Zinc‐Containing Bioactive Glass Programs Macrophage Polarization through Extracellular Traps Regulation for Enhanced Diabetic Wound Healing

ABSTRACT

Impaired diabetic wound healing is driven by immune dysregulation and microenvironmental disruptions induced by hyperglycemia, leading to excessive inflammation and defective macrophage polarization. Although macrophage extracellular traps (METs) play critical roles in chronic inflammatory diseases, their involvement in shaping the diabetic wound immune microenvironment and impeding macrophage polarization remains insufficiently understood. Here, we present zinc‐containing bioactive glass (ZnBG), in which zinc incorporation confers immunomodulatory properties. In a type 2 diabetic full‐thickness skin excision model, ZnBG significantly mitigates MET‐associated oxidative stress and inflammation. Mechanistic investigations reveal that ZnBG effectively suppresses MET formation by reducing reactive oxygen species levels, inhibiting PAD4 activation, and blocking the NLRP3/caspase‐1/GSDMD signaling pathway. Consequently, ZnBG facilitates macrophage transition from the pro‐inflammatory M1 phenotype to the reparative M2 phenotype in diabetic wounds, thereby alleviating inflammation, enhancing neovascularization, and ultimately promoting diabetic wound healing. These findings provide an innovative therapeutic strategy that integrates ZnBG with targeted modulation of macrophage function for the treatment of diabetic wounds.

Zinc‐containing bioactive glass (ZnBG) promotes diabetic wound healing by regulating macrophage extracellular traps (METs). Specifically, ZnBG reduces oxidative stress and inhibits the PAD4 and NLRP3/caspase‐1/GSDMD signaling pathways, thereby suppressing MET formation. This suppression facilitates the polarization of macrophages from the pro‐inflammatory M1 phenotype to the reparative M2 phenotype, thus alleviating inflammation and enhancing neovascularization. These findings demonstrate that ZnBG provides an innovative immunomodulatory strategy for treating diabetic wounds.

1. Introduction

Diabetes mellitus encompasses a group of metabolic disorders characterized by abnormally high blood sugar levels resulting from insulin dysfunction [1]. As one of the most serious complications of diabetes, diabetic foot ulcers (DFUs) affect approximately 18.6 million individuals worldwide each year and are associated with substantially increased risks of amputation and mortality [2]. In acute cutaneous wounds, the healing process occurs across four overlapping stages: hemostasis, inflammation, proliferation, and remodeling [3, 4]. However, in diabetic wounds, the persistent overexpression of pro‐inflammatory cytokines, elevated levels of proteases and reactive oxygen species (ROS), and impaired cellular functions collectively disrupt the wound healing cascade [5, 6, 7]. Macrophages are key regulators during the inflammatory phase of wound healing. They coordinate inflammation and repair through phagocytosis and cytokine secretion and undergo a phenotypic transition from pro‐inflammatory (M1, early) to anti‐inflammatory (M2, late) post‐injury to accelerate the healing process [8]. However, in diabetes, chronic hyperglycemia disrupts this polarization, trapping macrophages in the M1 state and impairing tissue repair [9, 10, 11]. Therefore, the development of biomaterials that regulate macrophage polarization and restore the immune balance is a promising therapeutic approach for treating chronic diabetic wounds.

Bioglass 45S5 (BG), a silicate‐based bioactive material, releases therapeutic ions (e.g., Si, Ca, and P) into tissue fluids with exceptional bonding capacity to both bone and soft tissues [12, 13]. Extensive literature has documented that bioactive glass enhances vascularization, thus facilitating wound repair [14, 15, 16, 17]. However, its potential for immunomodulation, particularly in modulating macrophage polarization, remains unclear. In recent years, strategies incorporating metal ions such as silver, magnesium, and cobalt to enhance the functional properties of bioactive glasses have demonstrated significant potential in fields such as bone regeneration [18, 19, 20]. Zinc, an essential trace nutrient, plays a critical role in immune regulation and wound healing [21]. Zinc participates in regulating all four phases of wound healing, ensuring precision and efficiency [22], and its deficiency impairs cicatrization [23, 24]. Moreover, zinc suppresses ROS production [25, 26, 27], thereby mitigating oxidative stress‐induced tissue damage. The immunomodulatory capability of zinc has prompted the development of zinc‐based biomaterials for diabetic wound healing. For example, a ZnDB/HBC hydrogel modulates inflammation via sustained zinc release, doubling the wound closure rate [28], while a zinc‐based nanozyme hydrogel shifts macrophages from the M1 to the M2 phenotype and reduces inflammation [29]. Therefore, incorporating zinc into bioactive glass would enhance its immunomodulatory and therapeutic potential in diabetic wound repair.

Extracellular traps (ETs), first identified in neutrophils in 2004, are formed through ETosis—a cell death process involving chromatin decondensation and the release of DNA and proteins, resulting in web‐like structures that trap and eliminate pathogens [30]. Excessive neutrophil extracellular trap (NET) formation has been observed in diabetic wounds, which exacerbates chronic inflammation and implicates NET in delayed wound healing [31, 32]. Recent studies have revealed that macrophages are capable of generating similar structures known as macrophage extracellular traps (METs) [33, 34]. METs have been implicated in the pathogenesis of various inflammatory diseases, including rheumatoid arthritis [35], liver inflammation [36], acute kidney injury [37], atherosclerosis [38], and COPD [39]. However, their role in diabetic wounds remains unclear. Emerging evidence suggests that under high‐glucose conditions, METs may impede wound healing by sustaining inflammatory responses and disrupting macrophage polarization. A recent study conducted in a spinal cord injury model documented the presence of METs and their impact on microglial polarization [40]. Another study revealed that METs exacerbate diabetes by promoting intestinal inflammation and pro‐inflammatory T‐cell migration [41]. These findings suggest that METs may serve as critical mediators of persistent chronic inflammation and dysregulated macrophage polarization in diabetic wounds.

Gasdermin D (GSDMD), which is widely recognized as a key executor of pyroptosis, is cleaved by inflammatory caspase to release its N‐terminal fragment (GSDMD‐NT‐p30), which oligomerizes and forms pores in the plasma membrane. This process leads to the loss of membrane integrity, release of inflammatory cytokines, cell swelling, and ultimately programmed inflammatory cell death—a process critical for host defense against intracellular pathogens [42, 43, 44]. GSDMD facilitates intricate crosstalk among multiple cell death modalities, including pyroptosis, apoptosis, necroptosis, and NETosis, enabling the dynamic and context‐specific regulation of cellular demise [45, 46]. GSDMD can form pore‐forming proteins in the granule, mitochondrial, nuclear, and plasma membranes of neutrophils, ultimately leading to the production and release of NETs [47, 48]. Furthermore, MET formation was observed following the activation of GSDMD. GSDMD plays a pivotal role in MET formation by forming membrane pores that enable the release of extracellular DNA. Pharmacological inhibition of GSDMD using agents such as disulfiram or blockade of caspase‐1 with pan‐caspase inhibitors effectively suppresses MET generation, underscoring the critical function of GSDMD in linking pyroptosis to METosis [49]. BG has been shown to promote wound healing by suppressing GSDMD‐mediated endothelial cell pyroptosis via regulation of the connexin 43/ROS pathway [50]. Moreover, zinc inhibits caspase‐1, thereby hindering inflammasome activation [51]. These findings suggest that zinc‐containing bioglass (ZnBG) may modulate MET formation by targeting the macrophage GSDMD pathway.

In this study, we developed a ZnBG and explored its ability to promote diabetic wound healing, along with its underlying regulatory mechanisms. We systematically compared the performance of BG and ZnBG in wound repair, with a focus on evaluating the regulatory effects of ZnBG on macrophage polarization and the diabetic wound microenvironment. Additionally, we explored the presence of excessive METs in diabetic wounds and determined whether ZnBG regulates MET generation via the NLRP3/caspase‐1/GSDMD pathway. This study not only reveals the crucial role of METs in diabetic wound healing for the first time but also provides new strategies for developing immunomodulatory biomaterials, demonstrating substantial translational potential.

2. Methods and Materials

2.1. Preparation of Bioglass

BG and ZnBG were prepared using the classical melt‐cooling method. For BG, oxide raw materials (SiO₂, 46.1 mol%; Na₂O, 24.4 mol%; CaO, 26.9 mol%; and P₂O₅, 2.6 mol%) were mixed in proportions as previously reported [52]. According to the literature [53], two types of ZnBG with different zinc concentrations were synthesized based on the composition of BG. Briefly, by adding 1.5 and 5 mol% ZnO to replace the corresponding molar percentages of CaO, a mixture of silicon dioxide, calcium oxide, sodium oxide, and zinc oxide was prepared. The mixture of raw materials was sintered in a platinum crucible in an electric furnace at 1200°C and then melted for 1 h at 1350°C to achieve complete homogenization. After melting, the molten glass was rapidly quenched in deionized water to solidify, and the resulting frit was dried and ground into a fine powder.

2.2. Characterization of Bioglass

The surface ultrastructure of the BG was examined using scanning electron microscopy (SEM; Hitachi, Japan). The elemental composition of ZnBG powder was analyzed using energy dispersive spectrometry (EDS; EDX‐Octane Elect Super, Thermo Fisher Scientific). One gram of BG was dissolved in 5 mL of double‐distilled water, incubated at 37°C on a constant temperature shaker for one week, and the supernatant was collected after filtration through a 0.22 µm membrane. The calcium and silicon contents in the BG ion extract were determined using inductively coupled plasma atomic emission spectrometry (ICP‐ES; ICPOES7200, Thermo Fisher Scientific). The zinc content in the BG ion extract was measured using inductively coupled plasma mass spectrometry (ICP‐MS; NexION 300D, Perkin Elmer).

2.3. Animal Model

Male ICR mice aged 6–8 weeks were used to establish a full‐thickness skin excision model. C57BL/6J male mice aged 6–8 weeks were used to develop a type 2 diabetes model. The diabetes modeling protocol involved feeding a high‐fat diet containing 60% fat for 4–6 weeks, followed by a 12 h fast and continuous intraperitoneal injection of 1% streptozotocin (STZ, SolarBio; 50 mg/kg) for 5 days. Blood glucose levels were monitored, and type 2 diabetes was diagnosed when fasting blood glucose exceeded 11.1 mm, random blood glucose surpassed 16.7 mm, accompanied by weight loss and polyuria. Mice that met these criteria underwent subsequent surgery after one week of stabilization. All experimental animals were purchased from Zhejiang Charles River Laboratory Animal Technology Co., Ltd., and the protocol was approved by the Animal Ethics Committee of Wenzhou Medical University (Approval No.: wydw2025‐0363).

2.4. Establishing a Whole‐Thickness Skin Excision Model

The full‐thickness skin excision model was constructed as follows. After the intraperitoneal injection of 1% pentobarbital sodium according to body weight for anesthesia, the dorsal hair was removed using a depilator. Annular spacer pads (0.5 mm thick) with outer and inner diameters of 16 and 8 mm, respectively, were placed on both dorsal sides and secured with 5‐0 sutures to prevent wound contraction. Subsequently, a full‐thickness skin defect was created within the spacer pads using surgical scissors. ICR mice were randomly divided into wound, 45S5, 1.5% ZnBG, and 5% ZnBG groups. Full‐thickness wounds were separately covered with three different types of bioglass, each applied at a density of approximately 4 mg/cm². The diabetic mice were randomly divided into four groups: wound, 45S5, 5% ZnBG, and DNase I. The bioglass application method followed that used in conventional wound models, while the DNase I group received immediate intravenous administration of DNase I (dissolved in normal saline; MCE; 5 mg/kg, as previously described by Zhang et al. [40]) after wound preparation. After administration, the wounds were covered with a 3M transparent film and bandages to prevent drug leakage. All mice were housed in identical environments, and wound photographs were taken on days 0, 3, 6, 9, 12, and 15 post‐injury. The wound area was quantified using the ImageJ software in a blinded manner by an independent researcher, and the relative wound area percentage was calculated as follows:

$$\left(\mathrm{wound}\mathrm{area}\mathrm{on}\mathrm{a}\mathrm{given}\mathrm{day}/\mathrm{area}\mathrm{on}\mathrm{day}0\right)\times 100\%$$

2.5. Histological Assessment

Tissue samples were collected on days 7 and 15 post‐injury, fixed with 4% polyformaldehyde, and embedded in paraffin after ethanol gradient dehydration. Sections were cut to a thickness of 5 µm. Collagen deposition and granulation tissue formation were assessed by hematoxylin‐eosin (H&E) staining and Masson's trichrome staining. Immunofluorescence (IF) and immunohistochemical (IHC) analyses were conducted to evaluate ROS, inflammatory markers, MET‐related indicators, macrophage polarization proteins, and angiogenic biomarkers. Tissue sections were sequentially incubated with primary antibodies: ROS fluorescent probe (1:1000, Beyotime), TNF‐α (1:200, Proteintech), IL‐1β (1:200, Proteintech), citrullinated histone H3 (CitH3, 1:2000; Abcam), myeloperoxidase (MPO, 1:500; Proteintech), neutrophil elastase (NE, 1:100; ZenBio), CD68 (1:50, Abcam), CD86 (1:500, Proteintech), CD206 (1:500, Proteintech), NLRP3 (1:500, Proteintech), N‐GSDMD (1:100, HuaBio), CD31 (1:200, Proteintech), and hypoxia‐inducible factor‐1α (HIF‐1α, 1:200; Proteintech). After secondary antibody incubation, nuclei were stained with DAPI. IHC images were captured using a positive microscope (Leica DM750), IF images were obtained using a laser confocal microscope (Leica STELLARIS5), and semiquantitative analysis was performed using the ImageJ software.

2.6. Real‐Time Quantitative Polymerase Chain Reaction (RT‐qPCR)

On day 7 post‐injury, wound tissue samples were collected, and total RNA was extracted using TianGen TRNzol Universal Reagent. The RNA concentration was quantified via NanoDrop, and 1 µg of total RNA was reverse‐transcribed into cDNA using the Yeasen Reverse Transcript Kit. cDNA was diluted 10‐fold in sterile nuclease‐free water and combined with the reaction components as specified in the Yeasen SYBR Green Kit protocol. The mixture was loaded into an 8‐well plate for RT‐qPCR analysis using a Roche LightCycler 96 system. β‐actin served as the internal reference gene for normalization, and mRNA expression levels were calculated using the relative quantification method (2^−ΔΔCt). Primer sequences were designed using the NCBI Primer‐BLAST tool, with details specified in Table S1.

2.7. RNA Sequencing Analysis

On day 7 post‐injury, the wound tissues from the diabetic mice were snap‐frozen on dry ice and transported to MagiGene for RNA sequencing. Eukaryotic mRNA was isolated using oligo(dT)‐enriched magnetic beads and processed into strand‐specific cDNA libraries. High‐throughput sequencing was performed on an Illumina platform (San Diego, California, USA), with differentially expressed genes (DEGs) screened under the criteria of |log2(fold change)| > 0 and P < 0.05. Functional annotation of the DEGs was conducted using Gene Ontology (GO) enrichment analysis, and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis was used to elucidate potential regulatory mechanisms.

2.8. Western Blotting

On day 7 post‐injury, the wound tissue was collected and then lysed using a mixture of RIPA buffer, protease inhibitors, and PMSF (100:1:1) to extract total protein. The protein concentration was quantified using a NanoDrop spectrophotometer. Equal amounts of proteins were separated by SDS‐PAGE and subsequently transferred to PVDF membranes. Membranes were blocked with 5% non‐fat milk in TBST for 2 h, followed by overnight incubation at 4°C with the following primary antibodies: Arg‐1 (1:5000, Proteintech), iNOS (1:1000, Proteintech), MPO (1:2000, Proteintech), MMP‐9 (1:1000, Proteintech), NLRP3 (1:1000, Abcam), caspase‐1 (total and cleaved forms; 1:1000, Abmart), GSDMD (total and N‐terminal forms; 1:1000, Abmart), and PADI4 (1:500, ZenBio). After three TBST washes, the membranes were incubated with appropriate HRP‐conjugated secondary antibodies for 2 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence substrate and quantified via densitometric analysis using the ImageJ software.

2.9. Detection of Cell Death

The lactate dehydrogenase (LDH) Cytotoxicity Assay Kit (Beyotime) was used to detect the toxicity of ZnBG at different concentrations on HUVECs, 3T3 cells, and RAW264.7 cells. After soaking one gram of 5% ZnBG in 5 mL of the basal medium for 24 h, the suspension was collected and diluted to different concentrations. Cells were seeded in 96‐well plates and treated with different concentrations of the bioactive glass extracts for 12 h. The plate was then centrifuged at 400 g for 5 min, and 120 µL of the supernatant was transferred to another plate, followed by the addition of 60 µL of the LDH assay solution. The mixture was incubated in the dark at room temperature for 30 min with gentle shaking. The absorbance at 490 nm was measured using a microplate reader.

2.10. Extraction of Mouse Bone Marrow‐Derived Macrophages

Bone marrow‐derived macrophages (BMDMs) were isolated from C57BL/6J mice. Following euthanasia, the femurs and tibias were dissected, and the bone marrow was flushed with cold PBS using a 25‐gauge needle. The suspension was filtered through a 40 µm strainer, red blood cells (RBCs) were hemolyzed with RBC Lysis Buffer, and the cells were centrifuged to collect the pellet. The pellet was resuspended in a complete medium (MEM‐alpha + 10% FBS + 20 ng/mL M‐CSF) and seeded at a density of 1 × 10⁸ cells/mL. Cells were cultured at 37°C in a 5% CO₂ incubator, with M‐CSF‐containing medium replenished on day 3, and differentiation was allowed to proceed for 6–7 days until adherent, spindle‐shaped macrophages formed. To verify that we extracted macrophages from the induced differentiation, fluorescence staining was performed using the macrophage surface marker CD68.

2.11. Induction and Clearance of METs in BMDMs

Following 7 days of differentiation in α‐MEM medium, mature BMDMs were cultured in media containing varying glucose concentrations (5.5, 15, 25, and 35 mm) for 24 h. MET formation was assessed by IF co‐staining for CitH3 and MPO. To evaluate the MET‐clearing effects of ZnBG and DNase I, high‐glucose‐treated cells were incubated with BG extract and DNase I solution (Thermo Scientific, 10 U/mL, as previously described by Weng et al. [35]) for 12 h, followed by METs quantification.

2.12. Laser Doppler Imaging

A laser Doppler imaging system (Perimed PSI HR model) was used to evaluate the wound perfusion status on day 15 post‐injury. Laser speckle contrast imaging was used for the visual observation of tissue blood flow perfusion in each wound. The near‐infrared laser imaging system scanned the wound surface at fixed distances and angles, thereby obtaining blood flow perfusion images of mouse skin tissues.

2.13. Statistical Analysis

Statistical analyses were performed using GraphPad Prism (version 10.0; GraphPad Software, La Jolla, California, USA). Quantitative data in this study are presented as the mean ± standard deviation (SD). All experiments were performed with at least three biological replicates. Prior to analysis, quantitative data were normalized to the corresponding internal control group within each experiment, with no other mathematical transformation applied. For multiple groups with one variable, one‐way analysis of variance (ANOVA) followed by post hoc Tukey's test was used to determine the differences. For multiple groups with two variables, comparison was performed using two‐way ANOVA followed by post hoc Tukey's test. P‐values were considered statistically significant as follows: ns: no significance, ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001.

3. Results

3.1. Characterization of ZnBG

A schematic of BG synthesis is shown in Figure 1A. The morphology, elemental composition, and ion release properties of ZnBG and 45S5 were systematically characterized. The SEM images revealed that all three bioactive glasses exhibited irregular particles (Figure 1B). Their particle size distributions were normal, with average particle sizes ranging from 7 to 9 µm (Figure S1). These results indicate that zinc incorporation (at doping levels of 1.5% or 5%) did not induce structural changes in the borosilicate bioactive glasses. EDS analysis (Figure 1C) detected the characteristic peaks of O, Na, Si, P, Ca, and Zn, confirming the successful incorporation of zinc. Previous studies suggested that zinc doping can inhibit ion release from bioactive glasses [53]. Consistently, our ion release analysis (Figure 1D) revealed that both the 1.5% and 5% Zn‐doped borosilicate glasses exhibited reduced dissolution rates of silicon and calcium compared to 45S5, which is attributable to their zinc content. Nevertheless, the release rates remained within the same order of magnitude, which is consistent with previous reports. Notably, controlled zinc ion release was achieved, with 5% ZnBG showing significantly greater zinc ion release than 1.5% ZnBG, establishing a positive correlation between the zinc doping concentration and the release amount. These results indicate the successful synthesis of ZnBG.

FIGURE 1.

ZnBG accelerates wound healing in ICR mice. (A) Schematic diagram of bioactive glass synthesis. (B) SEM images of bioactive glasses. (C) EDS analysis of ZnBG. (D) The concentration of ions in the BG extract. (E) Representative images on days 0, 3, 6, 9, 12, and 15. (F) Quantitative analysis of wound closure rates, n = 6, ^** P < 0.01, ^**** P < 0.0001. (G) H&E and Masson's staining of wound tissue on days 7 and 15, Scale bar: 500 µm, 50 µm. Data are presented as mean ± SD, and statistical significance was analyzed using two‐way ANOVA followed by Tukey's multiple comparison test.

3.2. ZnBG Accelerates Wound Healing in ICR Mice

A full‐thickness skin defect model was established on the dorsal side of normal ICR mice to evaluate the wound healing efficacy of ZnBG and to compare the therapeutic outcomes of two distinct ZnBG concentrations with 45S5. The wound healing rate (Figure 1F) demonstrated that all three types of bioactive glass exhibited significant pro‐healing properties on days 6, 9, 12, and 15. On day 15, the average remaining wound areas were 42.69%, 23.36%, 18.97%, and 8.77% in the wound, 45S5, 1.5% ZnBG, and 5% ZnBG groups, respectively. Notably, the 5% ZnBG group showed superior healing efficacy compared to the 45S5 and 1.5% ZnBG groups, whereas only a slight improvement was observed in the 1.5% ZnBG group relative to the 45S5 group. This clearly indicates that 5% ZnBG possesses enhanced wound‐healing capability relative to conventional BG. Consequently, 5% ZnBG was selected for subsequent experiments. Collagen deposition and granulation tissue formation were evaluated using H&E and Masson's trichrome staining (Figure 1G). The results showed that 5% ZnBG promoted more robust granulation tissue formation on days 7 and 15. Masson's trichrome staining revealed that compared to the 45S5 group, wounds treated with 5% ZnBG exhibited thicker and more organized collagen fibers on days 7 and 15 post‐injury, highlighting its potential to enhance cutaneous regeneration. Additionally, the hemocompatibility results demonstrated that all three bioactive glasses exhibited hemolysis rates below 3% (Figure S2), indicating their low biotoxicity and supporting their potential for in vivo applications.

3.3. ZnBG Accelerates Diabetic Wound Healing

Marked therapeutic efficacy of 5% ZnBG was observed in normal wounds, and subsequently, it was applied to more complex diabetic wounds to evaluate its efficacy. Type 2 diabetes was induced in C57/B6J mice using a high‐fat, high‐sugar diet combined with STZ injections, after which a full‐thickness wound model was established (Figure 2A). Both 45S5 and 5% ZnBG accelerated diabetic wound healing. Specifically, 5% ZnBG demonstrated a significantly higher healing rate than 45S5 on days 3, 6, 9, 12, and 15 (Figure 2C). On day 15, the residual wound areas were 26.93%, 17.65%, and 9.26% in the wound, 45S5, and 5% ZnBG groups, respectively. These results indicated that 5% ZnBG significantly promoted wound healing in diabetic mice compared to 45S5. Subsequent histological analyses using H&E and Masson's trichrome staining confirmed that 5% ZnBG enhanced granulation tissue formation and collagen deposition at the wound site (Figure 2D). Given the critical role of vascularization in wound healing, we conducted a comprehensive evaluation of angiogenesis in the wounds of diabetic mice. Laser Doppler imaging on day 15 post‐injury was used to evaluate blood flow in the wound area, with blue, green, and red indicating low, moderate, and high perfusion, respectively. The 5% ZnBG‐treated group exhibited a predominantly red coloration (Figure 2E), suggesting a robust blood supply to the wound. In addition, high‐resolution imaging was performed to capture the vascular distribution in the wound area. Quantitative analysis revealed that treatment with 5% ZnBG significantly increased the number of blood vessels in the wound bed (Figure 2G). To further validate angiogenesis, IF and IHC staining were conducted to measure the expression levels of vascular endothelial markers (CD31) and HIF‐1α. The results demonstrated significantly elevated CD31 and HIF‐1α expression levels in the 5% ZnBG group compared with the other groups (Figure 2F). These findings highlight the ability of 5% ZnBG to enhance diabetic wound healing by promoting angiogenesis.

FIGURE 2.

ZnBG accelerates diabetic wound healing. (A) Schematic diagram of the experimental design for the wound model in type 2 diabetic mice. (B) Representative images on days 0, 3, 6, 9, 12, and 15. (C) Quantitative analysis of wound closure rates, n = 6, ^* P < 0.05, ^**** P < 0.0001. (D) H&E and Masson's staining of wound tissue on days 7 and 15, Scale bar: 500, 50 µm. (E) Laser Doppler perfusion imaging and high‐resolution images of vasculature in the wound area of each group on day 15. (F) Immunofluorescence staining of CD31 (red) and immunohistochemical staining of HIF‐1α in each group at 15 days post‐injury, Scale bar: 40, 25 µm. (G) Doppler perfusion quantitative analysis and statistical analysis of vessel density, n = 3, ^* P < 0.05, ^*** P < 0.001. (H) CD31 immunofluorescence and HIF‐1α immunohistochemical quantitative analysis, n = 6, ^* P < 0.05, ^** P < 0.01^,**** P < 0.0001. Data are presented as mean ± SD, and statistical significance was analyzed using two‐way ANOVA (C) or one‐way ANOVA (G, H) followed by Tukey's multiple comparison test.

3.4. ZnBG Regulates Inflammation and Macrophage Polarization in Diabetic Wounds

To elucidate the molecular mechanisms by which 5% ZnBG accelerates wound repair, RNA sequencing analysis was performed on wound tissues collected on day 7. Heatmap visualization of DEGs revealed that treatment with 5% ZnBG significantly altered the gene expression profile of wound tissues (Figure 3A). GO enrichment analysis (Figure 3B) suggested that 5% ZnBG modulated immune and inflammatory responses, indicating its potential to remodel the diabetic wound microenvironment. KEGG pathway analysis of the DEGs revealed enrichment in the NOD‐like receptor signaling pathway (Figure 3C). These results suggest that 5% ZnBG participates in and regulates inflammatory and immune remodeling in diabetic wounds. The effect of 5% ZnBG on the wound inflammatory microenvironment was validated using IF and RT‐qPCR. IF analysis revealed that 5% ZnBG treatment downregulated the expression of TNF‐α and IL‐1β (Figure 3D). Meanwhile, RT‐qPCR (Figure 3F) confirmed that 5% ZnBG treatment reduced pro‐inflammatory cytokines (TNF‐α and IL‐1β) while upregulating anti‐inflammatory factors (TGF‐β and IL‐10). In diabetic wounds, impaired macrophage polarization causes excessive M1 macrophage infiltration, resulting in persistent local inflammation and impaired healing. To investigate the polarization state of the macrophages, IF staining was performed using CD86 (an M1 macrophage marker) and CD206 (an M2 macrophage marker). Treatment with 5% ZnBG decreased CD86 expression and increased CD206 expression (Figure 3G). Western blotting was performed to examine the protein levels of iNOS (secreted by M1 macrophages) and Arg‐1 (secreted by M2 macrophages). The 5% ZnBG group showed decreased iNOS protein expression and increased Arg‐1 protein expression (Figure 3H). Collectively, these results suggested that 5% ZnBG plays a pivotal role in regulating the levels of inflammatory factors and macrophage functional remodeling in diabetic wounds.

FIGURE 3.

ZnBG regulates inflammation and macrophage polarization in diabetic wounds. (A) Heatmap of significantly upregulated and downregulated genes in the wound group versus the ZnBG group in diabetic wounds. (B) Enrichment analysis and (C) KEGG pathway classification of identified differentially expressed genes. (D) Immunofluorescence analysis of inflammatory factor expression in diabetic wound areas on day 7 post‐injury, IL‐1β: green; TNF‐α: red; DAPI: blue, Scale bar: 40 µm. (E) Immunofluorescence quantitative analysis of IL‐1β and TNF‐α, n = 6, ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001. (F) Relative expression of inflammatory genes (IL‐1β, TNF‐α) and anti‐inflammatory genes (IL‐10, TGF‐β) in different treatment groups, n = 3, ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001. (G) Immunofluorescence analysis of macrophage polarization markers in diabetic wound areas on day 7 post‐injury, CD86: red; CD206: green; DAPI: blue, Scale bar: 40 µm. (H) Western blot analysis of macrophage polarization‐associated secretory protein expression in wound tissues. (I) Fluorescence quantification analysis of CD86 and CD206, n = 6, ^* P < 0.05, ^** P < 0.01, ^**** P < 0.0001. (J) Quantitative analysis of iNOS and Arg‐1 protein expression, n = 3, ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001. Data are presented as mean ± SD, and statistical significance was analyzed using one‐way ANOVA followed by Tukey's multiple comparison test.

3.5. ZnBG Regulates the Production of METs in Diabetic Wounds

Having elucidated the role of ZnBG in macrophage reprogramming, we further investigated the underlying mechanisms. Recently, Wang et al. demonstrated that METs directly regulate macrophage polarization in spinal cord injury [40]. Therefore, this study focused on the formation of METs in diabetic wounds and the potential role of ZnBG in this process. First, IF was used to investigate the formation of METs in diabetic wounds. Tissue sections were stained with CD68 to label macrophages and subsequently co‐stained with either CitH3 or MPO to label METs. The results revealed substantial MET infiltration in diabetic wounds, whereas 5% ZnBG significantly downregulated CitH3 and MPO expression (Figure 4A,B, respectively), indicating its regulatory role in MET formation. Furthermore, IHC analysis of NE in diabetic wounds revealed a marked reduction in NE protein levels following treatment with 5% ZnBG (Figure 4D). Consistent with the IF findings, western blotting revealed that 5% ZnBG suppressed MET formation in wound tissues, as evidenced by reduced MPO and MMP‐9 levels (Figure 4F). Collectively, these results demonstrate that 5% ZnBG effectively mitigated excessive MET formation under diabetic conditions.

FIGURE 4.

ZnBG regulates the production of METs in diabetic wounds. (A) Immunofluorescence analysis of CitH3 expression on macrophages, CitH3: red; CD68: green; DAPI: blue, Scale bar: 40 µm. (B) Immunofluorescence analysis of MPO expression on macrophages, CD68: red; MPO: green; DAPI: blue, Scale bar: 40 µm. (C) Immunofluorescence quantitative analysis of CitH3 and MPO, n = 6, ^* P < 0.05, ^*** P < 0.001, ^**** P < 0.0001. (D, E) Immunohistochemical images and quantification of NE in wounds of different groups, n = 6, ^* P < 0.05, ^**** P < 0.0001, Scale bar: 25 µm. (F, G) Western blot and quantification of MPO and MMP‐9 in different groups, n = 3, ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001. Data are presented as mean ± SD, and statistical significance was analyzed using one‐way ANOVA followed by Tukey's multiple comparison test.

3.6. Inhibiting METs can Regulate Macrophage Polarization and Accelerate Wound Healing

To elucidate the relationship between METs and diabetic wounds, as well as their potential role in delaying macrophage phenotype switching, we used DNase I, a recognized MET scavenger, for validation. Full‐thickness wounds were established in type 2 diabetic mice, and healing outcomes were compared between groups receiving local administration of 5% ZnBG and DNase I via tail vein injection. Compared with the untreated wound group, both the DNase I and 5% ZnBG groups exhibited accelerated wound healing (Figure 5A). On day 15, the residual wound areas were 27.26%, 13.45%, and 7.98% in the wound, DNase I, and 5% ZnBG groups, respectively (Figure 5C). These results suggest that the clearance of METs promotes diabetic wound healing and support the hypothesis that ZnBG facilitates diabetic wound repair by modulating METs. Subsequently, IF staining confirmed that in both the DNase I and 5% ZnBG groups, METs were significantly eliminated, and the transformation of macrophages from the M1 to M2 phenotype was promoted (Figure 5B). To validate these results in vitro, we first confirmed that a 1/1000 concentration of 5% ZnBG extract had no effect on the proliferation of HUVECs, 3T3, and RAW264.7 cells, as determined by the LDH assay (Figure S3). Furthermore, we assessed the biosafety of 5% ZnBG using live/dead staining, which also revealed that the 1/1000 concentration of ZnBG extract had no impact on RAW264.7 cell viability (Figure S4). Therefore, 1/1000 was selected as the optimal 5% ZnBG concentration for the treatment. BMDMs were isolated from mice and induced to mature using M‐CSF. After seven days, CD68 staining confirmed successful differentiation of the primary macrophages (Figure S5). When cultured under varying glucose concentrations (5.5, 15, 25, and 35 mm) to induce MET formation, the 35 mm glucose group (representing high‐glucose conditions) exhibited extensive MET production characterized by web‐like structures (Figure 5E). After 24 h of high‐glucose induction, the cells were treated with 5% ZnBG extract to evaluate their ability to eliminate pre‐formed METs. IF analysis revealed that the 5% ZnBG group exhibited a significant reduction in METs, with an efficacy comparable to that of the MET scavenger control (Figure 5G). Furthermore, IF staining for macrophage polarization markers demonstrated that MET clearance promoted a phenotypic transition from pro‐inflammatory M1 (CD86) to anti‐inflammatory M2 macrophages (CD206) (Figure 5I). These in vitro findings are consistent with our animal experimental observations, reinforcing the role of ZnBG in modulating macrophage polarization through MET regulation.

FIGURE 5.

Clearance of METs ameliorates macrophage polarization. (A) Representative images on days 0, 3, 6, 9, 12, and 15. (B) Immunofluorescence Analysis of METs Formation and Macrophage Polarization Status, CitH3: red; MPO: green; CD86: red; CD206, green; DAPI: blue, Scale bar: 40 µm. (C) Quantitative analysis of wound closure rates, n = 6, ^** P < 0.01, ^**** P < 0.0001. (D) Immunofluorescence quantitative analysis of CitH3, MPO, CD86 and CD206, n = 6, ns: no significance, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001. (E, F) Immunofluorescence analysis and quantitative analysis of METs formation in BMDMs under different glucose concentrations, CitH3: red; MPO: green; DAPI: blue, Scale bar: 40 µm, n = 6, ^* P < 0.05, ^** P < 0.01, ^**** P < 0.0001. (G, H) Immunofluorescence analysis and quantitative analysis of METs formation in BMDMs across different treatment groups, CitH3: red; MPO: green; DAPI: blue, Scale bar: 40 µm, n = 6, ^* P < 0.05, ^** P < 0.01, ^**** P < 0.0001. (I, J) Immunofluorescence and quantitative analyses of the polarization status in BMDMs across different treatment groups, CD86: red; CD206: green; DAPI: blue. Scale bar: 40 µm. n = 6. ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001. Data are presented as mean ± SD, and statistical significance was analyzed using two‐way ANOVA (C) or one‐way ANOVA (D, F, H, J) followed by Tukey's multiple comparison test.

3.7. ZnBG Regulates METs via NLRP3/caspase1/GSDMD Signal

Having established that ZnBG promotes wound healing by modulating macrophage polarization through the clearance of METs, we investigated the specific mechanisms by which ZnBG regulates METs. The chronic hyperglycemic environment in diabetic wounds leads to excessive ROS production, which drives chromosome decondensation and histone citrullination, which are the core mechanisms of MET formation [54]. The ROS levels in diabetic wounds were examined using dihydroethidium (DHE) as a fluorescent probe. Upon entering the cells, DHE is oxidized by superoxide anions to form ethidium cations, which emit red fluorescence proportional to intracellular oxidative stress. DHE staining revealed that 5% ZnBG significantly reduced the DHE fluorescence intensity (Figure 6A), indicating its ability to decrease ROS levels. Lee et al. reported that MET formation is associated with GSDMD activation, wherein N‐GSDMD forms membrane pores that facilitate the release of MET components [49]. Therefore, we used IF to examine whether 5% ZnBG reduced the expression of N‐GSDMD and its upstream NLRP3 inflammasome in macrophages. Treatment with 5% ZnBG significantly reduced the levels of NLRP3 and N‐GSDMD in macrophages (Figure 6C,D). Western blotting validated this finding, revealing a marked downregulation of NLRP3, cleaved‐caspase‐1, and N‐GSDMD in the 5% ZnBG treatment group (Figure 6G). Given that ROS overproduction activates peptidyl arginine deiminase 4 (PAD4), which drives MET formation via chromatin decondensation and histone citrullination [55, 56, 57], we examined PAD4 expression via western blotting. As expected, 5% ZnBG significantly suppressed PAD4 protein levels (Figure 6G). These findings collectively elucidated the mechanism by which ZnBG regulates MET formation by reducing ROS levels, subsequently inhibiting the NLRP3/caspase‐1/GSDMD pathway and PAD4 activation, and ultimately suppressing MET generation.

FIGURE 6.

ZnBG regulates METs via NLRP3/caspase1/GSDMD signal promotes diabetic wound healing. (A) Representative DHE staining images of wounds in different groups and (B) Quantitative analysis of DHE fluorescence intensity, n = 6, ^** P < 0.01, ^**** P < 0.0001, Scale bar: 500, 200 µm. (C) Immunofluorescence analysis of NLRP3 expression on macrophages, CD68: red; NLRP3: green; DAPI: blue, Scale bar: 40 µm. (D) Immunofluorescence analysis of N‐GSDMD expression on macrophages, N‐GSDMD: red; CD68: green; DAPI: blue, Scale bar: 40 µm. (E) Immunofluorescence quantitative analysis of NLRP3, n = 6, ^* P < 0.05, ^**** P < 0.0001. (F) Immunofluorescence quantitative analysis of N‐GSDMD, n = 6, ^* P < 0.05, ^** P < 0.01, ^**** P < 0.0001. (G, H) Western blot analysis of key NLRP3‐GSDMD signaling pathway protein expression in different groups and their quantitative analysis, n = 3, ns: no significant, ^* P < 0.05, ^** P < 0.01, ^*** P < 0.001, ^**** P < 0.0001. Data are presented as mean ± SD, and statistical significance was analyzed using one‐way ANOVA followed by Tukey's multiple comparison test.

4. Discussion

The core mechanism underlying the impaired healing of diabetic wounds is immune dysregulation caused by a chronic inflammatory microenvironment with abnormal macrophage polarization (persistent M1 polarization and impaired M2 transition). Bioactive glass enhances vascularization, suppresses inflammation, promotes cell proliferation and migration, and ultimately accelerates wound healing. However, the specific mechanisms through which it regulates the disrupted immune microenvironment in diabetic wounds remain poorly understood. Zinc ions possess unique immunomodulatory properties and can mitigate oxidative stress damage by suppressing ROS and improving wound repair by reprogramming macrophages from the pro‐inflammatory M1 phenotype to the anti‐inflammatory M2 phenotype. Therefore, it is reasonable to hypothesize that incorporating zinc into BG could enhance its ability to regulate immunity and inflammation, thereby fostering better wound healing. METs have been implicated in many chronic inflammatory diseases and are associated with shifts in macrophage polarization. Recent studies showed that MET formation is closely associated with GSDMD‐dependent pyroptosis. However, whether METs exist in diabetic wounds and how they influence the immune microenvironment of diabetic wounds remains unknown. In this study, two types of ZnBG were developed, and 5% ZnBG exhibited superior wound healing effects compared with conventional BG. Mechanistically, 5% ZnBG effectively suppressed MET formation by reducing ROS levels, inhibiting PAD4 enzyme activation, and blocking the NLRP3/caspase1/GSDMD signaling pathway. This intervention promoted a phenotypic shift from persistent M1 to M2 macrophages in diabetic wounds, ultimately achieving a coordinated improvement in inflammation resolution, angiogenesis, and wound repair (Figure 7).

FIGURE 7.

Schematic illustration of the mechanism by which ZnBG promotes diabetic wound healing. By reducing ROS levels, ZnBG effectively suppresses the overproduction of METs, thereby reprogramming macrophage polarization. This process ameliorates the inflammatory microenvironment in diabetic wounds, enhances vascularization, and ultimately accelerates wound healing. Created in BioRender. Wen, Z. (2026) https://BioRender.com/gj1pwef.

Bioactive glasses enhance wound healing primarily by releasing therapeutic ions [58]. Research on the mechanism by which bioactive glasses promote wound healing has focused on the augmentation of endothelial cell and fibroblast functions by certain therapeutic ions: silicon (Si⁴⁺) stimulates angiogenesis by upregulating pro‐angiogenic factors and improves skin strength via collagen and glycosaminoglycan synthesis [59, 60, 61, 62]; calcium (Ca²⁺) promotes fibroblast and keratinocyte proliferation, collagen production, ECM deposition, and accelerates granulation tissue formation and re‐epithelialization [63, 64, 65]; and phosphate ions (PO₄ ³⁻) contribute structurally to the ECM, support cellular signaling and metabolism [66], and stimulate angiogenesis through mediators such as bFGF and MMP‐2 [67, 68]. Bioactive glasses facilitate the polarization of macrophages from the M1 to the M2 phenotype to reduce excessive inflammation and establish a pro‐healing microenvironment [69]. Compared to conventional bioactive glasses, ZnBG releases Zn²⁺ ions, which effectively suppress inflammatory responses and regulate macrophage polarization through the NF‐κB pathway, thereby promoting bone tissue regeneration [70]. These findings suggest that ZnBG is a more suitable biomaterial for wound healing, particularly for chronic non‐healing wounds, compared to 45S5.

Notably, high‐concentration zinc doping significantly retards glass dissolution and almost completely suppresses the release of calcium and silicon ions. This is because zinc ions integrate into the silicate network as structural intermediates, effectively “locking” themselves in place, which markedly stabilizes the glass, slows its degradation, and reduces ion exchange with the surrounding medium, thereby further inhibiting the release of ions such as calcium, sodium, and phosphate [71, 72]. As a result, it impedes apatite formation and may completely prevent mineralization. In contrast, low zinc concentrations barely affect glass degradation, maintaining the release rates of therapeutic ions (Ca²⁺, PO₄ ³⁻, Si²⁺) of BG, slightly limiting apatite formation, and still harnessing the therapeutic advantages of zinc [53]. Therefore, two ZnBG with low Zn concentrations (1.5% and 5%) were designed to evaluate their structural characteristics and ion release behavior, while verifying whether the incorporation of additional zinc ions could amplify the pro‐healing capacity of bioactive glass.

This study revealed that the ZnBG microstructure was indistinguishable from that of 45S5, with zinc doping observed to reduce the release of calcium and silicon ions in a concentration‐dependent manner, which is consistent with previous literature (Figure 1). Subsequently, in acute wounds, 5% ZnBG significantly enhanced wound healing compared to 45S5, whereas 1.5% ZnBG provided a limited improvement in healing compared to 45S5 (Figure 1). These findings suggest that within a certain range, increasing the zinc doping concentration enhanced the wound‐healing performance of bioactive glasses. Furthermore, in diabetic mice, 5% ZnBG demonstrated superior wound healing compared to 45S5 (Figure 2). These observations demonstrate that ZnBG may possess greater therapeutic potential than 45S5, both in normal acute wounds and in chronic hard‐to‐heal wounds such as diabetic wounds.

A well‐balanced immune microenvironment and appropriate inflammatory response are generally conducive to wound healing [73, 74]. In diabetic wounds, excessive inflammatory accumulation and elevated ROS generation directly inhibit endothelial cell function [75], disrupt the balance of growth factors [76], and lead to excessive MMP activity, which degrades the extracellular matrix serving as a scaffold for angiogenesis [77], ultimately resulting in the formation of dysfunctional blood vessels. Therefore, modulation of the immune microenvironment is crucial. Although there was a potential reduction in the release of certain pro‐angiogenic therapeutic ions in vivo, the 5% ZnBG group exhibited increased neovascularization at the wound site compared with the 45S5 group (Figure 2). In addition to the direct pro‐angiogenic effect of zinc ions, this phenomenon may be attributed to the zinc‐induced remodeling of the wound microenvironment, which restores the function of endothelial cells. Subsequent evaluation of local wound inflammation levels confirmed that 5% ZnBG had better immunomodulatory ability than 45S5, as evidenced by reduced levels of inflammatory factors and improved macrophage polarization status (Figure 3). Furthermore, RNA sequencing confirmed that ZnBG primarily participates in regulating immune responses and inflammatory reactions. These results underscore the critical role of immune microenvironment remodeling in wound repair and demonstrate that the incorporation of zinc endows bioglass with an enhanced capacity to modulate the wound microenvironment.

Macrophages are a crucial component of the human immune system and play a central role in the immune response by combating infections and regulating tissue inflammation. Under various stimulating conditions, macrophages generate ETs, which are intricate web‐like structures composed of proteins and DNA. These structures not only capture and eliminate microorganisms but also contribute significantly to tissue damage, inflammatory processes, and the development of autoimmune diseases [78]. Importantly, METs have been identified as key pathogenic factors involved in the development of type 1 diabetes [41]. Therefore, it is reasonable to speculate that METs are involved in disrupting the immune microenvironment of diabetic wounds, thereby impairing the healing process. In this study, excessive METs were confirmed to be present in diabetic wounds, and ZnBG significantly downregulated MET formation (Figure 4). Recent studies have explored the relationship between METs and macrophage polarization. It has been reported that liraglutide promotes M2 polarization and reduces METs via the STAT3/6 pathway, while the MET inhibitor Cl‐amidine also affects polarization status, indicating a mutual regulatory relationship [79]. Additionally, in a calcium oxalate kidney injury model, METs and M1 polarization were shown to form a positive feedback loop, wherein M1 macrophages release METs, which further reinforce the M1 phenotype through the NF‐κB signal pathway [80]. In spinal cord injury, METs sustain M1 macrophage polarization through the LL37/P2×7R/NF‐κB pathway, exacerbating tissue damage [40].

Although emerging evidence suggests bidirectional interactions between METs and polarization states, the temporal hierarchy and mechanistic causality remain incompletely defined. In this study, definitive evidence was provided that METs function as primary drivers of polarization dynamics in diabetic wounds. DNase I, a widely recognized MET scavenger [36, 40, 81, 82], was used to validate the relationship between METs and polarization. In the diabetic wound model, DNase I was administered on day 0 post‐injury, resulting in reduced MET levels by day 7, accompanied by a decrease in M1 macrophages and an increase in M2 macrophages. More importantly, we validated this regulatory mechanism in a cellular model, in which the induction and subsequent clearing of METs led to significant shifts in macrophage polarization. In addition, the accelerated wound healing phenotype upon MET clearance functionally supported METs as critical upstream signals for inflammatory factors. Collectively, these multilayered findings establish METs as upstream regulators of macrophage polarization. While this study established METs as key regulators of macrophage polarization, we cannot rule out the potential feedback modulation of METs by polarized subsets, as suggested by prior reports of M1‐driven MET reinforcement loops. Future investigations employing time‐resolved co‐cultures to track secondary MET induction or polarization‐locking approaches to assess MET‐reprogramming capacity could clarify whether this interplay forms a self‐amplifying circuit or a context‐dependent hierarchy.

Chromosome decondensation and histone citrullination, driven by NADPH oxidase‐derived ROS, are the core mechanisms of MET formation [54]. ROS stimulate the release of MPO and NE, which are critical mediators of chromatin decondensation [83]. Meanwhile, ROS activates PAD4 and promotes its translocation into the nucleus, where PAD4 plays a central role in histone citrullination, a critical step in chromatin decondensation during MET formation. PAD4 destabilizes histones by mediating citrullination, thereby facilitating chromatin expansion and subsequent DNA release [54, 84]. In addition, ROS activates the NLRP3 inflammasome [85, 86, 87, 88], subsequently triggering caspase‐1‐mediated GSDMD activation and pore formation via its N‐terminal domain, which localizes to granule, nuclear, and plasma membranes to facilitate METs release [47, 48]. DHE staining revealed that 5% ZnBG exhibited superior ROS‐scavenging ability compared with 45S5 (Figure 6). Western blotting and IF demonstrated that 5% ZnBG led to a lower expression of MPO and NE (Figure 4), as well as downregulation of the NLRP3–caspase‐1–GSDMD axis (Figure 6). These results collectively account for the markedly reduced MET formation observed in the 5% ZnBG group. This phenomenon may be attributed, in part, to the direct regulatory role of zinc on ROS [25, 26, 27] and to the potential of zinc to bind directly to the His‐Cys‐Cys triad active site of caspase‐1, thereby inhibiting its protease activity and consequently hindering the cleavage of substrates such as GSDMD [51]. The formation of METs is a complex, multifaceted process, with the understanding of its underlying signaling pathways still in its nascent stages. Although this study demonstrated that 5% ZnBG significantly influenced the formation of METs, the precise mechanisms require further rigorous validation using transgenic animal models.

5. Conclusion

The successful development of ZnBG provides a favorable microenvironment for diabetic wound healing. By reducing ROS levels, inhibiting the PAD4 enzyme, and suppressing the NLRP3/caspase‐1/GSDMD signaling pathway, ZnBG effectively decreased the formation of METs. This facilitates a phenotypic shift from persistent M1 to M2 macrophages in diabetic wounds, alleviates inflammatory responses, accelerates wound vascularization, and promotes diabetic wound repair.

Author Contributions

RuiYang Sun: writing – original draft, visualization, and methodology. XueBo Wei: conceptualization, formal analysis, and project administration. ZhuoYang Song: visualization and investigation. JunJun Luo: software and visualization. JingYi Ye: validation and methodology. YinNan Zhang: investigation and data curation. LiFei Zhu: validation and data curation. Jie Gao: funding acquisition, conceptualization, and formal analysis. Hongyu Zhang: visualization and methodology. Hong Zhu: funding acquisition, supervision, visualization, and resources. Xin Wang: funding acquisition, supervision, and visualization. Ke Xu: funding acquisition, writing – review & editing, conceptualization, investigation, and supervision.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm70886‐sup‐0001‐SuppMat.docx.

Data Availability Statement

Additional data collected during this study are available from the corresponding author upon reasonable request.