Twenty adult male Sprague–Dawley (SD) rats (weighing 180 g–200 g) were obtained from the Shanghai Slack Laboratory Animal Center (license number: SCXK hu 2022-0004). All rats were housed in a standard specific pathogen-free (SPF) barrier environment at 20 °C–26 °C and 40%–70% humidity under a 12 h/12 h light–dark cycle. All animal experiments were approved by the Experimental Animal Ethics Center of Mengchao Hepatobiliary Hospital of Fujian Medical University (MCHH-AEC-2022-08). All experiments were designed and reported in accordance with the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guidelines 2.0.

The ADSCs were isolated and cultured according to previously published methods (Liao et al., 2017). In brief, adipose tissues were obtained from the inguinal region of male SD rats (n = 5) and washed with PBS solution. The tissues were then cut into small fragments and digested with 0.1% type I collagenase, followed by neutralization with α-MEM containing 10% FBS. Subsequently, the cells were cultured at a density of 1 × 10⁶ cells/mL in T-75 plates. ADSCs at passage 3 were collected and cultured at a density of 2 × 10⁶ cells per 10 cm plate. After overnight cell adhesion, the cultured ADSCs were washed with PBS solution to remove residual serum and then replaced with serum-free medium (YOCON, China) for 48 h. After incubation, the conditioned medium (10 mL/2 × 10⁶ ADSCs) was collected and centrifuged at 3,000 g for 5 min to remove any cell debris. Furthermore, the ADSC-conditioned medium was concentrated using ultrafiltration with a tangential flow filtration capsule (Pall, United States) containing a 3-kDa molecular weight cut-off membrane, following the manufacturer’s instructions. Finally, the concentration of the ADSC-conditioned medium was analyzed using a BCA assay kit (TransGen Biotech, China) and stored at −80 °C. A concentration of 100 ng/mL ADSC-conditioned medium was used in the present study.

The human umbilical vein endothelial cell (HUVEC) line was obtained from the National Institutes for Food and Drug Control (Beijing, China) and cultured with RPMI 1640 containing 10% FBS supplemented with 2% FBS, VEGF, IGF-1, and EGF at 37 °C/5% CO₂. For the tubule formation assay, 96-well plates were pre-coated with Matrigel (Corning, 10 mg/mL, 1:50 dilution in EGM-2) for 1 h at 37 °C to simulate the basement membrane matrix, as standardized in angiogenesis assays.

HUVECs were cultured at a density of 1 × 10⁴ cells per well in 96-well plates. After overnight cell adhesion, the cell supernatants were removed and replaced with 100 μL ADSC-conditioned medium, while the cells treated with serum-free medium were used as the negative control. After incubation for 24 h or 48 h, cell viability was evaluated using a CCK-8 assay kit (TransGen Biotech, China), according to the manufacturer’s instructions.

Total RNA was collected using a TRIzol reagent kit (TransGen Biotech, China) following the manufacturer’s instructions. Afterward, mRNA was reverse-transcribed into cDNA using a cDNA synthesis kit (Roche, Germany). The quantitative real-time PCR analysis was performed in an ABI StepOnePlus Real-time PCR System (Carlsbad, United States), and the PCR conditions were as follows: 95 °C for 15 s, 60 °C for 30 s, and 70 °C for 30 s, for a total of 40 cycles. The primer sequences are listed in Table 1. The 2^-△△Ct formula was used to analyze the relative gene expression.

The type 2 diabetic (T2D) model was established using a previously described method (Liao et al., 2017). In brief, the SD rats (n = 10) were fed with a high-fat diet (HFD) containing 66.5% normal chow, 20% sucrose, 10% lard, 2% cholesterol, and 1.5% cholate. After being fed with the HFD for 4 weeks, all rats were administered 25 mg/kg of streptozotocin (STZ) by intraperitoneal injection twice/week for 2 weeks. Rats treated with STZ and exhibiting a non-fasting blood glucose level ≥11.1 mmol/L were considered successful in establishing the T2D model. Next, the T2D rats were anesthetized with 40 mg/kg of pentobarbital sodium, and a full-thickness skin defect of 1 cm in diameter was created using a previously described method (Hur et al., 2017). Subsequently, the rats were randomly (random table method) divided into T2D skin-injured model and ACM groups (n = 5/group) and housed separately. Rats in the ACM group were treated daily with 100 μL ACM administered intradermally around the wound edges for 7 days, while model rats received an equal volume of serum-free medium. Normal rats (n = 5) were used as the negative control. On the 5th day after the last ACM treatment (the 12th day in total), all rats were euthanized with 100 mg/kg of pentobarbital sodium, and the wounds were harvested for further evaluation. All animals were selected at random for outcome assessment.

Tissues were collected and fixed in 4% paraformaldehyde for 24 h and then paraffin-embedded and sectioned into slices. Tissue sections were evaluated with hematoxylin and eosin (HE) staining and Masson staining, respectively. Finally, a double-blind histological examination was performed using an ortho-microscope (Zeiss, Germany) by two expert pathologists.

Total RNA from skin tissues was subjected to polyA-selected RNA-sequencing on the Illumina HiSeq X10 platform in a blinded manner. Using the DESeq2 package, RNA-seq analysis was carried out to determine the different gene expression (DEG) among three groups: normal vs. model and ACM vs. model. False discovery rate (FDR) of < 0.05 and fold change of ≥ 2 or ≤ 2 were the principles for DEG screening. Gene Ontology (GO) analysis was used to analyze the gene functions of the DEGs, and the Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis was used to target the DEGs’ enrichment pathway.

The skin wound sections were drenched in a citrate antigen retrieval solution (Beyotime Institute of Biotechnology, China) and heat-treated in a pressure cooker for 2 min, naturally cooled to RT, and washed with PBS buffer three times. Following incubation with 3% H₂O₂ for 10 min, the sections were blocked with 5% BSA for 30 min. The sections were then incubated overnight at 4 °C with primary antibodies against TNF-α, NF-κB, p-NF-κB, MAPK, p-MAPK, CXCL1, CXCL2, and CXCL8 at a dilution ratio of 1:200 . After washing three times with PBS, the sections were incubated with the secondary antibody at RT for another 2 h, followed by staining with DAB. The samples were observed using an ortho-microscope (Zeiss, Germany) in a blinded manner by the assessors.

All quantitative data were expressed as the mean ± standard deviation. GraphPad Prism version 9.0 (GraphPad Software, United States) was used for statistical analysis. The ANOVA was used to evaluate the significant differences among three independent groups, while the two-tailed paired sample Student’s t-tests were used to evaluate the significant differences between two groups. p < 0.05 was considered a statistically significant difference.

Flow cytometry analysis showed that ADSCs typically expressed CD73 and CD90 while lacking expression of CD45, CD19, and CD34 (Supplementary Figure S1), suggesting the successful isolation and culture of ADSCs in this study. The impaired skin wound healing in diabetic individuals is largely attributed to diabetic angiopathy, which is characterized by the dysfunction and impairment of the arteries throughout the body (Jin et al., 2022). We, therefore, investigated the effect of ACM on the angiogenesis and proliferation of vascular cells in vitro. After incubation with ACM for 24 or 48 h, the viability of HUVECs was significantly increased (Figure 1A), suggesting that ACM promoted HUVEC proliferation. After 2 days of continuous ACM incubation, HUVECs showed vascular-like morphological changes (Figure 1B), and the expression of genes associated with angiogenesis, including EGF, bFGF, VEGF, and KDR, was significantly upregulated after ACM treatment (Figure 1C), implying that ACM promotes HUVEC angiogenesis in vitro.

Based on the pro-angiogenic potential of ACM in vascular cells, we established a T2D skin wound model to further evaluate its therapeutic effects. As shown in Figures 2A, B, the skin wound healing rate of T2D rats was significantly improved by the continuous ACM treatment for 7 days compared to that of the model group. Moreover, increased tissue regeneration and decreased inflammatory infiltration were also observed in the ACM group compared to those in the model group (Figure 2C). Given the excellent performance of ACM on angiogenesis in vitro, we also investigated the beneficial effect of ACM on angiopathy in vivo. We found that the number of CD31⁺ cells and the expression of FGF-2 and VEGF were markedly increased in the ACM group compared to those in the model group (Figures 2D, E), suggesting that ACM could also improve angiopathy in vivo. Therefore, these data suggest that ACM accelerates T2D skin wound healing.

Given that excessive inflammation is a typical characteristic of skin wounds (Huang et al., 2022), we further analyzed the inflammatory genes in T2D skin wound tissues. As shown in Figure 3, the mRNA expression levels of TNF-α, IL-1β, IL-6, COX-2, IL-12, and IFN-γ were significantly increased in T2D skin wounds compared to those in normal skin tissues, indicating excessive inflammation. However, these levels were effectively decreased after ACM treatment compared to those in the model groups, suggesting that ACM could inhibit this excessive inflammation. Moreover, we found that ACM treatment reduced inflammatory cell infiltration. This included a reduction in both the CD3⁺ T cells and F4/80⁺ macrophages (Supplementary Figure S3). Furthermore, the expression of TNF-α, IL-1β, and IL-6 was significantly decreased in the ACM-treated groups (Figure 4). Taken together, these data suggest that ACM mitigates the inflammatory response in skin wounds of diabetic rats.

The potential molecular mechanism of ACM in accelerating T2D skin wound healing was further explored by RNA sequencing. Volcano plot analysis revealed differential expression of 10,655 genes was different between the normal and model groups (normal vs. model) and 5,287 genes between the ACM and model groups (ACM vs. model); a total of 4,269 genes were common to both comparisons (Figures 5A–C). GO annotation and pathway enrichment analysis showed that the upregulation of TNF and chemokine signaling was observed in the model group (compared with the normal group), while downregulation of TNF and chemokine signaling was clearly observed in the ACM group compared with the model groups (Figures 5D, E), suggesting that the potential molecular mechanism of ACM in T2D skin wound healing is by targeting the TNF and chemokine signaling pathway.

To confirm the RNA sequencing results, we further evaluated the protein expression of the main regulators in TNF and chemokine signaling. As shown in Figure 6, the TNF signaling-related proteins, including TNF-α, NF-κB, p-NF-κB, MAPK, and p-MAPK, and the chemokine signaling-related proteins, including CXCL1, CXCL2, and CXCL8, were all downregulated by ACM treatment in T2D skin wounds, suggesting that ACM accelerated T2D skin wound healing via the downregulation of TNF and chemokine signaling.