Introduction

Rapid advancements in reperfusion treatments have revolutionized the management of myocardial infarction; however, it remains a significant global cause of mortality [1]. Thrombolytic treatment or percutaneous coronary intervention is the preferred treatment for blocked arterial revascularization in patients of myocardial infarction. However, reperfusion can cause severe injury to the myocardium via inflammatory mediators and oxidative stress, commonly referred to as myocardial ischemia/reperfusion injury (MI/RI) [2]. Moreover, recent evidence suggests that disorders of energy metabolism play a key role in the pathogenesis of MI/RI and promote cardiomyocyte injury and myocardial dysfunction [3, 4]. Therefore, improving the energy metabolism of cardiomyocytes is of the utmost importance for mitigating MI/RI.

Glycolysis is a conserved and ancient metabolic pathway that predates the presence of atmospheric oxygen. In contrast to mitochondrial oxidative phosphorylation, tumor cells exhibit aerobic glycolysis, also known as the “Warburg effect.” This distinct metabolic behavior enables tumor cells to generate ATP and energy in an oxygen-independent manner [5]. This metabolic adaptation enables tumor cells to survive in oxygen-deprived environments and to rapidly generate ATP. Evidence suggests that ischemic glycolysis improves myocardial tolerance to anoxia and low-flow ischemia. GLUT4, a crucial regulator of enhanced glycolysis during ischemia, has been established as a significant protective factor against ischemic injury [6]. Pinocembrin, a natural cardioprotective compound, has been shown to alleviate acute ischemia-induced myocardial injury through the promotion of glycolysis [7]. Beltran et al. demonstrated that glycolysis confers protection against MI/RI by decreasing ROS production [8]. Recent studies have suggested that m6A demethylase, fat mass- and obesity-associated protein (FTO), attenuates cardiac dysfunction in mice by regulating glycolysis and glucose uptake [9]. However, several potential mechanisms related to glycolysis in the context of MI/RI have not been identified.

Molecular chaperones such as heat shock proteins (HSPs) play a pivotal role in preventing protein misfolding and aggregation, thereby maintaining protein homeostasis [10, 11]. HSPs facilitate the adaptation of cells to their surroundings and enhance their survival under lethal conditions [12]. HSPs can increase cellular resistance to stressors, including heat shock, oxidative conditions, cytotoxic drug exposure, and apoptosis-inducing factors [13]. Hsp27, also known as HSPB1, is a ubiquitous small heat shock protein expressed abundantly in the nucleus and cytoplasm [14]. It is essential for maintaining protein homeostasis, particularly during periods of stress [15]. Research has shown that Hsp27 provides cardioprotection against oxidative stress, apoptosis, and infarction during hypoxic stress or ischemia-induced myocardial injury [12]. Additionally, levodopa treatment has been found to induce cardioprotective effects by increasing Hsp27 activity [16]. Hsp27 overexpression protects against ischemia/reperfusion-induced cardiac dysfunction [17]. Although the cardioprotective role of Hsp27 is well-established, the underlying mechanisms remain poorly understood.

Eukaryotic genomic DNA forms chromatin structures through its interaction with histone and non-histone proteins [18], including post-translational histone modifications, such as phosphorylation, ubiquitination, and acetylation, which regulate chromatin dynamics and mediate chromatin-based nuclear processes [19, 20]. Histone H2A ubiquitination, which occurs on lysine residues K129 and K15, but primarily on H2AK119 (abbreviated as H2Aub), is a prevalent modification with 5-15% H2Aub occurring in mammalian cells [21, 22]. Current research suggests that H2Aub is involved in maintaining genomic stability, DNA repair, and suppression of specific transcriptional programs, whereas H2A deubiquitination is correlated with genomic instability, activating transcription, promoting target gene activation, and facilitating cell-cycle transition [23, 24]. The ubiquitin E3 ligase responsible for H2A ubiquitination is polycomb repressive complex 1 (PRC1) [25], the core components of which are BMI1, RING1A, and RING1B, with RING1B serving as the catalytic protein. PRC1-associated E3 ligase activity is modulated at multiple stages, and RING1B self-ubiquitination is crucial for its catalytic activity [26, 27]. Recent evidence suggests that H2Aub and its ubiquitinating enzyme PRC1 are responsible for the onset and progression of cardiovascular disorders [28, 29]. Although researchers are primarily investigating the relationship between H2Aub and tumors, the role of PRC1 in regulating H2Aub levels in chromatin and the involvement of transcriptional targets in cardiovascular disorders, specifically MI/RI, is yet to be determined.

This study established a strong association between H2Aub chromatin and the enrichment of Hsp27, suggesting an interaction between these two factors. Moreover, Hsp27 knockdown significantly downregulated PFKFB3 expression and worsened hypoxia-induced myocardial cell injury, suggesting that H2Aub is at least partially responsible for Hsp27-mediated glycolytic alterations in hypoxic cardiomyocytes. Additional research confirmed that Hsp27 maintained mitochondrial function in hypoxic-reoxygenated (H-R) cardiomyocytes by stabilizing COQ9. PCM-SH, a 21-mer peptide isolated through phage display, exhibited a 180-fold higher binding affinity for primary cardiomyocytes than the control, suggesting high specificity [30]. Based on this mechanism, PPMP-coated PRT (PRT@PPMP) nanoparticles were developed to inhibit PRC1-mediated H2Aub with the goal of further increasing glycolysis levels, improving mitochondrial function, and inhibiting MI/RI. This study suggests that the targeted manipulation of H2Aub levels could be a promising therapeutic strategy for MI/RI.

Results

Hsp27 is upregulated after hypoxia and enhances glycolysis in cardiomyocytes

The literature suggests that glycolysis occurs in MI [8]; however, the underlying mechanistic details, particularly regarding the metabolic reprogramming responses to ischemia in cardiomyocytes, are not fully understood. To establish the MI model in rats, myocardial ischemia was induced for 30 min, and electrocardiograms were monitored, which revealed significant ST-segment elevation in MI rats (Fig. S1A). Echocardiography results indicated a significant decrease in left ventricular fractional shortening (LVFS) and left ventricular ejection fraction (LVEF) in MI rats (Fig. S1B–D). Successful establishment of the MI model was confirmed by hematoxylin and eosin (HE) staining, H2AX immunofluorescence, and triphenyltetrazolium chloride (TTC) staining (Fig. S1E–G). ATP and secreted lactate levels were measured in the myocardial tissue, revealing a metabolic shift from oxidative phosphorylation to glycolysis due to ischemia, which resulted in reduced ATP levels and increased lactate secretion (Fig. S1H–I). Metabolic profiling confirmed the accumulation of glycolytic intermediates in ischemic rats (Fig. S1J). In vitro study, hypoxia was induced for 12 h in cardiomyocytes, and CCK-8 assay results demonstrated that hypoxia significantly reduced the viability of myocardial cells (Fig. S2A-B). ATP and lactate levels were measured to investigate glycolytic alterations under hypoxic conditions, revealing that hypoxia decreased ATP content and promoted lactate secretion (Fig. S2C-D). Additionally, Seahorse XF assays measuring the extracellular acidification rate (ECAR) indicated that hypoxia increased glycolysis in the myocardial cells (Fig. S2E-F).

RNA sequencing was performed in rats with and without MI to identify differentially expressed genes and signaling pathways. The results revealed that activation of the MAPK signaling pathway, which promotes glycolysis [31, 32], was associated with MI (Fig. 1A). Furthermore, a subset of ten significantly upregulated genes (log2 fold-change >2 and P < 0.01) in the MAPK signaling pathway was selected (Fig. 1B). qRT-PCR was used to validate changes in gene expression after hypoxia in vitro and in vivo, revealing significant increases in the expression of Il1b, Myc, Hspa1b, Pgf, Cd14, Map3k6, Dusp2, Hsp27, Flnc, and Il1α in the myocardial tissue of MI rats and hypoxic cardiomyocytes (Fig. S3A-B). The role of these genes in glycolysis was determined by individual gene knockdown in cardiomyocytes, revealing that only Hsp27 depletion significantly decreased acidification of the culture medium (Fig. S3C, Fig. 1C-D). Additionally, alterations in Hsp27 expression were detected in fibroblasts and endothelial cells, with more pronounced changes in cardiomyocytes (Fig. S4A). Metabolic profiling revealed increased production of glycolytic intermediates in hypoxic cardiomyocytes, which was reduced upon Hsp27 knockdown (Fig. 1E). Furthermore, Hsp27 knockdown reduced glycolytic flux in cardiomyocytes (Fig. 1F-G) and decreased cellular ATP production (Fig. 1H). Investigation of cardiomyocyte viability using CCK-8 revealed that Hsp27 knockdown worsened hypoxia-induced damage to cardiomyocytes by reducing their glycolytic capacity (Fig. 1I-J). These findings suggested the involvement of Hsp27 in glycolysis of hypoxic cardiomyocytes.

Fig. 1: Hsp27 knockdown inhibits glycolysis in hypoxic cardiomyocytes.
figure 1

A Pathway enrichment statistics for the identified mRNA. B Heat map showing upregulated mRNA of the MAPK signaling pathway in the rat heart after 30 min of ischemia. Color-coded representation of gene expression levels: Blue and red represented low and high expression, respectively. Data are representative of three independent experiments. C Immunofluorescence images of Hsp27 (red) staining in cardiomyocytes cultured with or without Hsp27 knockdown after 12 h of hypoxia. D Culture medium acidification following Hsp27 depletion in 12-h hypoxic cardiomyocytes. E Heat map showing glycolytic intermediates 12 h after hypoxia stimulation. Color-coded representation of gene expression levels: Green and red represented low and high expression, respectively. F–G Graphs showing the results of glycolytic stress analyses using a Seahorse XF analyzer to assess the response of cardiomyocytes to oligomycin, glucose, and 2-DG. H Intracellular ATP levels in cardiomyocytes. I–J Detection of cardiomyocyte viability. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Hypoxia group. Scale bars: (C) 50 μm; (I) 100 μm.

Full size image

Hsp27 enhances glycolysis by activating PFKFB3 transcription

The initial concern was whether changes in the expression of glycolytic pathway enzymes could explain the alterations in glycolytic flux observed during MI. RNA-seq analyses of these pathway enzymes revealed an increase in PFKFB3, HK3, TP53, and PFKL expression in MI rats (Fig. 2A). Analysis of the mRNA levels of PFKFB3, HK3, TP53, and PFKL in cardiomyocytes revealed that PFKFB3 expression was decreased upon Hsp27 knockdown, whereas the levels of HK3, TP53, and PFKL remained unchanged (Fig. 2B). Subsequently, PFKFB3 blockade was found to suppress glycolysis in cardiomyocytes. Figure 2C–E demonstrated that suppressing PFKFB3 caused a significant decline in ATP production and glycolytic capacity in hypoxic cardiomyocytes.

Fig. 2: Hsp27 promotes glycolysis by regulating PFKFB3 via NF-κB.
figure 2

A Schematic representation of upregulated glycolytic pathway enzymes. B Relative mRNA expression levels of PFKFB3, HK3, TP53, and PFKL detected using qRT-PCR in Hsp27 knockdown cardiomyocytes after 12 h of culture under hypoxic conditions. C, D Seahorse XF assay measuring the ECAR of cardiomyocytes cultured with or without PFKFB3 knockdown under control and hypoxic conditions for 12 h. E ATP levels in the cardiomyocytes. F Luciferase activity analysis in cells following co-transfection with luciferase reporters containing the NF-κB mimic and PFKFB3-wt. Data are presented as mean ± SD (n = 3). #P < 0.05 vs. the PFKFB3-wt group. G Statistical analysis of relative nuclear NF-κB content in cardiomyocytes. H Detection of IKK activity in cardiomyocytes cultured with or without Hsp27 knockdown and subjected to hypoxia for 12 h. I Immunoprecipitation using control IgG or anti-Hsp27 in 12-h hypoxic cardiomyocytes. Immunoblotting of precipitates for p-IKKα and Hsp27. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Hypoxia group; ns, not significant.

Full size image

Previous studies have highlighted that NF-κB activation may increase PFKFB3 expression [33]. To confirm direct binding between PFKFB3 and NF-κB, a luciferase reporter assay was performed using wild-type and mutant plasmids of PFKFB3. Co-transfection of luciferase plasmids with NF-κB mimic/NC into cells revealed that the NF-κB mimic significantly increased luciferase activity in the wild-type PFKFB3 plasmid, but did not affect the mutant PFKFB3 luciferase activity (Fig. 2F). NF-κB typically forms a heterodimer with P65 and P50 and is rendered inactive in the cytoplasm due to binding to the inhibitory protein IκB [34, 35], resulting in the formation of a trimer complex. Nuclear NF-κB content was increased in response to hypoxia, but this effect was reversed when Hsp27 was knocked down (Fig. 2G). These data suggests that Hsp27 may affect NF-κB activation in hypoxic cardiomyocytes. To further validate this hypothesis, IKK activity was assessed, and the results indicated that hypoxia promoted IKK activity, which was subsequently reversed by Hsp27 knockdown (Fig. 2H). The presence of p-IKKα in the immunoprecipitates was consistent with the high tendency of NF-κB activation in cardiomyocytes (Fig. 2I). These findings suggested that Hsp27 promoted PFKFB3 transcription under hypoxic conditions by binding and activating IKK.

Hsp27 improves mitochondrial function in H-R cardiomyocytes

To investigate the potential protective effects of Hsp27 during reperfusion, RNA sequencing was performed in rats with or without MI/RI. Interestingly, Hsp27 expression remained high in the MI/RI model (Fig. 3A). Literature indicates that Hsp27 maintains cell membrane integrity under various stresses [36], providing cardiac protection via the chaperone effect and facilitating cytoskeleton reconstruction under stress. Hsp27 also induces antioxidant mechanisms through the association of myocardial lesion generation with free radical production [37,38,39]. Furthermore, Hsp27 significantly inhibits ROS generation [40, 41]. Mitochondria are the primary source of ROS production in cardiomyocytes [42]. Mitochondrial membrane potential is commonly used to assess mitochondrial function, and a decrease in the mitochondrial membrane potential suggests mitochondrial dysfunction. Therefore, to confirm the significance of Hsp27 in improving mitochondrial function, the mitochondrial membrane potential was evaluated in H-R cardiomyocytes by Hsp27 knockdown. H-R cardiomyocytes exhibited significantly higher JC-1 monomer levels than the control cardiomyocytes. Moreover, Hsp27 knockdown further increased JC-1 monomer levels compared to those in H-R cardiomyocytes (Fig. 3B). To investigate the association between Hsp27 and H-R-induced ferroptosis, we assessed ROS levels, lipid peroxides, mitochondrial ROS, MDA content, and ptgs2 mRNA expression and found that Hsp27 knockdown increased these parameters in H-R cardiomyocytes (Fig. 3C–F). Hsp27 silencing further reduced the viability of H-R cardiomyocytes (Fig. 3G–H). These findings suggested that Hsp27 mitigated mitochondrial dysfunction and ferroptosis in H-R cardiomyocytes.

Fig. 3: Hsp27 improves mitochondrial function in hypoxic-reoxygenated cardiomyocytes.
figure 3

A, B Heat map depicting differentially expressed genes of the MAPK signaling pathway in cardiac tissue between the sham and MI/RI groups. Color-coded representation of gene expression levels: Green and red represented low and high expression, respectively. Data are representative of three independent experiments. B Representative images of JC-1 fluorescence in the H-R cardiomyocytes. C Detection of cellular ROS and lipid peroxide levels in the cardiomyocytes. D Statistical analysis of mitochondrial ROS levels. E Cellular MDA levels in cardiomyocytes. F Relative mRNA levels of ptgs2 in cardiomyocytes determined using qRT-PCR. G, H Viability of cardiomyocytes in the different groups. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Reoxygenation group. Scale bars: [C (top)] 50 μm; [B, C (bottom)] 25 μm; and (G) 100 μm.

Full size image

Hsp27 functions as a chaperone of COQ9 in H-R cardiomyocytes

The mitochondrial electron transport chain (ETC) uses a series of electron transfer reactions to generate ATP via oxidative phosphorylation. The generation of ROS during electron transfer can lead to oxidative stress and homeostatic signaling during disease pathology [43]. Thus, the activity of ETC complex I, whose dysfunction is primarily responsible for mitochondrial ROS production [44], was detected in this study. The results indicated that Hsp27 knockdown resulted in a further reduction in COQ9 protein expression compared to NDUFS1, ETFDH, ETFA, and NDUFB5 (Fig. 4A, B). However, the mRNA levels of COQ9 were not altered by Hsp27 knockdown (Fig. 4C). The reduction in COQ9 protein levels was independent of alterations in mRNA levels, excluding the possibility of transcriptional modifications. To investigate the effect of COQ9 on mitochondrial ROS production, we overexpressed COQ9 in H-R cardiomyocytes and observed a significant reduction in both mitochondrial and cellular ROS levels following COQ9 overexpression (Fig. 4D, G). Researchers have found that small HSPs function as molecular chaperones in the mitochondrial intermembrane space, and that the mitochondrial interactome of Hsp27 is enriched in transmembrane proteins of the inner mitochondrial membrane [45]. Therefore, we explored the potential role of Hsp27 as a molecular chaperone in maintaining COQ9 expression using Cluspro software, and the results confirmed a significant binding interaction between Hsp27 and COQ9 (Fig. 4H). COQ9 was detected in the immunoprecipitates, validating the prediction of the Cluspro software (Fig. 4I). These data showed that Hsp27 acted as a chaperone for COQ9 in H-R cardiomyocytes.

Fig. 4: Hsp27 stabilizes COQ9 levels in hypoxic-reoxygenated cardiomyocytes.
figure 4

A, B Western blot analysis of NDUFS1, ETFDH, COQ9, ETFA, and NDUFB5 in H-R cardiomyocytes. C Relative mRNA expression levels of COQ9 detected using qRT-PCR in Hsp27 knockdown cardiomyocytes. D, E Immunoblot analysis of COQ9 in cardiomyocytes transfected with COQ9. F Statistical analysis of mitochondrial ROS levels. G Detection of cellular ROS levels in H-R cardiomyocytes. H Diagram showing the binding site of Hsp27 (blue) and COQ9 (green). I Immunoprecipitation using control IgG or anti-Hsp27 in H-R cardiomyocytes. Immunoblotting of precipitates for COQ9 and Hsp27. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Reoxygenation group; ns, not significant. Scale bars: 50 μm.

Full size image

Hsp27 has been identified as a H2Aub nucleosome-associated gene in cardiomyocytes

The role of H2Aub was not well understood until recent studies revealed its association with DNA damage repair and gene suppression [46, 47]. H2Aub ChIP-seq analyses were conducted to identify H2Aub binding genes, and determine the corresponding transcriptional alterations in the genome of MI rats. Integration of H2Aub RNA-seq and ChIP-seq datasets facilitated the identification of 162 downregulated and 211 upregulated genes (Fig. 5A). Interestingly, KEGG analysis revealed the enrichment of H2Aub-regulated genes in the MAPK signaling pathway (Fig. 5B). In particular, the Venn diagram revealed that 23 genes were involved in the MAPK signaling pathway among the 211 upregulated genes with H2Aub occupancy. Additionally, Hsp27 was among the 23 upregulated genes with H2Aub occupancy (Fig. 5C). ChIP–seq of H2Aub revealed that ischemia increased H2Aub occupancy within Hsp27 gene (Fig. 5D). The ChIP-qPCR results also confirmed H2Aub binding to Hsp27 in both normal and hypoxic cardiomyocytes (Fig. 5E). Overexpression of RING1B in hypoxic cardiomyocytes increased H2Aub protein expression, but decreased the mRNA and protein levels of Hsp27 (Fig. 5F–J). These findings suggested the need for further mechanistic studies on H2Aub-regulated Hsp27, and indicated that RING1B-mediated H2Aub on Hsp27 inhibited transcriptional initiation.

Fig. 5: RING1B increases H2Aub occupancy of Hsp27.
figure 5

A Venn diagram illustrating the overlap of 1,289 genes with H2Aub binding and 211 upregulated genes. B KEGG pathway analysis of ChIP-seq data. C Venn diagram illustrating the overlap of 122 genes involved in the MAPK signaling pathway with H2Aub binding and 211 upregulated genes. D H2Aub ChIP–seq occupancy profiles at the Hsp27 loci in ischemic myocardial tissue. E ChIP–qPCR validation of H2Aub binding to Hsp27 in normal and hypoxic cardiomyocytes. Data are presented as mean ± SD, n = 6 independent replicates. #P < 0.05 vs. IgG group. F Immunofluorescence analysis of RING1B in cardiomyocytes treated with hypoxia for 12 h with or without RING1B or control vector (NC-RING1B) transfection. G, H Immunoblot analysis of H2Aub in hypoxic cardiomyocytes, with or without RING1B or control vector (NC-RING1B) transfection. I Quantification of Hsp27 expression in cardiomyocytes using qRT-PCR. J Representative images of Hsp27 staining (red) in cardiomyocytes counterstained with DAPI (blue). Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Hypoxia group. Scale bars: (F) 25 μm; (J) 50 μm.

Full size image

Design, synthesis, and characterization of PRT@PPMP

PRT effectively inhibits PRC1-mediated H2Aub, and if nanocomplexes successfully deliver PRT to the damaged myocardium, they could decrease H2Aub and provide additional protection to the myocardium. The overall synthesis strategy for PRT@PPMP was shown in Fig. 6A. Electron microscopy revealed that PRT@PPMP exhibited a uniform and spherical morphology, with a mean particle size of approximately 25 nm (Fig. 6B, C). The zeta potential was determined to analyze the charge variations and surface modifications of PPM. After loading PPMP with PRT, the zeta potential was altered to -20.54 ± 1.99 mV (Fig. 6D). The internalization of the PRT@PPMP nanospheres within cells is crucial for their therapeutic effectiveness. The PRT@PPMP nanoplatforms functionalized with PCM-SH are expected to demonstrate specificity in their homologous targeting capacity. To demonstrate the specific recognition and targeted delivery of PRT@PPMP to cardiomyocytes (CM), PRT@PPMP was administered to CM and cardiac microvascular endothelial cells (CMECs). The observation of weak PRT@PPMP signals in CMECs indicated the low efficiency of PRT@PPMP uptake. In contrast, the intracellular PRT@PPMP fluorescence signal in the CM was stronger than that in the CMECs (Fig. 6E). Next, the targeting ability of PRT@PPMP in H-R cardiomyocytes was analyzed using confocal laser scanning microscopy (CLSM). Cardiomyocytes incubated with PRT@PPMP exhibited a more intense green fluorescence than PRT@PPM nanospheres (Fig. 6F), suggesting that the PCM-SH-modification facilitated the intracellular uptake of nanocarriers. Moreover, prolonging the co-incubation duration further amplified the intensity of the green fluorescence. After 2 h of incubation, the PRT@PPMP nanosphere-treated group exhibited a higher rate of intracellular uptake than the PRT@PPM nanosphere-treated group. Flow cytometric analysis of the cellular uptake of PRT@PPM and PRT@PPMP by cardiomyocytes revealed that the fluorescence intensity of PRT@PPMP in the cells was significantly higher than that of the PRT@PPM and PBS groups (Fig. 6G). These findings suggested that PCM-SH promoted the intracellular uptake of nanocarriers, thereby exerting a more potent therapeutic effect.

Fig. 6: Preparation and characterization of PRT@PPMP.
figure 6

A Schematic illustrating the design of the PRT@PPMP. B Electron microscopic images of PRT@PPMP. C Size distribution. D The surface charge of PPM, PRT@PPM, and PRT@PPMP determined using the zeta potential and dynamic light scattering. E Fluorescence microscopy revealing the cellular uptake of PRT@PPMP by CMECs and CMs. F Immunofluorescence images showing intracellular localization of PRT@PPM and PRT@PPMP within CMs at different time points. Green-labeled PRT@PPM and PRT@PPMP with DAPI-stained nuclei (blue). G Cellular uptake of PRT@PPM and PRT@PPMP by CMs, observed through flow cytometry analysis. Data are presented as mean ± SD (n = 6). Scale bars: 25 μm.

Full size image

Therapeutic effects of PRT@PPMP on H-R cardiomyocytes

The effect of PRT@PPMP on cardiomyocyte viability was observed in vitro. At concentrations ≤20 μM, PRT@PPMP displayed minimal effects on cell viability, whereas at a concentration of 40 μM, cell viability was significantly decreased compared to that of untreated cells (Fig. S5A). In addition, investigating the effect of administration time on cell viability revealed no significant alterations in cell viability after 4 days of PRT@PPMP administration at concentrations of 5, 10, and 20 μM (Fig. S5B). PRT effectively suppressed RING1B expression in cardiomyocytes, and the inhibitory rate was achieved over 70% at 20 μM (Fig. S5C, E). The time axis scheme in Fig. 7A depicts the glycolytic capacity of cardiomyocytes subjected to 12-h of hypoxia. Cardiomyocytes were pretreated with PBS, PRT, PRT@PPM, or PRT@PPMP (20 μM PRT) before induction of hypoxia. Western blotting and immunofluorescence demonstrated that Hsp27 levels were significantly decreased in a time-dependent manner (Fig. S6A–C) in vivo. The results of immunofluorescence, western blotting, and qRT-PCR suggested that PRT@PPMP pretreatment effectively stabilized the high expression of Hsp27 in cardiomyocytes (Fig. S7A, Fig. 7B). Subsequent investigation revealed that PRT@PPMP, but not PRT and PRT@PPM, increased glycolytic flux (Fig. 7C, D) and ATP generation (Fig. 7E) in hypoxic cardiomyocytes.

Fig. 7: PRT@PPMP reverses myocardial injury by enhancing glycolysis during hypoxia and improving mitochondrial function during reoxygenation.
figure 7

A Scheme of the time axis representing the design of hypoxic cell research. B Immunofluorescence results showing Hsp27 expression in control cardiomyocytes and hypoxic cardiomyocytes treated with PRT, PRT@PPM, and PRT@PPMP. C, D Glycolytic stress analyses of cardiomyocytes using the Seahorse XF analyzer in response to oligomycin, glucose, and 2-DG. E Intracellular ATP levels in cardiomyocytes. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Hypoxia group. F Scheme of the time axis illustrating the design of H-R-induced cell study. G Representative images of JC-1 fluorescence in the control and reoxygenated cardiomyocytes treated with PRT, PRT@PPM, and PRT@PPMP. H Immunofluorescence detection of cellular ROS levels in cardiomyocytes. I Mitochondrial ROS levels in cardiomyocytes. J MDA levels in cardiomyocytes. K Relative mRNA levels of ptgs2 in cardiomyocytes determined using qRT-PCR. L–M Detection of cardiomyocyte viability. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Control group; *P < 0.05 vs. Reoxygenation group. Scale bars: (B, H) 50 μm; (G) 25 μm; and (L) 100 μm.

Full size image

Figure 7F depicts the time-axis scheme of the design for H-R cardiomyocytes. Cardiomyocytes were pre-treated with PBS, PRT, PRT@PPM, or PRT@PPMP (20 μM PRT). Compared with the control group, the JC-1 monomer exhibited a significant increase in H-R cardiomyocytes. However, this increase was reversed by the PRT@PPMP treatment (Fig. 7G). Moreover, the levels of ROS, MDA, and ptgs2 in the PRT and PRT@PPM treatment groups showed no obvious changes compared to the reoxygenated group. However, these levels were significantly reduced in the PRT@PPMP group (Fig. 7H–K). The CCK-8 assay results (Fig. 7L–M) revealed that the reoxygenated treatment decreased cell viability, while the PRT@PPMP exhibited increased cell viability, indicating the strong protective effect of PRT@PPMP on cardiomyocytes. These findings suggested that PRT@PPMP promoted glycolysis, improved mitochondrial function, and inhibited ferroptosis in H-R cardiomyocytes by upregulating Hsp27 expression.

PRT@PPMP protects the myocardium against ischemia/reperfusion injury

Figure 8A shows the experimental design of the animal studies. Analysis of cardiac function indicators revealed that the PRT@PPMP-4 and PRT@PPMP-8 (PRT administered at doses of 4 and 8 mg/kg) exhibited superior effectiveness in restoring LVFS and LVEF compared to the reperfusion group. This highlighted the significant treatment efficacy of both PRT@PPMP-4 and PRT@PPMP-8 in enhancing the function of the damaged myocardium (Fig. 8B–D). Therefore, PRT@PPMP-4 (PRT@PPMP) was selected for subsequent experiments. The echocardiographic results confirmed that PRT@PPMP did not affect cardiac function in sham rats (Fig. S8A–C). Next, the distribution of fluorescent signals of the nanocomplex in the heart was quantitatively analyzed using bioimaging techniques. In the PRT@PPM group, the weak fluorescence signals were observed in the heart, whereas in the PRT@PPMP group, the strong fluorescence signals were exhibited, demonstrating good target specificity of PRT@PPMP (Fig. 8E). PCM-SH facilitates the binding of PRT@PPMP to cardiomyocytes, and this interaction is specific to the heart because most of the other tissues or organs, such as the spleen, lungs, liver, and kidneys, exhibited significantly lower distribution of PRT@PPMP (Fig. 8F).

Fig. 8: Targeted release of PRT in the myocardium prevents cardiac dysfunction.
figure 8

A Scheme of the time axis representing the design of the animal study. B Representative echocardiographic images from each group. C, D Echocardiographic parameters, such as LVEF and LVFS, were calculated in sham and reperfusion rats treated with or without different concentrations of PRT@PPMP. E Immunofluorescence study depicting the delivery efficiency of Cy5.5-stained PRT@PPM or PRT@PPMP to the heart 6 h after individualized treatment. F Cy5.5-stained PRT@PPMP accumulated in the heart and other organs. G Representative images of the heart sections labeled with Hsp27 fluorescent probe. DAPI (blue, nuclei); Hsp27 positive (red). H Representative morphological analysis using HE staining. I–J TTC staining revealing the area of myocardial infarction in rats. K ROS levels in the cardiac tissue. L Electron microscopic analysis of myocardial cells revealing changes in mitochondrial morphology. M MDA levels in the cardiac tissue. N Relative mRNA levels of ptgs2 measured in cardiac tissue using qRT-PCR. Data are presented as mean ± SD (n = 6). #P < 0.05 vs. Sham group; *P < 0.05 vs. Reperfusion group. Scale bars: (G, H) 50 μm; (L) 2.0 μm.

Full size image

Moreover, immunofluorescence analysis revealed that PRT@PPMP pre-treatment effectively maintained an elevated level of Hsp27 in the heart (Fig. 8G). Only PRT@PPMP significantly increased the ATP production (Fig. S9A). To investigate the targeted therapeutic effect of PRT@PPMP on MI/RI rats, TTC and HE staining of hearts were performed to detect infarcted areas, and the results demonstrated a significant reduction in infarcted regions with PRT@PPMP (Fig. 8H–J). Ferroptosis markers in MI/RI hearts were examined to investigate the protective effects of PRT@PPMP in vivo. In the model group, exposure to PBS resulted in elevated levels of MDA, ROS, ptgs2, and atrophic mitochondria in the myocardial cells, as evidenced by qRT-PCR, ELISA, and electron microscopy. The ferroptosis indicators showed no changes in the PRT and PRT@PPM groups. However, in the PRT@PPMP group, there were significant reduction in ferroptosis indicators, indicating the potent anti-ferroptosis effect of PRT@PPMP on myocardial cells (Fig. 8K–N). Overall, the findings of this study confirmed that PRT@PPMP could specifically target the damaged myocardium and effectively enhance glycolysis while inhibiting ferroptosis, thereby conferring protection to the myocardium against MI/RI.

Discussion

Despite advancements in acute myocardial infarction (AMI) treatment and improved patient survival rates, ischemic heart disease remains a leading cause of mortality worldwide. Therefore, the development of novel cardioprotective strategies is crucial to mitigate the detrimental effects of AMI. Restoring coronary blood flow reperfusion is the most effective approach to improve AMI outcomes [48]. However, reperfusion can also exacerbate myocardial injury [49]. Therefore, identifying effective treatment targets for MI/RI inhibition is of utmost importance. This study reported a significant increase in Hsp27 levels through histone H2Aub in cardiac tissue during ischemia, followed by a decrease during reperfusion. Moreover, the upregulation of Hsp27 promoted glycolysis under ischemic conditions and stabilized the mitochondrial function during reperfusion. Further mechanistic investigations indicated that Hsp27 boosted glycolysis by promoting nuclear NF-κB translocation, leading to the upregulation of PFKFB3 expression during ischemia. In addition, Hsp27 inhibited ferroptosis by improving mitochondrial function through the stabilization of COQ9. The present study also introduced a noninvasive, targeted nanocomplex delivery platform using PCM-SH-coated PRT for the treatment of MI/RI. Myocardial cells phagocytized the accumulated PRT@PPMP in the damaged heart, resulting in therapeutic efficacy against MI/RI (Fig. 9). Furthermore, this study explored the intricate mechanism of PRT@PPMP treatment in MI/RI. First, target-specific delivery of PRT@PPMP was successfully accomplished. Upon uptake by cardiomyocytes, PRT@PPMP decomposition triggered the release of PRT into the cytoplasm, ultimately inhibiting PRC1. Additionally, reduced H2Aub expression stabilized the high Hsp27 expression, thereby augmenting glycolysis during ischemia and improving mitochondrial function during reoxygenation. This strategy provides a practical approach for implementing PRT treatment for MI/RI.

Fig. 9: Therapeutic mechanism of PRT@PPMP in MI/RI.
figure 9

PRT@PPMP targets injured cardiomyocytes, leading to the decomposition and release of PRT4165 into the cytoplasm. PRT4165 decreases the expression of RING1B and H2Aub and increases Hsp27 expression. Hsp27 activates the NF-κB/PFKFB3 signaling pathway, leading to increased glycolysis levels during hypoxia. Additionally, Hsp27 functions as a chaperone of COQ9 during reoxygenation to improve mitochondrial function and inhibit ferroptosis.

Full size image

HSPs, a highly conserved protein family activated in response to various physiological and environmental challenges such as hyperoxia and hypoxia, as well as emotional and mechanical stress, play a critical role in maintaining homeostasis of cells under stress or allowing cell survival in lethal environments [50,51,52]. Myocardial cells, with their high metabolic demands and abundant signal transduction pathways, are particularly reliant on chaperones for survival. Hsp27, a prominent member of the small HSP family, is highly expressed in the myocardium and exhibits cardioprotective properties [53]. Small HSPs regulate protein folding, provide significant protection against apoptosis and oxidative stress, and contribute to the maintenance of sarcomeric structure. Despite the well-established role of Hsp27 in the stress response and characterization, there is currently no practical approach for maintaining its activity in MI/RI for cardiac protection. In this study, Hsp27 was identified as a glycolysis-associated gene, and maintaining a high level of Hsp27 enhanced glycolysis via the NF-κB/PFKFB3 signaling pathway. Studies have confirmed that Hsp27 exhibits potent chaperone activity against amyloid and amorphous aggregation of proteins, such as Tau, α-synuclein, and SOD1 [54, 55]. Researchers have revealed that small heat shock proteins, including Hsp27, function as chaperone systems within the mitochondrial intermembrane space [56]. Similarly, the present study indicated that Hsp27 also enhanced mitochondrial function by stabilizing COQ9 protein expression rather than through changes in mRNA levels. Therefore, Hsp27 acted as a molecular chaperone, facilitated the proper protein folding of COQ9. These findings indicated that Hsp27 played a protective role in the myocardium by promoting glycolysis and improving mitochondrial function.

Published literature has identified covalent modification of histone core models in multiple DNA-based processes, including transcription, DNA repair, and replication [57]. Histone ubiquitination is implicated in various cellular processes. In particular, H2Aub, which primarily occurs at Lys119, is crucial because of its association with histones and its extensive range of modifications [58]. H2Aub is primarily correlated with gene repression in chromatin and is strongly associated with PRC1, the primary ligase enzyme for H2Aub [59]. Recent studies have also shown that H2Aub is linked to the emergence of multiple diseases. For example, BAP1 has been found to reduce H2Aub occupancy of the SLC7A11 promoter, leading to the suppression of SLC7A11 expression and contributing to elevated ferroptosis and lipid peroxidation [60]. Additionally, TRIM37 orchestrates the progression of renal cell carcinoma by regulating histone H2Aub [61]. However, the mechanisms underlying PRC1-regulated H2Aub suppression of target genes are not well understood in the context of cardiovascular disease. In this study, ChIP-seq analysis revealed that H2Aub regulated 1289 genes in the MI model. Moreover, KEGG analysis showed that H2Aub-regulated genes were enriched in the MAPK signaling pathway, which included Hsp27. The experimental results indicated reduced levels of H2Aub and RING1B in the hearts of MI rats and hypoxic cardiomyocytes. Enforcing RING1B expression in hypoxic cardiomyocytes increased H2Aub protein expression while simultaneously reducing the mRNA and protein levels of Hsp27. Furthermore, ChIP-qPCR results confirmed that H2Aub combined with Hsp27. These data suggested that H2Aub directly inhibited Hsp27 expression.

Recently, researchers have shifted their attention toward targeted therapies for MI/RI. This approach involves the use of a peptide-conjugated nanoparticle (PRT@PPMP) for the targeted delivery of a PPM-conjugated 21-mer peptide, namely PCM-SH, to the heart. In vitro and in vivo studies have demonstrated the preferential binding of PRT@PPMP to cardiomyocytes compared to the scrambled control, with the PRT@PPMP signal being prominently distributed in the rat heart. This indicated that PCM-SH could efficiently guide nanoparticles toward the heart. PRT accumulated in the injured heart was phagocytosed by cardiomyocytes, resulting in long-term treatment efficacy against MI/RI. Furthermore, the specific mechanism by which PRT@PPMP treatment affected MI/RI was investigated, revealing enhanced glycolysis, improved mitochondrial function, and reduced ferroptosis in cardiomyocytes. Therefore, this study provides a novel strategy for the clinical treatment of MI/RI.

In summary, this study reveals the involvement of H2Aub in the development of MI/RI. A novel regulatory pathway in glycolysis involving the H2Aub/Hsp27/NF-κB/PFKFB3 axis was established through mechanistic studies. Furthermore, the upregulation of Hsp27 expression improved mitochondrial function through its role as a chaperone for COQ9. More importantly, a noninvasive, targeted nanocomplex delivery platform was developed using PPMP-coated PRT to treat MI/RI. These findings not only expand our understanding of the potential therapeutic role of PRT in cardiovascular disease, but also highlight the importance of developing therapeutic strategies that target the H2Aub pathway.

Materials and methods

Micelle preparation and characterization

PCL-PEG-MAL or PCL-PEG-PCM (25 mg) was dissolved in 5 ml methanol, mixed with 15 mg of PRT, and subsequently dissolved in dichloromethane (5 ml). The thin solid film resulting from the evaporation of the solvent was dissolved in 10 ml PBS solution, gently agitated for 1 h, and then filtered using a 0.2 μm syringe filter to remove non-encapsulated or precipitated PRT. PRT-free blank micelles were also prepared.

Establishment of the MI/RI model

MI/RI was performed as previously described [62]. Briefly, male rats were anesthetized through isoflurane gas inhalation and placed in the supine position, where they were surgically assisted with breathing using an animal ventilator. Left thoracotomy was performed through a horizontal incision in the fourth intercostal space, and the left anterior descending coronary artery was ligated around the PE-10 tube with 6-0 sutures. Occlusion of the left anterior descending coronary artery was confirmed by elevation of the ST segment on electrocardiogram. After 30 min, the PE-10 tube was removed and the reperfusion was performed for 6 h. The hearts were harvested for subsequent experiments. The rats were randomly divided into five groups with six rats in each group, as follows: (1) sham group rats underwent a sham operation and received vehicle (PBS, caudal vein injection); (2) reperfusion group rats were subjected to MI/RI and received vehicle (PBS, caudal vein injection); and (3–5) PRT groups rats were received PRT, PRT@PPM, or PRT@PPMP (4 mg/kg, caudal vein injection). PRT, PRT@PPM, or PRT@PPMP was administered once daily for three days, and the ischemia-reperfusion injury model was established immediately after the last administration. In this study, the rats were subjected to MI/RI and treated with the above interventions randomly. Efficacy was evaluated based on morphology, transthoracic echocardiography, and biomarker analyses in a blinded fashion.

Echocardiographic evaluation

Echocardiography was performed following a previously described method [63]. The rats were anesthetized using isoflurane gas inhalation, and their chest hair was removed using a depilation agent. The chest was coated with ultrasound gel and scanned using a contact scanning head.

TTC and HE staining

The rats were euthanized after ischemia or reperfusion and their hearts were obtained. The hearts were subsequently stored at –20 °C for 25 min. The heart tissue was sectioned into five pieces and placed in 2% TTC at 37 °C for 15 min in the dark [64]. The infarct size was determined using ImageJ software. Moreover, several heart sections (4–5 μm thick) were prepared and stained with HE for histopathological analysis [65].

Electron microscopy

Mitochondrial alterations in cardiomyocytes during MI/RI were observed using electron microscopy as described previously [66].

Detection of reactive oxygen species (ROS), lipid peroxide, NF-κB content, and IKK activity

ROS, lipid peroxide, NF-κB content, and IKK activity were measured according to the manufacturer’s instructions.

Quantitative reverse-transcription PCR (qRT-PCR)

Total RNA from cardiomyocytes and cardiac tissues was isolated and reverse transcribed. The mRNA levels were detected using qRT-PCR and normalized to β-actin expression levels. Primer sequences used in this study are listed in Table S1.

Western blot

Myocardial tissue or cultured cardiomyocytes were lysed using RIPA lysis buffer for western blot analysis. Protein extracts (50 μg) were isolated using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to a nitrocellulose membrane, and probed with primary antibodies overnight at 4 °C (Table S2). Enhanced chemiluminescence (ECL) reagents were used to visualize the bands using an Imaging System (LI-COR Biosciences).

Immunofluorescence assay

Immunofluorescence assays were performed using antibodies against H2AX, Hsp27, and RING1B. The samples were incubated overnight at 4 °C with the primary antibodies. Next, the samples were rinsed three times with PBS and incubated with species-specific secondary antibodies.

The different uptake capacities of PRT@PPMP and PRT@PPM by cardiomyocytes were validated using immunofluorescence. PRT@PPM and PRT@PPMP were labeled with green dye.

Primary cardiomyocyte isolation

Myocardial tissue was isolated from 1–2-day-old rats. Cardiomyocyte isolation involved digestion of heart tissue with 2.5 mg/ml trypsin (Aladdin Co., Ltd., Shanghai, China) and perfusion with digestion buffer containing 2 mg/ml DNase I (Sigma, D7291). Cardiomyocytes were collected from the tissue piece suspension by centrifugation and cultured in DMEM (HyClone, C11995500BT) supplemented with 20% FBS and 40 IU/ml penicillin (Sigma, P3032).

Preparation of H-R cardiomyocyte model

Cardiomyocytes were subjected to a 12-h hypoxia treatment, followed by a 24-h reoxygenation to replicate MI/RI in vitro. Cells were cultured in a hypoxic incubator (5% CO2, 95% N2) at 37 °C. Following the standard culture method, cells were reoxygenated. Cardiomyocytes were randomly classified into five groups: (1) control group cells were incubated under normal conditions; (2) reoxygenated group cells were exposed to hypoxia for 12 h, followed by reoxygenation for 24 h; and (3–5) PRT group cells were treated with PRT, PRT@PPM, or PRT@PPMP, and then subjected to the same hypoxia and reoxygenation protocol.

Metabolite assays

Lactate and ATP levels were measured using the lactate production assay kit (Solarbio) and the ATP assay kit (Solarbio), respectively, following the manufacturer’s protocol.

Analysis of the extracellular acidification rate (ECAR)

The cells were inoculated into XF24 microporous cell culture plates. The culture medium was replaced following adherence of the cells, and the cells were washed and incubated for 60 min at 37 °C in a CO2-free incubator. Next, glucose, oligomycin, and 2-deoxyglucose were sequentially added. Measurement and software analyses were conducted using the XF glycolysis stress test report generator following the manufacturer’s protocol.

Cell viability assay

Cardiomyocytes were cultured and seeded into 96-well plates. Cell viability was assessed using the Cell counting kit-8 (CCK-8; Dojindo Molecular Technologies), following the manufacturer’s guidelines.

siRNA construction and transfection

Small interfering RNA (siRNA) sequences were synthesized by Gene Pharma (Shanghai Gene Pharma, China). The primer sequences used were listed in Table S3. Cardiomyocytes were incubated in 5 ml serum-free medium for 4–6 h. siRNA and X-treme GENE siRNA (Invitrogen, Carlsbad, CA, USA) were mixed separately with 300 μl serum-free medium. The two mixtures were combined and incubated at room temperature. Finally, the mixture was added to the cardiomyocytes and incubated at 37 °C for 24 h.

Luciferase reporter assays

The PFKFB3 3’-UTR full-length sequence was amplified. Cells were seeded in triplicate in 48-well plates. Each plate was co-transfected with 40 ng/well luciferase reporter vector and 10 pmol NF-κB mimic or mimic control using Lipofectamine 2000 (Invitrogen). Cells were lysed 24 h after transfection, and luciferase activity was determined.

Co-immunoprecipitation

Immunoprecipitation assay was performed according to the protocol described in a previous study [7]. The treated samples were incubated with control IgG or antibodies overnight at 4 °C, followed by a 4-h incubation with 50 μl protein G Sepharose. The samples were denatured, loaded onto an SDS-PAGE gel, and transferred to a nitrocellulose membrane. The membranes were then incubated with primary antibodies, followed by secondary antibodies.

Statistical analysis

Data analysis was performed using Statistical Product and Service Solutions (SPSS) 19.0 statistical software (SPSS Inc., Chicago, IL, USA) and Prism 9.0. Quantitative data are expressed as mean ± SD. Shapiro–Wilks test was used to explore whether the data were normally distributed. Brown-Forsythe test was used for group variances analysis. One-way analysis of variance (ANOVA) was used to analyze multiple group comparisons, and P < 0.05 was considered statistically significant.