Abstract Background Myocardial fibrosis is a pathological hallmark of heart failure post infarction, emphasizing the need for innovative treatment strategies. This research assesses the antifibrotic potential of a sodium alginate (SA) hydrogel loaded with extracellular vesicles (EVs) from bone marrow mesenchymal stem cells and PAP (p38α antagonistic peptides), aiming to interfere with fibrosis‐inducing pathways in myocardial tissue after infarction. Methods We induced fibrosis in mouse cardiac fibroblasts through hypoxia and disrupted the Mapk14 gene to study its contribution to fibrosis. Mesenchymal stem cell‐derived EVs, loaded with PAP, were encapsulated in the SA hydrogel (EVs‐PAP@SA). The formulation was tested in vitro for its effect on fibrotic marker expression and cell behavior, and in vivo in a murine model of myocardial infarction for its therapeutic efficacy. Results Map k14 silencing showed a decrease in the fibrotic response of cardiac fibroblasts. Treatment with the EVs‐PAP@SA hydrogel notably reduced profibrotic signaling, increased cell proliferation and migration, and lowered apoptosis rates. The in vivo treatment with the hydrogel post myocardial infarction significantly diminished myocardial fibrosis and improved cardiac performance. Conclusions The study endorses the SA hydrogel as an effective vehicle for delivering mesenchymal stem cell‐derived EVs and PAP to the heart post myocardial infarction, providing a novel approach for modulating myocardial fibrosis and promoting cardiac healing. Keywords: exosomes, fibrotic pathways, mesenchymal stem cells, myocardial fibrosis, myocardial infarction, p38α antagonistic peptides, sodium alginate hydrogel Subject Categories: Fibrosis __________________________________________________________________ Nonstandard Abbreviations and Acronyms CFs cardiac fibroblasts EDU 5‐ethynyl‐2′‐deoxyuridine EVs extracellular vesicles EVs@SA sodium alginate hydrogel loaded with extracellular vesicles EVs‐PAP extracellular vesicles loaded with p38α antagonistic peptides EVs‐PAP@SA sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides Mapk14 mitogen‐activated protein kinase 14 mBMSCs mouse bone marrow mesenchymal stem cells mBMSCs‐EVs extracellular vesicles derived from mouse bone marrow mesenchymal stem cells PAP p38α antagonistic peptide SA sodium alginate SRF serum response factor TGF‐β transforming growth factor beta TRPC6 transient receptor potential channel protein 6 Clinical Perspective. What Is New? * This study reveals that a novel sodium alginate hydrogel infused with extracellular vesicles derived from bone marrow mesenchymal stem cells and PAP (p38α antagonistic peptides) significantly reduces myocardial fibrosis in a murine model of myocardial infarction. * By targeting fibrosis‐inducing pathways and modulating profibrotic signaling, the extracellular vesicle‐PAP@sodium alginate hydrogel provides promising insights into myocardial repair mechanisms. What Question Should Be Addressed Next? * To advance clinical application, future studies should assess the long‐term safety, therapeutic efficacy, and adaptability of extracellular vesicle‐PAP@sodium alginate hydrogel in human cardiac fibrosis models, focusing on optimal delivery methods and possible impacts on arrhythmogenic risks associated with fibrotic remodeling. Cardiac fibrosis ensuing myocardial infarction (MI) represents a significant challenge in the continuum of cardiac disease management, with multifaceted repercussions documented in the literature.[32] ^1 , [33]^2 The fibrotic transformation of the myocardium not only alters cardiac architecture, manifesting as ventricular wall thickening and potential chamber dilation, but also compromises the myocardial contractile function, precipitating heart failure.[34] ^3 , [35]^4 The reversal of established fibrosis remains elusive with current therapeutic modalities, which at best, can only attenuate progression or alleviate symptoms rather than offering a cure.[36] ^5 , [37]^6 , [38]^7 Pharmacological interventions offer modest inhibition of fibrotic advancement, yet are limited in efficacy and not devoid of adverse effects.[39] ^8 Furthermore, the arrhythmogenic potential of post‐MI cardiac fibrosis adds a layer of complexity to the treatment regimen.[40] ^9 , [41]^10 , [42]^11 In recent years, regenerative medicine and tissue engineering have brought new hope for the treatment of MI. Sodium alginate (SA) has become a commonly used drug delivery system due to its advantages in long‐term storage, transportation, and use. Being a biocompatible material, it has a wide range of applications in the preparation of sustained‐release drugs and cell carriers.[43] ^12 , [44]^13 Additionally, alginate hydrogel is an effective decellularization strategy that can treat adverse cardiac remodeling and dysfunction in MI.[45] ^14 Extracellular vesicles (EVs) from stem cells have gained significant attention for their ability to maintain biological activity and undergo controlled release in pathological environments. Serving as signaling molecules for intercellular communication, they possess characteristics such as strong natural targeting, low immunogenicity, and can be taken up by cells to participate in regulating and promoting cell or tissue regeneration functions.[46] ^15 Mouse bone marrow mesenchymal stem cells (mBMSCs) have the potential for multilineage differentiation and the ability to promote tissue repair. The EVs derived from mBMSCs have been shown to regulate cell behavior, promote healing of damaged tissues, and suppress inflammation.[47] ^16 Furthermore, given the crucial role of signaling pathways in the management of MI, targeting these abnormal pathways to improve the pathological manifestations of MI is essential. Intervention in the TGF‐β (transforming growth factor beta)/SMAD (suppressor of mothers against decapentaplegic) signaling pathway can alter the expression of myocardial fibrosis markers α‐SMA (alpha‐smooth muscle actin) and collagen I genes, with p38α serving as a key mediator of TGF‐β1 action.[48] ^17 , [49]^18 , [50]^19 , [51]^20 Moreover, within fibroblasts, p38α can promote the transcriptional activity of serum response factor (SRF), induce the expression of TRPC6 (transient receptor potential channel protein 6), and activate (NFAT) nuclear factor of activated T cells (), thereby promoting cellular fibrosis.[52] ^21 , [53]^22 , [54]^23 Therefore, combining PAP with EVs (p38α antagonist peptide) may offer a new strategy for regulating fibrosis following MI.[55] ^21 , [56]^24 In this study, we used SA hydrogel loaded with EVs derived from mBMSCs and PAP to treat myocardial fibrosis. Acting as a biocompatible scaffold, the SA hydrogel loaded with EVs from mBMSCs and PAP may have regenerative effects, as well as individual benefits of anti‐inflammatory and antiapoptotic effects in heart infarct repair. This combination could synergistically enhance myocardial repair by providing structural support, promoting cell recovery, and inhibiting fibrotic pathways induction. Furthermore, we investigated whether PAP influences myocardial fibrosis post MI by modulating the TGF‐β1/SMAD and TRPC6/NFAT pathways. Through this in‐depth mechanistic study, we aim to offer more effective strategies for treating MI and establish a foundation for clinical applications. This interdisciplinary approach, combining regenerative medicine, drug delivery, and molecular biology, holds promise in bringing true healing prospects to patients with MI, improving their quality of life, reducing the risk of complications, and fostering innovation in clinical medicine. METHODS The data underlying this article will be shared on reasonable request to the corresponding author. Ethics Statement Our institution's animal experimentation guidelines were adhered to, and approval was obtained from the Institutional Animal Experiment Ethical Review Committee of Guangdong Provincial People's Hospital for all animal experiments conducted. Isolation and Culture of Neonatal Mouse Cardiac Fibroblasts The method for isolating and culturing neonatal male C57BL/6 mouse cardiac fibroblasts (CFs) is as follows: Neonatal C57BL/6 mice (213, purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd., Beijing, China) within 48 hours of birth were humanely euthanized to obtain heart tissues. The heart tissues were then cut into small pieces measuring 0.5 to 1 mm^3 using surgical scissors. These tissue fragments were digested for 10 minutes using a preheated 0.25% type IV collagenase‐trypsin EDTA solution (T4049, purchased from Sigma‐Aldrich, USA) at 37 °C. After digestion, the tissue solution was filtered through a 200‐mesh cell strainer (QN3036, purchased from Bio‐Link Co., Beijing, China) to obtain individual cells. The isolated cells were then placed in DMEM medium (11965092, purchased from Thermo Fisher, USA) supplemented with 10% FBS (10100147C, purchased from Thermo Fisher, USA) and 1% penicillin–streptomycin (15140163, purchased from Thermo Fisher, USA). The cells were cultured in a CO[2] incubator at 37 °C with 5% CO[2], and their status was monitored using an inverted microscope. After 72 hours of culture, cell viability was assessed using Trypan Blue staining (ST798, purchased from Beyotime, Shanghai, China). CFs were maintained at a density of 70% to 80% to prevent spontaneous transdifferentiation.[57] ^25 Establishment of an In Vitro Cardiac Fibroblast Fibrosis Model The purity of CFs was identified using immunohistochemistry to detect the expression of Vimentin. Sterilized and dried slides were coated with poly‐L‐lysine (ST509, purchased from Beyotime, Shanghai, China) for 30 minutes. After drying, the slides were placed in 6‐well culture plates, and 2 mL of cell suspension was added to each well. After 48 hours, the slides were washed twice with PBS for 1 minute each time. The cells were fixed with 4% paraformaldehyde (P0099, purchased from Beyotime, Shanghai, China) at room temperature for 30 minutes, then permeabilized with 0.5% Triton X‐100 (P0096, purchased from Beyotime, Shanghai, China) at room temperature for 30 minutes and washed with PBS (3 minutes×3 times). The slides were blocked with a 5% BSA blocking solution (ST023, purchased from Beyotime, Shanghai, China) at 37 °C for 20 minutes. Then, a rabbit monoclonal antibody Vimentin (5741, purchased from Cell Signaling Technology, USA), diluted 1:100, was added and incubated overnight at 4 °C in a humidified chamber. After washing with PBS, the slides were incubated with horseradish peroxidase‐labeled goat antirabbit IgG secondary antibody at room temperature for 1 hour, treated with DAB (Diaminobenzidine, P0203, purchased from Beyotime, Shanghai, China) solution for 3 to 5 minutes, counterstained with hematoxylin (ST2067, purchased from Beyotime, Shanghai, China) for 1 to 3 minutes, dehydrated, and mounted with neutral balsam. Images were visualized using a Nikon ECLIPSE Ti microscope system (Fukasawa Co., Japan).[58] ^26 Surface marker identification: CFs were incubated with the primary antibody Vimentin (ab209446, Abcam, UK) at 4 °C for 30 minutes, washed with PBS, and the surface marker expression was analyzed using the FACScan flow cytometry system (The FACSCalibur System, USA).[59] ^27 Cell Transfection The lentiviral packaging service was provided by Sangon Biotech (Shanghai, China). The pHAGE‐puro series plasmids and helper plasmids pSPAX2 and pMD2.G, along with the pSuper‐retro‐puro series plasmids and helper plasmids gag/pol and VSVG, were cotransfected into HKE293T cells (CRL‐11268, ATCC, USA). After 48 hours of cell culture, the supernatant was collected, filtered through a 0.45 μm filter, and centrifuged to collect the virus. After 72 hours, the supernatant was collected again, centrifuged, and concentrated. The 2 virus collections were then combined, and the titer was determined. When the cells reached the logarithmic growth phase, they were digested with trypsin, and 1 × 10^5 cells were seeded into each well of a 6‐well plate. After 24 hours of routine culture, with cell confluence reaching approximately 75%, the cells were infected with a medium containing an appropriate amount of packaged lentivirus (multiplicity of infection=10, with a working titer of approximately 5×10^6 TU/mL) and 5 μg/mL polybrene (Merck, TR‐1003, USA). After 4 hours of infection, an equal amount of culture medium was added to dilute the polybrene. After 24 hours of infection, the medium was replaced with a fresh medium. To construct a stable cell line, resistance screening was performed using an appropriate concentration of puromycin (Sangon Biotech, E607054, Shanghai, China) to obtain the stable transfected cell line. The silencing lentiviral sequences are listed in Table [60]S1, and the sequence with the most effective silencing was selected for subsequent experiments. Establishment of an In Vitro Cardiac Fibroblast Fibrosis Model The cultivation of CFs involved exposure to both normoxic (20% O[2]) and hypoxic (1% O[2]) conditions. To simulate myocardial injury and establish an in vitro model of cardiac fibrosis, CFs were cultured under hypoxic conditions. Primary CFs were maintained under normoxic conditions in a CO[2] incubator at 37 °C with 5% CO[2]. For the induction of cardiomyocyte fibrosis, primary CFs were cultured under hypoxic conditions in a 1% O[2], 5% CO[2], 37 °C incubator.[61] ^28 The CFs were initially cultured under normoxic conditions overnight until they adhered to the culture dish. Subsequently, various reagents were introduced to stimulate the cells, including a TGF‐β activator (SRI‐011381, SRI, HY‐100347, MedChemExpress, USA, dose: 10 μmol/L) and a TRPC6 activator (Hyperforin, Hyp, HY‐116330, MedChemExpress, USA, dose: 10 μmol/L), among others.[62] ^29 , [63]^30 Concurrently, the normal control group of CFs was cultured under normoxic conditions, while cells from the experimental group were cultured under hypoxic conditions. Follow‐up analysis and testing were performed 12 hours after administering the medication. Real‐Time Quantitative Polymerase Chain Reaction Total RNA was extracted from tissues and cells using Trizol reagent (16096020, Thermo Fisher, USA). Reverse transcription was carried out using a reverse transcription kit (RR047A, Takara, Japan) to synthesize the corresponding cDNA. The reaction system was prepared using the SYBR Premix Ex Taq II kit (DRR081, Takara, Japan), and the real‐time quantitative polymerase chain reaction (RT‐qPCR) reaction was conducted using a real‐time fluorescence quantitative PCR instrument (ABI 7500, Thermo Fisher, USA). The PCR program was designed as follows: initial denaturation at 95 °C for 30 seconds, followed by 40 cycles of 95 °C for 5 seconds and 60 °C for 30 seconds, with a final extension at 95 °C for 15 seconds and 60 °C for 60 seconds. The amplification curve was plotted. GAPDH was used as the internal reference, and all RT‐qPCR reactions were set up with 3 replicates, with the experiment repeated 3 times. The 2^−ΔΔCt method was used to quantify the fold change in gene expression between the experimental group and the control group, using the following formula: ΔΔCt=ΔCt (experimental group)−ΔCt (control group), where ΔCt=Ct (target gene)−Ct (reference gene). Ct represents the number of amplification cycles required for the fluorescence intensity to reach the set threshold, during which the amplification is in the exponential growth phase.[64] ^31 Primer design details are provided in Table [65]S2. Western Blot Tissues and cells were lysed using RIPA lysis buffer (P0013B, Beyotime, Shanghai, China) containing 1% phenylmethylsulfonyl fluoride to extract total protein. The extraction was performed according to the manufacturer's instructions. The supernatant was collected, and the total protein concentration of each sample was determined using a BCA assay kit (P0011, Beyotime, Shanghai, China), adjusting the protein concentration to 1 μg/μL. Each sample was set to a volume of 100 μL and boiled at 100 °C for 10 minutes to denature the proteins, then stored at −80 °C for future use. Based on the size of the target protein bands, an 8% to 12% SDS‐PAGE was prepared. Equal amounts of 50 μg protein samples were loaded into each lane using a micropipette, and electrophoresis was carried out at a constant voltage from 80 V to 120 V for 2 hours to separate the proteins. The proteins on the gel were then transferred to a PVDF membrane (1620177, Bio‐Rad, USA) using a constant current of 250 mA for 90 minutes. The membrane was blocked with 1× TBST containing 5% skim milk at room temperature for 1 hour, followed by washing with 1× TBST for 10 minutes. The membrane was incubated overnight at 4 °C with primary antibodies (Table [66]S3). After incubation, the membrane was washed three times with 1× TBST, each wash lasting 10 minutes. Next, the membrane was incubated at room temperature with a secondary antibody, either horseradish peroxidase‐labeled goat antirabbit IgG (ab6721, Abcam, Cambridge, UK) or goat antimouse IgG (ab205719, Abcam, Cambridge, UK), diluted at 1:5000, for 1 hour. The membrane was then washed 3 times with 1× TBST buffer at room temperature, each wash lasting 5 minutes. The membrane was immersed in ECL reagent (1705062, Bio‐Rad, USA) and incubated at room temperature for 1 minute. The liquid was removed, and the membrane was covered with plastic wrap for band exposure and imaging using the Image Quant LAS 4000C imaging system (GE, USA). The total cellular protein was normalized to GAPDH as the internal reference, and the relative expression levels of proteins were determined by calculating the ratio of the intensity of the target band to that of the reference band. Each protein's expression level was measured with 3 replicates per experimental group.[67] ^32 Apoptosis Detection in Cells Apoptosis assessment in primary mouse CFs involved the use of the Annexin V‐FITC/PI staining kit (C1062L, Beyotime, Shanghai, China). CFs were seeded into each well of a 6‐well plate at a density of 1×10^6 cells per well. Following cell collection, they were resuspended in 195 μL of Annexin V‐FITC binding solution, and subsequently, 5 μL of Annexin V/FITC solution and 10 μL of propidium iodide (PI) solution were added. The cells were then incubated at room temperature in darkness for 15 minutes. Flow cytometry analysis was conducted within a 20‐minute timeframe to determine the extent of cellular apoptosis, specifically assessing the combined proportion of apoptotic cells in the upper right and lower right quadrants.[68] ^33 5‐Ethynyl‐2′‐Deoxyuridine Assay The CFs were seeded into a 24‐well plate at a density of 1×10^5 cells per well, with triplicate wells allocated for each cell group. To achieve a concentration of 10 μmol/L, the EDU (5‐Ethynyl‐2′‐deoxyuridine) solution (ST067, Beyotime, Shanghai, China) was introduced into the culture medium, followed by incubation in a cell culture incubator for 2 hours. After removing the culture medium, the cells underwent fixation at room temperature for 15 minutes using a 4% paraformaldehyde solution in PBS. Subsequently, they were subjected to 2 PBS washes containing 3% BSA. The cells were then incubated at room temperature for 20 minutes in a PBS solution with 0.5% Triton‐100, followed by an additional two washes with PBS containing 3% BSA. A 100 μL staining solution was applied to each well and left to incubate at room temperature in the dark for 30 minutes. Following this, cell nuclei were stained with DAPI (C1002, Beyotime, Shanghai, China) for 5 minutes, covered with a lid, and subjected to random examination in 6 to 10 fields of view using a fluorescence microscope (FM‐600, Shanghai Pudan Optical Instrument Co., Ltd). Finally, the number of positive cells in each field of view was recorded. The EDU labeling rate (%) was calculated as the number of positive cells divided by the sum of positive and negative cells, multiplied by 100%. This experiment was repeated 3 times on separate occasions.[69] ^34 Cell Migration Assay In the transwell chamber (3428, Corning, USA) with an 8 μm pore size, 600 μL of culture medium enriched with 20% FBS was introduced into the lower chamber. The chamber was incubated at 37 °C for 1 hour to establish an equilibrium state. Cells subjected to different treatments were resuspended in a culture medium devoid of FBS. A concentration of 1 × 10^6 cells/μL was inoculated into the upper chamber and maintained at 37 °C with a 5% CO[2] concentration for 24 hours. After incubation, the transwell insert was carefully extracted and washed twice with PBS. The insert was subsequently fixed with 5% glutaraldehyde, stained with 0.1% crystal violet at 4 °C for 5 minutes, and then washed twice with PBS. Cells on the surface were gently removed using a cotton ball. The transwell insert was observed under a fluorescence microscope (TE2000, Nikon), with 5 random fields of view selected for photography. The average count of cells that traversed the insert was calculated for each experimental group.[70] ^35 This experiment was repeated 3 times on separate occasions. CCK‐8 Assay The cells infected with lentivirus from each group were digested and subsequently resuspended. The cell concentration was adjusted to 1 × 10^5 cells/mL, and 100 μL of this suspension was seeded per well in a 96‐well plate. Overnight incubation was carried out under standard culture conditions. The processing of cells followed the provided guidelines of the CCK‐8 kit (C0041, procured from Beyotime, Shanghai, China). Cell viability assessments were conducted at 12, 24, 36, and 48 hours after the initial incubation period using the CCK‐8 method. In each assay, 10 μL of CCK‐8 detection solution was added, and the samples were incubated at 37 °C in a 5% CO[2] incubator for 2 hours. Finally, an ELISA reader was employed to measure the absorbance at 450 nm, facilitating the calculation of cell viability.[71] ^36 Cytokine Expression Detection by ELISA Serum from each group of mice or supernatants from CFs were collected, and the expression of TNF‐α, IL‐1β (interleukin‐1beta), and collagen I in the mouse serum or CFs was detected using ELISA kits according to the manufacturer's instructions (Product numbers: E‐EL‐M3063 and E‐EL‐M0037; both from Elabscience, Wuhan, China; ab229425, Abcam, UK). Briefly, standards and samples were added to the wells and incubated to allow the proteins to bind to the immobilized antibodies. After washing, a biotinylated specific antibody was added. After a specific incubation period, the unbound biotinylated antibodies were washed away, and a horseradish peroxidase‐conjugate was added, followed by another wash. Then, 3,3′,5,5′‐Tetramethylbenzidine substrate was added. The addition of this substrate caused a blue color, which turned yellow after adding the stop solution. The optical density value was measured at 450 nm. After subtracting the value of the blank well, the expression levels of the inflammatory factors were calculated based on the standard protein concentration curve.[72] ^37 Isolation, Cultivation, and Surface Marker Identification of Mouse Bone Marrow‐Derived MSCs The isolation and cultivation of mBMSCs followed this procedure: Cervical dislocation was employed to euthanize 6‐ to 8‐week‐old male C57BL/6 mice (strain 213, procured from Beijing Vital River Laboratory Animal Technology Co., Ltd., Beijing, China). Subsequently, the hind limbs were collected and subjected to a thorough wash with 70% ethanol. The skin and muscle tissues were meticulously removed using forceps, and bone marrow was extracted while eliminating muscle and cell clusters through a 70 mm filter. The isolated cells were cultured in a DMEM medium that was supplemented with 10% FBS, 1% penicillin, and streptomycin. This culture was maintained in a 37 °C incubator with 5% CO[2]. After 3 hours, nonadherent cells were removed, and the culture medium was refreshed every 3 to 4 days. The cell condition was routinely monitored using an inverted microscope. Passage culture was performed when the cell density reached approximately 80%, using 0.25% trypsin. Upon reaching the third generation of mBMSCs, surface marker identification was carried out as follows: mBMSCs were incubated with primary antibodies targeting CD45, CD34, CD90, CD73, and CD105 (purchased from Abcam, UK) at a 1:500 dilution. This incubation occurred at 4 °C for 30 minutes. After a PBS wash, the surface marker expression was assessed using the FACScan flow cytometry system (The FACSCalibur System, USA).[73] ^27 , [74]^38 Extraction of mBMSCs‐EVs The procedure commenced by precentrifuging FBS at 200 000g for 18 hours, effectively eliminating EVs from the serum. Subsequently, cells were harvested and transferred to a DMEM medium enriched with 1% penicillin–streptomycin and 10% FBS, which had been meticulously cleared of EVs. Following a 48‐hour incubation, the culture medium underwent collection, and a subsequent centrifugation step at 3000g for 15 minutes effectively separated the cell debris. The resulting supernatant was carefully transferred to a sterile container and supplemented with the appropriate EVsQuick‐TC EVs precipitation solution (EVsQ5A‐1, procured from System Biosciences, USA). Gentle inversion or agitation of the sample tube facilitated mixing. After refrigerating the mixture at 4 °C for 24 hours, the tube was centrifuged at 1500g for 30 minutes, and the supernatant was removed. A second centrifugation at 1500g for 5 minutes facilitated the removal of the upper clear liquid, which was replaced with 100 μL of PBS. To resuspend the EVs particles, a sterile 27‐gauge needle (305 109, obtained from BD Biosciences, USA) was used to puncture the liquid 5 to 10 times. This process effectively suspended the EVs, which were then passed through a 0.2 μm filter (1784B, sourced from Sartorius, Germany) to prevent EVs aggregation and eliminate EVsQuick polymers. The determination of the total protein concentration of mBMSCs‐EVs was accomplished using the BCA protein assay kit.[75] ^39 , [76]^40 , [77]^41 Identification of mBMSCs‐EVs Adherence to the quantification assay kit's guidelines (FCET96A‐1, procured from System Biosciences, USA) was strictly maintained to precisely determine the protein concentration of EVs. The colorimetric assay was conducted at a wavelength of 405 nm, and the results were quantified employing a standard curve. To study the mBMSCs‐EVs particles, nanoparticle tracking analysis was performed. In this regard, 20 μg of mBMSCs‐EVs was dissolved in 1 mL of PBS, vigorously vortexed for 1 minute to ensure homogenous distribution, and then subjected to the ZetaView nanoparticle tracking analyzer (Particle Metrix, Germany). This analysis enabled the direct observation and measurement of the particle size distribution of EVs, thereby evaluating the average size and polymer dispersity index of the EVs. Additionally, the Zeta potential value of the EVs was determined.[78] ^42 For the observation with a transmission electron microscope, a freshly prepared mBMSCs‐EVs sample was loaded onto a carbon‐coated copper electron microscope grid in a volume of 20 μL and left for 2 minutes. Subsequently, the sample underwent negative staining using a phosphotungstic acid solution (procured from Sigma, USA, CAS number 12501‐23‐4) for 5 minutes. Afterward, it was rinsed 3 times with PBS, surplus phosphotungstic acid solution was removed using filter paper, and partial drying was allowed. The images were observed using a Hitachi H7650 transmission electron microscope (Hitachi, Japan) operating at 80 kV.[79] ^43 Surface markers on mBMSCs‐EVs were detected through Western blot analysis. Detailed procedures of Western blot experiments were referred to as outlined earlier. The mBMSCs‐EVs suspension was concentrated, and the protein concentration was assessed using the BCA assay kit. An SDS‐PAGE gel was prepared to denature and electrophorese the protein, followed by the transfer of proteins onto a transfer membrane. The detection focused on EVs‐specific marker proteins such as TSG101 (tumor susceptibility gene 101), CD9, CD63, and calnexin.[80] ^44 Synthesis of p38α Antagonist Peptide A previously documented study introduced a novel p38α antagonistic peptide known as PAP. Among the tested peptides, the peptide with the sequence Gln‐Gly‐Gln‐Val‐Val‐Ala‐Ala‐Gly‐Lys‐Ser‐Thr‐Asp‐Glu‐Gln‐Ser demonstrated the most potent inhibitory effect on p38α phosphorylation.[81] ^45 The synthesis of the p38α antagonist peptide was performed using the PS3 synthesizer (Protein Technology, USA). 9‐fluorenylmethyloxycarbonyl (Fmoc) protected amino acids, including Fmoc‐Gln, Fmoc‐Gly, Fmoc‐Val, Fmoc‐Ala, Fmoc‐Lys, Fmoc‐Ser, Fmoc‐Thr, Fmoc‐Asp, and Fmoc‐Glu (catalog numbers 8.52205, 8.52001, 8.52021, 8.52003, 8.52041, 8.52028, 8.52030, 8.52118, 8.52123; all sourced from Sigma, USA), were employed in the synthesis, along with Wang resin (16 318, procured from Sigma, USA). N, N‐dimethylformamide (DMF, PHR1553, purchased from Sigma, USA) served as the solvent for the solid‐phase chemical synthesis process. Wang resin was linked to the C‐terminus of each required amino acid during the peptide synthesis, and Fmoc protected the N‐terminus of the amino acid. The activation of the Fmoc‐protected amino acid attached to Wang resin (500 mg; 0.5 mmol) began with its initial deprotection, achieved by treatment with 20% piperidine (purchased from Sigma, USA) in DMF. Subsequently, the Fmoc group at the amino acid's N‐terminus (551 mg; 0.5 mmol) was activated using HBTU (455 mg; 1 mmol) in the presence of an NMM basic solution (0.4 mol/L). This activation led to ester formation, which was then coupled with another amino acid bound to the resin to yield a dipeptide. The remaining amino acids were added using the same approach to complete the peptide sequence. Finally, the Fmoc group was removed with 20% piperidine in DMF solvent. The peptide resin underwent washing with DMF, dichloromethane (Sigma, USA, catalog number 24856), and ethanol, followed by drying in a desiccator. To remove the resin, a mixture containing 500 mg of resin, 30 mL of trifluoroacetic acid, 1.5 mL of benzyl ether, 1.5 mL of methyl phenyl sulfide (purchased from Sigma, USA, with product numbers 8.08260, 8.01452, 8.20825), and 1.5 mL of water was stirred for 2 hours. Trifluoroacetic acid was removed using a rotary evaporator, and the peptide precipitate was obtained using anhydrous ether (200‐467‐2, procured from Nanjing Reagents, Nanjing, China). The peptide precipitate was subjected to multiple washes with anhydrous ether until complete solvent evaporation occurred. Subsequently, the resulting precipitate was dissolved in a 10% acetic acid solution (purchased from Sigma, USA; catalog number 27225) and freeze‐dried to obtain the peptide powder.[82] ^46 The Freeze–Thaw Cycle Method Loaded PAP into mBMSCs‐EVs The freeze–thaw cycle method is a simple and effective strategy for directly loading drugs into EVs. This process involves repeatedly freezing the EVs at low temperatures and then thawing them at room temperature to induce multiple ruptures and repairs of the EVsomal membrane. Through the continuous process of rupture and repair, the drugs enter the EVs, thereby achieving drug loading. The freeze–thaw cycle is a gentle process that does not damage the membrane structure of EVs, while also preserving the biological activity of the original EVs. In summary, purified mBMSCs‐EVs with a total protein concentration of 119 mg/mL and PAP at a concentration of 5000 IU/mL were mixed (final concentrations of mBMSCs‐EVs and PAP were 114.24 μg/mL and 200 IU/mL, respectively). The drug loading efficiency was calculated using the following formula: EE(%)= (Ct–Caq)/Ct, where Ct is the total amount of added PAP and Caq is the unloaded amount of PAP. The average drug loading efficiency was 4%.[83] ^47 Drug loading efficiency (%): The mixture was incubated at room temperature for 30 minutes, rapidly frozen at −70 °C, and then thawed at room temperature (this cycle was repeated 5 times). The resulting samples were centrifuged at 1789g for 8 minutes at 4 °C using an Amicon filter (Amicon Ultra‐15, with a maximum molecular weight of 100 kDa, UFC9100, purchased from Sigma, USA) to separate the EVs loaded with PAP (EVs‐PAP) from the unloaded PAP. The integrity of EVs‐PAP was characterized through nanoparticle tracking analysis, transmission electron microscope, and Western blot experiments.[84] ^47 , [85]^48 Preparation of Seaweed Saline Gel and Encapsulation of EVs‐PAP It has been established that biocompatible nanomaterials based on alginate demonstrate both biocompatibility and nonimmunogenicity. Furthermore, clinical trials have substantiated their safety and efficacy in the prevention of ventricular remodeling. This bionanomaterial holds promise for integration with EVs‐based treatments. To initiate the process, SA powder (S278630, Aladdin, Shanghai, China) was dissolved in PBS to create a 2% (wt/vol) SA solution. Subsequently, the collected EVs‐PAP or EVs were introduced into the alginate solution and thoroughly mixed. A 1% calcium chloride solution (C5670, Sigma, USA) was added in a volume ratio of 1:4. The resulting mixture was incubated at 37 °C for 10 minutes, leading to the formation of 2 distinct types of SA hydrogels: one containing EVs‐PAP (EVs‐PAP SA, EVs‐PAP@SA) and the other containing EVs (EVs SA, EVs@SA). The drug loading content in SA hydrogel was determined to be approximately 90% using a spectrophotometer.[86] ^49 Characterization and detection of EVs‐PAP@SA were carried out using a Fourier Transform Infrared Spectrometer and a scanning electron microscope. Initially, the EVs@SA and EVs‐PAP@SA hydrogels were frozen and then subjected to freeze‐drying for 48 hours. A frozen, dried sample (1 mg) was mixed with 80 mg of dried KBr (product number 221864, Sigma, USA). This mixture was subsequently ground into a fine powder and pressed into tablets. The resulting film was inserted into the infrared analyzer (Nicolet 6700, Thermo Fisher, USA) for scanning. The infrared spectrum was recorded using a single beam absorption mode with specific parameters, including a resolution of 4 cm^−1, wavelength range of 4000 to 400 cm^−1, scan speed of 0.15 cm/s, and 128 scans. During scanning, the air sample was purged with pure helium at a rate of 5 mL/min. Scanning electron microscope analysis involved several steps: The water gel and encapsulated EVs were examined using a scanning electron microscope (S‐4800, Hitachi, Japan) operated at an accelerated voltage of 3 kV. The sample was placed on the scanning electron microscope sample stub, mounted with conductive tape, and sputter‐coated for 60 seconds using a sputter coater (Cressington Scientific Instruments, Watford, UK) before observation. An in vitro drug release experiment was conducted, where a hydrogel containing 80 μg of EVs (100 μL) was immersed in 200 μL of PBS and incubated at 37 °C in a 5% CO[2] environment. The supernatant was replaced with fresh PBS every 2 days, and the number of released EVs into the culture medium was determined using the Bradford protein assay kit (P0006, Beyotime, Shanghai, China).[87] ^50 , [88]^51 PKH67 Tracing Experiment EVs were subjected to labeling with the membrane‐labeled dye PKH67 green fluorescence (HR8569, Bio‐Rad, Beijing, China) for the purpose of evaluating their uptake by fibroblasts, specifically CFs. A mixture comprising 10 μg/mL of EVs (1 mL) and 4 μL of PKH67 was prepared and incubated at room temperature for a duration of 15 minutes. Subsequently, the staining process was halted by the addition of 1 mL of 5% BSA. The mixture underwent centrifugation at 110 000g at 4 °C for 1 hour to facilitate the separation of the supernatant. The labeled EVs were then resuspended in prechilled PBS. CFs were cocultured with these labeled EVs for a duration of 24 hours. Following this coculture, the cells were fixed using 4% paraformaldehyde and subsequently stained with a 10 μg/mL DAPI staining solution (C1025, Beyotime, Shanghai, China) for 10 minutes. The uptake of EVs, which had been labeled and cocultured with CFs, was observed using the Nikon Eclipse fluorescence microscope (Nikon, Tokyo, Japan). Cells treated with untagged EVs emitted fluorescence and served as a negative control.[89] ^52 , [90]^53 Fluorescent Immunology Following the removal of the growth medium, the sample was subjected to 3 consecutive 2‐minute rinses with PBS. Subsequently, the sample was fixed by immersion in ice‐cold methanol at −20 °C for a duration of 30 minutes. After eliminating excess methanol, the sample underwent 3 5‐minute PBS rinses. The samples were then exposed to a 0.1% Triton X‐100 solution at room temperature for 15 minutes, followed by 3 PBS washes, each lasting 5 minutes. The procedure continued by introducing the BSA‐blocking solution, and the sample was incubated for 30 minutes. Subsequently, rabbit anti‐α‐SMA antibody (Catalog No. ab124964, diluted at 1:250, obtained from Abcam, UK), collagen I antibody (Catalog No. PA5‐95137, diluted at 1:1000, obtained from Thermo Fisher, USA), SRF antibody (Catalog No. 720240, diluted at 1:500, obtained from Thermo Fisher, USA), and TRPC6 antibody (Catalog No. PA5‐20256, diluted at 1:500, obtained from Thermo Fisher, USA) were added. The mixture was then incubated at 37 °C for 60 minutes, followed by a 5‐minute PBS wash, repeated 3 times. A goat antirabbit secondary antibody labeled with FITC was incubated at 37 °C in a dark environment for 60 minutes. Afterward, it was subjected to 3 PBS washes, each lasting 3 minutes. The sample was further incubated with DAPI staining for 10 minutes and then rinsed 3 times with PBS to eliminate any excess DAPI. Finally, 20 μL of the mounting medium was added for coverslipping, and immediate observation was conducted using a fluorescence microscope.[91] ^22 Store‐Operated Calcium Entry Measurement of Ca2^+ Influx CFs were cultured on 25 mm circular glass slides and incubated for a period ranging from 24 to 48 hours. Subsequently, they underwent washing with Ringer's solution, consisting of 145 mmol/L NaCl, 5 mmol/L KCl, 2 mmol/L CaCl[2], 1 mmol/L MgCl[2], and 10 mmol/L HEPES. The application of 5 M Fura‐2 AM (F1225, sourced from Thermo Fisher, USA) ensued, followed by a 20‐minute incubation. Within the next 20 minutes, the slides were subjected to 3 successive washes with Ringer's solution to eliminate esters. The slide was then inserted into the perfusion chamber, and the intensity of excitation light was assessed at a wavelength of 340/380 nm. Measurements were conducted employing an inverted microscope at 200× magnification, using a dual‐beam spectrophotofluorometer from Photon Technology located in Birmingham, NJ. The baseline measurement of Ca^2+ was carried out in Ca^2+ and Mg^2+ free Ringer's solution (CMF) for a duration of 3 minutes. Subsequently, Ringer's solution containing cyclopiazonic acid at a concentration of 10 μm (purchased from MedChemExpress, Shanghai, China; catalog number HY‐N6771) and EGTA at a concentration of 200 μm (purchased from MedChemExpress, Shanghai, China; catalog number HY‐D0861) was perfused for 20 minutes to establish a steady‐state condition and deplete intracellular Ca^2+. Finally, the Ca^2+‐rich Ringer's solution was introduced for a duration of 3 minutes. The quantification of calcium ion (Ca^2+) uptake was based on alterations in the intensity ratio (ΔRatio) of the excitation light at wavelengths 340/380 nm.[92] ^22 Hemolysis Test Fresh mouse whole blood containing sodium heparin as an anticoagulant was purchased, and 2.5 mL of saline with anticoagulant was added to 2 mL of the fresh whole blood, followed by gentle shaking of the tube to mix it evenly. The SA hydrogel and the gel material combined with exosomes and p38α peptide were placed in sterile centrifuge tubes containing 2 mL of saline and heated in a constant temperature incubator at 37 °C for 30 minutes. Next, 15 μL of diluted blood was added to each centrifuge tube and incubated at 37 °C for 1 hour. Finally, all the mixed tubes were centrifuged at 700g for 3 minutes, and the absorbance of the supernatant at 545 nm was measured using a UV–Vis spectrophotometer. For comparison, saline and ultrapure water without hydrogel were used as negative and positive controls, respectively. The hemolysis rate (%) was calculated using the following formula: [MATH: Hemolysis rate%=As−AnAp−An :MATH] ×100, where A [s], A [n], and A [p] represent the absorbance values of the gel, saline group, and ultrapure water group, respectively.[93] ^54 Construction of MI Mouse Model Sixty male specific pathogen‐free C57BL/6 mice aged 6 to 8 weeks (213, purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd., Beijing, China) were used in this study. They weighed between 18 and 25 g and were housed in individual cages within a specific pathogen‐free animal facility. The room was maintained under a 12‐hour light–dark cycle, with humidity levels kept between 60% to 65% and a temperature range of 22 to 25 °C. The mice had ad libitum access to food and water and underwent a 1‐week acclimatization period before the start of the experiment. Before initiation, the health status of the mice was assessed. The experimental procedures and animal protocols were approved by the Institutional Animal Care and Use Committee. The 50 mice were randomly divided into 6 groups as follows: the control group (n=10), PBS group (negative control, n=10), PAP group (injection of p38α antagonist peptide, n=10), EVs@SA group (injection of EVs encapsulated in SA hydrogel, n=10), EVs‐PAP group (injection of EVs‐PAP, n=10), and EVs‐PAP@SA group (injection of EVs loaded with p38α antagonist peptide encapsulated in SA hydrogel, n=10). Anesthesia was induced in the mice by intraperitoneal injection of sodium pentobarbital (60 mg/kg) followed by endotracheal intubation connected to a rodent ventilator for ventilation. The chest was opened to expose the heart, and the left anterior descending coronary artery was ligated using 8–0 nylon sutures. Successful induction of acute MI was confirmed by the appearance of paleness in the damaged area of the myocardium. One week post surgery, mice in the anesthesia model were evaluated for cardiac function using echocardiography, and those with left ventricular ejection fraction <55% were selected for subsequent treatment experiments. Each group of mice received the following treatments: the control group underwent open‐chest surgery without ligation and received a tail vein injection of PBS; the model control group (PBS group) received a tail vein injection of PBS after MI induction; the PAP treatment group received a tail vein injection of 200 μg/kg PAP; the EVs@SA treatment group (received a tail vein injection of an equal dose of PBS solution of empty EVs@SA; the EVs‐PAP treatment group received a tail vein injection of 200 μg/kg EVs‐PAP) (EVs‐PAP concentration was expressed as the concentration of PAP within the hydrogel); and the EVs‐PAP@SA treatment group received a tail vein injection of 200 μg/kg EVs‐PAP@SA (all subsequent EVs‐PAP@SA concentrations were expressed as the concentration of PAP within the hydrogel). The treatments were administered daily via tail vein injection, and cardiac function assessment was conducted every 7 days. After 28 days of treatment, the mice were euthanized, and subsequent biochemical experiments were performed. Echocardiogram and ECG Left ventricular systolic function was evaluated employing the Siemens Sequoia 512 ultrasound echocardiography system sourced from the United States. After anesthetizing the mice, they were maintained in a supine position, and a transducer with a frequency of 10 MHz was employed for the assessment. A range of parameters was quantified, encompassing the left ventricular (LV) ejection fraction, LV fractional shortening, tricuspid annular plane systolic excursion, LV diastolic inner dimension, LV systolic inner dimension, right ventricular area, and tibia length (TL). Concurrently, ECG was executed alongside the echocardiographic examination. The ECG was recorded over a duration of 1 minute using a dual amplifier (AD Instruments). ECG intervals, including PR, QRS, QT, and RR, were meticulously gauged and subjected to analysis via LabChart 7 software (AD Instruments). The calculation of the QTc interval was performed by dividing the QT interval (measured in milliseconds) by the square root of the RR interval (also in milliseconds), followed by dividing the resultant value by 100. The termination point of the T wave was ascertained by employing the maximum slope‐intercept method.[94] ^55 , [95]^56 Triphenyl Tetrazolium Chloride Staining Following the humane euthanasia procedure, the hearts were extracted from the mice and promptly rinsed with physiological saline. Subsequently, the hearts were subjected to freezing at −20 °C for a duration of 2 hours. The hearts were then horizontally sectioned from apex to base, generating cross‐sections of 2 mm thickness. These sections were expeditiously immersed in a 2% triphenyl tetrazolium chloride solution and placed within a water bath at 37 °C while shielded from light, and this incubation lasted for 15 to 30 minutes. To ensure thorough staining, the sections were flipped at 5‐minute intervals during incubation. Following the staining process, the specimens were preserved in 10% formaldehyde for fixation. The quantification process entailed assessing the ratio of the MI area (depicted by white staining) to the total area within the mice.[96] ^57 Hematoxylin and Eosin Staining The Sudan III staining kit obtained from Beyotime company in Shanghai, China, was employed to stain and analyze the histopathological modifications within the heart tissue. A heart tissue section was harvested and submerged in a 4% paraformaldehyde solution for fixation. Subsequent steps included dehydration, clearing, and embedding in paraffin wax. The specimen was then sliced to achieve 5 μm thick sections using a microtome. Following this, the sections underwent processes such as slide baking, dewaxing, and thorough water rinsing. Finally, they were subjected to staining with hematoxylin and eosin. The sample was rinsed with distilled water, followed by immersion in 95% ethanol, and ultimately stained with eosin. The experiment encompassed procedures like 70% hydrochloric acid ethanol differentiation, dehydration, transparency, and the application of neutral gum for slide sealing. Ultimately, the morphological alterations within mouse cardiac tissues were examined through the use of an optical microscope.[97] ^55 Masson Staining The assessment of atrial fibrosis degree used Masson's Trichrome staining kit (DC0032, Leagene Biotechnology, Beijing, China). Initially, a 4 μm‐thick tissue section underwent deparaffinization followed by water treatment. Subsequent steps involved Sudan III staining for 5 to 10 minutes, succeeded by a 5 to 15 seconds acid alcohol differentiation. After a water rinse, Masson's blue solution was applied for 3 to 5 minutes. Then, the samples underwent washing, and Sirius Red staining was administered for 5 to 10 minutes. Following another wash, the sections were treated with a phosphomolybdic acid solution for 1 to 2 minutes. Next, they were immersed in an aniline blue solution for 1 to 2 minutes. This was followed by ethanol dehydration and xylene clearing. Eventually, the sections were prepared for observation, and images were captured using an Olympus BX51 microscope (Tokyo, Japan) under 200× magnification. Subsequently, the ImagePro Plus 6.0 image analysis system was employed for analysis. For each mouse, 3 heart slices were randomly chosen from the apex to the base, and 5 fields per slice were observed under a high‐power microscope. The analysis adhered to a single‐blind principle. To quantify, researchers determined the ratio of the blue stained area to the total atrial area, referred to as collagen volume fraction.[98] ^53 , [99]^58 Sirius Red Staining Thin sections of mouse heart tissue, measuring 6 μm in thickness, were prepared. These sections underwent deparaffinization and were subjected to water treatment. Subsequently, hematoxylin staining was applied for a duration of 10 to 20 minutes. Following that, an acid differentiation solution was briefly used for a 10‐second differentiation step, and the samples were rinsed in tap water for 10 minutes. Staining was then carried out using a Sirius Red solution (365548‐5G, Sigma‐Aldrich, USA) at room temperature for 1 hour. After thorough rinsing with water, the samples underwent a process of dehydration, clearing, and mounting for observation. For each mouse, 3 random slices were selected, and within each slice, 5 randomly chosen fields of view were observed using a high‐power microscope. The semiquantitative analysis of the micro Sirius red‐positive area was conducted using ImagePro Plus 6.0 software.[100] ^26 TUNEL Detection Mouse cardiac tissue specimens were obtained and subjected to fixation using a 4% paraformaldehyde solution for a duration of 15 minutes. Subsequently, they underwent a triple wash with PBS and were permeabilized in a 0.1% Triton‐X 100 solution in PBS for 3 minutes. The cardiac tissue cells were then subjected to staining using the TUNEL staining kit (C1090, Beyotime, Shanghai, China). Specifically, a 50 μL biotin‐labeled solution was applied to the sample, which was incubated at 37 °C in the dark for 60 minutes. Following this, the sample underwent a thorough triple wash with PBS. A 0.3 mL labeling reaction stop solution was added, and the sample was washed with PBS 3 more times. Subsequently, 50 μL of streptavidin‐horseradish peroxidase working solution was introduced and incubated at room temperature for 30 minutes. The sample then underwent another triple wash with PBS. A 0.5 mL DAB chromogenic solution was added and incubated at room temperature for 5 minutes, followed by 3 additional washes with PBS. To conclude the staining process, the sample was counterstained with DAPI (10 μg/mL) for 10 minutes. Confocal microscopy was employed to capture images of each experimental group, and the apoptosis ratio for each group was determined using ImagePro Plus 6.0 software.[101] ^59 Immunohistochemical Staining The paraffin sections needed to be cooled either on ice or stored in a refrigerated environment at 4 °C before proceeding with the embedding process. Following overnight storage of the paraffin sections in a dry setting, they were transferred to an oven set at 60 °C and baked for a duration of 20 minutes. The subsequent step involved immersing the sections in xylene for 10 minutes, discarding the used xylene, and repeating the soaking process for an additional 10 minutes. The specimen was then immersed in anhydrous alcohol for 5 minutes, with a subsequent replacement of the alcohol and another 5‐minute hydration process. Subsequently, the specimen was sequentially immersed in 70% and 95% alcohol for 10 minutes each, followed by a thorough rinse with distilled water lasting 5 minutes. The slices were submerged in a citrate buffer solution with a pH of 6.0 and subjected to heating on the highest setting in a microwave for 8 minutes, after which they were allowed to cool down to room temperature. Three consecutive washes using PBS with a pH range of 7.2 to 7.6 were performed, with each wash taking 3 minutes. To inactivate endogenous enzymes, 3% H[2]O[2] was added and left to incubate at room temperature for a duration of 10 minutes, followed by an additional 3 washes with PBS, each lasting 3 minutes. The blocking solution, consisting of normal goat serum (E510009, Shenggong Biotechnology, Shanghai, China), was introduced and allowed to incubate at room temperature for 20 minutes. Subsequent to this incubation period, droplets of rabbit anti‐p38α (PA5‐17713, dilution ratio: 1:100, Thermo Fisher, USA), p‐p38α antibody (MA5‐15177, dilution ratio: 1:100, Thermo Fisher, USA), or TGF‐β1 antibody (ab215715, dilution ratio: 1:500, Abcam, UK) were added. Following an overnight incubation at 4 °C, 3 rounds of washing with PBS were carried out. After the introduction of goat antirabbit IgG secondary antibody, the mixture was incubated for 30 minutes. Subsequently, SABC (P0603, Beyotime, Shanghai, China) was introduced and allowed to incubate at room temperature for an additional 30 minutes. One drop each of DAB chromogenic reagent and chromogen was applied to the specimen and incubated for 6 minutes. Following this, the specimen was stained with hematoxylin for 30 seconds. Dehydration treatment was performed by sequentially introducing 70%, 80%, 90%, and 95% ethanol and anhydrous ethanol. Each type of ethanol was soaked for 2 minutes. Lastly, 2 rounds of clarification treatment lasting 5 minutes each were carried out using xylene. Subsequently, the samples were sealed with neutral resin. This experimental procedure was repeated 3 times, and the samples were observed under an upright microscope (BX63, Olympus, Japan).[102] ^60 , [103]^61 Sample Collection and Transcriptome Sequencing Heart tissue samples were randomly procured from mice belonging to both the PBS and EVs‐PAP@SA groups (comprising 4 mice per group). Subsequent to tissue collection, the extraction of total RNA was executed employing a Trizol reagent. The concentration of RNA samples was gauged through the assessment of the OD260/280 ratio using the Nanodrop ND‐1000 spectrophotometer (Thermo Fisher). Additionally, the concentration of RNA was determined using the Qubit RNA Assay Kit ([104]Q33221, Thermo Fisher, USA). The total RNA samples selected for downstream experiments adhered to specific criteria: RNA Integrity Number ≥7.0 and 28S:18S ratio≥1.5. Notably, the sequencing library was generated and sequenced by CapitalBio Technology, situated in Beijing, China, with each sample prepared using a total of 5 μg of RNA. To effectively eliminate ribosomal RNA from the total RNA pool, the Ribo‐Zero Magnetic Kit (MRZG12324) obtained from Epicentre, USA, was employed. Subsequently, the sequencing libraries were meticulously crafted using the NEB Next Ultra RNA Library Prep Kit (E7760S), which is fully compatible with the Illumina sequencing platform. The RNA fragments underwent fragmentation into approximately 300 base pairs fragments by leveraging the NEB Next First Strand Synthesis Reaction Buffer (5×). The process commenced with the synthesis of the first strand of cDNA, accomplished using a reverse transcriptase primer and random primers. This was followed by the synthesis of the second strand of cDNA in dUTP Mix's second strand synthesis reaction buffer (10×). Subsequently, the cDNA fragments underwent end repair, which involved the addition of a polyA tail and ligation of sequencing adapters. Post ligation of Illumina sequencing adapters, the second strand of cDNA underwent digestion with the USER Enzyme (M5508, NEB, USA), resulting in the construction of a strand‐specific library. The library DNA was further subjected to amplification, purification, and enrichment through PCR. Finally, the libraries were subjected to evaluation using Agilent 2100 and quantification using the KAPA Library Quantification Kit (kk3605, Merck, USA). The culminating step involved the execution of paired‐end sequencing through the use of the Illumina NextSeq CN500 sequencer.[105] ^62 , [106]^63 , [107]^64 Transcriptome Sequencing Data Analysis The evaluation of the quality of paired‐end reads within the raw sequencing data set was performed through the use of FastQC software version 0.11.8. Subsequently, preprocessing of the raw data was executed using Cutadapt software version 1.18, primarily to eliminate Illumina sequencing adapters and poly(A) tail sequences. Furthermore, a Perl script was employed to exclude reads containing an excess of 5% N content. Reads with a quality score >20 were selectively extracted, encompassing 70% of the bases, a process facilitated by FASTX Toolkit software, version 0.0.13. To ensure data integrity, paired‐end sequences were rectified using BBMap software. The ensuing set of filtered and high‐quality read fragments was subjected to alignment against the mouse reference genome employing HISAT2 software (version 0.7.12). Differential gene expression analysis was conducted by using the R package “edgeR,” which was applied to the mRNA read counts. Stringent filtering criteria for this analysis were established, specifically |log2FC|>1 and a P value <0.05. For further functional insights, Gene Ontology (GO) functional analysis and Kyoto Encyclopedia of Genes and Genomes pathway enrichment analysis were conducted on the pool of differentially expressed genes, leveraging the “ClusterProfiler” package within the R language. A significance threshold of P<0.05 was deemed as statistically significant. The exploration of protein–protein interaction relationships among key factors was conducted using the STRING database ([108]https://string‐db.org/) with a predefined minimum interaction score threshold set at 0.700. Visual representation of the resultant protein–protein interaction network was achieved through the use of Cytoscape 3.5.1 software, and the identification of core genes was accomplished using the integrated CytoHubba tool.[109] ^65 Statistical Analysis All statistical analyses were conducted using R (version 4.2.1) and GraphPad Prism (version 9.5.0). Quantitative data are expressed as mean±SD. The normality of each data set was assessed using the Shapiro–Wilk test, and homogeneity of variance was evaluated with Levene's test. For data sets that met the normality assumption and had a sample size of n≥6, comparisons between 2 groups were performed using a paired t test, and multiple group comparisons were conducted using 1‐way ANOVA, followed by Tukey's or Bonferroni‐adjusted post hoc tests. For data sets that did not meet the normality assumption or had a sample size of n<6, nonparametric tests were applied: the Mann–Whitney U test was used for 2‐group comparisons, and the Kruskal–Wallis test with Dunn's post hoc correction was used for multiple groups. Data from different time points were analyzed using the Friedman test. A P value <0.05 was considered statistically significant for all analyses. RESULTS Hypoxia‐Induced Fibrosis and Mapk14 Silencing Effects in CFs According to reports in the literature, MI is a contributor to global mortality. Although the short‐term mortality rate has decreased, complications following an MI continue to affect survival rates. Following MI, the extracellular matrix deposition contributes to the development of fibrotic scars, an essential process for healing acute injuries. Nevertheless, excess fibrosis could worsen disease progression and contribute to complications such as heart failure, ventricular arrhythmias, and sudden cardiac death. Hence, managing myocardial fibrosis following MI is critical in clinical practice.[110] ^18 We first isolated CFs from the hearts of neonatal mice. After 3 days of culture, optical microscopy revealed that the isolated cells exhibited a whirlpool or radiating pattern (Figure [111]S1A). Trypan blue staining showed that the cell viability exceeded 95% (Figure [112]S1B), and immunohistochemical staining demonstrated that more than 95% of the CFs expressed the vimentin protein marker (Figure [113]S1C). Additionally, flow cytometry results indicated that the percentage of vimentin‐positive cells reached 95.9% (Figure [114]S1D). In addition, >95% of CFs cells expressed the fibroblast markers α‐SMA and collagen I protein (Figure [115]S1E and [116]S1F). These results confirm that we successfully isolated and cultured CFs from mouse hearts in vitro. Subsequently, cardiac fibrosis was induced in CFs under hypoxic conditions. The results from the CCK‐8 assay, EDU staining, and Transwell assay showed a decrease in cell viability (Figure [117]S1G), proliferation ability (Figure [118]S1H), and migration ability (Figure [119]S1I) of CFs under hypoxic conditions. Furthermore, the flow cytometry analysis using Annexin V/PI double staining indicated an increase in the apoptosis rate of CFs under low oxygen conditions (Figure [120]S1J). Research has demonstrated that the occurrence of myocardial fibrosis is accompanied by the secretion of a considerable quantity of inflammatory factors.[121] ^66 Thus, we measured the expression of inflammatory factors in the supernatant of the cell culture medium using an ELISA kit. The findings demonstrated an increase in the expression of TNF‐α and IL‐1β inflammatory factors under hypoxic conditions (Figure [122]S1K). According to the literature, the expression of the Mapk14 gene is increased in MI mice,[123] ^67 and inhibiting the expression of the Mapk14 gene can prevent the differentiation of CFs into myofibroblasts, thereby inhibiting myocardial fibrosis.[124] ^21 Therefore, we detected the expression of the Mapk14 gene and the myocardial fibrosis marker α‐SMA encoding gene Acta2 by RT‐qPCR. The results showed that the expression of Mapk14 and Acta2 genes was increased in CFs cultured under hypoxic conditions (hypoxia group) (Figure [125]1A). In addition, Western blot results showed that the expression of p38α and its phosphorylated form p‐p38α increased in CFs cultured under hypoxic conditions, and the expression of myocardial fibrosis markers α‐SMA and collagen I was also significantly upregulated (Figure [126]1B). The expression of collagen I in the cell culture medium was detected by ELISA, and the results showed that collagen I expression was also significantly upregulated in CFs cultured under hypoxic conditions (Figure [127]1C). Figure 1. Influence of hypoxia induction and silencing or overexpression of the Mapk14 gene on CFs. Figure 1 [128]Open in a new tab A, RT‐qPCR detection of changes in Mapk14 and Acta2 mRNA expression in CFs after hypoxia induction; B, Western blot detection of p38α, p‐p38α, α‐SMA, and collagen I protein expression in CFs after hypoxia induction; C, ELISA detection of collagen I expression in the cell culture medium; D, RT‐qPCR detection of Mapk14 mRNA expression in CFs after Mapk14 gene silencing or overexpression; E, Western blot detection of p38α and p‐p38α protein expression in CFs after Mapk14 gene silencing or overexpression. Quantitative data in the figure are presented as mean±SD, with each cell experiment repeated 3 times. * indicates P<0.05 compared with the normal or sh‐NC group; # indicates P<0.05 compared with the oe‐NC group; F, CCK‐8 assay to detect the effect of Mapk14 silencing or overexpression on CF viability under hypoxic conditions; G, EDU assay to detect the effect of Mapk14 silencing or overexpression on CF proliferation under hypoxic conditions (scale bar: 25 μm); H, Transwell assay to detect the effect of Mapk14 silencing or overexpression on CF migration (scale bar: 25 μm); I, Annexin V/PI double staining flow cytometry to detect apoptosis in CFs under hypoxic conditions after Mapk14 silencing or overexpression. The bar chart shows the apoptosis rate in Q2 and Q3 regions; J, Immunofluorescence staining to detect α‐SMA protein expression in CFs under hypoxic conditions after Mapk14 silencing or overexpression. Green fluorescence indicates α‐SMA protein, and blue fluorescence indicates DAPI nuclear staining (scale bar: 25 μm); K, Immunofluorescence staining to detect collagen I protein expression in CFs under hypoxic conditions after Mapk14 silencing or overexpression. Red fluorescence indicates collagen I protein, and blue fluorescence indicates DAPI nuclear staining (scale bar: 25 μm); L, ELISA detection of collagen I expression in the cell culture medium. The comparison between the 2 groups was conducted using a Mann–Whitney U test, while multiple groups were analyzed using Kruskal–Wallis test. Data from different time points were analyzed using Friedman test. The quantitative data are represented as mean±SD, and each cell experiment was repeated 3 times (n=3). Compared with sh‐NC group, *P<0.05, **P<0.01; compared with oe‐NC group, ^# P<0.05, ^## P<0.01. α‐SMA indicates alpha‐smooth muscle actin; CF, cardiac fibroblasts; EDU, 5‐Ethynyl‐2′‐deoxyuridine; oe‐NC, overexpression negative control; PI, propidium iodide; RT‐qPCR, real‐time quantitative polymerase chain reaction; and sh‐NC, shRNA negative control. To further explore the inhibitory effect of suppressing the expression of p38α protein on CFs fibrosis, we constructed CFs with silenced or overexpressed Mapk14 gene using plasmid transfection. The Mapk14 gene and p38α protein expression were evaluated using RT‐qPCR and Western blot analysis. The results indicated that the expression levels of the Mapk14 gene, p38α, and p‐p38α protein were reduced in CFs with silenced Mapk14 gene compared with the negative control group. Conversely, overexpression of the Mapk14 gene increased the expression of p38α and p‐p38α protein (Figure [129]1D and [130]1E). We have successfully constructed CFs expressing either silenced or overexpressed Mapk14 genes. The transfection efficiency of plasmids was assessed using RT‐qPCR and Western blot analyses, and the most efficient sequence was selected for subsequent experimental investigations (Figure [131]S1L through [132]S1O). We cultured CFs under low oxygen conditions (1% O[2], 5% CO[2]) that either silence or overexpress the Mapk14 gene. Cell viability, proliferation, and migration ability of CFs were assessed using CCK‐8, EDU staining, and Transwell assays. Overexpression of the Mapk14 gene decreased these abilities compared with the control group. Conversely, silencing the Mapk14 gene increased these abilities compared with the sh‐NC group (Figure [133]1F through [134]1H). The flow cytometry assay demonstrated that the overexpression of the Mapk14 gene promoted apoptosis of CFs, whereas silencing of the Mapk14 gene inhibited CFs apoptosis (Figure [135]1I). Moreover, immunofluorescence staining results revealed a downregulation of α‐SMA and collagen I expression in CFs with suppressed Mapk14 gene. Conversely, CFs with overexpressed Mapk14 gene showed upregulation of α‐SMA and collagen I expression (Figure [136]1J and [137]1K). The expression of collagen I in the cell culture medium was detected by ELISA. The results showed that collagen I expression was significantly downregulated in CFs with silenced Mapk14 gene, whereas it was significantly upregulated in CFs with overexpressed Mapk14 gene (Figure [138]1L). Therefore, we postulate that under hypoxic conditions, there is an increase in the expression of fibrotic factors in CFs, leading to fibrosis, decreased vitality, and suppressed proliferation and migration ability of CFs. Additionally, silencing the Mapk14 gene can inhibit fibrosis in CFs by suppressing the expression of the p38α protein, further suppressing apoptosis in CFs. EVs‐PAP@SA Hydrogel: Development and p38α Targeting Previous studies have demonstrated that EVs possess unique characteristics such as stability, low immunogenicity, and high biocompatibility, which make them an ideal drug delivery platform (Figure [139]S2A). These characteristics allow EVs to efficiently transport EVsgenous proteins and nucleic acid drugs to target cells without being recognized or cleared by the immune system.[140] ^68 Numerous studies have convincingly shown that EVs derived from MSCs can promote tissue regeneration and repair in various organs, including myocardial diseases. Furthermore, they can effectively mitigate oxidative stress and fibrosis in myocardial cells.[141] ^69 Hence, we initially extracted mBMSCs from the bone marrow of 6‐ to 8‐week‐old C57BL/6 mice. Subsequently, we observed their fibroblast‐like morphology, appearing spindle shaped, through an optical microscope (Figure [142]S2B). Furthermore, flow cytometry analysis of mBMSCs from the third generation revealed high expression levels of positive surface markers CD90 (100.0%), CD105 (99.1%), and CD73 (100%), along with low expression levels of negative markers CD34 (0.78%) and CD45 (1%) (Figure [143]S2C). These results demonstrate the successful isolation of mBMSCs. Next, we isolated the secreted EVs from the conditioned medium of mBMSCs. We observed that these EVs, called mBMSCs‐EVs, appeared as circular or elliptical membrane‐bound vesicles through transmission electron microscopy (Figure [144]S2D). Western blot analysis of EVs markers revealed an increase in CD63, CD9, and TSG101 expression in mBMSCs‐EVs compared with the cell lysate. In contrast, calnexin expression was minimal (Figure [145]S2E). The Zeta potential of mBMSCs‐EVs was detected and analyzed using a nanoparticle size analyzer, revealing a value of −9.75 mV. The diameters of the EVs ranged from 30 to 300 nm, with an average diameter of 91.8 nm. The polymer dispersity index was calculated to be 0.285 (Figure [146]S2F and [147]S2G). Fresh whole mouse blood was used to assess the hemolysis ratio (%) of SA, EVs@SA, and ‐PAP@SA, and the results indicated that the hemolysis rate in all groups remained <6%, demonstrating excellent blood compatibility of the material (Figure [148]S2H). These results suggest that we have successfully extracted and separated relatively monodisperse mBMSCs‐EVs particles. According to reports, p38α is activated through self‐phosphorylation by binding specifically with the adaptor protein TAB1 (TGF‐β activated kinase 1 binding protein 1). This activation mechanism is unique to the p38α subtype. The TAB1‐induced p38α bypass pathway contributes to cardiomyocyte apoptosis.[149] ^70 These research findings led us to consult the literature of Pei Yujun et al., who reported novel PAP (Figure [150]2A). Among them, the peptide Gln‐Gly‐Gln‐Val‐Val‐Ala‐Ala‐Gly‐Lys‐Ser‐Thr‐Asp‐GluGln‐Ser exhibits the most effective inhibition of p38α phosphorylation. This peptide achieves inhibition of p38α phosphorylation by disrupting the interaction between p38α and TAB1, thereby suppressing the activity of p38α. First, PAP was successfully synthesized using the Fmoc method. Subsequently, PAP was loaded into mBMSCs‐EVs through the freeze–thaw cycle method and named EVs‐PAP. The nanoparticle size analyzer detected and analyzed EVs‐PAP, revealing a Zeta potential of −11.39 mV. The particles exhibited a diameter ranging from 20 to 350 nm, with an average diameter of 99.3 nm. The polymer dispersity index was 0.372, as shown in Figure [151]2B and [152]2C. Compared with EVs, the Zeta potential of EVs‐PAP slightly decreases whereas the diameter slightly increases. These results indicate that EVs‐PAP possesses better integrity. The Western blot analysis demonstrated that the expression levels of CD63, CD9, and TSG101 in EVs‐PAP were nearly indistinguishable from those in EVs (Figure [153]2D). These results indicate that we have successfully constructed EVs‐PAP. Figure 2. Characterization and uptake observation of EVs‐PAP and EVs‐PAP@SA. Figure 2 [154]Open in a new tab A, Amino acid sequence of p38α antagonistic peptide; B, Zeta potential analysis of EVs‐PAP nanoparticles; C, Particle size distribution of EVs‐PAP nanoparticles; D, Expression of CD63, CD9, and TGS101 in EVs and EVs‐PAP detected by Western blot; E, Morphology of EVs‐PAP@SA and SA observed by scanning electron microscopy (white arrows indicate EVs‐PAP, scale bar: 500 nm); F, Characterization of SA and EVs‐PAP@SA by Fourier‐transform infrared spectroscopy; G, Drug release experiment of EVs‐PAP@SA; H, Uptake of PKH‐67‐labeled EVs‐PAP@SA in CFs detected by immunofluorescence staining (scale bar: 25 μm), blue represents DAPI nuclear staining, and green represents PKH‐67‐labeled EVs‐PAP@SA. The comparison between the 2 groups was conducted using a Mann–Whitney U test, and multiple groups were analyzed using Kruskal–Wallis test. The bar graphs represent the data as mean±SD, and each cell experiment was repeated 3 times (n=3). EV indicates extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; PAP, p38α antagonistic peptide; PDI, polymer dispersity index; SA, sodium alginate; and TSG101, tumor susceptibility gene 101. SA was selected as the carrier material for EVs‐PAP. Prepared SA gel was mixed with EVs‐PAP in a calcium chloride solution and stirred to induce a reaction. The resulting compound was named EVs‐PAP@SA, where the EVs‐PAP is loaded into the SA gel. The EVs‐PAP@SA was characterized using scanning electron microscopy and Fourier‐transform infrared spectroscopy. The scanning electron microscopy results confirmed the presence of EVs‐PAP in SA, as shown in Figure [155]2E. The results of Fourier infrared spectroscopy showed no difference in wavelength between EVs‐PAP@SA and SA. However, the absorption peak of EVs‐PAP@SA was stronger than that of SA, indicating the presence of more active functional groups in EVs‐PAP@SA (Figure [156]2F). The results of EVs‐PAP@SA drug detection indicate that EVs‐PAP@SA is capable of gradual PAP peptide release (Figure [157]2G). The results indicate the successful construction of self‐healing hydrogels containing PAP and EVs. To further investigate whether CFs could internalize EVs‐PAP@SA, EVs‐PAP@SA was labeled with PKH‐67 (green) and cocultured with CFs for 24 hours. Fluorescence microscopy revealed green fluorescence in CFs (Figure [158]2H), indicating the uptake of EVs‐PAP@SA by CFs. The results indicate the successful construction of EVs‐PAP@SA, which could enter cells via CF uptake. EVs‐PAP@SA Hydrogel: Reducing Hypoxia‐Induced Fibrosis To further investigate the inhibitory effect of EVs‐PAP@SA on CFs fibrosis, we conducted a 24‐hour coculture of EVs‐PAP@SA, obtained under low oxygen conditions during preparation, with CFs. Initially, CFs cells under hypoxic conditions were treated with different concentrations of EVs‐PAP@SA, and cell viability was assessed using the CCK‐8 assay. The results showed that as the concentration of EVs‐PAP@SA increased, the viability of the hypoxic CFs cells gradually increased as well. Notably, when the concentration of EVs‐PAP@SA reached 20 μg/mL, the cell viability increased nearly 4‐fold. Therefore, subsequent experiments were conducted using a concentration of 20 μg/mL (Figure [159]3A). Figure 3. Effects of EVs‐PAP@SA on CFs fibrosis. Figure 3 [160]Open in a new tab A, The effect of different concentrations of EVs‐PAP@SA on the viability of CFs cells under hypoxic conditions was assessed using the CCK‐8 assay; B, Expression changes of proinflammatory factors TNF‐α and IL‐1β in the culture supernatant of CFs after different treatments for 24 hours measured by ELISA; C, Changes in cell viability of CFs after different treatments for 24 hours measured by the CCK‐8 assay; D, Changes in proliferation capacity of CFs after different treatments for 24 hours measured by the EDU assay (scale bar: 25 μm); E, Changes in migration ability of CFs after different treatments for 24 hours measured by the Transwell assay (scale bar: 50 μm); F, Changes in apoptosis of CFs after different treatments for 24 hours detected by Annexin V/PI double staining flow cytometry. The column chart represents the apoptosis rate of cells in the Q2 and Q3 regions; G, Expression changes of α‐SMA protein in CFs after different treatments for 24 hours detected by immunofluorescence staining. Green represents α‐SMA protein, and blue represents DAPI nuclear staining (scale bar: 25 μm); H, Expression changes of collagen I protein in CFs after different treatments for 24 hours detected by immunofluorescence staining. Red represents collagen I protein, and blue represents DAPI nuclear staining (scale bar: 25 μm). I, ELISA to detect the expression of collagen I in the cell culture medium. CFs in the normal group were cultured in a 5% CO[2], 37 °C incubator, whereas CFs in other treatment groups (PBS, PAP, EVs@SA, and EVs‐PAP@SA) were cultured in a 1% O[2], 5% CO[2], 37 °C incubator to induce cardiac fibrosis. The multiple groups were analyzed using Kruskal–Wallis while data from different time points were analyzed using Friedman test. The bar graphs represent the data as mean±SD, and each cell experiment was repeated 3 times (n=3). Compared with the Normal group, ^# P<0.05, ^## P<0.01; compared with the PBS group, *P<0.05, **P<0.01. α‐SMA indicates alpha‐smooth muscle actin; CF, cardiac fibroblasts; EDU, 5‐Ethynyl‐2′‐deoxyuridine; EV, extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; IL‐1β, interleukin‐1beta; OD, optical density; PAP, p38α antagonistic peptide; PI, propidium iodide; SA, sodium alginate; and TNF‐α, tumor necrosis factor‐alpha. Inflammatory factors in the supernatant of the cell culture medium were detected using an ELISA kit. The results indicated that EVs‐PAP@SA inhibited the expression of TNF‐α and IL‐1β inflammatory factors in CFs under low oxygen conditions (Figure [161]3B). Through experimentation involving EDU staining, Transwell assay, flow cytometry, and CCK‐8 assay, we have discovered a decrease in the viability, proliferation, and migration capabilities of CF cells in the PBS group under hypoxic conditions compared with the Normal group. Compared with the PBS group, treatment with EVs‐PAP@SA improves the cell viability, proliferation capacity, and migration ability of CFs during hypoxic conditions (Figure [162]3C through [163]3E). The results obtained from Annexin V/PI double staining flow cytometry demonstrated that EVs‐PAP@SA effectively inhibited apoptosis of CFs in hypoxic conditions (Figure [164]3F). Furthermore, EVs‐PAP@SA exhibited a stronger effect in enhancing the survival of CFs in hypoxic conditions compared with PAP and EVs@SA. The results suggest that EVs‐PAP@SA could enhance the survival of CFs in hypoxic conditions. Furthermore, immunofluorescence staining results demonstrated that the expression of α‐SMA and collagen I proteins in CFs was elevated under hypoxic conditions compared with the normal control group. Compared with the PBS group, the EVs‐PAP@SA group decreased the expression of α‐SMA and collagen I proteins (Figure [165]3G and [166]3H). The expression of collagen I in the cell culture medium was detected by ELISA. The results showed that collagen I protein expression was upregulated in CFs compared with the normal control group. However, compared with the PBS group, collagen I protein expression was decreased in the EVs‐PAP@SA group (Figure [167]3I). In conclusion, our study demonstrated that EVs‐PAP@SA could effectively alleviate hypoxia‐induced fibrosis in CFs and enhance their survival during hypoxic conditions. EVs‐PAP@SA: Improving Cardiac Function and Reducing Fibrosis in MI Mice To investigate the in vivo effects of EVs‐PAP@SA on myocardial fibrosis in mice, we first established an MI mouse model, followed by treatment with EVs‐PAP@SA via tail vein injection (Figure [168]S3A). Before conducting animal experiments, we verified the hemocompatibility of the material, with exo‐PAP@SA showing extremely low hemolysis rates, confirming its excellent blood compatibility (Figure [169]S2H). After 28 days of treatment, electrocardiogram results showed that, compared with the normal control group, MI mice exhibited significantly increased RR interval, PR interval, QSR interval, and corrected QT interval (QTc) (Figure [170]S3B and [171]S3C). Compared with the PBS group, the EVs‐PAP@SA treatment group showed significantly reduced RR interval, PR interval, QSR interval, and QTc (Figure [172]4A). Echocardiography results indicated that, compared with the normal control group, MI mice had reduced LV ejection fraction (LV ejection fraction %) and fractional shortening (LV fractional shortening %). In terms of cardiac structure, MI mice showed increased corrected LV end‐diastolic diameter (LV diastolic inner dimension/TL) and corrected LV end‐systolic diameter (LV systolic inner dimension/TL), indicating structural and functional damage to the heart. Compared with the PBS group, the EVs‐PAP@SA treatment group showed increased LV ejection fraction % and LV fractional shortening %, whereas LV diastolic inner dimension/TL and LV systolic inner dimension/TL were decreased. Additionally, compared with the PAP, EVs‐PAP, and EVs@SA treatment groups, the EVs‐PAP@SA treatment group exhibited reduced left ventricular injury and significantly improved cardiac contractile function (Figure [173]4B and [174]4C). This indicates that EVs‐PAP@SA treatment improved cardiac function in mice. Figure 4. Influence of EVs‐PAP@SA on the cardiac function of MI mice. Figure 4 [175]Open in a new tab A, Changes in the baseline RR, PR, QRS intervals, and corrected QT interval (QTc) after tail vein injection of different drugs for 28 days; B, Changes in the echocardiogram of mice in each group after tail vein injection of different drugs for 28 days; C, Changes in the baseline LVEF, LVFS, LVIDd/TL, and LVIDs/TL of mice in each group after tail vein injection of different drugs for 28 days. D, Triphenyl tetrazolium chloride staining to detect MI in mice after tail vein injection of different drugs for 28 days, quantified by the ratio of the white stained area (infarct area) to the total area (scale bar: 1 mm); E, H&E staining to detect myocardial necrosis in mice after 28 days of injection with different drugs (scale bar: 1 mm/100 μm); F, Masson staining to detect interstitial fibrosis deposition in the ventricular muscle of mice from each group after 28 days of injection (myocardial cells appear red, interstitium collagen appears blue‐green), collagen volume fraction represents the ratio of blue staining area to the total atrial area (scale bar: 1 mm); G, Sirius Red staining to detect fibrosis area in mice from each group after 28 days of injection (scale bar: 50 μm); H, TUNEL staining to detect apoptosis of myocardial cells in mice from each group after 28 days of injection, red fluorescence indicates apoptotic cells, and blue fluorescence represents DAPI nuclear staining (scale bar: 50 μm). The control group mice underwent thoracotomy without ligation; the other treatment groups (PBS group, PAP group, EVs‐PAP group, EVs@SA group, and EVs‐PAP@SA group) had MI induced by ligation surgery. The multiple groups were analyzed using Kruskal–Wallis test. The quantitative data in the figures are expressed as mean±SD, with 10 mice per group (n=10). # indicates a difference compared with the control group, ^# P<0.05, ^## P<0.01; * indicates a difference compared with the PBS group, *P<0.05, **P<0.01. CVF indicates collagen volume fraction; EV, extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; H&E, hematoxylin and eosin; LVEF, left ventricular ejection fraction; LVFS, left ventricular fractional shortening; LVIDd, left ventricular diastolic inner dimension; LVIDs, left ventricular systolic inner dimension; MI, myocardial infarction; PAP, p38α antagonistic peptide; SA, sodium alginate; and TL, tibia length. To further assess the impact of EVs‐PAP@SA on myocardial fibrosis in mice, we euthanized the mice after 28 days of treatment. Mouse heart tissues were extracted, and triphenyl tetrazolium chloride staining (Figure [176]4D) and hematoxylin and eosin staining (Figure [177]4E) were performed on the tissue slices to evaluate the degree of MI and ventricular wall thickness. The results revealed that in contrast to the normal control group, the mice with MI demonstrated clear infarct areas in the heart and decreased ventricular wall thickness. Additionally, the EVs‐PAP@SA treatment group showed reduced infarct areas in the heart tissue, increased ventricular wall thickness, and a diminished degree of myocardial necrosis compared with the PBS group. Furthermore, the inhibitory effect of EVs‐PAP@SA on MI in mice is greater than that of PAP, EVs‐PAP, and EVs@SA. Subsequently, we conducted additional investigations on the potential inhibitory impact of EVs‐PAP@SA on myocardial fibrosis in mice using Masson staining, Sirius Red staining, and TUNEL staining. Masson staining results (Figure [178]S3D) demonstrated an increase in the deposition area of interstitial fibrosis in the LV myocardium of MI mice, along with a notable decrease in the thickness of the ventricular wall when compared with the normal control group. Moreover, compared with the PBS group, the EVs‐PAP@SA treatment group exhibited a reduction in the deposition area of interstitial fibrosis in the left ventricular myocardium, a decrease in the collagen volume fraction value (the ratio of the deposition area of interstitial fibrosis to the total atrial area), and an increase in the thickness of the ventricular wall. Moreover, the deposition area of interstitial fibrosis in the left ventricle of mice was reduced in the EVs‐PAP@SA treatment group compared with the PAP and EVs@SA groups (Figure [179]4F). The results of the Tianlang scarlet staining revealed that the myocardial fibrosis area (micro Sirius red positive area) in MI mice was increased compared with the normal control group. However, treatment with EVs‐PAP@SA reduced the myocardial fibrosis area in mice (Figure [180]4G). The results of TUNEL staining confirmed an increase in the apoptosis of myocardial tissue cells in the PBS group compared with the normal control group. Additionally, the apoptosis of myocardial tissue cells was reduced in the PAP and EVs@SA groups compared with the PBS group. Furthermore, the apoptosis of myocardial tissue cells was further reduced after treatment with EVs‐PAP@SA compared with the PAP and EVs@SA groups (Figure [181]4H). The results confirm that EVs‐PAP@SA can inhibit heart necrosis induced by MI, improve cardiac fibrosis in MI mice, and suppress myocardial cell apoptosis. Transcriptomic Insights Into the Mechanism of EVs‐PAP@SA in Alleviating Myocardial Fibrosis Through TGF‐β1 and Mitogen‐Activated Protein Kinase Pathways To further investigate the mechanism of EVs‐PAP@SA on myocardial fibrosis in mice, heart tissues were randomly selected from 4 mice in the PBS group and 4 mice in the EVs‐PAP@SA group for transcriptome sequencing analysis to examine the mRNA expression levels in mice. After completing quality control and filtering the raw data, we conducted a differential analysis using the |log2FC|>1 and P value<0.05 criteria. In total, we identified 119 differentially expressed genes, with 79 genes downregulated and 40 genes upregulated (Figure [182]5A and [183]5B). We analyzed the protein–protein interaction network of key factors using the STRING database. The interaction relationships were visualized using Cytoscape 3.5.1 software. Additionally, we identified the core genes using the built‐in tool CytoHubba. The results indicated that several genes, including Mapk14, Ppang, Tgfb1, Mrpl2, Cox5a, and Rps11, are vital network components. Notably, the Mapk14 gene occupies a central position, as evidenced by its highest combined score (Figure [184]5C). Figure 5. Based on transcriptome sequencing analysis, possible mechanisms of EVs‐PAP@SA in improving myocardial fibrosis in mice. Figure 5 [185]Open in a new tab A, Heat map of differentially expressed genes between the PBS group (4 samples) and the EVs‐PAP@SA group (4 samples) in mouse heart tissue. Orange indicates upregulated genes, and blue indicates downregulated genes; B, Volcano plot of differentially expressed genes between the PBS group (4 samples) and the EVs‐PAP@SA group (4 samples). Orange dots represent upregulated genes, blue dots represent downregulated genes, and black dots represent nondifferentially expressed genes; C, PPI network diagram of Mapk14, with color grades from pink to red indicating increasing degree values of genes; D, KEGG pathway enrichment analysis results of differentially expressed genes; E, GO function enrichment analysis results of differentially expressed genes. BP indicates Biological Process; CC, Cellular Component; EV, extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; GO, Gene Ontology; KEGG, Kyoto Encyclopedia of Genes and Genomes; MAPK, mitogen‐activated protein kinase; MF, Molecular Function; PAP, p38α antagonistic peptide; PPAR, peroxisome proliferator–activated receptor; PPI, protein–protein interaction; SA, sodium alginate; and TGF, transforming growth factor. We performed Kyoto Encyclopedia of Genes and Genomes pathway enrichment analysis and Gene Ontology functional enrichment analysis based on the results of the differential gene analysis. The pathway analysis results indicated that these differentially expressed genes are primarily involved in several signaling pathways, such as Huntington's disease, MAPK (mitogen‐activated protein kinase) signaling pathway, diabetic cardiomyopathy, TGF‐β signaling pathway, cancer transcriptional abnormal regulation, and calcium signaling pathway (Figure [186]5D). The functional analysis results of Gene Ontology (Figure [187]5E) indicate that the differentially expressed genes primarily participate in various biological processes, including phosphorylation of the RNA polymerase C‐terminal domain, binding to sperm and acrosome, and mRNA transcription. Moreover, these genes are predominantly enriched in cellular components like proteins in mitochondria, membranes, and inner membrane protein complexes in mitochondria. Additionally, they are primarily associated with molecular functions such as growth factor activity, growth factor receptor binding, RNA polymerase II specificity, and DNA‐binding transcription factor binding. According to literature reports, TGF‐β1 receptor signaling could activate the transcription factor SMAD2/3 through the pathway mediated by the MAPK effector, thereby modifying the expression of myocardial fibrosis markers, such as α‐SMA and collagen I genes, and initiating the process of myocardial cell fibrosis. Additionally, p38α plays a crucial role as a mediator of TGF‐β1 action.[188] ^17 , [189]^18 , [190]^19 Therefore, we hypothesize that EVs‐PAP@SA could enhance myocardial fibrosis reduction in MI mice through the regulation of the TGF‐β1 signaling pathway. Inhibition of Cardiac Fibroblast Fibrosis by EVs‐PAP@SA Through Dual Modulation of TGF‐β1/SMAD and SRF/TRPC6 Signaling Pathways To further explore the potential of EVs‐PAP@SA in mitigating myocardial injury in mice by modulating the TGF‐β1 and SMAD signaling pathways, we analyzed the expression of TGF‐β1 and SMAD‐related genes and proteins in CFs from various groups using RT‐qPCR and Western blot techniques. RT‐qPCR results revealed that the expression of Mapk14, Tgfb1, Smad2, Smad3, and Acta2 genes in CFs under hypoxic conditions was upregulated when compared with the normal control group. Furthermore, EVs‐PAP@SA inhibited the expression of Mapk14, Tgfb1, Smad2, Smad3, and Acta2 genes under hypoxic conditions compared with the CFs treated with PBS. Notably, EVs‐PAP@SA exhibited a stronger inhibitory effect on the expression of genes associated with the TGF‐β1/SMAD signaling pathway compared with PAP and EVs@SA therapies (Figure [191]6A). Western blot analysis demonstrated that hypoxia upregulated the expression and phosphorylation levels of p38α protein, whereas EVs‐PAP@SA downregulated the expression and phosphorylation levels of p38α protein in CFs under hypoxic conditions (Figure [192]6B). Figure 6. Influence of EVs‐PAP@SA on the TGF‐β1/SMAD and TRPC6/NFAT pathways in the fibrosis of CFs. Figure 6 [193]Open in a new tab A, RT‐qPCR detection of changes in Mapk14, Tgfb1, Smad2, Smad3, and Acta2 mRNA expression in CFs after treatment with different reagents for 24 hours; B, Western blot detection of changes in p38α, p‐p38α, α‐SMA, and collagen I protein expression in CFs after treatment with different reagents for 24 hours; C, ELISA to detect the expression of collagen I in the cell culture medium; D, Western blot detection of changes in TGF‐β1, SMAD2, p‐SMAD2, SMAD3, and p‐SMAD3 protein expression in CFs after treatment with different reagents for 24 hours; E, RT‐qPCR detection of changes in Srf, Nfat1, and Trpc6 mRNA expression in CFs after treatment with different reagents for 24 hours; F, Western blot detection of changes in SRF, NFAT1, TRPC6, and whether PAP could modulate protein expression in CFs after treatment with different reagents for 24 hours; G, Changes in intracellular Ca^2+ influx in CFs after treatment with different reagents for 24 hours; H, Immunofluorescence staining to detect changes in SRF and TRPC6 protein expression in CFs after treatment with different reagents for 24 hours, with green fluorescence indicating SRF protein and red fluorescence indicating TRPC6 protein (scale bar: 25 μm). CFs in the normal group were cultured in a 5% CO[2], 37 °C incubator, whereas CFs in the other treatment groups (PBS group, PAP group, EVs@SA group, and EVs‐PAP@SA group) were cultured in a 1% O[2], 5% CO[2], 37 °C incubator to induce myocardial fibrosis. The multiple groups were analyzed using Kruskal–Wallis test. The bar graphs represent the mean±SD of quantitative data, and each cellular experiment was repeated 3 times (n=3). # represents a difference compared with the normal group, ^# P<0.05, ^## P<0.01; * represents a difference compared with the PBS group, *P<0.05, **P<0.01. CaN indicates calcium/calmodulin‐dependent protein kinase; CF, cardiac fibroblasts; EV, extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; NFAT, nuclear factor of activated T cells; PAP, p38α antagonistic peptide; RT‐qPCR, real‐time quantitative polymerase chain reaction; SA, sodium alginate; SMAD, suppressor of mothers against decapentaplegic; SRF, serum response factor; TGF‐β, transforming growth factor beta; and TRPC6, transient receptor potential channel protein 6. Furthermore, compared with the PBS group, the EVs‐PAP@SA group exhibited decreased expression of α‐SMA and collagen I proteins (Figure [194]6C). Compared with the normal control group, hypoxia could enhance the expression of TGF‐β1 and its downstream proteins, SMAD2 and SMAD3. Compared with the PBS group, the expression of TGF‐β1 protein and its downstream proteins, SMAD2 and SMAD3, as well as their phosphorylation levels, were downregulated in the EVs‐PAP@SA group (Figure [195]6D). The experimental results suggest that EVs‐PAP@SA effectively inhibits CF fibrosis under hypoxic conditions by suppressing the TGF‐β1/SMAD signaling pathway. Furthermore, our literature research has discovered that in fibroblasts, p38α can enhance the transcriptional activity of SRF. It also induces the expression of TRPC6, increasing intracellular Ca2+ influx through TRPC6. Consequently, this enhances the activity of CaN (calcium/calmodulin‐dependent protein kinase) and activates NFAT. The synergistic effect of SRF and NFAT promotes the expression of genes related to muscle fiber cell differentiation and fibrosis, ultimately leading to cell fibrosis.[196] ^21 , [197]^22 , [198]^23 Therefore, we assessed the expression of TRPC6/NFAT signaling pathway‐related proteins and genes using RT‐qPCR and Western blot analyses. The RT‐qPCR results demonstrated an upregulation of Srf, Nfat1, and Trpc6 mRNA in CFs under hypoxic conditions compared with the normal control group. In contrast, the expression of Srf, Nfat1, and Trpc6 mRNA was downregulated in the EVs‐PAP@SA group compared with the PBS group (Figure [199]6E). The Western blot analysis revealed that under hypoxic conditions, the treatment of EVs‐PAP@SA suppressed the expression of SRF, NFAT1, TRPC6, and whether PAP affects protein levels in CFs. Importantly, this inhibitory effect was stronger than that observed with PAP and EVs@SA treatments (Figure [200]6F). The results of the Ca2+ uptake experiment showed that, under hypoxic conditions, the intracellular Ca2+ influx of CFs increased compared with the normal control group. However, EVs‐PAP@SA inhibited the intracellular Ca2+ influx of CFs under hypoxic conditions compared with the PBS group. Moreover, this effect was stronger than the effects of PAP and EVs@SA (Figure [201]6G). Additionally, the results of immunofluorescence costaining for SRF and TRPC6 demonstrated increased expression of SRF and TRPC6 proteins in CFs under hypoxic conditions compared with the normal control group. However, the expression of SRF and TRPC6 proteins in the EVs‐PAP@SA group was downregulated compared with the PBS group (Figure [202]6H). This result suggests that EVs‐PAP@SA could decrease the fibrosis level of CFs under low oxygen conditions by inhibiting the TRPC6/NFAT signaling pathway. EVs‐PAP@SA Attenuates Myocardial Fibrosis and Inflammation in MI Mice Through Modulation of TGF‐β1/SMAD and TRPC6/NFAT Pathways To further explore the inhibition of fibrosis in CFs by EVs‐PAP@SA and examine its impact on the TGF‐β1/SMAD and TRPC6/NFAT signaling pathways, we costimulated CFs with EVs‐PAP@SA along with a TGF‐β activator (SRI‐011381, SRI) or a TRPC6 activator (hyperforin). EDU staining, Transwell assay, and flow cytometry analysis revealed that co‐treatment with SRI and Hyp, in comparison to treatment with EVs‐PAP@SA alone, decreased the proliferation and migration abilities of CFs (Figure [203]7A and [204]7B) and induced cell apoptosis (Figure [205]7C). The Western blot analysis demonstrated that co‐administration of SRI along with EVs‐PAP@SA enhanced the activation of SMAD2 and SMAD3 proteins compared with treatment with EVs‐PAP@SA alone (Figure [206]7D). Moreover, simultaneous administration of hyperforinresulted in increased expression of SRF, NFAT1, and whether PAP reduces elevated levels of α‐SMA and collagen I proteins (Figure [207]7E). This finding implies that the impact of EVs‐PAP@SA could be counteracted by either a TGF‐β activator or a TRPC6 activator. ELISA was employed to measure the expression of collagen I in the cell culture medium. The results indicated that the expression of collagen I protein was elevated when treated with SRI or hyperforinsimultaneously (Figure [208]7F). Figure 7. Influence of EVs‐PAP@SA on the TGF‐β1/SMAD and TRPC6/NFAT signaling pathways in CFs fibrosis. Figure 7 [209]Open in a new tab A, EDU assay to detect changes in the proliferative capacity of CFs after treatment with different reagents for 24 hours (scale bar: 25 μm); B, Transwell assay to detect changes in the migration capacity of CFs after treatment with different reagents for 24 hours (scale bar: 50 μm); C, AnnexinV/PI double‐staining flow cytometry to detect changes in apoptosis of CFs after treatment with different reagents for 24 hours, with the column chart representing the percentage of cells in the Q2 and Q3 regions as the apoptotic rate; D, Western blot detection of changes in SMAD2, p‐SMAD2, SMAD3, and p‐SMAD3 protein expression in CFs after treatment with different reagents for 24 hours; E, Western blot detection of changes in SRF, NFAT1, CaN, α‐SMA, and collagen I protein expression in CFs after treatment with different reagents for 24 hours. CFs in the normal group were cultured in a 5% CO[2], 37 °C incubator, whereas CFs in the other treatment groups (PBS group, PAP group, EVs@SA group, and EVs‐PAP@SA group) were cultured in a 1% O[2], 5% CO[2], 37 °C incubator to induce myocardial fibrosis. F, ELISA to detect the expression of collagen I in the cell culture medium; G, Immunohistochemical staining to detect changes in p38α, p‐p38α, and TGF‐β1 protein expression in mouse myocardial tissue after 28 days of injection treatment (scale bar: 50 μm); H, Western blot detection of changes in TGF‐β1, SMAD2, p‐SMAD2, SMAD3, and p‐SMAD3 protein expression in mouse myocardial tissue after 28 days of injection treatment. Mice in the normal group underwent thoracotomy without ligation; mice in the other treatment groups (PBS group, PAP group, EVs@SA group, and EVs‐PAP@SA group) underwent heart ligation to establish an MI model. The multiple groups were analyzed using 1‐way ANOVA. The bar graphs represent the mean ± SD of quantitative data, and each group consisted of 10 mice (n=10). # represents a difference compared with the normal group, ^# P<0.05, ^## P<0.01; * represents a difference compared with the PBS group, *P<0.05, **P<0.01. α‐SMA indicates alpha smooth muscle actin; CaN, calcium/calmodulin‐dependent protein kinase; CF, cardiac fibroblasts; EDU, 5‐Ethynyl‐2′‐deoxyuridine; EV, extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; MI, myocardial infarction; NFAT, nuclear factor of activated T cells; PAP, p38α antagonistic peptide; PI, propidium iodide; RT‐qPCR, real‐time quantitative polymerase chain reaction; SA, sodium alginate; SMAD, suppressor of mothers against decapentaplegic; SRF, serum response factor; and TGF‐β1, transforming growth factor beta1. Immunohistochemical staining was performed on heart tissue sections from mice to validate whether the EVs‐PAP@SA mechanism for improving myocardial fibrosis in MI mice aligns with its in vitro mechanism. It allowed us to examine the expressions of p38α, p‐p38α, and TGF‐β1 proteins. The experimental results indicated that in MI mice, the expression of p38α and its phosphorylated protein, along with TGF‐β1 protein, was upregulated compared with the normal control group. Compared with the PBS group, the EVs‐PAP@SA treatment group exhibited downregulation of p38α, p‐p38α, and TGF‐β1 protein expression in mice. Furthermore, the inhibitory effect of EVs‐PAP@SA on p38α, p‐p38α, and TGF‐β1 protein expression was more potent than that of PAP and EVs@SA (Figure [210]7G). Moreover, we performed Western blot analysis to homogenize cardiac tissue from mice. The results indicated that, compared with the normal control group, the expression of TGF‐β1 protein was increased in the myocardial tissue of MI mice. Moreover, the expression of downstream proteins, including SMAD2 and SMAD3, as well as their phosphorylated forms, was also elevated. In contrast to the PBS group, EVs‐PAP@SA inhibits the expression of TGF‐β1 and its downstream proteins SMAD2 and SMAD3 (Figure [211]7H). Immunofluorescence staining of mouse cardiac tissue revealed that the expression of α‐SMA, collagen I, SRF, and TRPC6 proteins was upregulated in MI mice compared with the normal control group. In contrast to the PBS group, the EVs‐PAP@SA treatment group mice showed downregulated expression of α‐SMA, Collagen I, SRF, and TRPC6 proteins (Figure [212]8A). The Western blot detection results demonstrated an upregulation of protein expression for SRF, NFAT1, and TRPC6 and could be in the myocardial tissue of MI mice, compared with the normal control group. In contrast to the PBS group, the expression levels of SRF, NFAT1, TRPC6, and proteins in the myocardial tissue of mice in the EVs‐PAP@SA group were found to be downregulated. This observation is supported by Figure [213]8B. Figure 8. Effects of EVs‐PAP@SA on regulating the TRPC6/NFAT pathway in myocardial fibrosis in mice. Figure 8 [214]Open in a new tab A, Immunofluorescent staining showing changes in the expression of α‐SMA and collagen I proteins (green fluorescence represents α‐SMA protein, red fluorescence represents collagen I protein) as well as SRF and TRPC6 proteins (green fluorescence represents SRF protein, red fluorescence represents TRPC6 protein) in the myocardial tissues of mice from different treatment groups after 28 days of injection therapy, with DAPI nuclear staining shown in blue (scale bar: 50 μm). B, Western blot analysis showed changes in the expression of SRF, NFAT1, TRPC6, and proteins in the myocardial tissues of mice from different treatment groups after 28 days of injection therapy. C, ELISA analysis shows changes in the expression of inflammatory factors TNF‐α and IL‐1β in the serum of mice from different treatment groups after 28 days of injection therapy. The normal group of mice underwent thoracotomy without ligation, whereas the other treatment groups (PBS group, PAP group, EVs@SA group, and EVs‐PAP@SA group) were subjected to heart ligation to induce MI. The multiple groups were analyzed using 1‐way ANOVA. The quantitative data in the figures are presented as mean±SD, with 10 mice in each group (n=10). # indicates statistical significance compared with the normal group, with ^# P<0.05, ^## P<0.01; * indicates statistical significance compared with the PBS group, *P<0.05, **P<0.01. α‐SMA indicates alpha smooth muscle actin; CaN, calcium/calmodulin‐dependent protein kinase; EV, extracellular vesicles; EVs‐PAP@SA, sodium alginate hydrogel loaded with extracellular vesicles and p38α antagonistic peptides; IL‐1β, interleukin‐1beta; MI, myocardial infarction; NFAT, nuclear factor of activated T cells; PAP, p38α antagonistic peptide; SA, sodium alginate; SRF, serum response factor; TNF‐α, tumor necrosis factor‐alpha; and TRPC6, transient receptor potential channel protein 6. Additionally, we detected the expression of inflammatory factors in the serum supernatant of each group of mice using ELISA. The results revealed that, in comparison to the normal control group, the expression of inflammatory factors TNF‐α and IL‐1β increased in MI mice. Compared with the PBS group, the expression of both TNF‐α and IL‐1β was decreased in the PAP and EVs@SA groups. Furthermore, the EVs‐PAP@SA group exhibited even lower expression of TNF‐α and IL‐1β (Figure [215]8C). In conclusion, the research findings demonstrate that EVs‐PAP@SA could enhance the extent of myocardial fibrosis in mice by modulating the TGF‐β1/SMAD and TRPC6/NFAT signaling pathways. DISCUSSION In recent years, myocardial fibrosis following MI has become a focal point of cardiovascular disease research.[216] ^20 Fibrosis not only leads to ventricular stiffness but also triggers arrhythmias, significantly affecting the survival rates of patients.[217] ^71 Despite numerous studies focusing on this pathological process, the specific mechanisms remain incompletely understood. Clarifying the pathogenesis can facilitate interventions targeting relevant pathways, thereby providing new therapeutic strategies. P38α plays a crucial role in cell signaling by engaging in various biological processes such as proliferation, differentiation, and apoptosis.[218] ^72 , [219]^73 , [220]^74 During the fibrotic process following MI, the activity of p38α is regulated, closely associated with fibrosis progression.[221] ^75 Other studies indicate that the TGF‐β/SMAD signaling pathway plays a significant role in cardiac fibrosis, with interventions altering the expression of fibrotic markers such as α‐SMA and collagen I genes, where p38α acts as a key mediator of TGF‐β1 activity.[222] ^17 , [223]^18 , [224]^19 , [225]^20 Additionally, p38α can promote the transcriptional activity of SRF, induce the expression of TRPC6, and activate NFAT to enhance cellular fibrosis.[226] ^21 , [227]^22 , [228]^23 Related myocardial fibrosis is a critical factor leading to patient mortality.[229] ^76 Interventions in this regard hold the potential to decrease mortality rates in patients with MI. The application of PAP offers a new strategy for regulating post‐MI fibrosis. The application of tissue engineering in disease treatment offers significant advantages. Hydrogels, as high‐molecular‐weight biomaterials, play a vital role in tissue repair. Biomaterials capable of forming self‐healing hydrogels include gelatin, collagen, hyaluronic acid, alginate, and chitosan.[230] ^77 Among these, alginate biomaterials have been proven to be biocompatible, nonimmunogenic, possess mechanical hardness, similar to the natural extracellular matrix, enhance EV repair capabilities, increase EV stability and bioavailability, and improve EV retention in the heart, making them an ideal material for treating MI.[231] ^51 This approach not only enhances drug release efficiency but also ensures its concentration at the target site. Compared with traditional drug delivery systems, SA hydrogels exhibit significant advantages. Furthermore, previous studies have demonstrated the critical role of EVs in the pathogenesis and treatment of various diseases.[232] ^69 , [233]^78 , [234]^79 Loading EVs into hydrogels can enhance their stability and facilitate sustained in situ release at defective sites.[235] ^68 This study used SA hydrogel‐encapsulated EVs as a delivery system for PAP to ensure the drug's stability and bioavailability in vivo. For the first time, a novel composite material incorporating SA hydrogel, EVs from murine BMSCs, and p38α antagonist peptide was employed to combat myocardial fibrosis, offering a new avenue for post‐MI therapy. The findings demonstrate that this novel composite material effectively inhibits myocardial fibrosis formation, exhibiting robust antifibrotic effects. Moreover, the EVs derived from BMSCs enhance cellular bioactivity and ameliorate myocardial structure and function post MI. Additionally, we confirmed the essential roles of the TGF‐β1/SMAD and TRPC6/NFAT pathways in myocardial fibrosis development, with the p38α antagonist peptide as a critical mediator; significant fibrosis reduction was achieved by inhibiting these pathways. This provides a strong target for developing novel therapeutic strategies, further confirming previous research on p38α in fibrosis of other diseases and the roles of TGF‐β1/SMAD and TRPC6/NFAT pathways in cardiovascular diseases. In conclusion, we successfully fabricated EVs‐PAP@SA. Through in vitro and in vivo mechanistic studies, we found that EVs‐PAP@SA suppresses the expression of TGF‐β1 protein and inhibits the phosphorylation of downstream proteins SMAD2 and SMAD3, subsequently suppressing myocardial fibrosis markers α‐SMA and collagen I protein transcription. Additionally, EVs‐PAP@SA inhibits p38α phosphorylation, reduces SRF transcriptional activity, lowers TRPC6 expression, decreases intracellular Ca2+ influx, subsequently diminishing the activity of CaN, inhibiting the signaling mediated by NFAT, downregulating SRF and NFAT expression, which further suppresses TRPC6 expression, forming a feedback loop to inhibit fibrosis and thus improving post‐MI myocardial fibrosis (Figure [236]S4). Additionally, our results not only demonstrate a significant effect of EVs‐PAP@SA on improving cardiac function but also show that the administration of SA hydrogel‐encapsulated EVs alone can enhance cardiac function post treatment. Studies indicate that EVs not only alleviate cardiac remodeling but also inhibit cardiomyocyte apoptosis, modulate immune cells and inflammation levels, and promote endothelial cell proliferation, migration, and vascular neogenesis.[237] ^80 , [238]^81 , [239]^82 EVs derived from mesenchymal stem cells contain functional proteins, mRNA, miRNAs, and tRNAs, which can suppress fibrosis and inflammation in the hearts of ischemic cardiomyopathy rat models by stimulating cardiomyocyte proliferation, reducing apoptosis, and inhibiting fibrosis.[240] ^83 This explains that besides serving as a delivery vehicle for PAP, BMSCs themselves have beneficial therapeutic effects on the heart. Furthermore, due to the poor stability and difficulty in retention of EVs, they are promptly eliminated by the immune system upon injection, necessitating a hydrogel delivery system to enhance their stability.[241] ^84 Fibrin hydrogel effectively improves the retention and stability of BMSC‐derived EVs in vivo, enhancing their delivery efficiency and dosing, thereby yielding higher efficacy and specificity in tissue injury treatment.[242] ^85 Coencapsulating PAP within EVs derived from mBMSCs using PAP in SA hydrogel instead of solely relying on SA hydrogel can ensure a more stable delivery and efficacy of PAP, preventing uncontrolled diffusion in the hydrogel matrix. Our study using SA hydrogel‐loaded EVs from BMSCs and PAP for MI treatment provides compelling evidence for the application of EVs in myocardial fibrosis therapy and offers a novel perspective for understanding its mechanisms. However, this study has certain limitations. Primarily, it is predominantly based on a mouse model. Given the substantial physiological and metabolic differences between humans and mice, the results may not directly translate to human application. Additionally, while the study shows that SA hydrogel can enhance the stability of PAP, it mainly focuses on short‐term cellular and tissue responses, neglecting the extended stability of stored materials, long‐term therapeutic effects after multiple uses, and possible side effects. Furthermore, because mBMSCs themselves have therapeutic properties, the optimal therapeutic ratio with PAP warrants further investigation. This therapeutic strategy may also be effective only for certain patients with MI, with individual responses varying. Furthermore, the detection of calreticulin as an endoplasmic reticulum marker indicates that our exosome preparations are not completely free of contamination. Such impurities could potentially affect the results and interpretations presented in this study. Refining exosome isolation techniques is crucial for future clinical applications to ensure the removal of these contaminants. Future research could delve into the mechanism of action of this composite material and assess its combined efficacy with other treatment strategies. Following stability and toxicity tests for the composite material along with exploration of potential side effects, early clinical trials can be conducted to validate the application value of this strategy in humans. Further refinement of this composite material to enhance its stability, efficacy, and suitability for large‐scale production and use can also be explored. In conclusion, although this study provides a new research direction and hope for myocardial fibrosis treatment post MI, further research is necessary to determine its clinical application value and assess its potential for clinical translation. CONCLUSIONS Ultimately, the composite therapy involving SA hydrogel‐loaded mBMSCs‐EVs and PAP holds promise in ameliorating the prognosis of patients with MI and enhancing their quality of life. The experimental findings of this study significantly augment our comprehension of the mechanisms underlying the treatment of myocardial fibrosis post MI, particularly with regard to the innovative approach of employing SA hydrogel for the delivery of mBMSCs‐EVs and PAP. Unmistakably, this therapeutic regimen proficiently modulates the TGF‐β1/SMAD and TRPC6/NFAT signaling pathways, thereby inhibiting myocardial fibrosis and promoting the repair and regeneration of myocardial cells, underscoring its substantial scientific merit and clinical potential. However, certain limitations are discernible in this research. Presently, the experimental outcomes primarily hinge on specific cell types and animal models. The broader applicability of this therapeutic modality to various stem cell categories, a more extensive array of animal models, or even human subjects requires further substantiation. Additionally, the long‐term stability of therapeutic outcomes and potential side effects warrant comprehensive exploration. Sources of Funding This study was supported by the Guangzhou Municipal Basic and Applied Basic Research Foundation (202201011721, 2023A04J0483). Disclosures None. Supporting information Tables S1–S3 Figures S1–S4 [243]JAH3-14-e036887-s001.pdf^ (1.5MB, pdf) Acknowledgments