Extracellular Vesicle–Associated miR-ERIA Exerts the Antiangiogenic Effect of Macrophages in Diabetic Wound Healing

The prevalence of diabetes among Chinese adults has reached 12.8% (1), and the lifetime risk of diabetic foot ulcers is an alarming 34.0%. Diabetic foot ulcers are a major cause of disability and death in patients with diabetes (2). Wound repair relies on the coordinated efforts of various cells (3). Therefore, understanding the mechanisms underlying cellular interactions in wound microenvironments is essential for promoting diabetic cutaneous wound healing.

During the initial phase, macrophages polarize to an M1-like phenotype, promoting the synthesis and release of inflammatory factors. During the proliferation stage, they shift to an M2-like phenotype, secreting growth factors that support granulation tissue formation. However, under diabetic conditions, an imbalance in polarization and impaired macrophage function occur, coinciding with decreased wound angiogenesis (4,5), which may be a crucial pathophysiological change underlying diabetic wound healing. Accumulating evidence has suggested an important role for extracellular vesicles (EVs) derived from macrophages in mediating metabolic disorders by transporting miRNAs (6–8). We hypothesized that abnormal macrophage polarization and changes in the secretion profile of EVs disrupt the cutaneous microenvironment and drive the development of nonhealing diabetic wounds. This study explored the specific molecular mechanism underlying the abnormal regulation of the diabetic skin wound microenvironment from the novel perspective of vesicle-mediated cell interactions. The aim was to provide a theoretical basis for clinical treatment strategies to improve skin microenvironment homeostasis.

Microcirculation disorders are key causes of diabetic refractory cutaneous ulcers. Impaired vascular endothelial cell function contributes to nonhealing diabetic wounds (9,10); however, the underlying mechanism remains unclear. Recent studies have demonstrated that EVs participate in the pathogenesis and progression of wound angiogenesis (11). However, there is limited evidence regarding the involvement of miRNAs carried by EVs in interactions between macrophages and vascular endothelial cells that are specifically related to refractory diabetic cutaneous wound healing. Therefore, in this study, we investigated the interactions between macrophages and vascular endothelial cells by exploring the functional roles of advanced glycation end products pretreated with EVs (AGEs-EVs) in vitro and in vivo. We discovered a novel miRNA, miRNA enriched in AGEs-EVs (miR-ERIA), and explored the mechanism by which miR-ERIA regulates wound angiogenesis and wound healing in diabetes.

Human acute monocytic leukemia cells (THP-1) and human umbilical vein endothelial cells (HUVECs) were purchased from Procell Company. THP-1 were cultured in RPMI medium (Gibco) containing 10% FBS. Phorbol myristate acetate (PMA) (100 ng/mL) was added to induce macrophage differentiation. Cells were treated with 100 mg/L AGEs or BSA for 48 h. HUVECs were cultured in endothelial cell medium (ScienCell Research Laboratories) containing 5% FBS and 1% endothelial cell growth supplement. Based on our preliminary experiments and other researchers’ published reports, we used 5 and 25 μg/mL as low and high concentrations of EVs, respectively, for coincubation with HUVECs for 48 h.

EV isolation was performed through ultracentrifugation. Culture media or wound exudates were centrifuged (300g for 10 min and 3,000g for 20 min) to remove dead cells and cell debris. The supernatants were concentrated through 100-kDa ultrafiltration (Millipore) and then centrifuged (10,000g for 30 min). Supernatants were further purified by filtering through a 0.22-μm filter (Millipore). The media were transferred to new ultracentrifugation tubes and centrifuged (100,000g for 70 min) twice. All steps were conducted at 4°C. Finally, the pellets were resuspended in 100 μL of cold PBS. The expression of EV protein markers, including CD9 (1:1,000; Abcam), CD63 (1:1,000; Abcam), and TSG101 (1:1,000; Abcam), was detected via Western blotting.

EVs were labeled using a PKH26 kit (Sigma-Aldrich) and coincubated with HUVECs overnight. Samples were observed and analyzed using a confocal laser scanning microscope (Nikon). All procedures were conducted without light irradiation.

Total RNA from cells or EVs was isolated using TRIzol reagent (Takara). Total RNA and miRNA were reverse-transcribed into cDNA using PrimeScript RT Master Mix (Takara) and Mir-X miRNA First Strand and Synthesis (Takara) kits, respectively. Actin and U6 were used as internal references for mRNA and miRNA, respectively, and the 2^−ΔΔCt method was used to calculate relative gene expression. Primer sequences are listed in Supplementary Table 1. Western blotting was performed as previously described (12). The primary and secondary antibodies are listed in Supplementary Table 2.

When cells reached 100% confluence, wounds were created using sterile 200-μL pipette tips. Images were captured under a microscope at the indicated time intervals. Wound areas were analyzed by ImageJ software, and wound healing rates were calculated using the following equation: wound healing rate (%) = ([A₀ – A_n] / A₀) × 100. For the transwell migration assay, cells were resuspended in serum-free medium and loaded into the upper chamber in the transwell system (Corning). The migrated cells in five different fields of each sample were photographed under a microscope and analyzed by ImageJ.

Matrigel (Corning) was redissolved at 4°C overnight, and 10 μL of Matrigel per well was coated on microslides (ibidi GmbH) for 30 min in an incubator at 37°C. Cells were resuspended in serum-free endothelial cell medium, and 2 × 10⁴ cells in 50 μL medium were loaded and incubated at 37°C for 6 h. Images were captured under a microscope, and the total tube numbers and tube lengths were measured using ImageJ.

Total RNA from EVs was collected as described above. Secondary sequencing was performed by Shanghai OE Biotech Company. Sequencing was performed on an Illumina instrument, and clean reads were mapped to data in an miRNA database, while novel miRNAs were identified using miRDeep2 software. Significant differences in miRNA expression were indicated by fold changes ≥2 and P < 0.05.

Total RNA extracted from HUVECs treated with miR-ERIA mimics or the negative control was used to establish a DNA library, constructed by BGI according to the manufacturer’s protocol, followed by sequencing on a BGISEQ-500 platform. Expression of mRNAs with fold changes ≥2 and P < 0.001 was considered statistically different.

Cells grown in 24-well plates were transfected with luciferase vectors containing wild-type or mutant helicase with zinc finger 2 (HELZ2) after transfection with miR-ERIA mimics or the negative control. After 48 h of incubation, the Dual-Glo Luciferase Assay System Kit (Promega) was used to measure luciferase activity according to the manufacturer’s instructions.

RNA samples were extracted and detected using the Highly Sensitive MiRNA Northern Blot Assay Kit (Signosis). Images were captured using a chemi-imaging system (iBright CL1000; Invitrogen). The cy3-labeled miRNA probe and negative control probe were synthesized by GenePharma (Shanghai, China). RNA molecules with a random sequence were used as the negative control probe, and an miRNA fluorescence in situ hybridization (FISH) assay kit (GenePharma) was used according to the manufacturer’s instructions. Images were acquired under a confocal laser microscope.

All experiments were approved by the Animal Research Committee of Sun Yat-sen University and were divided into two parts. First, male Sprague-Dawley rats (7 weeks old, weighing 200–250 g) were purchased from Guangdong Medical Laboratory Animal Center and randomly divided into three groups after 1 week of adjustment feeding. A cutaneous wound was created as described previously (12). Control EVs (Con-EVs) or AGEs-EVs (200 μg) in 100 μL PBS or an equal volume of PBS were injected at an average of four sites around the wound. Second, male Sprague-Dawley rats (4 weeks old, weighing 120–150 g) were randomly assigned to normal and diabetic groups. Diabetic Sprague-Dawley rats were induced by treatment with 65 mg/kg streptozotocin (STZ), as previously described (13). Cutaneous wounds were made, and miR-ERIA agomir/antagomir or a negative control purchased from Ribobio Co., Ltd. (Guangzhou, China) was injected around the wound sites. Images of wound healing were obtained, and weight, blood glucose, and rate of death were recorded on days 0, 4, and 7 postwounding. Rats were sacrificed on day 7 postwounding, and wound specimens were acquired for subsequent experiments.

EVs were labeled with DiR dye (Invitrogen) or PKH26 dye (Sigma-Aldrich) and subcutaneously injected around the cutaneous wounds of Sprague-Dawley rats. The distribution of DiR-labeled EVs in rats was recorded using an in vivo imaging system (NightOWL II LB983) on days 1, 4, and 7 postwounding. Rats injected with PKH26-labeled EVs were sacrificed on day 7 postwounding, and the wound specimens were embedded on frozen slides. These slides were fixed with cold methanol for 10 min and washed three times with PBS for 5 min. Subsequently, they were stained with DAPI for 10 min, and images were obtained under a fluorescence microscope.

Rats subjected to different treatments were anesthetized by intraperitoneal injection of 50 mg/kg pentobarbital sodium on days 4 and 7 postwounding. A microfil working solution (20 mL) was perfused continuously at a rate of 3 mL/min into the hearts of rats after perfusion with 100 mL heparinized saline. Subsequently, the animals were held at 4°C overnight, and wound tissues were collected the next day for fixation with 4% paraformaldehyde. Wound vessels were observed using micro–computed tomography (micro-CT), and three-dimensional images were reconstructed using the CTVol program (Bruker). Vessel volumes in percentages were analyzed using ImageJ.

Data analyses were performed using GraphPad Prism 8.0, and results are presented as mean ± SD. Unpaired Student t test was used for comparisons between two groups, whereas one-way ANOVA was used for comparisons among groups. Statistical significance was set at P < 0.05.

The data sets and resources generated and/or analyzed in the current study are available from the corresponding author upon reasonable request.

Wound tissues from STZ-induced diabetic Sprague-Dawley and normal rats, as well as from patients with or without diabetes, were analyzed through hematoxylin-eosin (H-E) staining. The granulation tissue was thinner in the diabetic groups than in the nondiabetic groups (Fig. 1A). CD31 expression in wound specimens from rats and patients was assessed by immunohistochemistry (IHC) staining to evaluate wound angiogenesis (Fig. 1B). The microvessel density in the diabetic groups was significantly decreased. Induced nitric oxide synthase (iNOS) and CD163 were used as markers for M1 and M2 macrophages, respectively. The percentages of iNOS⁺ cells in wound tissues of diabetic rats on days 4 and 7 postwounding were increased, while the percentages of CD163⁺ cells were decreased in the diabetic groups compared with the nondiabetic groups (Fig. 1C). THP-1 cells were treated with 100 mg/L of phorbol myristate acetate to induce macrophages in vitro, and CD68 expression, a macrophage marker, was observed using immunofluorescence (IF) staining, confirming successful macrophage induction (Supplementary Fig. 1A and B). AGEs were used to construct in vitro models of diabetes as described previously (14). The expression of several inflammatory factors was detected using quantitative RT-PCR (RT-qPCR) to determine the macrophage phenotype following AGEs treatment. Interferon-γ (IFN-γ) and interleukin-4 (IL-4) were used to induce macrophages into M1 and M2 types, respectively. The results showed that IL-1β and tumor necrosis factor-α (TNF-α) in AGE-treated macrophages were upregulated (Fig. 1D). Additionally, IF staining showed that the expression of iNOS increased in the AGE-treated group (Fig. 1E). These results demonstrate that diabetic conditions lead to macrophage polarization toward an M1 phenotype and a reduction in wound angiogenesis.

To explore the relationship between macrophage dysfunction and the suppression of wound angiogenesis under diabetic conditions, we first assessed the impact of macrophages on the biological behavior of HUVECs using a coculture system. Migration and tube formation of HUVECs were suppressed after incubation with AGE-pretreated macrophages (Fig. 2A–E), while there was no significant effect on HUVEC proliferation and apoptosis (Supplementary Fig. 1C–G). To evaluate how macrophages modulate the biological function of HUVECs under diabetic conditions, we isolated macrophage-derived EVs and explored their effect on HUVECs. Conditioned media from macrophages were collected for EV isolation. The morphology of extracted pellets was observed under transmission electron microscopy as cup-shaped bilayer membrane structures, with diameters ranging from 30 to 200 nm (Supplementary Fig. 2A). Protein markers of EVs, such as CD63, CD9, and Tsg101, were detected by Western blotting, and an endoplasmic reticulum protein marker, Grp94, was found undetectable in EVs, which aligns with the conclusions of other studies (15) (Supplementary Fig. 2B). Nanoparticle tracking analysis showed that the average diameter of the pellets was ∼177.8 nm and the concentration was 2.71 × 10¹⁰ particles/mL (Supplementary Fig. 2C). Moreover, the production of EVs derived from macrophages treated with AGE-EVs or the BSA control (Con-EVs) was similar between the groups (Supplementary Fig. 2D). These results indicated that macrophage-secreted EVs were successfully isolated. Macrophage-derived EVs were labeled with PKH26 and incubated with HUVECs. Laser confocal microscopy showed a gradual uptake of EVs by HUVECs (Supplementary Fig. 2E). Based on our preliminary studies, low and high doses (5 μg/mL and 25 μg/mL, respectively) of AGEs-EVs and Con-EVs were incubated with HUVECs for 24 h. CCK-8 and EdU assays were used to measure HUVEC proliferation, while cell apoptosis was detected by annexin V-FITC/propidium iodide staining and flow cytometry. Macrophage-derived EVs had no significant effect on the proliferation or apoptosis of HUVECs under diabetic conditions (Supplementary Fig. 2F–J). To evaluate the migration of HUVECs, wound healing and transwell assays were performed. As shown in Fig. 2F and G, 25 μg/mL AGEs-EV–treated HUVECs exhibited the slowest wound healing rate at the 48-h time point, aligning with the results of the transwell assay (Fig. 2H). The tube formation assay results showed that the tube numbers and total tube lengths of HUVECs decreased after incubation with 25 μg/mL AGEs-EVs (Fig. 2I–J). Thus, migration and tube formation of HUVECs were suppressed by macrophage-derived EVs under diabetic conditions.

We investigated the effects of macrophage-derived EVs on wound angiogenesis and healing in vivo (Fig. 3A). First, the distribution of EVs in the rats was determined. DiR dye was used to label the EVs because of its low autofluorescence. Images were obtained using an in vivo small animal imaging system on days 1, 4, and 7 after EV injection (Fig. 3B). PKH26-labeled EVs were injected subcutaneously around the wounds of rats, and after 1 week, wound specimens were collected for IF assays. As shown in Fig. 3C, red fluorescence was observed at the wound sites. Seven-week-old male rats were used to explore the effects of macrophage-derived EVs on wound angiogenesis. The rats were divided into three groups (AGEs-EVs, Con-EVs, and PBS). The wound healing rates of the AGEs-EV group were distinctly lower than those of the other groups (Fig. 3D and E). Meanwhile, we recorded the weight and blood glucose levels of rats at different time points, and there were no significant differences between the groups (Supplementary Fig. 3A and B). We also measured serum ALT, AST, blood urea nitrogen, and creatinine levels in rats, revealing minimal hepatic or renal toxicity caused by EVs (Supplementary Fig. 3C). Wound specimens on day 7 postwounding were collected for H-E staining, and the length of wounds after AGEs-EV treatment was increased compared with the other two groups (Fig. 3F). A CD31 expression assay using IHC staining was conducted to evaluate wound angiogenesis. The microvessel densities of the three groups were calculated, and the results indicated fewer microvessels at the wound sites after AGEs-EV treatment (Fig. 3G). As shown in Fig. 3H, the vessel volumes of cutaneous wound tissues from the AGEs-EV group were lower than those from the other two groups. The vessels in the AGEs-EV group were more fragile and finer, and perivascular leakage was observed. Therefore, AGEs-EVs delay cutaneous wound healing by suppressing angiogenesis.

miRNAs, vital components of EVs, have been shown to participate in diabetic wound angiogenesis (11,16). AGEs-EVs and Con-EVs were isolated for small RNA sequencing. A total of 640 miRNAs were detected, and 27 showed differential expression with fold changes ≥2 and P < 0.05, including 22 new predicted miRNAs. Twenty-one miRNAs were upregulated, while six were downregulated (Fig. 4A). The expression of these miRNAs in macrophage cell lysates and secreted EVs was measured by RT-qPCR (Fig. 4B and C). The miRNA expression profile observed in macrophages did not exactly match that of macrophage-secreted EVs. In EVs, the differential fold changes of miRNA novel_153, miRNA novel_195, and miRNA novel_197 ranked in the top three, while the expression of miRNA novel_197 was higher and, thus, selected as the candidate miRNA for further experiments. miRNA novel_197, which is located on chromosome 1, was predicted using miRDeep2 software. The pre-miRNA was predicted to contain a hairpin structure (Supplementary Fig. 4), and the sequence of mature miRNA was 5′-UCCGGAGGAAGGUGGGGA-3′. We named it miR-ERIA. The expression of miR-ERIA was not upregulated in HUVECs treated with AGEs (Fig. 4D); however, miR-ERIA was significantly upregulated in AGEs-EVs (Fig. 4E). The above results indicate that AGEs can induce specific miRNAs, including miR-ERIA, to be encapsulated in EVs secreted by macrophages and then taken up by HUVECs. Wound exudates from 24 patients with or without diabetes were collected, and EVs were isolated by ultracentrifugation. miR-ERIA was significantly upregulated in diabetic wound exudate–derived EVs (Fig. 4F). The expression level of miR-ERIA in EVs from the serum of patients with diabetes were also higher than those in patients without diabetes (Fig. 4G). The clinical characteristics of the patients are presented in Supplementary Tables 3 and 4. Moreover, another 30 serum samples from patients with diabetes with or without foot ulcers were collected, and serum-derived EVs were isolated. Patients with diabetic foot ulcers were divided into two groups: low Wagner grade (<3) and high Wagner grade (≥3). The expression level of serum EV–derived miR-ERIA was significantly increased in patients with diabetic foot ulcers, with higher levels in the high–Wagner grade subgroup than in the low–Wagner grade subgroup (Fig. 4H). Northern blotting and miRNA FISH were performed to evaluate the expression and distribution of miR-ERIA in HUVECs. As shown in Fig. 4I and J, expression was higher in HUVECs treated with miRNA mimics than in the negative control. miR-ERIA is located in both the cytoplasm and nucleus but mainly in the cytoplasm. Moreover, HUVECs were transfected with miRNA mimics and negative controls, and their biological behaviors were assessed. Following miR-ERIA overexpression, the migrated areas in the wound healing assay and the number of migrated cells in the transwell assay were significantly decreased, indicating that HUVEC migration was suppressed. In the tube formation assay, the total tube lengths and numbers in the miRNA mimic group significantly decreased (Fig. 4K). Moreover, after treatment with the miRNA inhibitor and AGEs-EVs, the wound healing areas of the HUVECs showed the opposite effect. A trend of increase in migrated cells in the transwell assay and total tube lengths, as well as tube numbers in the tube formation assay, was observed after transfection with the miRNA inhibitor in HUVECs treated with AGEs-EVs (Fig. 4L). The above results indicate that miR-ERIA could suppress HUVEC migration and tube formation.

The potential target genes of miR-ERIA were predicted using TargetScan 7.2 and miRDB (Supplementary Fig. 5A). To further confirm the candidate target genes, we overexpressed miR-ERIA in HUVECs and collected total RNA from cell lysates for transcriptome sequencing. A total of 561 differentially expressed mRNAs (145 upregulated and 416 downregulated) were detected with fold changes ≥2 and P < 0.001 (Fig. 5A). Among the downregulated genes, 358 were identified through database searches. We combined these 358 genes with the previously predicted target genes, and 5 candidate genes, DPP4, EDN1, KIAA0319, CXCL16, and HELZ2, were further validated (Fig. 5B). HELZ2 mRNA was downregulated in HUVECs treated with miRNA mimics, whereas the expression levels of the other four genes were not significantly changed (Fig. 5C). The protein level of HELZ2 was accordingly decreased in HUVECs treated with miRNA mimics (Fig. 5D). Bioinformatics analysis indicated that miR-ERIA can bind to the coding region of HELZ2 (Fig. 5E), and a dual luciferase reporter assay was performed for validation. Luciferase reporters containing wild-type or mutated sequences of HELZ were transfected into 293T cells. When cells were transfected with miRNA mimics, luciferase activity in the wild-type group significantly decreased, whereas the opposite effect was observed in the mutant group (Fig. 5F). These results indicate that HELZ2 is a direct target of miR-ERIA. To investigate whether miR-ERIA suppresses HUVEC migration and tube formation by targeting HELZ2, we transfected small interfering HELZ2 (si-HELZ2) into HUVECs. As shown in Fig. 5G, when HELZ2 expression was suppressed in HUVECs by siRNA transfection, cell migration and tube formation were both significantly affected. Conversely, HELZ2 overexpression reversed the suppression of cell migration and tube formation by miR-ERIA to some extent (Fig. 5H). In general, these results indicate that miR-ERIA suppresses the migration and tube formation of HUVECs by targeting HELZ2 mRNA.

Animal studies were performed to investigate whether miR-ERIA plays a significant role in cutaneous wound angiogenesis (Fig. 6A). As shown in Fig. 6B and C, wound healing was significantly delayed after treatment with the miRNA agomir in the nondiabetic group on day 7 postwounding. Wound closure rates recovered to some extent in the diabetic group treated with the miRNA antagomir compared with those in the negative control group. The wound lengths of all groups were calculated, and wider wounds were observed in the nondiabetic group treated with the miRNA agomir. The wounds in the diabetic group treated with the miRNA antagomir narrowed on day 7 postwounding (Fig. 6D). Next, we examined the protein expression of CD31 in the wounds of rats using IHC. In the nondiabetic group, rats injected with the miRNA agomir exhibited a lower microvessel density at the wound site than that in the negative control group. In the diabetic group, the microvessel density at the wound site in the miRNA antagomir group was higher than that in the negative control group (Fig. 6E). Microvessel morphology analysis was performed as described above. The vessel volume of cutaneous wound tissues was decreased in the miRNA agomir group compared with that in the negative control group. Conversely, it was slightly increased in the diabetic group treated with the miRNA antagomir (Fig. 6F). These findings suggest that miR-ERIA can negatively affect the biological function of HUVECs in vitro and suppress cutaneous wound angiogenesis, thereby contributing to a significant delay in wound healing in diabetic rats.

In this study, we found that macrophages polarized to the M1 type after AGE treatment and their secreted EVs could regulate the biology of vascular endothelial cells in vitro, corresponding to a decrease in wound angiogenesis and delayed wound healing in vivo. We identified a novel miRNA enriched in macrophage-derived AGEs-EVs, miR-ERIA, that suppressed the migration and tube formation of HUVECs by targeting HELZ2. Suppression of miR-ERIA increased wound angiogenesis and accelerated wound closure in diabetic Sprague-Dawley rats. Thus, our results indicate that macrophage-derived EVs carrying miR-ERIA play a significant role in diabetic wound healing.

Wound healing is a complex process in which various cells play different roles and collaborate (17,18). Angiogenesis is a limiting factor in wound healing, and the integrity of endothelial cell function, balance between proangiogenic factors and antiangiogenic factors, and effective coordination between endothelial cells and surrounding cells are crucial for the formation and maturation of new blood vessels (9,19). Studies have shown that under diabetic conditions, endothelial cells exhibit dysfunction and loss of integrity. An imbalance between proangiogenic and antiangiogenic factors involving other reparative cells in the microenvironment ultimately leads to impaired wound angiogenesis (20). Our current study shows that under diabetic conditions, macrophages polarize toward an M1 phenotype, leading to increased levels of proinflammatory cytokines while inhibiting endothelial cell function. However, an imbalance in the M1/M2 ratio hinders the transition from the inflammatory phase to the proliferative phase during the wound healing process, thus affecting wound closure (21,22). Chemokines and inflammatory factors, including CXCL1, CCL2, and NLRP3 inflammatory pathways, are involved in immune cell recruitment and macrophage polarization, ultimately impairing the repair process (23–25). Many studies have focused on promoting wound healing by enhancing the activity and numbers of M2-like macrophages (26,27). However, the continuous presence of proinflammatory M1-type macrophages without a transition to an anti-inflammatory phenotype does not favor wound healing. Recent research has shown that EVs derived from M2-type macrophages carrying factors such as CCL22, CCL24, and MFGE8 can effectively stimulate the polarization of M1 macrophages toward an M2 phenotype, indicating that enhancing the proangiogenic effects mediated by M2-type macrophages while reducing the inflammatory response induced by M1-type macrophages ultimately promotes wound healing. This suggests an effective therapeutic approach to various diseases associated with imbalances in pro- and anti-inflammatory immune responses (28).

Macrophages participate in angiogenesis not only by secreting cytokines and directly contacting endothelial cells but also by secreting different EVs that interact with vascular endothelial cells. In a model of coronary artery disease, EVs secreted by macrophages stimulated with oxidized LDL carried miR-4306 to target the VEGFA/Akt signaling pathway in endothelial cells, inhibiting their ability to form tubes (29). EVs have emerged as important players in the pathogenesis and progression of diabetes and its complications. They can carry various bioactive molecules, including miRNAs, exerting significant effects on target cells and tissues (30–33).

To our knowledge, this study is the first to demonstrate that under diabetic conditions, M1-type macrophage-derived EVs affect the migration and tube formation ability of vascular endothelial cells, thereby inhibiting wound angiogenesis and delaying cutaneous wound healing. We identified a novel miRNA enriched in AGEs-EVs, miR-ERIA, that suppresses wound angiogenesis by targeting HELZ2 in HUVECs. The HELZ2 protein is a transcriptional coactivator for peroxisome proliferator–activated receptor α (PPARα), which cooperates with PPARα to play a role in transcriptional regulation within nuclei (34,35). PPARα has been shown to participate in lipid metabolism and angiogenesis. Studies have shown that the PPARα agonist fenofibrate can improve endothelial progenitor cell function and promote angiogenesis through the NLRP3 inflammasome pathway (36). Moreover, PPARα and PPARγ increase the production of vascular endothelial growth factor by promoting the phosphorylation of NOS and Akt in endothelial cells, thus enhancing endothelial cell tube formation and angiogenesis in vivo, as observed in mouse corneal neovascularization (37). Significant progress has been made in diabetic wound healing studies. For example, amphibian-derived peptides and insulin are well studied for promoting wound healing by improving repair cell functions in diabetic skin environments (38–42). In this study, we have provided a new strategy to accelerate diabetic wound closure based on cell interactions between macrophages and vascular endothelial cells via EVs carrying therapeutic miRNA. Engineered EVs may offer a potential therapeutic method for promoting diabetic wound closure. For example, exosomes produced following miRNA-126 overexpression in human synovial mesenchymal stem cells enhance the biological functions of human dermal microvascular endothelial cells, promoting skin wound healing in diabetic rats (43). Moreover, EVs carrying certain noncoding RNAs, combined with therapeutic hydrogel, are effective for chronic wound healing (44–46).

However, there are certain limitations in this study. Given that EV isolation requires a large amount of cell supernatant and the isolation and cultivation of primary macrophages are challenging, THP-1–induced macrophages were used in this study. We have confirmed that miR-ERIA suppresses the biological functions of HUVECs by targeting HELZ2 under diabetic conditions, but further studies are needed to focus on HELZ2 downstream signaling pathways. The number of clinical cases in this study was limited. It is necessary to further increase clinical samples and follow-up time to clarify the relationship between miR-ERIA and diabetic cutaneous wound healing.

In conclusion, we identified a novel miRNA enriched in macrophage-derived EVs under diabetic conditions, called miR-ERIA, which is involved in diabetic cutaneous wound healing. miR-ERIA can inhibit the migration and tube formation ability of vascular endothelial cells by targeting HELZ2, leading to the suppression of wound angiogenesis and diabetic refractory wound healing. These results suggest that when seeking to improve endothelial cell function, it is necessary to disrupt abnormal interactions between macrophages and vascular endothelial cells. Induction of phenotypic transformation in macrophages and use of genetic engineering techniques to selectively edit the composition of EVs secreted by macrophages may be effective in modifying their negative regulatory effect on vascular endothelial cell function.

Acknowledgments. The authors thank eBioart for help with pattern diagram making, Editage for English-language editing, and Yangfu Lv for encouragement and assistance in this study.

Funding. This study was supported by National Natural Science Foundation of China grants 82200902 and 322171185, Guangdong Clinical Research Center for Metabolic Diseases grant 2020B1111170009, Guangzhou Key Laboratory for Metabolic Diseases grant 202102100004, National Natural Science Foundation of China Excellent Young Scientists Fund grant 82222014, the Key Laboratory of Human Microbiome and Chronic Diseases (Sun Yat-sen University), and Ministry of Education, China, Guangzhou Science and Technology Program grant 2023A03J0706.

Duality of Interest. No potential conflicts of interest relevant to this study were reported.

Author Contributions. T.Z. and K.S. designed and performed the study and wrote the manuscript. L.M. and X.Ho. performed the experiments and collected the data. X.He. analyzed the data and interpreted the results. W.L. designed the experiments and revised and approved the final version of the manuscript. S.C. and L.Y. are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and accuracy of the data analysis.