IDH2 lactylation promotes angiogenesis in murine diabetic myocardial infarction via blocking Cav1-eNOS interaction

Guangyao Zang; Siting Xu; Zhen Sun; Lili Zhang; Yongjiang Qian; Haipeng Yao; Lihua Li; Zhongqun Wang

doi:10.1038/s41467-025-67877-0

. 2025 Dec 19;17:1117. doi: 10.1038/s41467-025-67877-0

IDH2 lactylation promotes angiogenesis in murine diabetic myocardial infarction via blocking Cav1-eNOS interaction

Guangyao Zang ^1,², Siting Xu ^1,², Zhen Sun ^1,², Lili Zhang ^1,², Yongjiang Qian ^1,², Haipeng Yao ^1,², Lihua Li ³, Zhongqun Wang ^1,^2,^✉

PMCID: PMC12855196 PMID: 41419771

Abstract

Compensatory angiogenesis is critical for preserving left ventricular function after myocardial infarction; however, this process is severely impaired in diabetes, exacerbating adverse outcomes in diabetic myocardial infarction (DMI). This study employed liquid chromatography-tandem mass spectrometry to identify lactylated proteins in the infarct border zone of DMI male mouse hearts. Our findings revealed that IDH2 is lactylated at lysine 272, enhancing its binding to Cav1 while inhibiting the Cav1-eNOS interaction. This modification promotes eNOS activity and facilitates the proliferation, migration, and angiogenesis of cardiac microvascular endothelial cells under high glucose and hypoxic conditions. In endothelial cell-specific IDH2-K272R knock-in male mice, the loss of K272 lactylation impairs cardiac function and exacerbates pathological remodeling due to disrupted angiogenesis. Additionally, ACAT1 and HDAC1 act as lactyltransferase and delactylase, respectively, utilizing intracellular lactate transported via MCT1 as a substrate for IDH2 lactylation. Furthermore, pharmacologic enhancement of IDH2 lactylation, as demonstrated by empagliflozin mitigating post-DMI injury, supports its potential as a therapeutic target for DMI.

Subject terms: Post-translational modifications, Myocardial infarction

Angiogenesis following myocardial infarction is impaired in diabetes. Here the authors show that lactylation of IDH2 at lysine 272 promotes endothelial eNOS activation and angiogenesis, and enhancing this modification improved recovery in diabetic myocardial infarction.

Introduction

During myocardial infarction (MI), ischemic injury initiates a sterile inflammatory response, facilitating repair in the affected area and the formation of collagen-rich scar tissue to stabilize the structure of ischemic heart¹. Concurrently, ischemic injury triggers angiogenesis, enhancing angiogenesis repair within the damaged heart region². Angiogenesis extends from the infarct border zone towards the infarct core, augmenting blood supply to the affected region. Post-MI angiogenesis reduces infarct size, supports repair processes, and mitigates adverse myocardial remodeling and heart failure incidence³. Animal studies indicate that in MI mice, neo-capillary networks predominantly localize in the endocardium and epicardium of the infarct border zone. By the 7th day post-infarction, capillary proliferation subsides, reaching peak angiogenesis⁴. However, in diabetic microenvironments, heightened glucose metabolism inhibits post-myocardial ischemia angiogenesis, further impeding cardiac function recovery⁵. Additionally, diabetic neuropathy often leads to painless MI, delaying medical intervention⁶. Therefore, investigating angiogenic mechanisms post-diabetic myocardial infarction (DMI) holds significant clinical relevance for diagnosis and treatment strategies.

Under hypoxia and high glucose (HG) environment of DMI, increased activation of the glycolytic pathway leads to elevated lactate production. While lactate is a well-established metabolic substrate for oxidative metabolism in tissues such as the heart, its non-canonical roles in cell signaling remain incompletely characterized⁷. Recent studies emphasize its role as an epigenetic regulator by modifying specific lysine residues through protein lactylation, thereby regulating the functions of histone or non-histone proteins. It has been demonstrated that under diabetic conditions, lactate-mediated histone lactylation upregulates the expression of adiposity and obesity-related proteins, thereby enhancing endothelial cell proliferation, migration, and angiogenesis⁸., while hypoxia-induced lactylation of YY1-K183 in microglial cells enhances endothelial angiogenesis in proliferative retinopathy⁹ Histone lactylation activate repair genes shortly after MI, thereby promoting angiogenesis, and enhancing cardiac function¹⁰. Conversely, decreased myocardial α-MHC K1897 lactylation may disrupt myofilament interactions, potentially leading to heart failure¹¹. However, the role of protein lactylation in DMI is unclear.

Isocitrate dehydrogenase 2 (IDH2) is a form of IDH that catalyzes the oxidative decarboxylation of isocitrate to generate alpha-ketoglutarate. However, mutant forms of IDH have been observed to generate a metabolite called D-2-hydroxyglutarate, which induces global DNA hypermethylation and interferes with immunity, leading to stimulation of tumor growth¹². IDH2 is involved in regulating cellular activity by influencing mitochondrial function¹³. It has been demonstrated that IDH2 undergoes deacetylation by SIRT3 during cardiac hypertrophy preconditioning, contributing to cardioprotective effects¹⁴, while IDH2 desuccinylation provides cellular protection against oxidative stress and tumor suppression¹⁵. IDH2 has also been shown to improve the vascular function of endothelial progenitor cells in diabetic limb ischemia¹⁶. However, the role of IDH2 lactylation in post-DMI angiogenesis remains unknown.

In this study, we demonstrated that inhibition of IDH2 lactylation at lysine 272 (Lys272) exacerbates angiogenesis impairment and cardiac damage following DMI. Mechanistically, IDH2 lactylation enhances its binding to Caveolin-1 (Cav1) on cardiac microvascular endothelial cells (CMECs), thereby blocking Cav1 interaction with endothelial nitric oxide (NO) synthase (eNOS), preserving eNOS activity, and promoting angiogenesis. Moreover, we identified Acetyl-CoA acetyltransferase 1 (ACAT1) and Histone Deacetylase 1 (HDAC1) as lactyltransferase and delactylase involved in IDH2 lactylation, with intracellular lactate transported via Monocarboxylate transporter 1 (MCT1) playing a role in modulating IDH2 lactylation. Empagliflozin (EMPA) was identified to mitigate post-DMI injury by promoting IDH2 lactylation. In summary, our findings suggest that IDH2 lactylation represents a potential clinical target for DMI.

Results

Increased cardiac lactylation modification following DMI

Following sham surgery or MI induction in non-diabetic and diabetic mice, the impact of diabetes on mice after MI was investigated (Fig. 1a and Supplementary Fig. 1a, b). On the 7th day post-MI, cardiac tissues were procured from MI and DMI mice for 2, 3, 5-triphenyltetrazolium chloride (TTC) staining and unveiled a notably augmented infarct size in the hearts of DMI mice compared to MI (Supplementary Fig. 1c). Following MI, cardiac angiogenesis typically originates from sprouting of coronary artery endothelial cells¹⁷. To further explore the impact of diabetes on angiogenesis, we harvested aortas from mice with or without diabetes seven days after MI and implanted them into type I rat tail collagen for aortic ring assay to evaluate their endothelial cell sprouting capacity and found a reduced extent of endothelial cell sprouting in the DMI mice (Fig. 1b, c). In the end stages of MI, the formation of scars in the infarct zone reduced cardiac compliance, resulting in cardiac remodeling and fibrosis¹⁸. Masson’s trichrome staining results revealed a significant increase in collagen deposition in the hearts of the DMI mice compared to the MI mice thirty days post-MI (Fig. 1d). Furthermore, cardiac ultrasound was utilized to assess changes in left ventricular function in diabetes or MI-treated mice. The left ventricular ejection fraction (LVEF) and fractional shortening (FS) of the MI group decreased significantly compared to the Sham group, while the left ventricular end-diastolic diameter (LVIDd) and left ventricular end-systolic diameter (LVIDs) increased. Furthermore, the LVEF and FS of the DMI group were significantly lower than those of the MI group, while LVIDd and LVIDs were larger (Fig. 1e).

To investigate the overall status of cardiac lactylation modification following DMI, we utilized pan-lactylation antibody Pan Kla to assess lactylation levels in cardiac tissues. Western blot and immunofluorescence results of cardiac tissues revealed that compared to the Sham group, the MI group exhibited a significant increase in levels of protein lactylation in the infarct border zone tissue, while the levels of protein lactylation in the DMI group were significantly higher than those in the MI group (Fig. 1f, g). Taken together, these results indicate a significant increase in cardiac protein lactylation during diabetes-enhanced MI injury. Screening lactylated proteins as potential therapeutic targets for DMI warrants further investigation.

Proteomic profiling of lysine lactylation following diabetic myocardial infarction

To delve into the alterations of lactylation modification during the formation of DMI, on the 7th day after successfully establishing murine MI model with diabetes, we isolated infarct border zone tissues from Sham, diabetic Sham (Dsham), MI, and DMI mice. Equal amounts of protein were extracted, digested enzymatically, and subjected to enrichment of lactylated peptide segments, followed by mass spectrometry analysis for quantitative proteomic and lactylated proteomic investigations (Fig. 2a). Through mass spectrometric analysis, a total of 10,114 peptide segments were identified, among which 789 contained lactylation modifications. We identified 805 lactylation modification sites in 237 proteins, with 646 sites on 183 proteins having quantitative information (Supplementary Fig. 2a, b).

Fig. 2 — a Schematic representation of isolating myocardial infarction (MI) border zone tissue from each experimental group for lactylation proteomic analysis, created in BioRender. Zang, G. (2025) https://BioRender.com/95pgpau. b Venn diagrams illustrate the proteins and modification sites undergoing lactylation both in the MI/Sham groups, and in the DMI/MI groups. c Heatmap displays 27 lactylation modification sites in the DMI/MI comparison group, sorted by multiples of differential expressions. d Subcellular localization of lactylated proteins with differential modification between the MI group and DMI group. e GO functional enrichment analysis of differentially lactylated proteins in the DMI/MI comparison group. f KEGG pathway enrichment analysis of differentially lactylated proteins in the DMI/MI comparison group. g Schematic diagram depicting the lactylation modification status of tricarboxylic acid cycle (TCA) rate-limiting enzymes in DMI. Two-sided unpaired student’s t test was performed in (c). Fisher’s exact test was used in (e, f). Source data are provided as a Source data file.

We further delved into the process of protein lactylation modification during DMI while simultaneously exploring the impact of diabetes on MI. Analysis and selection of lactylation-modified proteins with differential expression between the MI and the Sham group, as well as between the DMI and the MI group, were conducted. To ascertain significant differences, only an average Kla enrichment or protein increase above 1.5 or below 0.67 was deemed a noteworthy alteration and subjected to further scrutiny. Following MI, there were significant alterations in lactylation expression in the infarct border zone tissue of MI mice compared to the Sham mice; 49 sites of lactylation were identified across 42 proteins. Specifically, the lactylation levels at 24 sites on 20 proteins were significantly elevated, whereas the lactylation levels at 25 sites on 22 proteins were markedly reduced. In investigating the impact of diabetes on MI-induced injury in mice, compared to the MI group, the DMI group exhibited lactylation at 90 sites of 62 proteins within the infarct border zone tissues. Specifically, the levels of protein lactylation at 23 sites of 21 proteins were significantly upregulated, while the extent of protein lactylation at 67 sites of 41 proteins were significantly downregulated (Supplementary Fig. 3a, b). Furthermore, the Sham and MI group shared five proteins displayed lactylation modifications at five sites, with a marked increase in lactylation levels observed within the MI group. Simultaneously, the DMI and MI group shared 27 lactylation sites across 17 proteins. Notably, two sites exhibited a significant increase in the DMI group, while lactylation levels decreased at 25 sites (Fig. 2b). Additionally, we utilized a heatmap to present the expression profile of differentially lactylated proteins among the groups, organizing them according to the fold change in expression. The substrate site showing the most pronounced lactylation alteration in DMI/MI comparison was IDH2-K272 (Fig. 2c and Supplementary Fig. 5c).

The subcellular localization and COG functional classification of differentially lactylated proteins were investigated. The results indicated that following MI, differentially lactylated proteins are predominantly localized in the mitochondria. In a diabetic environment, the enrichment of this lactylation modification in mitochondrial proteins significantly increases, with rates of 46% in the MI/Sham comparison and 57% in the DMI/MI comparison (Fig. 2d and Supplementary Fig. 4a). COG functional classification results demonstrated that the effects of MI and diabetes concentrate the function of differentially lactylated proteins in the heart on the regulation of energy production and conversion (Supplementary Fig. 4b, c). In the realm of biological processes illuminated by GO enrichment analysis results, the bubble chart of the MI/Sham group revealed that post-MI, differentially lactylated proteins primarily converge in the developmental processes of cardiovascular circulation and muscular organs. This signifies the potential involvement of lactylation in the cardiac repair post-MI (Supplementary Fig. 5a). Conversely, within the GO enrichment analysis of the DMI/MI comparison, it was evident that under the influence of diabetes, differentially lactylated protein modifications further engage in energy metabolism and immune regulation processes, suggesting a potential intervention of metabolism-associated protein lactylation in the cardiac repair post-MI within a diabetic milieu (Fig. 2e). Additionally, considering MI and its concomitant diabetes mellitus, we conducted KEGG pathway enrichment analysis of differentially lactylated proteins to delve into the signaling pathways potentially affecting cardiac function. The bubble plot results of the MI/Sham comparison indicated that post-MI, differentially lactylated proteins predominantly impact signaling pathways associated with dilated cardiomyopathy and hypertrophic cardiomyopathy, suggesting a focal influence of lactylation-modified proteins on adverse cardiac remodeling post MI (Supplementary Fig. 5b). Conversely, in the bubble plot of the DMI/MI comparison, it revealed that under the influence of diabetes mellitus, differentially lactylated proteins mainly affect the tricarboxylic acid (TCA) cycle process, hinting that the hyperglycemic environment induced by diabetes may influence post-MI cardiac function by modulating lactylation of key proteins in the TCA cycle (Fig. 2f). To delve deeper into the impact of lactylation on cardiac function in DMI mice, we conducted an analysis of the lactylation modification sites on key rate-limiting enzymes in the TCA cycle (Fig. 2g). The results indicated that under the diabetic MI conditions, nearly all rate-limiting enzymes in the TCA cycle underwent lactylation modification. Further cross-analysis with lactylation-modified proteins identified in the DMI/MI comparison revealed that IDH2 is the most significantly lactylated TCA-related protein in a diabetic MI environment.

Inhibition of IDH2 lactylation at Lys272 suppresses angiogenesis in high glucose and hypoxia environment

In the lactylation proteomics analysis, we observed a significant increase in levels of protein lactylation in the heart tissues of DMI mice during the early stage of infarct and identified the protein IDH2 as having significantly increased lactylation expression levels. To further validate the effects of IDH2 lactylation, primary CMECs were isolated from mice and subjected to HG and hypoxia treatments, divided into groups: normal glucose (NG), HG, normal glucose oxygen deprivation (NOD), and high glucose oxygen deprivation (HOD), to mimic the damage to CMECs during DMI. We found a significant increase in lactylation levels in HOD CMECs, and the lactylation levels gradually increased with the duration of hypoxia while the expression of IDH2 protein did not show significant changes (Fig. 3a–c). Furthermore, to validate the alterations in the lactylation levels of IDH2, we performed coimmunoprecipitation on heart tissues and CMECs. Our results indicated that the lactylation level of IDH2 was significantly elevated in the heart tissues of DMI mice and HOD CMECs (Fig. 3d, e). Additionally, the protein expression level of IDH2 did not exhibit significant changes under HG and hypoxic microenvironments, suggesting that IDH2 may modulate the function of CMECs in DMI primarily through alterations in its lactylation rather than changes in protein expression levels.

Fig. 3 — a Western blot analysis of global lactylation modification (Pan Kla) of cardiac microvascular endothelial cells (CMECs) after treatment with normal glucose (NG), high glucose (HG), normal glucose oxygen deprivation (NOD), and high glucose oxygen deprivation (HOD) (n = 6). b, c Western blot analysis of Pan Kla and IDH2 in CMECs with increasing hypoxia duration under high glucose conditions (n = 4). d Coimmunoprecipitation analysis of the lactylation level of IDH2 in the infarct border zone tissue of Sham, Dsham, MI, and DMI mice (immunoprecipitated by Pan Kla antibody). e Coimmunoprecipitation analysis of the lactylation levels of IDH2 in CMECs under NG, HG, NOD, and HOD conditions (immunoprecipitated by Pan Kla antibody). f Western blot analysis of the lactylation level of IDH2-K272 in the infarct border zone tissue of Sham, diabetic Sham (Dsham), myocardial infarction (MI), and diabetic myocardial infarction (DMI) mice (n = 6). g Western blot analysis of the lactylation level of IDH2-K272 in CMECs under NG, HG, NOD, and HOD conditions (n = 6). h Coimmunoprecipitation analysis of the lactylation status of IDH2 with lysine 272 mutation under NOD and HOD stimulation (immunoprecipitated by Pan Kla antibody). i Western blot analysis of the lactylation status of IDH2 with lysine 272 mutation under NOD and HOD stimulation (n = 6). j Proliferation capability of CMECs after transfected with WT or K272R plasmid under NOD and HOD stimulation was quantified by EdU assay (n = 6). Scale bar = 50 μm. k The angiogenesis capability of CMECs after transfected with WT or K272R plasmid under NOD and HOD stimulation was quantified by tube formation assay (n = 6). Scale bar = 100 μm. Data are presented as mean ± SEM. One-way ANOVA was used in (a, c, f, g, i–k). Source data are provided as a Source data file.

Through lactylation modification proteomic mass spectrometry analysis, we identified that the lactylation modification of IDH2 primarily occurs at its lysine 272 (K272) residue, which exhibits a high degree of conservation across multiple species, including Homo sapiens, Mus musculus, Rattus norvegicus, Oryctolagus cuniculus, Sus scrofa, and Bos taurus (Supplementary Fig. 6a, b). To validate this lactylation modification site, we generated a specific IDH2-K272 lactylation antibody, IDH2-K272la and observed a significant increase in the level of protein lactylation of IDH2-K272 in the infarct border zone tissue of DMI mice and in HOD CMECs (Fig. 3f, g). Moreover, by transfecting mutant plasmids targeting the K272 site of IDH2 (K272R) into CMECs, both the lactylation level of IDH2 and IDH2-K272 significantly decreased (Fig. 3h, i). while no detectable changes in acetylation or succinylation were observed (Supplementary Fig. 6c, d). Hyperosmotic stimulation also had no effect on IDH2 lactylation (Supplementary Fig. 6e). K272R mutation caused significantly reduced proliferation and migration (Fig. 3J and Supplementary Fig. 7a, b) of CMECs. Additionally, the tube formation ability of CMECs significantly decreased after transfection with the K272R (Fig. 3k). Collectively, these findings indicate that the Lys272 mutation of IDH2 impairs the angiogenesis of CMECs under HG and hypoxic microenvironments.

IDH2-K272 mutation in endothelial suppresses angiogenesis and cardiac function in DMI

We next sought to confirm the effect of IDH2-K272 lactylation in vivo by establishing murine model with endothelial cell-specific IDH2-K272R mutation (IDH2^EC-K272R; Fig. 4a and Supplementary Fig. 8a). Subsequently, we explored the influence of this mutation on outcome of post-DMI by inducing MI and DMI in both wild-type (IDH2^WT) and IDH2^EC-K272R mice (Fig. 4b). Initially, we examined the changes in IDH2 lactylation in the border zone tissue of MI and DMI mice in response to the endothelial cell-specific mutation at the IDH2-K272 site. In the border zone tissue of IDH2^EC-K272R MI and DMI mice, coimmunoprecipitation results demonstrated a significant decrease in level of IDH2 lactylation compared with the IDH2^WT mice, which was further confirmed by western blot analysis using IDH2-K272la antibody showing a marked reduction in lactylation levels (Fig. 4d, e). Immunofluorescence staining with IDH2-K272la revealed reduced expression of IDH2-K272la on endothelial in the hearts of IDH2^EC-K272R mice (Fig. 4f).

In the early phase of DMI, IDH2-K272R mutation in endothelial cells increased infarct area in IDH2^EC-K272R mice (Fig. 4c). To further validate the impact of the IDH2-K272R mutation on angiogenesis, the aortas of IDH2^WT mice and IDH2^EC-K272R mice from both MI and DMI groups were isolated for aortic ring assay and found a significant decrease in the sprouting ability of aortas from IDH2^EC-K272R mice (Fig. 4g). In the late stage of DMI, there was a significant increase in cardiac collagen deposition and myocardial fibrosis in DMI mice with the IDH2-K272R mutation. (Fig. 4h). Moreover, cardiac ultrasound results showed that the EF and FS were lower in IDH2^EC-K272R mice compared to IDH2^WT mice after DMI, while the LVIDd and LVIDs increased (Fig. 4i and Supplementary Fig. 8b). Taken together, these results suggest that the endothelial cell-specific IDH2-K272R mutation impairs cardiac function and aggravates adverse pathological remodeling from disruption of angiogenesis following DMI.

IDH2-K272 lactylation triggers eNOS activation by coupling with Cav1

eNOS activity is crucial for maintaining endothelial cell function in the post-ischemic revascularization process, and its phosphorylation is one of the main pathways regulating eNOS activity¹⁹. In the infarct border zone tissue of DMI mice and in CMECs subjected to HOD, compared to the MI and NOD groups, the expression levels of phosphorylated eNOS (p-eNOS) were significantly reduced, indicating that HG environments markedly inhibit eNOS activity under hypoxic conditions (Supplementary Fig. 9a, b). Moreover, in the heart tissue of IDH2^EC-K272R mice, the phosphorylation level of eNOS further decreased, leading to a reduction in eNOS activity (Fig. 5a). Transfection with K272R reduced lactylation of IDH2-K272 in CMECs treated with HOD, resulting in a significant decrease in the phosphorylation level of eNOS (Fig. 5b), subsequently leading to reduced eNOS activity and NO generation (Fig. 5c). To elucidate the mechanisms underlying the regulation of eNOS activity by IDH2 lactylation, we employed immunoprecipitation coupled with mass spectrometry (IP-MS) to detect intermediate interacting proteins involved in eNOS activity regulation after IDH2-K272 lactylation. CMECs were transfected with Flag-IDH2-WT and Flag-IDH2-K272R, followed by HOD treatment. Subsequently, immunoprecipitation using anti-Flag antibody was performed on lysates from treated CMECs, followed by mass spectrometry analysis (Fig. 5d). Through comprehensive analysis, 480 potential IDH2-binding proteins were identified, among which Cav1, a protein associated with eNOS activity negative regulation, was discovered (Supplementary Fig. 9c). The endogenous interaction between IDH2 and Cav1 was confirmed by coimmunoprecipitation in both cardiac tissues and CMECs (Supplementary Fig. 9d, e). To gain deeper insights into the spatial conformation of IDH2-Cav1 binding, we employed ZDOCK SERVER for molecular docking simulations (Fig. 5e). Previous research has indicated that the binding of Cav1 with eNOS can inhibit the activation of eNOS, thereby reducing the production of NO²⁰. This suggests that IDH2 lactylation might regulate eNOS activation mediated by Cav1 through affecting the interaction between IDH2 and Cav1. To further validate the effect of IDH2 binding to Cav1 on eNOS, coimmunoprecipitation was performed using Cav1 antibody. Our data demonstrated that IDH2-K272R mutation significant decreases the binding of Cav1 with IDH2, while increasing the binding of Cav1 with eNOS in HOD CMECs (Fig. 5f, g).

Fig. 5 — a, b Western blot analysis of the protein expression of eNOS and p-eNOS after the IDH2-K272R mutation both in vivo and in vitro experiments (n = 6). c NO content of cardiac microvascular endothelial cells (CMECs) after transfected with WT or K272R plasmid under normal glucose oxygen deprivation (NOD) and high glucose oxygen deprivation (HOD) stimulation was quantified by NO fluorescent probe (n = 6). Scale bar = 100 μm. d IP-MS analysis of CMECs treated with Flag-IDH2-WT and Flag-IDH2-K272R to screen for proteins interacting with IDH2, created in BioRender. Zang, G. (2025) https://BioRender.com/th0j3wz. e ZDOCK SERVER simulated the protein binding spatial conformation of IDH2 and Cav1, where yellow represents IDH2 and blue represents Cav1. f Coimmunoprecipitation experiment assessing the binding capacity of Cav1 with IDH2 and eNOS in CMECs after IDH2-K272R mutation under NOD and HOD stimulation (immunoprecipitated by Cav1 antibody, n = 3). g Schematic representation illustrating the effect of IDH2-Cav1 binding on the interaction between Cav1 and eNOS, created in BioRender. Zang, G. (2025) https://BioRender.com/1kizpt1. h Simulation using ZDOCK SERVER and PDBePISA to predict the spatial conformation of IDH2 binding to Cav1. Yellow represents IDH2, blue represents Cav1, and red indicates the peptide sequence of Cav1 binding to IDH2. i Schematic representation of truncated forms of Cav1 constructed based on its structural domains. Structures of Cav1 full-length (FL) and truncated plasmids (∆1–81, ∆82–101, ∆102–178). j CMECs were transfected with Flag-tagged IDH2 (Flag-IDH2) and truncated His-tagged Cav1 (His-Cav1) plasmids in i. Coimmunoprecipitation analysis of Flag-IDH2 and His-Cav1 interaction (immunoprecipitated by His antibody, n = 3). Data are presented as mean ± SEM. One-way ANOVA was used in (a–c). Source data are provided as a Source data file.

Subsequently, we endeavored to elucidate the mechanism by which the interaction between IDH2 and Cav1 influences the interaction between Cav1 and eNOS. We hypothesized that IDH2 competes with eNOS for binding to Cav1. To delineate the binding domain on Cav1 that IDH2 competes with eNOS for binding to, we employed the ZDOCK SERVER and PDBePISA for computational simulation, thereby predicting the spatial configuration of the interaction between IDH2 and Cav1. It was discerned that the binding peptide sequence of Cav1, 83-101 amino acids “GIWKASFTTFVTKYWFYR,” predominantly resides within the CSD domain of Cav1 (Fig. 5h). Reports indicate that Cav1 can inhibit eNOS activity, thereby suppressing the release of NO, through protein-protein binding in the form of eNOS interaction via amino acid residues within its CSD domain²¹. This suggests that IDH2 might competitively bind to the CSD domain of Cav1, influencing the interaction between Cav1 and eNOS. To validate this hypothesis, truncated plasmids of Cav1 protein bearing His-tags were constructed based on different structural domains of Cav1, following co-transfection of these truncated plasmids with Flag-IDH2-WT, coimmunoprecipitation was conducted using anti-His antibody. The results revealed that only the CSD domain truncated plasmid Cav1 ∆82-101 failed to bind with IDH2, whereas full-length Cav1 and two other truncated Cav1 could bind with IDH2, indicating an interaction between IDH2 and the CSD domain of Cav1 (Fig. 5i, j). Taken together, these data suggest that IDH2-K272 lactylation blocks the interaction between Cav1 and eNOS by increasing the binding of IDH2 with Cav1, ultimately triggering the activation of eNOS in CMECs.

ACAT1 and HDAC1 are responsible for lactylation and delactylation, respectively, at K272 of IDH2

In further exploring the potential mechanisms underlying the increased expression of IDH2 lactylation in DMI with the microenvironment of HG and hypoxia, we identified lactylation enzymes and delactylation enzymes capable of interacting with IDH2. Screening was conducted via IP-MS to identify proteins capable of interacting with IDH2 under HOD conditions. Initially, CMECs were subjected to NOD or HOD treatments, followed by immunoprecipitation using an IDH2-K272la antibody on cell lysates of CMECs, and subsequent mass spectrometry analysis (Supplementary Fig. 10a). A total of 897 proteins capable of binding to IDH2 were identified, and intersectional screening was performed with acyltransferases and deacylases from the Uniprot database, ultimately identifying two potential acyltransferases, N-acetyltransferase 10 (Nat10) and ACAT1, and one deacylase, HDAC1 (Supplementary Fig. 10b). We further examined the expression of Nat10, ACAT1, and HDAC1 under HOD conditions and found a significant increase in the expression of ACAT1, while the expression of Nat10 and HDAC1 decreased significantly (Supplementary Fig. 10c). The expression changes of the acyltransferase ACAT1 were consistent with the trend of level of IDH2 lactylation, while the expression changes of the acyltransferase Nat10 and the deacylase HDAC1 were opposite to the trend of IDH2 lactylation. These results suggest that ACAT1 and HDAC1 might be lactyltransferase and delactylase regulating IDH2 lactylation.

After filtering out potential acyltransferase ACAT1 for IDH2-K272 lactylation (Fig. 6a), ZDOCK SERVER was employed to simulate the spatial conformation of IDH2 bound to ACAT1 (Fig. 6b). We further validated its endogenous interaction with IDH2 in cardiac tissue and CMECs through coimmunoprecipitation (Supplementary Fig. 10d, e). Furthermore, under hypoxic conditions, HG stimulation promoted the interaction between ACAT1 and IDH2 or IDH2-K272la, consistent with the trend of IDH2 lactylation under HOD (Fig. 6c). These findings further support the regulatory role of ACAT1 in IDH2 lactylation. Therefore, we employed the ACAT1 inhibitor Avasimibe to investigate the influence of ACAT1 on IDH2 lactylation. The results revealed that treatment with Avasimibe in CMECs significantly reduces the expression of ACAT1 in both NOD and HOD conditions, accompanied by a decrease in lactylation level of IDH2 at the K272 site (Fig. 6d). Under the inhibition of ACAT1, the interaction between IDH2 and Cav1 decreases, allowing Cav1 to associate more with eNOS (Fig. 6e, f), consequently leading to reduced cell proliferation, NO generation, and tube formation of CMECs (Fig. 6g).

Meanwhile, after screening for potential delactylase HDAC1(Supplementary Fig. 11a), spatial conformation simulations of IDH2 binding with HDAC1 using ZDOCK SERVER was performed (Supplementary Fig. 11b). We further conducted coimmunoprecipitation to validate the endogenous interaction between HDAC1 and IDH2 in cardiac tissue and CMECs (Supplementary Fig. 11c, d). Under hypoxic conditions, HG stimulation inhibited the binding of HDAC1 with IDH2 and IDH2-K272la, consistent with the trend of IDH2 lactylation (Supplementary Fig. 11e). These experimental results further support the negative regulatory role of HDAC1 in IDH2 lactylation. Therefore, we employed the HDAC1 inhibitor Entinostat to investigate the effect of HDAC1 on IDH2 lactylation. Experimental results showed that treatment with Entinostat in CMECs significantly reduces the expression level of HDAC1, thereby increasing the lactylation level of IDH2-K272 (Supplementary Fig. 11f). Furthermore, it was found that under the inhibition of HDAC1, the binding between IDH2 and Cav1 increases through coimmunoprecipitation, leading to a decrease in the association between Cav1 and eNOS (Supplementary Fig. 11g, h), thereby enhancing the proliferation, NO generation and tube formation of CMECs (Supplementary Fig. 11i). Altogether, these results suggest that ACAT1 and HDAC1 function as the lactyltransferase and delactylase, respectively, for IDH2-K272 lactylation.

Lactate is transported intracellularly via MCT1 to facilitate lactylation of IDH2

Lactylation modification is a post-translational modification process using the metabolite lactate as a substrate²², and whether the lactate concentration can directly influence the generation of lactylation modification in CMECs under HOD conditions requires further investigation. Therefore, in HOD CMECs, exogenous sodium lactate (Nala) was added to explore the regulatory role of lactate on the lactylation of IDH2. The results demonstrated that the addition of exogenous Nala significantly increased the intracellular lactate concentration in CMECs (Supplementary Fig. 12a), further promoting the lactylation of IDH2-K272 (Supplementary Fig. 12b). Additionally, the upregulation of IDH2 lactylation level induced by exogenous Nala led to an increase in the binding between IDH2 and Cav1, while decreasing the binding between Cav1 and eNOS (Supplementary Fig. 12c). MCT1 is a crucial lactate transporter involved in lactate shuttling processes that transport lactate into cells²³. To further investigate the mechanism by which lactate regulates IDH2 lactylation, the MCT1 inhibitor AZD3965 was used, which was observed to reduce the intracellular lactate concentration increased by exogenous lactate in HOD, indicating that the MCT1 inhibitor reduced the entry of lactate into cells (Supplementary Fig. 12d). Additionally, AZD3965 decreased the lactylation of IDH2 (Supplementary Fig. 12e) and the binding between IDH2 and Cav1 promoted by Nala, allowing for an increased binding between Cav1 and eNOS (Supplementary Fig. 12f), thus reducing the proliferative capacity, NO production, and tube formation ability of CMECs facilitated by Nala (Supplementary Fig. 12g). Collectively, these findings suggest that lactate enters cells through the transport activity of MCT1, promoting the lactylation of the mitochondrial protein IDH2 and enhancing the binding between IDH2 and Cav1, while reducing the interaction between Cav1 and eNOS, thereby enhancing the activity of eNOS in CMECs.

Empagliflozin enhances angiogenesis and cardiac function by augmenting IDH2-K272 lactylation following DMI

To investigate whether existing antidiabetic medications promote IDH2-K272 lactylation, we tested several drugs and found that SGLT2 inhibitors, especially EMPA, showed the most significant enhancement in IDH2 lactylation. (Supplementary Fig. 13a). EMPA significantly enhanced the degree of IDH2-K272 lactylation in the HOD CMECs, and partially restore the impairment of eNOS activity induced by hypoxia and HG (Supplementary Fig. 13b). EMPA promoted endothelial cell proliferation, tube formation, and NO generation, suggesting that EMPA exerts a protective effect on HOD CMECs (Supplementary Fig. 13c). Additionally, the addition of EMPA to the WT group further promotes the lactylation of IDH2-K272 in HOD, but the addition of EMPA to the K272R group fails to restore the decreased level of IDH2-K272 lactylation (Supplementary Fig. 14a). EMPA further enhanced the binding between IDH2 and Cav1, leading to a reduction in the interaction between Cav1 and eNOS in HOD CMECs. However, the effect of EMPA disappears in the K272R group (Supplementary Fig. 14b).

To further investigate the impact of EMPA on IDH2 lactylation following DMI, administration of EMPA (30 mg/kg/day) to DMI mice via gavage 7 days before MI surgery revealed that EMPA could promote IDH2-K272 lactylation in the infarct border zone tissue of IDH2^WT DMI mice, but it could not restore the decreased IDH2-K272 lactylation in the infarct border zone tissue of IDH2^EC-K272R mice post-DMI (Fig. 7a, b and Supplementary Fig. 15a). Coimmunoprecipitation from cardiac tissue showed that administration of EMPA could enhance the binding between IDH2 and Cav1 in the infarct border zone tissue, leading to a reduction in the interaction between Cav1 and eNOS. However, this effect of EMPA was lost in IDH2^EC-K272R mice (Fig. 7c). Additionally, EMPA administration significantly reduces blood glucose levels in diabetic mice and decreases the myocardial infarct area in diabetic mice with MI, albeit to a lesser extent in IDH2^EC-K272R mice (Supplementary Fig.15b, c). We next explored the physiological effects of EMPA in IDH2^WT and IDH2^EC-K272R DMI mice. EMPA promoted angiogenesis in diabetic mice with MI, but this effect is attenuated in IDH2^EC-K272R mice (Fig. 7d). EMPA treatment also provided protection against aggravation of myocardial fibrosis in the IDH2^WT group. Nonetheless, the myocardial fibrosis alleviation caused by EMPA administration was partially offset by the IDH2-K272R mutation (Fig. 7e). Moreover, EMPA treatment ameliorated the decline in EF and FS and attenuated the increase in LVIDd and LVIDs in IDH2^WT DMI mice. However, the therapeutic effect of EMPA was partially attenuated in the IDH2^EC-K272R group compared to the IDH2^WT group (Fig. 7f, Supplementary Fig. 15d).

Fig. 7 — a Western blot analysis of IDH2-K272 lactylation levels in infarct border zone tissues of IDH2^WT and IDH2^EC-K272R mice following induction of myocardial infarction (MI) or DMI with gavage administration of EMPA (i.g. 30 mg/kg/day, 7 days; n = 6). b Representative Immunofluorescence staining of IDH2-K272la in heart from IDH2^WT and IDH2^EC-K272R mice following induction of MI or DMI with gavage administration of empagliflozin. CD31 is an endothelial cell marker (n = 6). Scale bar = 20 μm. c Coimmunoprecipitation experiment assessing the binding capacity of Cav1 with IDH2 and eNOS in infarct border zone tissues of IDH2^WT and IDH2^EC-K272R mice following induction of MI or DMI with gavage administration of empagliflozin (immunoprecipitated by Cav1 antibody). d Arterial ring experiments were conducted in IDH2^WT and IDH2^EC-K272R mice following induction of MI or DMI with gavage administration of empagliflozin (n = 6). Scale bar = 100 μm. e Representative images and quantification of Masson’s trichrome staining for cardiac fibrosis in the infarct border zone from IDH2^WT and IDH2^EC-K272R mice following induction of MI or DMI with gavage administration of empagliflozin (n = 6). Scale bar = 20 μm. f Representative images and ejection fraction (EF) of cardiac function evaluated by echocardiography in IDH2^WT and IDH2^EC-K272R mice following induction of MI or DMI with gavage administration of empagliflozin (n = 6). Data are presented as mean ± SEM. One-way ANOVA was used in (a, b, d, e, f). Source data are provided as a Source data file.

In summary, EMPA exhibited therapeutic effects on angiogenesis and cardiac function following DMI. However, endothelial IDH2-K272R mutation partially attenuated the protective effects of EMPA against DMI. Collectively, our findings underscore the significant role of IDH2-K272 lactylation after DMI.

Discussion

Endothelial cells within the myocardial microvasculature play a vital role in the early stages of myocardial repair post-MI, facilitating angiogenesis and the formation of capillary networks^3,17. The efficacy of these processes, however, is compromised in hyperglycemic conditions⁵. Our findings reveal that inhibition of IDH2 lactylation in endothelial cells exacerbates angiogenesis impairment in DMI, underscoring its indispensable role as a compensatory mechanism to mitigate vascular injury. Specifically, we pinpoint Lys272 on IDH2 as the pivotal lactylation site dictating the impact of the diabetic microenvironment on MI. Mechanistically, heightened lactylation on IDH2-K272 under HOD enhances the interaction between IDH2 and Cav1 protein, consequently blocking the binding between Cav1 and eNOS. This cascade diminishes the impairment to eNOS function and fosters angiogenesis. These findings suggest that pharmacologic amplification of IDH2 lactylation, such as through SGLT2 inhibitors, may serve as a potential therapeutic strategy to enhance vascular repair and mitigate adverse outcomes in post-DMI management.

eNOS is a membrane protein mainly located on plasma vesicles or the Golgi apparatus. In the cardiovascular system, eNOS is primarily expressed on vascular endothelial cells²⁴. The activity of eNOS and the bioavailability of NO are crucial for maintaining endothelial cell function. The biological effects of eNOS are influenced by its coupling state. Plasma membrane caveolar protein Cav1 interacts with eNOS located on plasma vesicles through its CSD domain, inhibiting eNOS activation and reducing NO synthesis^20,21, indicating Cav1 is a key negative regulator of eNOS activation. Recent research indicates that the binding of Cav1 to eNOS regulates eNOS activity and NO synthesis, playing a significant role in cardiovascular and cerebrovascular diseases. Excessive shear stress on BACE1 lead to Cav1 accumulation on the membrane, mediating eNOS endocytosis and increasing Cav1 binding to eNOS, thereby deactivating eNOS and subsequently affecting cerebrovascular endothelial cell function²⁵. Additionally, deletion of Sp1/Sp3 increases Cav1 binding to eNOS, decreases eNOS activity, and leads to endothelial dysfunction and hypertension²⁶. In our study, we identified IDH2 as a Cav1-binding protein. IDH2-K272 lactylation promotes the binding of IDH2 to the CSD domain of Cav1, reducing the interaction between Cav1 and eNOS, ultimately mitigating the damage of eNOS activity, and promoting angiogenesis in a HG hypoxic microenvironment. Thus, it is evident that IDH2 lactylation plays a crucial role in maintaining endothelial cell angiogenesis function during DMI.

To elucidate the impact of IDH2 lactylation on DMI, we employed diverse methodologies to manipulate the extent of IDH2 lactylation. Apart from modulating the lysine residue implicated in IDH2 lactylation, we delved into the regulation of enzymes participating in IDH2 lactylation. Protein acylation modification constitutes a reversible and dynamic covalent modification process chiefly governed by enzymes²⁷. In the realm of protein acylation modification, virtually all identified short-chain acylation alterations are recognized to be regulated by acyltransferases and deacylases alike²⁸. Through IP-MS analysis, we identified acyltransferase and deacylase, ACAT1 and HDAC1, which interact with IDH2. The ACAT1 is a key regulatory enzyme that controls fatty acid oxidation and ketogenesis and promotes cholesterol esterification²⁹. Additionally, as an acetyltransferase, ACAT1 also facilitates acetylation modifications of multiple proteins. Studies have shown that acetylation modification of ACAT1 enhances the activity of ME1 and participates in the development of colorectal tumors³⁰. Moreover, ACAT1 acetylates mutated IDH2, thereby regulating its activity to control the production of 2-HG, ultimately impacting the development of AML³¹. This suggests that ACAT1 has the potential to lactylate IDH2. On the other hand, the histone deacetylase family plays a significant role in regulating protein acetylation modification. Research has revealed that HDAC1 not only deacetylates histones but also deacetylates non-histone proteins such as PTEN, inhibiting its activity and regulating the PI3K/Akt signaling pathway³². In cardiovascular diseases, HDAC1 located in cardiomyocyte mitochondria promotes myocardial ischemia-reperfusion injury³³. Moreover, Class I HDACs (HDAC 1-3) are considered among the most effective lysine lactylation erasers³⁴. Our study confirms that ACAT1 and HDAC1 serve as modifier enzymes for IDH2 lactylation. Intervening with ACAT1 and HDAC1 in HOD microenvironment significantly regulates IDH2-K272 lactylation, affecting the binding of IDH2 to Cav1, regulating the interaction between Cav1 and eNOS, thereby influencing NO generation, as well as the proliferation and angiogenesis capacity of CMECs.

Beyond its classical role as an energy metabolite and gluconeogenic precursor, lactate is now recognized as a pleiotropic signaling molecule. Its differential produced in various diseases significantly regulates disease functions. Recent studies have shown that through the MCT family and the receptor GPR81, lactate activates cellular energy metabolism, neuroprotection, angiogenesis, and inflammation modulation pathways, thereby regulating the physiological and pathological functions of the body^35–37. Furthermore, similar to other fatty acids, lactate also forms acyl-CoA, lactyl-CoA, providing lactyl groups as donors for lactylation modifications²². This confirms the potential of lactate to exert biological activity through protein lactylation and regulate disease progression. Previous studies have reported that the addition of exogenous lactate significantly increases intracellular lactate levels and histone H3K18 lactylation levels in bone marrow-derived macrophages, enhancing endothelial cell angiogenic capacity. Intraperitoneal injection of sodium lactate also promotes angiogenesis after MI in mice, improving cardiac dysfunction¹⁰. However, the role of lactylation in DMI is still unclear. Our study found that the addition of exogenous sodium lactate significantly increases lactate levels in CMECs under HOD and promotes lactylation levels of IDH2-K272, further impacting the binding of IDH2 to Cav1. This regulates the interaction between Cav1 and eNOS, thereby protecting endothelial function under HG and hypoxic injury. MCT1 plays a crucial role in cellular lactate uptake and induction of the healing processes, with its expression observed in endothelial cells, cardiomyocytes, and fibroblasts according to the Human Protein Atlas³⁸. Although cardiomyocytes exhibit the highest levels of MCT1, endothelial cells have double the expression levels compared to fibroblasts. Cells with high MCT1 expression may be the first to uptake lactate from the circulation. Based on the abundance of MCT1 expression, the heart may prioritize self-repair (affecting cardiomyocytes) and promote angiogenesis (affecting endothelial cells). Our study also revealed that the use of the MCT1 inhibitor AZD3965 significantly reduces the increase in lactate levels in endothelial cells induced by exogenous sodium lactate. It inhibits the lactylation of IDH2-K272 promoted by sodium lactate under HOD and further suppresses eNOS activity and endothelial cell function. This confirms that lactate enters cells through the transport protein MCT1, promotes the intracellular lactylation of proteins, and consequently protects the function of microvascular endothelial cells following DMI.

Diabetes represents the most prevalent and consequential comorbidity in cardiovascular disease, with cardiovascular complications standing as a primary cause of mortality and morbidity among diabetic patients. Notably, recent studies indicate that certain antidiabetic medications may confer cardiovascular benefits for both diabetic and non-diabetic individuals. SGLT2 is a sodium-glucose cotransporter located in the proximal tubule of the kidney, responsible for reabsorbing approximately 90% of glucose from urine. Inhibiting SGLT2 promotes urinary glucose excretion, thereby reducing blood glucose levels³⁹. Clinical studies including EMPA-REG OUTCOME, CANVAS, and DECLARE-TIMI 58 have demonstrated that SGLT2 inhibitors such as EMPA, canagliflozin (CANA), and dapagliflozin (DAPA) significantly reduce the relative risk of major adverse cardiovascular events^40–42. Clinical research indicates that treatment with SGLT-2 inhibitors has beneficial clinical outcomes for diabetic patients hospitalized for acute MI and reduces the prognostic gap between diabetic and non-diabetic patients⁴³. This suggests that anti-diabetic drugs may have a reparative effect on MI damage in diabetic patients, but the specific mechanisms are not fully understood. In this study, using three classic SGLT2 inhibitors, canagliflozin, EMPA, and dapagliflozin, it was found that EMPA significantly promoted the expression of IDH2-K272 lactylation in the infarct border zone of DMI mice and in HOD CMECs. By regulating IDH2-K272 lactylation, EMPA reduced the damage from DMI and promoted the repair of endothelial cell function. In reports of EMPA treatment for heart failure, studies have indicated that EMPA activates eNOS activity, promotes NO generation, and improves diastolic function of the left ventricle in a non-diabetic heart failure pig model⁴⁴. However, the specific mechanism by which EMPA regulates eNOS activity in endothelial cells is not yet clear. Our study found that EMPA promotes the binding of IDH2 to Cav1 through lactylation of IDH2, thereby reducing the interaction between Cav1 and eNOS, mitigating the damage of eNOS activity in the DMI microenvironment, which has not been reported in previous studies. However, less discussion has been undertaken in our research regarding the impact of IDH2 lactylation on IDH2 enzyme activity. Therefore, the effect of IDH2 lactylation on IDH2 enzyme activity and its metabolite generation will be verified through further in vivo and in vitro experiments. This will help explore the relationship between protein lactylation and metabolic reprogramming in DMI. Moreover, whether IDH2 lactylation is protective without sex bias remains to be defined by more studies in females.

In conclusion, our study found that during the repair process following DMI injury, lactate produced in a HG hypoxic environment was transported into cells via MCT1 as a substrate for lactylation modification. Additionally, the diabetic HG microenvironment upregulates the expression of the lactyltransferase ACAT1 and downregulates the expression of the delactylase HDAC1 under hypoxic conditions, thereby promoting IDH2-K272 lactylation. Following IDH2-K272 lactylation, it facilitates the binding of IDH2 to the Cav1 protein, reducing the interaction between Cav1 and eNOS, thereby mitigating the impairment of endothelial cell eNOS activity and promotes angiogenesis in a HG and hypoxia environment. Furthermore, our findings indicate that the diabetes treatment drug EMPA promotes angiogenesis after DMI by regulating IDH2-K272 lactylation. This finding helps to elucidate the mechanism behind the cardiovascular protective effects of EMPA and suggests that targeting protein lactylation may be a therapeutic strategy for DMI.

Methods

Animal study

C57BL/6J male mice and Idh2^K272R-flox knock-in mice were obtained from GemPharmatech (Nanjing, China). Idh2^K272R-flox knock-in mice, in which Lysine 272 of IDH2 was mutated to arginine (K272R) using CRISPR/Cas9 technology. Idh2^K272R-flox knock-in mice were crossbred with Tek-Cre mice to obtain Idh2^EC-K272R (IDH2^EC-K272R) mice. Only male mice were used to reduce potential biological variability. All mice were housed in a SPF environment with a 12:12 h light/dark cycle. All animal studies were conducted with the approval of the Animal Research Committee of Jiangsu University (UJS-IACUC-2023101002).

Diabetes model

After acclimation, streptozotocin (STZ) was injected into the peritoneal cavity of the male mice at a dose of 40 mg/kg/day for 5 consecutive days to establish the diabetes model. Blood glucose levels of the mice were measured 2 weeks later, and mice with random blood glucose levels above 16.7 mmol/L in three consecutive measurements were defined as successfully constructed diabetes models.

Myocardial infarction model

Male mice were anesthetized with 2% isoflurane and positioned supine. The surgeon was blinded to the genotype and treatment group of the mice during all surgical procedures. After confirming anesthesia with a stable II lead electrocardiogram, the chest hair was disinfected with 75% ethanol, and a 1 cm incision was made along the left edge of the sternum. The subcutaneous tissue and muscles were bluntly separated. A curved vascular clamp was used to gently extrude the heart through an incision between the 3rd and 4th ribs, exposing the chest cavity. The left anterior descending coronary artery was ligated at a standardized anatomical position (approximately 2–3 mm from its origin) with a 7-0 silk suture, ensuring appropriate depth and width without penetrating the heart. After ligation, the heart was gently returned to the chest cavity, and the chest incision was closed with care. Post-surgery, mouse electrocardiograms were monitored closely for elevated T waves, ST segment elevation, and widened QRS waves indicating successful MI induction^45,46. Mice typically regained consciousness within 2 min after anesthesia cessation. Sham surgery involved similar procedures, except the coronary artery was pierced without ligation. Mice were classified as follows: Sham; diabetic sham (Dsham); MI; DMI.

TTC staining

Hearts were washed in PBS to remove blood, then briefly frozen at −80 °C for 10 min and sliced perpendicular to the long axis in 1 mm intervals from apex to base. Slices were thawed and stained with 2% 2, 3, 5-TTC solution (G3005, Solarbio, China) at 37 °C in darkness for 15 min, followed by fixation in 4% paraformaldehyde for 30 min. Images were captured using a dissecting microscope; ischemic areas appeared white and non-ischemic areas red. Infarct size was quantified using ImageJ software. For each heart, the infarcted (white) area and left ventricular (LV) area were measured across all slices, and the final infarct size was expressed as the percentage of the infarct area relative to the total LV area^46,47.

Microfil cardiac vascular imaging

1000 units of heparin were injected into the abdominal cavity of mice at 5-min intervals. Mice were anesthetized, positioned supine, and disinfected with 75% alcohol. The chest cavity was exposed, and a catheter needle was inserted into the left ventricle (3–4 mm) while cutting the inferior vena cava. A PBS solution with 0.1 mM sodium nitroprusside perfused the heart to dilate blood vessels. Blood color changes from red to transparent indicated circulation drainage (5–8 min). Subsequently, 1% paraformaldehyde stiffened vessel walls. A yellow dye contrast agent (diluent: curing agent, 5:2:3; Flow Tech, USA) was perfused within 10 min, followed by PBS rinsing. Hearts were stored overnight at 4 °C, then fixed in 4% paraformaldehyde. Dehydration in alcohol gradients (25–100%) and immersion in methyl salicylate preceded heart vessel photography using a dissecting microscope.

Aortic ring assay

The mice were euthanized using excess carbon dioxide and immersed in 75% alcohol for 10 min. Using sterile ophthalmic surgical instruments, the aortic sheath was dissected under a dissecting microscope to isolate blood vessels, which were subsequently washed in sterile PBS. The vessels were then transferred to culture medium and carefully peeled to remove the outer layer. After rinsing in PBS, the vessels were sectioned into 1 mm rings and starved overnight in serum-free Opti-MEM medium. The following day, plates were prepared using type I mouse tail collagen gel (354236, Corning, USA) by mixing with NaOH and PBS to achieve a concentration of 2 mg/ml. The arterial rings were embedded in this collagen gel in a 96-well plate. After solidification for 1 h at 37 °C, medium containing 2.5% fetal bovine serum DMEM was added and changed every other day. On the 7th day, blood vessels sprouting from the aortic rings were observed under a microscope.

Cardiac ultrasound

Cardiac function was evaluated using cardiac ultrasonography with a Vevo 2100 High-Resolution Micro-Ultrasound System (FUJIFILM Visual Sonics Inc., Japan). Mice were anesthetized with 2% isoflurane gas and positioned on the examination table to maintain their heart rate within a specific range (400–550 beats per minute). Cardiac ultrasonography included M-mode and B-mode analysis to measure parameters such as LVEF, FS, LVIDd, and LVIDs, which are widely applied methods for assessing cardiac function^48,49. Measurements were averaged over three cardiac cycles to assess the mice’s cardiac function.

Masson’s trichrome staining

Masson’s trichrome staining was conducted according to the manufacturer’s protocol (G1340; Solarbio, China). Paraffin sections were deparaffinized twice in xylene for 5–7 min each and then hydrated in a graded series of ethanol solutions (100%, 100%, 95%, 85%, 80%, 75%) for 5 min each. Sections were rinsed twice in distilled water to prevent dehydration. Staining began with Weigert’s iron hematoxylin solution for 10 min, followed by differentiation in acidic ethanol solution for 10 s, rinsing in distilled water for 1 min, counterstaining with Biebrich scarlet for 5 min, and final rinsing in running water and distilled water. Subsequently, sections were stained with Light Green Scarlet solution for 12 min, followed by rinsing in 0.1% acetic acid solution for 1 min. Further steps included washing in phosphomolybdic acid solution for 2 min, direct staining in Aniline Blue solution for 1 min and 30 s, cleaning with 0.1% acetic acid solution, and final observation under a microscope.

Immunofluorescence staining

The heart tissue paraffin sections underwent initial deparaffinization and hydration, followed by antigen retrieval using boiling EDTA solution in a pressure cooker. After cooling and PBS washing, the sections were circled with an immunohistochemistry pen. Subsequently, 0.3% Triton X-100 was applied and incubated, followed by blocking with 5% goat serum. The primary antibodies used were anti-CD31 (ab7388, Abcam, USA), Anti-L-Lactyl Lysine Rabbit mAb (anti-Pan Kla, PTM-1401RM, PTMBIO, China) and IDH2-K272la (PTMBIO, China), applied overnight at 4 °C. After washing, sections were incubated with Alexa Fluor® 488 or 594 conjugated secondary antibodies (Abcam, USA) at 37 °C for 45 min. DAPI staining of cell nuclei followed, and after washing, sections were observed using a fluorescence microscope. Quantification of fluorescence intensity was performed using Image J software.

Western blot and immunoprecipitation

Infarct border zone tissues from hearts and CMECs lysates were lysed using a lysis buffer. Protein samples were quantified and adjusted based on target protein expression for Western blot (WB) analysis. Protein A/G magnetic beads (22202, BEAVERBIO, China) were utilized for coimmunoprecipitation and lactylation immunoprecipitation, incubating them with primary antibodies (anti-Pan Kla, PTM-1401RM, PTMBIO; anti-pan-Acetyllysine, PTM-101, PTMBIO; anti-pan-Succinyllysine, A22228, Abclonal; anti-Cav1, 16447-1-AP, Proteintech; anti-His, 66005-1-Ig, Proteintech; anti-ACAT1, 16215-1-AP, Proteintech; anti-HDAC1, 10197-1-AP, Proteintech) for 30 min at room temperature. Subsequently, the complexes underwent overnight incubation with the protein lysate at 4 °C. After washing with lysis buffer, complexes were eluted using sodium dodecyl sulfate (SDS) loading buffer. Following electrophoresis on acrylamide gels and transfer onto 0.22 µm PVDF membranes, the membranes were incubated with BSA, primary antibodies (overnight at 4 °C), and secondary antibodies (1 hour at room temperature). Primary antibodies against the following target proteins were used: Pan Kla (PTM-1401RM, PTMBIO, China); IDH2 (A7190, ABclonal, China); IDH2-K272la (PTMBIO, China); eNOS (A1548, ABclonal, China); p-eNOS (AP0421, ABclonal, China); Cav1 (16447-1-AP, Proteintech, China); ACAT1(16215-1-AP, Proteintech, China); HDAC1(10197-1-AP, Proteintech, China); Tubulin (AC007, ABclonal, China). Signal intensities were analyzed using ImageJ software, with protein expression normalized against Tubulin.

IDH2 in vitro binding assay

Immunoprecipitation coupled with mass spectrometry (IP-MS) facilitates the identification of proteins interacting with IDH2. Initially, proteins interacting with IDH2 under various stimuli are enriched through immunoprecipitation experiments. Next, proteins influenced by IDH2-K272 lactylation are screened by transfecting CMECs with Flag-IDH2-WT and Flag-IDH2-K272R plasmids, followed by enrichment using anti-Flag antibody (66008-4-Ig, Proteintech, China). Additionally, under HG hypoxic conditions, enzymes regulating IDH2 lactylation and de-lactylation are screened by culturing CMECs treated with low or HG and enriching proteins interacting with IDH2-K272la using IDH2-K272la antibody-bound beads. Eluted proteins from immunoprecipitation undergo SDS-PAGE for separation, followed by trypsin digestion of gel bands. Further analysis involves chromatographic separation of samples using an Easy-nLC 1000 system and tandem mass spectrometry on a QE mass spectrometer. Mascot 2.2.2 software is utilized for protein identification, and subsequent protein-protein binding spatial conformation prediction is performed using ZDOCK SERVER (https://zdock.wenglab.org/), with visualization using PyMOL (https://www.pymol.org/). PDBePISA (https://www.ebi.ac.uk/pdbe/pisa/) analyzes binding sequences between proteins, enabling screening for peptide sequences with the strongest binding affinity.

Lactylation proteomics

The protein extraction process from isolated hearts involved enzymatic digestion based on the protein concentrations, with equal amounts of protein digested and volumes normalized using lysis buffer (8 M urea, 1% protease inhibitor cocktail, 3 μM Trichostatin A, 50 mM nicotinamide). Trichloroacetic acid was added slowly, followed by agitation and a 2-h incubation at 4 °C, before centrifugation at 4500 × g for 5 min at 4 °C, removal of the supernatant, and washing of the pellets with pre-chilled acetone. The dried pellets were resuspended in a tetraethylammonium bromide solution, sonicated, and digested overnight with trypsin. Subsequent steps included reduction with dithiothreitol, alkylation with iodoacetamide, and incubation in the dark. For enriching lactylation-modified peptide segments, an immunoprecipitation experiment using anti-L-lactyllysine antibody conjugated agarose beads (PTM-1404, PTMBIO, China) was conducted, followed by desalting with C18 ZipTips and analysis using liquid chromatography-tandem mass spectrometry (LC-MS/MS). The LC-MS analysis involved peptide separation using a high-performance liquid chromatography system, ionization in the Capillary source, and analysis with the timsTOF Pro mass spectrometer, with data acquisition in parallel accumulation serial fragmentation mode. Database searches of the experimental MS/MS data were performed using MaxQuant(v1.6.15.0) software with specific parameters for enzyme specificity, mass tolerances, variable modifications, and false discovery rate (FDR) control. The relative quantification values of each lysine lactylation site were averaged across the biological replicates to determine the fold change (FC). Differential lactylation was defined as FC cutoff of >1.5 for significant upregulation and <0.67 for significant downregulation. Additionally, a t-test was performed on the relative quantification values between the comparison groups, with a p-value threshold of <0.05. The comprehensive workflow encompassed multiple intricate steps to extract proteins, enrich lactylation-modified peptide segments, and analyze the samples using cutting-edge mass spectrometry techniques for in-depth characterization and identification of proteins and post-translational modifications.

Bioinformatics analysis

Differential lactylation protein bioinformatics analysis involved pathway enrichment significance analysis using Fisher’s exact test, with a P value < 0.05 considered significant. Gene Ontology (GO) analysis was performed using eggnog-mapper software (v2.0) based on the EggNOG database, categorizing proteins by cellular component, molecular function, and biological process. Protein structural domain annotation was conducted using the Pfam database and the PfamScan tool. Subcellular localization predictions were analyzed using PSORTb software (v3.0). EggNOG provided comprehensive homologous classification and functional annotation. GO enrichment analysis classified protein GO annotations into three main categories, with Fisher’s exact test for significance analysis. The Kyoto Encyclopedia of Genes and Genomes (KEGG) database was used for pathway enrichment analysis.

Cell culture

Neonatal C57 BL/6J mice aged 1–3 days were euthanized with excess carbon dioxide, disinfected in 75% ethanol for 10 s, and then transferred to a sterile workstation for the isolation of primary mouse CMECs. After securing the mice, a midline incision along the left clavicle was made to extract the heart. The heart was minced into approximately 1 mm³ tissue blocks, digested with a 0.2% type II collagenase (17101015, Gibco, USA) solution in serum-free DMEM, and subsequently washed and resuspended in HG DMEM medium containing 10% FBS. Following digestion, the cell suspension was filtered, centrifuged, and resuspended in culture medium before being plated and incubated. After 2 h, the supernatant containing suspended endothelial cells was transferred to another dish with ECM medium (1001, ScienCell, USA) on gelatin-coated dishes for 2–3 days. Upon culturing primary CMECs for 2–3 passages, cell experiments were initiated. To prepare a HG culture medium, glucose powder was added to normal-glucose (NG) ECM medium with a glucose concentration of 5.5 mmol/L, reaching a final concentration of 25 mmol/L. The medium was then filtered to ensure sterility. To simulate a hypoxic environment resembling MI injury, cells treated with normal-glucose and HG medium were cultured in a 37 °C, 5% CO₂, 95% N₂ incubator to induce oxygen deprivation (OD). Various compounds including ACAT1 inhibitor Avasimibe (HY-13215, 20 µM), HDAC1 inhibitor Entinostat (HY-12163, 250 nM), sodium lactate (HY-B2227B, 25 mM), MCT1 inhibitor AZD3965 (HY-12750, 0.5 µM), Canagliflozin (HY-10451, 20 µM), EMPA (HY-15409, 1 µM), Dapagliflozin (HY-10450, 20 µM) obtained from MedChemExpress (USA), or corresponding vehicles were added to the medium for 24 hours. Plasmids obtained from Zebrafish Biotech were transfected into CMECs, encoding wild-type (WT) IDH2, single lysine-to-arginine IDH2 mutants (K272R), and various structural domains of Cav1: Cav1 FL (full length), Cav1 ∆1-81 (deletion of amino acids 1-81), Cav1 ∆82-101 (deletion of amino acids 82-101), and Cav1 ∆102-178 (deletion of amino acids 102-178).

CCK8 assay

The treated CMECs were neutralized with trypsin, centrifuged to collect cells, and then resuspended in ECM medium. They were seeded into a 96-well plate and placed in a 37 °C, 5% CO₂ incubator overnight. Upon reaching approximately 80% confluence, the medium was changed based on the different experimental conditions. After removing the supernatant, 10% CCK8 working solution (CK04, Dojindo, Japan) was added, prepared with fresh complete medium while ensuring no bubble formation and working in the dark. The absorbance at 450 nm was measured every half hour using a microplate reader at 37 °C for subsequent statistical analysis.

EdU cell proliferation assay

The EdU stock solution (C0078, Beyotime, China) was diluted to a 2× working solution and mixed with cell culture medium at a 1:1 ratio to achieve a final concentration of 1×. After 4 h of incubation, cells were fixed with 4% paraformaldehyde, permeabilized with PBS containing 0.25% Triton X-100, and then subjected to the Click reaction mixture for 30 min. Following a wash, cells were stained with Hoechst solution for 12 min. Finally, capture images under a fluorescence microscope.

Wound healing assay

When cells reach 80% confluency in a 6-well plate, create a vertical scratch using a sterile pipette tip. Clean the scratch area by washing away floating cells with PBS, then treat cells with fresh culture medium according to experimental groups. Incubate the plate at 37 °C; for the oxygen deprivation group, incubate at 37 °C with 5% CO₂ and 95% N₂. Capture microscope images at 0 and 24 h and analyze migration rate using Image J software.

Tube formation assay

Pre-cool a 96-well plate and 200 µl pipette tips at 4 °C the day before, and thaw Matrigel matrix gel (356234, Corning, USA) at 4 °C from −20 °C. Inject 75 µl of the gel into each well on ice, then allow it to solidify at 37 °C for 30 min. Digest CMECs with 0.25% trypsin, collect them, and resuspend at a concentration of 3 × 10⁵ cells/mL. Gently add 100 µl of the cell suspension per well onto the solidified gel. Incubate the plate at 37 °C and monitor tube formation hourly under a microscope.

NO activity assay

After treating cells with different stimuli, discard the culture medium and wash three times with PBS solution. Prepare a fresh solution of DAF-FM DA (BL767A, Biosharp, China) in serum-free and phenol red-free medium at a final concentration of 5 µmol/L. Add this solution to the culture plate, ensuring even coverage of the cells. Incubate the plate at 37 °C for 20 min. After incubation, wash three times with PBS for 5 min each to remove excess probe that has not been taken up by the cells. Finally, observe the fluorescence intensity of the probe using a fluorescence microscope.

Lactate measurement

The concentrations of intracellular lactate in CMECs were measured using the Lactic Acid Content Assay Kit (BL868B, Biosharp, China).

Statistical analyses

Data are presented as mean ± standard error of the mean (SEM). Replicate numbers for each experiment and results of the statistical analyses are mentioned in the Figure legends. Two-sided unpaired Student’s t test was employed to compare two groups, while one-way analysis of variance (ANOVA) followed by Turkey’s post-hoc test was used for multiple groups. P values are indicated in the Figures. P < 0.05 was considered statistically significant. All statistical analyses were performed, and graphs were generated with GraphPad Prism 9.5.0.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Supplementary information

Supplementary Information^{(2.7MB, pdf)}

Reporting Summary^{(86KB, pdf)}

Transparent Peer Review file^{(1MB, pdf)}

Source data

Source data^{(21.2MB, xlsx)}

Acknowledgements

We thank W. Zhang from the Department of Cardiology, Qilu Hospital of Shandong University. This work was supported by National Natural Science Foundation of China (82370457 to Z.W.); Jiangsu Provincial Key Research and Development Program (BE2022780 to Z.W.).

Author contributions

Z.W. conceived and supervised the overall project. L.L. revised the manuscript. G.Z. designed and performed the majority of the in vitro and in vivo studies. G.Z. and S.X. analyzed the data and drafted the manuscript. L.Z. and H.Y. provided assistance in the in vitro experiments. Y.Q. and Z.S. provided guidance on data analysis and graphing. All authors have read and approved the article.

Peer review

Peer review information

Nature Communications thanks Dharendra Thapa and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Data availability

Data supporting the findings of this study are available in the article and its Supplementary information. Source data are provided as Source data file and may be obtained from the corresponding authors upon request. The proteomics data have been deposited in the ProteomeXchange Consortium via the iProX partner repository with the dataset identifier PXD056842. Source data are provided with this paper.

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-025-67877-0.

References

1.Prabhu, S. D. & Frangogiannis, N. G. The biological basis for cardiac repair after myocardial infarction: from inflammation to fibrosis. Circ. Res.119, 91–112 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
2.Hausenloy, D. J. et al. The coronary circulation in acute myocardial ischaemia/reperfusion injury: a target for cardioprotection. Cardiovasc. Res.115, 1143–1155 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
3.van der Laan, A. M., Piek, J. J. & van Royen, N. Targeting angiogenesis to restore the microcirculation after reperfused MI. Nat. Rev. Cardiol.6, 515–523 (2009). [DOI] [PubMed] [Google Scholar]
4.Dubé, K. N. et al. Recapitulation of developmental mechanisms to revascularize the ischemic heart. JCI Insight2, e96800 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]
5.Bao, X. L. et al. Vasostatin-2 associates with coronary collateral vessel formation in diabetic patients and promotes angiogenesis via angiotensin-converting enzyme 2. Eur. Heart J.44, 1732–1744 (2023). [DOI] [PubMed] [Google Scholar]
6.Elliott, M. D. et al. Prevalence and prognosis of unrecognized myocardial infarction in asymptomatic patients with diabetes: a two-center study with up to 5 years of follow-up. Diabetes Care42, 1290–1296 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
7.Li, X. et al. Lactate metabolism in human health and disease. Signal Transduct. Target. Ther.7, 305 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
8.Chen, X. et al. Lactylation-driven FTO targets CDK2 to aggravate microvascular anomalies in diabetic retinopathy. EMBO Mol. Med.16, 294–318 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Wang, X. et al. YY1 lactylation in microglia promotes angiogenesis through transcription activation-mediated upregulation of FGF2. Genome Biol.24, 87 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
10.Wang, N. et al. Histone lactylation boosts reparative gene activation post-myocardial infarction. Circ. Res.131, 893–908 (2022). [DOI] [PubMed] [Google Scholar]
11.Zhang, N. et al. α-myosin heavy chain lactylation maintains sarcomeric structure and function and alleviates the development of heart failure. Cell Res.33, 679–698 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
12.Rudà, R., Horbinski, C., van den Bent, M., Preusser, M. & Soffietti, R. IDH inhibition in gliomas: from preclinical models to clinical trials. Nat. Rev. Neurol.20, 395–407 (2024). [DOI] [PubMed] [Google Scholar]
13.Wang, Y. et al. IDH2 reprograms mitochondrial dynamics in cancer through a HIF-1α-regulated pseudohypoxic state. FASEB J.33, 13398–13411 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
14.Ma, L. et al. Hypertrophic preconditioning attenuates myocardial ischemia/reperfusion injury through the deacetylation of isocitrate dehydrogenase 2. Sci. Bull.66, 2099–2114 (2021). [DOI] [PubMed] [Google Scholar]
15.Zhou, L. et al. SIRT5 promotes IDH2 desuccinylation and G6PD deglutarylation to enhance cellular antioxidant defense. EMBO Rep.17, 811–822 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]
16.Dai, X. et al. Nrf2 transcriptional upregulation of IDH2 to tune mitochondrial dynamics and rescue angiogenic function of diabetic EPCs. Redox Biol.56, 102449 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
17.Jamaiyar, A. et al. The essential role for endothelial cell sprouting in coronary collateral growth. J. Mol. Cell. Cardiol.165, 158–171 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
18.Frantz, S., Hundertmark, M. J., Schulz-Menger, J., Bengel, F. M. & Bauersachs, J. Left ventricular remodelling post-myocardial infarction: pathophysiology, imaging, and novel therapies. Eur. Heart J.43, 2549–2561 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Luque Contreras, D., Vargas Robles, H., Romo, E., Rios, A. & Escalante, B. The role of nitric oxide in the post-ischemic revascularization process. Pharmacol. Ther.112, 553–563 (2006). [DOI] [PubMed] [Google Scholar]
20.Feron, O. & Balligand, J. L. Caveolins and the regulation of endothelial nitric oxide synthase in the heart. Cardiovasc. Res.69, 788–797 (2006). [DOI] [PubMed] [Google Scholar]
21.Lim, J. E., Bernatchez, P. & Nabi, I. R. Scaffolds and the scaffolding domain: an alternative paradigm for caveolin-1 signaling. Biochem. Soc. Trans.52, 947–959 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]
22.Zhang, D. et al. Metabolic regulation of gene expression by histone lactylation. Nature574, 575–580 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]
23.Felmlee, M. A., Jones, R. S., Rodriguez-Cruz, V., Follman, K. E. & Morris, M. E. Monocarboxylate transporters (SLC16): function, regulation, and role in health and disease. Pharmacol. Rev.72, 466–485 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]
24.Shaul, P. W. Regulation of endothelial nitric oxide synthase: location, location, location. Annu. Rev. Physiol.64, 749–774 (2002). [DOI] [PubMed] [Google Scholar]
25.Zhou, H. et al. Endothelial BACE1 impairs cerebral small vessels via tight junctions and eNOS. Circ. Res.130, 1321–1341 (2022). [DOI] [PubMed] [Google Scholar]
26.Lu, H. et al. Endothelial Sp1/Sp3 are essential to the effect of captopril on blood pressure in male mice. Nat. Commun.14, 5891 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]
27.Gillette, T. G. & Hill, J. A. Readers, writers, and erasers: chromatin as the whiteboard of heart disease. Circ. Res.116, 1245–1253 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]
28.Shang, S., Liu, J. & Hua, F. Protein acylation: mechanisms, biological functions and therapeutic targets. Signal Transduct. Target. Ther.7, 396 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
29.Tomita, K. et al. Acyl-CoA:cholesterol acyltransferase 1 mediates liver fibrosis by regulating free cholesterol accumulation in hepatic stellate cells. J. Hepatol.61, 98–106 (2014). [DOI] [PubMed] [Google Scholar]
30.Zhu, Y. et al. Dynamic regulation of ME1 phosphorylation and acetylation affects lipid metabolism and colorectal tumorigenesis. Mol. Cell77, 138–149.e5 (2020). [DOI] [PubMed] [Google Scholar]
31.Chen, D. et al. Lysine acetylation restricts mutant IDH2 activity to optimize transformation in AML cells. Mol. Cell81, 3833–3847.e11 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]
32.Sun, J. et al. Valproic acid targets HDAC1/2 and HDAC1/PTEN/Akt signalling to inhibit cell proliferation via the induction of autophagy in gastric cancer. FEBS J.287, 2118–2133 (2020). [DOI] [PubMed] [Google Scholar]
33.Herr, D. J. et al. HDAC1 localizes to the mitochondria of cardiac myocytes and contributes to early cardiac reperfusion injury. J. Mol. Cell. Cardiol.114, 309–319 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]
34.Moreno-Yruela, C. et al. Class I histone deacetylases (HDAC1-3) are histone lysine delactylases. Sci. Adv.8, eabi6696 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
35.Halestrap, A. P. The SLC16 gene family - structure, role and regulation in health and disease. Mol. Aspects Med.34, 337–349 (2013). [DOI] [PubMed] [Google Scholar]
36.Ahmed, K. et al. An autocrine lactate loop mediates insulin-dependent inhibition of lipolysis through GPR81. Cell Metab.11, 311–319 (2010). [DOI] [PubMed] [Google Scholar]
37.Brown, T. P. & Ganapathy, V. Lactate/GPR81 signaling and proton motive force in cancer: Role in angiogenesis, immune escape, nutrition, and Warburg phenomenon. Pharmacol. Ther.206, 107451 (2020). [DOI] [PubMed] [Google Scholar]
38.Uhlén, M. et al. Proteomics. Tissue-based map of the human proteome. Science347, 1260419 (2015). [DOI] [PubMed] [Google Scholar]
39.O’Hara, D. V. et al. Applications of SGLT2 inhibitors beyond glycaemic control. Nat. Rev. Nephrol. 20, 513–529 (2024). [DOI] [PubMed]
40.Zinman, B. et al. Empagliflozin, cardiovascular outcomes, and mortality in type 2 diabetes. N. Engl. J. Med.373, 2117–2128 (2015). [DOI] [PubMed] [Google Scholar]
41.Neal, B. et al. Canagliflozin and cardiovascular and renal events in type 2 diabetes. N. Engl. J. Med.377, 644–657 (2017). [DOI] [PubMed] [Google Scholar]
42.Sattar, N., Anker, S. D., Butler, J. & Verma, S. Obesity, heart failure, and SGLT2 inhibition: DECLARE-TIMI 58 provides insights. Eur. Heart J.43, 2968–2970 (2022). [DOI] [PubMed] [Google Scholar]
43.Paolisso, P. et al. Infarct size, inflammatory burden, and admission hyperglycemia in diabetic patients with acute myocardial infarction treated with SGLT2-inhibitors: a multicenter international registry. Cardiovasc. Diabetol.21, 77 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]
44.Santos-Gallego, C. G. et al. Empagliflozin ameliorates diastolic dysfunction and left ventricular fibrosis/stiffness in nondiabetic heart failure. JACC14, 393–407 (2021). [DOI] [PubMed] [Google Scholar]
45.Gao, E. et al. A novel and efficient model of coronary artery ligation and myocardial infarction in the mouse. Circ. Res.107, 1445–1453 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]
46.Pei, J. et al. LPA(2) contributes to vascular endothelium homeostasis and cardiac remodeling after myocardial infarction. Circ. Res.131, 388–403 (2022). [DOI] [PubMed] [Google Scholar]
47.Wang, Z. et al. Nuclear Tkt promotes ischemic heart failure via the cleaved Parp1/Aif axis. Basic Res. Cardiol.117, 18 (2022). [DOI] [PubMed] [Google Scholar]
48.Chen, Q. et al. Nrf3-Mediated mitochondrial superoxide promotes cardiomyocyte apoptosis and impairs cardiac functions by suppressing Pitx2. Circulation151, 1024–1046 (2025). [DOI] [PubMed] [Google Scholar]
49.Wang, X. et al. STING aggravates ferroptosis-dependent myocardial ischemia-reperfusion injury by targeting GPX4 for autophagic degradation. Signal Transduct. Target. Ther.10, 136 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Information^{(2.7MB, pdf)}

Reporting Summary^{(86KB, pdf)}

Transparent Peer Review file^{(1MB, pdf)}

Source data^{(21.2MB, xlsx)}

Data Availability Statement

[CR1] 1.Prabhu, S. D. & Frangogiannis, N. G. The biological basis for cardiac repair after myocardial infarction: from inflammation to fibrosis. Circ. Res.119, 91–112 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR2] 2.Hausenloy, D. J. et al. The coronary circulation in acute myocardial ischaemia/reperfusion injury: a target for cardioprotection. Cardiovasc. Res.115, 1143–1155 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR3] 3.van der Laan, A. M., Piek, J. J. & van Royen, N. Targeting angiogenesis to restore the microcirculation after reperfused MI. Nat. Rev. Cardiol.6, 515–523 (2009). [DOI] [PubMed] [Google Scholar]

[CR4] 4.Dubé, K. N. et al. Recapitulation of developmental mechanisms to revascularize the ischemic heart. JCI Insight2, e96800 (2017). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR5] 5.Bao, X. L. et al. Vasostatin-2 associates with coronary collateral vessel formation in diabetic patients and promotes angiogenesis via angiotensin-converting enzyme 2. Eur. Heart J.44, 1732–1744 (2023). [DOI] [PubMed] [Google Scholar]

[CR6] 6.Elliott, M. D. et al. Prevalence and prognosis of unrecognized myocardial infarction in asymptomatic patients with diabetes: a two-center study with up to 5 years of follow-up. Diabetes Care42, 1290–1296 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR7] 7.Li, X. et al. Lactate metabolism in human health and disease. Signal Transduct. Target. Ther.7, 305 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR8] 8.Chen, X. et al. Lactylation-driven FTO targets CDK2 to aggravate microvascular anomalies in diabetic retinopathy. EMBO Mol. Med.16, 294–318 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR9] 9.Wang, X. et al. YY1 lactylation in microglia promotes angiogenesis through transcription activation-mediated upregulation of FGF2. Genome Biol.24, 87 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR10] 10.Wang, N. et al. Histone lactylation boosts reparative gene activation post-myocardial infarction. Circ. Res.131, 893–908 (2022). [DOI] [PubMed] [Google Scholar]

[CR11] 11.Zhang, N. et al. α-myosin heavy chain lactylation maintains sarcomeric structure and function and alleviates the development of heart failure. Cell Res.33, 679–698 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR12] 12.Rudà, R., Horbinski, C., van den Bent, M., Preusser, M. & Soffietti, R. IDH inhibition in gliomas: from preclinical models to clinical trials. Nat. Rev. Neurol.20, 395–407 (2024). [DOI] [PubMed] [Google Scholar]

[CR13] 13.Wang, Y. et al. IDH2 reprograms mitochondrial dynamics in cancer through a HIF-1α-regulated pseudohypoxic state. FASEB J.33, 13398–13411 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR14] 14.Ma, L. et al. Hypertrophic preconditioning attenuates myocardial ischemia/reperfusion injury through the deacetylation of isocitrate dehydrogenase 2. Sci. Bull.66, 2099–2114 (2021). [DOI] [PubMed] [Google Scholar]

[CR15] 15.Zhou, L. et al. SIRT5 promotes IDH2 desuccinylation and G6PD deglutarylation to enhance cellular antioxidant defense. EMBO Rep.17, 811–822 (2016). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR16] 16.Dai, X. et al. Nrf2 transcriptional upregulation of IDH2 to tune mitochondrial dynamics and rescue angiogenic function of diabetic EPCs. Redox Biol.56, 102449 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR17] 17.Jamaiyar, A. et al. The essential role for endothelial cell sprouting in coronary collateral growth. J. Mol. Cell. Cardiol.165, 158–171 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR18] 18.Frantz, S., Hundertmark, M. J., Schulz-Menger, J., Bengel, F. M. & Bauersachs, J. Left ventricular remodelling post-myocardial infarction: pathophysiology, imaging, and novel therapies. Eur. Heart J.43, 2549–2561 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR19] 19.Luque Contreras, D., Vargas Robles, H., Romo, E., Rios, A. & Escalante, B. The role of nitric oxide in the post-ischemic revascularization process. Pharmacol. Ther.112, 553–563 (2006). [DOI] [PubMed] [Google Scholar]

[CR20] 20.Feron, O. & Balligand, J. L. Caveolins and the regulation of endothelial nitric oxide synthase in the heart. Cardiovasc. Res.69, 788–797 (2006). [DOI] [PubMed] [Google Scholar]

[CR21] 21.Lim, J. E., Bernatchez, P. & Nabi, I. R. Scaffolds and the scaffolding domain: an alternative paradigm for caveolin-1 signaling. Biochem. Soc. Trans.52, 947–959 (2024). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR22] 22.Zhang, D. et al. Metabolic regulation of gene expression by histone lactylation. Nature574, 575–580 (2019). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR23] 23.Felmlee, M. A., Jones, R. S., Rodriguez-Cruz, V., Follman, K. E. & Morris, M. E. Monocarboxylate transporters (SLC16): function, regulation, and role in health and disease. Pharmacol. Rev.72, 466–485 (2020). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR24] 24.Shaul, P. W. Regulation of endothelial nitric oxide synthase: location, location, location. Annu. Rev. Physiol.64, 749–774 (2002). [DOI] [PubMed] [Google Scholar]

[CR25] 25.Zhou, H. et al. Endothelial BACE1 impairs cerebral small vessels via tight junctions and eNOS. Circ. Res.130, 1321–1341 (2022). [DOI] [PubMed] [Google Scholar]

[CR26] 26.Lu, H. et al. Endothelial Sp1/Sp3 are essential to the effect of captopril on blood pressure in male mice. Nat. Commun.14, 5891 (2023). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR27] 27.Gillette, T. G. & Hill, J. A. Readers, writers, and erasers: chromatin as the whiteboard of heart disease. Circ. Res.116, 1245–1253 (2015). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR28] 28.Shang, S., Liu, J. & Hua, F. Protein acylation: mechanisms, biological functions and therapeutic targets. Signal Transduct. Target. Ther.7, 396 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR29] 29.Tomita, K. et al. Acyl-CoA:cholesterol acyltransferase 1 mediates liver fibrosis by regulating free cholesterol accumulation in hepatic stellate cells. J. Hepatol.61, 98–106 (2014). [DOI] [PubMed] [Google Scholar]

[CR30] 30.Zhu, Y. et al. Dynamic regulation of ME1 phosphorylation and acetylation affects lipid metabolism and colorectal tumorigenesis. Mol. Cell77, 138–149.e5 (2020). [DOI] [PubMed] [Google Scholar]

[CR31] 31.Chen, D. et al. Lysine acetylation restricts mutant IDH2 activity to optimize transformation in AML cells. Mol. Cell81, 3833–3847.e11 (2021). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR32] 32.Sun, J. et al. Valproic acid targets HDAC1/2 and HDAC1/PTEN/Akt signalling to inhibit cell proliferation via the induction of autophagy in gastric cancer. FEBS J.287, 2118–2133 (2020). [DOI] [PubMed] [Google Scholar]

[CR33] 33.Herr, D. J. et al. HDAC1 localizes to the mitochondria of cardiac myocytes and contributes to early cardiac reperfusion injury. J. Mol. Cell. Cardiol.114, 309–319 (2018). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR34] 34.Moreno-Yruela, C. et al. Class I histone deacetylases (HDAC1-3) are histone lysine delactylases. Sci. Adv.8, eabi6696 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR35] 35.Halestrap, A. P. The SLC16 gene family - structure, role and regulation in health and disease. Mol. Aspects Med.34, 337–349 (2013). [DOI] [PubMed] [Google Scholar]

[CR36] 36.Ahmed, K. et al. An autocrine lactate loop mediates insulin-dependent inhibition of lipolysis through GPR81. Cell Metab.11, 311–319 (2010). [DOI] [PubMed] [Google Scholar]

[CR37] 37.Brown, T. P. & Ganapathy, V. Lactate/GPR81 signaling and proton motive force in cancer: Role in angiogenesis, immune escape, nutrition, and Warburg phenomenon. Pharmacol. Ther.206, 107451 (2020). [DOI] [PubMed] [Google Scholar]

[CR38] 38.Uhlén, M. et al. Proteomics. Tissue-based map of the human proteome. Science347, 1260419 (2015). [DOI] [PubMed] [Google Scholar]

[CR39] 39.O’Hara, D. V. et al. Applications of SGLT2 inhibitors beyond glycaemic control. Nat. Rev. Nephrol. 20, 513–529 (2024). [DOI] [PubMed]

[CR40] 40.Zinman, B. et al. Empagliflozin, cardiovascular outcomes, and mortality in type 2 diabetes. N. Engl. J. Med.373, 2117–2128 (2015). [DOI] [PubMed] [Google Scholar]

[CR41] 41.Neal, B. et al. Canagliflozin and cardiovascular and renal events in type 2 diabetes. N. Engl. J. Med.377, 644–657 (2017). [DOI] [PubMed] [Google Scholar]

[CR42] 42.Sattar, N., Anker, S. D., Butler, J. & Verma, S. Obesity, heart failure, and SGLT2 inhibition: DECLARE-TIMI 58 provides insights. Eur. Heart J.43, 2968–2970 (2022). [DOI] [PubMed] [Google Scholar]

[CR43] 43.Paolisso, P. et al. Infarct size, inflammatory burden, and admission hyperglycemia in diabetic patients with acute myocardial infarction treated with SGLT2-inhibitors: a multicenter international registry. Cardiovasc. Diabetol.21, 77 (2022). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR44] 44.Santos-Gallego, C. G. et al. Empagliflozin ameliorates diastolic dysfunction and left ventricular fibrosis/stiffness in nondiabetic heart failure. JACC14, 393–407 (2021). [DOI] [PubMed] [Google Scholar]

[CR45] 45.Gao, E. et al. A novel and efficient model of coronary artery ligation and myocardial infarction in the mouse. Circ. Res.107, 1445–1453 (2010). [DOI] [PMC free article] [PubMed] [Google Scholar]

[CR46] 46.Pei, J. et al. LPA(2) contributes to vascular endothelium homeostasis and cardiac remodeling after myocardial infarction. Circ. Res.131, 388–403 (2022). [DOI] [PubMed] [Google Scholar]

[CR47] 47.Wang, Z. et al. Nuclear Tkt promotes ischemic heart failure via the cleaved Parp1/Aif axis. Basic Res. Cardiol.117, 18 (2022). [DOI] [PubMed] [Google Scholar]

[CR48] 48.Chen, Q. et al. Nrf3-Mediated mitochondrial superoxide promotes cardiomyocyte apoptosis and impairs cardiac functions by suppressing Pitx2. Circulation151, 1024–1046 (2025). [DOI] [PubMed] [Google Scholar]

[CR49] 49.Wang, X. et al. STING aggravates ferroptosis-dependent myocardial ischemia-reperfusion injury by targeting GPX4 for autophagic degradation. Signal Transduct. Target. Ther.10, 136 (2025). [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

IDH2 lactylation promotes angiogenesis in murine diabetic myocardial infarction via blocking Cav1-eNOS interaction

Guangyao Zang

Siting Xu

Zhen Sun

Lili Zhang

Yongjiang Qian

Haipeng Yao

Lihua Li

Zhongqun Wang

Abstract

Introduction

Results

Increased cardiac lactylation modification following DMI

Fig. 1. Increased cardiac lactylation modification following diabetic myocardial infarction (DMI).

Proteomic profiling of lysine lactylation following diabetic myocardial infarction

Fig. 2. Proteomic profiling of lysine lactylation following diabetic myocardial infarction (DMI).

Inhibition of IDH2 lactylation at Lys272 suppresses angiogenesis in high glucose and hypoxia environment

Fig. 3. Inhibition of IDH2 lactylation at Lys272 suppresses angiogenesis in high glucose and hypoxia environment.

IDH2-K272 mutation in endothelial suppresses angiogenesis and cardiac function in DMI

Fig. 4. IDH2-K272 mutation in endothelial suppresses angiogenesis and cardiac function after diabetic myocardial infarction (DMI).

IDH2-K272 lactylation triggers eNOS activation by coupling with Cav1

Fig. 5. IDH2-K272 lactylation triggers eNOS activation by coupling with Cav1.

ACAT1 and HDAC1 are responsible for lactylation and delactylation, respectively, at K272 of IDH2

Fig. 6. ACAT1 is the lactyltransferase for IDH2-K272.

Lactate is transported intracellularly via MCT1 to facilitate lactylation of IDH2

Empagliflozin enhances angiogenesis and cardiac function by augmenting IDH2-K272 lactylation following DMI

Fig. 7. Empagliflozin (EMPA) induces angiogenesis and ventricular remodeling by augmenting IDH2-K272 lactylation following diabetic myocardial infarction (DMI).

Discussion

Methods

Animal study

Diabetes model

Myocardial infarction model

TTC staining

Microfil cardiac vascular imaging

Aortic ring assay

Cardiac ultrasound

Masson’s trichrome staining

Immunofluorescence staining

Western blot and immunoprecipitation

IDH2 in vitro binding assay

Lactylation proteomics

Bioinformatics analysis

Cell culture

CCK8 assay

EdU cell proliferation assay

Wound healing assay

Tube formation assay

NO activity assay

Lactate measurement

Statistical analyses

Reporting summary

Supplementary information

Source data

Acknowledgements

Author contributions

Peer review

Peer review information

Data availability

Competing interests

Footnotes

Supplementary information

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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