Lactylation and lactate: insights into muscle cell epigenetic regulators in exercise

Abstract

Lactate was once regarded as a metabolic waste. However, recent studies have established its dual role as both a versatile signaling molecule and a substrate for a novel post-translational modification (PTMs)—lysine lactylation. Exercise, as a primary physiological stimulus for lactate production, provides an ideal context for exploring lactate-mediated epigenetic regulation. This regulatory mechanism is particularly relevant in muscle cells, which serve as both the main producers and consumers of lactate. This review focuses on three core muscle cell types: skeletal muscle cells, cardiomyocytes and vascular smooth muscle cells (VSMCs). We systematically elucidate the functional roles of the lactate-lactylation in these cells, examining how lactate can regulate cellular adaptation and pathological processes both independently and in concert with other PTMs. Furthermore, we analyze the distinct effects of diferent exercise modalities (ranging from moderate training to overtraining) on the lactylation dynamics, revealing its the dual functions of lactylation in exercise physiology and pathology. By integrating these mechanisms, this review provides a novel theoretical framework for understanding exercise-mediated muscle cell function and offers fresh perspectives for intervention strategies targeting metabolic and cardiovascular diseases.

Graphical Abstract

Introduction

Muscle cells are constantly exposed to changes in energy demand, particularly during exercise, when contraction, ion transport, repair, and remodeling require rapid adjustments of substrate use. Although skeletal muscle cells, cardiomyocytes, and VSMCs differ in structure and function, they share a reliance on flexible metabolic regulation (Hjorth et al. 2016). Under conditions of increased workload, cellular stress, or altered oxygen availability, glycolysis is activated and lactate production increases. Once viewed primarily as a glycolytic end product, lactate is now recognized as an important participant in energy transfer and cellular signaling (Clark and Pyne-Geithman 2005). This change is particularly relevant to muscle-related tissues. Skeletal muscle is a major source of circulating lactate during exercise, whereas the heart can take up and oxidize lactate as a fuel (Brooks 2021). VSMCs are also metabolically linked to lactate, especially during the transition from a contractile phenotype to a synthetic or pathological phenotype (Yang et al. 2017). Together, these observations indicate that lactate should not be discussed merely as a metabolic byproduct. In muscle-related cells, lactate production, transport, uptake, and utilization can collectively influence cellular adaptation under both physiological and disease conditions.

The discovery of lysine lactylation (Kla) provides a further reason to revisit the role of lactate in muscle biology (Zhang et al. 2019). Kla is a lactate-derived post-translational modification that occurs on histone and non-histone proteins. Histone lactylation affects chromatin state and gene transcription, whereas non-histone lactylation potentially modulates enzyme activity, protein stability, and protein interactions. Lactate can thus influence muscle-related cells as an energy substrate, a signaling molecule, and a source of protein lactylation. This multifunctional role makes the lactate–lactylation axis a productive framework for examining how metabolic changes are connected with muscle cell function. Although lactylation has been widely investigated in cancer, immune regulation, inflammation, and neurological disorders, the evidence in muscle-related cells remains scattered. Lactylation has been implicated in processes as diverse as skeletal muscle autophagy, myogenesis, mitochondrial substrate metabolism, cardiac hypertrophy, myocardial ischemia–reperfusion injury, sarcomeric stability; and VSMC senescence, calcification, phenotypic switching, and apoptosis. However, these observations typically derive from isolated disease or cell-type models. A direct comparison among skeletal muscle cells, cardiomyocytes, and VSMCs is still lacking, which limits the ability to distinguish shared mechanisms from cell-type-specific regulatory patterns.

Exercise adds another layer of complexity. Although exercise is a primary physiological driver of lactate production, exercise-induced lactylation does not follow a simple linear dose–response relationship. Moderate training, acute exhaustive exercise, and overtraining can produce distinct lactate dynamics and distinct lactylation responses. In some settings, lactylation appears to support adaptive remodeling, such as autophagy activation, mitochondrial regulation, and stress resistance. In others, excessive or prolonged lactate accumulation may be associated with maladaptive outcomes. Species differences further complicate the interpretation. Rodent studies have documented exercise-responsive lactylation in skeletal muscle and heart, whereas available human studies have not consistently detected skeletal muscle lactylation changes after acute or chronic exercise. This discrepancy may stem from differences in biopsy timing, exercise modality, detection sensitivity, tissue specificity, or species-dependent lactate kinetics.

A structured literature search was conducted in PubMed and Web of Science from database inception to March 31, 2026, using combinations of terms related to lactylation (e.g., “lysine lactylation,” “histone lactylation,” “Kla”), lactate metabolism, muscle cell types (skeletal muscle, cardiomyocyte, VSMC), exercise (training, overtraining), and post-translational regulation. Additional records were identified by manually screening reference lists of relevant original articles and reviews. Studies were included if they met at least one of the following criteria: (i) investigation of lactylation in skeletal muscle cells, cardiomyocytes, or VSMCs; (ii) examination of lactate-dependent regulation of muscle cell function with mechanistic relevance to lactylation; (iii) assessment of exercise-induced changes in lactate signaling or lactylation; or (iv) characterization of lactylation writers, erasers, readers, or crosstalk with other PTMs. Studies were excluded if they focused exclusively on non-muscle tissues without mechanistic relevance to muscle cells, reported lactate metabolism without connection to lactylation, lacked full-text availability, or were conference abstracts, editorials, or non-peer-reviewed reports.

Unlike previous reviews that discuss lactylation mainly in cancer, immunity, or general metabolic disease, this review focuses specifically on muscle-related cell types and exercise physiology. The conceptual contribution of this work is not to propose a new lactylation enzyme or pathway, but to organize currently scattered findings into a comparative framework across skeletal muscle cells, cardiomyocytes, and VSMCs. This framework emphasizes three underexplored dimensions: cell-type specificity, exercise dose dependence, and translational uncertainty between rodent and human studies. By distinguishing adaptive from maladaptive lactylation responses and direct from indirect evidence, this review aims to clarify where the lactate–lactylation axis is mechanistically supported and where further validation is required.

Lactate: from metabolic byproduct to signaling molecule

Lactate production

Lactate accumulates in the cytoplasm under increased energy demand, including strenuous exercise or tissue hypoxia. Lactate biosynthesis proceeds primarily through glycolysis and lactate dehydrogenase (LDH) reaction, with a secondary contribution from glutamine-derived pathways (DeBerardinis et al. 2007). L-lactate, which accounts for over 95% of total lactate, is responsible for most metabolic and signaling functions (Connor et al. 1983). Once dismissed as a mere anaerobic waste product (Ferguson et al. 2018; Rogatzki et al. 2015), lactate is now recognized as a versatile fuel and signaling molecule (Gladden 2004). Circulating lactate flux exceeds that of glucose, consistent with the broad tissue distribution of LDH and monocarboxylate transporters (MCTs) relative to glucose transporters (Jang et al. 2022). Under aerobic conditions, pyruvate enters the TCA cycle; when oxygen availability is limited, accelerated glycolysis diverts pyruvate toward LDH-catalyzed lactate production, regenerating NAD⁺ in the process (Rabinowitz and Enerback 2020). LDH exists as tetramers composed of M-subunits (encoded by LDHA) and H-subunits (encoded by LDHB); LDHA-dominant isoforms, which are enriched in skeletal muscle, favor net lactate generation, whereas LDHB-dominant isoforms in cardiac tissue preferentially catalyze lactate oxidation (de la Cruz-López 2019).

The lactate shuttle

The “lactate shuttle” concept, introduced by Brooks (Brooks 2009; Hashimoto et al. 2006), describes the coordinated transport of lactate at intercellular and intracellular levels. At the intercellular level, lactate is exchanged between glycolytic producer cells and oxidative consumers; across muscle fiber types; between skeletal muscle and organs, such as the heart, liver, and brain (Nishima and Tanaka 2024), and bidirectionally between astrocytes and neurons (Hagihara et al. 2021). Intracellularly, lactate is shuttled between the cytosol and organelles, including the mitochondria and peroxisomes (McClelland et al. 2003). Transmembrane lactate transport is mediated primarily by MCTs, a subfamily of solute carriers (SLCs) comprising 14 known isoforms (Felmlee et al. 2020). MCT1, a high-affinity isoform, facilitates lactate uptake in oxidative tissues such as the myocardium (Gizak et al. 2020; Halestrap 2013). Meanwhile, MCT4, with lower substrate affinity, mediates lactate efflux from glycolytically active cells (Brown and Ganapathy 2020). Lactate also signals through the G protein-coupled receptor GPR81/HCAR1, modulating energy homeostasis, lipid turnover, and inflammatory tone across multiple tissues (Laroche 2021; Wu et al. 2022).

The lactate shuttle is particularly relevant during exercise, when the lactate produced by contracting muscle is distributed to distal organs for oxidative disposal or gluconeogenic recycling (Yan et al. 2024). Aerobic oxidation represents the principal clearance route during and after exercise. Impairment of shuttle function has been associated with reduced exercise capacity and motor-unit instability (Bisetto 2019), underscoring the physiological importance of inter-organ lactate coordination.

Physiological functions of lactate

Lactate serves as an oxidizable fuel, a signaling mediator, and an immunomodulatory agent (Pucino et al. 2018). As a substrate, it feeds into the TCA cycle and contributes to ATP generation across diverse tissues (Hui et al. 2017; Jin et al. 2019). Lactate also triggers endoplasmic reticulum Mg²⁺release; this cation flux acts as a second messenger linking cytosolic lactate levels to mitochondrial bioenergetics (Daw et al. 2020). During hypoglycemia, lactate sustains cerebral energy supply and neuronal function (Brooks 2023; Dienel 2019; Gómez-Valadés et al. 2021). Within the tumor microenvironment, lactate supports metabolic symbiosis among heterogeneous cell populations (Morrot et al. 2018).

Lactate contributes to cellular redox homeostasis by functioning as an intercellular redox-signaling molecule that coordinates oxidative metabolism across tissues (Titov et al. 2016). Increased lactate concentrations increase the NAD⁺/NADH ratio, thereby attenuating glycolytic flux and mitochondrial respiration (Quinn et al. 2020; Wang et al. 2016). When oxidative NAD⁺ demand exceeds its glycolytic regeneration, particularly under mitochondrial respiratory limitation, MCT-dependent intercellular lactate exchange supports NAD⁺ recycling. Disruption of MCT1/MCT4-mediated transport or impairment of LDH activity compromises NAD⁺ regeneration, precipitating ATP depletion and cell death (Benjamin et al. 2018).

Lactate also modulates fatty acid metabolism. Increased circulating lactate suppresses the availability of free fatty acid (Jiang et al. 2021; Querol et al. 2017), an effect mediated largely through GPR81 activation, which inhibits lipolysis in a pH-independent manner (Wu et al. 2022; Liu et al. 2017). Downstream signaling through HCAR1 appears to involve cAMP and CREB-dependent pathways (Li et al. 2020; Vaupel and Multhoff 2021). In CD4⁺T cells, lactate upregulates the transporter SLC5A12, which, in turn, facilitates further lactate uptake and promotes fatty acid synthesis through a positive-feedback loop (Pucino et al. 2019). Exercise-induced increases in circulating lactate have been inversely associated with fat oxidation rates (San-Millán and Brooks 2018). The relationship between lactate and fatty acid oxidation is likely bidirectional, and the molecular mechanisms governing this interplay remain incompletely defined. The pathways linking lactate metabolism to protein lactylation are summarized in Fig. 1, which illustrates the two routes of lactate entry into cells and the subsequent conversion to lactyl-CoA for histone and non-histone lactylation.

Fig. 1.

Intracellular lactate metabolism and lactylation

Lactate enters cells via two main routes. First, extracellular lactate signals through the G protein-coupled receptor GPR81 or is transported into the cell via MCTs. Once inside, lactate can be oxidized to pyruvate, which enters the mitochondria and is metabolized through the TCA cycle. Second, glucose is broken down by glycolysis to produce pyruvate, which is then converted to L-lactate and D-lactate by LDH. Lactate is subsequently converted to lactyl-CoA, which serves as the direct substrate for Kla of both histone and non-histone proteins. This figure illustrates the metabolic pathways that link glycolysis and lactate production to epigenetic regulation via lactylation. (Created with BioRender.com.)

Lactylation: a novel epigenetic regulatory mechanism

Discovery and characterization of protein lactylation

Kla was first identified by Zhang et al. in 2019 as a histone modification linking lactate metabolism to gene regulation (Martin and Zhang 2005; Millán-Zambrano et al. 2022; Park et al. 2022). The enzymatic and non-enzymatic pathways that regulate Kla are depicted in Fig. 2. In addition to the role of histone lactylation, Kla was found on thousands of non-histone proteins across diverse tissues and disease models (Zhang et al. 2019; Fan et al. 2023; Pan et al. 2022; Wei et al. 2023; Xiong et al. 2022).

Fig. 2.

Regulatory mechanisms of lysine lactylation

Lysine lactylation (Kla) is regulated by both enzymatic and non-enzymatic pathways. In the enzymatic pathway, L-lactate is converted to L-lactyl-CoA by acetyltransferases such as p300/CBP. Lactyl-CoA then serves as a donor for the transfer of a lactyl group to specific lysine residues on histones (e.g., H3K18la), thereby regulating gene transcription. In the non-enzymatic pathway, methylglyoxal reacts with glutathione to form lactylglutathione, which spontaneously modifies lysine residues. Together, these mechanisms link cellular metabolic states to epigenetic control of gene expression

The regulation of lactylation system comprises three components: writers that install the modification, erasers that remove it, and readers that recognize it and transduce downstream signals. Current evidence supports p300/CBP, KAT5/TIP60, HBO1, and AARS1/2 as potential writers, whereas class I HDACs and sirtuins have been implicated as erasers (Zhang et al. 2019). Figure 3 provides an overview of the complete lactylation modification systemand highlights how dysregulation of this system contributes to disease pathogenesis. Because many of these enzymes also regulate acetylation or other lysine acylations, distinguishing lactylation-specific effects from broader PTM changes remains a major methodological challenge.

Fig. 3.

The lactylation modification system

The lactylation regulatory system comprises three functional modules: writers, readers, and erasers. Writers (e.g., p300/CBP) catalyze the addition of lactyl groups to lysine residues. Readers (e.g., YEATS domain-containing proteins) recognize and interpret lactylation marks to modulate downstream gene expression. Erasers (e.g., HDACs, particularly HDAC1-3) remove lactyl groups, reversing the modification. This dynamic system orchestrates key cellular processes by linking metabolic flux (lactate availability) to epigenetic regulation. Dysregulation of this system has been implicated in cardiovascular disease, metabolic disorders, and cancer. The figure highlights the major players and their roles in the lactylation cycle

The lactylation regulatory system comprises three functional modules: writers, readers, and erasers. Writers (e.g., p300/CBP) catalyze the addition of lactyl groups to lysine residues. Readers (e.g., YEATS domain-containing proteins) recognize and interpret lactylation marks to modulate downstream gene expression. Erasers (e.g., HDACs, particularly HDAC1-3) remove lactyl groups, reversing the modification. This dynamic system orchestrates key cellular processes by linking metabolic flux (lactate availability) to epigenetic regulation. Dysregulation of this system has been implicated in cardiovascular disease, metabolic disorders, and cancer. The figure highlights the major players and their roles in the lactylation cycle.

Lactylation can be generated through lactoyl-CoA-dependent enzymatic pathways and non-enzymatic reactions. L-lactylation was first identified by mass spectrometry and isotope tracing, whereas D-lactylation appears to arise mainly through non-enzymatic acyl transfer (Wang et al. 2023). Although global lactylomic studies have identified thousands of lactylation sites in different tissues and disease models, the functional relevance of many sites remains unknown. Therefore, site-specific validation is essential for determining whether lactylation is merely associated with metabolic stress or directly regulates protein function (Yang et al. 2022).

Crosstalk between lactylation and other PTMs

Protein function is rarely controlled by a single PTM. Lactylation interacts extensively with classical modifications, such as acetylation, methylation, and ubiquitination, either competitively on shared lysine residue or synergistically on adjacent sites. This section synthesizes current evidence to elucidate lactylation should be interpreted as part of a broader PTM network rather than as an isolated modification (Fig. 4).

Fig. 4.

Crosstalk Between Lactylation and Other PTMs

Lactylation does not function in isolation but interacts with other PTMs, such as acetylation, phosphorylation and succinylation. These interactions can be competitive (when different modifications target the same lysine residue) or synergistic (when modifications on adjacent residues cooperate to regulate protein function). Key examples illustrated include: (i) competition between lactylation and acetylation for shared lysine residues on histones, regulated by p300 and HDACs; (ii) lactylation-dependent regulation of PARP1 activity through competitive inhibition of acetylation; and (iii) synergistic effects between lactylation (H3K18la) and methylation (H3K4me3) at active gene promoters. This network allows integrated cellular responses to metabolic and environmental cues

Competitive interplay with acetylation

Lactylation and acetylation are closely related because both occur on lysine residues and can be regulated by overlapping enzymes, including p300/CBP and HDACs (Wu et al. 2024; Rho et al. 2023). This creates potential competition between lactoyl-CoA- and acetyl-CoA-dependent modifications (Raju and Sankaranarayanan 2025). For example, Double knockdown of p300/HDAC1 selectively reduced the level of lactylation, and acetylation showed characteristic changes (decreased upon p300 knockdown but increased upon HDAC1 knockdown), revealing a substrate competition mechanism (Meng et al. 2025).

A paradigmatic example is poly (ADP-ribose) polymerase 1 (PARP1), whose auto-modification domain contains a lysine residue serving as a shared site for both Kla and Kac. Lactylation restores PARP1’s ADP-ribosyl transferase activity by competitively inhibiting acetylation at this site, thus promoting DNA repair and regulating pluripotent gene expression (Sun et al. 2022). Similarly, in macrophages, MCT-mediated lactate uptake promotes p300/CBP-dependent HMGB1 lactylation, whereas GPR81 signaling activates YAP and recruits p300/CBP through β-arrestin2, simultaneously inhibiting SIRT1 to drive acetylation (Yang et al. 2022).

This interaction exhibits cell type specificity (Yang et al. 2022; Wu et al. 2023). During mouse oocyte maturation and early embryonic development, histone Kla and Kac showed distinct temporal and spatial distribution patterns. Under hypoxia, H3K23la and H3K18la in blastocysts decreased significantly, whereas acetylation at corresponding sites remained stable (Yang et al. 2021). However, lactylation and acetylation do not always act antagonistically. In some contexts, lactate can promote both HMGB1 lactylation and acetylation through p300/CBP-dependent mechanisms. Histone lactylation may also coexist with acetylation marks at active chromatin regions. These findings suggest that the relationship between lactylation and acetylation is context-dependent rather than uniformly competitive (Meng et al. 2021).

Crosstalk with methylation and phosphorylation

Lactylation also interacts with methylation and phosphorylation. H3K18la correlates positively with transcription activation markers such as H3K27ac and H3K4me3 (Galle et al. 2022). Through this chromatin context, lactylation can influence downstream signaling pathways, including STAT3 phosphorylation and METTL3-mediated RNA methylation (Xiong 2022). RNA methylation participates by METTL15 deletion inducing oxidative stress and metabolic reprogramming, promoting lactate production, and increasing H4K12la and H3K9la modifications (Lv et al. 2024). In DNA damage repair, ubiquitination of H2BK123 promotes H3K4/46/79 methylation and regulates lactylation levels, maintaining genome stability (Liu et al. 2020).

In disease contexts, lactylation often appears together with changes in other regulatory marks rather than acting alone. Promoter lactylation may increase the expression of genes related to inflammation or cell proliferation, and lactylation of signaling proteins may further influence phosphorylation-dependent pathways (Liu et al. 2025; Chen et al. 2024; Wang et al. 2023). These observations suggest that lactylation is better viewed as part of a coordinated regulatory network involving methylation and phosphorylation, instead of as a separate modification with a fixed biological effect.

Broader PTM network interactions

Studies showed that lactylation interacts with crotonylation, butyrylation, and succinylation, finely regulating gene expression (Liu et al. 2022; Xie et al. 2022). Crotonylation and lactylation occur on almost all core histones and share multiple modification sites (e.g., H3K9 and H3K18) with classical Kac (Xie et al. 2022), suggesting dynamic competition on the same residue yielding different transcriptional outputs. Lactylation generation may also relate to butyrylation mediated by butyrate metabolism, indicating crosstalk between short-chain fatty acid and lactate metabolic pathways at the epigenetic level. In a study of catalpol-treated breast cancer, a unique PTM spectrum was observed with significant increases in Kla, Kac, and 2-hydroxyisobutyrylation, whereas succinylation, propionylation, and phosphorylation decreased significantly (Liu et al. 2022), demonstrating that catalpol’s antitumor effect involves reprogramming the PTM network.

Therefore, understanding the role of lactylation in other PTM network interactions is important for comprehensively expounding its functions in physiological and pathological processes.

Pathophysiological roles of lactylation in disease

Lactylation has been implicated in diverse pathological processes, including metabolic reprogramming, inflammation (Pan et al. 2022), immune regulation (Lopez Krol et al. 2022; Wang et al. 2023), fibrosis (Mejias et al. 2020), angiogenesis (Wang et al. 2023; Gordon-Weeks et al. 2017), cell cycle progression (Jin et al. 2023), and cell fate determination (Yu et al. 2021). In many disease contexts, increased lactate production provides substrate for lactylation, whereas lactylation-dependent gene regulation or protein modification further reinforces pathological remodeling. However, most mechanistic studies have been conducted in cancer, immune, neurological, or fibrotic models. Evidence in muscle-related cells is more limited and remains dispersed across skeletal muscle, cardiac, and vascular studies. The following sections therefore focus on the regulatory roles of the lactate–lactylation axis in skeletal muscle cells, cardiomyocytes, and VSMCs.

The lactate-lactylation axis: a metabolic hub orchestrating muscle cell function

Skeletal muscle cell

Skeletal muscle is a major source and consumer of lactate during exercise (Westerblad et al. 2010). Its metabolic flexibility allows rapid switching between glycolytic and oxidative substrate use (Aoi and Tanimura 2021; Waldemer-Streyer et al. 2022; Hoffmann and Weigert 2017). This makes skeletal muscle an important tissue for studying whether lactate-derived lactylation connects exercise-induced metabolic changes with muscle adaptation.

The regulatory scope of lactylation in skeletal muscle cell extends beyond epigenetic modifications to encompass key signaling pathways and metabolic processes. For example, lactate activates the MEK/ERK signaling pathway by binding to GPR81 and promotes hypertrophy of C2C12 myotubes (Ohno et al. 2018). This anabolic effect is further supported by findings that the dietary antioxidant morin reduces high-fat diet-induced lactate production and protein lactylation, with the LDHA–lactate axis playing a central role in mitochondrial functional improvement (Wang 2025). In the context of metabolic disease, skeletal muscle cell lactylation has been directly linked to insulin resistance, which is a key pathological feature of Type 2 diabetes. Studies confirmed the presence of lactate-derived modifications in human skeletal muscle and their association with metabolic dysregulation (Maschari et al. 2022; Hong et al. 2017). Based on these metabolisms and signalings, lactylation is also related to sarcopenia and regenerative ability. During aging, glycolytic activity is progressively inhibited, leading to a gradual decline in histone lactylation. This decline further impairs DNA repair mechanisms and protein homeostasis, thereby accelerating muscle functional decline (Meng et al. 2025). Dai et al. showed that lactate-induced H3K9 lactylation at the Neu2 promoter, which upregulates Neu2 transcription and promoted myoblast differentiation. Neu2 encodes a cytosolic sialidase involved in desialylation of cell surface receptors required for myoblast fusion, linking metabolic lactate signaling to a specific differentiation program (Dai et al. 2023). This histone lactylation-mediated transcriptional mechanism contrasts with the non-histone enzymatic regulation of VPS34 (Jia et al. 2023), indicating that lactylation engages fundamentally different effector pathways depending on the target protein class. However, H3K9 is also subject to acetylation and methylation, both of which regulate myogenic genes. Whether H3K9la cooperates with or displaces these competing modifications at the Neu2 locus was not examined. Desgeorges et al. found that histone lactylation in infiltrating macrophages correlates with the transcriptional program of ischemia-induced skeletal muscle regeneration (Desgeorges et al. 2024). This finding suggests that lactylation may regulate muscle repair indirectly through macrophage reprogramming within the injury niche, rather than only through cell-autonomous effects in myofibers or myoblasts. Thus, lactylation may influence regeneration at the muscle cell and microenvironment levels. However, this study provided correlative rather than causal evidence. It remains unclear whether macrophage lactylation is required for regeneration, or whether the observed transcriptional changes merely reflect other lactate-dependent pathways.

In addition to these known mechanisms, emerging evidence revealed the functional interaction between lactylation and autophagy, which is the key process to maintain cell homeostasis. Jia et al. identified an enzymatic mechanism linking lactylation to autophagy. TIP60/KAT5 catalyzes VPS34 lactylation at K356 and K781. This modification enhances VPS34 lipid kinase activity, leading to increased PI3P production at the phagophore membrane and subsequent autophagosome nucleation (Jia et al. 2023). The authors validated causality by using lactylation-deficient VPS34²ᴷᴿ mutants, which abolished lactylation-induced autophagy activation without affecting basal autophagic flux, thus providing one of the few loss-of-function demonstrations in the muscle lactylation field. Given the established role of VPS34 in autophagy induction (Jaber and Zong 2013) and skeletal muscle atrophy (Yamada et al. 2012), this mechanism positions lactylation as a metabolic sensor that couples glycolytic status to proteostatic maintenance. However, VPS34 activity is regulated by AMPK-mediated phosphorylation and TRAF6-mediated ubiquitination. Whether lactylation acts independently of these modifications or requires their coordination was not addressed. Additionally, TIP60 acetylates multiple substrates involved in DNA damage response and chromatin remodeling; whether its lactylation activity is substrate-selective or broadly promiscuous in skeletal muscle cell remains an open question that affects the specificity of any TIP60-targeted intervention. Lactylation may also directly regulate mitochondrial function in skeletal muscle cell. Increased plasma lactate is an early marker of mitochondrial dysfunction (Sciaccaluga et al. 2022), and chronic lactate exposure increases reactive oxygen species (ROS) production while reducing coupling efficiency (San-Millan et al. 2022). Beyond its metabolic roles, lactate directly targets mitochondrial proteins. Mao et al. identified AARS2-dependent lactylation of PDHA1 (K336) and CPT2 (K457/458) in skeletal muscle mitochondria. These modifications inhibit the pyruvate dehydrogenase complex and fatty acid β-oxidation, respectively (Mao et al. 2024). Both modifications limit mitochondrial substrate oxidation under high lactate, forming a potential negative feedback loop: when glycolysis exceeds oxidative capacity, accumulated lactate triggers AARS2-mediated lactylation, throttling oxidative phosphorylation and possibly limiting excessive ROS. This is plausible because AARS2 uses lactate directly without lactoyl-CoA, enabling rapid modification within the mitochondrial matrix. However, whether this feedback operates under physiological conditions or only during pathological accumulation remains unclear. The lactate threshold required to activate this pathway in skeletal muscle cell is currently unknown.

Cardiomyocyte

Cardiomyocytes have high and continuous energy demands and can use lactate as an oxidative substrate, especially under increased workload or stress (Bertero and Maack 2018). Under pathological conditions, however, excessive glycolysis and lactate accumulation may alter protein lactylation and contribute to cardiac remodeling (Chen et al. 2024; Dai et al. 2020). Current studies suggest that cardiomyocyte lactylation acts through both histone-dependent transcriptional regulation and non-histone protein modification.

In cardiomyocytes, lactate accumulation induced-lactylation changes the spatial conformation of proteins and then regulates the maintenance of cardiac structure and function. Zhang et al. identified lactylation of α-MHC at K1897 as a modification that maintained α-MHC interaction with titin. The K1897R lactylation-deficient mutant exhibited impaired sarcomere organization and reduced contractility, establishing direct causality (Zhang et al. 2023). In failing hearts, α-MHC K1897la was significantly reduced. This structural mechanism is distinct from histone lactylation-driven transcriptional regulation seen in cardiac hypertrophy (Zhao et al. 2024), indicating that cardiac lactylation operates through the following pathways: chromatin remodeling and direct contractile protein modification. Whether these two arms are activated at different lactate thresholds or disease stages has not been investigated. Moreover, HSPA12A stabilizes HIF‑1α by recruiting SMURF1, preventing its ubiquitination during reperfusion. The stabilized HIF‑1α then activates glycolytic genes, which sustains lactate production and drives histone H3 lactylation at pro‑survival promoters (Yu et al. 2024). This positions lactylation as a downstream epigenetic executor within the HSPA12A-HIF-1α protective axis. However, the specific H3 lactylation sites and target genes were not identified, and HIF-1α drives H3K27 acetylation, making it difficult to attribute cardioprotection to lactylation specifically. Zhao et al. reported increased H3K18 lactylation in hypertrophic cardiomyocytes and attenuation of hypertrophic growth by p300 inhibitor C646 (Zhao et al. 2024). However, C646 inhibits the acetyltransferase and lactylation activities of p300 with comparable potency. Given that H3K18 acetylation drives hypertrophy-associated gene expression, the individual contribution of lactylation cannot be determined from pharmacological inhibition alone. This limitation extends to all studies relying on p300/CBP inhibitors as evidence for lactylation-specific function.

Lactylation offers a potential strategy for the treatment of cardiac diseases by influencing gene expression related to metabolic reprogramming, mitochondrial function, and metabolism. MI/R injury is related to metabolic reprogramming, which leads to the increase in lactate level (Bei et al. 2024). MI/R injury enhanced Serpina3k lactylation at K351, stabilizing the protein and conferring cardioprotection (Wang et al. 2025). A similar stabilization mechanism has been reported for Snail1 in cardiac EndMT (Fan et al. 2023). Notably, Serpina3k lactylation is protective and Snail1 lactylation is pathological, demonstrating that the same modification type produces opposing outcomes depending on the substrate protein. This duality complicates the global regulation of cardiac lactylation. She et al. showed that dexamethasone inhibited PDK4, reducing lactate and MDH2 K241 lactylation (She et al. 2024). K241la impaired MDH2 enzymatic activity in the TCA cycle, creating a feedforward loop where glycolysis-derived lactate further compromises mitochondrial oxidation. However, the causal hierarchy remains ambiguous. Whether MDH2 lactylation directly triggers ferroptosis or acts indirectly through worsening mitochondrial dysfunction requires lactylation-site mutant validation. Another study showed that X-ray irradiation induced P4HB K311 lactylation in cardiomyocytes, enhancing P4HB-PTGS2 interaction and promoting mitoROS-driven mitophagy that exacerbated radiation injury (Ouyang et al. 2024). Aloe emodin attenuated this by reducing P4HB lactylation, but whether this compound directly inhibits the writers or indirectly lowers lactate availability was not distinguished, which is an ambiguity common to studies using natural compounds as lactylation modulators. Recently, bioinformatics analysis identified BRD4 as a key lactylation‑related gene in heart failure (Li et al. 2025). The findings preliminarily confirmed that targeted lactylation modification has therapeutic potential for cardiac health.

Lactylation may also be involved in cardiac regeneration, a capacity that is transient in newborn mice and humans and declines rapidly after birth (Mahmoud et al. 2013). The metabolic state of cardiomyocytes changes during the development of the heart. Fetal heart mainly relies on glycolysis for energy supply, but after birth, it gradually turns to fatty acid oxidation to meet high energy demand (Lopaschuk et al. 2010; Alaynick et al. 2007). This metabolic transformation is closely related to the loss of heart regeneration ability, and metabolic reprogramming has been proved to be highly important for the proliferation of cardiomyocytes in neonatal hearts (Ma et al. 2024; Bae et al. 2021). Zhang et al. demonstrated that 5SM increased lactate and stimulated cardiomyocyte proliferation through AARS2-mediated lactylation of PDHA1 (K336) and CPT2 (K457/K458), suppressing oxidative phosphorylation and recapitulating the fetal glycolytic state (Zhang et al. 2024). This mechanism parallels the findings on skeletal muscle cell (Mao et al. 2024), suggesting a conserved mitochondrial lactylation program across striated muscles. Whether these modifications are reversible upon stimulus withdrawal is critical because persistent metabolic suppression could compromise the contractile function of regenerated cardiomyocytes. By conducting a comprehensive analysis of the Kla profile in the hearts of newborn mice (Du et al. 2022), researchers were able to systematically characterize their lactylation-related features, laying an important foundation for exploring the mechanisms of cardiac regeneration and cardiac diseases.

To sum up, lactylation modification of cardiomyocytes is a field worthy of attention. In-depth exploration of lactylation modification of cardiomyocytes not only helps understand its role in cardiac physiology and pathology but also may provide new targets and strategies for the diagnosis and treatment of cardiac diseases.

VSMC

VSMCs are the main components of vascular media, which are mainly responsible for regulating vascular tension, blood flow, and blood pressure (Salabei and Hill 2013; Sarkar et al. 2024). Unlike striated muscle cells, most of ATP in VSMCs comes from glycolysis, and only about 30% comes from mitochondrial oxidation (Tuo et al. 2022; Xu et al. 2021). A remarkable feature of VSMCs is its outstanding phenotypic plasticity (Lee et al. 2019). Under physiological conditions, VSMCs maintain a differentiated “contractile” phenotype, which is characterized by low proliferative activity. However, after vascular injury, they can transform into a dedifferentiated “synthetic” phenotype, which is characterized by increased proliferation. This phenotypic transformation drives pathological vascular remodeling, which leads to diseases such as atherosclerosis and hypertension (Bennett et al. 2016; Park et al. 2021; Shi et al. 2020). The enhancement of the proliferation ability of synthetic VSMCs leads to increased energy demand, and the enhancement of glycolysis metabolism meets this demand (Park et al. 2021; Werle et al. 2005). This metabolic adaptation occurs in a microenvironment rich in lactate, and the accumulation of lactate further promotes the formation of synthetic phenotype by enhancing cell proliferation, migration, and the expression levels of synthetic markers, partly by regulating MCT (Yang et al. 2017). LDHA is upregulated in aortic dissection (AD) tissues. Its inhibitor, oxamate, reverses VSMC dedifferentiation and reduces AD incidence (Wu et al. 2022). While these findings established lactate as a phenotypic switching driver, the mediating role of lactylation remains undefined. VSMC identity is canonically regulated by myocardin/SRF and KLF4. Whether lactylation modifies these regulators directly or alters their promoter accessibility has not been investigated.

The complex relationship between lactylation and the function of VSMC has been proposed, and it is related to cell aging, atherosclerosis, vascular remodeling, and vascular calcification. High glucose increased lactate in VSMCs, which promotes H3K18 lactylation at the CHI3L1 promoter and activating IL-13Rα2/JAK1/STAT3-dependent osteogenic transdifferentiation (Zhu et al. 2025). However, H3K18 is also subject to acetylation. Whether H3K18la displaces H3K18ac under hyperglycemia or both marks coexist with additive effects was not examined. This distinction is important: if lactylation replaces acetylation, the transcriptional outcome depends on their relative activating potencies at this specific locus. During the aging process of VSMC, lactylation affects the expression levels of key genes involved in cell aging and inflammation. Li et al. revealed that TRAP1 impairs mitochondrial respiration and increases lactate in VSMCs. H4K12 lactylation required prior HDAC3-mediated deacetylation at the same residue, establishing an ordered two-step process. VSMC-specific TRAP1 knockout reduced plaque area and SASP markers in vivo (Li et al. 2024). This sequential model implies that HDAC inhibitors could paradoxically reduce lactylation by blocking the prerequisite deacetylation. Whether this logic extends beyond H4K12 remains untested. In pulmonary hypertension, Chen et al. showed that mROS-mediated HIF-1α stabilization drove glycolytic shift in PASMCs, promoting histone lactylation and proliferative remodeling (Chen et al. 2023). However, the specific lactylation sites and target genes were not characterized. HIF-1α simultaneously promotes acetylation, making it difficult to attribute the proliferative phenotype to lactylation alone, a methodological challenge shared with the cardiac hypertrophy findings (Zhao et al. 2024). Tumor necrosis factor α-induced SOX10 lactylation activates the PI3K/AKT signaling pathway in a phosphorylation-dependent manner, which drives the transcription program of transdifferentiation of VSMC and eventually leads to cell pyroptosis (Xu et al. 2023). This crosstalk suggests that lactylation can amplify inflammatory signaling by priming proteins for subsequent activating modifications, though whether these modifications occur at proximal sites enabling direct conformational coupling was not examined. These studies emphasized the potential of targeted lactylation as a therapeutic strategy in relieving vascular diseases characterized by VSMC dysfunction.

Nuclear receptor subfamily 4 group A member 3 (NR4A3), an orphan nuclear receptor, critically mediates histone lactylation in VSMCs, particularly during vascular media calcification. NR4A3 enhances glycolytic activity, leading to an increase in lactate production, which, in turn, promotes histone lactylation and eventually leads to vascular calcification (Mao et al. 2024). Thus, lactylation not only regulates gene expression but also participates in metabolic reprogramming of VSMCs, linking metabolic changes to epigenetic modifications. Li et al. showed that March2 loss maintained PKM2 in its dimeric form, which has reduced kinase activity but gains nuclear translocation capacity as a transcriptional co-activator. This metabolic reprogramming increased H3K18 lactylation at the p53 promoter, activating pro-apoptotic transcription (Li et al. 2025). However, nuclear PKM2 dimers also promote H3 phosphorylation and acetylation, and the specific contribution of lactylation was not isolated. Recent studies have revealed that ischemic/hypoxic injury induces VSMC damage through lactate–lactylation–mitochondrion cascade. Specifically, hypoxia promotes lactate accumulation and H3K18Ia modification and upregulates the expression of arginase 1 (Arg1), thus destroying the structure of mitochondrial crista and damaging the contact site of the mitochondria and the tissue of protein of subunit 10 (Mic10) in crista tissue system. Meanwhile, lactylation of VDAC1 at K224 synergistically promotes mitochondrial DNA release. This activates the cGAS‑STING pathway and triggers cell apoptosis (She et al. 2025). This process establishes a positive feedback loop between lactate production and mitochondrial damage, which eventually aggravates vascular dysfunction.

In a word, lactylation is an important regulatory mechanism in VSMCs, affecting their function, aging process, and response to pathological stimuli. A comprehensive understanding of the molecular pathway and mechanism of lactylation in blood vessels may be helpful to develop new therapeutic strategies for treating vascular diseases and improving vascular health. A summary of lactylation mechanisms across the three muscle cell types (skeletal muscle cells, cardiomyocytes, and VSMCs) is presented in Fig. 5 and Table 1, which highlight cell-specific lactylation targets and their associated functional outcomes in physiology and disease.

Fig. 5.

Mechanism of lactylation in muscle cells

This figure summarizes the current understanding of lactylation in skeletal muscle cells, cardiomyocytes, and VSMCs. In skeletal muscle cells, lactylation regulates muscle homeostasis, regeneration, and autophagy. Key examples include VPS34 lactylation (K356/K781) promoting autophagy flux, and histone H3K9la activating Neu2 expression to promote myogenesis. In cardiomyocytes, lactylation is involved in myocardial I/R injury, cardiac protection, and heart failure. For instance, H3K18la is elevated in pathological cardiac hypertrophy, while α-MHC lactylation (K1897) preserves sarcomeric structure. In VSMCs, lactylation is linked to pathological processes including arterial calcification (H3K18la upregulating CHI3L1), apoptosis, proliferation, and inflammation. Each cell type is shown with its representative lactylation targets and associated functional outcomes

Table 1.

Summary of studies on lactylation in muscle cells

Organ/Tissue	Cell type	Species	Disease/Pathological condition	Lactylation site	Effects	Reference
Skeletal muscle	Skeletal muscle cell	Human	Insulin resistance	Pan Kla	Positive associated with insulin resistance	(Maschari et al. 2022)
Skeletal muscle	Skeletal muscle cell	Mice	NR	VPS34 K356 and K781	Induce autophagy	(Jia et al. 2023)
Skeletal muscle	Skeletal muscle cell	Human Mice	NR	H3K18la	A marker for tissue-specific active enhancers	(Galle et al. 2022)
Skeletal muscle	Skeletal muscle cell C2C12	Mice	NR	PDHA1 K336 CPT2 K457/8	Catalyzed by AARS2	(Mao et al. 2024)
Skeletal muscle	Myoblast	Mice	Skeletal muscles injury	H3K9la	Promote myoblast differentiation and muscle regeneration	(Dai et al. 2023)
Skeletal muscle	Soleus muscle cell	Mice	NR	Pan Kla	Kla levels peak at 24 h after HIIT	(Huang et al. 2023)
Skeletal muscle	Fibroblasts; Endothelial cells	Mice	Senescence	H3K9la H3K14la H3K18la	Upregulate key genes in the DNA repair and proteostasis pathways	(Meng et al. 2025)
Heart	Cardiomyocyte	Mice	Myocardial I/R	Serpina3k K351La	Protect cardiomyocytes from reperfusion-induced apoptosis	(Wang et al. 2025)
Heart	Cardiomyocyte	Mice	Pathological cardiac hypertrophy	H3K18la	Induce cardiac hypertrophy	(Zhao et al. 2024)
Heart	Cardiomyocyte	Mice	NR	ACADL HADHA ACAT1 ENO3 ALDOA PKM	Offer a comprehensive Kla map in the neonatal mouse heart	(Zhang et al. 2024)
Heart	Cardiomyocyte	Mice	Myocardial I/R	H3K56la	Promote cardiomyocyte survival	(Yu et al. 2024)
Heart	Cardiomyocyte	Mice	Heart failure	α-MHC K1897	Preserve sarcomeric structure and function	(Zhang et al. 2023)
Heart	Cardiomyocyte H9c2	Rat	Myocardial I/R	MDH2 K241	Promote myocardial injury and mitochondrial damage	(She et al. 2024)
NR	Cardiomyocyte H9c2	Rat	ROS accumulation	P4HB K311	A crucial pathophysiological process to regulate RIHD	(Ouyang et al. 2024)
Heart	Cardiomyocyte	Mice	NR	PDHA1 CPT2	Facilitate cardiomyocytes proliferation	(Du et al. 2022)
Aorta	VSMC	Mice	Senescent VSMC model Atherosclerosis	H4K12la	Promote cell senescence and atherosclerosis	(Li et al. 2024)
Aorta	VSMC	Mice	Arterial calcification	H3K18la	Promote vascular calcification	(Mao et al. 2024; Zhu et al. 2025)
Aorta	VSMC	Mice	Chronic inflammation	SOX10	Promote cell transdifferentiation	(Xu et al. 2023)
Aorta	VSMC	Mice	Aortic aneurysm/dissection	H3K18la	Promote cell apoptosis	(Li et al. 2025)
Mesenteric artery	VSMC	Mice	Ischemia/hypoxia injury	VDAC1la H3K18la	Promote mitochondrial cristae remodeling and PANoptosis	(She et al. 2025)
Pulmonary artery	Pulmonary artery smooth muscle cell	Mice	Pulmonary hypertension	H3K18la H4K5la	Promote cell proliferation	(Chen et al. 2023)

Abbreviations: NR not reported; other abbreviations can be found in the attached table “Abbreviation”.

Therefore, the interaction between lactylation and other PTMs must be studied in the future to reveal the complex regulatory network that regulates muscle adaptation. Deciphering these networks not only helps clarify the mechanism of muscle fatigue and recovery but also reveals a series of new pathogenic drivers and therapeutic targets of musculoskeletal diseases (Table 1 and Fig. 5).

Exercise-induced modulation of lactate signaling and lactylation in muscle cells

Exercise induces extensive metabolic remodeling and epigenetic regulation in muscle tissues. Lactylation has been proposed as a potential mediator linking lactate metabolism to transcriptional and functional adaptation during exercise. However, the current evidence remains preliminary. The extent and direction of exercise-associated lactylation appear to differ in accordance with exercise type, intensity, and duration and may depend on species and tissue specificity. Available studies focus mainly on skeletal muscle cells, cardiomyocytes, and VSMCs. These studies distinguish acute from chronic exercise, human from animal models, and different exercise modalities (Fig. 6 and Table 2).

Fig. 6.

Exercise induced lactylation modulating responses in muscle cells

This figure illustrates how exercise and its mimetics (exogenous lactate or hypoxia) modulate lactylation-dependent pathways across multiple cell types. In cardiomyocytes, chronic swimming training reduces H3K18la levels and suppresses YTHDF2 expression, thereby alleviating myocardial ischemia-reperfusion injury. In skeletal muscle cells, acute exhaustive exercise promotes VPS34 lactylation (via KAT5/TIP60), enhancing autophagy flux; moderate training alters the lactylome of mitochondrial ATP synthase subunits; overtraining induces SORBS3 lactylation, leading to formation of lactate bodies that cause liver fibrosis. In VSMCs, lactate potentially drives phenotypic switching via PI3K-AKT signaling, although direct evidence for lactylation in this context remains unclear. The figure distinguishes between acute and chronic exercise, moderate and overtraining, and highlights established versus proposed mechanisms

Table 2.

Physical exercise mediates lactylation modification in muscle cells

Cell type	Subjects	Exercise intervention	Lactylation and protein expression responses	Ref.
Skeletal muscle cell	8-week-old male mice	Treadmill training: Moderate Training: 15 m/min, 60 min/day, 5 days/week (12 weeks) Overtraining (Progressive): Phase I (Weeks 1–4): Speed 15→22.5 m/min, duration 20→60 min Phase II (Weeks 5–8): Fixed at 25 m/min, 60 min/session Phase III (Weeks 9–12): Sessions 1→4/day (25 m/min, 60 min each)	Muscle lactate accumulation→SORBS3 lactylation→LLPS→FBXO2 sorting into lactate bodies→hepatocyte apoptosis→hepatic stellate cell activation→liver fibrosis	(Liu et al. 2025)
	Young and healthy participants(18-40 years; 18.5≤BMI≤23.9 kg/m2)	Cycling training: ①Contol group: PRE: ≥7 (>80% of sessions); ACWR Range:>1.3(7weeks), >1.5(5weeks) ②Overtraining group: PRE:<80% sessions at RPE≥7; ACWR Range:>1.3(7weeks), >1.5(5weeks)
	Male B57BL/6 mice	Acute swimming exercise: 3h (Immediate access)	Exercise-induced Vps34 lactylation regulates muscle autophagy and correlates with cancer progression.	(Jia et al. 2023)
	Male B57BL/6 mice	Treadmill high-intensity fatigue exercise: Speed increased 5 m/min/5min to 20 m/min, then incline rose 5%/5min to exhaustion	Hypoxia-induced AARS2 accumulation promotes PDHA1/CPT2 lactylation, inhibiting Ac-CoA production and forming an OXPHOS-limiting negative feedback loop post-exercise.	(Mao et al. 2024)
	8-week-old male mice	Moderate intensity treadmill training: Run at 6 m/min initially, increased to target speed, 60 min/day, 5 days/week for 6 weeks; each mouse runs 22 km	Moderate exercise altered 159 lactylation sites on 78 proteins. Atp5mg and Atp5po lactylation enhanced ATP hydrolysis, while Mtatp8, Atp5mg, and Atp5po showed decreased lactylation but increased protein levels.	(Chang et al. 2024)
	Young and healthy participants (25±4 years old, 24.0±3.8 kg/m2)	Acute and chronic resistance training: 4×9-12RM unilateral leg extensions, 90s rest, 2-3×/week (24 sessions/10 weeks) Acute Endurance Training Study: 60-min cycling at 70% HRR (avg HR 161±9 bpm)	Exercise induces more dynamic changes in skeletal muscle protein acetylation, but not lactylation.	(Mattingly et al. 2024)
	College-aged men	Resistance training: Barbell bench press, pronated grip barbell row, barbell stiff-leg deadlift, 3 days/weeks for 6 weeks	Acute resistance exercise raised lactate 7.2-fold but not lactylation, while chronic training caused hypertrophy without lactylation changes.	(Mattingly et al. 2023)
Cardiomyocyte	8-week-old male mice	Swimming exercise: increased from 10min twice daily by 10 min daily until reaching 90min/day over 4 weeks	Exercise training mitigates cardiac ischemia-reperfusion injury and remodeling by suppressing lactylation and YTHDF2, offering cardioprotection.	(Xu et al. 2024)

Abbreviations: ACWR acute: chronic workload ratio, PRE Perceived rate of exertion, SORBS3 SH3 domain-containing 3, LLPS liquid-liquid phase separation, FBXO2 F-box protein 2

Lactylation of skeletal muscle cell in exercise

Most of the current evidence for exercise-associated lactylation in skeletal muscle comes from rodent studies, whereas human data remain limited. Existing findings suggest that the extent and direction of lactylation changes depend on the exercise protocol, tissue sampling time, and the specific protein or site examined.

In murine skeletal muscle, lactylation responds to both chronic training and acute exercise, although the affected targets differ by exercise paradigm. Chronic moderate-intensity treadmill training remodels the skeletal muscle lactylome, with enrichment of changes in mitochondrial proteins. In one study, 6 weeks of treadmill running altered 159 lactylation sites across 78 proteins, including reduced lactylation of the ATP synthase subunits Mtatp8, Atp5mg, and Atp5po, accompanied by increased protein expression (Chang et al. 2024). These findings raise the possibility that chronic exercise reorganizes mitochondrial lactylation in a manner favorable to oxidative metabolism. Acute exhaustive exercise appears to induce a different pattern, characterized by stress-responsive and site-specific modifications. Mao et al. showed that exhaustive treadmill running promoted mitochondrial AARS2-dependent lactylation of PDHA1 (K336) and CPT2 (K457/K458), with consequent suppression of oxidative phosphorylation under hypoxic conditions. This effect may reflect a transient adaptive mechanism during severe metabolic stress rather than a stable training response (Mao et al. 2024). In addition to mitochondrial regulation, lactylation has been implicated in exercise-induced autophagy. Li et al. identified lactylation of mTOR at K2066 as a direct exercise-responsive event in skeletal muscle and linked this modification to enhanced autophagic signaling (Li et al. 2025). Jia et al. found that prolonged swimming induced KAT5/TIP60-dependent lactylation of VPS34 at K356 and K781, thereby facilitating autophagic flux; mutation of these sites impaired the autophagic response (Jia et al. 2023). Taken together, these studies suggest that exercise-associated lactylation in skeletal muscle is context-dependent and may participate in mitochondrial and autophagic adaptation.

The relationship between exercise load and lactylation is unlikely to be linear. Evidence from rodent studies suggests that moderate training, exhaustive exercise, and overtraining may engage distinct lactylation programs with different physiological consequences. Using a progressive overtraining model, Liu et al. found that 12 weeks of excessive treadmill running (progressing to four 60‑min sessions per day at 25 m/min) caused sustained lactate accumulation and induced SORBS3 lactylation in skeletal muscle. This modification promoted liquid–liquid phase separation and the formation of “lactate bodies.” These structures facilitated FBXO2 transport to the liver and were associated with hepatocyte apoptosis and liver fibrosis (Liu et al. 2025). Notably, this response was absent in animals subjected to moderate training, indicating that the downstream effects of lactylation may shift from adaptive to maladaptive under conditions of excessive exercise stress.

Exercise may also modulate lactylation in the setting of skeletal muscle aging. Meng et al. reported that 2 months of treadmill training in aged mice restored histone lactylation marks, including H3K9la, H3K14la, and H3K18la, at the promoter regions of genes involved in DNA repair and proteostasis. These changes were accompanied by improved muscle function and attenuation of age-related decline (Meng et al. 2025). The same study further suggested that aging skeletal muscle is characterized by reduced glycolytic activity and loss of histone lactylation, whereas exercise partially reverses these alterations (Meng et al. 2025). Together, these findings support a link between exercise-driven metabolic remodeling and maintenance of epigenetic regulation during aging.

Human studies, however, have not yet provided comparable evidence for exercise-induced lactylation in skeletal muscle. Mattingly et al. reported that acute resistance exercise (four sets of unilateral leg extensions at 9–12 repetition maximum) increased circulating lactate by about 7.2‑fold in young healthy adults, but did not alter skeletal muscle protein lactylation. Likewise, neither 10 weeks of resistance training nor a single 60 min cycling bout at 70% heart rate reserve produced detectable changes in muscle lactylation (Mattingly et al. 2024). Similar results were reported by Mattingly et al., who observed muscle hypertrophy after 6 weeks of resistance training in college-aged men, without measurable changes in lactylation (Mattingly et al. 2023).

In human studies, muscle biopsies are often taken more than 24 h after the last exercise bout, potentially missing transient lactylation responses that occur during exercise or early recovery. In contrast, rodent studies commonly analyze tissues immediately after exercise. Moreover, resistance exercise produces short‑lived, intermittent lactate increases, which may be less effective at driving detectable lactylation than the sustained lactate exposure seen in some animal protocols. Analytical approaches also differ: most human studies rely on pan-Kla immunodetection, whereas rodent studies have more often used mass spectrometry-based lactylomics with greater site specificity and sensitivity. Species differences in lactate metabolism and modification dynamics may contribute to the discrepancy. Thus, skeletal muscle lactylation is responsive to exercise in rodent models (exhaustive exercise, chronic training, aging, and overtraining), but human studies have not consistently detected such changes. Whether this difference reflects biological differences, sampling windows, or analytical limitations remains unresolved.

Cardiomyocyte lactylation in exercise

Evidence for exercise-associated lactylation in the heart is more limited than that in skeletal muscle. However, the available studies provide initial mechanistic clues as to how lactylation may participate in exercise-induced cardioprotection.

Xu et al. examined chronic swimming training in mice using a 4‑week protocol that increased from 10 min twice daily to 90 min per day. This intervention reduced myocardial lactate levels and decreased H3K18 lactylation (Xu et al. 2024). Mechanistically, reduced H3K18la suppressed YTHDF2 expression and subsequently inhibited G3BP1-mediated hypertrophic and apoptotic signaling. This alleviated myocardial ischemia–reperfusion injury. This study suggests that chronic endurance exercise can remodel cardiac histone lactylation and that such remodeling may contribute to myocardial stress resistance (Xu et al. 2024). The observation is consistent with reports showing that pathological cardiac hypertrophy is associated with increased H3K18la, implying that exercise may exert beneficial effects not simply by increasing lactylation but by restoring an appropriate lactylation state in a disease- and tissue-specific manner.

A recent work further indicated that exercise-associated lactylation in the heart may not be restricted to cardiomyocytes. Sun et al. reported that exercise induced histone lactylation in monocyte-derived cardiac macrophages. This improved cardiac immune homeostasis in a mouse model of sepsis-induced cardiomyopathy. In that study, exercise was associated with the emergence of iNOS⁺Arg1⁺ monocyte-derived cardiac macrophages displaying pro-inflammatory and pro-reparative features. Lactylation was proposed to contribute to this immune reprogramming (Sun et al. 2025). These data extend the scope of exercise-induced lactylation to non-myocyte cardiac cell populations and point to a potential role in shaping the cardiac immune microenvironment.

However, the cardiac literature remains at an early stage. Direct evidence in human myocardium is lacking. Moreover, it is unknown to what extent findings from murine swimming or disease models translate to human exercise physiology. Moreover, few studies have systematically compared different exercise modalities, intensities, or sampling windows in cardiac tissue. Thus, although rodent studies support a role for lactylation in exercise-induced cardiac adaptation, its relevance to human cardioprotection remains to be established.

VSMCs lactylation in exercise

Evidence for exercise-induced lactylation in VSMCs is currently sparse. To date, no study has directly tested whether acute or chronic exercise alters lactylation in VSMCs at defined histone or non-histone sites. As a result, any discussion of VSMC lactylation in the context of exercise remains largely inferential.

However, some indirect evidence makes this question biologically plausible. Lactate influences VSMC phenotype. Yang et al. reported that lactate promotes a synthetic VSMC phenotype characterized by increased proliferation and migration, partly through regulation of MCTs (Yang et al. 2017). Exercise transiently or chronically alters circulating lactate depending on intensity and training status. Therefore, exercise‑derived lactate may affect VSMC phenotype, but whether this involves lactylation‑dependent transcriptional regulation remains unknown. Indirect support comes from studies in other vascular cell types. Chen et al. demonstrated that MeCP2 K271 lactylation-mediated M2 macrophage polarization can inhibit atherosclerosis (Chen et al. 2024). Wang et al. further reported that exercise-induced endothelial MeCP2 lactylation suppresses atherosclerosis through the Ereg/MAPK signaling pathway (Wang et al. 2023). Although these studies do not directly address VSMCs, they indicate that lactylation can participate in vascular remodeling and may be responsive to exercise in vascular-associated cells.

The absence of direct VSMC data represents an important gap. Future studies should examine whether exercise modifies VSMC lactylation at histone marks, such as H3K18la or H4K12la, and non-histone sites implicated in calcification, senescence, and phenotypic switching. It is important to determine whether lactylation promotes contractile maintenance, maladaptive remodeling, or context‑dependent transitions between these states. Such work could help clarify whether lactylation is merely associated with vascular metabolic changes or serves as an active regulator of exercise-induced vascular adaptation.

Translational potential and therapeutic opportunities

Lactylation in skeletal muscles, cardiomyocytes, and VSMCs is now linked to metabolic disease, positioning the lactylation pathway as a potential therapeutic target. Three intervention strategies have been explored, as summarized in Table 3: modulating lactate supply, targeting lactylation enzymes, and using exercise as a physiological regulator. Nearly all current evidence is preclinical.

Table 3.

Translational strategies targeting the lactate-lactylation axis

Target/Intervention	Agent/Modality	Muscle cell type	Disease/Pathological condition	Effect on lactylation	Mechanism	Functional outcome	Reference
Lactate metabolism
LDHA	Oxamate	VSMC	Aortic dissection	↓ Pan-Kla	Inhibits LDHA; reduces lactate production	Reverses VSMC phenotype switching	(Wu et al. 2022)
LDHA	Oxamate	Skeletal muscle	Insulin resistance	↓ Pan-Kla	Inhibits LDHA	Improves insulin sensitivity	(Maschari et al. 2022)
LDHA	FX-11	Cardiomyocyte	Myocardial infarction	↓ Pan-Kla	LDHA inhibition	Cardioprotection	(Wang et al. 2022)
LDHA	GSK2837808A	Skeletal muscle	Embryonic development	↓ Pan-Kla	LDHA inhibition	Developmental effects	(Yang et al. 2021)
Lactate transport
MCT1/4	CHC	Cardiomyocyte	Myocardial infarction	↓ Pan-Kla	MCT inhibitor	Cardioprotection	(Fan et al. 2023)
MCT1	AZD3965	Cardiomyocyte	Myocardial infarction	↓ Pan-Kla (inferred)	Selective MCT1 inhibition	Cardioprotection	(Wang et al. 2022)
MCT1	7ACC2	Immune cells (NKT)	Malignant pleural effusion	↓ Histone lactylation	MCT1 inhibitor	Reduces FOXP3 expression	(Wang et al. 2023)
Lactylation (Writers)
p300/CBP	C646	Cardiomyocyte	Cardiac hypertrophy	↓ H3K18la	p300/CBP inhibitor	Attenuates pathological cardiac hypertrophy	(Zhao et al. 2024; Shi et al. 2025)
p300/CBP	C646	Endothelial cell	Sepsis	↓ H3K18la	p300/CBP inhibitor	Reduces endothelial permeability; improves survival	(Yang et al. 2022)
p300/CBP	A-485	Neuronal cell	Ischemic stroke	↓ H3K18la	p300/CBP inhibitor	Reduces brain injury; suppresses inflammation	(Huang et al. 2024; Xiong et al. 2024)
TIP60/KAT5	MG149	Skeletal muscle	Autophagy regulation	↓ VPS34 lactylation	TIP60 acetyltransferase inhibitor	Suppresses autophagy	(Jia et al. 2023)
Lactylation (Erasers)
HDAC1/3	MS-275	Neuronal cell	Neural development	↑ H4K12la	Class I HDAC inhibitor	Modulates gene expression	(Dai et al. 2022)
HDACs (pan)	TSA	Colorectal cell	Colorectal cancer	↑ Pan-Kla	Pan-HDAC inhibitor	Alters gene expression	(Xiong 2022)
SIRT3	Honokiol	Hepatocyte	Hepatocellular carcinoma	↓ CCNE2-K348la	SIRT3 activator	Induces apoptosis	(Jin et al. 2023)
Lactylation upstream modulators (Glycolysis)
Hexokinase	2-DG	Skeletal muscle	Insulin resistance	↓ Pan-Kla	Inhibits glycolysis	Improves insulin sensitivity	(Maschari et al. 2022)
Hexokinase	2-DG	Neuronal cell	Cerebral infarction	↓ Pan-Kla	Inhibits glycolysis	Reduces infarct volume	(Zhang et al. 2023)
PDH	Rotenone	Cardiomyocyte	Myocardial infarction	↑ Pan-Kla	Complex I inhibitor	Context-dependent effects	(Wang et al. 2022)
PDK	DCA	Cardiomyocyte	Myocardial infarction	↓ Pan-Kla	PDK inhibitor	Cardioprotective	(Wang et al. 2022)
PDK	DCA	Immune cell	Inflammation	↓ Pan-Kla	PDK inhibitor	Reduces lactate; anti-inflammatory	(Lopez Krol et al. 2022)
Lactylation physiological modulators (Exercise)
moderate exercise	Swimming (4 weeks)	Cardiomyocyte	Cardiac I/R injury	↓ H3K18la	Reduces myocardial lactate; suppresses YTHDF2	Cardioprotection	(Xu et al. 2024)
moderate exercise	Treadmill (2 months)	Skeletal muscle cell	Muscle aging	↑ H3K9/14/18la	Restores histone lactylation at DNA repair genes	Reverses age-related decline	(Meng et al. 2025)
Acute exercise	Swimming (3 hours)	Skeletal muscle cell	Exercise stress	↑ VPS34 K356/K781la	KAT5/TIP60-mediated lactylation	Promotes autophagy flux	(Jia et al. 2023)
Overtraining	Progressive treadmill (12 weeks)	Skeletal muscle cell	Overtraining syndrome	↑ SORBS3 lactylation	LLPS → lactate body formation	Liver fibrosis (inter-organ)	(Liu et al. 2025)

Abbreviations: AARS1 Alanyl-tRNA synthetase 1, BSO Buthionine sulfoximine, CBP CREB-binding protein, CCNE2 Cyclin E2, CD147 Cluster of differentiation 147 (Basigin), CFZ Carfilzomib, CHC α-Cyano-4-hydroxycinnamate, DCA Dichloroacetate, FGS Fasentin + Glutamine + Selenium, FOXP3 Forkhead box P3, GCLC Glutamate-cysteine ligase catalytic subunit, HK Hexokinase, LLPS Liquid-liquid phase separation, MCT Monocarboxylate transporter, MVA Mevalonate, NKT Natural killer T cell, PD-1 Programmed cell death protein 1, PDH Pyruvate dehydrogenase, PDK Pyruvate dehydrogenase kinase, PD-L1 Programmed death-ligand 1, PKM2 Pyruvate kinase M2, ROS Reactive oxygen species, SAHA Suberoylanilide hydroxamic acid (Vorinostat), SIRT3 Sirtuin 3, SORBS3 Sorbin and SH3 domain containing 3, TCA Tricarboxylic acid, TIP60 Tat-interacting protein 60, TSA Trichostatin A, TTM Tetrathiomolybdate, VB124 MCT4 inhibitor

Targeting lactate metabolism and transport

Lactylation uses lactyl-CoA as its direct substrate, and its abundance is driven by intracellular lactate concentrations. Thus, pharmacological targeting of lactate production or transport can indirectly regulate lactylation at its source.

LDHA is the rate-limiting enzyme for lactate production. Oxamate reversed VSMC synthetic phenotype switching in a murine AD model (Wu et al. 2022)and improved skeletal muscle insulin sensitivity in insulin-resistant mice (Maschari et al. 2022). FX-11 reduced myocardial lactate and protected against infarction injury (Wang et al. 2022). These agents remain at proof-of-concept stage for muscle diseases.

In the lactate transport process, MCT1 and MCT4 shuttle lactate across membranes. CHC and AZD3965 protect the heart in infarction models by limiting lactate accumulation in cardiomyocytes (Fan et al. 2023; Wang et al. 2022). Meanwhile, 7ACC2 reduced histone lactylation in NKT cells and reversed immunosuppression in malignant pleural effusion (Wang et al. 2023). Lactate drives VSMC synthetic switching via MCT-dependent uptake (Yang et al. 2017), but MCT inhibition has not been directly tested in vascular remodeling.

Systemic lactate inhibition may disrupt energy supply to lactate-dependent organs, including the brain and heart. Tissue-selective delivery or intermittent dosing may improve the therapeutic window.

Targeting lactylation enzymes

Directly targeting the enzymes that catalyze lactylation can regulate site-specific lactylation more precisely than metabolic-level interventions. However, this approach also faces the technical challenge of insufficient modification selectivity.

p300 and CBP are the most extensively studied lactylation writers. They transfer the lactoyl group from lactoyl-CoA to lysine residues. Moreover, p300/CBP inhibitors have shown therapeutic potential in various disease models. For example, C646 reduces cardiomyocyte hypertrophy (Zhao et al. 2024) and suppresses HMGB1 lactylation in sepsis, thereby lowering endothelial permeability and improving survival (Yang et al. 2022). A-485 attenuated ischemic brain injury (Huang et al. 2024; Xiong et al. 2024). The core limitation is that p300/CBP also catalyzes acetylation through the same active site, and no existing inhibitor distinguishes between the two modifications.

With regard to the erasers of lactylation, class I HDACs function as delactylases (Zhang et al. 2023; Dai et al. 2022). A similar selectivity issue exists: HDAC inhibitors target both acetylation and lactylation. Kac equilibrates within ~6 h, whereas Kla accumulates over 24 h (Zhang et al. 2019), pointing to distinct kinetics that may create temporal competition at shared sites. SIRT3 is a mitochondrial delactylase. Honokiol, a SIRT3 activator, reduced CCNE2 K348 lactylation in hepatocellular carcinoma (Jin et al. 2023). SIRT3 expression correlates with skeletal muscle lactylation during aging (Meng et al. 2025), but whether SIRT3 activation restores muscle function in aged animals has not been tested. In addition, the TIP60/KAT5 inhibitor MG149 reduced VPS34 lactylation and suppressed autophagy (Jia et al. 2023). The AARS1/2 inhibitor β-alanine showed antitumor activity (Zong et al. 2024), but it is untested in muscle.

Exercise as a physiological modulator

Exercise regulates lactylation through endogenous lactate production, offering noninvasive benefits. However, clinical application requires clarification of dosage and intervention protocols.

Current evidence suggests that moderate-intensity exercise exerts a protective effect through lactylation: four weeks of swimming training reduced myocardial H3K18la and alleviated I/R injury (Xu et al. 2024), and 2 months of treadmill exercise restored H3K9/14/18la in aged skeletal muscle and reversed functional decline (Meng et al. 2025). Overtraining, however, induces SORBS3 lactylation, triggering phase separation and liver fibrosis (Liu et al. 2025). Inactivity is equally detrimental—decreased histone lactylation in skeletal muscle leads to downregulation of DNA repair pathways (Meng et al. 2025).

Exercise mode influences lactylation effects. Endurance exercise produces sustained lactate increase, which more readily drives lactylation modifications. In contrast, resistance exercise generates intermittent lactate peaks, and human studies have not detected sustained lactylation changes (Mattingly et al. 2024). Acute exercise induces transient, site-specific lactylation accumulation such as VPS34 K356/K781la involvement in autophagy activation (Jia et al. 2023). Chronic training, however, reshapes the baseline lactylation profile, involving mitochondrial functional reprogramming (Chang et al. 2024).

Limitations remain at the translational level. While rodent data are relatively consistent, human studies have not yet reliably detected post-exercise lactylation changes (Mattingly et al. 2024, 2023). Possible reasons include delayed biopsy timing (> 24 h, missing the peak window) and insufficient sensitivity of pan-Kla antibodies. Based on animal evidence, moderate-intensity aerobic exercise (3–5 times/week, 30–60 min) can serve as an initial regimen, whereas prolonged excessive training should be avoided. Future studies should establish the time-dependent dynamics of post-exercise lactylation accumulation in humans. They should also employ targeted mass spectrometry for detection.

Lactylation in clinical translation

The lactate–lactylation axis is increasingly showing potential for clinical translation. Combined with bioinformatics and multi-omics analyses, lactylation levels may reflect disease-related metabolic states and serve as candidate biomarkers for early diagnosis and disease progression monitoring in cancer, metabolic disorders, and inflammatory diseases. With further clinical validation, Lactylation may also help link exercise load with disease risk (Certo et al. 2022). At present, several drugs targeting lactate production or transport already have translational relevance. For example, the MCT1 inhibitor AZD3965 has entered phase I clinical trials and has shown good safety and tolerability, suggesting the clinical feasibility of regulating transmembrane lactate transport (Halford et al. 2023). The HDAC inhibitor MS-275 has also entered phase III clinical trials and has shown antitumor activity (Ryan et al. 2005). In the cardiovascular system, AZD3965 and the MCT4 inhibitor VB124 may improve cardiomyopathy, myocardial infarction, and heart failure by modulating lactate metabolism. By suppressing lactate production, the LDHA inhibitors FX11 and oxamate have also shown therapeutic potential in myocardial injury models (Chen et al. 2023). In addition, sodium lactate has been approved by the FDA, and DCA has entered phase II clinical trials, further supporting the clinical feasibility of targeting lactate metabolism.

Direct targeting of lactylation itself is also emerging as a potential strategy. For example, hemin can enhance METTL3 lactylation and induce ferroptosis through m6A regulation, whereas curcumol can inhibit H3K18 lactylation (Zhang et al. 2023). These findings hint that specific lactylation sites could be druggable. However, these studies remain largely mechanistic, and direct evidence in exercise-induced adaptation or muscle function is still lacking. MCT inhibitors may also cause fatigue, gastrointestinal discomfort, and metabolic disturbances (Cluntun et al. 2021). More importantly, current human exercise studies have not consistently detected significant changes in skeletal muscle lactylation, leaving uncertainty about its feasibility as an exercise-related intervention target. Therefore, the application of the lactate–lactylation axis in exercise-related muscle adaptation and clinical therapy still requires validation in human tissues, site-specific detection, and more selective lactylation-targeted modulators.

Limitations and perspectives

Despite rapid progress, several methodological limitations and unresolved questions still limit our understanding of lactylation in muscle cells.

From a methodological standpoint, the widespread use of pan‑Kla antibodies raises specificity concerns. These reagents may cross‑react with structurally similar acyl modifications, such as succinylation and β‑hydroxybutyrylation, potentially leading to overestimation of lactylation signals in some experimental settings. Many studies have drawn functional conclusions using p300/CBP inhibitors (e.g., C646), or LDHA inhibitors (e.g., oxamate). However, these agents also affect acetylation or global lactate metabolism, making it difficult to attribute observed phenotypes solely to lactylation. The situation is further complicated by the discovery that two distinct enzymatic systems can catatlyze lactylation: the lactoyl-CoA-dependent p300/CBP pathway and the lactoyl-CoA-independent AARS1/AARS2 pathway. To date, no study has compared their relative activities in the same tissue. Thus, it remains unclear which system predominates under different physiological or pathological conditions.

At the biological level, seemingly contradictory findings have emerged. For instance, chronic swimming exercise reduced myocardial H3K18la, which was associated with cardioprotection (Xu et al. 2024). However, in aged skeletal muscle, exercise increased the same histone mark with apparently beneficial transcriptional consequences (Meng et al. 2025). Similarly, lactylation can stabilize proteins toward divergent outcomes within the same organ: Serpina3k K351la contributed to cardioprotection after I/R injury (Wang et al. 2025), whereas Snail1 lactylation promoted pathological EndMT (Fan et al. 2023). These apparent contradictions may reflect differences in cell type, metabolic milieu, local lactoyl-CoA/acetyl-CoA balance, or eraser availability. However, such variables have rarely been controlled for in individual studies. The sequential HDAC3-then-p300 model for H4K12la in VSMCs adds another layer of complexity, suggesting that modification order rather than simple substrate competition may govern lactylation at certain loci. It remains unclear how lactylation interacts with established transcriptional regulators of muscle identity, such as the myocardin/SRF axis in VSMCs or MyoD/myogenin in skeletal muscle cells. Furthermore, whether it functions independently or modulates these existing programs is unknown. The lactoyl-CoA synthetase in mammalian tissues has yet to be identified, and muscle-specific delactylases remain poorly defined, both limitations hinder mechanistic interpretation.

From a translational perspective, the evidence base relies heavily on rodent models, often using exercise protocols—such as forced swimming to exhaustion or prolonged treadmill running—that differ substantially from typical human training in terms of intensity control, psychological stress, and metabolic context. To our knowledge, few studies have examined lactylation in human skeletal muscle or vascular tissue, and none have incorporated exercise interventions. The lactate concentrations needed to trigger specific lactylation events have not been defined in any species. Perhaps more importantly, the relationship between exercise dose (intensity, modality, duration, and training history) and lactylation magnitude has only been inferred indirectly. This inference comes from comparing results across separate studies using different protocols, species, and detection methods. It remains unknown whether lactylation responses to exercise follow a linear, threshold‑dependent, or biphasic pattern. This information could be fundamental for any future attempt to incorporate lactylation biology into exercise prescription. Addressing these gaps will likely require prospective human studies that pair standardized exercise protocols with site-resolved lactylation measurement techniques (e.g., targeted mass spectrometry). It will also require the development of more selective pharmacological tools capable of distinguishing lactylation from other acyl modifications at individual substrate sites.

Conclusions

In this review, we systematically examined the lactate–lactylation axis in skeletal muscle cells, cardiomyocytes, and VSMCs, revealing that lactylation functions as a metabolic sensor linking glycolytic flux to epigenetic reprogramming and direct protein functional regulation in muscle tissues.

In skeletal muscle cells, lactylation governs autophagy, myogenesis, and mitochondrial substrate selection via both histone-dependent transcriptional control and non-histone enzymatic modulation. In cardiomyocytes, lactylation regulates cardiac function at multiple levels–from sarcomere structural integrity to hypertrophic gene programs and ischemia–reperfusion responses. The same modification type can produce protective or pathological outcomes, depending on the target protein. In VSMCs, lactylation participates in phenotypic switching, senescence, calcification, and apoptosis. Emerging evidence suggests that the interplay between deacetylation and lactylation at shared lysine residues introduces an additional layer of regulatory complexity beyond simple substrate competition.

Several regulatory enzymes govern lactylation, including writers (p300/CBP and AARS1/AARS2), erasers (HDAC1-3 and SIRT1-3), and metabolic machinery for lactate production and transport (LDHA and MCTs). These all represent potential intervention targets. The crosstalk between lactylation and other PTMs, particularly acetylation at shared lysine sites, further underscores the integration of lactylation into broader epigenetic regulatory networks in muscle cells. Exercise, as the most potent physiological stimulus for lactate production, modulates these pathways in a manner that appears to depend on intensity, duration, and training status, though the precise parameters governing beneficial versus detrimental lactylation responses require further definition.

Collectively, lactate has transcended its traditional role as a metabolic byproduct and now functions as both a signaling molecule and a substrate for lactylation in muscle cells. Exercise-induced lactate production directly links metabolic flux to epigenetic regulation via lactylation, both on histone and non-histone proteins. The biological outcome of lactylation is context-dependent, varying across skeletal muscle cells, cardiomyocytes, and VSMCs, and is further influenced by exercise intensity and duration. The lactate-lactylation axis may offer therapeutic possibilities for metabolic and cardiovascular diseases. However, current evidence derives largely from rodent models. This mechanistic foundation awaits validation in human studies before translational applications can be reliably assessed.

Abbreviations

ATP

Adenosine triphosphate

Kla

Lactylation

LDH

Lactate dehydrogenase

MCT

Monocarboxylic acid transporter

lactyl-CoA

Lactoyl coenzyme A

NADH

Nicotinamide adenine dinucleotide

SLC

Solute carrier

GPRs

G protein-coupled receptors

VEGF

Vascular endothelial growth factor

TCA

Cycle tricarboxylic acid cycle

HIF-1α

Hypoxia-inducible factor-1α

GLUT

Glucose transporter protein

FFA

Free fatty acids

HCAR-1

Hydroxycarboxylic acid receptor 1

TCA cycle

Tricarboxylic acid cycle

ACADL

Long-chain specific acyl-CoA dehydrogenase

HADHA

Trifunctional enzyme subunit alpha

ACAT1

Acetyl-CoA acetyltransferase 1

ABCF1

ATP binding cassette subfamily F member 1

IL

Interleukin

FGF2

Fibroblast growth factor 2

cAMP

Cyclic AMP

CREB

CAMP response element-binding protein

PTMs

Post-translational modifications

PK

Pyruvate kinase

GK

Glycerol kinase

HDACs

Histone deacetylases

HATs

Histone acetyltransferases

HPLC–MS/MS

High-performance liquid chromatography-tandem mass spectrometry

CBP

CREB-binding protein C

HMGB1

High mobility group protein 1

SIRT

Sirtuin

CIRI

Cerebral ischemia–reperfusion injury

TLR

Toll-like Receptor

Arg1

Arginase 1

PDGFA

Platelet-derived growth factor A

THBS1

Thrombospondin 1

NK

Natural killer

Foxp3

Forkhead transcription factor p3

PKM2

Pyruvate kinase M2

TGF-β

Transforming growth factor-β

METTL3

Methyltransferase-like 3

JAK

Janus kinase

ROS

Reactive oxygen species

MEK/ERK

Mitogen-activated protein kinase kinase/Extracellular signal-regulated kinase

T2D

Type 2 diabetes

VPS34

Vacuolar protein sorting 34

KAT5/TIP60

Lysine acetyltransferase 5/61 kDa TBP-interacting protein

AARS1

Alanyl-tRNA synthetase

TIP60

Tat-interacting protein 60

HBO1

Histone acetyltransferase binding to origin 1

PDHA1

Pyruvate dehydrogenase E1 alpha 1

CPT2

Carnitine palmitoyltransferase 2

myokines

Muscle factors

β-OHB

β-Hydroxybutyrate

MI

Myocardial infarction

HSPA12A

Heat shock protein A12A

MI/R

Myocardial ischemia/reperfusion

ATPIF1

ATP synthase

PDK4

Pyruvate Dehydrogenase Kinase 4

NDRG

N-myc downregulated gene family

α-MHC

α-Actin heavy chain

Titin

Elastin

SMURF1

Smad ubiquitin regulatory factor 1

Dex

Dexamethasone

PDK4

Pyruvate dehydrogenase kinase 4

NR3C1

Nuclear receptor subfamily 3 group C member 1

MDH2

Malate dehydrogenase 2

P4HB

Protein disulfide-isomerase

mitoROS

Mitochondrial reactive oxygen species

PTGS2

Prostaglandin G/H synthase 2

5SM

Five small molecules

VSMCs

Vascular smooth muscle cells

PFKFB3

6-Phosphofructo-2-kinase/fructose-2,6-bisphosphatase 3

AD

Aortic dissection

TRAP1

Tumour necrosis factor receptor-associated protein 1

SASP

Senescence-associated secretory phenotype

mROS

Mitochondrial reactive oxygen species

PASMC

Pulmonary artery smooth muscle cell

TNF-α

Tumor necrosis factor α

SRY

Sex-determining region Y

SOX10

SRY-related HMG-box 10

NR4A3

Nuclear receptor subfamily 4 group A member 3

Ca²⁺

Calcium ions

DOMS

Delayed onset muscle soreness

HCAR-1

Hydroxycarboxylic acid receptor 1

NLRP3

NOD-like receptor family pyrin domain containing 3

ARR-β2

Arrestin β2

PGC-1α

Peroxisome proliferators-activated receptor γ coactivator α

COx

Cyclooxygenase

Olfr78

Olfactory receptors 78

NICD

Notch intracellular domain

YTHDF2

YTH domain-containing RNA-binding protein 2

HIIT

High-intensity interval training

iWAT

White adipose tissue

BAT

Brown adipose tissue

mPFC

Medial prefrontal cortex

SNAP91

Synaptic proteins like synaptosomal-associated protein 91

G3BP1

GTPase-activating proteinbinding protein 1

March2

Membrane-associated RING [really interesting new gene] finger protein 2 AAD: aortic aneurysm/dissection

CHI3L1

Chitinase-3-like protein 1

Arg1

Arginase 1

Mic10

Mitochondrial contact site and cristae organizing system subunit 10

VDAC1

Voltage-Dependent Anion Channel

mtDNA

Mitochondrial DNA

GAS-STING

GMP-AMP Synthase- STimulator of INterferon Genes

Authors’ contributions

M.H. contributed to project administration, funding acquisition, and writing—review, and editing. J.W.L. contributed to conceptualization, supervision, and writing— review and editing. B.C. and J.B.X contributed to conceptualization, writing—original draft preparation, and visualization. C.G.W contributed to writing—original draft preparation. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by: the National Key Research and Development Program of China (2020YFC2002904); National Natural Science Foundation of China (82202815); Natural Science Foundation of Guangdong Province (2025A1515012478 and 2026A1515010379); Guangzhou Basic Research Plan, Basic and Applied Basic Research Project (SL2022A04J00488 and SL2022A04J00446); Characteristic Innovation Project of Guangdong Provincial Education Department (2024KTSCX128); the Program for Key Research Areas of University in Guangdong Province (2024ZDZX2062), the Special Funds for the Cultivation of Guangdong College Students' Scientific and Technological Innovation (72301602 and 72400102); Beijing Sport University 2026 Graduate Research Excellence and Innovation Program (2026026).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.