Empagliflozin-pretreated mesenchymal stem cell-derived extracellular vesicles facilitate cardiac repair via AGGF1

Abstract

Background

Current treatment modalities can only reap a limited efficacy in reversing myocardial damage and promoting functional recovery in patients with myocardial infarction. Therefore, there is an urgent need to explore new therapeutic approaches to enhance cardiac repair.

Methods

MSCs were pretreated with empagliflozin to obtain EMPA-EVs, with non-pretreated MSC-EVs serving as controls. In vitro experiments were conducted to evaluate the effects of the two types of EVs on macrophage phenotypic transformation, cardiomyocyte apoptosis, and vascular endothelial cell lumen formation. In vivo experiments, a rat model of myocardial infarction was constructed, and EMPA-EVs, non-pretreated MSC-EVs, or phosphate-buffered saline were administered via myocardial injection. Post-treatment assessments included left ventricular ejection fraction, myocardial infarction area, and levels of pro-inflammatory cytokines in myocardial tissue. In addition, AGGF1 was depleted in EMPA-EVs to observe their anti-apoptotic, anti-inflammatory, and pro-angiogenic effects.

Results

Compared with non-pretreated MSC-EVs, EMPA-EVs increased the proportion of macrophages transforming into anti-inflammatory phenotypes, decreased the rate of apoptotic cardiomyocytes, and promoted the formation of vascular endothelial cell lumen. Animal experiments showed that the left ventricular ejection fraction of rats in the EMPA-EVs group was higher than that in the control group, the myocardial infarction area was reduced, and the level of pro-inflammatory factors in myocardial tissue decreased. Moreover, AGGF1 expression was significantly upregulated in EMPA-EVs, and the anti-apoptotic, anti-inflammatory, and pro-angiogenic effects of EMPA-EVs were inhibited upon AGGF1 knockdown.

Conclusion

EMPA-EVs can promote cardiac repair in myocardial infarction, and this therapeutic effect involves the upregulation of AGGF1 and may promise a cell-free therapeutic strategy.

Graphical Abstract

MSC-EVs pretreated with empagliflozin exerted myocardial protective functions after myocardial infarction at least partially by activating the AGGF1/AKT signaling pathway to synergistically inhibit myocardial cell apoptosis, regulate the transformation of macrophages to pro-repair phenotypes, and promote angiogenesis

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1186/s12951-026-04747-x.

Introduction

Cardiovascular diseases remain the leading cause of mortality globally, with the annual related deaths having increased by approximately 60% over the past three decades. In today’s global population expanding and meanwhile aging, the burden of ischemic heart disease continues to escalate, with 254 million cases reported in 2021 [1]. As the most serious manifestation of ischemic heart disease, acute myocardial infarction (AMI) typically results from sudden occlusion or abrupt reduction in coronary blood flow, and culminates in irreversible ischemic necrosis of the affected myocardium.

Timely revascularization and pharmacological interventions can partially preserve cardiac function following infarction, but cannot enable the regeneration of necrotic cardiomyocytes [2]. Stem cell therapy has emerged to overcome this issue, but its application is hindered by several events, including immune rejection, a poor survival in ischemic microenvironments, and a limited capacity of cardiomyogenic differentiation. Increasing evidence suggests that the regenerative effects of stem cells are primarily mediated by paracrine factors that exert pleiotropic effects on such processes as cardiac regeneration, metabolic homeostasis, inflammation, angiogenesis, and fibrosis [3, 4].

Among various paracrine mediators, extracellular vesicles (EVs)—nanosized, lipid bilayer-enclosed particles—have shown a mediatory effect on intercellular communication. EVs are rich in bioactive molecules derived from their parent cells, including proteins, lipids, and nucleic acids [5], and serve as key effectors in cell-to-cell signaling [6]. Notably, EVs possess low immunogenicity and cytotoxicity, a strong tissue-penetrating capacity, and a high feasibility of bioengineering modifications [7]. Accumulating evidence supports the therapeutic potential of mesenchymal stem cell-derived EVs (MSC-EVs) in the treatment of cardiovascular diseases, including atherosclerosis [8], AMI [9, 10], ischemia/reperfusion injury [11, 12], and heart failure [13, 14].

Empagliflozin (EMPA), a selective sodium-glucose cotransporter 2 inhibitor, has demonstrated robust cardioprotective and renoprotective effects beyond its glucose-lowering activity [15, 16]. The cardioprotective effect of EMPA involves molecular regulation on multiple processes, including apoptosis, oxidative stress [17], inflammation [18], autophagy [19], and myocardial fibrosis [20]. Previous studies have also reported that pharmacological pretreatment can enhance the efficacy of EVs [21–23]. These findings inspired us to systematically investigate the reparative effects of EMPA-pretreated MSC-derived EVs (EMPA-EVs) against myocardial infarction.

Methods

Cell culture and drug pretreatment

Human bone marrow-derived mesenchymal stem cells (RRID: CVCL_A9JT), rat cardiomyoblast cell line H9c2 (H9c2 (2 − 1), RRID: CVCL_0286), human umbilical vein endothelial cells (HUVECs, RRID: CVCL_2959), and mouse monocyte-macrophage cell line RAW264.7 (RRID: CVCL_0493) were obtained from commercial suppliers. All the cells used were identified and free from mycoplasma contamination before the experiment, and were cultured in accordance with the instructions. Mesenchymal stem cells (MSCs) were cultured in α-MEM (Gibco, USA) supplemented with 10% fetal bovine serum (FBS, Gibco, USA) and 1% penicillin-streptomycin. H9c2, HUVECs, and RAW264.7 cells were maintained in Dulbecco’s modified Eagle medium (DMEM, Gibco, USA) containing 10% FBS and 1% penicillin-streptomycin. Culture medium was refreshed every 1–3 days, and all cells were incubated at 37 °C in a humidified atmosphere with 5% CO₂. In the hypoxia experiment, the cells were placed in a hypoxia incubator (1% O₂, 5% CO₂, and 94% N₂) at 37 °C for subsequent culture.

In preliminary experiments, the optimal concentration and duration of EMPA (Beyotime, SD2411, purity ≥ 98%) were determined using Cell Counting Kit-8 (CCK-8; Vazyme, A311) and JC-1 (5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide, Beyotime, C2003S) staining. To induce a pro-inflammatory phenotype, RAW264.7 cells were stimulated with 1 µg/mL lipopolysaccharide (LPS; Beyotime, S1732) for 24 h. In formal experiments, the cells were treated with phosphate buffered saline (PBS), MSC-EVs (1 µg/µL), or EMPA-EVs (1 µg/µL).

Cell viability assay

Cell viability was assessed using the CCK-8 assay. MSCs were seeded into 96-well plates at a consistent density. Upon reaching 50% confluence, the cells were exposed to various concentrations of EMPA for 24–48 h. Subsequently, 10 µL of CCK-8 reagent was added into each well and incubated for 2 h at 37 °C. Absorbance was measured at 450 nm using a microplate reader.

Mitochondrial membrane potential assessment

Mitochondrial membrane potential (MMP) was evaluated using JC-1 dye. At a high MMP, JC-1 aggregates in mitochondria and emits red fluorescence; at a low MMP, JC-1 remains in a monomeric form emitting green fluorescence. Cells were seeded in confocal dishes, treated under experimental conditions, and stained with JC-1 working solution at 37 °C for 20 min. Mitochondrial polarization was evaluated according to red-to-green fluorescence ratios under a fluorescence microscope.

Additionally, the TMRE MMP assay kit (tetramethyl rhodamine ethyl ester, Beyotime, C2001S) was used to detect mitochondrial dysfunction. A reduction in TMRE fluorescence indicated MMP loss and mitochondrial permeability transition, characteristics of early apoptosis.

Identification of MSC surface markers

MSCs were analyzed by flow cytometry for surface markers. The cells were harvested and then incubated with fluorophore-conjugated antibodies (CD45-FITC, CD44-FITC, CD31-PE, CD105-PE, CD34-APC, CD73-APC; 1:100; eBioscience) at 4 °C for 30 min. Samples were analyzed using BD FACS Aria II flow cytometer (BD Biosciences, China) to confirm MSC identity.

Cell proliferation assay

Cell proliferation was measured via EdU (5-ethynyl-2′-deoxyuridine, Vazyme, C10310) incorporation. After treatment, cells were stained with EdU working solution and Apollo fluorescent dye to visualize DNA synthesis, and counterstained with Hoechst 33,342 for nuclei. The percentage of EdU-positive cells was calculated under a fluorescence microscope.

Cell migration assay

Cells were seeded in 6-well plates and cultured in the complete medium until the cell monolayer showed a 100% confluence. A straight wound was created in the center of the cell layer using a 200 µL pipette tip, and then gently washed with PBS to remove cell debris. The medium was replaced with basal medium containing drugs or corresponding controls, and the cells were further cultured for 12 h. The areas of the scratch before and after culture were compared under a microscope to assess cell migration ability.

Senescence-associated β-galactosidase Staining

After cell fixation, SA-β-Gal (senescence-associated β-galactosidase, Beyotime, C0602) staining was performed in a CO₂-free incubator at 37 °C overnight. The cumulative optical density values of the positive staining areas were observed and counted under an optical microscope to evaluate the degree of cell senescence.

Cell cycle analysis

One stage of cell cycle can be indicated by the DNA content in cells. The distribution of the cell cycle can be analyzed by propidium iodide (PI) staining combined with flow cytometry. Cells were collected and fixed with absolute ethanol, and then cell cycle staining solution (sharebio, SB-C6031) was added and incubated in the dark at 37 °C for 30 min. The flow cytometry histograms were analyzed using FlowJo software to calculate the proportions of cells in G0/G1, S, and G2/M phases.

Transcriptome sequencing and bioinformatics analysis

MSCs were divided into EMPA (500 nM) pretreatment group and vehicle (DMSO) control group, with three independent biological replicates per group. Total RNA was isolated using Magzol reagent and DNase I digestion. Samples with RNA integrity values > 8 were further processed. mRNA was enriched using Oligo (dT) magnetic beads according to the NEBNext^® Poly(A) mRNA Magnetic Isolation Module. The library was constructed through mRNA fragmentation, cDNA synthesis, adaptor ligation, and low-cycle enrichment. The final library was subjected to quality control and paired-end sequencing with a read length of 150 bp. Differentially expressed genes (DEGs) were identified using edge R, and their overall distribution was presented by volcano plots. Kyoto encyclopedia of genes and genomes (KEGG) enrichment analysis was performed on the DEGs using a cluster profiler.

Isolation, identification and uptake of EVs

MSCs were pretreated with EMPA or control until the confluence reached approximately 80%, and then cultured in EV-free complete medium for 48 h (EV-free fetal bovine serum was purchased from VivaCell). After discarding the culture supernatant, cell debris and impurities were removed by sequential differential centrifugation at 300 g for 10 min, 2000 g for 10 min, and 10,000 g for 30 min (4 °C). The supernatant was then ultracentrifuged at 100,000 g for 90 min to obtain crude EV pellets. To eliminate residual protein contaminants, the crude EVs were resuspended in PBS and washed via an additional ultracentrifugation at 100,000 g for 90 min. Finally, the purified EVs were resuspended in 100 µL PBS and stored at −80 °C. The total protein content of EVs was determined using a BCA kit (bicinchoninic acid assay, Keygen, KGP2101). The absorbance was detected at 562 nm using a microplate reader, and the total protein concentration of each sample was calculated from the standard curve. The morphology of EVs was observed by transmission electron microscopy, and total particle concentration of EVs were evaluated by nanoparticle tracking analysis after each batch of samples had been diluted to an appropriate concentration. The particle-to-protein ratio was calculated as the total particle concentration divided by the corresponding total protein concentration. The surface markers Tetraspanin-Associated Protein 1 (CD81), Cluster of Differentiation 63 (CD63), and Tumor Susceptibility Gene 101 (TSG101) of EVs were detected by Western blot.

To verify their uptake ability, EVs were labeled with the red fluorescent dye Dil (1,1’-dioctadecyl-3,3,3’,3’-tetramethylindocarbocyanine perchlorate, Beyotime, C1036), and the unbound dye was removed. Then, the EVs were co-incubated with H9c2 rat cardiomyocytes, HUVECs, and mouse monocyte - macrophage leukemia cell line RAW 264.7 cells for 24 h. Subsequently, the cytoskeleton was stained with Actin-Tracker Green-488 (Beyotime, C2201S), and the nuclei with DAPI (4′,6 - diamidino −2 - phenylindole, Beyotime, C1005). The uptake was visualized and analyzed by confocal microscopy.

Quantification of empagliflozin by UPLC-MS/MS

EMPA concentrations in culture supernatant, EMPA-MSC pellets and EMPA-EVs were quantified via validated UPLC-MS/MS MR assay. All samples were subjected to methanol extraction combined with three cycles of freeze-thaw plus ice-bath ultrasonication to fully disrupt EV lipid bilayers and release intravesicular components prior to quantification. Chromatographic and mass spectrometry parameters are summarized in Supplementary Table 1. Serial EMPA standard solutions were prepared for calibration curve establishment, and compound concentration was calculated based on validated external standard formulas. Samples with signals below the lower limit of quantification were defined as undetectable.

Apoptosis detection

H9c2 cells were seeded in 6-well plates, cultured overnight, treated with PBS or EVs and subjected to hypoxia/serum deprivation (H/SD) stimulation. The cells were collected and stained with annexin V-FITC and PI (Keygen, KGA1102). The apoptotic cell ratios were detected by flow cytometry, and the analysis was performed using FlowJo software.

Tube formation assay

To evaluate the effect of EVs on the angiogenic ability of endothelial cells, HUVECs were starved for 12 h, uniformly seeded in 24-well plates pre-coated with matrigel (absin, abs9491), and cultured in fresh medium containing PBS or EVs for 12 h. The tubes were imaged, and the tube length, branch nodes, and ring structures were analyzed using ImageJ software.

Macrophage polarization detection

To investigate the regulatory effect of EVs on macrophage polarization in an inflammatory environment, RAW264.7 cells were induced by LPS (1 µg/mL, 24 h) and treated with PBS or EVs. Anti-mouse cluster of differentiation 80 (CD80) antibody (Invitrogen, 12–0801-82) was added and incubated at room temperature in the dark for 30 min. After fixation and membrane disruption, the same incubation with anti-mouse cluster of differentiation 206 (CD206) antibody (Invitrogen, 17–2061-82) was carried out. The proportion of M1-type and M2-type macrophages were detected by flow cytometry. Un-treated RAW264.7 cells were used as M0-type control.

Total RNA extraction and real-time quantitative PCR (RT-qPCR)

Total RNA was extracted from cells using the centrifugal column method (Beyotime, R0027). cDNA was synthesized through reverse transcription using HisyGo RT Red SuperMix (Vazyme, RT101). RT-qPCR was performed using ChamQ Blue SYBR qPCR Master Mix (Vazyme, Q312). The relative gene expression levels were calculated using the 2^−∆∆Ct method. Primer information is provided in Supplementary Table 2.

Nitric oxide (NO) content detection

Inducible nitric oxide synthase (iNOS) catalyzes the production of NO that plays a crucial role in the development of inflammatory macrophages. The inflammatory level in RAW264.7 macrophages was evaluated using a NO content detection kit (Beyotime, S0021S). The cell culture supernatants were collected, centrifuged at 1000 g for 10 min, and added with Griess Reagent I and II. The absorbance was measured at 540 nm, and the concentration of nitrite, an indirect product of NO, was calculated using a standard curve.

Construction of myocardial infarction (MI) animal model and in vivo tracing of EVs

All animal experiments were approved by the Ethics Committee of Nanjing Medical University (No. IACUC-2308024). Male sprague-dawley (SD) rats aged 6–8 weeks were randomly divided into groups. A permanent ligation of the left anterior descending artery was performed to establish the MI model. The pale anterior wall of the left ventricle and weakened myocardial contraction indicated a successful modeling.

EVs were diluted to a working concentration of 1 µg/µL based on the measured protein concentration. Immediately after the surgery, 50 µL of PBS, EVs (approximately 6 × 10⁹ particles per rat), or DiR (1,1’-dioctadecyl-3,3,3’,3’-tetramethylindotricarbocyanine iodide, UELandy, D4006) labeled EVs were injected into multiple points in the myocardium at the infarct border zone. On the first day after the surgery, the rats were randomly selected from the surgery group and the sham group, and their hearts were collected, cut into slices approximately 2 mm thick, and stained with TTC (2,3,5-triphenyltetrazolium chloride, Beyotime, C0651). Red areas indicated viable myocardium, while white areas indicated infarcted regions and could indicated the success or failure of model construction. The biodistribution of DiR-EVs was tracked from day 1 to day 12 after surgery using an in vivo imaging system.

Histological analysis and cardiac function assessment

On the third day after surgery, immunofluorescence staining was performed on tissue sections to detect the pro-inflammatory macrophage marker iNOS and the pro-reparative macrophage marker cluster of differentiation 163 (CD163). Immunohistochemical staining was used to detect the expression of cytokines interleukin-1β and the pro-reparative macrophage marker CD206.

On the 28th day after surgery, M-mode echocardiography was used to record cardiac parameters and determine left ventricular ejection fraction (LVEF) and left ventricular fractional shortening (LVFS). Tissue samples were stained with Masson’s trichrome, Sirius Red, and HE to assess infarct size, fibrosis, and inflammatory cell infiltration. At the same time, the liver, spleen, lung, and kidney of rats in each group were collected, fixed with 4% paraformaldehyde, embedded in paraffin, and sectioned for HE staining to observe histopathological changes and assess the in vivo biosafety of EMPA-EVs.

Western Blot

For extracellular vesicle samples, purified EV pellets were resuspended in pre-chilled lysis buffer supplemented with protease and phosphatase inhibitor cocktails, followed by incubation on ice for 1 h. For cell samples, cells were washed twice with ice-cold PBS and lysed with the same lysis buffer on ice for 15 min. After incubation, all lysates were centrifuged at 14,000×g for 10 min at 4℃, and the supernatants were collected as total protein extracts. Protein concentration was determined using a BCA protein assay kit (Keygen, KGB2101). Total protein was mixed with loading buffer and denatured at 95℃ for 5 min. Equal amounts of total protein (30 µg) were separated by SDS-PAGE and transferred to a polyvinylidene fluoride membrane (Millipore, IEVH07850). After blocking with 5% skim milk, the following primary antibodies were added: CD81 (AF2428, Beyotime), CD63 (AF1471, Beyotime), TSG101 (AF8259, Beyotime), B-cell lymphoma-2 (Bcl-2, 26593-1-AP, Proteintech), Bcl-2-associated X protein (BAX, 50599-2-Ig, Proteintech), β-actin (20536-1-AP, Proteintech), iNOS (80517-1-RR, Proteintech), arginase-1 (ARG-1, 16001-1-AP, Proteintech), angiogenic factor 1 (AGGF1, 11889-1-AP, Proteintech), GAPDH (60004-1-Ig, Proteintech), AKT (60203-2-Ig, Proteintech), and phosphorylated AKT (p-AKT, 28731-1-AP, Proteintech). After incubation with horseradish peroxidase-labeled secondary antibodies, enhanced chemiluminescence (Vazyme, E422) was used for detection.

Protein sequencing and bioinformatics analysis

MSC-EV and EMPA-EV were isolated and extracted, then enzymatically digested for liquid chromatography-mass spectrometry analysis. DIA data were analyzed using the DIA-NN software with a non-prespecified spectral library strategy, and matched against the UniProt human protein database. Protein quantification was standardized by the MaxLFQ algorithm. Significantly differentially expressed proteins (SDPs) between groups were defined as those with fold change > 1.2, Student’s t-test P < 0.05, and Benjamini-Hochberg correction q < 0.05. The overall distribution of proteins was visualized using a volcano plot. Gene Ontology functional enrichment analysis was further conducted.

Plasmid transfection

When the cell confluence reached 60%, the human AGGF1 overexpression plasmid (AGGF1-OE, RefSeq: NM_018046.5) and empty vector negative control (NC-OE) were transfected into MSCs using riboFECT^™ CP transfection reagent. Meanwhile, small interfering RNA targeting AGGF1 (siAGGF1) and non-targeting negative control siRNA (siNC) were transfected into EMPA-preconditioned MSCs. The AGGF1-OE plasmid, siAGGF1 and their corresponding negative controls were obtained from Origene (SC127530, China) and RiboBio (China). Transfection was performed using riboFECT^™ CP transfection reagent following the manufacturer’s instructions. The culture supernatants were then collected for subsequent EV isolation. The target sequences are listed in Supplementary Table 3.

Statistics

Continuous variables were compared between two groups using the independent sample t-test. For the comparison among three or more groups, one-way analysis of variance was used. All data were expressed as mean ± standard error of the mean. Statistical analyses were performed using GraphPad Prism software, and P < 0.05 was considered statistically significant.

Results

EMPA pretreatment enhances MSCs proliferation and inhibits senescence

Flow cytometry showed high expression of CD44, CD105 and CD73 (positive rate > 95%), but not expression of CD45, CD31, or CD34 in MSCs cultured in vitro (Fig. 1a). CCK-8 and mitochondrial membrane potential detection results indicated that a treatment with 500 nM EMPA for 48 h could significantly enhance the viability of MSCs and maintain mitochondrial function (Fig. 1b-d). Therefore, this dosage was established as the standard for subsequent experiments. EdU staining and scratch assays found that compared with the control group, EMPA-pretreated MSCs (EMPA-MSCs) presented significantly stronger abilities to proliferate and migrate (P < 0.01; Fig. 1e-f, i-j). Flow cytometry cycle analysis revealed that the proportion of cells in the G0/G1 phase increased significantly in the EMPA group (Fig. 1g, k). SA-β-GAL activity detection showed that EMPA could significantly inhibit cellular senescence (Fig. 1h, l). RNA sequencing analysis indicated that in EMPA-MSCs, the differentially expressed genes (DEGs) were mainly enriched in processes such as cell cycle, senescence regulation, and DNA replication, and signaling pathways including Hippo and AKT (Fig. 1m-o). The above results consistently confirm that EMPA can promote the proliferation and inhibit the senescence of MSCs.

Fig. 1.

EMPA exhibits pro-proliferative, pro-migratory, and anti-senescence effects on MSCs. (a) Flow cytometry analysis of surface markers on MSCs. (b) CCK-8 assay for cell viability of MSCs treated with different concentrations of EMPA at 24 h and 48 h (n = 3). (c-d) JC-1 fluorescence staining for mitochondrial membrane potential and quantitative analysis of the red/green fluorescence intensity ratio to measure mitochondrial depolarization in MSCs treated with different concentrations of EMPA for 48 h (n = 3) (scale bar = 50 μm). (e, i) EdU immunostaining for cell proliferation markers and quantitative analysis of EdU-positive cell ratio in the Ctrl group and EMPA group (n = 5) (scale bar = 50 μm). (f, j) Scratch assay for cell migration ability and quantitative analysis of migration area in the Ctrl group and EMPA group (n = 5) (scale bar = 500 μm). (g, k) Flow cytometry for cell cycle distribution and quantitative analysis of cell percentages in each phase in the Ctrl group and EMPA group (n = 3). (h, l) β-galactosidase staining for senescence degree and quantitative analysis of cumulative optical density value in the Ctrl group and EMPA group (n = 5) (scale bar = 200 μm). (m) Volcano plot showing differentially expressed genes between the Ctrl group and EMPA group, with the x-axis representing log₂(fold change) and the y-axis representing -log₁₀ (P value); Red represents upregulated genes, green represents downregulated genes, and blue represents genes with no statistically significant difference. (n) Heatmap showing representative differentially expressed genes in the Ctrl group and EMPA group (red represents high expression, blue represents low expression). (o) KEGG pathway bubble chart of differentially expressed genes (x-axis represents the statistical significance of gene expression change difference, y-axis and bubble size represent the number of enriched genes, and the bubble color indicates the corresponding functional pathway name and overall classification). Continuous variables are shown as mean ± SEM. *P < 0.05, ***P < 0.001, ****P < 0.0001

Preparation and characterization of extracellular vesicles

EVs were isolated from MSCs supernatants using differential centrifugation (Fig. 2a). Transmission electron microscopy revealed that both groups of EVs exhibited a typical cup-shaped morphology, and nanoparticle size detection indicated that the size and concentration distribution of EVs were similar between two groups (Fig. 2b-c). Furthermore, no significant differences were observed in total protein concentration, and particle-to-protein ratio between MSC-EVs and EMPA-EVs (Fig. 2d-f, Supplementary Table 4). Western blot analysis showed that specific marker proteins, such as TSG101, CD63, and CD81, were expressed in both groups of samples (Fig. 2e). After incubating Dil-labeled EVs with H9c2 cells, HUVECs, and RAW264.7 cells for 24 h, respectively, the fluorescent signal was localized in the perinuclear cytoplasmic region (Fig. 2f-h), confirming that EVs had been effectively internalized by the cells. These results suggest that the EVs were successfully extracted, and they can be absorbed and taken up by target cells, and also distribute in the cytoplasm to exert biological functions.

Fig. 2.

Preparation, characterization, and cellular uptake of EVs. (a) Workflow for the preparation of MSC-EVs and EMPA-EVs by differential ultracentrifugation (created with BioGDP.com). (b) Transmission electron microscopy showing that MSC-EVs and EMPA-EVs have similar typical cup-shaped structures (Scale bars: 500 nm (upper), 100 nm (lower)). (c-d) Nanoparticle tracking analysis for particle size and concentration of MSC-EVs and EMPA-EVs. (e-f) Comparisons of protein concentration and the particle-to-protein ratio of MSC-EVs and EMPA-EVs. (g) Western blot for characteristic marker proteins of MSC-EVs and EMPA-EVs (n = 3). (h-j) Dil-EVs were co-cultured with H9c2 cells, HUVECs, and RAW264.7 macrophages for 24 h. Actin-Tracker Green was used to stain the cytoskeleton and DAPI to stain the nucleus, demonstrating successful uptake of EVs by cells (scale bar = 50 μm). Continuous variables are shown as mean ± SEM (ns: Not significant)

Quantitative UPLC-MS/MS was performed to determine EMPA concentrations in culture supernatant, EMPA-preconditioned MSCs and EMPA-EVs. The quantitative results showed that EMPA reached approximately 0.2 µg/mL in cell culture supernatant, and the intracellular EMPA level was around 0.05 ng per 10⁶ MSC cells, confirming successful drug uptake by MSCs during preconditioning. Notably, EMPA signals were undetectable and below the lower limit of quantification in EMPA-EVs (Supplementary Fig. 1a–e, Supplementary Table 5), effectively ruling out the possibility that EMPA itself directly exerts pharmacological effects.

EMPA-EVs inhibit cardiomyocyte apoptosis, promote angiogenesis, and regulate macrophage phenotypic transformation

Based on the established therapeutic efficacy ranges from in vitro cardiovascular studies (ranging from 2 to 100 µg/mL) [24, 25, 26], we selected EVs at a concentration of 50 µg/mL in cell-based experiments. Previous studies have confirmed that extracellular vesicles derived from MSCs can protect cardiomyocytes from ischemia and hypoxia by inhibiting apoptosis [27]. In our in vitro experiments, H9c2 cardiomyocytes were treated with H/SD and co-cultured with MSC-EVs or EMPA-EVs for 24 h. Flow cytometry showed that the rates of apoptotic cardiomyocytes in both MSC-EVs and EMPA-EVs groups were significantly lower than that in the PBS group (Fig. 3a-b). Western blot analysis revealed that compared with the PBS group, the expression of pro-apoptotic protein BAX was significantly down-regulated, while the expression of anti-apoptotic protein Bcl-2 was significantly up-regulated in both MSC-EVs and EMPA-EVs groups (Fig. 3c, f-g). JC-1 and TMRE detections confirmed that both MSC-EVs and EMPA-EVs groups showed significantly increased mitochondrial membrane potential, compared with the PBS group (Fig. 3d-e, h-i). Notably, EMPA-EVs were significantly superior to MSC-EVs in inhibiting apoptosis and improving mitochondrial function (P < 0.001), suggesting that EMPA-EVs have a stronger anti-cardiomyocyte apoptosis effect.

Fig. 3.

EMPA-EVs can inhibit the apoptosis of H9c2 cells and improve their mitochondrial function, while EMPA-EVs can also promote therapeutic angiogenesis. (a-b) Quantitative analysis of apoptosis levels and apoptosis rates of H9c2 cells in the control group, MSC-EVs and EMPA-EVs treatment groups under H/SD conditions by flow cytometry (n = 6). (c, f-g) Western blot and quantitative analysis of apoptosis-related protein expression levels in H9c2 cells of the control group, MSC-EVs and EMPA-EVs treatment groups (n = 3). (d, h) JC-1 fluorescence staining for quantitative analysis of mitochondrial membrane potential and the ratio of red to green fluorescence intensity as a measure of mitochondrial depolarization in the control group, MSC-EVs and EMPA-EVs treatment groups (n = 3) (scale bar = 50 μm). (e, i) TMRE staining for quantitative analysis of mitochondrial function and mean fluorescence intensity in the control group, MSC-EVs and EMPA-EVs treatment groups (DAPI staining for nucleus) (n = 3) (scale bar = 50 μm). (j, m) Quantitative analysis of EdU staining for proliferation cells and the proportion of EdU-positive cells in the control group, MSC-EVs and EMPA-EVs treatment groups under serum starvation conditions (n = 6) (scale bar = 50 μm). (k, n) Quantitative analysis of cell migration ability and migration area in the control group, MSC-EVs and EMPA-EVs treatment groups by scratch assay (n = 5) (scale bar = 200 μm). (l, o) Quantitative analysis of vascular network formation and tube length in the control group, MSC-EVs and EMPA-EVs treatment groups by tube formation assay (n = 4) (scale bar = 200 μm). Continuous variables are shown as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

Under serum starvation conditions, compared with PBS, either MSC-EVs or EMPA-EVs significantly increased the proportion of EdU-positive HUVECs (Fig. 3i, m) and enhanced cell migration ability (Fig. 3k, n). In addition, compared with PBS, EVs significantly increased the number of tube-forming junctions and grids of HUVECs and promoted the formation of tubular structures (Fig. 3l, o). Meanwhile, quantitative analysis found that EMPA-EVs were significantly superior to MSC-EVs in increasing cell proliferation, migration and angiogenesis indicators (P < 0.001), suggesting that EMPA-EVs have a stronger ability to promote angiogenesis.

RAW264.7 cells were stimulated with LPS to establish an inflammatory model. Flow cytometry showed that both EV groups significantly reduced CD80⁺ M1 macrophages and increased CD206⁺ M2 macrophages, with the most prominent effect in the EMPA-EV group (Fig. 4a-f). The opposite changes in CD80⁺ and CD206⁺ populations confirm that EMPA-EVs promote macrophage polarization toward the reparative M2 phenotype. RT-qPCR analysis revealed that compared with PBS, both EVs treatments repressed the expression of pro-inflammatory genes, including iNOS, tumor necrosis factor -α (TNF-α), and interleukin-6 (IL-6), but boosted the expression of pro-repair genes, including ARG-1 and transforming growth factor-β (TGF-β) (Fig. 4g-k). Supernatant detection showed that the release of NO after EVs treatment was significantly reduced, compared with that in the PBS group (Fig. 4l). Similarly, Western blot analysis indicated that compared with PBS, both EVs treatments significantly decreased iNOS expression and increased ARG-1 expression (Fig. 4m-o). Our experiments revealed that EMPA-EVs had a significantly stronger effect than MSC-EVs in inhibiting pro-inflammatory phenotypic transformation.

Fig. 4.

EMPA-EVs facilitate the transformation of macrophages into a reparative phenotype. (a-f) Flow cytometry detection and quantitative analysis of the proportion of CD80-positive and CD206-positive cells in RAW264.7 macrophages of the control group, MSC-EVs and EMPA-EVs treatment groups under LPS-induced stimulation (n = 3). (g-k) RT-qPCR detection of the expression of pro-inflammatory and pro-reparative related genes in macrophages of the control group, MSC-EVs and EMPA-EVs treatment groups (n = 3 for each group). (l) Kit colorimetric assay for the detection of the level of NO oxidation products in the cell supernatant of the control group, MSC-EVs and EMPA-EVs treatment groups at 12 h and 24 h (n = 3). (m-o) Western blot detection and quantitative analysis of the protein expression of pro-inflammatory (iNOS) and pro-reparative (ARG-1) related genes in the control group, MSC-EVs and EMPA-EVs treatment groups (n = 3). Continuous variables are shown as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

In summary, these in vitro experiments confirmed that EMPA-EVs are significantly superior to MSC-EVs in inhibiting cardiomyocyte apoptosis, enhancing mitochondrial function, promoting angiogenesis, and driving the transformation of macrophages to anti-inflammatory phenotypes, thus realizing more effective myocardial protection.

EMPA-EVs promotes myocardial repair through shifting macrophages from pro-inflammatory to reparative in rats

To investigate the cardioprotective role of EMPA-EVs, we established a myocardial infarction model in SD rats by permanently ligating the left anterior descending coronary artery. Thirty minutes after model establishment, the model rats were divided into the control group treated with PBS and the experimental groups injected with MSC-EVs or EMPA-EVs (Fig. 5a). Consistent with validated EV doses from past cardiovascular studies, 1–5 × 10⁸ particles [28, 29] or 5 µg per mouse [30] is the standard intramyocardial dose for myocardial infarction models, while a therapeutic dose of 10–50 µg per rat [24, 26] has been established in preclinical rat models. We therefore selected 50 µg EVs in 50 µL for intramyocardial injection, aligning with literature-validated effective doses. At 24 h after model establishment, TTC staining revealed clearly demarcated infarcted areas in the myocardial tissue, confirming the successful establishment of the model (Fig. 5b-c). Meanwhile, DiR-labeled EVs exhibited strong fluorescence signals in the infarcted myocardium, indicating that EVs can be effectively retained at the injured site after intramyocardial injection (Fig. 5d).

Fig. 5.

EMPA-EVs reprogram macrophages toward a reparative phenotype in rats. (a)Schematic diagram of the in vivo experimental process (created with BioGDP.com). (b) TTC staining to verify myocardial changes 24 h after myocardial infarction modeling in rats, red represents viable myocardial tissue and grayish-white represents necrotic myocardial tissue. (c) Quantitative analysis of myocardial infarction area in rats of the myocardial infarction group and the sham operation group (n = 3). (d) The optical imaging system in small animals showed the distribution of EVs or Dir-EVs in different groups of rats at different times after surgery. (e)Representative images of immunohistochemistry analysis for IL-1β and CD206 expression at 3 days in different groups and (h-i) quantification analysis of IOD (n = 5) (Scale bar = 50 μm). (f-g, j-k) Representative images of iNOS and CD163 immunofluorescence staining in myocardial tissues of rats from different groups on the 3rd day after surgery and the corresponding quantitative analysis of average fluorescence intensity (n = 5). Continuous variables are shown as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

At post-operation 72 h, the immunohistochemical results remained consistent. After EV treatment, the pro-inflammatory factor IL-1β in the infarct area decreased significantly, and the proportion of CD206⁺ reparative macrophages increased significantly (Fig. 5e, h-i).Meanwhile, immunofluorescence detection showed that, compared with the PBS group, the number of infiltrating iNOS⁺ pro-inflammatory macrophages in the infarcted areas decreased significantly in the MSC-EVs and EMPA-EVs groups, while the proportion of infiltrating CD163⁺ reparative macrophages increased significantly (Fig. 5f-g, j-k). Further quantitative analysis showed that the regulatory effect of EMPA-EVs on macrophage phenotypic transformation was significantly stronger than MSC-EVs, suggesting that EMPA-EVs have a superior effect in regulating the local inflammatory microenvironment of myocardial infarction and promoting the polarization of macrophages toward the reparative phenotype.

To evaluate the long-term effects of EMPA-EVs on cardiac function and tissue repair in rats with myocardial infarction, echocardiography was performed on rats in each group at post-operation 28 days. The results showed that, compared with the PBS control group, the LVEF and LVFS in the MSC-EVs and EMPA-EVs groups were significantly increased (Fig. 6a-c), indicating that EVs could effectively improve myocardial systolic function. The echocardiographic parameters for all groups are showed in Supplementary Table 6. Further Masson staining and Sirius red staining showed that the myocardial infarction area in the EVs groups was significantly smaller than that in the control group, collagen fiber deposition was significantly reduced, and fibrosis was effectively inhibited (Fig. 6d-g). Meanwhile, the range of necrotic tissue in the infarcted area was reduced, and the infiltration of inflammatory cells around the area was also significantly alleviated (Fig. 6h). Notably, quantitative comparisons showed that EMPA-EVs were significantly superior to MSC-EVs in improving cardiac function (higher LVEF and LVFS levels) and inhibiting fibrosis (less collagen deposition and smaller infarct area), indicating that EMPA can enhance the cardioprotective effect of MSC-EVs. HE staining of major organs (liver, spleen, lung, and kidney) showed no obvious inflammatory infiltration, tissue damage, or other pathological abnormalities in all treatment groups, indicating that EMPA-EVs have favorable in vivo biosafety (Fig. 6i).

Fig. 6.

EMPA-EVs reduce infarct size and enhance cardiac function in MI rats. (a) Representative images of the parasternal short-axis view of the left ventricle by echocardiography on the 28th day after surgery in sham-operated, control, MSC-EVs and EMPA-EVs treated groups. (b-c) LVEF and LVFS evaluated by echocardiography in different groups (n = 3). (d, f) Masson staining showing the degree of infarction in the border and infarct zones and quantitative analysis of the infarct area in each group (muscle fibers are red, collagen fibers are blue) (n = 3) (scale bars = 2000 μm, 200 μm). (e, g) Sirius red staining showing the degree of fibrosis in the border and infarct zones and quantitative analysis of collagen fibers in each group (muscle fibers are yellow, collagen fibers are red) (n = 3) (scale bars = 2000 μm, 200 μm). (h) Hematoxylin-Eosin staining showing the degree of inflammatory cell infiltration in the border and infarct zones (n = 3) (scale bar = 500 μm). (i) Hematoxylin-Eosin staining of major organs, including lung, liver, spleen, and kidney tissues from rats in each group (scale bar = 50 μm). Continuous variables are shown as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

Collectively, these results indicated that EMPA-EVs can transform the inflammatory microenvironment, promote macrophage phenotypic switching, and inhibit fibrosis to restore cardiac function.

AGGF1 encapsulated in EMPA-EVs At least partially mediates their cardioprotective effects

To further elucidate the specific molecular mechanisms through which EMPA-EVs promote cardiac repair, we performed 4D proteomic analyses on MSC-EVs and EMPA-EVs. Hierarchical clustering revealed that the protein profiles of EMPA-EVs were markedly distinct from those of MSC-EVs (Fig. 7a). A volcano plot demonstrated that there were 49 differentially expressed proteins between the MSC-EVs and EMPA-EVs groups (P < 0.05, fold change > 1.2), with 30 proteins up-regulated and 19 down-regulated (Fig. 7b). Compared with the down-regulated proteins and the overall proteome (Supplementary Fig. 2a–b), the enrichment analysis of biological processes associated with up-regulated proteins was more consistent with the anticipated research direction. Functional enrichment analysis indicated that these differentially expressed proteins were significantly enriched in biological processes such as negative regulation of inflammatory response, negative regulation of T cell activation, negative regulation of lymphocyte activation, negative regulation of leukocyte cell-cell adhesion, regulation of endothelial cell proliferation and some potentially related biological processes. (Fig. 7c). It is worth noting that AGGF1 (a vascular growth factor with a G domain and FHA domain) is a potent vascular growth factor and is gradually emerging as a new type of anti-inflammatory factor in the fields of cardiovascular and nervous systems. The function of AGGF1 is highly correlated with the biological direction we are exploring, and the expression level of this protein in EMPA-EVs is more than twice that of MSC-EVs in proteomics analysis. Western blot confirmed this expression of AGGF1 in two groups (Fig. 7d-e), which was consistent with the proteomic results.

Fig. 7.

Proteomics reveals the key role of AGGF1, and EMPA-EVs regulate macrophage transition through the AGGF1/AKT axis. (a) Heatmap showing the representative protein expression with significant differences between the MSC-EVs and EMPA-EVs groups. (b) Volcano plot showing the significantly differentially expressed proteins between the MSC-EVs and EMPA-EVs groups (threshold: 1.2-fold change, P < 0.05). (c) The biological process enrichment results for up-regulated and overall differentially expressed genes were visualized using a Gene Ontology functional enrichment bubble plot. (x-axis: The fold enrichment of differentially expressed genes, y-axis: The GO term descriptions; the size of the dots in the figure represents the number of differentially expressed proteins enriched, and the color represents the P value). (d-e) Western blot and quantitative analysis confirm that EMPA-EVs are rich in AGGF1 protein expression. (f) RT-qPCR detection of the expression of AGGF1 in control-overexpressing and AGGF1-overexpressing MSCs (n = 3). (g) RT-qPCR detection of the expression of AGGF1 in control-silenced and AGGF1-silenced EMPA-MSCs (n = 3). (h-k) Western blot analysis and quantification of AGGF1 protein levels in different groups (n = 3). (l-m) AGGF1 overexpression plasmid (AGGF1-OE) and corresponding negative control were transfected into MSCs, and the related EVs were co-cultured with RAW264.7 macrophages. The protein expression levels of AGGF1, inflammatory markers (iNOS, ARG-1), and AKT, p-AKT were detected by Western blot and quantitative analysis. (n-o) AGGF1 silencing plasmid (siAGGF1) and corresponding negative control were transfected into EMPA-MSCs, and co-cultured with macrophages. The protein expression levels of AGGF1, iNOS, ARG-1, AKT, and p-AKT were detected by Western blot and quantitative analysis. (p-q) MSC-EVs^AGGF1−OE were co-cultured with macrophages with or without AKT inhibitor, and the protein expression levels of the above proteins were detected by Western blot and quantitative analysis. (r-s) EMPA-EV^siAGGF1 were co-cultured with macrophages with or without AKT agonist, and the protein expression levels of the above proteins were detected by Western blot and quantitative analysis. Continuous variables are shown as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, ns: Not significant

To further investigate whether the highly expressed AGGF1 in EMPA-EVs mediates the anti-inflammatory, anti-apoptotic, and pro-angiogenic effects of EMPA-EVs, we first validated the efficiency of AGGF1 overexpression and knockdown in MSCs via RT‑qPCR and Western blot. RT‑qPCR analysis revealed that AGGF1 mRNA expression was significantly upregulated in the AGGF1-OE group compared with the NC-OE group (Fig. 7f). Western blot further confirmed the marked upregulation of AGGF1 protein expression in the AGGF1-OE group (Fig. 7h-i). Three independent siRNA sequences targeting AGGF1 were screened, and siAGGF1−2 with the optimal silencing efficiency was selected for subsequent experiments (decreasing AGGF1 mRNA to approximately 32% of the siNC group) (Fig. 7g). Western blot analysis further verified that siAGGF1−2 markedly downregulated AGGF1 protein expression (Fig. 7j-k). Subsequently, EVs were isolated from these genetically modified MSCs and co-cultured with macrophages, cardiomyocytes, and human umbilical vein endothelial cells (HUVECs), respectively, to explore the role of AGGF1 in EMPA-EVs-mediated cardioprotective effects.

In the LPS-induced M1-type macrophage model, Western blot results showed that compared with the control, MSC-EVs overexpressing AGGF1 (MSC-EVs^AGGF1−OE) significantly inhibited iNOS expression, increased ARG-1 level, and enhanced AKT phosphorylation. This effect was reversed by the AKT inhibitor Ipatasertib, as shown by the rebounded expression of pro-inflammatory factors (Fig. 7l-m, p-q). The anti-inflammatory effect of EMPA-EVs silencing AGGF1 (EMPA-EV^siAGGF1) was weakened, and AKT phosphorylation was inhibited (Fig. 7n-o). The anti-inflammatory effect of EMPA-EV^siAGGF1 was salvaged after treatment with AKT agonists, evidenced by a significant enhancement of AKT phosphorylation (Fig. 7r-s). In conclusion, in an inflammatory environment, EMPA-EVs turn macrophages from pro-inflammatory M1 to pro-repair M2 by activating the AKT pathway through AGGF1.

Further experiments were conducted to explore the functional effects of EVs rich in AGGF1 on cardiomyocytes and endothelial cells. MSC-EVs^AGGF1−OE significantly improved the mitochondrial function of H9c2 cells under hypoxia/serum deprivation conditions (Fig. 8a-b), and inhibited their apoptosis (Fig. 8e-f), which was further verified at the protein level, accompanied by the upregulation of a p-AKT level (Fig. 8i-j). EMPA-EV ^siAGGF1 significantly discounted the protective effect of EMPA-EVs, as displayed by their weaker mitochondrial function (Fig. 8c-d) and anti-apoptotic ability (Fig. 8g-h), as well as a lower level of p-AKT (Fig. 8k-l). MSC-EVs ^AGGF1−OE significantly enhanced the tube-forming ability (Fig. 8m-n) and proliferative activity (Fig. 8q-r) of serum-starved HUVECs, while EMPA-EV^siAGGF1 inhibited the angiogenic (Fig. 8o-p) and proliferative ability (Fig. 8s-t) of EMPA-EVs. Previous studies have found that AGGF1 activates the FAK-Src-AKT angiogenesis pathway through interacting with the functional angiogenic domains of integrins α5 and β1³¹, while AKT phosphorylation synergistically promotes the angiogenesis by enhancing endothelial cell viability and lumen-forming ability [32]. In Western blot analysis, we also found that MSC-EVs^AGGF1−OE significantly upregulated the expression of p-AKT and its downstream anti-apoptotic protein Bcl-2 (Fig. 8u-v). EMPA-EV^siAGGF1 significantly inhibited the protein levels of p-AKT and Bcl-2 (Fig. 8w-x).

Fig. 8.

EMPA-EVs inhibit cardiomyocyte apoptosis and enhance therapeutic angiogenesis through the AGGF1/AKT axis. (a-d) Quantitative analysis of mitochondrial function and mean fluorescence intensity of H9c2 cells co-cultured with MSC-EVs^AGGF1−OE, MSC-EVs^NC−OE, EMPA-EV ^siAGGF1, and EMPA-EV^siNC groups, respectively, as detected by TMRE staining (n = 5) (scale bar = 100 μm). (e-h) Quantitative analysis of cardiomyocyte apoptosis levels and apoptosis rates in each group as detected by flow cytometry (n = 4) (scale bar = 100 μm). (i-k) Western blot and quantitative analysis of AGGF1, apoptosis-related proteins, AKT, and p-AKT protein expression levels in each group (n = 3). (m-p) Tube formation assay to detect the formation of vascular networks and quantitative analysis of tube length in each group after co-culture with MSC-EVs^AGGF1−OE, MSC-EVs^NC−OE, EMPA-EVs ^siAGGF1, and EMPA-EVs^siNC groups and HUVECs (n = 5) (scale bar = 200 μm). (q-t) EdU staining to detect proliferating endothelial cells in each group and quantitative analysis (n = 4) (scale bar = 50 μm). (u-x) Western blot and quantitative analysis of AGGF1, Bcl-2, AKT, and p-AKT protein expression levels in endothelial cells of each group (n = 3). Continuous variables are shown as mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

Based on these in vitro mechanism research results, we further verified the protective effect of AGGF1 on the heart in vivo. The MI rats were randomly divided into three groups: MI + PBS, MI+EMPA-EVs^siAGGF1, and MI+EMPA-EVs^siNC.

At 28 days post-surgery, echocardiographic analysis was performed to evaluate cardiac function (Fig. 9a). Compared with the MI + PBS group, EMPA-EV^siNC treatment significantly improved LVEF and LVFS, confirming the cardioprotective efficacy of EMPA-EVs (Fig. 7b-c). Notably, knockdown of AGGF1 in EMPA-EVs (EMPA-EV^siAGGF1) markedly reversed these functional improvements. Consistently, Masson staining revealed that EMPA-EV^siNC treatment drastically reduced myocardial infarct size compared with the MI + PBS group, while this reduction was largely abolished in the EMPA-EV^siAGGF1 group (Fig. 9d-e). These in vivo data further confirm that AGGF1 is a critical functional cargo of EMPA-EVs that mediates their cardioprotective effects in the rat MI model.

Fig. 9.

AGGF1 mediates the cardioprotective effects of EMPA-EVs in myocardial infarction and workflow diagram of this research. (a) Representative images of echocardiography on the 28th day after surgery in MI + PBS group, EMPA-EV^siNC and EMPA-EV^siAGGF1groups. (b-c) LVEF and LVFS evaluated by echocardiography in different groups (n = 3). (d-e) Masson staining showing the degree of infarction in the border and quantitative analysis of the infarct area in each group (n = 3) (scale bars = 2000 μm). (f) MSC-EVs pretreated with empagliflozin exerted myocardial protective functions after myocardial infarction at least partially by activating the AGGF1/AKT signaling pathway to synergistically inhibit myocardial cell apoptosis, regulate the transformation of macrophages to pro-repair phenotypes, and promote angiogenesis

Taken together, AGGF1 encapsulated in EMPA-EVs mediate their cardioprotective effects through activating the AKT signaling pathway.

Discussion

Myocardial infarction, a cardiovascular emergency with high morbidity, disability and mortality, has a complex pathogenesis often involving acute coronary occlusion that leads to ischemic and hypoxic necrosis of cardiomyocytes, triggering cardiac dysfunction or even heart failure [33]. Existing measures, such as reperfusion and drug therapies, have significantly improved patients’ short-term prognosis [34], but post-MI issues, such as excessive inflammatory responses, massive cardiomyocyte apoptosis, and insufficient angiogenesis, still hinder cardiac repair and functional recovery [35, 36], underscoring the necessity of exploration for safe and effective new strategies.

This study systematically investigated the role of EMPA-EVs in myocardial repair, with the main findings including: (1) EMPA pretreatment enhances the proliferation of MSCs and inhibits their senescence; (2) EMPA-EVs inhibit cardiomyocyte apoptosis, promote angiogenesis, and regulate macrophage phenotypic transformation; (3) EMPA-EVs shift macrophages from pro-inflammatory to reparative in rats after MI; and (4) AGGF1 encapsulated in EMPA-EVs at least partially mediates their cardioprotective effect (Fig. 9f). These findings not only refresh the understanding of the roles of EMPA and extracellular vesicles in myocardial repair, but also provide experimental basis and theoretical support for the development of EMPA-EVs-based new therapies for myocardial infarction.

The biological activity and function of MSCs, as the primary donor cells for EVs, directly determine the quality and therapeutic efficacy of their secreted EVs. In the present work, we demonstrated that EMPA preconditioning remarkably remodels the intrinsic phenotype of MSCs: it enhances MSC proliferation and inhibits cellular senescence, accompanied by the regulation of cell cycle. Correspondingly, EMPA-EVs acquire enhanced functions: they suppress cardiomyocyte apoptosis, facilitate angiogenesis and modulate macrophage polarization, leading to improved cardioprotective effects.

Beyond their direct involvement in tissue repair, MSCs also participate in intercellular communication via EV-mediated transfer of bioactive molecules, such as growth factors, cytokines, lipids, and RNAs [5, 37]. In various pathological conditions, including brain injury [38], acute lung injury [39], inflammatory bowel disease [40], liver fibrosis [41], acute kidney injury [42], and bone injury [43], MSC-derived EVs contribute to immune modulation, tissue protection, and regenerative processes. Recent advances in EV engineering, including small molecule pretreatment [21], hypoxic preconditioning [44], nanoparticle modification [45], and modulation of EV cargo [46], can ensure a higher therapeutic efficacy and a targeted delivery [47]. Our findings suggest that by activating signaling pathways, such as Hippo and AKT, to regulate cell cycle and senescence-related genes, EMPA can be introduced to improve the in vitro expansion of MSCs and extend their active lifespan, thereby offering a possibility for large-scale production of highly active EVs.

During the early phase of MI, necrotic cardiomyocytes release damage-associated molecular patterns that can rely on Toll-like receptors to activate macrophages and polarize them toward the pro-inflammatory M1 phenotype. This polarization elevates the secretion of pro-inflammatory cytokines such as TNF-α and IL-6, along with reactive oxygen species [48]. While this response facilitates the clearance of necrotic tissue, the silence of adequate anti-inflammatory factors may sustain inflammation and impair tissue healing. In the acute and subacute stages of MI, the enrichment of IL-4 and TGF-β in the cardiac microenvironment, along with Galectin-9 secretion by damaged cardiomyocytes, promote macrophage polarization towards the reparative M2 phenotype via PI3K/AKT signaling [49]. M2 macrophages then release anti-inflammatory mediators, such as IL-10 and ARG-1, to resolve inflammation, promote angiogenesis, and remodel extracellular matrix [50]. Persistent or excessive inflammation can lead to cardiomyocyte apoptosis and infarct expansion, thereby impeding the healing process. Thus, the transition from M1 to M2 macrophages is expected to facilitate myocardial recovery. Here, our study revealed the multiple roles of EMPA-EVs in inhibiting cardiomyocyte apoptosis, promoting angiogenesis, and regulating macrophage phenotypic transformation, all rendering their advantages as a candidate therapy for myocardial repair. Compared with non-pretreated MSC-EVs, EMPA-EVs bring with comprehensive effects on the pathological processes after MI by synergistically regulating inflammatory balance, cell survival, and angiogenesis. In particular, they drive the transformation of macrophages to set up a favorable microenvironment for tissue repair, which may particularly explain their superiority over traditional MSC-EVs.

Despite ongoing advancements in clinical management of AMI, long-term cardiac recovery is still challenged by irreversible cardiomyocyte loss, microvascular dysfunction, and progressive ventricular remodeling. In the present rat model of MI, EMPA-EVs improved cardiac contractile function, reduced fibrosis, and transformed the immune microenvironment, further verifying their therapeutic potential in vivo. The increase in left ventricular ejection fraction and reduction in fibrosis area directly reflected the restorations in cardiac structure and function, while the anti-inflammatory transition of macrophage phenotypes allowed a long-term inhibition on disease progression. These results support EMPA-EVs as a novel multi-targeted therapeutic tool for MI, with anti-inflammatory, anti-apoptotic, and pro-regenerative properties.

Our results further support that AGGF1, a component enriched in EMPA-EVs, mediates cardioprotective effects by activating the AKT signaling pathway across various cell types. AGGF1 (formerly VG5Q) is a novel angiogenic regulator originally identified as the first disease-causing gene in Klippel-Trenaunay syndrome, a rare congenital vascular disorder characterized by capillary malformation, venous abnormality, and tissue hypertrophy [51]. As a potent angiogenic regulator, AGGF1 exhibits stronger pro-angiogenic activity than vascular endothelial growth factor (VEGF) in endothelial cells, and specifically binds to integrin α5β1 to trigger the downstream FAK-Src-AKT signaling cascade [31]. Mechanistically, the AGGF1/PI3K-AKT axis serves as the core signaling pathway for its biological functions, independent of the VEGF pathway [52]. Beyond angiogenesis, AGGF1 exerts therapeutic effects by activating autophagy [53], maintaining vascular integrity [52], and producing anti-inflammatory effects by suppressing the NF-κB signaling pathway [54, 55]. In cardiovascular disease models, AGGF1 acts as a critical protective factor: it attenuates myocardial infarction and ischemia-reperfusion injury by promoting angiogenesis and inhibiting cardiomyocyte apoptosis [52, 56] and ameliorates cardiac hypertrophy and heart failure by alleviating cardiomyocyte apoptosis caused by endoplasmic reticulum stress [57]. Interestingly, the therapeutic benefits of AGGF1 have also been observed in models of ischemic retinopathy [58], diabetic vascular dysfunction [59], cachexia-induced muscle wasting [60], cerebral ischemic injury [61], and pulpitis [55], advocating its broad anti-inflammatory and anti-apoptotic capabilities.

Of note, the present study focuses on EMPA-preconditioned MSC-EVs with remodeled endogenous cargo, rather than engineered EVs designed as drug delivery nanocarriers. As verified by UPLC-MS/MS, EMPA was undetectable in purified EVs, indicating that MSCs were mainly phenotypically modulated during EMPA incubation, and no intentional drug loading into EVs occurred. Functionally, the cardioprotective capacities of EMPA-EVs are at least partially mediated by the upregulation of AGGF1 in EV cargo. As widely acknowledged, therapeutic cargo loading strategies are categorized into passive loading and active loading. Passive loading relies on the spontaneous incorporation of the drug into the vesicles during the vesicle formation process, either by interacting with the parental cells or by directly co-culturing with the EVs; this approach is simple but suffers from low efficiency, poor controllability, and non-specific packaging [62]. In contrast, active loading achieves direct cargo encapsulation into isolated EVs by enhancing membrane permeability through sonication, electroporation, heat shock, or detergent treatment, with higher efficiency and better dosage control [63, 64]. Collectively, these methods aim to develop EVs as targeted drug delivery systems, which is a distinct research direction beyond the scope of the current study.

This study has certain limitations: first, commercial cell lines cannot fully simulate the physiological characteristics of primary cardiomyocytes, and the use of primary cardiomyocyte or organoid models could enhance the reliability of in vitro studies. Second, proteomics suggests the presence of pleiotropic cardioprotective proteins in EMPA-EVs, but their specific functions have not been verified, and the downstream effector molecules and regulatory networks of the AGGF1/AKT axis still require in-depth analysis. Moreover, the current research is based on animal models, and the pharmacokinetic characteristics, optimal dosage and long-term safety of EMPA-EVs in humans need to be further verified by preclinical and clinical studies.

Conclusion

EMPA-EVs exert therapeutic effects against MI by upregulating AGGF1, underscoring their potential as a cell-free therapeutic strategy. Future research should optimize their preparation and delivery to explore the translational potential of EMPA-EVs in the treatment of MI.

Abbreviations

AMI

Acute myocardial infarction

EVs

Extracellular vesicles

MSC-EVs

MSC-derived extracellular vesicles

EMPA

Empagliflozin

EMPA-EVs

EMPA-pretreated MSC-derived extracellular vesicles

HUVECs

Human umbilical vein endothelial cells

MSCs

Mesenchymal stem cells

FBS

Fetal bovine serum

CCK-8

Cell counting kit-8

LPS

Lipopolysaccharide

PBS

Phosphate buffered saline

MMP

Mitochondrial membrane potential

JC-1

5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide

TMRE

Tetramethyl rhodamine ethyl ester

CD45

Protein tyrosine phosphatase receptor type c

CD44

Cluster of differentiation 44

CD31

Platelet endothelial cell adhesion molecule −1

CD105

Endoglin

CD34

Cluster of differentiation 34

CD73

Ecto-5’-nucleotidase

EdU

5-ethynyl-2′-deoxyuridine

SA-β-Gal

Senescence-associated β-galactosidase

PI

Propidium iodide

DEGs

Differentially expressed genes

KEGG

Kyoto encyclopedia of genes and genomes

BCA

Bicinchoninic acid assay

CD81

Tetraspanin-associated protein 1

CD63

Cluster of differentiation 63

TSG101

Tumor susceptibility gene 101

Dil

1,1’-dioctadecyl-3,3,3’,3’-tetramethylindocarbocyanine perchlorate

DAPI

4′,6 - diamidino −2 - phenylindole

H/SD

Hypoxia/serum deprivation

CD80

Cluster of differentiation 80

CD206

Cluster of differentiation 206

RT-qPCR

Quantitative real-time PCR

iNOS

Inducible nitric oxide synthase

NO

Nitric oxide

SD

Sprague-dawley

DiR

1,1’-dioctadecyl-3,3,3’,3’-tetramethylindotricarbocyanine iodide

TTC

2,3,5-triphenyltetrazolium chloride

CD163

Cluster of differentiation 163

LVEF

Left ventricular ejection fraction

LVFS

Left ventricular fractional shortening

Bcl-2

B-cell lymphoma-2

BAX

Bcl-2-associated X protein

ARG-1

Arginase-1

AGGF1

Angiogenic factor 1

p-AKT

Phosphorylated AKT

EMPA-MSCs

EMPA-pretreated MSCs

TNF-α

Tumor necrosis factor-α

IL-6

Interleukin-6

TGF-β

Transforming growth factor-β

MSC-EVs^AGGF1-OE

MSC-EVs overexpressing AGGF1

EMPA-EV^siAGGF1

EMPA-EVs silencing AGGF1

Author contributions

Q.C., Q.G., and J.W. performed the experiments, analyzed the data, and wrote the original draft. L.D. and Y.D. assisted in experimental work and data collection. J.Y.W. contributed to technical support and resource provision. J.H. conducted statistical analysis. T.S. and L.P. prepared figures and assisted in data visualization. W.M., Q.W., R.W., and L.S. supervised the study, acquired funding, and reviewed and edited the manuscript. All authors reviewed and approved the final manuscript.

Funding

This study was supported by grants from National Natural Science Foundation of China [Grant No.82270328], Key Project of Jiangsu Provincial Health Commission [2024103606], Construction Project of High Level Hospital of Jiangsu Province [GSPJS202404, GSPJS202518], Changzhou High Level Medical Talents Training Project [2022CZBJ054], Young Talents Program of Wuxi Medical Center to Nanjing Medical University [WMCQ202403], General Projects of Wuxi Medical Center to Nanjing Medical University [WMCG202408], Clinical Research Project of Wuxi Medical Center to Nanjing Medical University [WMCC202505], Changzhou Talent Program [CQ20240067] and Medical Key Strategic Project of Wuxi Municipal Health Commission [2025TH-GD-SL-2025].

Data availability

No datasets were generated or analysed during the current study.

Declarations

Competing interests

The authors declare no competing interests.