GelMA‐Based Hydrogel Encapsulating Astragalus Polysaccharides and Umbilical Cord Mesenchymal Stem Cell Exosomes Promotes Full‐Thickness Skin Wound Healing

ABSTRACT

Background

Wound healing involves excessive oxidative stress and cellular dysfunction, while current single therapies remain inadequate. Astragalus polysaccharides (APS) and exosomes (Exos) possess antioxidative and regenerative properties, and gelatin methacryloyl (GelMA) hydrogel provides a biocompatible platform for localized delivery. Here, we developed an APS–Exos@GelMA hydrogel to promote wound repair.

Methods

The biological effects of APS and Exos were assessed through in vitro cellular assays, while the physicochemical properties and biocompatibility of APS–Exos@GelMA were characterized. Its therapeutic efficacy was further evaluated in a full‐thickness skin defect model through macroscopic observation and histological analysis.

Results

APS–Exos treatment showed enhanced effects compared with APS or Exos alone, improving fibroblast (130.90% ± 2.98%, p < 0.05) and keratinocyte (124.50% ± 3.18%, p < 0.01) viability, promoting fibroblast migration (74.15% ± 7.72%), and reducing oxidative damage (72.11% ± 2.13%). APS–Exos@GelMA maintained favorable hydrogel properties and biocompatibility while enabling localized incorporation and application of APS and Exos. In vivo, APS–Exos@GelMA accelerated wound closure, promoted epidermal reconstruction and extracellular matrix deposition, and exhibited good biosafety.

Conclusion

The combination of APS and Exos within an appropriate concentration range demonstrated improved therapeutic potential compared with individual treatments. The APS–Exos@GelMA hydrogel served as a biocompatible localized delivery platform to facilitate tissue repair, highlighting its potential for wound‐healing applications.

Highlights

• GelMA hydrogel was engineered as a localized delivery platform for APS and HUCMSC‐Exos.

• The combined application of APS and HUCMSC‐Exos enhanced cell viability, promoted cell migration, and partially alleviated cellular resistance to oxidative stress in vitro.

• APS–Exos@GelMA hydrogel accelerated wound healing and enhanced tissue regeneration in a full‐thickness skin defect model.

1. Introduction

The skin serves as a critical barrier protecting the body against external environmental insults. Upon injury, it undergoes a highly coordinated healing process consisting of hemostasis, inflammation, proliferation, and remodeling [1]. However, non‐healing wounds fail to proceed through the normal healing timeline due to underlying conditions such as diabetes, vascular diseases, or infection, exemplified by diabetic foot ulcers and pressure ulcers [2]. These wounds are typically characterized by oxidative stress imbalance and impaired cellular functions, which severely compromise patients’ quality of life and impose substantial healthcare burdens [3]. In chronic wounds, the physiological healing cascade is profoundly disrupted, with the wound microenvironment remaining in a dysregulated state and failing to transition into effective tissue repair. Therefore, regulating the wound microenvironment, restoring cellular function, and promoting tissue regeneration are critical challenges in chronic wound management.

Recently, natural bioactive compounds derived from medicinal plants have gained increasing attention in chronic wound therapy. Astragalus polysaccharides (APS), one of the major active components of Astragalus membranaceus, exhibits anti‐inflammatory [4], antioxidant [5], and pro‐angiogenic properties [6]. Accumulating evidence indicates that APS possesses anti‐inflammatory and antioxidant properties, enhances fibroblast function, and promotes wound healing, demonstrating promising therapeutic potential [7].

Meanwhile, exosome‐based cell‐free therapies have emerged as a promising strategy in regenerative medicine. Exosomes are nanosized vesicles (30–150 nm) secreted by cells, carrying proteins, lipids, and microRNAs, and playing a pivotal role in intercellular communication. Mesenchymal stem cell‐derived exosomes (MSC‐Exos) have been reported to facilitate skin wound healing through multiple biological activities, including enhancing cellular proliferation and migration, regulating oxidative stress responses, and promoting tissue regeneration [8, 9, 10]. However, their clinical application is limited by rapid diffusion and degradation in vivo, resulting in poor retention and low bioavailability at wound sites [11].

To address these limitations, biomaterial‐based delivery systems provide a promising approach to enhance therapeutic efficacy [12]. Currently, various delivery systems have been developed and widely applied in the fields of tissue engineering and regenerative medicine, including natural polymer‐based hydrogels such as gelatin and hyaluronic acid, synthetic polymer‐based hydrogels such as polyethylene glycol (PEG) and its derivatives, as well as poly(lactic‐co‐glycolic acid) (PLGA) nanoparticle‐based delivery systems [13]. Notably, although these delivery platforms have partially addressed the limitations associated with bioactive molecules, including rapid degradation and insufficient retention at target sites, several challenges remain. For instance, natural polymer‐based hydrogels often exhibit relatively weak mechanical properties and excessive swelling behavior, which may restrict their broader applications [14]. In contrast, synthetic hydrogels generally offer excellent tunability but often lack the inherent bioactive signals present in natural extracellular matrix components [12].

Among these biomaterials, gelatin methacryloyl (GelMA) hydrogels have attracted increasing attention due to their combination of natural bioactivity and tunable properties [15]. GelMA is derived from methacryloyl‐modified gelatin, retaining bioactive motifs such as RGD sequences and matrix metalloproteinase‐sensitive sites while enabling the formation of ECM‐like three‐dimensional networks through photocrosslinking. These structures provide a favorable microenvironment for cell adhesion, proliferation, and functional maintenance. Moreover, GelMA exhibits excellent biocompatibility, controllable degradability, and tunable mechanical properties, making it a promising platform for localized delivery of bioactive molecules in tissue engineering and regenerative medicine applications [14, 16, 17, 18].

Based on this, a GelMA‐based composite hydrogel was constructed and loaded with human umbilical cord mesenchymal stem cell‐derived exosomes (HUCMSC‐Exos) and APS, aiming to achieve their combined application, with GelMA serving as an effective platform for localized delivery. This system was designed to improve the wound microenvironment by enhancing cellular functions, alleviating oxidative stress, and promoting tissue regeneration. Its therapeutic efficacy was systematically evaluated through in vitro cellular experiments and in vivo wound‐healing models, aiming to provide a promising strategy and theoretical basis for the development of advanced biomaterial‐based approaches for wound repair.

2. Materials and Methods

2.1. Cells and Animal

Human umbilical cord mesenchymal stem cells (HUCMSCs) were obtained from BioAngle (Xiamen) Technology Co., Ltd. and cultured in a specialized stem cell medium according to the manufacturer's instructions. Human keratinocyte HaCaT cells and mouse fibroblast L929 cells were purchased from the American Type Culture Collection (ATCC, USA). Both cell lines were maintained in Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS; Gibco, USA) and 1% penicillin–streptomycin. Cells were cultured in a humidified incubator at 37°C with 5% CO₂. Specific pathogen‐free (SPF) male C57BL/6J mice were purchased from Jiangsu Jicui Yaokang Biotechnology Co. Ltd.

2.2. Culture and Characterization of HUCMSCs

HUCMSCs (passages 3–5) were cultured in 60 mm dishes and observed under an inverted microscope. When reaching 85%–90% confluence, cells were digested, resuspended, and adjusted to 1 × 10⁶ cells/tube. Surface markers were analyzed by flow cytometry after incubation with antibodies against CD73, CD90, CD105 (Elabscience, Wuhan, China) and CD34, CD45, HLA‐DR, CD19, CD11b (Elabscience, Wuhan, China), using isotype controls.

For differentiation assays, cells were induced toward adipogenic, osteogenic, and chondrogenic lineages using standard induction media. Adipogenesis was confirmed by Oil Red O (Solarbio, Beijing, China) staining after 10–14 days. Osteogenesis was evaluated by Alizarin Red S (Solarbio, Beijing, China) staining after 21–28 days. Chondrogenesis was assessed using pellet culture followed by Alcian Blue (Solarbio, Beijing, China) staining after 21–28 days.

2.3. Isolation and Characterization of HUCMSC‐Exosomes

HUCMSC‐derived exosomes (HUCMSC‐Exos) were isolated from conditioned medium by differential ultracentrifugation. The supernatant was sequentially centrifuged at 2000 × g for 10 min and 10,000 × g for 30 min to remove cells and debris, followed by ultracentrifugation at 100,000 × g for 75 min. The pellet was washed with PBS and re‐centrifuged under the same conditions. The final exosome pellet was resuspended in PBS and stored at −80°C.

Nanoparticle tracking analysis (NTA; Particle Metrix GmbH, Germany) was used to determine particle size and concentration, and transmission electron microscopy (TEM; HT7700, Hitachi, Japan) was used to observe morphology.

For protein characterization, exosomal proteins were extracted using RIPA buffer (Beyotime, Shanghai, China). Proteins were separated by SDS‐PAGE and transferred to PVDF membranes. After blocking with 5% skim milk, membranes were incubated overnight at 4°C with primary antibodies against CD9, CD81, TSG101, and Calnexin (1:1000; ABclonal, Wuhan, China), followed by HRP‐conjugated secondary antibodies (1:3000; ZSGB‐BIO, Beijing, China). Protein bands were visualized using ECL reagents (Solarbio, Beijing, China) and detected with a UVITEC imaging system.

2.4. HUCMSC‐Exos Uptake Assay

HUCMSC‐Exos were labeled with PKH67 fluorescent dye (Beyotime, Shanghai, China) following the manufacturer's protocol. After labeling and quenching, exosomes were incubated with HaCaT and L929 cells. Cellular uptake was evaluated using a confocal laser scanning microscope after incubation with labeled exosomes for 6 h in HaCaT cells and 24 h in L929 cells, respectively.

2.5. Screening of APS and HUCMSC‐Exos Concentrations

Cell viability was assessed using a CCK‐8 assay (Beyotime, Shanghai, China). L929 fibroblasts (5 × 10³ cells/well) were seeded in 96‐well plates and allowed to attach overnight. Cells were then treated with different concentrations of APS (0–200 μg/mL; Maclin, Shanghai, China) or HUCMSC‐Exos (0–40 μg/mL) for 24 h. For comparison, cells were also treated with DMEM, APS, Exos (10 μg/mL), Exos (20 μg/mL), or APS‐Exos. Subsequently, 10 μL of CCK‐8 reagent was added to each well and incubated for 2 h. Absorbance was measured at 450 nm using a microplate reader.

2.6. Cell Migration Assay

Cell migration was evaluated using a scratch wound assay. L929 cells were seeded in 24‐well plates and cultured to confluence at 37°C and 5% CO₂. Cells were serum‐starved for 12 h prior to scratching. A linear scratch was created using a sterile 200 μL pipette tip, and detached cells were removed with PBS washing. Cells were then treated with different media (DMEM, APS, Exos, or APS‐Exos). Images were captured at 0 and 24 h using an inverted microscope (Olympus, Tokyo, Japan). Wound area was quantified using ImageJ software (NIH, Bethesda, MD, USA), and migration rate was calculated.

2.7. ROS Scavenging Assay

L929 cells were seeded at 1 × 10⁵ cells/mL in 24‐well plates and cultured at 37°C, 5% CO₂ until adherence. Oxidative stress was induced by treating cells with Rosup reagent (Beyotime, Shanghai, China) for 30 min. After washing with PBS, cells were incubated with DMEM, Exos, APS, or APS‐Exos for 6 h. Intracellular ROS levels were detected using DCFH‐DA fluorescent probe (Beyotime, Shanghai, China) and imaged with an inverted fluorescence microscope.

2.8. HaCaT Cell Viability and Antioxidant Assay

HaCaT cells were seeded in 96‐well plates and cultured at 37°C, 5% CO₂ until attachment. To evaluate the effects of APS and Exos on cell viability, cells were treated with APS (25, 75, 200 μg/mL), Exos, or APS‐Exos for 24 h. Cell viability was assessed using a CCK‐8 assay by adding 10 μL reagent per well, incubating for 2 h in the dark at 37°C, and measuring absorbance at 450 nm using a microplate reader.

Antioxidant capacity under oxidative stress was evaluated using CCK‐8 and Calcein‐AM live‐cell staining (Beyotime, Shanghai, China). Cells were pretreated with DMEM, Exos, APS, or APS‐Exos for 6 h, followed by H₂O₂ (1 mmol/L) for 4 h. Cell viability was measured by CCK‐8 at 450 nm, and live‐cell fluorescence was visualized using an inverted fluorescence microscope.

2.9. Preparation and Characterization of GelMA Hydrogels

GelMA hydrogels (EFL, Suzhou, China) were prepared by photopolymerization. Lithium phenyl‐2,4,6‐trimethylbenzoylphosphinate (LAP; EFL, Suzhou, China) was dissolved in sterile PBS and mixed with GelMA to prepare 10% (w/v) prepolymer solution at 60°C–70°C. The solution was sterilized through a 0.22 μm filter and stored protected from light. For functionalized hydrogels, APS or Exos were gently mixed into GelMA prepolymer, followed by UV crosslinking (405 nm, 30 s) to form composite hydrogels.

Hydrogel microstructure was observed by scanning electron microscopy (SEM; SU8010, Hitachi, Japan) after lyophilization and gold sputtering. Swelling behavior was evaluated by immersing dried hydrogels in PBS at 37°C for 24 h. Swelling ratio was calculated as:

Swelling ratio (%) = (W_t  − W ₀)/W ₀ × 100%,

where W_t represents the wet weight after 24 h of swelling, and W ₀ represents the initial dry weight.

For degradation assessment, hydrogels were incubated in PBS at 37°C. Samples were collected on days 1, 4, 7, and 10, lyophilized, and weighed (W_t ) to evaluate the degradation rate. Remaining weight was calculated as:

Remaining weight (%) = W_t /W ₀ × 100%,

where W_t represents the dry weight after t days of degradation, and W ₀ represents the initial dry weight.

To mimic the enzymatic degradation environment in vivo, APS‐Exos@GelMA hydrogels were incubated in PBS containing 1 U/mL collagenase type II. At predetermined time points, the hydrogels were removed and weighed to assess their degradation behavior. Degradation rate was calculated as:

Degradation rate (%) = (W ₀ − W_t )/W ₀ × 100%,

where W_t represents the dry weight after t hours of degradation, and W ₀ represents the initial dry weight.

2.10. Exosome Release Profile

PKH67‐labeled HUCMSC‐Exos were prepared according to the instructions, followed by ultracentrifugation to remove excess dye. The purified PKH67‐labeled HUCMSC‐Exos were then incorporated into GelMA hydrogels to evaluate exosome release behavior. The prepared hydrogels were immersed in PBS and incubated at 37°C under gentle shaking. At predetermined time intervals, the release medium was collected and replaced with an equal volume of fresh PBS. The fluorescence intensity of the collected samples was measured using a microplate reader to quantify the released exosomes.

2.11. Hemolysis Assay

Hydrogels (0.2 g/mL) were incubated in sterile PBS at 37°C for 24 h to obtain hydrogel extracts. Fresh anticoagulated whole blood was washed with PBS and centrifuged to remove plasma and leukocytes, then diluted to 2% erythrocyte suspension. Equal volumes of hydrogel extract and erythrocyte suspension were mixed and incubated at 37°C for 3 h. After centrifugation (845 × g, 10 min), the supernatant was transferred to a 96‐well plate, and absorbance at 545 nm was measured using a microplate reader to calculate hemolysis.

2.12. Live/Dead Cell Staining

To evaluate cytocompatibility, L929 cells were seeded in 24‐well plates and indirectly co‐cultured with pre‐crosslinked hydrogels at 37°C, 5% CO₂ for 48 h. Cells were stained using a Calcein‐AM/PI viability/cytotoxicity kit (Beyotime, Shanghai, China) and imaged with an inverted fluorescence microscope to assess live (green) and dead (red) cells.

2.13. Animal Wound Model and Treatment

Eight‐ to ten‐week‐old male C57BL/6 J mice were acclimated for 1 week and randomly divided into four groups (n = 6): PBS, GelMA, Exos@GelMA, and APS‐Exos@GelMA. The final concentration of HUCMSC‐Exos in the GelMA hydrogel was adjusted to 20 μg/mL for animal studies. Mice were anesthetized and dorsal hair removed under sterile conditions. Two full‐thickness skin wounds (6 mm diameter) were created on the back using a sterile biopsy punch. Treatments were applied immediately: PBS group received equal volume PBS; hydrogel groups received 60 μL of corresponding hydrogel applied to the wound and crosslinked in situ under 405 nm light for 30 s. Wounds were covered with sterile dressings. Wound healing was photographed on days 0, 7, and 12. Wound area was quantified using ImageJ software, and healing rate was calculated as:

Wound closure (%) = (A ₀ − A_t )/A ₀ × 100%,

where A ₀ represents the wound area on day 0, and A_t represents the wound area on day t.

2.14. Tissue Collection and Histological Analysis

On day 12, mice were euthanized, and wound tissues with surrounding skin were collected. Major organs (heart, liver, spleen, lung, and kidney) were harvested for biosafety evaluation. Tissues were fixed in 4% paraformaldehyde for 24 h, washed, dehydrated through graded ethanol (70%–100%), cleared with xylene, and embedded in paraffin. Section (4–5 μm) were prepared, mounted on slides, and dried. For H&E staining, sections were deparaffinized with xylene and rehydrated through graded ethanol. Slides were stained with hematoxylin for 5–10 min, differentiated with acid alcohol, blued with ammonia water, and counterstained with eosin for 1–3 min. After dehydration and clearing, sections were mounted with neutral resin and imaged under a light microscope.

Masson's trichrome staining was performed as follows: sections were stained with Weigert's iron hematoxylin for 5–10 min, washed, differentiated, and blued. Ponceau‐fuchsin was applied for 5–10 min, followed by differentiation with phosphomolybdic or phosphotungstic acid and counterstaining with aniline blue for 5 min. Sections were treated with 1% acetic acid to enhance staining, dehydrated, cleared with xylene, mounted with neutral resin, and imaged under a light microscope.

2.15. Statistical Analysis

All experimental data were obtained from three independent replicate experiments. Flow cytometry data were analyzed using FlowJo software, while cell migration assays and in vivo wound healing quantitative analyses were performed using ImageJ software. Statistical analyses were performed using GraphPad Prism 9.1.0 software (GraphPad Software, San Diego, CA, USA). Data were first tested for normality and homogeneity of variance. For data with a normal distribution and equal variances, one‐way ANOVA was applied, followed by Tukey's post hoc test for multiple comparisons, and results were presented as mean ± SD. For non‐normally distributed data, the Kruskal–Wallis test followed by Dunn's post hoc multiple comparisons test was used, and results were expressed as median (interquartile range) [M (Q1, Q3)]. Asterisks (*p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001) indicate that the differences between comparison groups are statistically significant.

3. Result

3.1. HUCMSC Identification

Passage 3–5 HUCMSCs exhibited adherent short spindle or fibroblast‐like morphology (Figure 1A). Flow cytometry analysis revealed high expression of CD73, CD90, and CD105 (≥ 95%), while CD34, CD45, HLA‐DR, CD19, and CD11b were expressed at ≤ 2% (Figure 1B), consistent with the characteristic surface markers of HUCMSCs.

Figure 1.

Identification of HUCMSCs. (A) Morphology of HUCMSCs after adhesion (×100). Scale bar = 200 μm. (B) Flow cytometry analysis of HUCMSCs surface markers (CD73, CD90, CD105, CD34, CD45, HLA‐DR, CD19, and CD11b). (C) Chondrogenic differentiation (Alcian blue staining, ×50). Scale bar = 200 μm. (D) Adipogenic differentiation (Oil Red O staining, ×400. Scale bar = 50 μm. (E) Osteogenic differentiation (Alizarin Red staining, ×100). Scale bar = 200 μm.

Tri‐lineage differentiation assays confirmed their multipotency: chondrogenic induction resulted in blue extracellular matrix by Alcian blue staining (Figure 1C); adipogenic induction showed intracellular red lipid droplets by Oil Red O staining (Figure 1D); osteogenic induction produced orange–red calcium nodules by Alizarin Red staining (Figure 1E). These results confirm that HUCMSCs possess multi‐lineage differentiation potential and are suitable for subsequent exosome isolation.

3.2. HUCMSC‐Exos Characterization

The physicochemical and biological characteristics of HUCMSC‐derived exosomes were comprehensively evaluated. Western blot analysis demonstrated strong expression of exosomal markers CD9, CD81, and TSG101, while the endoplasmic reticulum marker Calnexin was not detected (Figure 2A), indicating high purity of the isolated exosomes. TEM revealed typical cup‐shaped vesicles with clear boundaries, intact lipid bilayers, and uniform dispersion (Figure 2B). NTA showed that the particle size was mainly distributed between 30 nm and 150 nm, with an average diameter of approximately 130.4 nm (Figure 2C), confirming successful and stable exosome isolation. Cellular uptake of exosomes was assessed using PKH67 fluorescent labeling. After 6 h incubation, obvious green fluorescence was observed in the cytoplasm of HaCaT cells (Figure 2D). At 24 h, stronger fluorescence signals were detected in L929 cells (Figure 2D), indicating efficient uptake and intracellular accumulation of exosomes in skin‐related cells. These findings suggest that exosomes can be internalized by cells, supporting their potential role in intercellular communication and cargo delivery.

Figure 2.

Characterization of HUCMSC‐Exos. (A) Western blot analysis of exosomal marker proteins, including positive markers CD9, CD81, and TSG101, and the negative marker Calnexin. (B) Morphology of exosomes observed by TEM. Scale bar = 500 nm. (C) Size distribution of exosomes measured by NTA. (D) HaCaT cells exhibited exosome uptake at 6 h (×10). Scale bar = 50 μm. L929 cells exhibited exosome uptake at 24 h (×4). Scale bar = 100 μm.

3.3. Effects of APS and HUCMSC‐Exos on L929

CCK‐8 assay results showed that APS exhibited no cytotoxicity to L929 cells within the concentration range of 0–200 μg/mL, with cell viability exceeding 100%. Accordingly, 75 μg/mL APS was selected as the working concentration for subsequent cell experiments (Figure 3A). Treatment with HUCMSC‐Exos at concentrations ranging from 10 to 40 μg/mL improved L929 cell viability (Figure 3B). Combined treatment with APS and HUCMSC‐Exos further enhanced L929 cell viability. Notably, the APS–Exos group (10 μg/mL, 130.90% ± 2.98%) increased L929 cell viability compared with the APS group, and the difference between the two groups was statistically significant (p < 0.05) (Figure 3C). Increasing the Exos concentration to 20 μg/mL did not result in a further improvement in L929 cell viability, and the APS–Exos (20 μg/mL) group showed no significant difference compared with the APS–Exos (10 μg/mL) group (p > 0.05). Considering that 10 μg/mL Exos achieved a comparable biological effect to 20 μg/mL, it was selected as the working concentration for subsequent in vitro cell experiments.

Figure 3.

Screening of APS and exosome concentrations and evaluation of L929 cellular functions. (A) Cell viability of L929 cells treated with different concentrations of APS measured by CCK‐8 assay (n = 3). (B) Cell viability of L929 cells treated with different concentrations of HUCMSC‐Exos measured by CCK‐8 assay (n = 3). (C) Selection of HUCMSC‐Exos concentration for subsequent experiments (n = 3). (D) Representative images of L929 cells in scratch wound assay at 0 and 24 h. Scale bar = 200 μm. (E) Quantitative analysis of scratch migration rate (n = 3). (F) DCFH‐DA staining shows the expression of ROS in L929 cells under different treatment conditions. Scale bar = 200 μm. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, statistically significant differences between the two groups.

To evaluate cell migration capacity through scratch experiments. After 24 h, all treatment groups exhibited certain cell migration capacity, with the APS‐Exos group showing the most significant cell migration (Figure 3D). Quantitative analysis revealed that the cell migration rate of the APS‐Exos group was 74.15% ± 7.72%, higher than that of the APS group (47.66% ± 2.61%, p < 0.001), Exos group (39.71% ± 2.39%, p < 0.0001), and control group (15.82% ± 2.54%, p < 0.0001) (Figure 3E), indicating its combined enhancing effect in promoting fibroblast migration.

Regarding the regulation of oxidative stress, ROS fluorescence analysis demonstrated that Rosup treatment increased intracellular ROS levels, whereas treatment with APS, Exos, and APS‐Exos for 6 h reduced ROS accumulation (Figure 3F). Among these groups, APS‐Exos treatment resulted in lower intracellular ROS levels compared with individual treatments, indicating enhanced regulation of oxidative stress.

3.4. Effects of APS and HUCMSC‐Exos on HaCaT

Re‐epithelialization is a critical process in wound healing and may be impaired when HaCaT cells are exposed to a pro‐inflammatory microenvironment. CCK‐8 assay results showed that 75 μg/mL APS enhanced the viability of HaCaT cells (p > 0.05) without exhibiting cytotoxicity (Figure 4A). Co‐treatment with 10 μg/mL HUCMSC‐Exos (APS–Exos) further increased cell viability to 124.50% ± 3.18%, which was higher than that observed with APS or HUCMSC‐Exos alone (p < 0.01) (Figure 4B).

Figure 4.

Effects of APS and HUCMSC‐Exos on HaCaT cells. (A) Cell viability of HaCaT cells treated with different concentrations of APS measured by CCK‐8 assay (n = 3). (B) Effects of APS and HUCMSC‐Exos on HaCaT cell viability determined by CCK‐8 assay (n = 3). (C) Cell viability of HaCaT cells under H₂O₂‐induced oxidative stress after treatment with APS and HUCMSC‐Exos, measured by CCK‐8 assay (n = 3). (D) Live cell staining of HaCaT cells after H₂O₂‐induced oxidative injury using Calcein‐AM. Scale bar = 100 μm. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, statistically significant differences between the two groups.

In an H₂O₂‐induced oxidative stress model, cell viability in the H₂O₂ group decreased to 31.79% ± 3.79%, accompanied by morphological damage. Pretreatment with APS or HUCMSC‐Exos partially alleviated H₂O₂‐induced cytotoxicity, whereas APS‐Exos treatment further partially improved the viability of injured cells, reaching 72.11% ± 2.13% (p < 0.05, Figure 4C). Calcein‐AM staining showed improved cellular morphology in the treatment group compared with the H₂O₂‐treated group (Figure 4D), suggesting that the combined treatment partially alleviated oxidative stress‐induced cellular damage.

Collectively, APS and HUCMSC‐Exos improved the viability of fibroblasts and keratinocytes, enhanced fibroblast migration, reduced intracellular ROS levels and partially alleviated oxidative stress, thereby contributing to improved cellular responses. These findings provide an experimental basis for the development of multifunctional wound repair strategies and support their potential application in wound healing.

3.5. Structural Characteristics and Biocompatibility of Composite GelMA Hydrogels

The structural and physicochemical properties of GelMA and its composite hydrogels (APS@GelMA, Exos@GelMA, APS‐Exos@GelMA) were systematically evaluated. GelMA prepolymer rapidly crosslinked under 405 nm UV light to form stable hydrogels, demonstrating good operability and in situ gelling potential (Figure 5A). Adhesion tests on dynamic surfaces indicated stable attachment without noticeable detachment, reflecting excellent mechanical compliance (Figure 5B).

Figure 5.

Preparation and characterization of GelMA hydrogels. (A) Morphological changes of GelMA hydrogel before and after UV irradiation (405 nm). (B) Adhesion performance of GelMA hydrogel on skin tissue. (C) Microstructure of GelMA, APS@GelMA, Exos@GelMA, and APS‐Exos@GelMA hydrogels observed by SEM (500×). Scale bar = 50 μm. (D) Distribution of exosomes within the hydrogel. (E) Average pore size of hydrogels in each group (n = 3). (F) Equilibrium swelling ratio of hydrogels in PBS at 24 h (n = 3). (G) Degradation profiles of hydrogels in PBS at days 1, 4, 7, and 10 (n = 3). (H) Degradation profiles of hydrogels in Type II collagenase solution (n = 3). (I) Cumulative release curve of exosomes in APS‐Exos@GelMA hydrogel (n = 3).

SEM and confocal laser scanning microscopy revealed that all hydrogels exhibited uniform, interconnected 3D porous networks (Figure 5C). Exosomes were evenly distributed without aggregation (Figure 5D). Pore size analysis showed no significant differences among groups (GelMA: 47.30 ± 5.88 μm; APS@GelMA: 43.33 ± 4.27 μm; Exos@GelMA: 51.84 ± 1.57 μm; APS‐Exos@GelMA: 47.60 ± 6.71 μm; Figure 5E), indicating that incorporation of APS or exosomes did not disrupt the hydrogel's basic structure, which is conducive to cell adhesion, migration, and nutrient exchange.

Swelling tests demonstrated that all hydrogels had high water absorption capacity, reaching equilibrium within 24 h, and degradation tests exhibited environment‐dependent degradability, showing relatively slow degradation in PBS but accelerated degradation under collagenase‐mediated conditions (Figure 5F–H), suggesting that the hydrogel can maintain a moist wound environment while exhibiting controlled and enzyme‐responsive degradability. Overall, APS and HUCMSC‐Exos incorporation had minimal impact on the fundamental physicochemical properties of GelMA, preserving structural integrity and stability. Furthermore, the hydrogel exhibited a gradual exosome release profile, while the incorporation of APS and HUCMSC‐Exos established a multifunctional bioactive platform for local wound healing (Figure 5I).

Hemocompatibility was assessed via hemolysis assays. No visible red coloration was observed in the supernatant of any group (Figure 6A), and hemolysis rates were below 5% (Figure 6B), meeting biomaterial safety standards and indicating negligible damage to red blood cells.

Figure 6.

Biocompatibility of GelMA‐based composite hydrogels. (A) Hemolysis assay to evaluate the blood compatibility of hydrogels. (B) Quantitative analysis of hemolysis (%) in different hydrogel groups (n = 3). (C) Live/dead staining of L929 cells after co‐culture with GelMA‐based composite hydrogels. Scale bar = 100 μm.

Meanwhile, cytocompatibility was evaluated by co‐culturing L929 cells with hydrogels for 48 h followed by live/dead staining. No obvious dead cells (red fluorescence) were observed in any group (Figure 6C).

3.6. In Vivo Wound‐Healing Evaluation

A full‐thickness skin defect model was established in mice to evaluate the in vivo wound healing efficacy of the hydrogel. Compared with the PBS group, all hydrogel‐treated groups exhibited accelerated wound closure (Figure 7A). Among them, the APS‐Exos@GelMA group demonstrated the most pronounced therapeutic effect, characterized by rapid migration of wound edges toward the center and extensive coverage of the wound surface by newly regenerated tissue. Quantitative analysis revealed that the wound closure rate in the APS‐Exos@GelMA group reached 89.89% ± 3.17% on day 7, showing a statistically significant improvement compared with the PBS group (p < 0.0001) (Figure 7B).

Figure 7.

Analysis of wound healing in mice. (A) Representative macroscopic images of wound healing in mice. (B) Quantitative analysis of wound closure on day 7 (n = 6). (C) Quantitative analysis of wound closure on day 12 (n = 6). **p < 0.01, ****p < 0.0001, statistically significant differences between the two groups.

By day 12, wounds treated with APS‐Exos@GelMA were nearly completely healed, while the other groups also exhibited enhanced wound closure. The wound closure rates of the PBS, GelMA, and Exos@GelMA groups were 88.63% ± 2.96%, 91.17% ± 2.55%, and 97.22% ± 2.40%, respectively (Figure 7C). Notably, the GelMA group showed a slight improvement compared with the PBS group, which may be attributed to the ability of GelMA hydrogel to absorb wound exudate and maintain a moist wound environment [19]. These findings indicate that the APS‐Exos@GelMA composite hydrogel provides enhanced therapeutic efficacy by integrating the beneficial effects of APS, HUCMSC‐Exos, and the GelMA delivery matrix, thereby promoting skin wound healing.

While accelerated wound closure reflects the overall healing progress, complete skin regeneration involves not only surface restoration but also reconstruction of epidermal structure and extracellular matrix organization. To further determine whether the improved wound closure was accompanied by enhanced tissue regeneration, H&E, and Masson's trichrome staining were performed on day 12.

H&E staining revealed that the APS‐Exos@GelMA group exhibited a continuous and well‐organized epidermal layer, with tightly arranged keratinocytes and clear stratification, indicating enhanced re‐epithelialization (Figure 8A). In contrast, the PBS and GelMA groups showed less mature epidermal structures, with relatively poor epidermal–dermal organization and incomplete tissue remodeling. The Exos@GelMA group formed a continuous epidermis with improved thickness and cellular arrangement compared with the GelMA group, although, compared with the APS‐Exos@GelMA group, the epidermis remained slightly thickened and residual erythrocytes were present in the underlying tissue.

Figure 8.

Histological analysis of wound tissues. (A) H&E and Masson's trichrome staining of wound tissues (Original magnification: ×10). Scale bar = 100 μm. (B) H&E staining of major organs (heart, liver, spleen, lung, and kidney) in mice (×20). Scale bar = 200 μm.

Masson's trichrome staining demonstrated enhanced collagen deposition in the APS‐Exos@GelMA group, with more abundant collagen distribution in the tissue (Figure 8A). In contrast, the PBS and GelMA groups exhibited relatively limited collagen accumulation, indicating insufficient extracellular matrix remodeling. The Exos@GelMA group showed increased collagen deposition compared with the GelMA group, accompanied by improved collagen distribution; however, the collagen matrix remained less organized. These findings suggest that APS‐Exos@GelMA promoted extracellular matrix formation and facilitated tissue remodeling during wound healing.

In vivo biosafety was evaluated by H&E staining of major organs (heart, liver, spleen, lung, and kidney). Histological analysis revealed intact tissue structures and normal cellular morphology in all groups, with no obvious inflammatory infiltration, necrosis, or pathological abnormalities observed (Figure 8B). These findings indicate that the composite hydrogel exhibits good biocompatibility and safety in vivo.

4. Discussion

Compared with normal wound healing, chronic hard‐to‐heal wounds are frequently characterized by imbalanced oxidative stress and impaired cellular functions, which subsequently lead to disrupted tissue repair processes and extended healing periods [20]. At present, therapeutic strategies for chronic wounds have shifted from single‐modality approaches to multi‐component and integrated treatments [21]. Based on existing studies, combination therapies are expected to regulate the wound microenvironment at multiple levels, including alleviating oxidative stress, restoring cellular function, and ultimately promoting the healing of refractory wounds. In this context, we designed a composite biomaterial composed of multiple bioactive components, in which GelMA hydrogel was used as a local delivery platform to incorporate HUCMSC‐Exos and APS for wound repair.

GelMA is synthesized via methacryloyl modification of gelatin, retaining the excellent cellular biocompatibility of gelatin while enabling tunable hydrogel properties [22]. Its three‐dimensional porous network structure not only supports cell migration and tissue regeneration but also allows controlled release of bioactive molecules through physical or chemical encapsulation, thereby prolonging their retention at the wound site and enhancing therapeutic efficacy. In this study, the tissue adhesiveness, porous structure, and tunable properties of GelMA were utilized to establish a local delivery platform for HUCMSC‐Exos and APS. The hydrogel facilitated local retention and gradual release of HUCMSC‐Exos, while providing a suitable matrix for APS incorporation, thereby enhancing the availability of therapeutic components at the wound site. Meanwhile, both HUCMSC‐Exos and APS were physically incorporated into GelMA. Characterization of the physicochemical properties revealed no significant changes after loading, indicating good compatibility between GelMA and the bioactive components, which preserved the intrinsic properties of the hydrogel while endowing it with bioactivity.

Building upon this, the degradation behavior of the composite hydrogel was further evaluated. Ideally, the degradation rate of hydrogels should be synchronized with the process of tissue regeneration. During the early stage of wound healing, the hydrogel is expected to provide a stable structural framework to support cell adhesion and protect the wound site, followed by gradual degradation to facilitate cell infiltration, extracellular matrix deposition, and tissue remodeling [23]. In this study, the GelMA‐based hydrogel exhibited environment‐dependent degradation behavior, characterized by relatively slow degradation in PBS and accelerated degradation in the presence of collagenase type II. These results indicate that the hydrogel undergoes both hydrolytic and enzyme‐responsive degradation, consistent with previously reported findings [24]. Although the in vivo degradation kinetics were not directly characterized, the in vivo findings demonstrated that APS–Exos@GelMA promoted wound closure, epidermal regeneration, and extracellular matrix deposition without observable adverse effects. These outcomes indirectly suggest that the degradation behavior of the hydrogel is, to some extent, compatible with the process of tissue regeneration and does not hinder tissue repair. However, it should be noted that the current in vitro degradation model, based on a single enzyme system, cannot fully replicate the complexity of the in vivo wound microenvironment, which involves the coordinated action of multiple enzymes, dynamically evolving biochemical cues, and various physical factors [25]. Therefore, further studies are warranted to systematically investigate the in vivo degradation behavior of the hydrogel and its relationship with tissue regeneration, which will facilitate the rational optimization of the hydrogel system.

APS, as a natural bioactive polysaccharide, exhibit potent anti‐inflammatory, antioxidant, and immunomodulatory properties and have been widely applied in adjunctive therapy for tumors [26], diabetes [27], and its complications [28], as well as liver and kidney injuries [29]. Wound healing is often associated with excessive oxidative stress and impaired cellular function, which subsequently disrupt tissue repair processes. Therefore, the antioxidative and potential immunomodulatory properties of APS may contribute to wound repair. In this study, both in vitro and in vivo experiments further demonstrated its effects on skin‐related cells and wound healing. Within a certain concentration range, APS enhanced the viability and migration of L929 fibroblasts and HaCaT keratinocytes, partially reduced oxidative stress‐induced cellular damage, and maintained normal cellular physiological functions, which is consistent with previous reports.

Exosomes, as one of the most widely studied cell‐free therapeutic strategies, contain various proteins, RNAs, and signaling molecules and exhibit high biological activity and functional specificity. They can be efficiently internalized by recipient cells to mediate intercellular communication and have been extensively applied in various pathological studies, including wound healing, liver and kidney injury, and bone regeneration. In the context of wound repair, exosomes have been reported to enhance cell proliferation and migration by regulating cell cycle–related proteins and multiple signaling pathways, including the PI3K/Akt and MAPK pathways [30]. They have also been widely reported to exert antioxidant effects by downregulating the expression of pro‐oxidant genes, thereby reducing excessive ROS accumulation [31]. In this study, exosomes improved the viability and migration of L929 fibroblasts and partially improved HaCaT keratinocyte viability under oxidative stress conditions, which is consistent with previous reports, further supporting their potential role in regulating cellular behaviors associated with tissue regeneration.

However, studies directly combining APS and exosomes for wound healing remain limited. The combined application of these two components to investigate their complementary effects on wound repair may further improve our understanding of therapeutic strategies for chronic hard‐to‐heal wounds. At present, most studies have focused on the combination of APS with stem cells or APS preconditioning of stem cells, which has been shown to promote stem cell proliferation, enhance immunomodulatory capacity both in vitro and in vivo, and improve homing ability without altering cellular morphology or surface marker expression, thereby supporting subsequent pathological investigations [32]. In contrast, leveraging the bioactivities of both APS and exosomes to regulate the wound microenvironment and improve cellular functions may represent a promising approach for wound healing and provide further insights into cell‐free therapeutic strategies.

In this study, the combined use of APS and exosomes showed improved therapeutic outcomes compared with either treatment alone. This may be attributed to their complementary biological activities. First, within the complex wound microenvironment, both APS and exosomes can contribute to microenvironmental improvement, thereby creating favorable conditions for sustained cellular regulation and tissue repair. Second, APS and exosomes regulate cell proliferation and migration through partially distinct signaling pathways. For instance, APS has been reported to promote angiogenesis by activating the PI3K/Akt pathway and upregulating the expression of CD31, CD34, and VEGFR, whereas exosomes promote angiogenesis not only through activation of multiple signaling pathways but also by delivering functional proteins, RNAs, and other bioactive molecules to recipient cells [6, 33]. These indirect and direct mechanisms provide complementary effects, thereby promoting angiogenesis at multiple levels. Furthermore, during tissue remodeling in wound healing, APS has been reported to inhibit fibrosis by suppressing the TLR4/NF‐κB signaling pathway and reducing inflammatory cytokine activation, while exosomes may regulate gene expression to modulate cellular behavior and suppress excessive fibrosis [34, 35]. Together, their distinct mechanisms enable a dynamic balance between promoting wound repair and preventing excessive fibrotic remodeling in wound healing.

In summary, this study presents a multifunctional GelMA‐based therapeutic platform incorporating APS and exosomes for enhanced wound repair. GelMA provides a favorable delivery matrix, while APS and exosomes contribute biological activities, including attenuation of oxidative stress, regulation of cellular behaviors, and tissue regeneration. By integrating local delivery with multiple bioactive components, this strategy provides a potential approach to addressing the limitations of conventional single‐component therapies and offers new insights into the development of multifunctional biomaterials for tissue engineering applications.

Despite these promising findings, several limitations should be acknowledged. First, this study demonstrated the therapeutic potential of APS‐Exos@GelMA hydrogels for wound repair, the controlled delivery mechanism of APS within the hydrogel matrix remains to be further elucidated due to the lack of systematic evaluation of APS release behavior. Future studies should investigate the release kinetics of APS from GelMA hydrogels and optimize hydrogel properties, including crosslinking density, network structure, and physicochemical characteristics, to achieve more precise spatiotemporal control of APS delivery and further enhance therapeutic efficacy. Second, the complexity of APS should be considered, as variations in extraction and processing methods may lead to differences in pharmacological activity, and its molecular weight can also influence biological efficacy [28]. Therefore, the establishment of standardized sources and extraction protocols is essential to facilitate further clinical translation of APS and to ensure the stability and reproducibility of therapeutic outcomes in future studies. In addition, the administration route of APS may also affect therapeutic efficacy, as different delivery methods such as transdermal administration and intravenous injection may yield distinct biological effects. Third, the processing and isolation methods of exosomes may also influence their biological activity [36]. Although the present study demonstrates that the combined application of APS and exosomes yields enhanced therapeutic effects, dosage optimization based on their respective bioactivities is still required. In addition, previous studies have indicated that both APS‐ and MSC‐derived exosomes possess immunomodulatory properties; however, whether their combination produces enhanced or complementary effects on inflammatory regulation remains unclear. The specific mechanisms underlying the interactions between APS and exosomes, particularly their roles in regulating inflammatory responses, cellular behavior, and oxidative stress responses, require further investigation. Future studies should incorporate comprehensive analyses of inflammatory cytokines, macrophage polarization profiles, and multi‐omics approaches, together with clinically relevant impaired‐healing models, to elucidate the mechanistic basis of the APS–Exos@GelMA system. Such efforts will provide stronger theoretical support for optimizing this composite platform and advancing its potential clinical translation.

Author Contributions

Jingwen Tian: conceptualization, data curation, formal analysis, investigation, methodology, writing – original draft, writing – review and editing. Xiangjing Zhao: conceptualization, data curation, formal analysis, investigation, methodology, writing – original draft, writing – review and editing. Jingxian Xie: data curation, formal analysis, methodology, software. Zheng Huang: data curation, formal analysis, methodology. Pengbo Song: project administration, supervision, visualization, writing – review and editing. Suzhuang Hong: supervision, visualization, writing – review and editing. Cailing Tong: project administration, supervision, visualization, writing – review and editing. Zhongquan Qi: funding acquisition, project administration, resources, supervision, writing – review and editing.

Ethics Statement

All animal experiments in this study were approved by the Animal Ethics Committee of Mogengel Biotechnology Co. Ltd. (Xiamen, China, IACUC: XMMGLAC202510002).

Conflicts of Interest

The authors declare no conflicts of interest.

Tian J., Zhao X., Xie J., et al., “GelMA‐Based Hydrogel Encapsulating Astragalus Polysaccharides and Umbilical Cord Mesenchymal Stem Cell Exosomes Promotes Full‐Thickness Skin Wound Healing,” Chronic Diseases and Translational Medicine 0 (2026): 1‐15, 10.1002/cdt3.70072.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.