Jaceosidin Attenuates Sepsis-Induced Myocardial Dysfunction by Promoting SIRT2-Mediated Inhibition of Histone H3K18 Lactylation

Abstract

Background: Sepsis-induced myocardial dysfunction (SIMD) is a life-threatening complication with limited therapeutic options. Jaceosidin (JAC), a natural flavonoid from Folium Artemisiae Argyi, shows potential in cardiovascular diseases, but its role and mechanism in SIMD remain unclear. This study aims to investigate the protective effects of JAC against SIMD and explore the underlying molecular mechanisms. Methods: In vitro, AC16 human cardiomyocytes were stimulated with TNF-α and treated with JAC. Cell viability and apoptosis were assessed using CCK−8 and flow cytometry, respectively. Transcriptomic and metabolomic analyses were performed to identify altered pathways. Molecular docking evaluated JAC’s interaction with SIRT2. The SIRT2 inhibitor AGK2 was used to validate its role. Chromatin immunoprecipitation quantitative PCR (ChIP-qPCR) determined H3K18la enrichment on target gene promoters. In vivo, a murine SIMD model was established via LPS injection, and cardiac function was evaluated by echocardiography. Serum markers (cTnT, CK−MB) and myocardial lactylation levels were measured. Results: JAC significantly attenuated TNF-α−induced injury in AC16 cells by enhancing viability and reducing apoptosis. Multi-omics analyses revealed JAC suppressed glycolysis and lactate production. JAC specifically inhibited histone H3K18 lactylation (H3K18la), and molecular docking indicated strong binding affinity with SIRT2. AGK2 treatment reversed JAC-mediated suppression of H3K18la. ChIP-qPCR confirmed H3K18la directly regulates IL-6, BAX, and BCL-2 expression. In vivo, JAC improved cardiac function (LVEF, LVFS, LVDd, LVDs), reduced serum cTnT and CK−MB levels, and decreased myocardial H3K18la in LPS−treated mice. Conclusions: JAC alleviates SIMD by activating SIRT2, which inhibits H3K18la, thereby modulating inflammatory and apoptotic pathways. This study identifies JAC as a novel metabolic-epigenetic therapeutic agent for SIMD.

1. Introduction

Sepsis is a life-threatening organ dysfunction syndrome caused by a dysregulated host response to infection, with the heart being one of the key target organs highly vulnerable to such dysregulation and injury [1]. Sepsis-induced myocardial dysfunction (SIMD) is a frequent and severe complication in septic patients, associated with increased mortality [2]. Substantial evidence indicates that approximately 50% of septic patients exhibit signs of myocardial dysfunction [3]. The pathogenesis of SIMD is highly complex, entailing multifactorial synergism and collaborative effects among dysregulated inflammatory responses, mitochondrial dysfunction, calcium dyshomeostasis, apoptosis/pyroptosis, and metabolic derangements [4,5]. Notably, this multifactorial pathogenesis aligns with the traditional medicine concept of “toxic heat invading the heart collaterals”, wherein systemic infection triggers inflammatory storm and oxidative stress, leading to qi stagnation, blood stasis, and phlegm turbidity obstructing the heart vessels. Pathogen-associated molecular patterns (PAMPs), such as lipopolysaccharide (LPS)—an exotoxin from Gram-negative bacteria, serve as pivotal initial triggers [6]. They activate innate immune receptors on host immune cells, cardiomyocytes, endothelial cells, and other cell types, eliciting a robust systemic inflammatory response that generates abundant pro-inflammatory mediators [7,8]. As tissue damage progresses, endogenous damage-associated molecular patterns (DAMPs) are abundantly released [9]. These DAMPs synergize with PAMPs to further amplify inflammatory signaling [9], and induce significant cardiac dysfunction through both direct actions on cardiomyocytes−such as stimulating production of myocardial depressant factors (including TNF-α and IL-1β) and impairment of the cardiac microenvironment [10]. Importantly, injured cardiomyocytes themselves become a source of new DAMPs, thereby establishing a self-perpetuating cycle of persistent inflammatory stimulation and progressive myocardial injury [11]. Despite advances in modern critical care, specific therapies for SIMD remain scarce. Current management focuses on source control, fluid resuscitation, vasoactive agents, and selected anti-inflammatory strategies [12]. While these can partially stabilize hemodynamics and restore cardiac function, they fail to effectively reverse myocardial injury progression or significantly improve long-term prognosis. Many experimental or clinical cardioprotective agents demonstrate suboptimal efficacy. Thus, in−depth dissection of SIMD pathogenesis, identification of key druggable targets, and development of highly effective and safe therapeutics constitute an urgent unmet clinical and scientific need.

In recent years, monomers or extracts derived from traditional Chinese medicine (TCM) have demonstrated distinct advantages in modulating cardiovascular pathologies, garnering increasing recognition within the scientific community [13,14]. Their multi-target and multi−pathway properties hold substantial promise for regulating core pathological processes−including inflammation, oxidative stress, apoptosis, and metabolic dysregulation [15]. This polypharmacological approach stands in contrast to conventional single-target therapies, enabling simultaneous intervention across interconnected pathological axes that drive disease progression. Systematic investigation of TCM constituents not only facilitates novel therapeutic discovery but also provides important clues for understanding disease pathogenesis and identifying new targets [16].

Artemisia carvifolia has been historically employed in traditional medicine to treat “toxic−heat” syndrome, wherein “toxicity” conceptually equates to contemporary inflammatory pathology. Modern phytochemical studies confirm that this syndrome reflects systemic hyperinflammation driven by cytokine storms and oxidative burst. As the core anti−inflammatory constituent identified in this herb, Jaceosidin (JAC) is a natural flavonoid compound with demonstrable biological activities. Contemporary pharmacological research reveals that JAC exhibits broad−spectrum anti-inflammatory, antioxidant, antitumor, and antifibrotic effects. Studies demonstrate that JAC exerts cardioprotective effects by significantly ameliorating myocardial oxidative stress, suppressing inflammatory responses, and reducing cardiomyocyte apoptosis [17]. These findings suggest JAC’s potential value in mitigating inflammatory myocardial injury; however, its specific role in SIMD and underlying molecular mechanisms-particularly whether it involves emerging epigenetic regulatory mechanisms-remain elusive.

Accumulating evidence indicates that severe metabolic derangements during sepsis-particularly dysregulated glycolysis and lactate accumulation-represent not merely manifestations of energy imbalance but may directly participate in gene expression regulation and organ injury through modulating epigenetic modifications [3,18,19]. Mounting evidence highlights the crosstalk between metabolic reprogramming and newly emerging epigenetic regulation across diverse diseases, with lactylation-based modifications playing a pivotal role in inflammatory and cardiovascular pathologies [20,21]. Building on this paradigm, our integrated transcriptomic and metabolomic profiling revealed profound glycolytic pathway reprogramming in TNF-α and LPS induced septic cardiomyopathy models, accompanied by significant elevations of key metabolites lactate and acetate in both in vitro and in vivo systems. Importantly, lactate and acetate serve as essential substrates for lactylation, linking metabolic reprogramming to epigenetic alterations. Mechanistic investigations revealed that JAC’s cardioprotective effects are centrally linked to its potent suppression of sepsis-induced lactylation. This study therefore delineates JAC−mediated regulation of the metabolic and epigenetic axis in SIMD, which likely represents the pivotal mechanism underlying its myocardial preservation.

2. Results

2.1. In Vitro Protective Effects by JAC Against TNF-α-Induced Injury in AC16 Cells

JAC treatment demonstrated significant protection against TNF-α−induced cardiomyocyte injury in AC16 cells. Morphological analysis revealed severe cellular damage in TNF-α−treated groups, characterized by membrane rupture and cytosolic shrinkage (Figure 1a), whereas JAC treatment preserved normal cell architecture. CCK−8 assays indicated that 20 ng/mL TNF-α reduced cell viability, while 50 ng/mL treatment caused more pronounced reduction, confirming that 20 ng/mL was principally manifested in TNF-α−induced cardiomyocyte injury rather than nonspecific drug toxicity, with an IC₅₀ of 60.92 ng/mL (Figure 1b,c). Flow cytometry detected apoptosis levels in AC16 cells. The results indicated apoptosis in the majority of cardiomyocytes at both early and late stages. JAC treatment significantly attenuated TNF-α−induced apoptosis (Figure 1d,e). The CCK−8 assay demonstrated that within the tested concentration range (particularly at the 20 μM JAC concentration used in this study), JAC alone did not exhibit any significant toxic effects on AC16 cells, with no statistically significant difference in cell viability compared to the control group, and its half-maximal inhibitory concentration (IC₅₀) was 196.04 μM. (Figure 1f). Western blotting analysis further revealed that JAC treatment modulated expression of inflammatory and apoptotic proteins, indicating anti-inflammatory and anti−apoptotic effects (Figure 1g–m). Additionally, PCR assays showed altered expression levels of inflammation- and apoptosis−related gene expression after JAC administration, further evidencing its protective mechanism (Figure 1n–s). Collectively, these results demonstrate that JAC effectively protects AC16 cardiomyocytes against TNF-α−induced injury through regulation of inflammatory and apoptotic pathways.

2.2. Gene Expression Profiles of mRNA Sequences in JAC-Treated Cells Under TNF-α Induction

Transcriptome profiling of TNF-α−stimulated cells treated with JAC identified DEGs. A volcano map was constructed to visualize the DEGs between the Model vs. Control (Model vs. Control) and JAC vs. Model (JAC vs. Model) groups, with criteria of |log2 (fold change) (FC)| > 1 and false discovery rate (FDR) < 0.001 (Figure 2a,b). The Venn diagram revealed the intersection of DEGs from both comparisons, indicating the common genes affected by TNF-α induction and JAC treatment (Figure 2c). KEGG pathway enrichment analysis was performed to elucidate the biological pathways associated with these DEGs, with lower Q values suggesting higher degrees of enrichment (Figure 2d). Additionally, GO enrichment analysis was conducted to categorize the common DEGs into biological processes, cellular components, and molecular functions (Figure 2e). These analyses collectively provided elucidating the molecular mechanisms underpinning JAC’s therapeutic effects in modulating gene expression in response to TNF-α−induced myocardial injury.

2.3. JAC Remodels the Cellular Metabolic Profile and Alleviates Metabolic Dysfunction Through Gly-Colysis Regulation

To fully elucidate the molecular mechanisms of JAC’s metabolic regulation, we performed metabolomic analysis. Metabolomic profiling revealed profound alterations in cellular metabolites upon TNF-α induction and JAC treatment. The total ion flow plots provided an overview of the metabolic landscape (Figure 3a). PCA and OPLS-DA diagrams illustrated the distinct metabolic clustering between groups, reflecting the metabolic shifts induced by JAC (Figure 3b,c). Pathway enrichment analysis pinpointed key metabolic pathways, such as pyruvate metabolism and glycolysis/gluconeogenesis, with VIP > 1 indicating significant metabolic perturbations (Figure 3d). A heatmap imaged the differentially expressed metabolites (DEMs) between the JAC−treated and the Model (Figure 3e). Notably, JAC significantly suppressed the levels of acetic acid and lactic acid, as revealed by top differential metabolite analysis (Figure 3g,h). ECAR measurements and glycolysis detection further corroborated JAC’s impact on glycolytic activity and capacity (Figure 3f,i,j).

2.4. JAC’s Specific Inhibition of H3K18la and Its Impact on Metabolic-Epigenetic Coupling

Our investigation delved into the epigenetic mechanisms by which JAC exerts its effects. We found that JAC specifically inhibited intracellular lactate levels in AC16 cells (Figure 4a). Correlation analyses unveiled significant associations between lactate levels and the expression of IL-6, BAX, and BCL-2, with corresponding R² and p values underscoring these relationships (Figure 4b,d). Western blotting analyses screened pan lactylation expression levels and revealed that JAC selectively attenuated H3K18la without significantly affecting H3 and H4 histone acetylation sites (Figure 4e–m). These findings implicated H3K18la as a critical epigenetic modification targeted by JAC in the context of myocardial injury.

2.5. JAC Regulates SIMD Through SIRT2 Activation and H3K18la Inhibition

Molecular docking studies demonstrated that JAC exhibited optimal binding energy with SIRT2, suggesting a potential interaction (Figure 5a). Western blotting analyses confirmed that JAC treatment restored SIRT2 expression levels, which were diminished in the modeling process (Figure 5b,c). The introduction of the SIRT2 inhibitor AGK2 reversed JAC’s effects on H3K18la and SIRT2 expression, further validating the involvement of SIRT2 (Figure 5d,e). qPCR analysis also revealed that AGK2 impacted SIRT2 mRNA levels (Figure 5f). IF staining localized SIRT2 and H3K18la expression predominantly to the nucleus (Figure 5g). ChIP-PCR analyses provided evidence of direct regulatory relationships between the H3K18 site and genes such as BCL-2, IL-6, and BAX (Figure 5h). Overall, our results demonstrate that JAC activates SIRT2 to inhibit H3K18la, thereby attenuating SIMD.

2.6. JAC Rescues Cardiac Function and Mitigates Myocardial Injury in LPS-Induced In Vivo SIMD in Mice

In the in vivo component of our study, we employed a murine model of LPS−induced SIMD to evaluate JAC’s efficacy. The experimental design encompassed Control, Model (LPS−treated), and JAC treatment groups (Figure 6a). The chemical structure of JAC was presented, highlighting its molecular formula (C₁₇H₁₄O₇) and CAS number (18085−97−7) (Figure 6b). Echocardiographic assessments, including M-mode imaging, illustrated that JAC preserved cardiac function, as evidenced by improved LVEF and LVFS, and reduced LVDd and LVDs values (Figure 6c,d). Histopathological examination via HE staining revealed that JAC mitigated LPS−induced myocardial injury (Figure 6e). TUNEL staining revealed significantly decreased apoptotic cell populations within myocardial tissues of JAC−treated mice compared to the SIMD model group (Figure 6f). Subsequent immunoblotting and PCR analyses confirmed attenuated expression of inflammatory mediators and apoptosis-associated markers in cardiac specimens from JAC−administered animals (Figure 6g–i). Consistent with these findings, serum quantification of biomarkers of cardiac injury cTnT and CK−MB demonstrated markedly lowered concentrations following JAC intervention (Figure 6j,k).

2.7. JAC Alleviates SIMD In Vivo by Modulating Lactate Metabolism and SIRT2-H3K18la Axis

In our in vivo validation studies, we Demonstrated a Considerable reduction in lactate levels in both cardiac tissue and blood serum following JAC treatment. Specifically, heart tissue lactate quantification revealed pronounced reductions in the JAC intervention group (Figure 7A). Similarly, blood lactate levels exhibited a pronounced reduction, further corroborating the lactate-lowering effects of JAC (Figure 7B). Western blotting analysis of cardiac tissue revealed a significant suppression of H3K18la in JAC–treated mice (Figure 7C). This result aligns with our in vitro observations, reinforcing the consistency of JAC’s mechanism in targeting H3K18la across experimental models. Pan lactylation analysis demonstrated a broad reduction in lactylation across cardiac proteins in JAC–treated mice (Figure 7D). Western blotting showed that JAC treatment restored SIRT2 expression levels, which were downregulated in the LPS–induced model group (Figure 7E), and the PCR results similarly confirmed this conclusion (Figure 7F). To further investigate the expression of SIRT2 in cardiac tissues, we performed IF staining on cardiac tissue sections. IF imaging demonstrated increased nuclear expression of SIRT2 following JAC treatment (Figure 7G), with co–localization analysis confirming that both SIRT2 and H3K18la co–localized within cardiomyocyte nuclei (Figure 7H). The in vivo results were highly consistent with our in vitro findings, confirming that JAC inhibits SIMD in vitro and in vivo by promoting SIRT2-mediated lactylation of histone H3K18.

3. Discussion

JAC, a naturally derived bioactive compound, has demonstrated therapeutic potential across multiple disease models. Previous studies revealed that JAC significantly suppresses LPS induced pulmonary inflammation while mitigating oxidative stress and tissue damage [22]. In metabolic regulation, this compound reduces oxidative stress and inflammation by activating insulin signaling pathways, effectively ameliorating metabolic dysregulation and organ injury [23]. In osteoarthritis models, JAC inhibits NF–κB signaling transduction through blocking IκB degradation, thereby attenuating inflammatory responses and retarding cartilage degeneration [24]. Furthermore, JAC exhibits potent metabolic modulatory effects in obesity models: reducing body weight, blood glucose, and insulin levels; improving glucose tolerance and insulin sensitivity; concurrently decreasing serum triglycerides and cholesterol. Its hepatoprotective mechanism involves suppressing hepatic lipid accumulation, downregulating lipogenic gene expression, and improving liver function parameters [25]. In cardiovascular protection, JAC operates through dual pathways [26]: activating the nuclear factor Nrf2 pathway to enhance antioxidant enzyme expression, reduce ROS and lipid peroxidation, thereby alleviating cardiomyocyte oxidative stress and apoptosis; while concurrently inhibiting NF–κB mediated inflammatory cascades to reduce proinflammatory cytokine levels and mitigate cardiac injury. This study uncovers a novel cardioprotective mechanism of JAC in SIMD, featuring specific targeting of the metabolic epigenetic axis through upregulating deacetylase SIRT2 to selectively suppress H3K18la. Compared to conventional broad spectrum anti-inflammatory agents, JAC demonstrates superior therapeutic advantages by precisely modulating sepsis associated metabolic derangements and epigenetic dysregulation.

During sepsis, significant alterations occur in the glycolytic pathway within both cardiomyocytes and macrophages [27]. PAMPs activate TLR4 signaling, triggering p38 MAPK pathway activation. This upregulates the key glycolytic regulator PFKFB3, driving accelerated glycolytic flux, which is critically associated with myocardial dysfunction. Notably, this metabolic perturbation instigates a pathogenic cascade wherein pathogen invasion activates the innate immune response, exacerbating inflammation and apoptosis in cardiomyocytes while concurrently disrupting their metabolic and functional integrity. The substantial release of inflammatory and pro-apoptotic factors induces a metabolic shift towards glycolysis in both cardiomyocytes and macrophages, resulting in lactate accumulation. Beyond its role as a metabolic byproduct of glycolysis, lactate serves important functions in modulating processes such as T helper cell and macrophage regulation [28]. Furthermore, lactate actively participates in regulating histone lactylation [20]. This multifaceted role of lactate provides a novel perspective for in–depth mechanistic analysis of the interplay between metabolic and epigenetic regulation in SIMD. In our study, profound alterations in the glycolytic pathway were observed in TNF-α and LPS–induced models of septic cardiomyopathy, characterized by significant elevations in key metabolites lactate and acetate. These findings align with previous reports emphasizing the central role of dysregulated glycolytic flux in the pathophysiology of sepsis. Dysregulation of metabolic pathways not only disrupts energy homeostasis within cardiomyocytes but also acts as a critical driver of inflammatory signaling and oxidative stress, thereby further exacerbating myocardial injury. By targeting metabolic reprogramming, JAC offers a novel therapeutic strategy to disrupt the vicious cycle of metabolic dysfunction and inflammation characteristic of SIMD.

Lactate is not only a metabolic waste product and prognostic marker in sepsis, but also a key regulatory molecule mediating immune imbalance and organ injury. Its pathological accumulation drives multi–organ injury by inducing histone lactylation [19]. On one hand, in sepsis-associated acute lung injury (ALI), p300–mediated H3K18la directly upregulates the methyltransferase METTL3, activating the ACSL4-dependent ferroptosis pathway leading to alveolar epithelial cell injury [29]. On the other hand, H3K18la levels positively correlate with disease severity [30], and targeted intervention can significantly improve survival rates. This study found that under pathological conditions induced by TNF-α/LPS, lactate accumulation driven by glycolytic disruption specifically promotes an increase in H3K18la, while H3K18ac remains stable, suggesting that lactate preferentially occupies modification sites and drives lactylation in the metabolic stress environment of SIMD. SIRT2, as a key delactylase, possesses structural adaptability in its substrate-binding domain for the lactyl group (long–chain fatty acyl group), conferring selective catalytic advantages [31]. Research shows that SIRT2 specifically inhibits the cuproptosis pathway by regulating METTL16 K229 lactylation [32]. Notably, SIRT2 prioritizes the processing of lactylation at the expense of its acetyl group removal capability, thereby avoiding perturbation of the physiological acetylation network. Based on this, this study found that unlike broad–spectrum epigenetic modulators, under inflammatory stress, JAC precisely and preferentially targets H3K18la by activating SIRT2 without disrupting the physiological acetylation network. This specificity avoids potential off–target effects, highlighting its therapeutic advantage for septic myocardial injury.

Lactylation, as a key intersection of metabolism and epigenetics, exhibits dual protective and pathogenic roles in cardiovascular diseases by regulating inflammatory balance, energy metabolism, and cellular remodeling [33,34]. Studies show that the extracellular matrix protein Lumican significantly promotes histone lactylation at H3K14 and H3K9 sites by enhancing glycolysis and lactate production, thereby driving the progression of aortic valve calcification lesions [13]. Conversely, targeted inhibition of the key lactylation enzyme p300 effectively reduces H3 histone lactylation levels and alleviates valvular calcification injury [20]. In atherosclerosis, lactylation improves plaque stability and the inflammatory microenvironment by inhibiting RUNX1 transcriptional activity and enhancing chromatin accessibility [21]. Notably, lactate accumulation in the myocardial hypoxic microenvironment triggers histone lactylation, activating the “lactate clock” mechanism to precisely coordinate anti-inflammatory responses and tissue repair signaling networks [35]. Our research indicates that in vitro, TNF-α stimulation activates the glycolytic pathway, leading to lactate accumulation, which subsequently drives H3K18la, exacerbating cardiomyocyte inflammation and apoptosis. JAC, by upregulating SIRT2 expression, specifically targets H3K18la, effectively reversing cardiomyocyte damage and alleviating SIMD. In vivo experiments similarly confirmed the protective effects of JAC, as it significantly improved cardiac function in a septic mouse model, reduced myocardial injury, and lowered levels of inflammation and apoptosis markers. These findings suggest JAC as a potential therapeutic agent that may offer new strategies for the clinical treatment of SIMD.

We comprehensively assessed the pharmacological effects of JAC on SIMD and mechanistically confirmed its functional involvement in a metabolic-epigenetic regulatory pathway through integrated in vivo and in vitro models. Our investigation revealed that lactate accumulation during SIMD drives pathological reprogramming of histone lactylation, with H3K18 identified as the pivotal modification site via Western blot screening. Significantly, JAC alleviated SIMD by modulating SIRT2–dependent delactylation to inhibit H3K18la, consequently suppressing transcriptional activation of pro-inflammatory IL-6 and pro–apoptotic BAX genes while restoring BCL-2 expression. This study establishes a novel mechanistic framework for JAC’s cardiovascular protection, positioning it as a promising therapeutic candidate for SIMD through targeting the newly identified SIRT2–H3K18la regulatory axis.

This study systematically reveals a novel mechanism by which JAC improves SIMD through the SIRT2–H3K18la axis, but it has certain limitations. Although the experimental models primarily employed TNF-α/LPS–induced AC16 cells and murine sepsis systems, these platforms inadequately recapitulate sepsis–specific immune-cardiomyocyte crosstalk, particularly the complex interaction networks between immune cells and cardiomyocytes. Future research should establish immune cell-cardiomyocyte co–culture systems or animal models to enhance pathological simulation comprehensiveness. Mechanistically, although AGK2 inhibition experiments confirmed the necessity of SIRT2, no cardiomyocyte specific SIRT2 gene knockout animal model was used to further validate its cardiac specific role. Subsequent studies could construct conditional knockout models to clarify SIRT2’s cell autonomous function. Additionally, while this study mainly focused on H3K18la regulation of BCL-2, BAX, and IL-6, histone lactylation may affect broader gene networks, such as metabolic reprogramming related genes. Future work could employ ChIP–seq technology to comprehensively map the genome-wide binding profile of H3K18la for deeper mechanistic understanding. Furthermore, the pharmacokinetic properties of JAC (e.g., cardiac tissue distribution, metabolic stability) remain unevaluated, which are critical unresolved issues for clinical translation planning. These limitations provide clear directions for future research, but the core conclusions of this study remain reliable, offering important theoretical foundations and experimental support for treating septic myocardial injury.

4. Materials and Methods

4.1. Cell Culture Protocols and Experimental Treatments

AC16 cells (Human-derived cardiomyocytes, NO: CL–0790, Procell Life Science & Technology Co., Ltd., Wuhan, China) were cultured in Minimum Essential Medium (MEM, NO: PM150411, Procell Life Science & Technology Co., Ltd.) supplemented with 10% fetal bovine serum (FBS; Cell Box, Changsha, China) under standard conditions (37 °C, 5% CO₂). Until the cells reached approximately 80% confluence, cells were subjected to concurrent stimulation with 20 ng/mL TNF-α (PeproTech, Cranbury, NJ, USA, NO: 300-01A) and 20 μM JAC for 24 h [36]. AGK2, a selective SIRT2 inhibitor, suppresses cellular proliferation and growth in a dose-dependent manner while exhibiting no cytotoxicity at low concentrations. To understand more about the role of SIRT2, we treated cells with AGK2 (MedChemExpress, Monmouth Junction, NJ, USA, Cat: HY-100578 CAS No: 304896–28–4). A statement to confirm that all methods were carried out in accordance with relevant guidelines and regulations.

4.2. Cell Viability Assay

AC16 cells were seeded in 96–well plates (5 × 10³ cells/well) and exposed to designated treatments. Cell viability was assessed using a CCK-8 (Cell Counting Kit-8) assay kit (NO: K1018, APExBIO, Houston, TX, USA) according to manufacturer’s protocol. Cells were incubated with CCK-8 reagent for 2 h at 37 °C, after which absorbance was measured at 450 nm using a microplate reader. Data from three independent biological replicates were normalized to untreated controls and analyzed by one-way ANOVA to determine statistically significant differences (p < 0.05) in cell viability across TNF-α or JAC treatment groups.

4.3. Flow Cytometric Analysis of Apoptosis

AC16 cells were seeded in 6-well plates and incubated with 20 μM JAC for 24 h. Subsequently, cells were harvested, subjected to Annexin V–FITC/propidium iodide (PI) double staining, and analyzed by flow cytometry.

4.4. RNA Sequencing (RNA-Seq)

Changes in cellular mRNA profiles across experimental treatments were investigated through quantitative RNA–seq methodology. Cellular material was harvested with TRIzol reagent (Vazyme, Nanjing, China, NO: R401-01) followed by RNA extraction. Purified RNA samples were subsequently transferred to BGI Ltd. (Wuhan, China) for RNA sequencing. Raw sequencing reads underwent computational processing using R software (version 3.5.1) to determine differentially expressed genes (DEGs). Enrichment analyses of identified DEGs were conducted for Gene Ontology (GO) annotations and KEGG pathway databases [37].

4.5. Gas Chromatography-Mass Spectrometer (GC–MS) Metabolome Analysis

AC16 cells were rinsed thrice with ice-cold PBS and lysed in pre–chilled HPLC-grade methanol (−80 °C). Cell lysates underwent three freeze–thaw cycles (liquid nitrogen/37 °C water bath, 10 min per cycle) followed by centrifugation (12,000× g, 4 °C, 10 min). The clarified supernatant was subsequently dried under nitrogen gas at 30 °C.

Dried extracts were derivatized with 80 μL methoxyamine hydrochloride (20 mg/mL in pyridine) at 37 °C for 90 min, followed by addition of 80 μL N,O–bis(trimethylsilyl)trifluoroacetamide (BSTFA) with 1% trimethylchlorosilane (TMCS). Samples were vortex–mixed (1 min), centrifuged (10,000× g, 1 min), and incubated at 70 °C for 1 h. Derivatized extracts were centrifuged (12,000× g, 4 °C, 10 min), and supernatants were transferred to GC vials.

GC–MS analysis was performed within 48 h using a Trace 1300 GC system coupled to ISQ QD mass spectrometer (Thermo Scientific, Waltham, MA, USA) equipped with a TG–5MS capillary column (30 m × 0.25 mm × 0.25 μm). Injection volume: 1 μL in splitless mode. At the final stage of the experiment, data processing was performed using the Chromatographic Fingerprint Similarity Evaluation System (2012 edition) and SIMCA-P+ 14.1 (Umetrics, Umeå, Sweden).

4.6. Real-Time Metabolic Flux Analysis Using Seahorse XF Technology

AC16 cell suspensions were seeded at 6 × 10⁴ cells/well in Matrigel™-precoated Seahorse XF96 cell culture microplates (250 μL/well) and incubated for 12 h at 37 °C/5% CO₂. For drug-treated groups, 20 μM JAC was administered after initial incubation, followed by continued culture for 24 h total. Prior to assay, cells were washed thrice with prewarmed (37 °C) XF assay medium (pH 7.4) and maintained in 500 μL/well of the same medium. Microplates were equilibrated for 60 min in a non–CO₂ incubator at 37 °C. 10 mM glucose, 1 μM oligomycin and 50 mM 2-DG were added to the cell culture plates. Extracellular acidification rate (ECAR) was measured over 100 min (3 measurement cycles per injection, 5 min mix/2 min wait/3 min measure). Following experimental completion, datasets were processed using Agilent Seahorse Wave analytical software (Seahorse Wave 2.6.1).

4.7. Lactate Acid Detection

Cardiac tissues (50 mg) were homogenized in 500 μL ice-cold Lactate Assay Buffer using a Polytron homogenizer. Homogenates were centrifuged at 12,000× g (4 °C, 5 min), and supernatants were collected for analysis. For cellular lactate measurement, AC16 cells grown in 6-well plates were trypsinized, washed twice with ice–cold PBS, and resuspended in 200 μL Lactate Assay Buffer per 1 × 10⁶ cells. Cell suspensions underwent ice–bath sonication (200 W, 3 s pulse/7 s interval, 30 cycles; Sonics Vibra–Cell VCX130) followed by centrifugation (12,000× g, 4 °C, 5 min).

Supernatants from tissues or cells were assayed using a Lactate Content Detection Kit (NO: KTB1100, ABBkine, Atlanta, GA, USA). Aliquots (50 μL) were incubated with 50 μL assay reagent at 37 °C for 30 min in the dark. Absorbance at 450 nm was measured.

4.8. Molecular Docking

The 3D structures of the compounds were retrieved from the PubChem database (https://pubchem.ncbi.nlm.nih.gov/ (accessed on 19 July 2025)). Preprocessing of these structures, including energy minimization and conversion to PDBQT format, was carried out using OpenBabel–2.3.2 and AutoDock4.2.6 tools. The crystal structure of the target protein was obtained from the UniProt database (https://www.uniprot.org/ (accessed on 19 July 2025)). To prepare the protein for docking, dehydration was performed to remove all water molecules, and the native ligand was extracted using PyMOL 2.4.0. The binding energy calculations and molecular docking simulations were performed using AutoDock. The docking process involved an automated search for the optimal binding conformation of each compound within the predefined binding site of the protein. The results were visualized and analyzed using PyMOL’s visualization tools to assess the interaction between the compounds and the target protein.

4.9. Immunofluorescence (IF) Assay

AC16 cells were seeded on glass coverslips at 1 × 10³ cells/coverslip. After 24 h incubation, cells were washed twice with PBS, fixed with 4% paraformaldehyde for 20 min at room temperature, and permeabilized with 0.2% Triton X–100/PBS for 10 min. Non–specific binding was blocked with 5% bovine serum albumin (BSA)/PBS for 30 min. Samples were incubated overnight at 4 °C with primary antibodies against: SIRT2 (1:200; Immunoway, San Jose, CA, USA) and H3K18la (1:200, Zen-Bio, Durham, NC, USA). After PBS washes, cells were incubated with goat anti-rabbit IgG and goat anti-mouse IgG secondary antibodies for 60 min at 37 °C in the dark. Nuclei were counterstained with 4′,6-diamidino–2–phenylindole for 5 min, and then imaged under microscope.

4.10. Chromatin Immunoprecipitation Quantitative Polymerase Chain Reaction (ChIP-qPCR) Assay

Assays were performed utilizing a ChIP Assay Kit (Beyotime, Wuhan, China, NO: P2078) in strict accordance with the manufacturer ’s instructions. In detail, cells were initially subjected to cross–linking with 1% formaldehyde to establish covalent linkages between DNA and associated proteins. Subsequently, the harvested cell lysates underwent sonication to yield chromatin fragments within the size range of 200–300 bp, ensuring optimal accessibility for downstream immunoprecipitation procedures. The prepared chromatin samples were then subjected to immunoprecipitation employing an anti-P65 antibody, with IgG serving as a negative control to assess background nonspecific binding.

In order to evaluate the enrichment of specific genomic regions, qPCR was conducted on the immunoprecipitated DNA samples. The qPCR primers were meticulously designed to specifically amplify the proximal promoter regions of the IL-6, BCL-2 and BAX genes, which harbor putative H3K18la binding sites. Following quantification, the percent input values derived from each experiment were normalized and converted into fold changes relative to an untreated control group. Ultimately, the average fold changes garnered from three independent experimental replicates were calculated and plotted to present the experimental outcomes in a comparative and quantifiable manner.

ChIP-qPCR primer sequences employed in this study were:

BAX: F: 5′-TCGCGATCGGCCTGGTTC-3′, R: 5′-TAAAGGGACAGCAACTGT-3′;

IL-6: F: 5′-TGGAACTGCCAGCGGCGG-3′, R: 5′-TCCTTTTTGTCCCCCGGG-3′;

BCL-2: F: 5′-TCGAGAGGAGTTATAATA-3′, R: 5′-GTTTTTAGACTTTCTTAT-3′;

4.11. Animal Model and Treatment

Male C57BL/6J mice (6–8 weeks old, 22 ± 2 g) were obtained from Hangzhou Ziyuan Experimental Animal Technology Co., Ltd., Hangzhou, China, (License SYXK(E) 2023-0067) and maintained under standardized conditions (24 ± 2 °C, 12 h light/dark cycle). All experimental protocols were approved by the Animal Ethics Committee of Hubei University of Chinese Medicine (Approval No. HUCMS 202311047) and emphasis on minimizing animal suffering and sample sizes. Animals were randomly assigned to three experimental groups (n = 6/group): normal control (Control), LPS (Model)–induced model, and JAC-treated. SIMD was induced via i.p. of 12 mg/kg LPS (Sigma, Cream Ridge, NJ, USA, USA, NO: L2880) [38]. Two hours post-modeling, the JAC-treated group received 80 mg/kg JAC (purity ≥ 98%, ChemFaces, Wuhan China, CFN90386; CAS 18085-97-7) dissolved in saline via intraperitoneal injection, while Control groups received equivalent saline volumes. The murine dosage was determined using validated interspecies dose translation (human dose [mg/kg] × 70 kg × 0.0026/0.02 kg) [39,40]. Twenty-four hours after LPS administration, euthanasia was performed by intravenous injection of sodium pentobarbital (100 mg/kg). After euthanasia, the excised hearts were perfused retrogradely via the aorta with ice-cold phosphate-buffered saline (PBS) to remove residual blood. For molecular analyses (Western blotting and qRT–PCR), whole hearts were flash-frozen in liquid nitrogen and pulverized to ensure homogeneity. For histological examinations (H&E, TUNEL) and immunofluorescence (IF), whole hearts were immersion–fixed in 4% paraformaldehyde, followed by standard paraffin embedding. Transverse sectioning was performed at the mid-ventricular level to ensure consistent anatomical orientation and inclusion of the left ventricular free wall, interventricular septum, and right ventricle in all analyses. All histological and IF assessments were conducted on sections obtained from this standardized anatomical plane. Terminal blood collection was executed via cardiac puncture, with plasma isolated following 2 h coagulation at room temperature and centrifugation (3000 rpm, 15 min, 4 °C), then stored at −80 °C. Excised hearts were perfused with ice-cold PBS to remove residual blood, with one portion immersion-fixed in 4% paraformaldehyde for histology and the remainder flash-frozen in liquid nitrogen for molecular analyses at −80 °C. A statement to confirm that all methods were carried out in accordance with relevant guidelines and regulations.

4.12. Mouse Echocardiography

Prior to tissue collection, transthoracic echocardiography was performed under isoflurane anesthesia. Following chest hair removal, mice were positioned supine on a temperature–controlled platform (37 °C) to maintain normothermia. A standardized layer of acoustic coupling gel was applied to the precordial region before cardiac imaging using the Vevo 2100 system (FUJIFILM VisualSonics, Inc., Toronto, ON, Canada). Comprehensive evaluation included two-dimensional, M–mode, and color Doppler imaging at the aortic valve level. To ensure objective assessment, operators were blinded to group assignments during image acquisition and analysis. Hemodynamic parameters-specifically left ventricular ejection fraction (LVEF), left ventricular fractional shortening (LVFS), Left ventricular end–diastolic internal diameter (LVDd), left ventricular end–systolic internal diameters (LVDs) and peak aortic flow velocity-were measured across three consecutive cardiac cycles with reported values representing averaged measurements. This protocol evaluated the effects of JAC on cardiac performance in the experimental model.

4.13. Histopathological Analysis

Cardiac tissues from mice were fixed in 4% paraformaldehyde, dehydrated through a graded ethanol series, embedded in paraffin, and sectioned at 5 μm thickness. TdT-mediated dUTP nick end labeling (TUNEL) staining and Hematoxylin and eosin (H&E) staining were subsequently performed to assess histopathological changes and early cardiomyocyte apoptosis as a means of evaluating the cardioprotective efficacy of JAC.

4.14. Enzyme Linked Immunosorbent Assay (ELISA)

Cardiac tissue and serum specimens were harvested from experimental mice. Serum levels of cardiac injury biomarkers—cardiac troponin T (cTnT) and creatine kinase–MB isoenzyme (CK–MB)—were measured using Elabscience kits (CK–MB: E–EL–M0355; cTnT: E–EL–M1801) per manufacturer’s protocols to evaluate JAC’s protective effects on myocardial tissue.

4.15. Western Blotting

For the preparation of protein samples, mice heart tissues were homogenized at 4 °C and collected using RIPA lysis buffer (Servicebio, Wuhan, China, NO: G2002-100ML). Similarly, AC16 cells cultured were harvested by scraping and collected using the same RIPA lysis buffer. The lysates were thoroughly lysed on ice, followed by centrifugation at 12,000× g for 10 min at 4 °C. The resulting supernatants were collected and stored at −20 °C until further use. For Western blotting analysis, the proteins were denatured by heating at 100 °C for 10 min. Gel electrophoresis was performed to separate the proteins based on their molecular weights, followed by transferring onto a membrane for immunoblotting. The membrane was incubated with the primary antibody at 4 °C overnight with gentle shaking. After washing the membrane three times with TBST buffer, the HRP–conjugated secondary antibody diluted in TBST was applied, and the membrane was incubated at 37 °C for 1 h. Following additional TBST washes to remove unbound antibodies, the protein bands were visualized using an ECL detection system. The band intensities were quantified and analyzed using Quantity One software V4.6.2.

The antibody for Western blotting used for the experiments are listed in

Table 1. Antibody for Western Blotting.

Antibody	Cat No	Producer
TNF-α	346654	ZenBio, Chengdu, China
IL-1β	660092	ZenBio, China
IL-6	347023	ZenBio, China
BAX	R380709	ZenBio, China
BCL-2	381702	ZenBio, China
Caspase-9	680095	ZenBio, China
Caspase-3	R22842	ZenBio, China
β-Actin	R51031	ZenBio, China
Pan Kla	PTM-1425	PTM BIO, Hangzhou, China
Anti-L-Lactyl-Histone H4 (Lys16)	PTM-1417RM	PTM BIO, China
Histone H4	PTM-1015RM	PTM BIO, China
Anti-L-Lactyl-Histone H4 (Lys5)	PTM-1409	PTM BIO, China
Anti-L-Lactyl-Histone H4 (Lys12)	PTM-1411	PTM BIO, China
Histone H3	PTM-6613	PTM BIO, China
Anti-L-Lactyl-Histone H3 (Lys14)	PTM-1414RM	PTM BIO, China
Anti-L-Lactyl-Histone H3 (Lys18)	PTM-1406RM	PTM BIO, China
Anti-L-Lactyl-Histone H3 (Lys9)	PTM-1419RM	PTM BIO, China
Anti-L-Lactyl-Histone H3 (Lys23)	PTM-1413RM	PTM BIO, China
Anti-L-Lactyl-Histone H3 (Lys56)	PTM-1421RM	PTM BIO, China
Anti-L-Lactyl-Histone H3 (Lys27)	PTM-1428	PTM BIO, China
Anti-Acetyl-Histone H4 (Lys16)	PTM-122	PTM BIO, China
Anti-Acetyl-Histone H4 (Lys5)	PTM-119	PTM BIO, China
Anti-Acetyl-Histone H4 (Lys12)	PTM-121RM	PTM BIO, China
Anti-Acetyl-Histone H3 (Lys14)	PTM-113RM	PTM BIO, China
Anti-Acetyl-Histone H3 (Lys18)	PTM-114RM	PTM BIO, China
Anti-Acetyl-Histone H3 (Lys9)	PTM-112RM	PTM BIO, China
Anti-Acetyl-Histone H3 (Lys23)	PTM-115RM	PTM BIO, China
Anti-Acetyl-Histone H3 (Lys56)	PTM-162	PTM BIO, China
Anti-Acetyl-Histone H3 (Lys27)	PTM-116RM	PTM BIO, China
SIRT7 Polyclonal antibody	12994-1-AP	Proteintech, Rosemont, IL, USA
SIRT4 Rabbit pAb	862208	ZenBio, China
SIRT2 Rabbit mAb	R25722	ZenBio, China
SIRT6 Rabbit mAb	R381408	ZenBio, China
SIRT3 Rabbit mAb	R25724	ZenBio, China
SIRT5 Rabbit mAb	R381473	ZenBio, China
HDAC8 Rabbit mAb	R381478	ZenBio, China
HDAC4 Rabbit pAb	R381467	ZenBio, China
KAT8 Rabbit mAb	R383056	ZenBio, China
KAT1 Rabbit mAb	R383080	ZenBio, China
KAT5 Mouse mAb	221268	ZenBio, China
KAT2A Rabbit mAb	R24422	ZenBio, China

.

4.16. Quantitative Real-Time PCR (qRT–PCR)

Primer sequences were designed using the NCBI Primer-BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi (accessed on 23 July 2025)) tool. Total RNA was extracted from mouse heart tissues and AC16 cells, followed by reverse transcription to synthesize cDNA. The target genes were then amplified using a real-time PCR system. The relative mRNA expression levels were calculated using the 2−ΔΔCt method and normalized to the internal control β-actin. The primers for the target genes were synthesized by Beijing Qingke Biotechnology Co., Ltd. (Wuhan, China).

The sequences of the primers are listed in

Table 2. Primer sequences for real-time RT-PCR.

Gene	Primer (5′-3′)
H-IL-1β	F: AGCACCTTCTTTCCCTTCATCTT R: CACCACTTGTTGCTCCATATCCT
M-IL-1β	F: CAGCACATCAACAAGAGCTTCAG R: GAGGATGGGCTCTTCTTCAAAGA
H-IL-6	F: AACATGTGTGAAAGCAGCAAAGA R: CTCTGGCTTGTTCCTCACTACTC
M-IL-6	F: GTATGAACAACGATGATGCACTTG R: CTCTCTGAAGGACTCTGGCTTTG
H-TNF-α	F: CTGTAGCCCATGTTGTAGCAAAC R: TTGAAGAGGACCTGGGAGTAGAT
M-TNF-α	F: GACCCTCACACTCAGATCATCTT R: CCTTGAAGAGAACCTGGGAGTAG
H-BAX	F: GCTTCAGGGTTTCATCCAGGATC R: ATCCTCTGCAGCTCCATGTTACT
M-BAX	F: ATCCTCTGCAGCTCCATGTTACT R: TCATCCTCTGCAGCTCCATATTG
H-Bcl-2	F: GGATTGTGGCCTTCTTTGAGTTC R: CTTCAGAGACAGCCAGGAGAAAT
M-Bcl-2	F: GGATTGTGGCCTTCTTTGAGTTC R: CTTCAGAGACAGCCAGGAGAAAT
H-caspase-9	F: TGTCCTACTCTACTTTCCCAGGT R: CCCTTTCACCGAAACAGCATTAG
M-caspase-9	F: CCCGTGGACATTGGTTCTGG R: GAGGAAGGGCAGAAGTTCACA
H-caspase-3	F: TGAGCCATGGTGAAGAAGGAATAA R: CCCGGGTAAGAATGTGCATAAAT
M-caspase-3	F: CAGCCAACCTCAGAGAGACATT R: TTTCAGTTCAACAGGCCCATTTG
H-SIRT2	F: TGCGGAACTTATTCTCCCAGA R: GGAGAGCGAAAGTCGGGGAT
M-SIRT2	F: GCAGAACATAGACACGCTGG R: CCCTGGGAGTTGCTTCTGAGA
H-β-actin	F: CTAGGCGGACTGTTACTGAGC R: ATGTTTGCTCCAACCAACTGC
M-β-actin	F: AAATCTGGCACCACACCTTCTAC R: CAGCCTGGATAGCAACGTACAT

.

4.17. Statistical Analysis

Statistical analyses were conducted using SPSS 22.0 and GraphPad Prism 8.0 software. Parametric tests were applied according to experimental design: one–way ANOVA for multiple group comparisons and Student’s t-test for pairwise comparisons. Post hoc analyses proceeded only when ANOVA yielded significant F-values (p < 0.05) with confirmed variance homogeneity. Results are presented as mean ± standard deviation (SD), with statistical significance defined as p < 0.05.