Synergistic Immunomodulation and Angiogenesis‐Driven Microenvironment Remodeling via Bioactive Hydrogel/Nanofiber Composites for Enhanced Infectious Wound Healing

ABSTRACT

Numerous types of wounds are at risk for infection, such as burn wounds, traumatic wounds, surgical wounds, chronic ulcer wounds, and radiologic wounds. The repair of infectious wounds depends on specific microenvironments, including moderate hydration, reactive oxygen scavenging, antimicrobial, anti‐inflammatory, and angiogenic conditions. To effectively promote the repair of infectious wounds and reduce the risk of infection spread, we developed a multifunctional bilayer wound dressing (MBWD) that simultaneously meets these criteria. The inner layer (DMOG@PCL/ASC) consists of a polycaprolactone (PCL)/fish collagen (ASC) nanofiber decorated with coaxial microparticles containing dimethyloxalylgcine (DMOG). The three‐dimensional pore structure of nanofibers offers numerous adhesion sites for cells, and the gradual release of DMOG can promote angiogenesis. The outer layer consists of a glycidyl methacrylate‐modified carboxymethyl chitosan hydrogel loaded with cerium oxide nanozymes (M‐CMCS/CeO₂). This M‐CMCS hydrogel provides a moist microenvironment and exhibits antimicrobial properties. The localized release of CeO₂ can deliver antioxidant effects, combat microbes, and modulate inflammation. In vivo studies confirmed that the MBWD dressing can enhance the healing of infectious and diabetic wounds by providing rapid antimicrobial, anti‐inflammatory, and pro‐angiogenic effects. Therefore, this combined system of nanofibers and hydrogels, each loaded with specific functional components, offers an effective strategy for treating infectious wounds.

A multifunctional bilayer wound dressing (MBWD) was constructed via electrospinning/electrospraying and photocrosslinking technologies for synergistic treatment of infectious and diabetic wounds. The inner layer (DMOG@PCL/ASC) consists of polycaprolactone (PCL)/fish collagen (ASC) nanofibers decorated with DMOG‐loaded coaxial microparticles, providing a 3D porous structure for cell adhesion and sustained DMOG release to stabilize HIF‐1α and promote angiogenesis. The outer layer (M‐CMCS/CeO₂) is a glycidyl methacrylate‐modified carboxymethyl chitosan hydrogel loaded with cerium oxide (CeO₂) nanozymes, exerting antibacterial, antioxidant, and anti‐inflammatory effects by scavenging reactive oxygen species (ROS) and regulating macrophage polarization.

1. Introduction

When skin integrity is compromised, the ability to resist pathogen invasion is significantly reduced, leading to wound infection and systemic infection [1, 2, 3, 4]. The presence of pathogens promotes excessive release of inflammatory factors and further activates matrix metalloproteinases, prolonging the inflammatory response cycle and hindering a smooth transition from the inflammatory to the proliferative phase [5, 6, 7]. During the inflammatory response, large numbers of pro‐inflammatory macrophages accumulate, leading to a significant increase in reactive oxygen species (ROS). This process exacerbates local inflammation and inhibits normal angiogenesis. Wound healing is a complex biological process that requires adequate nutritional support, along with the orderly proliferation and migration of fibroblasts and epithelial cells, as well as the synthesis and remodeling of the extracellular matrix [8, 9, 10]. Current treatment methods used in clinical practice often struggle to achieve rapid, efficient, and high‐quality wound healing, typically lacking effective regulation of the microenvironment. There is an urgent need to develop novel multifunctional dressing systems capable of precisely regulating the wound microenvironment, timely modulating the inflammatory response, promoting angiogenesis, and accelerating cell proliferation and matrix remodeling.

Electrospun nanofibers and hydrogels are two core dressing materials in the treatment of infectious and chronic wounds, each boasting unique functional advantages [11, 12, 13, 14]. Electrospun nanofibers enable efficient drug loading and controlled release, greatly expanding the interaction interface between the material and wound tissue to effectively facilitate cell adhesion, proliferation, and migration. These nanofibers feature high porosity and large specific surface area, which easily lead to rapid water evaporation. Excessive air permeability can leave the wound overly dry, failing to maintain the optimal humidity required for wound healing. Modern wound care concepts emphasize maintaining an appropriate humidity balance in the wound, and the limitations of electrospun dressings in this regard make them unable to support moist wound healing. Hydrogels possess excellent hydrophilic properties, enabling them to efficiently absorb substantial amounts of wound exudate. They help maintain the optimal humidity balance of the wound, effectively preventing scab formation and tissue adhesion that can occur due to wound dehydration. The inherent flexibility and elastic deformation capacity of hydrogels enable them to conform well to irregularly shaped wounds and adapt to human physiological activities. Some hydrogels can effectively inhibit the colonization and proliferation of pathogenic bacteria in wounds by loading antibacterial active ingredients or by relying on their inherent antibacterial properties. Hydrogels can also serve as intelligent carrier systems for bioactive molecules, regulating the biological behavior of local cells to synergistically promote tissue regeneration and functional reconstruction.

Compared with single‐structure materials, the composite of electrospun fibers and hydrogels can construct an ideal bifunctional wound dressing system and fully leverage the complementary advantages of the two materials. With its ultrafine fibrous structure and high porosity, the electrospun layer can quickly and efficiently absorb wound exudate and fluid accumulation, thereby preventing bacterial growth and tissue maceration caused by excessive fluid buildup. Researchers developed a multifunctional nerve conduit that is responsive to ultrasound [15]. The inner layer is composed of poly(vinylidene fluoride‐trifluoroethylene) (P(VDF‐TrFE)) electrospun nanofibers doped with barium titanate piezoelectric nanoparticles (BTNPs). Both the piezoelectric properties and the oriented arrangement of these fibers have been optimized. The outer layer is a thermosensitive poly(N‐isopropylacrylamide) (pNIPAM) composite hydrogel capable of encapsulating bioactive drugs. Experimental results showed that this nerve conduit can significantly accelerate functional recovery and axonal regeneration in a rat model of long sciatic nerve defects. Chen et al. developed an injectable electrospun fiber‐hydrogel composite drug delivery system, aiming to achieve long‐term and selective analgesia targeting injurious sensations [16]. Poly (ε‐caprolactone) (PCL) electrospinning nanofibers were used to load QX‐314, while F127 hydrogel was utilized to deliver capsaicin. This composite material was shown to significantly prolong the duration of sciatic nerve block in rats by sequentially releasing capsaicin and QX‐314. Wu et al. developed a novel electrospun composite dressing made of PCL fibers and gelatin methacrylamide (GelMA) hydrogel, featuring a reticular surface structure and enabling controlled release of deferoxamine [17]. In a diabetic rat model treated with this dressing, wound healing occurred significantly faster, and the incidence of wound infection was reduced. Therefore, composite biomaterials incorporating electrospun nanofibers and hydrogels demonstrate great potential for applications in nerve repair, controlled drug delivery, and wound healing.

Dimethyloxalylglycine (DMOG), a prolyl hydroxylase inhibitor, stabilizes hypoxia‐inducible factor‐1α (HIF‐1α) and enhances the expression of angiogenesis‐related genes, thereby inducing neovascularization [18, 19]. Applying DMOG directly to the wound may lead to rapid drug degradation and excessively high local concentrations, which can trigger inflammatory responses or cytotoxicity. Electrospinning technology can facilitate the sustained release of DMOG, creating a microenvironment that promotes neovascularization at the wound site. Research has demonstrated that electrospun nanofibers made from PCL and type I collagen, when loaded with DMOG, can significantly enhance angiogenesis and support epithelial regeneration during the wound healing process [20]. Essentially, cerium oxide (CeO₂) nanozymes are nanomaterials with enzymatic catalytic activity, primarily in the form of nanoscale CeO₂ nanoparticles. Multiple studies have incorporated CeO₂ nanozymes into hydrogels, using them as carriers to achieve sustained release. This approach not only enhances their utilization efficiency but also imparts hydrogels with antioxidant, anti‐inflammatory, and antibacterial properties [21, 22, 23, 24, 25]. The combination of electrospun nanofibers and hydrogel platforms is expected to enable sustained local release of various therapeutic agents, thereby accelerating the healing of infected wounds.

This study has successfully developed an multifunctional bilayer wound dressing (MBWD) that employs a layered, functionalized design to efficiently meet the therapeutic needs of infectious and diabetic wounds. The dressing features a bilayer composite structure that fully leverages the synergistic effects of its structural components. The inner layer (DMOG@PCL/ASC) is fabricated via a composite process combining electrospinning and electrospraying. Its lies in encapsulating DMOG within the core layer of coaxial microspheres, a structure that enables controlled, sustained release, ensuring continuous, stable promotion of angiogenesis throughout the entire wound‐healing process. The outer layer (M‐CMCS/CeO₂) is constructed using a functional hydrogel. CeO₂ with enzyme‐mimetic activity is uniformly loaded into a glycidyl methacrylate‐modified carboxymethyl chitosan hydrogel matrix (M‐CMCS). This design endows the outer layer with excellent antibacterial, antioxidant, and anti‐inflammatory properties, providing comprehensive protection for the wound. The bilayer three‐dimensional porous network structure of the MBWD not only exhibits superior hemostatic effects but also precisely regulates the wound microenvironment through the synergistic release of DMOG and CeO₂. Validation in SD rat models of infected full‐thickness skin defects and diabetic skin defects has shown that the dressing significantly accelerates wound healing, demonstrating favorable therapeutic outcomes. By virtue of the organic combination of its unique bilayer structural design and multifunctional active components, the MBWD developed in this study offers a bioactive material solution with broad application prospects for treating infectious wounds.

2. Experimental Section

2.1. Materials and Reagents

Carboxymethyl chitosan (CMCS), glycidyl methacrylate (GMA), chloroform, and hexafluoroisopropanol (HFIP) were purchased from Macklin (Shanghai, China). PCL and ASC were sourced from Sigma‐Aldrich (USA). Lithium phenyl‐2,4,6,‐trimethylbenzoylphosphinate (LAP) was obtained from Engineering For Life (Suzhou, China). CeO₂ nanozymes were acquired from Nanjing Jike Biotechnology Co., Ltd. (Nanjing, China). Dialysis tubing MD44, hydrogen peroxide (H₂O₂) assay kits, live/dead staining kits, 4,6‐diamidino‐2‐phenylindole (DAPI), phosphate‐buffered saline (PBS), 4% paraformaldehyde, and fluorescein isothiocyanate‐labeled bovine serum albumin (FITC‐BSA) were purchased from Solarbio (Beijing, China). Penicillin/streptomycin, 0.25% trypsin, and 0.1% crystal violet were sourced from Biosharp (Beijing, China). Matrigel was obtained from Corning (USA). DMOG and Cell counting kit‐8 (CCK‐8) assay kits were purchased from Glpbio (USA). Fetal bovine serum (FBS) was sourced from Gibco (USA). Macrophage colony‐stimulating factor (M‐CSF) was obtained from PeproTech (USA). Antibodies for CD68, CD86, CD206, and VEGF were purchased from Hua'an Biotechnology (Hangzhou, China). HIF‐1α, iNOS (Inducible Nitric Oxide Synthase), and Arg‐1 (Arginase‐1) were sourced from Abcam (USA). Enzyme‐linked immunosorbent assay (ELISA) kits for interleukin‐4 (IL‐4), interleukin‐6 (IL‐6), tumor necrosis factor‐α (TNF‐α), and interleukin‐10 (IL‐10) were purchased from Hua'an Biotechnology (Hangzhou, China). Mouse fibroblasts (L929) and human umbilical vein endothelial cells (HUVECs) were provided by the Plastic Surgery Department of Xinqiao Hospital, Army Medical University. Primary bone marrow macrophages (BMDMs) were isolated and characterized from the femurs of SD rats.

2.2. Screening for DMOG Concentration

To investigate the impact of various concentrations of DMOG on HIF‐1α protein expression, this study designed relevant experiments. HUVECs were uniformly seeded into 24‐well plates at a density of 5 × 10³ cells per well. A total of five groups were established, including a control group and four treatment groups with different concentrations of DMOG. Specifically, no DMOG was added to the control group, whereas the treatment groups received 250, 500, 750, and 1,000 µM DMOG, respectively. Each group was given 400 µL of culture medium containing the corresponding DMOG concentration to culture HUVECs, which were then incubated in a CO₂ incubator. At the same time points on Days 1, 3, and 5, a 10% CCK‐8 solution was added to each well and incubated for 2 h in a CO₂ incubator. The absorbance was measured at 450 nm using a microplate reader (Thermo, USA) to assess the effect of varying DMOG concentrations on HUVEC proliferation.

The Western blot was used to clarify how different DMOG concentrations affect HIF‐1α protein expression. HUVECs exposed to DMOG at 250, 500, 750, or 1000 µM were collected and lysed to extract total protein. Equivalent amounts of protein extracts were separated by SDS‐PAGE (sodium dodecyl sulfate‐polyacrylamide gel electrophoresis) and transferred onto PVDF (polyvinylidene difluoride) membranes. PVDF membranes were blocked with 5% nonfat milk for 2 h to reduce nonspecific binding. Membranes were incubated overnight at 4°C with a beta‐actin primary antibody (1:2000) as an internal control and a specific HIF‐1α antibody (1:1000) to ensure adequate binding. Membranes were rinsed with TBST (Tris‐Buffered Saline and Tween‐20) to remove unbound primary antibodies and then incubated with the corresponding secondary antibody (1:5000) at room temperature for 1 h. Immunoblotted proteins were visualized using an enhanced chemiluminescence (ECL) detection system, and the resulting bands were quantitatively analyzed with ImageJ software.

2.3. Construction and Characterization of DMOG@PCL/ASC, M‐CMCS/CeO₂, SLM and the MBWD Scaffolds

PCL was dissolved in HFIP at a concentration of 10% w/v and uniformly mixed with 1 mg/mL of Rhodamine B as the shell layer solution marker. Simultaneously, ASC was dissolved in HFIP at 7% w/v as the core layer solution. Subsequently, these two solutions were separately injected into a coaxial injector and fixed onto a syringe pump. The needle‐to‐roller distance was set at 10 cm, a flow rate of 1 and 0.5 mL/h for both the outer and inner solutions. The applied voltages were set to 15 and 3 kV at the positive and negative poles, respectively, to generate a stable electric field. A roller was used as the receiving device, with a rotational speed of 500 rpm/min. The obtained PCL/ASC coaxial nanofibers were placed in a vacuum oven for 24 h to thoroughly remove non‐volatile organic solvents. Next, the PCL solution concentration was adjusted to 2% w/v, the solvent was changed to chloroform, and 1 mg/mL of FITC‐BSA was added as a shell marker. DMOG was dissolved in chloroform as the core solution. The pump speeds of the shell and core solutions were 2 mL/h and 0.2 mL/h, respectively. The previously prepared PCL/ASC nanofibers served as the receiving substrate for collecting the coaxial microparticles containing DMOG. Through the methods, we successfully fabricated the inner layer (DMOG@PCL/ASC) of the dressing. The morphology and structure of the nanofibers and microparticles were observed using an S‐4800 scanning electron microscope (SEM, Hitachi, Japan). The diameters were measured using ImageJ (n = 100). The wettability was determined using a contact angle tester (DSA10, Kruss, Hamburg, Germany).

CMCS was dispersed in deionized water, and GMA was added dropwise over 48 h while stirring continuously. A total of 1 mol/L hydrochloric acid aqueous solution was added, and the mixture was dialyzed for 3 days, filtered through a filter membrane, frozen, and lyophilized to obtain solid M‐CMCS. Solid M‐CMCS was dissolved in LAP initiator to prepare a 5% w/v hydrogel precursor solution. CeO₂ nanoparticles were evenly dispersed into it to form M‐CMCS‐CeO₂ hydrogel mixtures with concentrations of 25, 50, 100, 200, and 300 mg/mL, respectively, which were cured by photocrosslinking. The obtained hydrogel scaffolds were named M‐CMCS‐CeO₂‐25, M‐CMCS‐CeO₂‐50, M‐CMCS‐CeO₂‐100, M‐CMCS‐CeO₂‐200, and M‐CMCS‐CeO₂‐300, respectively. The DMOG@PCL/ASC scaffold was treated with plasma for 5 min and assembled layer‐by‐layer with the M‐CMCS‐CeO₂ scaffold to construct the MBWD composite scaffolds. Specifically, the inner DMOG@PCL/ASC nanofiber layer was subjected to plasma treatment to introduce carboxyl (–COOH) functional groups on its surface, while the outer M‐CMCS hydrogel contains glycidyl methacrylate (GMA)‐derived epoxy groups. During photo‐crosslinking, ring‐opening reactions occur between the carboxyl and epoxy groups, forming covalent bonds between the two layers.

The morphologies and structures of scaffolds and CeO₂ were observed by S‐4800 scanning electron microscope (SEM, Hitachi, Japan) and JEM‐2100 high‐resolution transmission electron microscope (TEM, Jeol, Japan). Wettability was determined using a contact angle tester. Pore diameters were measured using ImageJ. The chemical compositions of the scaffolds were analyzed using Fourier transform infrared spectroscopy (FTIR). A 0.02 g hydrogel sample was weighed (W₁ ) and immersed in PBS. After 24 h, the sample was removed, and excess water on the hydrogel surface was blotted with absorbent paper and weighed again (W₂ ). The swelling ratio was calculated as: Swelling Ratio = (W₂ −W₁ ) / W₁ × 100%. The initial mass of the scaffolds was recorded as M₀ . Then, the scaffolds were placed in PBS. The scaffolds were retrieved at different time points and freeze‐dried, and their weight (M_t ) was recorded. The degradation rate of the scaffolds was calculated as: degradation rate = (M₀ −M_t ) / M₀ × 100%. Using FITC‐BSA as a model load, the MBWD scaffold was placed in a 15 mL centrifuge tube, and 10 mL of PBS was added to each tube. The tubes were incubated in a horizontal shaker at 37°C and 120 rpm/min. At 4 h, 8 h, 12 h, 1 d, 3 d, 5 d, 7 d, 14 d, and 21 d of incubation, 1 mL of PBS was aspirated from each tube. The absorbance of the PBS was measured, and the cumulative release rate was calculated according to the instructions of the Bicinchoninic Acid kit.

To prepare a control group with identical chemical compositions but a simplified structure, all functional components of the MBWD were integrated into a monolithic hydrogel matrix. Briefly, M‐CMCS was dissolved in deionized water containing 0.25% (w/v) LAP initiator to form a 5% (w/v) precursor solution. Subsequently, CeO₂ nanoparticles (200 mg/mL) and free DMOG (at an equivalent dose to the total loading in MBWD, 250 µM) were uniformly dispersed into the solution. To maintain consistency in polymer composition, an equivalent proportion of pre‐pulverized PCL/ASC polymer components, prepared via high‐speed shearing, was blended into the mixture. The resulting slurry was cast into a mold and photo‐crosslinked under 405 nm UV light for 60 s to form the monolithic Single‐Layer Mixed (SLM) scaffold. Unlike MBWD, the DMOG in the SLM group was incorporated directly in its free form, without coaxial microparticle encapsulation.

2.4. Biocompatibility and H₂O₂ Scavenging Activity of M‐CMCS Hydrogel

To assess the biocompatibility of M‐CMCS, M‐CMCS‐CeO₂‐25, M‐CMCS‐CeO₂‐50, M‐CMCS‐CeO₂‐100, M‐CMCS‐CeO₂‐200, and M‐CMCS‐CeO₂‐300, the materials were immersed in complete culture medium at a ratio of 0.1 g/mL for 48 h. Cell compatibility was then determined using the CCK‐8 assay after co‐culturing the scaffold extracts with L929 cells. The RO scavenging capability was evaluated using an H₂O₂ detection kit. After a 10‐minute reaction time, the absorbance at 415 nm was measured using a microplate reader. To investigate intracellular ROS levels in RAW264.7 and L929 cells, the DCFH‐DA assay was used. RAW264.7 and L929 cells were seeded at a density of 1 × 10⁴ cells/mL in 24‐well plates and cultured for 24 h. Cells were then stimulated with 500 ng/mL LPS for 24 h, with hydrogel extracts added to a subset of samples. Following stimulation, cells were stained with DCFH‐DA (10 µM) for 30 min in the dark. Intracellular ROS levels were quantified by measuring fluorescence intensity under an inverted fluorescence microscope.

2.5. Live/Dead Staining, Cell Migration, and Antibacterial Properties

To evaluate the biocompatibility of the MBWD scaffold, DMOG@PCL/ASC (with a DMOG dose of 250 µM) and M‐CMCS‐CeO₂‐200 (abbreviation in M‐CMCS/CeO₂) scaffolds were used as experimental controls. The various scaffolds were immersed in complete culture at a ratio of 0.1 g/mL for 48 h, respectively, and the scaffold extracts were collected. L929 cells and HUVECs were inoculated in 24‐well culture plates at a density of 5 × 10³ cells per well, respectively. They were then co‐cultured with scaffold extracts for 24 h. Subsequently, the cells were stained according to the instructions for the live/dead staining kit and incubated at 37°C for 30 min. Cell viability was observed using an inverted microscope (Olympus, Japan).

In vitro L929 cell migration assays were carried out to assess the effect of different scaffold samples on wound healing. L929 cells were seeded in 24‐well culture plates at a density of 5 × 10⁴ cells/well. Cells were allowed to reach about 90% confluency before a vertical scratch was made at each well's center with a sterile 200 µL pipette tip. Cells were washed with sterile PBS to remove cellular debris, then co‐incubated with various scaffold extracts. At 0 h (W ₀) and 24 h (W _t), cells were stained with calcein using a live/dead cell staining kit and observed under an inverted microscope. Cell migration distances were measured using ImageJ software, and the wound closure rate was calculated with the formula: Migration Rate = (W ₀ −W _t) / W ₀ × 100%.

To assess the antibacterial activities of DMOG@PCL/ASC, M‐CMCS‐CeO₂, and MBWD scaffolds, they were placed in 24‐well plates under the specified conditions. Escherichia coli (E. coli) and Staphylococcus aureus (S. aureus) in log phase were seeded on sample surfaces at 1 × 10⁵ CFU/mL (100 µL) and incubated at 37°C for 4 h. After incubation, bacteria were rinsed with sterile PBS, and 100 µL of the rinse solution was spread onto agar plates. These plates were further incubated at 37°C for 24 h, after which the colonies were imaged and quantified using ImageJ. Each group of samples was soaked in 10 mL of bacterial suspension and incubated with shaking for 24 h. Bacteria were stained with a live/dead staining kit and examined under an inverted fluorescence microscope to observe growth.

A clear indication of antibacterial activity is the scaffolds’ ability to break down bacterial biofilms. A total of 1 mL of overnight broth‐cultured S. aureus (1 × 10⁶ CFU/mL) was added to the wells of a 48‐well microtiter plate and incubated at 37°C for 48 h to induce static biofilm formation. Various scaffolds were added and incubated at 37°C for 24 h before bacterial biofilms were fixed with 4% paraformaldehyde and stained with 0.3% w/v crystal violet for 20 min. The biofilms were washed with PBS to remove unbound dye and then photographed digitally. They were subsequently dissolved in anhydrous ethanol for absorbance measurement at 570 nm using a microplate reader.

2.6. The Regulatory Role of HUVECs In Vitro

The effects of DMOG@PCL/ASC, M‐CMCS‐CeO₂ and MBWD scaffolds on cell migration were further analyzed by HUVECs scratch assay and transwell migration assay to elucidate the wound healing. The HUVECs scratch assay was performed as described in Section 2.5. The steps of the transwell migration assay were as follows: scaffold extracts were added to the lower chamber, and the upper chamber was inoculated with HUVECs at 5 × 10⁴ cells/ well. After 24 h of co‐culture, the upper chamber was removed. The cells in the lower chamber were fixed with 4% paraformaldehyde for 20 min, stained with 0.1% crystal violet for 30 min, and imaged under an inverted microscope.

The angiogenic capacity of the scaffolds was evaluated via an angiogenesis assay with HUVECs. A total of 150 µL Matrigel was dispensed into each well of a 24‐well culture plate and incubated at 37°C for 30 min to achieve solidification. A density of 1 × 10⁵ cells/well was seeded onto the solidified Matrigel surface and incubated together with scaffold extracts until HUVECs underwent vascularization. Angiogenesis‐related images were acquired using an Olympus inverted microscope, and vascularization parameters were quantified using ImageJ.

To verify whether various scaffolds could upregulate VEGF expression, we used immunofluorescence staining and Western blot experiments. HUVECs were inoculated in 24‐well plates at a density of 1 × 10⁴ cells/well and co‐cultured with various scaffold extracts for 48 h. Cells were fixed with 4% paraformaldehyde for 15 min and rinsed 3 times with PBS. The cells were then permeabilized with 0.3% Triton X‐100 for 10 min at room temperature. To reduce nonspecific binding, cells were blocked with 1% bovine serum albumin for 60 min at room temperature. Anti‐VEGF primary antibody solution (1:200) was added and incubated overnight at 4°C. Subsequently, cells were washed 3 times with PBST buffer (PBS containing 0.1% Tween 20) to remove unbound primary antibody. Secondary antibody (1:1000) was added and incubated for 1 h at room temperature in the dark. The cells were observed under a fluorescence microscope (Nikon, Japan). HUVECs were inoculated in 6‐well plates and co‐cultured with various scaffold extracts at 37°C until the cell density exceeded 90%. Total protein was extracted from each cell group and denatured by heating at 95°C–100°C for 10 min. Western blot analysis was then performed to test whether the scaffolds could up‐regulate VEGF (1:1000) expression by up‐regulating HIF‐1α. The experimental procedure was described in Section 2.2.

2.7. In Vitro Anti‐Inflammatory Evaluation

SD rat femurs were taken and placed in 6‐well plates containing antibiotic‐supplemented medium. The bone marrow was flushed with a sterile syringe containing culture medium, then centrifuged. The obtained BMDMs were resuspended in complete medium containing 40 ng/mL M‐CSF and cultured until they reached 90% confluence. Macrophage marker CD68 was used for immunofluorescence staining.

To investigate how different scaffolds influence the polarization state of BMDMs, specifically whether they promote a shift towards M1 (Macrophage Type I) or M2 (Macrophage Type II), we conducted immunofluorescence staining and Western blot experiments as outlined in section 2.6. Additionally, we performed an ELISA assay to validate the expression of key inflammatory factors. Immunofluorescence staining was used to visualize the changes in the polarization state of BMDMs. BMDMs were inoculated in 24‐well plates at a density of 1 × 10⁴ cells/well, stimulated with LPS at 1 µg/mL for 24 h, and then co‐cultured with various scaffold extracts for 48 h. Primary antibody concentrations were CD68 (1:500), iNOS (1:50) and Arg‐1 (1:200). Western blot assay was performed to detect the expression levels of key inflammatory cytokines: BMDMs were inoculated in 6‐well plates, stimulated with LPS at a concentration of 1 µg/mL for 24 h and then co‐cultured with various scaffold extracts at 37°C for Western blot analysis. ELISA was used to determine the concentrations of key inflammatory factors TNF‐α, IL‐4, IL‐6, and IL‐10. Briefly, BMDMs were inoculated into 12‐well plates at a density of 1 × 10⁵ cells/well and co‐incubated with various composite scaffold extracts for 24 h. After incubation, the supernatants were collected. The concentration of inflammatory factors was determined using ELISA kits. By comparing cytokine concentrations across treatment groups, we aimed to evaluate the effects of scaffolds on cellular inflammatory responses and their potential immunomodulatory mechanisms.

2.8. In Vitro Whole Blood Clotting, Platelet Adhesion, Hemolysis, and In Vivo Hemostasis

Whole blood coagulation was used to assess the coagulation effect of 35 mg gauze, and samples (M‐CMCS‐CeO₂ and MBWD scaffolds) were each added with 100 µL whole blood and incubated at 37°C for 10 min. 10 µL of 0.2 M CaCl₂ solution and 10 mL of PBS were added sequentially, and blood diffusion was observed. The absorbance of the PBS solution at 540 nm after blood diffusion was recorded as As. Whole blood mixed with PBS was used as a negative control (Ac reference value), and BCI was calculated by the formula: BCI (%) = As / Ac × 100%.

Platelet adhesion tests were done with samples (M‐CMCS‐CeO₂ and MBWD scaffolds) placed in 24‐well plates and rabbit whole blood dropped into each well. All samples were incubated at 37°C for 1 h, then washed with PBS to remove adhering blood cells physically. Samples were fixed with 4% paraformaldehyde for 2 h and dehydrated using a series of ethanol gradients (30%, 50%, 70%, 80%, 90%, and 100%) for 10 min each. Samples were dried, and platelet and blood cell adhesion was observed under a scanning electron microscope.

Fresh rabbit blood was used for scaffolds (DMOG@PCL/ASC, M‐CMCS‐CeO₂ and MBWD scaffolds) hemolysis assay. Red blood cells (RBCs) were obtained by centrifugation (1500 rpm, 10 min) and diluted with PBS to obtain RBCs suspension. The scaffolds were respectively immersed in RBCs suspension for 2 h at 37°C. After centrifugation at 1500 rpm for 10 min, the supernatant (H_t ) was collected, and its absorbance was measured at 541 nm. DI (H_p ) and PBS (H _n) served as the positive and negative controls, respectively. The hemolysis ratio was quantified using the formula: Hemolysis Ratio (%) = (H_t—H_n ) / (H_p —H_n ) × 100%.

The hemostatic performance of scaffolds on non‐compressible wounds was evaluated using SD rats (200–250 g, male) with liver incisions and tail amputations. Gauze, M‐CMCS‐CeO₂ and MBWD scaffolds were studied without any treatment as a control group. After anesthetizing the SD rats with sodium pentobarbital, an 8 mm incision was made on the left lobe of the liver, and after allowing the blood to flow for 5 s, the sample was applied with gentle pressure to cover the wound and stop the bleeding. The blood loss was recorded (n = 6). One‐third of the rat's tail was amputated using surgical scissors, exposed to the air for 5 s, and the sample was applied to the bleeding site to measure blood loss (n = 6).

2.9. Evaluation of Wound Healing and Pathological Analysis In Vivo

Wound models (infectious and diabetic) were established in SD rats, and the animal experiments were approved by the Animal Ethics Committee of Army Medical University (Ethical Approval Number: AMUWEC20245251). Regarding the infectious wound, SD rats were randomly divided into four groups (n = 4): an untreated control group, a DMOG@PCL/ASC group, an M‐CMCS/CeO₂ group, and an MBWD group. After anesthesia with sodium pentobarbital, a 15 mm circular wound was created on the back of each rat, and 50 µL of a bacterial suspension containing 1 × 10⁷ CFU of S. aureus was added to the wound. As the culture medium evaporated and bacteria adhered, an infectious wound model was established. After adapting to the standard diet for 1 week, rats were placed on a high‐fat diet. 4‐6 weeks after starting a high‐fat diet, diabetes was induced by a single intraperitoneal injection of 35 mg/kg of 2% streptozotocin (STZ) solution. On Day 3, 7, and 14 after STZ injection, blood glucose levels were measured using a blood glucose meter and a blood sampling device at the tail end of the rat. SD rats with blood glucose levels ≥16.67 mmol/L and weight loss were diagnosed as diabetic. Subsequently, a full‐layer circular skin wound with a diameter of 15 mm was formed on the back of the rat. Each wound was then coated with the scaffolds. Wounds were photographed on Day 0, 3, 7, 10, and 14, and wound areas were accurately measured using ImageJ.

To comprehensively understand histological changes and immune responses during wound healing, wound tissue and major internal organs (heart, liver, spleen, lungs, and kidneys) were collected on Day 3, 7, and 14. Following standard protocols, samples were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned. H&E staining and Masson staining were used to observe the microstructure of wound healing and collagen fiber deposition. All sections were analyzed and recorded using an upright microscope. Furthermore, to explore the immunoregulatory mechanisms of regenerated skin, immunofluorescence staining and immunohistochemistry analysis were performed. On Day 14, excised regenerated skin tissue was used to detect the expression of key inflammatory cytokines IL‐10, IL‐1β, Arg‐1, and TNF‐α. The expression patterns of these cytokines were observed under an inverted fluorescence microscope using specific antibodies and DAPI nuclear staining.

2.10. Statistical Analysis

GraphPad Prism 6 software was used for comprehensive data analysis to ensure the accuracy and reliability of the results. All experiments were independently repeated at least three times to verify the stability and reproducibility of the results. Experimental data were presented as mean ± standard deviation. Statistical comparisons among multiple groups were performed using one‐way ANOVA followed by Tukey's post hoc test. Statistical significance levels were set at *p < 0.05, **p < 0.01, and ***p < 0.001 to clearly distinguish whether differences in efficacy between different treatment groups were statistically significant.

3. Result and Discussion

3.1. The Determination of DMOG

We initially evaluated the impact of DMOG concentrations (250, 500, 750, and 1000 µM) on HUVEC proliferation using the CCK‐8 assay to identify an effective concentration. As depicted in Figure S1, on Day 1, 3, and 5, all groups exhibited a trend of cell proliferation. On Day 3 and 5, HUVEC proliferation decreased gradually with increasing DMOG concentration. However, on both Day 3 and 5, the proliferation effect of the 250 µM‐treated group was better than that of the other concentration groups, and was significantly different (p < 0.001). Furthermore, we investigated the effect of varying DMOG concentrations on HIF‐1α expression levels. Western blot analysis (Figure S2A,B) revealed that as DMOG concentration increased, HIF‐1α expression correspondingly increased. Subsequent statistical analysis indicated that at concentrations of 750 and 1000 µM, HIF‐1α accumulation approached saturation, with no significant difference between the two groups (p > 0.05). Based on these dual experimental outcomes, to ensure that the endothelial cells maintained optimal proliferation and simultaneously promoted appropriate levels of HIF‐1α accumulation to aid in angiogenesis [26], we chose 250 µM as the effective dose and incorporated it into the designed delivery system (abbreviation: DMOG@PCL/ASC).

3.2. Characterization of DMOG@PCL/ASC, M‐CMCS/CeO₂, and the MBWD Scaffolds

As shown in Figure 1A,B, PCL/ASC coaxial nanofibers were produced by coaxial electrospinning, with their shell layers labeled with Rhodamine B to exhibit clear red fluorescence. This method resulted in core‐shell fibers with relatively uniform diameters of 650–750 nm (Figure 1C). Figure 1D,E shows coaxial microparticles containing DMOG, generated by electrospray, with their shells labeled with FITC‐BSA, which emits green fluorescence. SEM images revealed that these microparticles have uniform sizes, nearly spherical morphologies, and diameters of about 4–6 µm (Figure 1F). These two techniques were combined to successfully build the DMOG@PCL/ASC scaffold as seen in the SEM image in Figure 1G.

FIGURE 1.

Characterization of the MBWD scaffold. (A) SEM image, (B) fluorescence image, and (C) frequency distribution histogram of PCL/ASC nanofibers. (D‐E) SEM and fluorescence images and (F) frequency distribution histogram of coaxial microparticles containing DMOG. (G) SEM image of DMOG@PCL/ASC. (H) TEM image of CeO₂ nanozymes. (I) Photocured form of M‐CMCS and M‐CMCS/CeO₂ hydrogels. (J) Injectability of M‐CMCS/CeO₂ in air and under water. (K) Representative SEM images and quantitative pore size analysis of freeze‐dried M‐CMCS, M‐CMCS/CeO₂ hydrogels, and MBWD. (L) Porosity diameter statistics. (M) Swelling ratio. (N) Contact angle. (O) FTIR spectra. (P) Degradation ratio. (Q) The release of DMOG.

CeO₂ nanozyme morphology was observed with TEM (Figure 1H), measuring 2–5 nm in diameter. The vial‐inversion method was used to assess the photocuring performance of M‐CMCS and M‐CMCS/CeO₂ hydrogels. Figure 1I shows that both hydrogels were fluid at room temperature and solidified quickly under 405 nm UV light, showing excellent photocurable properties. This is achieved through UV‐induced radical polymerization of carbon‐carbon double bonds, resulting in a crosslinked network generated by the reaction between the epoxy groups of glycidyl methacrylate and the amino groups of carboxymethyl chitosan, thereby improving mechanical strength and making it suitable for wound defect repair [27]. Figure 1J demonstrates that the M‐CMCS/CeO₂ hydrogel exhibits good injectability in both air and underwater environments, enabling it to treat a wide range of wet and irregularly shaped wounds that meet various clinical needs. SEM images (Figure 1K) revealed that M‐CMCS, M‐CMCS/CeO₂, and MBWD scaffolds all had irregular porous network structures, with M‐CMCS and M‐CMCS/CeO₂ exhibiting pore sizes of approximately 200 µm (Figure 1L). Photographs of the front and reverse sides of the bilayer construct after 1, 3, and 5 days of incubation demonstrate no delamination or separation between the hydrogel and nanofiber layers, confirming the structural integrity of the bilayer design (Figure S3). The inner and outer layers of the MBWD scaffold bond tightly. Plasma treatment of the inner DMOG@PCL/ASC nanofibers introduces carboxyl groups (‐COOH) on the surface, thereby improving hydrophilicity and introducing additional active functional groups. The outer M‐CMCS hydrogel contains glycidyl groups (epoxy groups) from GMA modification and the carboxyl groups undergo an ring‐opening reaction with these glycidyl groups to form covalent bonds.

Figure 1M shows that adding CeO₂ had little effect on the swelling ratio of M‐CMCS hydrogel, while the MBWD scaffold's swelling ratio increased significantly. This may relate to the enhanced hydrophilicity of DMOG@PCL/ASC scaffold after plasma treatment, and the hydrogel's porosity, combined with a high swelling ratio, promotes nutrient penetration and metabolite excretion to support cell and vessel growth [28, 29, 30, 31]. Figure 1N demonstrates that DMOG@PCL/ASC has a smaller contact angle, enabling tight attachment to the wound bed and reducing the available space for bacterial survival. M‐CMCS and M‐CMCS/CeO₂ hydrogels are highly hydrophilic, retaining moisture to form a moist environment that favors cell growth and wound healing. Figure 1O indicates that in the 3200‐3500/cm wavenumber range, M‐CMCS's hydroxyl vibrations show a distinct broad absorption peak. Integrating CeO₂ into the M‐CMCS hydrogel shifts this peak slightly to lower frequencies due to hydrogen‐bond formation, resulting in increased peak complexity that reflects interactions between M‐CMCS molecules and hydroxyl or adsorbed water molecules on the CeO₂ surface. In the 1600–1750/cm wavenumber range, M‐CMCS's carboxyl vibrations exhibit a characteristic absorption peak. However, adding CeO₂ may make the peak in this region more complex. This complexity comes from M‐CMCS's own carboxyl vibrations, potential contributions from hydroxyl or adsorbed water molecules on the CeO₂ surface, and vibrations of possible chemical bonds between M‐CMCS and CeO₂. Scaffold degradation should coincide with the growth of new wound tissue. The M‐CMCS and MBWD groups degrade relatively rapidly, reaching nearly 80% degradation in 4 weeks (Figure 1P). Fluorescein isothiocyanate‐labeled bovine serum albumin was released from the particles within 0–3 days, with cumulative release reaching 71.26% it increased to 75.66% by Day 7 and was slowly released continuously until Day 21, as shown in Figure 1Q. To clarify the structural advantage of the bilayer architecture, we compared DMOG release kinetics between the MBWD and the SLM scaffold containing identical active components. As shown in Figure S4, the SLM exhibited a pronounced burst release (46% at 4 h), whereas MBWD maintained a sustained release profile (20% at 4 h), confirming the regulatory role of spatial separation.

3.3. Biocompatibility and H₂O₂ Scavenging Activity

As shown in Figure 2A, all groups of L929 cells’ viability gradually increased over time, demonstrating excellent biocompatibility. However, the cell proliferation trend in the M‐CMCS/CeO₂‐300 group was consistently lower than in all other groups at Day 3 and 5, and was significantly different from that in the control group (p < 0.001), indicating that the cell proliferation rate may be inhibited when the CeO₂ concentration exceeds a certain range. In many cellular experiments, low concentrations of certain substances may be harmless or even beneficial to cells, but when their concentration exceeds a threshold, they can interfere with normal cellular physiological processes. Further analysis of Figure 2B showed that the H₂O₂ scavenging rates of the M‐CMCS/CeO₂‐100, M‐CMCS/CeO₂‐200, and M‐CMCS/CeO₂‐300 groups were 81.84%, 82.49%, and 87.01%, respectively, and the differences among the three groups were not statistically significant (p > 0.05). CeO₂ has been proven to have antioxidant function in studies of wound healing and neurodegenerative disorders, among others. When CeO₂ encounters H₂O₂, it is able to decompose H₂O₂ into water and oxygen through redox reactions. In this process, Ce^{4 +} is reduced to Ce^{3 +}, while H₂O₂ is reduced to water, which reduces the content of H₂O₂ and serves to remove H₂O₂. Therefore, based on the results of cell proliferation and H₂O₂ scavenging assays, the M‐CMCS/CeO₂‐200 scaffold (M‐CMCS/CeO₂) was selected for subsequent studies.

FIGURE 2.

Biocompatibility and H₂O₂ Scavenging Capability. (A) Biocompatibility assessment of L929 cells and (B) determination of H₂O₂ scavenging rates. (C) Live/dead staining of L929 cells and (D) HUVECs. (E) RAW264.7 and (F) L929 cell ROS staining. (G) Migration ability of L929 cells co‐cultured with various scaffold extracts for 24 h and (H) statistical graph of migration rate. (*p < 0.05, **p < 0.01, ***p < 0.001, compared with control group).

3.4. In Vitro Biocompatibility, ROS Scavenging, and Antibacterial Properties of MBWD Scaffold

We used live/dead staining to check the effects of DMOG@PCL/ASC, M‐CMCS‐CeO_2, and MBWD scaffolds on cell viability. Figure 2C,D shows that live cells (green) were the predominant type, with very few dead cells (red) in both L929 cells and HUVECs, indicating that the three constructed scaffolds have good biocompatibility. After 3 days of treatment, L929 cells in each group showed no morphological changes compared with the control group (Figure S5). DCFH‐DA fluorescence probe assays (Figure 2E,F) revealed that all treatment groups exhibited varying degrees of ROS scavenging in both the LPS‐induced RAW 264.7 macrophage inflammation model and in L929 cells, and green fluorescence intensity correlated positively with ROS content. Experimental results show the MBWD and M‐CMCS‐CeO₂ groups had the weakest fluorescence, demonstrating the strongest antioxidant capacity. The DMOG@PCL/ASC group showed less effective ROS scavenging, and the positive control group displayed the most intense fluorescence. This finding was validated across distinct cell line types, confirming MBWD's intracellular ROS‐scavenging capacity. Additionally, we investigated the impact of the three scaffolds on cell migration using an L929 cell scratch test. As depicted in Figure 2G,H, after 24 h of incubation, the healing rates for the DMOG@PCL/ASC, M‐CMCS/CeO₂, and MBWD scaffolds were 42.78%, 43.01%, and 41.1%, respectively, while the control group was only 41.76%. No significant differences were observed among groupsHowever, there was no significant difference in migration rates among the scaffold groups (p > 0.05), indicating that the three scaffolds were equally effective, with none showing significantly better migration‐promoting properties than the others. Both experiments demonstrated that these scaffolds have good cell biocompatibility.

This study selected E. coli and S. aureus to test the antibacterial activity of different scaffolds in vitro. Figure 3A,B shows the experimental results after scaffold‐bacteria co‐culture, with the control group having the most bacterial colonies, followed by the DMOG@PCL/ASC group. This relates to the nanofibers’ unique physical network, characterized by a large specific surface area and porosity, which partly inhibits bacterial activity and slows their proliferation. Notably, bacterial colony numbers decreased sharply in the M‐CMCS/CeO₂ and MBWD groups, with no significant difference (p > 0.05), and antibacterial efficiency exceeding 95% against both E. coli and S. aureus, as verified by the quantitative statistics in Figure 3D,E. The antibacterial activity mainly results from electrostatic interactions between the cationic groups of CMCS and negatively charged bacteria, which destabilize bacterial cells and lead to bacterial death [32, 33, 34]. CeO₂ nanozymes damage bacterial cell membranes and intracellular biomolecules by catalyzing reactions that generate ROS (such as hydroxyl radicals and superoxide anions), resulting in bacterial death, which forms CeO₂’s antibacterial mechanism.

FIGURE 3.

In vitro antibacterial activity against E. coli and S. aureus. (A) Digital camera images of bacterial colonies of E. coli and S. aureus. (B) Fluorescent images of bacterial live/dead staining (green: live bacteria, red: dead bacteria). (C) Digital photographs of S. aureus biofilm formation. (D‐E) Statistical graphs of antibacterial rates for S. aureus and E. coli. (F) Statistical graph of bacterial biofilm formation. (*p < 0.05, **p < 0.01, ***p < 0.001, compared with each other).

To further check bacterial viability after treating with the three scaffolds, we used live/dead bacterial staining (Figure 3B). Results showed high bacterial survival in the untreated control group (prominent green fluorescence). Treatment with DMOG@PCL/ASC caused a slight increase in the proportion of dead bacteria (stronger red fluorescence) and almost no live bacteria were found in the M‐CMCS/CeO₂ and MBWD groups (notable red fluorescence), which matches the quantitative results (Figure 3D,E). Resistant bacteria forming biofilms pose a significant challenge to antibiotic treatment. This study examined how well different scaffolds can break down biofilms in a bacterial infection environment by establishing an S. aureus biofilm model (Figure 3C). Using chitosan's ability to penetrate biofilms, the M‐CMCS/CeO₂ and MBWD groups effectively disrupted biofilms and performed better than the control and DMOG@PCL/ASC groups, with a significant difference between the MBWD and control groups (p < 0.001), as supported by the biofilm quantitative results in Figure 3F.

3.5. Modulation of HUVECs Behavior by MBWD Scaffold In Vitro

The efficiency of angiogenesis significantly influences the time and outcome of wound healing [35, 36, 37]. HUVECs migration results (Figure 4A,B) demonstrated that the migration rates of the DMOG@PCL/ASC and MBWD groups after 24 h were 90.01% and 83.65%, respectively, which were much higher than those of the control group (53.68%) and the M‐CMCS/CeO₂ group (60.04%). EdU assay results (Figure S10) indicated that DMOG@PCL/ASC and MBWD groups promoted HUVECs proliferation compared to the other two groups, and were significantly different from the control group (p < 0.01). Transwell migration assay results (Figure 4C,D) also showed that the DMOG@PCL/ASC and MBWD groups had the highest number of migrated HUVECs, and were significantly different from the control group (p < 0.001), consistent with the EdU assay results.

FIGURE 4.

In vitro HUVECs behavior. (A) Migration ability of HUVECs co‐cultured with various scaffold extracts for 24 h and (B) corresponding statistical graph. (C) Transwell migration assay and (D) statistical graph. (E) VEGF immunofluorescence staining and (F) statistical graph. (G) Angiogenesis assay and (H) statistical graphs of various angiogenic indicators. (I) Western blot analysis of HUVECs protein levels (HIF‐1α and VEGF) and (J‐K) corresponding statistical graphs. (*p < 0.05, **p < 0.01, ***p < 0.001, compared with each other).

We used angiogenic assays to check how the MBWD scaffold affects angiogenesis. Figure 4G,H shows that, compared with the control and M‐CMCS/CeO₂ groups, the DMOG@PCL/ASC and MBWD groups promoted greater vascular growth, with more node junctions, master junctions, and meshes. These two groups also increased the expression of angiogenesis‐related proteins HIF‐1α and VEGF, demonstrating a stronger angiogenic capacity than the control and M‐CMCS/CeO₂ groups (Figure 4I–K). VEGF immunofluorescence staining (Figure 4E,F) had the same results as VEGF Western blot experiments, and VEGF intensity in both DMOG@PCL/ASC and MBWD groups was higher than the control group with significant differences (p < 0.05). These findings demonstrate that the DMOG@PCL/ASC and MBWD groups enhance the vasogenic function of HUVECs, as both contain DMOG [38, 39, 40]. DMOG is a competitive inhibitor of prolyl hydroxylase (PHD); it stabilizes HIF‐1α and mimics hypoxia responses even under normoxic conditions, speeding up angiogenesis. Microvascular system formation during tissue regeneration takes around 2‐4 weeks, so steady release of DMOG supports angiogenesis.

We further compared the release rates of SLM and MBWD, and VEGF secretion showed a distinct temporal pattern. The SLM group reached a peak at 48 h (330 pg/mL), then decreased at 72 h (310 pg/mL), whereas the MBWD group continued to increase to 472 pg/mL at 72 h, consistent with sustained DMOG release. This differentiation further supports the stage‐dependent functional regulation of the bilayer structure (Figures S6). Interestingly, the SLM group showed slightly higher EdU incorporation than MBWD, reflecting the transient burst‐release effect at 24 h (Figures S7A). However, at 48 h, MBWD surpassed SLM, indicating sustained proliferative stimulation enabled by controlled release (Figures S7B). At 48 h, although wound closure approached saturation in all groups, MBWD still demonstrated a significantly higher migration rate (91.9%) compared to SLM (83.2%), suggesting prolonged pro‐migratory activity (Figures S8). In the 48‐hour tube formation experiment (Figures S9), although SLM has formed a relatively complete vascular network, MBWD still shows a slight advantage in NB Junctions and NB Meshes. It is worth noting that the functional gap is smaller than the VEGF secretion gap, suggesting that vascular network formation is regulated by multiple factors and is not a simple linear response. In conclusion, SLM performs outstandingly in the early stage, while MBWD achieves continuous regulation and functional enhancement in the middle and later stages. The double‐layer structure achieves synergy through structure‐driven optimization of release timing.

3.6. In Vitro Anti‐Inflammatory Effects of MBWD

We studied how DMOG@PCL/ASC, M‐CMCS/CeO₂ and MBWD scaffolds influence the polarization of BMDMs in vitro. Figure 5A,B show that we identified isolated cells expressing the CD68 marker, and all groups exhibited positive green fluorescence, clearly confirming that the cells were primary BMDMs. Observation revealed that the proportion of iNOS‐positive cells in the M1 phenotype marker in the M‐CMCS/CeO₂ and MBWD groups was considerably lower than in the other two groups. The proportion of M2 phenotype marker Arg‐1‐positive cells was notably higher (Figure 5C,D). These findings suggest that the active components in M‐CMCS/CeO₂ and MBWD may exert significant anti‐inflammatory effects, thereby effectively enhancing the expression of anti‐inflammatory factors. Western blot results confirmed this. In an LPS‐induced inflammatory environment, the expression of pro‐inflammatory factors CD86 and iNOS increased significantly. M‐CMCS/CeO₂ and MBWD notably decreased the protein levels of these two pro‐inflammatory factors and elevated the M2 phenotype markers CD206 and Arg‐1 protein expression (Figure 5E–I), with significant differences compared with the LPS‐induced group (p < 0.05).

FIGURE 5.

Regulation of macrophage immune properties. (A‐B) Immunofluorescence staining images of primary macrophage markers CD68, iNOS, and Arg‐1. (C) Statistical graph of iNOS and (D) Arg‐1 immunofluorescence staining. (E) Western blot analysis of the impact of different scaffolds on the protein expression level of BMDMs. (F‐I) Statistical graphs of western blot analysis for iNOS, Arg‐1, CD86, and CD206. (J) ELISA detection of pro‐inflammatory cytokines (TNF‐α and IL‐6) and anti‐inflammatory cytokines (IL‐4 and IL‐10). (*p < 0.05, **p < 0.01, ***p < 0.001, compared with each other).

The ELISA results (Figure 5J) also supported the above conclusions. In the M‐CMCS/CeO₂ and MBWD groups, the expression levels of M1 phenotype markers TNF‐α and IL‐6 were significantly lower than in the other two groups, while the expression levels of M2 phenotype markers IL‐4 and IL‐10 were significantly higher, and the difference was significant compared with the control group (p < 0.001). Previous studies have shown that Ce³⁺ and Ce⁴⁺ ions in CeO₂ can undergo reversible redox conversion, mimicking the functions of superoxide dismutase (SOD) and catalase (CAT) through their abundant surface oxygen vacancies, thereby effectively reducing or eliminating ROS production. Reduction of ROS can inhibit the activation of the nuclear factor κB (NF‐κB) pathway, thereby down‐regulating the transcription of pro‐inflammatory cytokines such as TNF‐α and IL‐6 [41, 42].

3.7. In Vitro Coagulation, Blood Compatibility, and In Vivo Hemostasis

We performed whole blood coagulation tests to quantitatively confirm the scaffolds’ hemostatic efficacy. Previous studies have clearly demonstrated that BCI exhibits a negative correlation with coagulation performance—higher BCI values signify weaker coagulation [43, 44]. Figure 6A,B shows that M‐CMCS/CeO₂ and MBWD groups have notably lower BCI values than the gauze group (p < 0.001), fully reflecting the two scaffolds’ superior coagulation capability. This excellent performance stems from M‐CMCS's intrinsic porous structure that effectively absorbs blood, while MBWD's bilayer design—its inner DMOG@PCL/ASC nanofiber layer with a suitable three‐dimensional structure—promotes platelet adhesion aggregation and the adsorption and activation of surface coagulation factors triggering the coagulation cascade. The remarkable hemostatic ability of M‐CMCS scaffolds is closely linked to their porous structure and surface amino groups, which stimulate platelet aggregation and erythrocyte adhesion, thereby forming an interlinked network that absorbs more exudates [27, 45]. Figure S11 shows strong adhesion of blood cells and platelets to the scaffolds, directly demonstrating the outstanding hemostatic performance of the M‐CMCS/CeO₂ and MBWD groups. We also evaluated the blood compatibility of DMOG@PCL/ASC, M‐CMCS/CeO_2, and MBWD scaffolds using a hemolysis test (Figure 6C) results show all scaffold groups have hemolysis rates of less than 0.4% with no notable differences between groups (p > 0.05), well below the 2.0% international standard [45], confirming good blood compatibility.

FIGURE 6.

In vitro and in vivo blood compatibility. (A) Photos of gauze and various scaffolds after contacting with blood. (B) BCI statistical results. (C) Optical images and statistical graph of hemolysis. (D) Representative images and schematic illustration of hemostatic performance of different scaffolds in liver resection and tail amputation models. (E) Blood loss quantification for different scaffolds in tail amputation and (F) liver resection models. (*p < 0.05, **p < 0.01, ***p < 0.001, compared with each other).

To further verify the hemostatic effect of various scaffolds in a real physiological environment, we designed two hemorrhage models: the rat liver incision model and the tail amputation hemostatic experiment. As shown in Figure 6D–F, in the liver incision model, the blood loss of the M‐CMCS/CeO₂ group and the MBWD group was (51 ± 3) mg and (45 ± 6) mg, respectively, which was significantly lower than that of the control group (332 ± 3) mg. The results of the tail amputation model were consistent. Compared with the control group, the hemostatic effects of the M‐CMCS/CeO₂ and MBWD groups were more significant (p < 0.001). This may be because CMCS not only activates red blood cells and induces platelet aggregation, but also its three‐dimensional porous structure acts as a physical barrier, helping to stop bleeding.

3.8. Accelerating the Healing of Infectious Wounds

This study aims to thoroughly evaluate the infectious wound healing ability of DMOG@PCL/ASC, M‐CMCS/CeO₂ and MBWD scaffolds by setting up an S. aureus‐infected full‐thickness skin defect model in rats. Figure 7A,B shows that during continuous treatment, the M‐CMCS/CeO₂ and MBWD groups accelerated wound healing more than the other groups. The MBWD group showed especially notable effects. On treatment Day 3, the MBWD group achieved a 35% wound closure rate, which was higher than that of the control and DMOG@PCL/ASC groups (p > 0.05). We also measured wound bacterial loads on Day 3 (Figure 7C). Results showed that the M‐CMCS/CeO₂ and MBWD groups had notably lower bacterial numbers than the control and DMOG@PCL/ASC groups. As treatment continued, the MBWD group exhibited the most prominent healing effect on Day 7 and 10, with wound closure rates of 76% and 93%, respectively. By Day 14, wounds in this group were nearly completely healed (approximately 95%), demonstrating their exceptional healing capability. At Day 14, we found that the wound‐healing rate in the M‐CMCS/CeO₂ group was also close to 93%. It was observed that wounds heal faster and better in a moist environment provided by hydrogels, which is due to this environment facilitating cell proliferation and neovascularization, among other benefits [46, 47, 48]. When excessive wound exudate is present, using a hydrogel dressing to absorb it can effectively manage wound wetness, maintain a suitable moist environment, and prevent exudate accumulation. The introduction of the M‐CMCS hydrogel layer in the MBWD scaffold is precisely the addition of this wet environment management. In addition, during wet wound healing, the wound area at the beginning of healing may even be larger than at the beginning due to the higher water content of the wound tissue, which is a normal phenomenon.

FIGURE 7.

Infectious wound healing in vivo. (A) Views of wound closure on Day 0, 3, 7, 10, and 14 for each group. (B) Statistical graph of wound healing rates. (C) Digital images of S. aureus colonies of wounds on Day 3. (*p < 0.05, **p < 0.01, ***p < 0.001, compared with MBWD group).

To gain insights into histological changes during wound healing, skin samples were collected on Day 3, 7, and 14 post‐surgery for H&E and Masson staining. On Day 3 (Figure S12), both H&E and Masson staining revealed that the epidermis, granulation tissue, and collagen production in the injured skin were at an early growth stage. Notably, the MBWD group exhibited lower levels of inflammation compared to other groups, potentially due to the excellent antibacterial and ROS‐scavenging properties of its constituents. On Day 7 (Figure 8A), the control and DMOG@PCL/ASC groups’ wounds were heavily populated by inflammatory cells, blocking the formation of neonatal epidermis. The M‐CMCS/CeO₂ and MBWD groups had not completed full epithelialization; however, the notable reduction in inflammatory cell infiltration facilitated wound healing. By Day 14 (Figure 8B), all scaffold‐treated wounds had fully re‐epithelialized. The MBWD group exhibited favorable regeneration of skin appendages, including hair follicles and sebaceous glands, indicating optimal re‐epithelialization. Collagen deposition served as a marker of wound‐healing quality. Masson staining on Day 14 (Figure 8C,D) showed that the DMOG@PCL/ASC group exhibited greater collagen accumulation than the control group, with a significant statistical difference between them (p < 0.01), likely because DMOG release speeds up tissue blood vessel development and provides damaged tissues with sufficient oxygen and nutrients. The control and M‐CMCS/CeO₂ groups showed sparse collagen accumulation compared with the other groups. The MBWD group exhibited the highest collagen deposition, with statistical significance compared to the other groups (p < 0.001), and its collagen fibers were more neatly arranged, aligning with the structural characteristics of normal skin tissue. This phenomenon arises from DMOG's pro‐angiogenic activity as well as the M‐CMCS hydrogel and its loaded CeO₂’s capacity to establish an antibacterial, antioxidant, and anti‐inflammatory microenvironment.

FIGURE 8.

Tissue staining analysis in vivo. (A) H&E staining of wound tissue on Day 7 and 14 for each group. (B) Statistical graph of epidermal thickness on Day 14. (C) Masson staining of wound tissue on Day 7 and 14 for each group. (D) statistical graph of collagen deposition on Day 14. (E) TNF‐α immunofluorescence staining and (F) statistical graph. (G) IL‐10 immunofluorescence staining and (H) statistical graph. (*p < 0.05, **p < 0.01, ***p < 0.001, compared with each other).

During physiological skin healing, M1 macrophages dominate the initial stage (Day 1–3) to eliminate pathogens and necrotic debris through phagocytosis. In the proliferative phase (Day 3–7), macrophages gradually polarize toward the M2 phenotype to initiate anti‐inflammatory repair processes. In pathological microenvironments such as infected or diabetic wounds, prolonged M1 polarization sustains persistent inflammation, thereby hindering wound healing. On Day 3 post‐treatment, the MBWD scaffold restrained excessive inflammatory responses while retaining baseline M1 macrophage polarization. H&E staining and bacterial quantification confirmed that the reserved M1 macrophages efficiently eliminated S. aureus and ensured antibacterial activity. Following infection clearance on Day 7, the sustained release of CeO₂ nanozymes remodeled the wound microenvironment from a pro‐inflammatory state toward a reparative phenotype. Unlike most existing scaffolds that force M2 polarization within 48 h, MBWD recapitulates the physiological immune cascade. It preserves early antibacterial defense and achieves favorable M2 polarization in the late healing stage, ultimately improving the overall quality of wound regeneration.

For comprehensive evaluation of the biosafety of each scaffold, we performed histopathological examinations on the major organs of experimental animals, with no significant pathological changes observed in the heart, liver, spleen, lungs or kidneys of animals in all groups (Figure S13), confirming good biocompatibility of each scaffold. On Day 14 post‐wounding, we conducted immunofluorescence analysis on wound tissues focusing on clarifying the expression characteristics of two key immunoregulatory factors, tumor necrosis factor‐α (TNF‐α) and interleukin‐10 (IL‐10), where TNF‐α is a pro‐inflammatory cytokine mainly produced by macrophages and monocytes involved in normal inflammatory responses and immune processes [49], and IL‐10 is a potent anti‐inflammatory cytokine capable of inhibiting pro‐inflammatory cytokine production and promoting immune cell apoptosis that plays a core role in inflammation regulation and tissue repair [50]. Figure 8E–H shows the expression of TNF‐α and IL‐10 in different groups, with quantitative analysis results (Figure 8E,F) demonstrating the lowest TNF‐α expression level in the MBWD group with significant differences from other groups (p < 0.05) followed by the M‐CMCS/CeO₂ group, indicating the anti‐inflammatory activity of CeO₂ in these two groups, while the control and DMOG@PCL/ASC groups exhibited significantly elevated TNF‐α expression suggesting a high inflammatory state possibly due to lack of effective immune regulation. The MBWD group showed significantly higher IL‐10 expression levels than the other three groups (Figure 8G,H), with statistically significant differences (p < 0.001), confirming the remarkable efficacy of MBWD in regulating inflammation and providing strong support for its application in the treatment of inflammation‐related diseases.

3.9. Accelerating the Healing of Diabetic Wounds

The regenerative potential of MBWD was further evaluated in a diabetic wound model and compared with that in an infectious wound model to delineate shared and distinct therapeutic mechanisms across diverse pathological landscapes. Consistent with observations in the infectious model, Figure 9A,B demonstrates that MBWD significantly expedited closure, achieving near‐complete re‐epithelialization with minimal scarring by Day 14, whereas the control group exhibited persistent tissue defects. These findings underscore that maintaining a moist microenvironment, coupled with ROS scavenging, antimicrobial activity, and immunomodulation, constitutes the universal foundation for MBWD. With the different pathological mechanisms of the two wound models, the therapeutic focus of MBWD shifted accordingly. In infectious wounds, where the primary hurdles are bacterial colonization and acute inflammation, MBWD primarily functions as a protective shield, focusing on rapid sterilization and inflammation control. In contrast, diabetic wounds are characterized by chronic low‐grade inflammatory stasis, severe angiogenic impairment, and disorganized extracellular matrix deposition, which collectively stall the healing progression. Histological evidence further substantiated these divergent roles. While MBWD primarily attenuated inflammatory infiltration and localized damage in infectious wounds (Figure 8), its impact on epithelial integrity, granulation tissue thickness, and collagen regularity was even more pronounced in the diabetic model (Figure 9C,D). Masson staining revealed that MBWD not only enhanced collagen quantity but also orchestrated a more ordered fiber arrangement. Immunohistochemical analysis (Figure 9E) confirmed that MBWD significantly downregulated IL‐1β and upregulated Arg‐1, paralleling the M2‐polarization trend observed in infectious models. The ability of MBWD to disrupt this dysfunctional immune niche is critical for propelling the wound into the reparative phase. Notably, morphometric quantification (Figure 9F–H) indicated that the relative improvements in granulation thickness, epidermal restoration, and collagen deposition rate were quantitatively higher in diabetic wounds than in infectious ones. This indicates that under the pathology of diabetes, the synergistic interaction of pro‐angiogenic, antioxidant and immunomodulatory signals in MBWD provides a superior microenvironment and effectively promotes the regenerative process.

FIGURE 9.

Wound healing in vivo. (A) Views of wound closure on Day 0, 3, 7, 10, and 14. (B) Statistical graph of wound healing rates. (C) H&E and (D) Masson on Day 7, 14 days for each group. (E) IL‐1β and Arg‐1 immunohistochemistry staining. (F‐H) statistical graph. (***p < 0.001, compared with each other).

It is well established that diabetic wounds are characterized by a prolonged inflammatory phase, during which excessive ROS levels sequester macrophages into a persistent M1 phenotype. Within this pathological milieu, MBWD effectively reignites the stalled immune regulatory program. By Day 14, the marked up‐regulation of Arg‐1 and the concomitant down‐regulation of IL‐1β demonstrate that MBWD treatment successfully disrupts the pro‐inflammatory loop. Mechanistically, this transition is primarily attributable to the early quenching of high‐glucose‐induced ROS by the M‐CMCS/CeO₂, which lowers the metabolic threshold for BMDMs polarization toward the reparative M2 state. Concurrently, the sustained release of DMOG from the inner nanofibrous layer stabilizes HIF‐1α, ensuring that the microenvironment is not only anti‐inflammatory but also pro‐angiogenic.

The advantages of MBWD in accelerating re‐epithelialization and collagen maturation stem from the synergistic effect between its bilayer components, a performance that single‐layer intervention methods cannot match. The limitation of single‐layer M‐CMCS/CeO₂ lies in its inability to provide long‐term structural guidance and continuous biological induction signals for tissue reconstruction. Although it can effectively manage water balance and microbial load, the lack of fiber scaffolds can lead to disordered collagen deposition and insufficient depth of angiogenesis. On the contrary, monolayer DMOG@PCL/ASC nanofibers, despite their excellent cell adhesion and angiogenic advantages, remain vulnerable in the hostile microenvironment of infected or diabetic wounds. In the absence of external antibacterial shielding, the stability of HIF‐1α induced by DMOG is often covered by excessive pro‐inflammatory cytokines, which drive cells towards apoptosis rather than proliferation.

In contrast, the MBWD bilayer structure achieves an ordered healing process. The outer hydrogel first acts as a barrier for bacterial microorganisms, consuming excessive ROS and preventing secondary infections, thereby effectively reducing the repair threshold of the wound bed. This creates a stable microenvironment, enabling the nanofibers internally loaded with DMOG to fully realize their regenerative potential. The multi‐level pore structure in the inner layer facilitates HUVEC migration, while the sustained release of DMOG supports angiogenesis.

3.10. RNA Sequencing Analysis

Under LPS‐induced inflammatory conditions, the molecular mechanisms in BMDMs co‐cultured with MBWD for 24 h were assessed via RNA sequencing (RNA‐seq, Figure 10). Differential expression analysis, Gene Ontology (GO) enrichment studies, and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analyses were conducted to compare the transcriptomes of BMDM cells between the M1 group (LPS‐induced positive control) and the MBWD group. Principal Component Analysis (PCA) revealed significant transcriptional differences between the M1 and MBWD groups (Figure 10A). In the volcano plot, differentially expressed genes (DEGs) exhibited significant differences between the M1 and MBWD groups (Figure 10B). A total of 3429 genes were detected in both the M1 and MBWD groups, with 1633 upregulated and 1796 downregulated genes.

FIGURE 10.

Analysis of RNA sequencing results for MBWD composite scaffold treatment. (A) PCA analysis presenting differentially expressed genes as identified by RNA‐seq in the M1, MBWD groups. (B) Volcano plots illustrating differentially expressed genes (gray signifies non‐significant genes; red indicates upregulated genes; blue stands for downregulated genes). (C) Expressed genes in the KEGG pathway. (D) Differential gene cluster analysis of M1 and MBWD groups. (E) GO enrichment analysis presenting the DEGs. (F) A chord diagram showing GO enrichment terms along with their corresponding top 10 downregulated genes. (F) Increased/decreased expression of genes during the immune system. (G) Analysis of the differential gene enrichment within the KEGG pathway.

To gain deeper insights into the pathways activated by MBWD, Gene Set Enrichment Analysis (GSEA) was employed to explore relevant signaling pathways (Figure 10C). The results indicated downregulation of NOD‐like receptor and TNF signaling pathways in MBWD‐treated cells. A clustered heatmap in Figure 10D illustrated the expression patterns between the M1 and MBWD groups, highlighting similarities and differences. The distinct expression patterns indicated transcriptome‐level differences between the two groups. The chord diagram in Figure 10E displayed downregulated genes and their corresponding biological processes, such as inflammatory response, immune system process, and immune response. Among the 22 DEGs related to the immune system, expression factors related to validation (TNF, IL‐1β, IL‐12β) exhibited a decreasing trend in the MBWD group (Figure 10F), explaining the downregulation of immune and inflammation‐related genes by the MBWD scaffold. KEGG pathway enrichment analysis identified representative signaling pathways, as shown in Figure 10G. According to previous reports [51, 52, 53], ROS activates the TNF‐α signaling pathway.

4. Conclusion

To address the clinical challenges of infectious skin wounds, this study developed a novel MBWD scaffold with an inner layer of DMOG@PCL/ASC nanofibers, enabling localized and sustained DMOG release to activate HIF‐1α expression in HUVECs, increase VEGF levels, and promote neovascularization. An outer layer of M‐CMCS/CeO₂ loaded with CeO₂ nanozymes that efficiently scavenge excess ROS, regulate inflammatory responses, and exert antibacterial activity to create a favorable microenvironment for tissue regeneration. This MBWD scaffold combines the dual advantages of nanofibers and hydrogels, significantly accelerating the repair of infected wounds and holding great promise for widespread application in the treatment of various wound infections.

Author Contributions

Ziyi Zhou: Methodology, Data Curation, Investigation, Formal analysis, Writing – original draft. Dengjun Zhang: Methodology, Investigation. Qingxia Guo: Methodology, Data Curation. Linbo Jin: Investigation, Supervision. Yuanfei Wang: Investigation, Supervision, Writing – review and editing. Jiaping Zhang: Methodology, Project administration, Supervision. Tong Wu: Conceptualization, Resources, Supervision, Funding acquisition, Writing – review & editing. Yiming Zhang: Resources, Methodology, Project administration, Supervision, Funding acquisition, Writing – review & editing.

Funding

This research was supported by the Chongqing Natural Science Foundation (CSTB2024NSCQ‐MSX0538), and the Textile Vision Basic Research Program of China National Textile and Apparel Council (J202404).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm71354‐sup‐0001‐SuppMat.docx.

Data Availability Statement

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