An injectable multifunctional hydrogel for cardiac spatiotemporal repair via modulating oxidative stress and the inflammatory microenvironment

Abstract

Myocardial infarction (MI), a leading cause of global cardiovascular mortality, is characterized by a vicious cycle of oxidative stress and inflammatory responses, resulting in irreversible myocardial damage and ventricular remodeling. To address the limitations of current therapies in comprehensively targeting the post-MI pathological microenvironment, this study developed an injectable hydrogel system, termed CPH (DS/CMCS), through the rational integration of carboxymethyl chitosan (CMCS), dextran sulfate (DS), and oxidized dextran (ODex) as a dynamic crosslinker. The CPH hydrogel not only mimicked the mechanical properties of the native myocardial extracellular matrix but also integrated multifunctional capabilities, including antioxidant activity, anti-inflammatory effects, pro-angiogenic potential, and enhanced electrical signal conduction. Through both cellular and animal studies, it was conclusively shown that the CPH hydrogel effectively scavenged reactive oxygen species (ROS), protected cardiomyocytes from oxidative damage, modulated macrophage polarization to mitigate inflammatory cascades, and promoted vascular regeneration and myocardial remodeling. In the rat MI model, the CPH hydrogel significantly improved cardiac function and achieved comprehensive structural restoration of infarcted myocardium. This study introduces an innovative acellular spatiotemporal approach for the treatment of MI and advances the rational design of cardiac tissue-engineered biomaterials, highlighting its substantial clinical translation potential for regenerative medicine.

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1. Introduction

Myocardial infarction (MI), a life-threatening global disease, results from acute coronary occlusion leading to sustained myocardial ischemia, hypoxia, and irreversible cardiomyocytes (CMs) necrosis.¹ According to the World Health Organization, MI and its complications account for approximately 9 million deaths annually, representing over 40% of total cardiovascular mortality.^2,3 The pathological progression is driven by a vicious cycle of oxidative stress and inflammatory responses, which disrupts myocardial repair.⁴ Following coronary occlusion, mitochondrial electron transport chain uncoupling and NADPH oxidase activation trigger reactive oxygen species (ROS) overproduction, amplifying inflammatory cascades via nuclear factor-κB (NF-κB) and mitogen-activated protein kinase pathway activation.^5–7 Although reperfusion therapies restore blood flow, subsequent oxidative stress and cytokine storms exacerbate adverse ventricular remodeling, manifesting as dyssynchrony in the electrical conduction of CMs, vascular network disruption, and aberrant collagen deposition, ultimately leading to heart failure.^8–10 Current interventions, including pharmacological agents and stem cell transplantation,¹¹ provide partial symptomatic relief but fail to achieve integrated repair by concurrently modulating the microenvironment, restoring electromechanical coupling, and promoting tissue regeneration. Consequently, the development of bioactive and functionally adaptive myocardial repair materials has emerged as a pivotal research frontier to address these multidimensional challenges.

In the field of cardiac tissue engineering, hydrogels derived from natural polysaccharides have recently attracted considerable interest owing to their exceptional biocompatibility, biodegradability, and tailorable functionality.¹² Carboxymethyl chitosan (CMCS), a water-soluble derivative obtained through chitosan carboxylation, retains the inherent bioactivity of chitosan while offering tunable properties via chemical modification. The coexistence of amino and carboxyl groups in its molecular structure confers pH responsiveness, ion-binding capacity, and characteristics of multiple biological interactions,¹³ positioning CMCS as a uniquely advantageous material for myocardial regeneration. Research demonstrates that CMCS promotes cardiac repair through synergistic mechanisms encompassing physical support, biochemical regulation, and dynamic responsiveness.¹⁴ As a functional scaffold, CMCS can be combined with gelatin or polycaprolactone to fabricate biomimetic myocardial matrices. These constructs mimic native cardiac mechanical microenvironments in elastic modulus and porosity, guiding CMs alignment and enhancing electromechanical synchronization.¹⁵ Concurrently, CMCS mitigates pathological oxidative stress and inflammation via its intrinsic antioxidant and anti-inflammatory properties.¹⁶ Notably, its dual pH/ROS-responsive behavior enables on-demand therapeutic delivery: protonation of carboxyl groups in ischemic regions enhances drug dissociation efficiency, dynamically adapting to myocardial repair processes.^17,18

Dextran sulfate (DS), an anionic polysaccharide derived from sulfated dextran, holds significant promise in cardiac tissue engineering due to its biocompatibility, biodegradability, and anticoagulant activity. The sulfate groups confer strong negative charges, which not only reduce thrombotic risk by inhibiting clotting factors but also enable electrostatic interactions with vascular endothelial growth factor (VEGF), fibroblast growth factor,¹⁹ facilitating sustained release to promote angiogenesis and myocardial repair. DS further modulates inflammatory microenvironments by suppressing pro-inflammatory cytokine expression and enhancing hydrogel mechanical properties when composited with gelatin or hyaluronic acid, achieving elastic modulus values akin to native myocardium.²¹ In practical applications, DS is utilized as 3D scaffold materials to support CMs or stem cell adhesion/differentiation, or as carriers for drug/gene delivery to regulate fibrotic progression.²² When integrated with conductive materials, DS-based cardiac patches improve electrical signal propagation and cardiac functional recovery.¹⁰

However, conventional single-component systems remain inadequate to address the multifaceted pathological microenvironment post-MI, necessitating rational integration of multifunctional components. Herein, we innovatively engineered an injectable hydrogel termed CPH (DS/CMCS) by compositing CMCS with DS and incorporating oxidized dextran (ODex) as a dynamic crosslinker. The hydrogel formed a homogeneous porous network through Schiff base reactions and electrostatic interactions (Fig. S1), with a 3D topology mimicking the mechanical properties of native myocardial extracellular matrix to provide structural support for CMs migration and alignment.²³ The unique electroactivity of the CPH hydrogel stemmed from ionic conduction synergy between CMCS and DS, significantly enhancing intercellular electromechanical synchronization to mitigate post-MI conduction block caused by prolonged QRS intervals.²³ This study systematically investigated the hydrogel's multidimensional biofunctions, including ROS scavenging capacity, macrophage polarization modulation, pro-inflammatory cytokine suppression, and anti-fibrotic efficacy (As shown in Fig. 1). The experimental data confirmed that the CPH hydrogel not only ameliorated ventricular remodeling but also augmented tissue repair via pro-angiogenic mechanisms. These findings underscore the clinical potential of the CPH hydrogel as a transformative intervention for ischemic myocardial diseases.

Fig. 1 In the rat MI model, the CPH hydrogel was intramyocardially injected into ischemic tissue. This intervention alleviated oxidative stress, modulated macrophage polarization, enhanced angiogenesis, and suppressed fibrosis in the infarct zone, ultimately improving myocardial repair.

2. Materials and methods

2.1 Materials

Dextran (Dex, M_w = 40 kDa) and dextran sulfate (DS, M_w = 100 kDa) were purchased from Aladdin (Shanghai, China). 3,4-Ethylenedioxythiophene (EDOT, 97%), sodium periodate (NaIO₄, ≥99.8%), ethylene glycol (99.8%), ammonium persulfate (APS, ≥98%), and iron(II) sulfate heptahydrate (FeSO₄·7H₂O, ≥99.0%) were purchased from Sigma-Aldrich (USA). Carboxymethyl chitosan (CMCS, carboxylation degree ≥80%) was purchased from Yuanye (Shanghai, China).

Synthesis of ODex. First, Dex (5 g) was dissolved in water (50 mL). Then, NaIO₄ solution (0.25 mol L⁻¹, 20 mL) was added to the solution. The reaction proceeded for a duration of 6 h. Subsequently, ethylene glycol (30 mL) was introduced to effectively terminate the reaction process, followed by a three-day dialysis process against a deionized aqueous medium. The resulting ODex product was collected via lyophilization.

For the synthesis of the CPH:(DS/CMCS), an initial step involved dissolving DS (0.5 g) in aqueous solution (25 mL) under an argon atmosphere with ice-water bath cooling. Thereafter, the EDOT (600 µL) was introduced dropwise into the mixture. Subsequently, an aqueous solution containing APS (1.2 g) and FeSO₄·7H₂O (30 mg) in 5 mL of water was prepared and incorporated into the reaction system. This mixture was maintained at 0 °C for 48 h. After completing dialysis against deionized water for 3 days, the CPH:DS complex was acquired through lyophilization. To formulate the CPH:(DS/CMCS) aqueous dispersion, the CPH:DS (0.5 g) was uniformly distributed in the CMCS solution (10 mL) using ultrasonic treatment. Finally, the ODex and the CPH : (DS/CMCS) solutions at predetermined concentrations were combined in a 1 : 1 volume ratio employing a dual-chamber syringe system to yield the CPH hydrogel.

The H9C2 cell line was purchased from Sunncell Biotechnology (Wuhan, China). The CCK-8 kit was purchased from Solarbio Technology (Beijing, China), and the acridine orange (AO) kit was purchased from Beyotime Biotechnology (Shanghai, China). Dihydroethidium (DHE) and 2′,7′-dichlorofluorescein diacetate (DCFH-DA) were purchased from Beyotime Biotechnology (Shanghai, China). Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) was purchased from Life-iLab (Shanghai, China). Dulbecco's modified Eagle medium (DMEM) was purchased from Sigma Aldrich (Germany). Masson's trichrome solution was purchased from Beyotime Biotechnology (Shanghai, China). Fetal bovine serum (FBS), penicillin, and streptomycin were purchased from Thermo Fisher (USA). Actin, alpha-actinin 2 (ACTN2), cardiac troponin T (cTnT), and connexin 43 (Cx43) were purchased from Huaan Biotechnology (Hangzhou, China). 8-Hydroxydeoxyguanosine (8-OHdG) was purchased from Arigo biolaboratories (Shanghai, China). CD34 and alpha-smooth muscle actin (α-SMA) were purchased from Cell Signaling Technology (USA). CD68 and F4/80 were purchased from Biolegend (USA). Inducible nitric oxide synthase (iNOS) and CD206 were purchased from Santa Cruz (USA). Interleukin-10 (IL-10) and tumor necrosis factor-α (TNF-α) were purchased from Huaan Biotechnology (Hangzhou, China). Wheat germ agglutinin (WGA) was purchased from VECYOR (USA). 4′,6-Diamidino-2-phenylindole (DAPI) was purchased from Cell Signaling Technology (USA). Fluorescent secondary antibodies were purchased from Invitrogen (USA).

2.2 Animals

All animal subjects used in this study were supplied by the Laboratory Animal Center at North China University of Science and Technology. The rats were maintained under specific pathogen-free conditions within a barrier facility, with a controlled 12-h light/dark photoperiod. They received ad libitum access to drinking water and were fed a standard rodent diet throughout the experimental period. All procedures involving animals were conducted in strict accordance with China's Guidelines for the Ethical Treatment of Laboratory Animals. The study protocol received formal review and approval from the Ethics Committee of North China University of Science and Technology (Approval number: SCXK (Beijing) 2019–0008) and was carried out under comprehensive oversight.

2.3 Isolation and culture of CMs and BMDMs

CMs isolation and culture. Neonatal SD rats (within 24 h post-birth) were euthanized following ethanol disinfection. Hearts were aseptically excised, rinsed with saline to remove residual blood, and minced into 1 mm³ fragments. Tissue fragments were digested in 0.2% collagenase II solution under 37 °C agitation (5 min per cycle, repeated 5–6 times). The first digestion supernatant was discarded, while subsequent supernatants were collected. Pooled supernatants were centrifuged at 1500 rpm for 5 min, and the pellet was resuspended in high-glucose DMEM, filtered through a 100-mesh sieve, and subjected to differential adhesion in culture flasks for 2 h. Non-adherent cells were collected, centrifuged (1500 rpm, 5 min), and resuspended in DMEM for 72 h culture (37 °C, 5% CO₂). To enhance purity, adherent cells were maintained for 4–5 days until reaching >90% confluency with synchronized beating before experimentation.

Bone marrow-derived macrophages (BMDMs) isolation and culture. Femurs and tibiae from Eight-week-old male C57BL/6J mice were aseptically isolated, and bone marrow cells were flushed with RPMI-1640. After centrifugation (1500 rpm, 5 min, 4 °C), the pellet was treated with RBC lysis buffer (2 min, 37 °C), neutralized with PBS, and centrifuged again. Cells were resuspended in RPMI-1640 supplemented with 10% FBS and cultured for 7 days (37 °C, 5% CO₂). Macrophage purity was confirmed by F4/80 and CD68 positivity (analyzed by flow cytometry, >95% purity) before downstream assays.

2.4 AO/PI staining

Cell viability was detected by AO/PI fluorescence staining. Cells underwent two washing cycles using PBS and were subsequently resuspended in 500 µL of staining buffer, followed by the addition of 5 µL AO reagent and 10 µL PI reagent. After gentle mixing, the samples were maintained under dark conditions at 37 °C for a duration of 20 min. Following this incubation, the cells underwent four successive washes with PBS, each lasting 5 min, and were imaged under a fluorescence microscope. Viable cells exhibited green fluorescence.

2.5 CCK-8 detection

To prepare the hydrogel extract, 10 mg of the CPH hydrogel was incubated in 10 mL of complete culture medium over a 48-h period. H9C2 cells were then plated onto 96-well plates at a density of 1 × 10⁴ cells per well, with three replicate wells allocated per experimental group. All cellular cultures were subsequently maintained under controlled conditions at 37 °C with 5% CO₂ atmosphere, using fresh culture medium as the control treatment. H9C2 cells were randomly divided into three groups: control, H₂O₂, and H₂O₂ + CPH. Cells in the H₂O₂ group were treated with 200 µmol L⁻¹ H₂O₂ for 24 h, while the H₂O₂ + CPH group received co-treatment with 1 mg mL⁻¹ CPH hydrogel and 200 µmol L⁻¹ H₂O₂ for 24 h. After that, to each well, 10 µL of CCK-8 reagent was added, and the plates were subsequently incubated for 2 h. The absorbance values were then quantified at a wavelength of 450 nm employing a microplate reader. Cell viability was calculated using eqn (1):where A_sample and A_control represent the absorbance of CCK-8 solution in the hydrogel sample and control group, respectively, A_blank represents the absorbance of CCK-8 solution in the absence of cell culture.

2.6 ROS level detection

CMs in optimal condition were seeded onto pre-coated cell climbing slides in 24-well plates at 2 × 10⁵ mL⁻¹ density and cultured for 12 h. DHE and DCFH-DA probes were prepared as 10 µmol L⁻¹ working solutions using the antibody dilution buffer. Following fixation with 4% paraformaldehyde, the cells underwent three washes with PBS. Subsequently, each well was treated with 100 µL of probe working solution and maintained in darkness at 37 °C for 30 min. After three PBS rinses (5 min each), samples were mounted with anti-fade DAPI mounting medium and imaged via laser confocal microscopy. Mean fluorescence intensity was quantified using ImageJ software to evaluate intracellular ROS levels.

2.7 Injection procedure of the rat MI model and the CPH hydrogel

The MI model was constructed by ligation of the anterior descending artery of the left coronary artery in rats. Specific steps: sixty male SD rats (8 weeks, weighing 250 ± 10 g) were fasted for 12 h, and then tracheal intubation and mechanical ventilation were performed after anesthesia. A thoracic incision was made along the fourth and fifth intercostal spaces adjacent to the cardiac apex, followed by ligation of the left anterior descending coronary artery using a 6–0 suture. Successful ligation is judged by partial pallor ischemia of the myocardium, which may be accompanied by abnormal wall motion. The rats subjected to MI modeling were randomly allocated to three experimental groups: Sham, MI, and CPH. For the Sham group, a myocardial puncture procedure was conducted without subsequent coronary ligation. In the MI group, 100 µL of PBS was injected into the myocardium. In the CPH group, 100 µL of the CPH hydrogel (1 mg mL⁻¹) was injected into the myocardial region around the infarction at three points. After the operation, the residual gas in the chest cavity was discharged, and the chest was closed layer by layer, and the rats continued to be reared after waking up.

2.8 Electrocardiogram

At postoperative day 28, rats were anesthetized and immobilized for electrocardiogram (ECG) recording. Subcutaneous needle electrodes were positioned at the left forelimb and right hindlimb, and signals were acquired using a PowerLab system (AD Instruments, Australia). Continuous ECG waveforms were recorded for 3 minutes at a paper speed of 50 mm s⁻¹via LabChart software. ST-segment alterations were analyzed, and the QRS interval was measured to evaluate cardiac electrophysiological recovery.

2.9 Echocardiography

Left ventricular function was assessed using the Visual Sonics Vevo 2100 high-resolution small animal ultrasound imaging system. Key parameters, including left ventricular ejection fraction (LVEF), fractional shortening (LVFS), end-diastolic diameter (LVDD), and end-systolic diameter (LVDS), were measured to quantify systolic performance and pump efficiency. Myocardial injury severity was evaluated through comparative analysis of these functional indices.

2.10 Histopathological and immunostaining analysis

Rats were anesthetized and euthanized at postoperative days 1, 3, 5, and 28. Hearts were rapidly excised, fixed, and embedded. Oxidative stress was assessed via DHE, DCFH-DA, and 8-OHdG fluorescence to label reactive ROS. Apoptosis was quantified using the TUNEL assay, while immunofluorescence staining was performed to evaluate inflammatory markers (TNF-α, IL-10, iNOS, CD206) and DNA oxidative damage marker (8-OHdG). At day 28, myocardial pathology was examined by H&E staining, and collagen deposition was quantified using Masson's trichrome staining. Potential histopathological alterations in the liver, spleen, lung, and kidney of rats in the three groups were investigated to assess in vivo biocompatibility. Myocardial structural integrity was assessed via Actin and Cx43 labeling, angiogenesis density was determined by dual CD34/α-SMA immunofluorescence, and cardiomyocyte cross-sectional area was analyzed using WGA membrane staining combined with DAPI nuclear counterstaining. All fluorescent images were acquired using a laser confocal microscope (Andor Dragonfly, Oxford Instruments, UK) and analyzed for fluorescence intensity and morphometric parameters with ImageJ software.

2.11 Statistical analysis

All quantitative results were expressed as mean ± standard deviation (n = 3 or n = 5). Comparisons between two groups were carried out with Student's t-test, while differences among multiple groups were evaluated through one-way analysis of variance. Data analyses were performed using GraphPad Prism 10 and ImageJ software, with a threshold of P < 0.05 indicating statistical significance.

3. Results

3.1 Biological properties of the CPH hydrogel and the regulation of ROS microenvironment

To investigate the direct antioxidant activity of the CPH hydrogel, we conducted radical scavenging assays against DPPH˙, O₂⁻, and ˙OH.^20,24 The results revealed that the CPH hydrogel exhibited significant antioxidant efficacy, demonstrating scavenging rates of 45.12 ± 2.89%, 53.03 ± 2.05%, and 50.17 ± 2.15% for DPPH˙, O₂⁻, and ˙OH, respectively (Fig. S2). Then, in vitro ROS microenvironment was simulated, CMs were subjected to oxidative stress using 200 µmol L⁻¹ H₂O₂, with normal medium-cultured CMs as controls.²⁵ To systematically evaluate the ROS clearance capacity of the CPH hydrogel, dihydroethidium (DHE) and 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA) fluorescence were used to detect intracellular superoxide anion and total ROS levels. On the first day after H₂O₂ stimulation, cells without the CPH hydrogel (0 mg mL⁻¹) showed significant accumulation of DHE red fluorescence and DCFH-DA green fluorescence around the nuclei, indicating a large accumulation of ROS. As the CPH concentration increased to 0.2 mg mL⁻¹ and 0.5 mg mL⁻¹, the fluorescence intensity of both decreased in a concentration-dependent manner. Under treatment with 1 mg mL⁻¹ CPH, the fluorescence signal was significantly reduced by the first day and was nearly completely gone by the third day, with only weak background fluorescence visible (Fig. 2(A)). Quantitative fluorescence analysis further indicated that, on the third day, the signal intensity of DHE and DCFH-DA in the 1 mg mL⁻¹ CPH group was 11.9% and 41.7% of the Control group, respectively, significantly lower than the other concentration groups (***P < 0.001, Fig. 2(B) and (C)). The above results clearly demonstrated that CPH hydrogel could dose-dependently clear intracellular ROS, with optimal antioxidant effects at a concentration of 1 mg mL⁻¹. Therefore, this concentration was selected for subsequent experiments to further validate the ROS clearance capability and cardioprotective effects in vivo.

Fig. 2 Biological properties of the CPH hydrogel and the regulation of ROS microenvironment. (A) Representative fluorescence images of intracellular superoxide anion radical activity (DHE) and intracellular total ROS (DCFH-DA) on days 1 and 3 of the CPH hydrogel in the ROS microenvironment. Statistical evaluation of (B) DHE and (C) DCFH-DA on days 1 and 3. (D) AO/PI staining of H9C2 cells. (E) CCK-8 detection results of H9C2 cells cultured in the CPH hydrogel extract for 48 h. (F) Confocal laser microscopy of Actin (red) and Cx43 (green) immunostaining of CMs. (G) Statistical evaluation of fluorescence intensity of Actin. (H) Statistical evaluation of fluorescence intensity of Cx43. (n = 3, *P < 0.05, **P < 0.01, ***P < 0.001).

To assess the biological properties of the CPH hydrogel, we used the AO/PI staining method to detect the cell viability of the rat myocardial cell line H9C2. According to Fig. 2(D), the H₂O₂ group demonstrated sparse green fluorescence (viable cells), whereas the CPH group demonstrated a significant increase in green fluorescent cells, indicating effective suppression of oxidative stress-induced injury. Assessment with the CCK-8 assay indicated that H₂O₂ markedly reduced H9C2 cells’ viability (Fig. 2(E)), whereas the CPH hydrogel markedly improved H9C2 cells’ survival under oxidative stress, suggesting its potential to scavenge ROS and mitigate oxidative damage. To assess in vivo biocompatibility, H&E staining of major organs (liver, spleen, lung, kidney) revealed intact histological architectures across all groups (Fig. S3), including ordered hepatic lobules, clear alveolar spaces, normal glomerular morphology, and distinct splenic red/white pulp boundaries, with no pathological abnormalities such as inflammatory infiltration, cellular edema, or fibrosis, demonstrating that the CPH hydrogel had good biocompatibility. Furthermore, the CPH hydrogel demonstrated excellent hemocompatibility with a hemolysis rate of 1.84 ± 0.04%, which is well below the 5% safety threshold for biomedical materials (Fig. S4).

To investigate the cardioprotective effects of the CPH hydrogel on CMs, immunofluorescence analysis was employed to assess the localization and expression levels of key structural proteins in myocardial tissue, specifically Actin and connexin 43 (Cx43). Actin maintains cellular morphology and mechanical integrity as a core cytoskeletal component, while Cx43 mediates intercellular electrical coupling. As shown in Fig. 2(F), H₂O₂ treatment reduced Actin red fluorescence intensity and disrupted filament alignment, whereas the CPH group maintained intact Actin architecture with fluorescence intensity close to the Control group (Fig. 2(G)). Concurrently, Cx43 green fluorescence at lateral membranes was markedly attenuated in the H₂O₂ group, whereas the CPH group exhibited Cx43 expression intensity and a typical stepwise distribution close to the Control group (Fig. 2(H)). These findings indicated that the CPH hydrogel effectively decreased oxidative stress-induced suppression of Cx43 and Actin expression, preserving cytoskeletal stability and electromechanical synchronization to protect CMs against oxidative damage.

3.2 Anti-apoptotic effects of the CPH hydrogel in vivo

Based on in vivo experimental results, we further validated the CPH hydrogel efficacy in the rat MI model. DHE staining (Fig. 3(A)) revealed a superoxide anion surge in the infarct zone of the MI group as early as day 1. The CPH group exhibited 1.3-fold (***P < 0.001) and 2.0-fold (***P < 0.001, Fig. 3(B)) reductions in ROS levels at days 1 and 3, respectively, compared to the MI group, confirming sustained ROS scavenging capacity.

Fig. 3 The CPH hydrogel intervention in oxidative stress-induced cellular damage and apoptosis. (A) Representative fluorescence images of DHE in the infarct zone on day 1 and day 3. (B) Statistical evaluation of DHE staining. (C) Immunofluorescence staining images of 8-OHdG (red) and cTnT (green) in the infarct zone on day 3 post-surgery. (D) Statistical evaluation of fluorescence intensity of 8-OHdG. (E) Statistical evaluation of fluorescence intensity of cTnT. (F) Representative fluorescence image of TUNEL staining in the infarct zone on day 3 post-surgery. (G) Statistical evaluation of the fluorescence area of TUNEL staining. (n = 3, ***P < 0.001).

To assess the impact of the CPH hydrogel on oxidative DNA damage, immunofluorescence staining was performed to assess 8-hydroxydeoxyguanosine (8-OHdG, an oxidative stress marker) expression in myocardial infarct zones.²⁶ As shown in Fig. 3(C), the Sham group exhibited intact myocardial architecture with minimal 8-OHdG fluorescence (intensity: 77.2 ± 0.4), whereas the MI group displayed structural disruption and intense 8-OHdG signals, indicating aggravated oxidative damage. The CPH group demonstrated a 29.5% reduction in 8-OHdG signal compared to the MI group (***P < 0.001, Fig. 3(D)), while the cTnT signal increased by 48.7% (***P < 0.001, Fig. 3(E)), with preserved myocardial organization. Furthermore, TUNEL staining revealed that apoptotic CMs accounted for 20.30 ± 0.36% of the infarct zone in the MI group at postoperative day 3, which was reduced to 1.50 ± 0.05% (***P < 0.001) in the CPH group (Fig. 3(F) and (G)). Molecular mechanism studies have shown that ROS-induced 8-OHdG accumulation activated the p53-Bax pathway to trigger mitochondria-dependent apoptosis, concurrent with NF-κB pathway activation, exacerbating inflammatory responses.^27,28 By scavenging ROS and reducing 8-OHdG levels, the CPH hydrogel abolished these cascades, ultimately attenuating CMs apoptosis and limiting infarct expansion.

3.3 Regulation of macrophage polarization and inflammation by the CPH hydrogel

During the initial stages of MI, the activation of oxidative stress and inflammatory responses serves as a central mechanism contributing to ischemic damage in cardiac tissue.²⁹ Macrophages, as key immune effector cells, are rapidly recruited and activated in infarcted myocardium, releasing pro-inflammatory cytokines to exacerbate tissue damage. Pro-inflammatory M1 macrophages dominate inflammatory cascades, whereas reparative M2 macrophages facilitate tissue remodeling.^30,31 This study evaluated the regulatory effects of the CPH hydrogel on macrophage polarization using an in vitro ROS microenvironment model (200 µmol L⁻¹ H₂O₂ treatment). Bone marrow-derived macrophages (BMDMs) were isolated from mice and subsequently underwent a 7-day differentiation protocol (Fig. S5). Flow cytometry and immunofluorescence confirmed BMDM purity: F4/80⁺ cells accounted for 96.1% of the population, with CD68⁺ cells at 94.9% (Fig. S6), consistent with macrophage phenotypic characteristics. Parallel immunofluorescence staining demonstrated robust F4/80 and CD68 signals (Fig. S7), validating successful BMDM isolation. BMDMs were treated with H₂O₂ or H₂O₂ and 1 mg mL⁻¹ CPH hydrogel for 24 h. The fluorescence imaging (Fig. S8) revealed that the H₂O₂ treatment significantly upregulated iNOS expression. In contrast, the CPH hydrogel reduced iNOS expression by 1.3-fold (***P < 0.001, Fig. S9) and elevated CD206 expression by 1.8-fold (***P < 0.001, Fig. S10) in comparison with the H₂O₂ group. These data demonstrated that the CPH hydrogel suppressed ROS-induced M1 polarization while promoting macrophage transition toward the M2 phenotype.

To elucidate the role of cardiac macrophages in MI pathophysiology, this study focused on their phenotypic dynamics and intervention strategies. Resident cardiac macrophages constitute 7%–8% of non-cardiomyocytes and play dual roles in immune surveillance and homeostasis regulation, while also serving as pivotal regulators of post-MI inflammation and fibrosis.³² During the acute phase (days 1–3), pro-inflammatory M1 macrophages dominate the immune response by secreting TNF-α, interleukin-6, and matrix metalloproteinases to clear necrotic debris and degrade extracellular matrix (ECM). Although M1 polarization is initially necessary for tissue repair, its sustained activation exacerbates infarct expansion and delays inflammation resolution, leading to pathological scar formation.³⁰ During the transitional phase (days 3–5), anti-inflammatory M2 macrophages increase significantly, promoting inflammation resolution and tissue remodeling via IL-10 and transforming growth factor-β secretion.³³ The M1-to-M2 phenotypic switch is a critical hallmark of post-MI inflammation resolution, making macrophage polarization modulation a promising therapeutic target to mitigate myocardial injury. To assess the CPH hydrogel's impact on macrophage polarization, immunofluorescence at day 5 post-MI revealed that the CPH hydrogel induced a 2.4-fold increase in CD206⁺ cells compared to the MI group (***P < 0.001, Fig. 4(A) and (B)), and an 11.7-fold increase in iNOS⁺ cells in the infarct zone of the MI group versus the Sham group (***P < 0.001), whereas the CPH group exhibited an 81.8% reduction compared to the MI group (***P < 0.001, Fig. 4(A) and (C)), indicating robust M2 polarization. Immunohistochemistry further confirmed that the CPH hydrogel reduced TNF-α expression by 61.2% (***P < 0.001, Fig. 4(D) and (E)) and upregulated IL-10 by 67.1% (***P < 0.001, Fig. 4(D) and (F)) relative to the MI group. These findings demonstrated that the CPH hydrogel attenuated early-phase hyperinflammation by suppressing M1 polarization and promoting M2 conversion, offering a novel strategy to enhance myocardial repair.

Fig. 4 Regulation of macrophage polarization and inflammation by the CPH hydrogel. (A) Representative fluorescence images of iNOS (green) and CD206 (red) in the infarct zone on day 5. (B) Statistical evaluation of CD206 expression. (C) Statistical evaluation of iNOS expression. (D) Immunohistochemical detection of TNF-α and IL-10 in the infarct zone on day 5. (E) Statistical evaluation of TNF-α expression. (F) Statistical evaluation of IL-10 expression. (n = 3, *P < 0.05, **P < 0.01, ***P < 0.001).

3.4 The CPH hydrogel contribution to myocardial repair

The pathological progression of MI exhibits dynamic spatiotemporal characteristics (Fig. 5(A)), necessitating stage-specific therapeutic interventions. During the acute phase (days 1–5), priority is given to mitigating oxidative stress and suppressing CMs apoptosis via ROS scavenging, macrophage polarization modulation, and anti-inflammatory mechanisms.^31,34 In the chronic phase (>28 days), the focus shifts to tissue remodeling and functional recovery, emphasizing fibrosis suppression and angiogenesis to synergistically improve cardiac outcomes.^35,36 To evaluate the long-term efficacy of the CPH hydrogel, cardiac morphology and molecular pathology were analyzed at 28 days post-MI. The CPH group exhibited a 60.5% reduction in infarct size (***P < 0.001, Fig. 5(B) and (C)) and increased left ventricular wall thickness (from 0.960 ± 0.011 mm to 2.150 ± 0.042 mm) in comparison with the MI group (***P < 0.001, Fig. 5(B) and (D)). H&E staining confirmed attenuated ventricular remodeling in the CPH group, with myocardial architecture resembling the Sham controls. Regarding the degree of myocardial fibrosis, the three groups showed significant differences. As shown by Masson's trichrome staining, the Sham group had intact myocardial tissue structure with only minimal collagen fiber deposition, and the fibrotic area accounted for 6.90 ± 0.59%. In stark contrast, the MI group's infarct region displayed extensive blue collagen deposition, with the fibrotic area significantly increased to 33.97 ± 0.58%, indicating successful model establishment and the induction of severe pathological fibrosis. The CPH group, however, exhibited a notable anti-fibrotic effect, with the fibrotic area reduced to 18.95 ± 0.43%, representing an approximately 44% decrease compared with the MI group (Fig. S11).

Fig. 5 Histological analysis of cardiac structure 28 days postoperatively. (A) Timeline of myocardial repair process of the CPH hydrogel in the MI region. (B) Characteristic morphological features revealed by H&E and Masson trichrome staining. (C) Statistical evaluation of myocardial infarction area. (D) Statistical evaluation of left ventricular wall thickness. (n = 5, ***P < 0.001).

In addition, Immunofluorescence further revealed that in the CPH group (Fig. 6(A)), the striated structure of cardiomyocyte α-actinin 2 (ACTN2) was clear, with an expression level 1.19-fold that of the MI group (***P < 0.001, Fig. 6(B)). In addition, the expression level of cTnT in the CPH group was 1.22-fold that of the MI group (***P < 0.001, Fig. 6(C) and (D)), and the expression level of Cx43 was 1.67-fold that of the MI group (***P < 0.001, Fig. 6(C) and (E)), suggesting that it promoted cardiac repair by remodeling the intercellular electrical conduction network. These therapeutic outcomes likely stemmed from the CPH hydrogel's biphasic mechanisms: early-phase ROS clearance preserved CMs viability, while macrophage polarization modulation and enhanced gap junction formation synergistically drove myocardial repair. WGA staining (Fig. 6(F)) revealed that the CPH hydrogel restored cardiomyocyte cross-sectional area to the Sham levels (***P < 0.001, Fig. 6(G)), with preserved membrane integrity, effectively suppressing pathological hypertrophy. To assess multidimensional effects, vascularization was evaluated. Immunofluorescence (Fig. 6(H)) showed a 95.9% increase in CD34⁺ neovessel density (***P < 0.001, Fig. 6(I)) and a 2.42-fold elevation in α-SMA⁺ mature microvessels (***P < 0.001, Fig. 6(J)) in the CPH group, suggesting robust angiogenic maturation, potentially mediated by VEGF upregulation. Collectively, the CPH hydrogel promoted myocardial repair and mitigated adverse remodeling by orchestrating angiogenesis, fibrosis resolution, and electromechanical restoration.

Fig. 6 Myocardial repair at 28 days after surgery. (A) Representative fluorescence image of ACTN2 (red) in the MI region. (B) Statistical evaluation of fluorescence intensity of ACTN2. (C) Representative fluorescence images of the MI region, cTnT (red) and Cx43 (green). (D) Statistical evaluation of fluorescence intensity of cTnT. (E) Statistical evaluation of fluorescence intensity of Cx43. (F) Representative fluorescence images of WGA (red) and Actin (green) in the MI region. (G) Statistical assessment of cardiomyocyte cross-sectional area. (H) Representative fluorescence images of α-SMA (red) and CD34 (green) in the MI region. (I) Statistical evaluation of fluorescence intensity of α-SMA. (J) Statistical evaluation of fluorescence intensity of CD34. (n = 3, *P < 0.05, ***P < 0.001).

3.5 Recovery of cardiac function by the CPH hydrogel

MI disrupts coronary blood flow, inducing ischemia, CMs death, and inflammatory cascades, ultimately leading to fibrotic scar formation, electrical conduction impairment, and irreversible cardiac dysfunction.^37,38 At postoperative day 28, electrocardiography (Fig. 7(A)) revealed significant ST-segment elevation in the MI group, and the QRS interval was prolonged to 45.0 ± 2.2 ms (Fig. 7(B)), suggesting the presence of conduction block. In contrast, the CPH group exhibited normalized QRS intervals, demonstrating improved electrophysiological stability. The echocardiography (Fig. 7(C)) further revealed that in the MI group, the left ventricular ejection fraction (LVEF) and left ventricular fractional shortening (LVFS) decreased to 37.13 ± 4.97% and 20.28 ± 3.66%, with left ventricular end-diastolic diameter (LVDD) and end-systolic diameter (LVDS) significantly expanded to 8.68 ± 0.41 mm and 7.25 ± 0.76 mm, respectively. Following the CPH hydrogel intervention, LVEF improved to 59.33 ± 3.45% (**P < 0.01), LVFS recovered to 36.11 ± 2.08% (**P < 0.01), while LVDD and LVDS decreased to 6.96 ± 0.19 mm (**P < 0.01) and 4.91 ± 0.20 mm (**P < 0.01, Fig. 7(D)–(G)). These results demonstrated that the CPH hydrogel achieved comprehensive myocardial spatiotemporal repair, attenuating ventricular dilation, enhancing electromechanical synchrony, and restoring pump function. In addition, the CPH hydrogel offered a novel cell-free therapeutic strategy for cardiac regeneration, without reliance on stem cells or drug delivery.

Fig. 7 Cardiac function assessment at 28 days postoperatively. (A) Electrocardiographic recordings of distinct groups of rats. (B) Echocardiographic imaging of rats from distinct groups. (C) Duration of the QRS complex. (D) LVEF measurement and analysis. (E) LVFS measurement and analysis. (F) LVDD measurement and analysis. (G) LVDS measurement and analysis. (n = 5, **P < 0.01, ***P < 0.001).

4. Conclusion

To address the clinical challenge of insufficient myocardial repair post-MI, this study developed an injectable CPH hydrogel based on the integration of CMCS and DS, designed to synergistically intervene in post-infarction pathological cascades through multidimensional mechanisms. The CPH hydrogel, constructed via dynamic crosslinking technology, formed a biomimetic three-dimensional scaffold that replicated the mechanical properties of the native cardiac extracellular matrix, providing structural support for CMs alignment and synchronized electrical signal conduction. Its core innovation lay in the integration of multifunctional modules, including antioxidant activity, anti-inflammatory effects, pro-angiogenic potential, and enhanced electroactive properties. In vitro experiments demonstrated that the CPH hydrogel effectively scavenged ROS, significantly improved the viability of CMs under oxidative stress, and preserved cytoskeletal integrity and electrical coupling by maintaining Cx43 and Actin expression. Concurrently, the CPH hydrogel regulated macrophage phenotypic polarization through inhibition of pro-inflammatory M1 macrophages and simultaneous enhancement of anti-inflammatory M2 macrophages, resulting in effective mitigation of inflammatory cascade reactions. In the rat MI model, the CPH hydrogel exhibited robust therapeutic efficacy: early-stage intervention reduced CMs apoptosis and oxidative DNA damage by ROS clearance, effectively limiting infarct expansion; late-stage therapy enhanced myocardial structural integrity and electrophysiological stability through angiogenesis and collagen remodeling. By postoperative day 28, echocardiography revealed that the CPH hydrogel elevated LVEF and LVFS to 59.33% and 36.11%, respectively, with marked reversal of ventricular dilation, culminating in cardiac spatiotemporal repair. This study not only presents a novel cell-free therapeutic strategy with dual bioactivity and functional adaptability for MI treatment but also establishes a theoretical framework for the rational design of cardiac tissue engineering materials, underscoring its significant potential for clinical translation.

5. Discussion

Following MI, a substantial proportion of CMs undergo irreversible injury or death within a short timeframe. Ischemia-induced hypoxia triggers an overproduction of ROS, leading to excessive oxidative stress. This pathological state is characterized by oxidative modifications to proteins, peroxidation of lipids, and damage to DNA structures, which collectively exacerbate inflammatory cascades.^39–41 Persistent post-MI inflammation promotes aberrant collagen deposition, aggravates fibrosis and adverse left ventricular remodeling, and significantly impedes cardiac functional recovery.^42,43 Thus, mitigating oxidative stress and reprogramming the inflammatory microenvironment represent pivotal therapeutic targets. The pathological progression in the infarct zone originates from ROS overgeneration. ROS activates the Toll-like receptor 4 and NF-κB pathway, triggering the excessive release of pro-inflammatory cytokines and establishing a self-perpetuating inflammatory cycle.⁴⁴ During the inflammatory peak (3–5 days post-MI), monocyte and macrophage infiltration exacerbates cell death and ECM degradation,^45,46 further amplifying tissue injury. Modulating macrophage polarization emerges as a critical strategy to attenuate inflammation: pro-inflammatory M1 macrophages exacerbate oxidative stress via ROS secretion, while reparative M2 macrophages secrete anti-inflammatory factors and pro-angiogenic mediators, facilitating ECM reconstruction, cellular proliferation, and angiogenesis to drive tissue repair.³⁰

The CPH hydrogel developed in this study integrates CMCS, DS, and ODex to achieve synergistic regulation of antioxidant activity, anti-inflammatory effects, enhanced electrical signal conduction, and vascular regeneration. Unlike stem cell-based therapies, the CPH hydrogel operates as a cell-free platform, circumventing immune rejection and ethical concerns while directly modulating the pathological microenvironment through physical support and dynamic biochemical regulation, offering a streamlined pathway for clinical translation. However, limitations persist: the rat MI model may overestimate therapeutic efficacy due to the absence of human-like coronary collateral circulation, and the long-term biological impact of hydrogel degradation byproducts remains uncharacterized. Future studies should validate efficacy in large animal models, elucidate hydrogel degradation kinetics in relation to tissue remodeling, and delineate molecular mechanisms underlying macrophage polarization via transcriptomic or proteomic approaches.⁴⁷

Future research should prioritize clinical translation and functional optimization, including the development of minimally invasive catheter-based delivery systems, incorporation of photothermal-responsive moieties or gene carriers for spatiotemporal control of ROS scavenging, anti-inflammatory action, and regeneration,⁴⁸ as well as the design of stratified hydrogels for stage-specific release of VEGF, microRNA, or antifibrotic agents tailored to pathological phases (acute ischemia, subacute inflammation resolution, chronic fibrosis).⁴⁹ Furthermore, synergistic strategies combining the CPH hydrogel with cardiac patches, bioprinting, or electrical stimulation could overcome current therapeutic bottlenecks.^48,50 Integrating drug delivery, gene editing, and exosome-based therapies may unlock the full potential of the CPH hydrogel as a multifunctional platform, paving the way for personalized approaches in myocardial regenerative medicine.

Author contributions

Jiajun Lu: data curation, formal analysis, writing – original draft. Jing Chen: methodology, conceptualization. Linken Lu: methodology. Yating Zhao: conceptualization. Ruiqi Liu: data arrangement. Yanru Li: conceptualization. Chenguang Liu: conceptualization. Pengcheng Che: conceptualization. Hong Sun: project administration, funding acquisition, writing, review, and editing.

Conflicts of interest

The authors declare no competing financial interest.

Data availability

The data supporting this article have been included in the experimental part.

Supplementary information is available. See DOI: https://doi.org/10.1039/d5tb02290h.

Acknowledgements

This work was financially supported by the National College Student Innovation and Entrepreneurship Training Program (202410081057). Hebei Key Laboratory for Rehabilitation Engineering and Regenerative Medicine (SZX202327). Natural Science Foundation of Hebei Province (no. H2024209013 & H2023209021).

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