HSPA12B May Help Prevent Aging in Blood Vessel Cells by Controlling STING Breakdown

Cellular senescence has emerged as a key contributor to the pathogenesis of age‐related cardiomyopathy (Childs et al. 2018; Evangelou et al. 2023; Gevaert et al. 2017; Hemanthakumar et al. 2021; Marino et al. 2022). To assess cellular senescence in the aging heart, we evaluated the expression of senescence marker expression in cardiac endothelial cells isolated from young (3–5 m) and aged (22–24 m) mice, and measured key components of the SASP, including chemokine (C‐X‐C motif) ligand (CXCL)2, CXCL9, Interleukin‐1 alpha (IL‐1A), and tumor Necrosis Factor Alpha (TNFα) of young and aged mouse heart tissues (Figure 1A,B and Figure S1A–C). Aged mouse hearts exhibited a significant increase in senescence‐associated β‐galactosidase (SA‐β‐gal) positive staining and elevated expression of senescence and SASP markers, accompanied by enhanced fibrotic remodeling (Figure 1C,D and Figure S1D,E) and impaired cardiac function, as evidenced by reduced ejection fraction (EF) and fractional shortening (FS) compared with young controls (Figure 1F,G). To further define the identity of senescent cell populations, we analyzed publicly available single‐cell RNA sequencing (scRNA‐seq) data from human cardiac tissues of elderly individuals (ages 60–75 years) (Kanemaru et al. 2025). scRNA‐seq analysis revealed that cardiac endothelial cells exhibited significantly higher expression of senescence‐associated genes compared with other cardiac cell types (Figure 1E). Additionally, we compared the senescence signature of cardiac endothelial cells from young (20–60 years) and aged (60–75 years) individuals. Compared with young, cardiac endothelial cells from aged individuals showed increased expression of senescence‐related genes. To validate these findings in mice, we performed immunofluorescent co‐staining of the senescence markers cyclin‐dependent kinase inhibitor 1A (Cdkn1a/p21) and cyclin‐dependent kinase inhibitor 2A (Cdkn2a/p16) alongside the endothelial marker CD31 in heart tissues from young (3–5 months) and aged (22–24 months) mice. As expected, the majority of p21‐ and p16‐positive cells in aged hearts co‐localized with CD31, confirming that endothelial cells represent the predominant senescent population in the aging heart (Figure 1H,I, Figure S1F,G). Collectively, these results indicate that endothelial cells represent the predominant senescent cell population in the aged heart.

HSPA12B is a member of the HSP70 family of proteins that is predominantly expressed in endothelial cells (Steagall et al. 2006). Our previous studies have shown that endothelial cell HSPA12B plays a critical role in regulating angiogenesis and modulating immune responses (Tu et al. 2020; Zhang et al. 2020). To explore the potential role of HSPA12B in cardiac aging, we analyzed publicly available single‐cell RNA sequencing datasets and observed a significant reduction in HSPA12B expression in cardiac endothelial cells from aged humans and mice (Figure 2A,B) (Litviňuková et al. 2020). Consistently, cardiac endothelial cells isolated from aged mice also exhibited markedly lower HSPA12B expression compared to those from young mice (Figure 2C). To determine whether reduced endothelial HSPA12B expression contributes to age‐related cardiac dysfunction, we generated endothelial‐specific HSPA12B‐deficient (eHSPA12B^−/−) mice and assessed their cardiac phenotype during aging (Figure S2A). Remarkably, by 16 months of age, eHSPA12B^−/− mice exhibited pronounced cardiac fibrotic remodeling and significant functional decline compared to age‐matched wild‐type (WT) controls (Figure 2D,G,H and Figure S2B). Notably, these pathological features typically appeared in WT mice only at 24 months of age or older, suggesting that HSPA12B deficiency accelerates cardiac aging.

We next evaluated whether HSPA12B deficiency contributes to age‐related endothelial senescence. Hearts from eHSPA12B^−/− mice showed significantly increased endothelial cell senescence as indicated by elevated expression of the senescence markers p21, p16, and β‐gal (Figure 2E and Figure S2C,D), along with increased levels of SASP cytokines and chemokines compared to WT controls (Figure 2F). To further define the role of HSPA12B in endothelial senescence, we transfected human umbilical vein endothelial cells (HUVECs) with control siRNA (siRNA‐Con) or siRNA targeting HSPA12B (siRNA‐HSPA12B) and exposed them to from young (3–5 m) and aged (22–24 m) mice for 24 h. Knockdown of HSPA12B exacerbated aged serum–induced endothelial senescence, as evidenced by increased expression of p16, p21, and acetylated p53, along with enhanced SA‐β‐gal staining (Figure 2K,L, Figure S3D,E). Furthermore, aged HUVECs (P14) transfected with siRNA targeting HSPA12B exhibited accelerated replicative senescence, further supporting the link between reduced HSPA12B expression and enhanced endothelial cell senescence (Figure 2I,J and Figure S3A–C). Collectively, these findings demonstrate that reduced HSPA12B expression exacerbates endothelial cell senescence, which contributes to age‐related cardiac dysfunction.

The cGAS–STING signaling pathway plays a central role in sensing cytosolic DNA and driving cellular senescence, particularly in the context of aging (Glück et al. 2017; Guo et al. 2021; Liu et al. 2023). We first examined the activation status of this pathway in cardiac endothelial cells isolated from young and aged mice, as well as in young (passage <P6) and aged (passages P12–16) HUVECs. Heart tissues and endothelial cells from aged mice exhibited increased phosphorylation of STING, TANK‐binding kinase 1 (TBK1), and Interferon regulatory factor 3 (IRF3) compared to young controls, indicating activation of the pathway (Figure 3A and Figure S4A–C). Similarly, aged HUVECs showed elevated levels of phosphorylated STING, TBK1, and IRF3, along with increased expression of downstream target genes, Interferon alpha and beta (IFNα and IFNβ) (Figure 3B,C and Figure S4D).

Mitochondrial DNA (mtDNA) is a key activator of cGAS–STING signaling, and mitochondrial dysfunction is a well‐recognized hallmark of aging (He et al. 2022; Jiménez‐Loygorri et al. 2024; Zhong et al. 2022). To assess whether cytosolic accumulation of mtDNA occurs in aged endothelial cells, we quantified cytosolic DNA levels in young and aged HUVECs using quantitative PCR. Compared to young cells, aged HUVECs exhibited a significant increase in cytosolic mtDNA (Figure 3E). Consistently, immunofluorescent staining for double‐stranded DNA (dsDNA) revealed a marked accumulation of cytosolic dsDNA in aged HUVECs (Figure 3D), supporting enhanced mtDNA leakage into the cytosol during aging. We further evaluated cGAS expression in endothelial cells. Aged HUVECs showed significantly increased levels of cGAS compared with young cells (Figure S4E). However, modulation of HSPA12B (via Ad‐HSPA12B overexpression or si‐HSPA12B knockdown) did not significantly alter cGAS expression (Figure S4F,G). In contrast, HSPA12B deficiency specifically enhanced downstream signaling, as evidenced by increased levels of both total and phosphorylated STING, phosphorylation of TBK1, and IRF3, and upregulation of IFNα and IFNβ expression (Figure 3F,G). Together, these findings demonstrate that reduced HSPA12B expression exacerbates aging‐related cGAS–STING pathway activation in endothelial cells and likely contributes to the enhanced senescence phenotype observed during vascular aging.

To further investigate the role of HSPA12B in regulating age‐associated cGAS–STING activation and endothelial senescence, we increased HSPA12B expression using an adenoviral vector (Ad‐HSPA12B) in replicative‐aged HUVECs (Figure S3F). Ad‐GFP was used as a control. In parallel, young HUVECs were transduced with Ad‐HSPA12B for 24 h and subsequently exposed to aged mouse serum to mimic systemic aging. As shown in Figure 3H,I, increased expression of HSPA12B significantly attenuated activation of the cGAS–STING pathway under both replicative aging and aged serum conditions. This was evidenced by reduced phosphorylation of STING, TBK1, and IRF3 compared to control groups.

We next examined whether HSPA12B overexpression could mitigate endothelial senescence in the context of replicative aging and exposure to aged serum. As shown in Figure 3, increased HSPA12B expression significantly reduced the expression of key senescence markers, including p16, p21, and acetylated p53 (Figure 3J,K and Figure S3J,K). Furthermore, SA‐β‐galactosidase staining was decreased in cells with increased expression of HSPA12B compared to GFP controls (Figure S3G–J). To further investigate the role of STING activation in HSPA12B deficiency‐induced endothelial cell senescence, we inhibited STING activation with C‐176 in HUVECs with and without HSPA12B knockdown by its specific siRNA. As shown in Figure S4H–I, HSPA12B knockdown led to increases in both total and phosphorylated STING, as well as phosphorylation of TBK1 and IRF3. These changes were accompanied by elevated levels of senescence markers p16 and p21. Inhibition of STING activation reduced this pathway activation and attenuated cellular senescence caused by HSPA12B knockdown. Together, these findings suggest that HSPA12B exerts protective effects against age‐related STING activation, which contributes to endothelial senescence.

We then asked whether reducing STING activation could rescue age‐related endothelial senescence and cardiac dysfunction caused by HSPA12B deficiency in vivo. To test this, we treated aged WT and eHspa12b^−/− mice with the STING inhibitor C‐176 (Chen et al. 2025) every other day for 1 month. Cardiac function was assessed via echocardiography. C‐176 treatment significantly improved ejection fraction (EF) and fractional shortening (FS) in both aged WT and eHSPA12B^−/− mice compared with vehicle‐treated controls (Figure 4A,B).

To further confirm whether C‐176 effectively inhibits STING pathway activation in cardiac endothelial cells, we performed immunofluorescent co‐staining of phosphorylated STING (p‐STING), TBK1 (p‐TBK1), and IRF3 (p‐IRF3) with the endothelial marker CD31. As shown in Figure S5, compared with aged WT mice, eHSPA12B^−/− mice displayed strong activation of the STING pathway within cardiac endothelial cells, as indicated by increased phosphorylation of STING, TBK1, as well as elevated nuclear levels of p‐IRF3. However, C‐176 treatment significantly attenuated this activation in cardiac endothelial cells in both aged WT and eHSPA12B^−/− mice.

To assess whether STING inhibition reduces age‐related cardiac endothelial senescence in vivo, we performed immunofluorescent co‐staining of the endothelial marker CD31 with the senescence markers p21 and p16. As shown in Figure 4C,D and Figure S6A,B, C176 treatment markedly reduced the number of senescent endothelial cells in both aged (22–24 months) WT and eHSPA12B^−/− mice, indicating that STING inhibition alleviates endothelial senescence in aged mice.

To further determine whether STING activation directly contributes to endothelial senescence, HUVECs were treat with C‐176 in the context of replicative aging and aged serum exposure. In aged HUVECs, C‐176 treatment significantly reduced the expression of senescence markers p16, p21, and acetylated p53 (Figure 4E,F and Figure S6E,F). SA‐β‐galactosidase staining also revealed a substantial reduction in senescent cell burden following STING inhibition (Figure 4G,H and Figure S6C,D). Furthermore, quantitative PCR analysis demonstrated downregulation of key SASP‐associated cytokines and chemokines, further supporting the anti‐senescent effect of STING inhibition in aged endothelial cells (Figure S6G).

Together, these results demonstrate that inhibition of STING alleviates age‐related cardiac endothelial senescence and preserves cardiac function in both WT and HSPA12B‐deficient mice, highlighting the therapeutic potential of targeting STING signaling in vascular aging.

To elucidate the mechanism by which HSPA12B regulates cGAS–STING activation, we first examined STING protein expression in aged mouse hearts and senescent endothelial cells. STING protein levels were significantly elevated in both, and this increase was further augmented by HSPA12B deficiency (Figure 3A,B,F). In contrast, overexpression of HSPA12B significantly reduced STING protein levels (Figure 3H,I). We next investigated whether HSPA12B regulates STING transcriptionally. qPCR analysis revealed no significant differences in STING mRNA levels following either HSPA12B knockdown or overexpression, suggesting that HSPA12B modulates STING expression at a post‐transcriptional level (Figure 5A).

Given that STING protein turnover is regulated by SEL1L/HRD1‐mediated ER‐associated degradation (ERAD) (Ji et al. 2023), we examined whether HSPA12B affects the expression of SEL1L and HRD1. Both SEL1L and HRD1 were downregulated in aged endothelial cells, and their expression was further reduced in cells with HSPA12B knockdown (Figure 5B,J). In contrast, HSPA12B overexpression significantly upregulated SEL1L and HRD1 expression (Figure 5C,H).

As SEL1L and HRD1 are transcriptional targets of XBP1 (Kaneko et al. 2007; Yamamoto et al. 2008), we hypothesized that HSPA12B may influence XBP1 activity. Structural modeling using AlphaFold predicted a direct interaction between HSPA12B and XBP1, which we validated by co‐immunoprecipitation, confirming a physical interaction (Figure 5D,G,I).

These results suggest that HSPA12B enhances XBP1 transcriptional activity, thereby upregulating SEL1L and HRD1 expression to promote ERAD‐mediated degradation of STING. This regulatory mechanism reveals a previously unrecognized role for HSPA12B in limiting aberrant cGAS–STING signaling activation and preserving endothelial homeostasis during aging.

To further validate the role of ERAD‐mediated STING degradation in the protective effects of HSPA12B against age‐related cGAS–STING activation and endothelial senescence, we performed both genetic (siRNA‐mediated) and pharmacological inhibition of ERAD in aged endothelial cells. We first silenced SEL1L, an adaptor protein for the ERAD E3 ubiquitin ligase Hrd1 (Ji et al. 2023; Lin et al. 2024), in aged HUVECs using siRNA, followed by transduction with either Ad‐HSPA12B or Ad‐GFP as a control. As shown in Figures 3I and 5D, HSPA12B overexpression increased SEL1L expression and attenuated cGAS–STING activation, as evidenced by decreased phosphorylation of STING, TBK1, and IRF3, compared to the Ad‐GFP control group. However, knockdown of SEL1L abolished the ability of HSPA12B to suppress cGAS‐STING signaling, leading to a significant increase in STING expression and phosphorylation (Figure 6A). These results suggest that SEL1L is essential for HSPA12B‐mediated STING degradation.

Consistent with these findings, pharmacological inhibition of ERAD using the HRD1 inhibitor LS‐102 (Fujita et al. 2015) in aged HUVECs led to a significant increase in STING protein levels and abolished the suppressive effects of HSPA12B on cGAS–STING pathway activation (Figure 6B). Importantly, both SEL1L knockdown and ERAD inhibition reversed the anti‐senescent effects of HSPA12B in aged HUVECs. Specifically, we observed increased expression of senescence markers p16, p21, and acetylated p53, enhanced SA‐β‐galactosidase activity, and elevated p21‐positive staining in cells treated with ERAD‐disrupting interventions despite HSPA12B overexpression (Figure 6C–F, Figures S7A–D and S8A,B). Together, these findings indicate that ERAD‐mediated degradation of STING is critical for the protective effects of HSPA12B against aberrant STING activation and endothelial senescence during aging.

XBP1 is a key regulator of ER‐associated degradation (ERAD) by promoting the expression of SEL1L and HRD1 (Kaneko et al. 2007; Yamamoto et al. 2008). To investigate the role of XBP1 in age‐related cGAS–STING activation and endothelial senescence, aged HUVECs were treated with the XBP1 activator IXA4 (Cabral‐Miranda et al. 2022; Sheng et al. 2019) (Figure S8C,D). IXA4 treatment significantly upregulated SEL1L and HRD1 expression and attenuated cGAS–STING activation, as indicated by decreased phosphorylation of STING, TBK1, and IRF3 (Figure 7A,B and Figure S8F). Additionally, IXA4 markedly attenuated endothelial senescence, as demonstrated by decreased levels of p16, p21, and acetylated p53 (Figure 7C,D), as well as reduced p21 immunostaining and SA‐β‐galactosidase activity compared with vehicle‐treated controls (Figure 7E–G).

To further determine whether XBP1 is required for the protective effects of HSPA12B, aged HUVECs were treated with the XBP1 inhibitor MKC8866 (Sheng et al. 2019) following Ad‐HSPA12B transduction. Inhibition of XBP1 abrogated the HSPA12B mediated upregulation of SEL1L and HRD1 and abolished its suppressive effects on STING activation (Figure 7H,I). Furthermore, XBP1 inhibition negated the anti‐senescent effects of HSPA12B, leading to increased expression of p16, p21, and acetylated p53, as well as elevated p21 staining and SA‐β‐gal positivity (Figure 7H, Figure S8E,G–I). These findings indicate that XBP1 plays a critical role in mediating the protective effects of HSPA12B on age‐associated cGAS–STING activation and endothelial senescence and highlight XBP1 as an important regulator of the cGAS–STING signaling and vascular aging.

Wild‐type C57BL/6 mice were obtained from Jackson Laboratory and housed at the Institutional Animal Care and Use Facility at East Tennessee State University. Endothelial cell specific HSPA12B knockout mice (eHSPA12B^−/−) were generated by crossing HSPA12B^fl/fl with Tie2‐Cre (JAX, Stock: 008863). Male mice aged 3–5 months were categorized as young, while aged mice (22–24 months old) were bred in our laboratory and obtained from the National Institute on Aging (NIA). All animal care and experimental procedures were approved by the ETSU Committee on Animal Care and Use and adhered to NIH guidelines to ensure humane treatment. Mice were euthanized by isoflurane overdose followed by cervical dislocation. Hearts were excised immediately, rinsed in cold PBS, and transversely sectioned at the mid‐ventricular level to ensure consistent orientation across samples for histological and molecular analyses.

Both wild‐type (WT) and endothelial‐specific HSPA12B knockout (eHSPA12B^−/−) mice, including young and aged cohorts, were anesthetized, and heart tissues were harvested. Tissues were fixed in 4% paraformaldehyde (PFA) for 12 h at 4°C, washed three times with cold PBS, and then immersed in 30% sucrose overnight for cryoprotection. Samples were then embedded in OCT compound and sectioned at 10 μm thickness using a Leica CM3050S cryostat. For in vitro studies, HUVECs were cultured on chamber slides, washed with PBS, and fixed in 2% PFA for 20 min at room temperature. Both cryosections and HUVECs were washed three times with PBS, permeabilized with 0.5% Triton X‐100 in PBS for 20 min at room temperature, and then blocked with 10% goat serum for 1 h. Samples were incubated overnight at 4°C with the following primary antibodies: anti‐CD31 (endothelial marker, R&D, Cat #AF3628, 1:100), anti‐p16 (Cell Signaling Technology, Cat #80772S, 1:100), anti‐p21 (Cell Signaling Technology, Cat #2947S, 1:100), anti‐dsDNA (Novus, Cat #4565, 1:100), anti‐TOM20 (mitochondrial marker, Proteintech, Cat# 66777‐1‐Ig, 1:100), and anti‐STING (ABclonal, Cat #A21051, 1:100). Following primary incubation, sections and cells were washed once with PBST and twice with PBS, then incubated with appropriate Alexa Fluor–conjugated secondary antibodies for 1 h at room temperature. After three final PBST washes, samples were mounted with DAPI‐containing fluorescence mounting medium (Thermo Fisher Scientific, Cat #P36981) to stain nuclei. Confocal images were captured using a Leica TCS SP8 microscope and processed using ImageJ software (NIH) for quantitative and qualitative analysis.

Human umbilical vein endothelial cells (HUVECs) were obtained from the American Type Culture Collection (ATCC, PCS‐100‐013) and cultured in Vascular Cell Basal Medium (ATCC, PCS‐100‐030) supplemented with the Endothelial Cell Growth Kit–VEGF (ATCC, PCS‐100‐041). Cells were maintained at 37°C in a humidified incubator with 5% CO₂. HUVECs at passages below P6 were considered young, while those at passage P12 or higher were classified as aged. For treatment experiments, HUVECs were exposed to the following stimuli and inhibitors, either alone or in combination as indicated: Ad‐GFP (Abm, Cat: A000541, MOI: 10), Ad‐HSPA12B (Abm, Cat: A123995, MOI: 10), C‐176 (STING inhibitor, MCE, Cat: HY‐112906, 2.5 μM), LS‐102 (HRD1 inhibitor, MCE, Cat: HY‐135844, 10 μM), MKC8866 (XBP1 inhibitor, MCE, Cat: HY‐104040, 10 μM), IXA4 (MCE, Cat: HY‐139214, 10 μM), siRNA‐control (QIAGEN, Cat: 1022076, 40 nM), siRNA‐HSPA12B (QIAGEN, Cat: SI04165637, 40 nM), and siRNA‐SEL1L (QIAGEN, Cat: SI02664494, 40 nM). As well as serum from young or aged wild‐type mice (50 μL/mL). The serum used for cell treatments was collected from young (3–5 months) and aged (22–24 months) WT mice during tissue collection, following cardiac functional assessment by echocardiography. Aged mice exhibited cardiac aging phenotypes, including endothelial cell senescence, fibrosis, and cardiac dysfunction. For each treatment, serum from 6 to 10 mice within the same group was pooled to ensure consistency. Treatments were administered for the indicated durations to evaluate changes in signaling pathways and expression of senescence‐associated markers.

Senescence‐associated β‐galactosidase (SA‐β‐Gal) activity was assessed using commercial staining kits (Cell Signaling Technology, Cat# 9860S) following the manufacturers' protocols.

For immunoprecipitation (IP) assays, HUVECs were harvested, washed with PBS, and lysed on ice for 15 min in lysis buffer containing [1% NP‐40, 150 mM NaCl, 25 mM Tris–HCl pH 7.5, protease/phosphatase inhibitors]. Cell lysates were pre‐cleared with Protein G Magnetic Beads for 1 h at 4°C to remove nonspecific binding proteins. Following centrifugation to remove the beads, the cleared lysates were incubated overnight at 4°C with the indicated primary antibodies. Immune complexes were captured by incubation with fresh Protein G Magnetic Beads for 1 h at 4°C and collected by centrifugation. The immunoprecipitants were washed three times (5 min each) with wash buffer and separated by SDS‐PAGE for further analysis.

For Western blotting, protein was extracted from heart tissues, and in vitro cultured cells were performed using RIPA lysis buffer (Thermo Fisher Scientific, Cat: 89900) supplemented with protease and phosphatase inhibitors (Sigma‐Aldrich, Cat: PPC2020). Protein concentrations were measured using a BCA protein assay kit (Thermo Fisher Scientific, Cat: 23223). Equal amounts of protein were separated by electrophoresis on Bis‐Tris protein gels and transferred onto nitrocellulose membranes. Membranes were blocked with 5% BSA in TBS containing 0.5% Tween‐20 (TBST) for 1 h at room temperature, followed by incubation with specific primary antibodies overnight at 4°C. After washing with TBST, membranes were incubated with HRP‐conjugated secondary antibodies (Cell Signaling Technology, Cat: 7074) for 1 h at room temperature, followed by additional washes with TBST to remove unbound antibodies. Protein bands were visualized using the SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Fisher Scientific, Cat: 34096), optimized for detecting low‐abundance targets, and SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo Fisher Scientific, Cat: 34580) for higher‐abundance targets. Imaging was conducted with the G:Box Chemi gel documentation system (GeneSys Version: 1.8.10.0), and band intensities were quantified using Image J software.

The following primary and secondary antibodies were used in this study: Anti‐p21 (Cat: EPR18021), anti‐p16INK4a (Cat: EPR20418) were purchased from Abcam. Anti‐CD31 (Cat: 50274) was purchased from Biosciences. Anti‐STING (Cat: 13647S), anti‐p‐STING (Cat: 72971S), anti‐TBK1 (Cat: 3504S), anti‐p‐TBK1 (Cat: 5483S), anti‐IRF3 (Cat: 4302S), anti‐p‐IRF3 (Cat: 4947S), anti‐p21 (Cat: 2947S) were purchased from Cell Signaling Technology. Anti‐p21 (Cat: A22460), anti‐p16 (Cat: A23882), anti‐A‐p53 (Cat: A19836), anti‐SEL1L (Cat: A26877), anti‐HRD1 (Cat: A2605) were purchased from Abclonal. Anti‐TOM20 (Cat: 66777–1‐Ig), anti‐XBP1S (Cat: 24168–1‐AP) were purchased from Proteintech. All primary antibodies were validated for species cross‐reactivity and used at a dilution of 1:100 for immunofluorescence or 1:1000 for Western blotting.

Alexa Fluor 488 donkey anti‐rabbit IgG (H + L) (Cat: A‐21206), Alexa Fluor 488 donkey anti‐mouse IgG (H + L) (Cat: A‐10680), Alexa Fluor 488 donkey anti‐rat IgG (H + L) (Cat: A‐21208), and Alexa Fluor 594 donkey anti‐rabbit IgG (H + L) (Cat: SA5‐10080) were purchased from Invitrogen. HRP‐conjugated goat anti‐rabbit IgG (H + L) (Cat: 7074) and HRP‐conjugated goat anti‐mouse IgG (H + L) (Cat: 7076) were purchased from Cell Signaling Technology. All secondary antibodies were validated for species cross‐reactivity and used at a dilution of 1:500 for immunofluorescence or 1:5000 for Western blotting.

Total RNA was isolated from mouse heart tissues and cultured cells using RNAzol RT reagent (Molecular Research Center, Cat: RN 190) according to the manufacturer's protocol. cDNA was synthesized from 0.1 to 1 μg of total RNA using the High‐Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific, Cat: 4368814). Quantitative real‐time PCR (qRT‐PCR) was performed using SYBR Select Master Mix on a Bio‐Rad CFX96 real‐time PCR detection system. The following PCR primers were used: Human genes: CDKN1A (forward, 5′‐TGTCCGTCAGAACCCATGC‐3′; reverse, 5′‐AAAGTCGAAGTTCCATCGCTC‐3′), CDKN2A (forward, 5′‐GGAGGCCGATCCAGGTCAT‐3′; reverse, 5′‐ATGGTTACTGCCTCTGGTGC‐3′), IFNA1 (forward, 5′‐GCCTCGCCCTTTGCTTTACT‐3′; reverse, 5′‐CTGTGGGTCTCAGGGAGATCA‐3′), IFNB1 (forward, 5′‐ATGACCAACAAGTGTCTCCTCC‐3′; reverse, 5′‐GGAATCCAAGCAAGTTGTAGCTC‐3′), CXCL2 (forward, 5′‐AGATCAATGTGACGGCAGGG‐3′; reverse, 5′‐TCTCTGCTCTAACACAGAGGGA‐3′), CXCL1 (forward, 5′‐TGCTGCCACTAATGCTGATGT‐3′; reverse, 5′‐CTCAGGAACCAATCTTTGCACT‐3′), IL1A (forward, 5′‐TGGTAGTAGCAACCAACGGGA‐3′; reverse, 5′‐ACTTTGATTGAGGGCGTCATTC‐3′), STING1 (forward, 5′‐GGCCCGGATTCGAACTTACA‐3′; reverse, 5′‐TCATCTGCAGGTTCCGCTG‐3′), SYVN1 (forward, 5′‐GCTCACGCCTACTACCTCAAA‐3′; reverse, 5′‐GCCAGACAAGTCTCTGTGACG‐3′), SEL1L (forward, 5′‐AAACCAGCTTTGACCGCCAT‐3′; reverse, 5′‐GTCATAGGTTGTAGCACACCAC‐3′), XBP1 (forward, 5′‐CCCTCCAGAACATCTCCCCAT‐3′; reverse, 5′‐ACATGACTGGGTCCAAGTTGT‐3′). Mouse genes: Cdkn1a (forward, 5′‐CCTGGTGATGTCCGACCTG‐3′; reverse, 5′‐CCATGAGCGCATCGCAATC‐3′), Cdkn2a (forward, 5′‐CGCAGGTTCTTGGTCACTGT‐3′; reverse, 5′‐TGTTCACGAAAGCCAGAGCG‐3′), Cxcl2 (forward, 5′‐ATTCTGTGACCATCCCCTCAT‐3′; reverse, 5′‐TGTATGTGCCTCTGAACCCAC‐3′), Il1a (forward, 5′‐CGAAGACTACAGTTCTGCCATT‐3′; reverse, 5′‐GACGTTTCAGAGGTTCTCAGAG‐3′). Primer specificity was confirmed by melting curve analysis showing single distinct peaks.

Mitochondria were isolated using the Cell Mitochondria Isolation Kit (Thermo Fisher Scientific, Cat: 89874) according to the manufacturer's instructions. DNA was extracted from fractionated cytosolic fractions or isolated mitochondria using the DNeasy Blood and Tissue Kit (Qiagen, Cat: 69504) following the manufacturer's instructions. Mitochondrial DNA (mtDNA) levels were quantified by qRT‐PCR using specific primers for Cytochrome c oxidase I (forward: 5′‐GCCCCAGATATAGCATTCCC‐3′, reverse: 5′‐GTTCATCCTGTTCCTGCTCC‐3′). Results were normalized to 18S rRNA levels (forward: 5′‐TAGAGGGACAAGTGGCGTTC‐3′, reverse: 5′‐CGCTGAGCCAGTCAGTGT‐3′) to account for variation in sample input (Li et al. 2025).

All experiments were performed at least three independent times and representative data are shown. Statistical analysis between two groups was performed by a two‐tailed Student's t‐test, while one‐way ANOVA with Tukey's post hoc test was used for multiple comparisons. A p‐value less than 0.05 was considered statistically significant. Data are expressed as mean ± SD. Analyses were performed using GraphPad Prism 8.4.3 software.