SCAP‐Exo‐Integrated CA/HACC Hydrogel Promotes Diabetic Wound Repair via miR‐122‐Driven Macrophage Phenotypic Transition

ABSTRACT

Chronic diabetic wounds are characterized by persistent inflammation and a disrupted transition from the inflammatory to the reparative phase, largely driven by macrophage dysregulation and metabolic imbalance within the wound microenvironment. Here, we developed an exosome‐integrated hydrogel system based on citrate‐crosslinked hydroxypropyltrimethyl ammonium chloride chitosan (HACC), incorporating exosomes derived from stem cells of the apical papilla (SCAP‐Exo) to modulate immune responses and promote diabetic wound repair. The SCAP‐Exo–loaded HACC hydrogel exhibited favorable physicochemical properties and enabled sustained local exosome release. In vitro studies demonstrated efficient uptake of SCAP‐Exo by macrophages, leading to a pronounced shift toward an anti‐inflammatory, pro‐reparative phenotype. MicroRNA profiling and functional validation identified miR‐122‐5p as a key bioactive component enriched in SCAP‐Exo, mediating macrophage reprogramming through the regulation of inflammatory signaling and metabolism‐associated pathways. Consequently, the remodeled immune microenvironment enhanced fibroblast proliferation and migration. In a diabetic wound model, treatment with the SCAP‐Exo–functionalized HACC hydrogel significantly accelerated wound closure, improved granulation tissue formation, and promoted collagen deposition. Collectively, this study demonstrates that SCAP‐Exo–based HACC hydrogel therapy facilitates diabetic wound healing through miRNA‐driven immunometabolic modulation, highlighting a promising strategy for the treatment of chronic wounds.

A SCAP‐derived exosome–integrated HACC‐based hydrogel provides sustained delivery of bioactive miR‐122‐5p to diabetic wounds. The system reprograms macrophages toward a pro‐regenerative phenotype, enhances fibroblast migration and proliferation, and promotes angiogenesis, thereby orchestrating inflammation resolution and tissue regeneration within the impaired diabetic wound microenvironment.

1. Introduction

According to recent reports, in 2021, there were 529 million people with diabetes worldwide, and it is projected that by 2050, this number will exceed 1.31 billion [1]. Diabetic wounds, which are often accompanied by a complex pathological microenvironment, including hyperglycemia, oxidative stress, excessive inflammation, vascular network damage, and even large skin defects, making it one of the most challenging clinical treatment problems [2]. Diabetic wound healing proceeds through four overlapping phases: hemostasis, inflammation, proliferation, and remodeling [3]. However, wound healing in patients with diabetes is profoundly disrupted, particularly during the inflammatory phase, where excessive and prolonged inflammation predominates. This impairment is primarily driven by the persistent retention of pro‐inflammatory macrophages, which compromises the orderly transition from inflammation to tissue repair [4]. Macrophages play a pivotal role in wound repair by orchestrating the transition from the inflammatory to the proliferative phase. Within the diabetic microenvironment, chronic hyperglycemia and oxidative stress sustain inflammatory cytokine production, thereby reinforcing M1 macrophage polarization and establishing a self‐amplifying inflammatory loop [5]. Accumulating evidence indicates that this pathological macrophage phenotype is fundamentally encoded by metabolic programming, as classically activated macrophages rely on glycolysis to support inflammatory gene transcription and cytokine synthesis [6, 7]. Hypoxia‐inducible factor‐1α (HIF‐1α) acts as a central transcriptional hub linking inflammatory signaling to glycolytic metabolic reprogramming by regulating glucose transport and glycolysis‐related enzymes, thereby maintaining a pro‐inflammatory metabolic state [6, 8]. Under diabetic conditions, aberrant stabilization of HIF‐1α enhances glycolytic flux in macrophages, locking them into a sustained M1‐like phenotype, whereas inhibition of HIF‐1α attenuates inflammatory metabolism and facilitates tissue repair [5, 8, 9, 10]. Collectively, these findings identify glycolysis as an essential metabolic program for pro‐inflammatory macrophage activation and position the HIF‐1α–glycolysis–macrophage axis as a critical pathological determinant of impaired diabetic wound healing. This dysregulation commonly drives macrophages toward a pro‐inflammatory M1 phenotype and impairs their transition to the anti‐inflammatory M2 phenotype. As a result, the inflammatory response persists, delaying tissue regeneration and contributing to the chronic nature of diabetic wounds.

In addition to abnormal macrophage function, the proliferation and migration capacity of fibroblasts are also suppressed in the diabetic environment, further exacerbating the delay in diabetic wound healing. Fibroblasts are integral to both the inflammatory and proliferative phases of wound healing, with their principal activity occurring during the proliferative stage, where they synthesize new fibrous connective tissue and facilitate wound closure [11]. In diabetic patients, fibroblast dysfunction worsens wound healing delay, highlighting that the impact of diabetes on wound healing is multifaceted, particularly in the disruption of cellular responses during the inflammatory phase. This underscores the urgent need for new therapeutic strategies to restore normal immune responses and promote tissue repair [12, 13].

Currently, commonly used systemic diabetes treatments include metformin, sulfonylureas, and insulin. These treatments mainly aim to control blood glucose levels by increasing insulin secretion or overcoming insulin resistance, thereby alleviating the diabetic condition. However, they are often insufficient to address the specific issues in diabetic wound healing. For local diabetic wound treatment, current approaches predominantly rely on debridement, negative pressure wound therapy, and traditional dressings [14], which are complex and have limited clinical efficacy [15].

In this context, exosomes have garnered widespread attention as a novel therapeutic tool in recent years. Exosomes, which are extracellular lipid bilayer vesicles at the nanoscale, are secreted by almost all cell types under both physiological and pathological conditions [16]. They are involved in a variety of fundamental physiological processes, such as neuronal communication [17], antigen presentation [18], immune responses [19], and organ development [20]. Specifically, exosomes provide a natural lipid bilayer structure that protects miRNAs from rapid degradation, enhances cellular uptake, and enables efficient intracellular delivery. In addition, exosomes carry a complex cargo of bioactive molecules, allowing coordinated and multi‐target regulation that cannot be readily achieved by a single synthetic miRNA. Exosomes, particularly those secreted by stem cells, are rich in bioactive molecules such as proteins, lipids, and small RNAs (such as miRNAs), allowing coordinated and multi‐target regulation that cannot be readily achieved by a single synthetic miRNA [21]. Studies have shown that stem cell‐derived exosomes can significantly enhance wound healing by regulating immune responses and promoting cell migration and differentiation [22]. Among different stem cell sources, dental‐derived stem cells have attracted increasing attention due to their strong regenerative and immunomodulatory potential. In particular, stem cells from the apical papilla (SCAPs) possess robust proliferative and tissue‐regenerative capacity and have been shown to exert superior pro‐regenerative effects compared with other dental stem cell populations, making them a functionally relevant source for exosome‐based therapy in wound repair [23]. Notably, exosomes can modulate macrophage polarization, promoting the conversion of the M1 phenotype to the M2 phenotype, which is crucial for resolving inflammation and facilitating tissue repair. Among these, miRNAs, which are non‐coding RNA molecules typically 18–24 nucleotides in length, have received significant attention due to their role in post‐transcriptional gene regulation [24]. miRNAs can exert powerful effects on cellular pathways and are involved in various diseases, making them promising targets for therapeutic intervention [25]. The therapeutic potential of extracellular vesicles offers considerable promise for clinical translation, however, several challenges remain. Direct injection of exosomes is often limited by their short half‐life, insufficient targeting capacity, and poor control over release kinetics [26, 27]. To address these limitations, hydrogel‐based delivery systems for exosome incorporation have been developed as a promising strategy.

In recent years, hydrogels have become one of the most widely used wound dressings for diabetic wounds in clinical practice. Due to the use of single active compounds, some hydrogel dressings exhibit poor anti‐inflammatory, antibacterial, and regenerative properties. To overcome these limitations, composite hydrogels have emerged as a promising candidate. Their good biocompatibility and customizable physical structure make hydrogels an ideal wound dressing material, and they also possess multifunctional properties, including bacterial prevention and tissue regeneration [28, 29]. The porous structure and appropriate swelling ratio of the hydrogel matrix can effectively absorb exudate, providing a moist environment for diabetic wound treatment and promoting wound healing [30, 31]. Hydrogels can also be loaded with cytokines, mesenchymal stem cells, or extracellular vesicles, further accelerating wound repair by modulating macrophage polarization [32, 33, 34, 35]. In this study, we selected hydroxypropyltrimethyl ammonium chloride chitosan (HACC) as a derivative of chitosan. By grafting cationic quaternary ammonium groups, HACC exhibits superior antibacterial properties and water solubility, with antibacterial activity primarily achieved by disrupting the cell walls of Gram‐positive bacteria [36].

Considering the key roles of macrophage M2 phenotype polarization and fibroblast function in wound healing, developing a novel functional hydrogel dressing based on exosomes to modulate the diabetic wound microenvironment and promote diabetic wound repair is of great significance. In this study, we developed a composite hydrogel dressing using HACC and citric acid as raw materials, incorporating apical papillary stem cell‐derived exosomes. This hydrogel facilitates the sustained release of therapeutic factors, offering a novel treatment strategy for diabetic wound healing(Figure 1).

FIGURE 1.

Schematic illustration of the therapeutic mechanism by which the SCAP‐Exo–loaded CA/HACC hydrogel promotes macrophage phenotypic switching and fibroblast proliferation and migration in diabetic wound healing.

2. Results and Discussion

2.1. Isolation and Identification of Exosomes

To obtain SCAP‐derived exosomes (SCAP‐Exos), apical papilla tissues were isolated from rat molars, and stem cells from the apical papilla (SCAPs) were subsequently harvested. A stable exosome‐producing system was then established using differential ultracentrifugation. (Figure 2A). Under optical microscopy, SCAPs exhibited a good growth state, showing spindle‐shaped adherent cells, which is consistent with the morphological characteristics of mesenchymal stem cells (Figure 2B). Flow cytometry (FCM) analysis revealed that SCAPs expressed mesenchymal stem cell markers CD29, CD44, and CD73, but did not express hematopoietic markers CD34, CD45, or immune‐related marker HLA‐DR, confirming their typical mesenchymal stem cell phenotype (Figure 2F). Subsequently, as shown in Figure 2A, SCAP‐Exos were isolated from SCAPs using differential centrifugation, and exosomes were characterized by transmission electron microscopy (TEM), Western blotting (WB), and nanoparticle tracking analysis (NTA) (Figure 2C–E). These results confirm the successful isolation and extraction of both SCAPs and SCAP‐Exos.

FIGURE 2.

Isolation and characterization of SCAP‐Exos. (A) Schematic illustration of SCAP‐Exo isolation and extraction. (B) Representative images of SCAPs at passages 0 and 5 (scale bar = 200 µm). (C) TEM imaging of SCAP‐Exos (scale bar = 100 nm). (D) Western blot analysis of exosomal and cellular markers. (E) Particle size distribution and mean diameter of exosomes measured by NTA. (F) Flow cytometric analysis of SCAP surface markers (positive: CD29, CD44, CD73; negative: CD34, CD45, HLA‐DR).

2.2. Preparation and Characterization of the Hydrogel

In this study, hydroxypropyltrimethyl ammonium chloride chitosan (HACC) and citric acid (CA) were cross‐linked through an amide bond reaction mediated by EDC/NHS. Subsequently, the incorporation of SCAP‐derived exosomes led to the formation of Exo/CA/HACC hydrogels (Figure 3A). The material exhibited distinct morphological transitions before and after gelation (Figure 3B) and demonstrated favorable injectability and fluidity (Figure 3C). The hydrogel could be smoothly extruded from a sterile syringe and used as ‘ink’ to write letters, indicating its ability to conform to irregular skin defects.

FIGURE 3.

Preparation and characterization of Exo/CA/HACC hydrogels. (A) Schematic of Exo/CA/HACC hydrogel fabrication. (B) Photographs of the hydrogel before (left) and after (right) gelation. (C) Injectable and printable properties of the hydrogel visualized with red dye. (D) The self‐healing performance of hydrogels (E) Hydrogel adhesion on skin and organ surfaces. (F) Adhesion performance test of CA/HACC hydrogel. (G) Rheological characterization: from top to bottom, frequency‐dependent storage (G′) and loss (G″) moduli; self‐healing behavior under destructive shear and recovery; shear‐thinning behavior; and yield stress. (H) SEM images of the CA/HACC hydrogel (scale bars = 100 µm and 50 µm). (I) In vitro release profile of Exos from CA/HACC hydrogel (n = 3).

To assess the self‐healing capability, the hydrogel was divided, one portion was stained with red ink, and the two segments were reassembled. After a short period, the rejoined hydrogel could be lifted under its own weight (Figure 3D), suggesting the presence of dynamic and reversible crosslinking that enhances mechanical stability and enables continuous coverage on uneven wound surfaces, which is particularly relevant for diabetic ulcers.

The CA/HACC hydrogel also adhered firmly to skin and organ surfaces (Figure 3E) and maintained adhesion and stretchability during positional changes. Quantitative adhesion testing demonstrated that the hydrogel exhibits consistent adhesive performance across various substrates (Figure 3F), maintaining effective adhesion even on moist biological tissues, which is critical for wound applications. Although the measured adhesion strength is moderate rather than high, it is sufficient for secure dressing fixation while minimizing the risk of secondary tissue damage upon removal. The cationic nature of HACC likely contributes to this behavior by forming electrostatic interactions with anionic components on tissue surfaces [37]. Such adhesion is advantageous for wound dressings, as it prolongs local exosome retention, minimizes loss under fluid disturbance, and improves stability in moist environments.

Rheological analysis (Figure 3G) showed that the hydrogel formed a stable three‐dimensional crosslinked network with typical solid‐like behavior. G′ exceeded G″ across the tested range, indicating predominantly elastic responses and limited frequency dependence, which supports mechanical integrity when applied to wound beds. The time‐dependent evolution of G′ and G″ demonstrated rapid network formation and stabilization. The viscosity decreased with increasing shear rate, confirming shear‐thinning properties that facilitate injectability. Variations in shear stress further supported the capacity of the internal network to withstand deformation.

SEM imaging revealed a uniform porous architecture with an average pore size of approximately 50 µm (Figure 3H), a scale conducive to maintaining mechanical strength while enabling efficient exosome loading. The porous structure also promotes nutrient exchange and metabolic waste removal, supporting biocompatibility and microenvironmental stability [38, 39].

As shown in Figure 3I, Exo/CA/HACC exhibited a sustained exosome release profile. Exosomes were gradually liberated as the hydrogel degraded, extending their functional duration and preventing rapid loss of bioactivity associated with bolus administration. Then we conducted a test on the water retention performance of the hydrogel (Figure S1), the hydrogel exhibited stable water retention behavior under both conditions, with a gradual and controlled decrease over time, indicating its ability to maintain a hydrated microenvironment. Notably, the retention profile at 37°C remained comparable to that at room temperature.

2.3. Effects of SCAP‐Exos on Macrophages and Fibroblasts

Exosome‐mediated miRNA transfer is recognized as a critical mechanism by which stem cells modulate host immune responses, particularly in chronic inflammatory microenvironments such as diabetic wounds, where exosomal miRNAs exhibit greater stability and targeting capacity compared with free miRNAs [40, 41]. To investigate the biological function of SCAP‐Exos, exosomes were labeled with PKH26 and visualized under fluorescence microscopy, confirming efficient uptake by target cells and successful delivery of functional molecules (Figure 4A). In macrophages under a high glucose environment, SCAP‐Exos modulated macrophage polarization by suppressing M1 markers CD86 and iNOS while upregulating M2 markers CD206 and Arg‐1, indicating promotion of an anti‐inflammatory, reparative phenotype and providing a potential molecular mechanism for inflammation resolution in diabetic conditions (Figure 4B). Additionally, we conducted an immunofluorescence experiment at the protein level (Figure 4C). The results showed that compared with the DM group, the fluorescence signal of iNOS in the DM + Exos group was significantly reduced, while the expression of CD206 was significantly enhanced, and the expression of HIF‐1α protein was synchronously downregulated.

FIGURE 4.

Effects of SCAP‐exos on macrophages and fibroblasts (A) The uptake of PKH26 (red) ‐labeled SCAP‐Exo by L929 was observed by laser confocal microscopy. Scale = 25 µm.(B)The effect of SCAP‐Exos on macrophage polarization under high glucose conditions showed M1 labeling (CD86, iNOS) and M2 labeling (CD206, Arg‐1) (C)Representative images of immunofluorescence detection and 3D Plot analysis (blue = DAPI, green = iNOS/CD206/HIF‐1α; Scale bar = 25 and 50 µm) (D) CCK‐8 assay of L929 cell proliferation. (E) Representative fluorescent images of live/dead staining of L929 cells in each group. (F) Schematic of the L929 migration assay. (G) Representative images and (H) corresponding quantitative analysis of L929 transwell migration in each group. (scale bar = 100 µm). (I) Representative images of the L929 scratch assay (scale bar = 500 µm). (J) Quantitative analysis of scratch closure in each group at different time points.

Given the central role of macrophage‐mediated inflammation in shaping the wound microenvironment, we further investigated the effects of the SCAP‐Exo–based composite hydrogel system on fibroblast behavior. Effective repair and remodeling of damaged skin critically depend on the migration of fibroblasts into the wound site, followed by their proliferation and differentiation. To evaluate the biocompatibility of the composite hydrogel and its influence on cellular migratory behavior, L929 fibroblasts were used as an in vitro model. Four experimental groups were established: an untreated control group, an exosome‐treated group (Exos), a hydrogel‐only group (CA/HACC), and an exosome‐loaded hydrogel group (Exos/CA/HACC).

CCK‐8 assay results (Figure 4D) showed that after 1, 3, and 5 days of culture, all treatment groups maintained cell viability and proliferative capacity comparable to those of the control group. Notably, the Exos/CA/HACC group exhibited the highest proliferation rate at day 5, suggesting that this composite system provides a more stable and sustained microenvironment for the release of bioactive signals. Live/dead staining further confirmed that cell viability exceeded 95% in all four groups (Figure 4E), indicating negligible cytotoxicity and excellent biocompatibility of the materials. These findings are consistent with the growing body of literature highlighting chitosan derivatives as promising skin repair materials due to their low immunogenicity, biodegradability, and ability to promote tissue regeneration [42].

The migratory capacity of L929 fibroblasts was further assessed using Transwell assays (Figure 4F). The number of cells that migrated through the membrane in the Exos/CA/HACC group was significantly higher than in the other groups, representing an approximately twofold increase compared with the control group. This result indicates that the exosome‐loaded hydrogel most effectively promoted directional fibroblast migration (Figure 4G, H). Consistently, scratch wound assays demonstrated that the Exos/CA/HACC group exhibited the most stable and robust migratory performance (Figure 4I and J). The wound closure rate was significantly increased in this group, reaching the highest level at 24 h and achieving nearly complete closure by 48 h, markedly outperforming both the exosome‐only and hydrogel‐only groups. These results collectively confirm the superior migration‐promoting capability of the Exos/CA/HACC composite system.

Importantly, the integrated advantages of Exos/CA/HACC observed in vitro are closely associated with the anti‐inflammatory and immunomodulatory effects identified in our earlier macrophage studies. Exosome‐induced M2 polarization of macrophages attenuates local inflammatory burden, thereby creating a more permissive microenvironment for fibroblast proliferation and migration. Consequently, the composite hydrogel integrates immunomodulation, structural support, and pro‐regenerative bioactivity, constituting a multifunctional wound healing strategy that aligns well with current trends in the development of advanced biomaterials for tissue repair.

2.4. In Vivo Evaluation of Diabetes Wound Healing Effect

The above results highlight the potent ability of Exos/CA/HACC in modulating inflammation and skin fibroplasia, as well as its reassuring biocompatibility. Next, we evaluated their feasibility as an effective wound dressing for chronic diabetic wounds. For this purpose, in Figure 5A, we selected male SD rats (4–6 weeks old) and induced type 1 diabetes by injecting streptozotocin (STZ) to explore the dynamics of wound healing in a diabetic injury model. After STZ injection, diabetic rats displayed typical diabetic symptoms, including thirst, hunger, increased urination, and significantly reduced activity levels. Additionally, random blood glucose levels progressively increased and remained hyperglycemic throughout the experiment (Figure 5B).

FIGURE 5.

In Vivo evaluation of diabetic wound healing. (A) Schematic illustration of diabetic rat model establishment and hydrogel treatment. (B) Blood glucose levels of rats during the wound healing process. (C) Representative digital images of wounds in each group at selected time points. (D) Wound area changes over time. (E) Representative H&E and Masson's trichrome staining images of each group (scale bars = 1000 µm and 100 µm). (F) Wound length at the wound site on days 7 and 14 (n = 3). (G) Epithelial thickness at the wound site on day 14 (n = 3). (H) The immunofluorescence images and 3D Plot analysis of macrophages in the regenerated tissue on the seventh day after treatment with iNOS and Arg‐1 staining. (scale bar = 50 µm). (I) Immunofluorescence images of macrophages stained for HIF‐1α in regenerated tissues on day 14. Green: HIF‐1α; blue: DAPI (scale bar = 50 µm).

Full‐thickness circular wounds (18 mm in diameter) were created on the dorsal surface of the rats. Then, CA/HACC, exosomes, and Exos/CA/HACC were applied to the wounds, followed by simple bandaging. Figure 5C presents representative wound images captured at preset time intervals for each group, along with overlaid wound healing analysis. All treatment groups exhibited a similar trend of gradual wound contraction over time, but with significant differences in contraction rates. On day 14 post‐surgery, the wounds in the control and diabetic groups were still visibly open. The wound healing area changes shown in Figure 5D indicate that the Exos/CA/HACC group achieved nearly complete wound healing by day 14, whereas wound healing in the diabetic group was notably delayed. Although the healing efficiency in the CA/HACC group was not significantly higher than that in the Exos/CA/HACC group, its therapeutic effect was relatively better in the early stages of healing (day 7). Moreover, at all monitored time points, the healing rate of diabetic wounds treated with Exos/CA/HACC was significantly faster than in the diabetic group. From a macroscopic perspective, wounds in the diabetic group treated with Exos/CA/HACC remained unhealed at the time points corresponding to complete closure. Throughout the healing process, exosomes deliver miR‐122‐5p to macrophages within the wound microenvironment, mediating immunomodulation and inflammation resolution, while the hydrogel network maintains a local moist environment, stabilizes exosome localization, and provides sustained release support. This enables the system to function effectively during both the early inflammatory phase and the tissue proliferative phase. The pro‐healing effects of Exos/CA/HACC are consistent with its immunomodulatory and regenerative functions observed in vitro, demonstrating a clear advantage of this hydrogel in accelerating wound closure.

Wound healing requires complex biological and molecular processes and interactions, including specific biological events such as hemostasis, inflammation, cell proliferation, and tissue reconstruction. To assess the therapeutic effects of the composite hydrogel dressing from a histological perspective, wound skin tissues collected at fixed time points were subjected to Hematoxylin and Eosin (H&E) staining and Masson's trichrome staining. According to the H&E staining (Figure 5E), after 14 days, the Exosomes and Exos/CA/HACC groups showed reduced inflammatory cell infiltration and narrower wound widths (Figure 5F). However, in the diabetic group, inflammatory cell infiltration was still present on day 14, indicating that the wound remained in the inflammatory phase. This indicates that the composite hydrogel successfully facilitates the dynamic transition of the wound microenvironment from a pro‐inflammatory state to a reparative state. Additionally, the epithelial thickness in the diabetic, Exosomes, and CA/HACC groups was significantly greater than in the Exos/CA/HACC group, suggesting that the exosome‐loaded hydrogel group had progressed beyond the inflammatory stage and completed reepithelialization, forming mature thin epithelium (Figure 5G). When compared at the same time point, the tissue structure in the Exos/CA/HACC group appeared more organized than in the diabetic group. Masson's staining (Figure 5E) showed increased collagen density in the wound tissues across all groups. In the exosome hydrogel treatment group, abundant new epithelial structures, skin appendages, and granulation tissue were clearly observed.

The immunofluorescence staining of macrophage markers (iNOS and Arg‐1) indicated that the polarization of macrophages had changed (Figure 5H). In the Exos/CA/HACC group, the M1 type cells decreased while the M2 type cells increased, suggesting a shift in the immune microenvironment towards a reparative direction. Immunofluorescence analysis of wound tissues revealed that HIF‐1α fluorescence intensity was markedly elevated in the diabetic microenvironment (Figure 5I), whereas treatment with Exos/CA/HACC significantly reduced its intensity. As a central regulator of oxygen metabolism and inflammatory responses, HIF‐1α is aberrantly activated under high‐glucose and hypoxic conditions, promoting pro‐inflammatory M1 polarization and impairing normal tissue repair. The suppression of HIF‐1α expression or activity by Exos/CA/HACC provides a molecular basis for its ability to drive macrophages toward a reparative phenotype through immunometabolic pathways [43].

2.5. MiRNA Profiling of SCAP‐Exos Identifies miR‐122‐5p as a Key Immunometabolic Regulator

In recent years, accumulating evidence has revealed that stem cell–derived exosomes are not merely passive carriers of paracrine signals, but instead exert pivotal regulatory functions through their selectively enriched miRNA cargos, orchestrating inflammation, metabolism, and tissue repair at multiple levels [44]. Exosomes derived from stem cells deliver abundant bioactive molecules, particularly miRNAs, to modulate the functions of recipient cells. By binding to the 3′ untranslated region (3′UTR) of target mRNAs, miRNAs play a central role in maintaining metabolic homeostasis by inducing mRNA degradation or translational repression [45, 46].

The volcano plot (Figure 6A) illustrates differentially expressed miRNAs in SCAP‐Exo, revealing a substantial number of significantly upregulated miRNAs (orange) compared with the control group. These findings indicate that SCAP‐Exo does not randomly encapsulate miRNAs, but instead selectively enriches a distinct set of biologically active miRNAs. Given that the therapeutic efficacy of exosomes is largely attributed to upregulated miRNAs, we further focused on the highly expressed miRNAs in SCAP‐Exo. A heatmap (Figure 6B) was generated to display the top 10 upregulated miRNAs in SCAP‐Exo relative to the SCAP group, including miR‐150‐5p, miR‐126‐5p, and miR‐122‐5p. Although miR‐122‐5p was initially recognized as a liver‐specific miRNA involved in metabolic regulation [47, 48], increasing evidence has demonstrated that it participates in immune cell functional reprogramming by modulating metabolism‐related signaling pathways and inflammatory mediator expression, and is closely associated with macrophage metabolic status and inflammatory phenotypes [49]. Accordingly, we further predicted the target genes of these miRNAs (Figure 6C) and subjected them to Gene Ontology (GO; Figure 6D) and Kyoto Encyclopedia of Genes and Genomes (KEGG; Figure 6E) enrichment analyses.

FIGURE 6.

miRNA Profiling of SCAP‐Exos and Functional Validation of miR‐122‐5p (A) Volcano plot showed downregulated and upregulated miRNAs in SCAP‐Exo. (B)Heat map exhibited top 10 upregulated miRNAs in SCAP‐Exo compared with SCAP. Orange and blue denote high and low expression. (C)Target gene prediction network around miR‐122‐5p (D) GO enrichment of miR‐122‐5p (E) KEGG enrichment of miR‐122‐5p.

The predicted target genes were broadly involved in HIF‐1α–related signaling pathways, glycolytic metabolism, regulation of inflammatory cytokine expression, and signaling pathways governing cell migration and proliferation. GO enrichment analysis revealed that these miRNA targets were significantly enriched in biological processes that cooperatively regulate inflammatory responses and fibroblast behavior in diabetic wounds. Furthermore, KEGG pathway analysis enabled the construction of an interaction network among enriched pathways, highlighting coordinated involvement in metabolic reprogramming, restoration of cell–cell junctions and migratory capacity, and amplification of intercellular signaling mediated by exosomes. Collectively, these pathways suggest that SCAP‐Exo mitigates the dysregulated “metabolism–inflammation–migration” axis in diabetic wounds through a concerted miRNA regulatory network.

Based on these observations, miR‐122‐5p was selected as a representative candidate miRNA for functional validation. Quantitative PCR analysis showed that miR‐122‐5p expression in macrophages was significantly increased 8 h after SCAP‐Exo treatment, indicating that miR‐122‐5p is a functional miRNA efficiently delivered by SCAP‐Exos (Figure 7A). In addition, transfection with miR‐122 mimics markedly elevated intracellular miR‐122‐5p levels, confirming the successful establishment of an overexpression model (Figure 7B).

FIGURE 7.

miRNA Profiling of SCAP‐Exos and Functional Validation of miR‐122‐5p (A) qRT‐PCR analysis for the expression of miR‐122‐5p in SCAP and SCAP‐Exo, respectively. qRT‐PCR analysis for the expression of miR‐126‐5p in RAW264.7(B)The expression level of miR‐122‐5p in RAW264.7 was detected by qRT‐PCR after incubation with SCAP‐Exo for 8 h. (C) Expression of phenotypic genes (CD86, CD206, iNOS, Arg‐1) in macrophages after transfection with miR‐122‐5p mimics. (D)DCFH‐DA fluorescence probe detection and 3D Plot analysis of reactive oxygen species (ROS) levels in RAW264.7 transfected with miR‐122 mimics. (Scale bar = 50 µm) (E)Representative immunofluorescence staining images show the expression of DAPI (blue) and HIF‐1α (green). (Scale bar = 50 µm.) (F)Bioinformatics prediction and the target sequence of the dual‐luciferase reporter gene system demonstrated the binding relationship between the miR‐122‐5p sequence and the wild‐type HIF‐1α (HIF‐1α‐WT) (G)The expression levels of HIF‐1α, GLUT1 and LDHA in macrophages after transfection with miR‐122‐5p mimics were analyzed by qPCR. (H) Effects of miR‐122‐5p mimic and HIF‐1α inhibition on macrophage inflammatory polarization. (I) Effects of miR‐122‐5p mimic and HIF‐1α inhibition on glycolysis‐related genes.

Given that macrophage phenotypic imbalance—characterized by persistent M1 activation and impaired M2 transition—is a hallmark of delayed diabetic wound healing, we next evaluated the role of miR‐122‐5p in macrophage polarization. qPCR analysis revealed that miR‐122 mimics significantly downregulated the expression of M1‐associated markers CD86 and iNOS, while markedly upregulating M2‐associated markers CD206 and Arg‐1. Notably, CD86 and iNOS expression levels were even lower than those observed in the LPS‐treated group, indicating a pronounced anti‐inflammatory regulatory effect (Figure 7C). Consistent with these findings, DCFH‐DA fluorescence staining and three‐dimensional intensity analysis demonstrated that intracellular ROS levels in the miR‐122 mimics group were significantly reduced compared with the LPS and NC mimics groups (Figure 7D). These results suggest that miR‐122‐5p promotes the maintenance of an anti‐inflammatory, reparative macrophage phenotype by alleviating oxidative stress. Considering the well‐established positive feedback between ROS accumulation and pro‐inflammatory signaling pathways such as HIF‐1α [50], suppression of ROS may represent an early event in miR‐122‐5p–mediated immune regulation.

To further elucidate the anti‐inflammatory signaling pathways involved, immunofluorescence staining was performed to assess HIF‐1α expression in macrophages. As shown in Figure 7E, LPS treatment markedly increased HIF‐1α protein levels, whereas co‐treatment with miR‐122 mimics significantly attenuated HIF‐1α expression. These findings indicate that miR‐122‐5p may regulate macrophage polarization by interfering with the HIF‐1α–dependent pro‐inflammatory signaling axis. This regulatory relationship was further confirmed by bioinformatic prediction and dual‐luciferase reporter assays, which revealed a complementary binding site between miR‐122‐5p and the 3′UTR of wild‐type HIF‐1α (HIF‐1α‐WT). Mutation of this binding region (HIF‐1α‐MUT) abolished the interaction, demonstrating that miR‐122‐5p directly targets the HIF‐1α 3′UTR (Figure 7F).

Given the central role of HIF‐1α in macrophage immunometabolism—where HIF‐1α–driven GLUT1‐mediated glucose uptake and LDHA‐dependent lactate production collectively support the metabolic program of M1 macrophages—we further examined hypoxia‐ and glycolysis‐related genes. qRT‐PCR analysis showed that miR‐122 mimics significantly reduced the mRNA levels of HIF‐1α, GLUT1, and LDHA compared with other groups (Figure 7G), indicating that miR‐122‐5p suppresses glycolytic flux and thereby limits the persistence of pro‐inflammatory signaling at its metabolic source. To further strengthen the mechanistic link between miR‐122‐5p and HIF‐1α signaling in macrophage polarization, an HIF‐1α pharmacological inhibition group (PX‐478) was included for comparison. Gene transcript level analysis of macrophages showed that both miR‐122‐5p mimic and PX‐478 treatment significantly reversed LPS‐induced M1 polarization, and increased expression of CD206 and Arg‐1(Figure 7H). Meanwhile, both interventions markedly reduced the expression of HIF‐1α and its downstream glycolytic targets GLUT1 and LDHA compared with the LPS group (Figure 7I). These results further support the critical role of the miR‐122‐5p/HIF‐1α in regulating macrophage polarization.

Taken together, these findings demonstrate that miR‐122‐5p markedly reduces intracellular oxidative stress in macrophages, a phenomenon that is intrinsically linked to HIF‐1α inhibition. On the one hand, attenuation of glycolysis may decrease mitochondrial electron leakage and subsequent ROS generation [51]; on the other hand, reduced ROS levels may reciprocally destabilize HIF‐1α, forming a positive regulatory loop that further favors macrophage polarization toward the M2 reparative phenotype [52]. Therefore, the miR‐122‐5p/HIF‐1α axis identified in this study not only modulates inflammatory gene expression but also governs macrophage fate at the metabolic level (Figure 8).

FIGURE 8.

The miR‐122‐5 derived from SCAP‐Exo may be involved in the signaling pathway that regulates macrophage phenotypic differentiation.

In summary, miRNA sequencing and functional validation demonstrate that SCAP‐Exos selectively enrich functional miRNAs, including miR‐122‐5p, thereby establishing a coordinated molecular network that counteracts the dysregulated microenvironment of diabetic wounds. Through the integrated regulation of oxidative stress, metabolic reprogramming, and inflammatory signaling, this network orchestrates macrophage phenotypic switching toward a reparative state. Such miRNA‐mediated fine‐tuning likely constitutes a fundamental mechanistic basis for the pronounced pro‐healing effects of SCAP‐Exos in chronic diabetic wounds. Nevertheless, the intrinsic complexity of exosomal cargo should be taken into consideration. Exosomes are heterogeneous nanovesicles containing diverse classes of biomolecules—including multiple miRNAs, proteins, and lipids—that may act in a coordinated and context‐dependent manner rather than through a single dominant effector. Our previous work has demonstrated that miR‐126‐5p plays a critical role in angiogenic regeneration, supporting its potential involvement in the observed therapeutic effects [23]. Therefore, the therapeutic effects observed here are unlikely to be attributed exclusively to miR‐122‐5p, and potential synergistic interactions among exosomal components warrant further mechanistic investigation. In this context, future work will incorporate controlled delivery systems for synthetic miRNAs, including hydrogel‐based platforms, to enable direct comparison with exosome‐mediated delivery and to better delineate the relative contributions of individual RNA species versus the vesicular carrier.

Furthermore, diabetic wound healing is a dynamic, multicellular process requiring coordinated interactions among diverse cell populations. While this study mainly focuses on macrophages and fibroblasts, other key cell types, such as endothelial cells and keratinocytes, play indispensable roles respectively in angiogenesis and re‐epithelialization processes. Although accelerated wound closure was observed, these processes were not directly evaluated. Future studies integrating cell‐specific functional assays and in vivo tracking approaches will be essential to define the broader impact of SCAP‐Exos‐loaded hydrogel on the wound microenvironment and to further elucidate its multi‐target mechanisms.

3. Conclusions

The Exos/CA/HACC composite hydrogel developed in this study demonstrates coordinated advantages in both structural performance and biological functionality, offering a practical material strategy for the repair of diabetic chronic wounds. The hydrogel exhibits favorable gelation properties and exosome‐loading capacity, enabling stable release of SCAP‐Exos while preserving their bioactivity, which in vitro significantly promotes fibroblast proliferation and migration, mitigates inflammatory responses, and facilitates macrophage polarization toward a reparative phenotype. In vivo, the material effectively alleviates prolonged inflammation in diabetic wounds, accelerates re‐epithelialization and collagen deposition, and restores tissue structural integrity and organization. Mechanistic investigations suggest that the highly expressed miR‐122‐5p within exosomes may downregulate HIF‐1α and its glycolysis‐related pathways, providing critical regulation for the transition from inflammation to the proliferative phase. Overall, the Exos/CA/HACC hydrogel enhances diabetic skin repair through the synergistic effects of its material properties and biological functions, providing a promising foundation for the development of materials for chronic wound therapy.

4. Materials and Methods

4.1. Cell Isolation and Culture

Ten 1‐week‐old SD rats were selected, and 2% pentobarbital (40–50 mg/g) was injected intraperitoneally. After anesthetizing the rats, they were euthanized by cervical dislocation. The apical papillary tissue of the rat's tooth root was isolated and cut into small pieces. The tissue was then digested in type I collagenase (Solabio, China) solution and incubated in a 37°C water bath for approximately 30 min. After sufficient digestion, the collagenase solution was removed, and the tissue suspension was mixed in Alpha Minimum Essential Medium (α‐MEM; SIGMA) supplemented with 10% fetal bovine serum (FBS; AusGeneX, Australia) and 1% penicillin‐streptomycin (Gibco). The cells were cultured in 25 cm² culture flasks and incubated at 37°C in 5% CO₂. Early passage stem cells from apical papilla (SCAP) s (P3‐P8) were used for subsequent experiments. RAW264.7 and L929 cells were cultured in DMEM medium containing 10% FBS. Cells were subcultured every 2–3 days.

4.2. Exosome Isolation and Purification

When SCAPs reached 60%–70% confluence, they were washed with PBS and cultured in medium containing 10% exosome‐depleted FBS (normal FBS was centrifuged at 11,000 g for 16 h to deplete exosomes). After 48–72 h of culture, the medium was collected and stem cells from apical papilla‐derived exosomes (SCAP‐Exo) isolated by gradient ultracentrifugation. Briefly, the supernatant was centrifuged at 300 g for 10 min to remove cells, at 2000 g for 15 min, and at 12,000 g for 45 min to remove debris and large vesicles. The supernatant was further filtered through a 0.22 µm filter, then ultracentrifuged at 11,000 g for 70 min at 4°C. SCAP‐Exos were resuspended in PBS and stored at ‐80°C for further use.

4.3. Exosome Characterization

Exosome solutions were dropped onto carbon‐coated grids and air‐dried overnight. The morphology of SCAP‐Exo and DPSC‐Exo was observed by transmission electron microscopy (Hitachi, Japan). Particle size and concentration were measured by nanoparticle tracking analysis (NTA) using a ZetaView PMX 110 (Particle Matrix, Germany) and ZetaView 8.04.02 software. Exosome markers were detected by western blot. Exosomes were lysed in ice‐cold RIPA buffer, separated by SDS‐PAGE, and transferred to PVDF membranes. After blocking with 5% skim milk, membranes were incubated overnight with primary antibodies at 4°C, then with secondary antibodies at room temperature. Bands were detected using chemiluminescence (Bio‐Rad).

4.4. Preparation of CA/HACC Hydrogels Loaded With SCAP‐Exos

Hydroxypropyltrimethyl ammonium chloride chitosan (HACC), citric acid (CA), N‐(3‐dimethylaminopropyl)‐N'‐ethylcarbodiimide hydrochloride (EDC), and N‐hydroxysuccinimide (NHS) were obtained from Shanghai Macklin Biochemical Technology Co., Ltd. (Shanghai, China). All reagents were of analytical grade and used without further purification.

To prepare the hydrogel, 1.25 g of HACC was dissolved in deionized water, followed by the addition of 0.125 g of CA after complete dissolution. Subsequently, 0.5 g of EDC and 0.1 g of NHS were weighed and slowly added dropwise into the solution under magnetic stirring. The mixture was maintained at 25°C for 2 days, washed several times with anhydrous ethanol, and the resulting white flocculent precipitate was rinsed with deionized water. SCAP‐Exos were simultaneously introduced during the rinsing process. The final hydrogel containing SCAP‐Exos was stored at 4°C.

4.5. Characterization of CA/HACC Hydrogels

Clear photographs were taken before gelation (solution state) and after complete gel formation to visually demonstrate the transition in physical morphology. Red ink was incorporated into the hydrogel, mixed uniformly, loaded into a sterile syringe, and extruded under gravity or constant pressure. Images of the extrusion process were captured to assess whether the hydrogel could pass continuously and uniformly through the needle while maintaining a stable filament.

The hydrogel was cut into two halves, with one portion mixed evenly with red ink, and photographs were taken before and after contact to evaluate its self‐healing capacity. The adhesive properties were examined by attaching the hydrogel to the skin surface and photographing it under upright, inverted, and flexed finger positions. For this experiment involving the contact of hydrogel with human skin, written informed consent forms have been obtained from all participants. Additional adhesion tests were conducted by using tweezers to place the hydrogel onto the surface of excised organs, followed by photographing its attachment under static gravitational conditions.

Rheological measurements were performed using a Discovery Hybrid Rheometer (DHR; TA Instruments) at 25°C with a working gap of 1000 µm. The viscoelastic properties were assessed through oscillatory frequency sweeping (0.1 rad/s, 0.5% constant strain). Self‐healing behavior was evaluated by monitoring changes in the storage modulus (G′) and loss modulus (G″) during destructive shear (1 Hz frequency, strain increased from 0.1% to 1000%) and subsequent recovery (0.5% strain, 1 Hz frequency, 200 s oscillation). Viscosity was measured over shear rates ranging from 0.1 to 50 s⁻¹ to determine shear‐thinning characteristics. Yield stress was assessed through oscillatory stress sweeps at 1 Hz, and the yield point was defined as the crossover where G″ exceeded G′.

After freeze‐drying, hydrogel samples were collected to preserve their microstructure. The fractured cross‐sections were sputter‐coated with gold, and the microscopic morphology was examined at various magnifications using a field‐emission scanning electron microscope (SEM, JSM‐7001F, JEOL).

4.6. Exosome Release Rate From Hydrogel

The in vitro release of Exo@CS‐Gel was studied using a Transwell system [53]. DiO (100 mg)‐labeled Exo/CA/HACC was placed in the upper chamber of a 24‐well Transwell, allowing Exo to pass freely through the semi‐permeable membrane. Release medium (PBS containing 10% FBS) was added to the lower chamber (1.2 mL). The system was incubated at 37°C in a shaking incubator at 60 rpm. At designated time points, 400 µl of the lower chamber solution was collected (with fresh release medium added to replenish the volume). Fluorescence intensity was measured using an excitation wavelength of 488 nm and emission wavelength of 525 nm to calculate the cumulative release rate.

$$\begin{array}{cc}& \mathrm{Cumulative}\mathrm{release}\mathrm{rate}(\%)\hfill \\ & =\mathrm{Cumulative}\mathrm{amount}\mathrm{of}\mathrm{released}\mathrm{Exo}/\mathrm{Total}\mathrm{amount}\mathrm{of}\mathrm{Exo}\hfill \\ & \times 100\%\hfill \end{array}$$

4.7. MiRNA Transfection Method

RAW264.7 cells were cultured to 60%–70% confluence and transfected with miR‐122‐5p mimic using lipofection according to the manufacturer's instructions. The cells were incubated with the miR‐122‐5p mimic at a final concentration of 50 nM for 6 h, after which the medium was replaced with regular culture medium to terminate transfection. After 48 h, gene expression levels were analyzed using quantitative reverse transcription PCR.

4.8. SCAP‐Exo Uptake by Cells

SCAP‐Exo solution was thawed on ice and mixed with 0.25 mL Diluent C and 2 µl PKH26 for 5 min, then added to α‐MEM medium. The labeled SCAP‐Exo solution was incubated with L929 cells for 24 h. After incubation, the cells were washed twice with PBS, stained with DAPI for nuclei, and analyzed by fluorescence microscopy to observe SCAP‐Exo uptake.

4.9. Detection of Intracellular ROS

DCFH‐DA (S0033S, Biyuntian, China) was used to measure intracellular ROS levels. LPS (1 µg/mL) was added to the medium to induce an inflammatory microenvironment. RAW264.7 cells were seeded in a 24‐well plate, and incubated under different conditions (NC mimics, miR‐122 mimics) to reach about 80% confluence. The cells were washed three times with PBS, then loaded with DCFH‐DA (10 mM) for 30 min at 37°C. After incubation, cells were washed three times with PBS and analyzed under fluorescence microscopy.

4.10. Immunofluorescence Assay

RAW264.7 cells were transfected with miR‐122 mimics at a density of 4×10⁵/mL and co‐cultured for 48 h. Cells were fixed with 4% paraformaldehyde for 30 min at room temperature, washed twice with PBS, and permeabilized with Triton X‐100 for 15 min. Blocking was performed for 60 min, followed by incubation with HIF‐1α (A15863, 1:100) at 4°C overnight. The cells were then incubated with a fluorescence‐conjugated secondary antibody at room temperature for 1 h in the dark, washed with PBS, and stained with DAPI for 5 min. Imaging was performed using confocal laser scanning microscopy (CLSM).

4.11. RNA Extraction and qRT‐PCR Analysis

Total RNA from RAW264.7 or L929 cells was extracted using RNAiso Plus reagent (Takara, Japan) for mRNA analysis. Then, 500 ng of total RNA was reverse transcribed into complementary cDNA using the 5×PrimeScript RT Master Mix (Takara, Japan). Quantitative reverse transcription PCR (qRT‐PCR) was performed using a 10 µL reaction mixture containing cDNA, TB Green PreMix Ex Taq (Takara, Japan), forward and reverse primers, and RNase‐free water. The M1 markers CD86 and iNOS, and the M2 markers CD206 and Arg‐1, were quantified, were validated, with GAPDH as a housekeeping gene. MiRNA was extracted using the miRcute miRNA Isolation Kit (TIANGEN, China), followed by reverse transcription into cDNA using the miRcute Plus miRNA First‐Strand cDNA Kit (TIANGEN, China). The diluted cDNA, 2×miRcute Plus miRNA PreMix (TIANGEN, China), and primers were mixed for qRT‐PCR. The primers were designed by Takara (Japan). Let‐7a‐5p was used as a housekeeping miRNA, and the relative expression levels of miRNA‐122‐5p were analyzed. qRT‐PCR was performed on a CFX96 system (Bio‐Rad). Gene expression levels were calculated relative to the blank control (fold change).

4.12. Cytotoxicity Assay

L929 cells (5×10⁴) were seeded in 96‐well plates overnight. After treatment with different groups (control, exosome, hydrogel, exosome‐loaded hydrogel), cells were incubated for 1, 3, and 5 days. The medium was discarded, and the cells were washed with PBS. Then, 10% CCK‐8 reagent in basal medium was added to each well. After several hours, absorbance at 450 nm was measured using a microplate reader. Cell proliferation rate was calculated as:

$$\begin{array}{cc}& \text{Cell}\text{Proliferation}\text{Rate}\hfill \\ & =\text{(Experimental}\text{group}\text{OD}\mathrm{/}\text{Control}\text{group}\text{OD)}\times 100\%.\hfill \end{array}$$

4.13. Live/Dead Assay

L929 cells (1×10⁵) were seeded in 24‐well plates overnight. After treatment with different groups (control, exosome, hydrogel, exosome‐loaded hydrogel), cells were incubated for 24 h. The cells were washed with PBS, then stained with live/dead staining solution for 15 min. After washing with PBS, cells were observed under an optical microscope.

4.14. Scratch Assay

A linear scratch model was used to assess L929 cell migration. L929 cells (6×10⁴) were seeded in 6‐well plates overnight, then linear scratches were made with a sterile P200 pipette. Exosome, hydrogel extract, and exosome‐loaded hydrogel extract were added to the culture medium. Cells were incubated for 0, 24, and 48 h. After washing with PBS, cells were observed under an optical microscope. Scratch area was measured using ImageJ software, and healing rate was calculated as:

$$\begin{array}{cc}& \text{Wound}\text{Healing}\text{Rate}\hfill \\ & =\text{[(0}\mathrm{h}\text{wound}\text{width)}\hfill \\ & -\text{(wound}\text{width}\text{at}\text{each}\text{time}\text{point)]}\mathrm{/}\text{(0}\mathrm{h}\text{wound}\text{width)}\hfill \\ & \times 100\%.\hfill \end{array}$$

4.15. Transwell Migration Assay

L929 cells (1×10⁵ cells/well) were seeded in the upper chamber of a Transwell plate overnight. The medium in the lower chamber was replaced with complete medium containing exosomes, hydrogel extract, or exosome‐loaded hydrogel extract. After 24 h, cells were fixed with 4% paraformaldehyde for 15 min and stained with 1% crystal violet. Non‐migrated cells in the upper chamber were removed with a cotton swab. Migration was observed under an optical microscope.

4.16. Diabetes Wound Healing Experiments In Vivo

Animal experiments were performed according to ARRIVE guidelines and approved by the Shanxi Medical University Animal Ethics Committee (License No.: CGSC‐2025‐010). Thirty male SD rats (4‐6 weeks old, 150–200 g) were housed under standard light/dark conditions with free access to water and a balanced diet. All procedures were carried out in accordance with the UK Animals (Scientific Procedures) Act of 1986. Briefly, male SD rats were injected with streptozotocin (STZ, 50 mg/kg) to induce diabetes. Blood glucose levels were monitored weekly via tail vein blood sampling, and T1DM was considered established when blood glucose exceeded 16.7 mmol/L and stabilized. After anesthesia, four full‐thickness circular wounds (approximately 18 mm in diameter) were made on the dorsal area of each rat. The rats were divided into five groups: blank, diabetes, exosome, hydrogel, and exosome‐loaded hydrogel. After sterilizing the materials, they were placed on the wounds. Photographs of the wounds were taken on days 1, 4, 7, and 17 post‐surgery. Wound healing was assessed using ImageJ software, and healing rate was calculated using:

$$\mathrm{Wound}\mathrm{Healing}\mathrm{Rate}=(\mathrm{R}0-\mathrm{Rx})/\mathrm{R}0,$$

where R₀ was the wound area on day 0 and Rx was the wound area on day x (x = 3, 7, 10, 14).

4.17. Histological and Immunofluorescence Analysis

On days 7 and 17, residual wound tissue was collected, fixed in 4% paraformaldehyde, and embedded in paraffin. Sections (5 µm) were cut longitudinally for H&E and Masson's staining to assess inflammation and collagen deposition. Immunofluorescence staining was used to evaluate HIF‐1α expression in skin tissue and assess inflammation and tissue repair in the wound. ImageJ was used for quantitative analysis of immunofluorescence staining.

4.18. MiRNA Sequencing Analysis

To comprehensively characterize the miRNA expression landscape of SCAP and SCAP‐derived exosomes (SCAP‐Exo), small RNA sequencing (miRNA‐seq) was performed. Small RNA libraries were constructed using the NEBNext Multiplex Small RNA Library Prep Set for Illumina (NEB) and subsequently sequenced on an Illumina platform.

After quality filtering, clean reads were aligned to the miRBase database, and only perfectly matched sequences were retained for the identification and quantification of known miRNAs. MiRNA expression levels were normalized using the transcripts per million (TPM) method.

Differentially expressed miRNAs were identified using the DESeq2 algorithm with the criteria of |log₂ fold change| > 1 and a false discovery rate (FDR) < 0.05. Putative target genes of the differentially expressed miRNAs were predicted by integrating results from TargetScan, miRDB, and StarBase databases. Functional annotation and pathway enrichment analyses were subsequently conducted using Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) analyses to elucidate the biological processes and signaling pathways potentially regulated by these miRNAs.

4.19. Statistical Analysis

Statistical analysis was performed using GraphPad Prism 9 software. Data were presented as mean ± standard deviation (n ≥ 3). Final data were generated using GraphPad Prism. Statistical significance was determined by *p < 0.05 and **p < 0.01. One‐way or two‐way ANOVA with Tukey's multiple comparisons test was used for data analysis.

Funding

National Natural Science Foundation of China (82470959), Research Project Supported by Shanxi Scholarship Council of China (2024‐076), Scientific Activities of Selected Returned Overseas Professionals in Shanxi Province (20240016), Scientific research project of Shanxi Traditional Chinese Medicine Administration (2024ZYYB050), the Key Open Project of the National Key Laboratory for Oral Disease Prevention and Treatment (SKLOD2025OF07), Doctor to Jin work project of Shanxi Province (SD2333), Shanxi Medical University doctoral start‐up fund project (XD2233), the Fundamental Research Program of Shanxi Province, China (No. 202503021211038), Joint Construction of Shanxi Key Laboratory of Functional Proteins (No. RZ2500002801), Fundamental Research Program of Shanxi Province (202303021222140).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm71338‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.