and collagen deposition, thereby exerting an anti-fibrotic effect.

Fig. 6.

The effects of HILPDA expression on mitochondrial dysfunction and myocardial fibroblast activation in vivo. MI mice were administered with sh-NC/ sh-HILPDA via tail vein injection 4 weeks before MI surgery. Cardiac tissues were collected 7 days after MI surgery. In vivo experimental groups: Sham, MI, MI + sh-NC, MI + sh-HILPDA. (A-B) The expression levels of HILPDA, METTL1, and MYC in cardiac tissues were quantified by qRT-PCR and WB. (C) IF was performed to assess ROS levels. (D) Enhanced ATP assay kit was employed to measure ATP production rate. (E-F) qRT-PCR and WB were utilized to analyze the expression of Collagen I and Collagen III in cardiac tissues. (G) The S100A4 and α-SMA expression levels in cardiac tissues were visualized by IF. Data are presented as mean ± SD. n = 3. ^*p <0.05, ^**p <0.01, ^***p <0.001 vs. Sham/ MI + sh-NC

Discussion

Myocardial fibrosis represents a core pathological process in cardiac remodeling following MI, yet its post-transcriptional regulatory mechanisms remain incompletely understood. This study reveals the pivotal role of the MYC-METTL1-HILPDA axis in the development of myocardial fibrosis. This finding not only advances our understanding of epitranscriptional regulation in cardiac diseases but also reveals novel potential therapeutic targets for myocardial fibrosis.

In this study, we observed that HILPDA expression began to increase at 24 h post-MI and peaked at 7 d. This temporal pattern aligns with the well-established pathophysiology of post-infarction cardiac remodeling: the inflammatory phase occurs within the first 3 d, the proliferative phase spans from 3 to 14 d, characterized by cardiac fibroblast activation and extracellular matrix deposition, and the maturation phase extends beyond 2 weeks [24]. Specifically, the 24 h time point falls within the acute inflammatory phase, during which HILPDA upregulation was already detectable. The 7 d time point corresponds to the proliferative fibrotic phase, which also represents the peak of HILPDA expression. Therefore, selecting the 7 d time point for fibrosis-related analysis is justified. Although the public dataset GSE153494 only provided 24 h post-MI data for preliminary screening, our time-course experiments bridged this gap by demonstrating that HILPDA upregulation at 24 h progressively increased and was sustained through 7 d, supporting the relevance of the 24 h screening data to later fibrotic events.

HILPDA, a hypoxia-induced regulator of lipid metabolism, has been reported to participate in disease development by promoting lipid accumulation in organs such as the liver and kidneys [25, 26]. This study found that HILPDA expression was significantly upregulated in the MI model and in hypoxia-induced MCFs. Furthermore, mounting evidence indicates that mitochondrial dysfunction plays a crucial role in the pathogenesis of various fibrotic diseases, with lipid metabolism disorders recognized as a key factor inducing mitochondrial injury [27, 28]. Under pathological conditions of lipid metabolism disorders, imbalanced energy supply and oxidative stress may drive metabolic reprogramming in fibroblasts, triggering mitochondrial dysfunction. This, in turn, promotes their transformation into activated myofibroblasts, characterized by α-SMA overexpression and abnormal extracellular matrix deposition, ultimately accelerating myocardial fibrosis progression [29, 30]. This study confirms that HILPDA overexpression induced mitochondrial dysfunction, characterized by excessive ROS production, reduced mitochondrial membrane potential, and impaired ATP synthesis, thereby promoting fibroblast-to-myofibroblast transition. Conversely, knocking down HILPDA expression reverses these changes. This finding is consistent with reports by Liu et al. in renal fibrosis [17] and underscores the central role of metabolic dysfunction in the fibrotic process.

Unlike previous research primarily focusing on METTL1’s function in tumor progression or neurological diseases [31, 32], this study reveals its equally significant role in the myocardial fibrosis process. Experimental results demonstrate that both METTL1 expression and its mediated m7G modification levels are significantly upregulated in myocardial tissue following MI. This phenomenon aligns with the m7G modification trend observed by Ma et al. in heart failure models [33], suggesting m7G modification may represent a common mechanism for cardiac response to pathological stress. Further mechanistic studies revealed that METTL1, as a core m7G methyltransferase, specifically binds to HILPDA mRNA and enhances its stability through m7G modification, thereby promoting HILPDA protein expression. This discovery not only uncovers a novel function of METTL1 in myocardial fibrosis but also establishes a direct link between m7G modification and metabolic regulation. Notably, He et al. discovered that the m6A methyltransferase METTL3 promotes liver fibrosis by regulating the TGF-β signaling pathway [34], revealing potential synergistic regulatory networks involving different RNA modifications in organ fibrosis.

This study further elucidated the role of MYC in myocardial fibrosis. Consistent with its established pro-fibrotic role in multiple organs [21, 22, 35], MYC expression was significantly upregulated in our MI model. To elucidate novel MYC-mediated fibrotic mechanisms, bioinformatics analysis, ChIP, and dual luciferase reporter assays confirmed that MYC directly binds to specific sequences within the METTL1 promoter region, thereby enhancing its transcriptional activity. This establishes a novel link between MYC, a classic pro-fibrotic factor, and epitranscriptional regulation. Notably, the MYC-METTL1-HILPDA axis identified in this study corresponds with previously reported MYC regulation of HILPDA expression in renal clear cell carcinoma research [36], but further reveals the specific regulatory mode of this axis within the myocardial system. Pearson correlation analyses of both public datasets and our WB data confirmed significant positive correlations among MYC, METTL1, and HILPDA, further supporting this regulatory axis. Subsequent functional experiments demonstrated that MYC ultimately influences fibroblast proliferation, migration, and activation by regulating the METTL1-HILPDA axis. These findings establish MYC as a pivotal hub in the fibrosis regulatory network, providing novel targets for multi-level intervention strategies.

Finally, the therapeutic potential of targeting HILPDA was validated through in vivo experiments. Knockdown of HILPDA in the MI model significantly improved mitochondrial function and reduced expression of fibrosis markers. This finding is corroborated by Wang et al.‘s report that fibroblast-specific knockdown of METTL1 alleviates myocardial fibrosis [9], collectively supporting the central role of the MYC-METTL1-HILPDA pathway in cardiac remodeling. These findings have significant translational value, suggesting that the MYC-METTL1-HILPDA axis may serve as a promising therapeutic target. This provides novel insights for clinical intervention in the progression of heart failure following MI. These discoveries not only deepen our understanding of the molecular network underlying myocardial fibrosis but also point to new directions for subsequent translational medical research.

The innovation of this study lies in establishing a complete signaling pathway from transcription factors to epigenetic modifications to metabolic regulation, systematically elucidating a novel regulatory mechanism of fibrosis following MI. However, this study has several limitations: First, the specific m7G modification site on HILPDA mRNA mediated by METTL1 and its functional significance remain unclear. Additionally, the precise downstream mechanisms by which HILPDA induces mitochondrial dysfunction, particularly its potential association with lipid metabolism and ferroptosis, require further investigation. Based on these findings, future research should focus on the following areas: precisely identifying m7G modification sites on HILPDA mRNA using mass spectrometry and gene editing technologies; elucidating the molecular mechanisms by which HILPDA regulates mitochondrial function through lipidomics and metabolic flux analysis; exploring the universality of this pathway in other cardiac disease models; and developing small-molecule inhibitors targeting this pathway to provide novel therapeutic strategies for clinical applications.

Conclusion

In summary, this study reveals that following MI, upregulation of the transcription factor MYC directly activates transcription of the m7G methyltransferase METTL1. This, in turn, mediates m7G modification of the mRNA for the key metabolic factor HILPDA, enhancing its stability and thereby promoting HILPDA protein expression. The elevated HILPDA ultimately drives myocardial fibroblast activation and the progression of myocardial fibrosis by inducing mitochondrial dysfunction. These results elucidate a novel MYC-METTL1-HILPDA regulatory axis, providing a theoretical framework and promising therapeutic targets for combating cardiac remodeling and heart failure.

Electronic Supplementary Material

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Abbreviations

α-SMA

α-smooth muscle actin

ATGL

Adipose triglyceride lipase

CCK-8

Cell counting Kit-8

CFs

Cardiac fibroblasts

ChIP

Chromatin immunoprecipitation

H&E

Hematoxylin and eosin

HILPDA

Hypoxia-inducible lipid droplet-associated protein

IF

Immunofluorescence

IHC

Immunohistochemistry

LAD

Left anterior descending coronary artery

LVEF

Left ventricular ejection fraction

LVFS

Left ventricular fractional shortening

MCFs

Mouse cardiac fibroblasts

m7G

N7-methylguanosine

MI

Myocardial infarction

Postn

Periostin

qRT-PCR

Quantitative real-time PCR

RIP

RNA Immunoprecipitation

SD

Standard deviation

TGF-β1

Transforming growth factor-β 1

TSA

Tyramide signal amplification

WB

Western blot

Author contributions

Yue Liu: Conceptualization, Investigation, Data curation, Formal analysis, Writing-original draft, Writing-review & editing. Kai Li: Conceptualization, Investigation, Formal analysis, Writing-review & editing. Yi Chen: Formal analysis, Methodology, Writing-review & editing. Huasong Xia: Conceptualization, Project administration, Resources, Supervision, Writing-review & editing.

Funding

This study was supported by Jiangxi Provincial Health Commission Scientific Research Project (202610430).

Data availability

All data generated and/or analyzed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

All experimental protocols were approved by The Institutional Animal Care and Use Committee of Nanchang Royo Biotech Co., Ltd (RYE2025070602). All mouse experimental procedures involved in this study were conducted in accordance with international guidelines for the care and use of laboratory animals.

Competing interests

The authors declare no competing interests.