Polyethylene Glycol Loxenatide speeds diabetic wound healing by reducing inflammation and improving blood vessel repair cells

To evaluate the effect of PEG-loxe on systemic metabolism, body weight data were collected from the initiation of treatment, and ITTs and OGTTs were conducted at 5 and 6 weeks, respectively. Diabetic mice exhibited increased body weight, while PEG-loxe treatment resulted in a slight reduction in weight (Figure 1A). Insulin tolerance tests (ITTs) and oral glucose tolerance tests (OGTTs) revealed impaired glucose clearance in db/db mice, as evidenced by elevated blood glucose levels at 60 and 120 min post-challenge (p < 0.01 vs. db/m controls). PEG-loxe treatment markedly improved insulin sensitivity, reducing the area under the curve (AUC) (p < 0.001; Figure 1B–E). These results suggest that PEG-loxe effectively improved the glucose metabolism of diabetic mice.

To investigate the role of PEG-loxe in diabetic wound healing, a wound model was established by creating dorsal skin wounds in db/db and control mice. Wound closure rates were measured every other day up to day 10 [17]. Diabetic mice exhibited delayed wound healing compared to the control group, while PEG-loxe treatment notably accelerated wound closure (Figure 1F,G). Hypoxia-inducible factor-1 alpha (HIF-1α), vascular endothelial growth factor (VEGF) and stromal cell-derived factor-1 (SDF-1) are critical mediators of vascular regeneration. The HIF-1α/VEGF/SDF-1 pathway has been shown to facilitate EPC mobilization and accelerate neovascularization [18]. HIF-1α is essential for the expression of SDF-1, which in turn promotes the homing of EPCs to ischemic regions [19]. In diabetic conditions, circulating levels of EPCs and plasma concentrations of SDF-1 are reduced [20]. Animal and in vitro studies have demonstrated that the HIF-1α/VEGF/SDF-1 axis exhibits impaired responsiveness in diabetic contexts, which subsequently hampers neovascularization [18,21]. Consistent with prior studies, the levels of HIF-1α, VEGFA, and SDF-1 in diabetic wound tissues were significantly lower than in controls but were notably upregulated following PEG-loxe treatment (Figure 1H).

Using Olink proteomics, we analyzed serum protein expression during wound healing. A heatmap of all differentially expressed proteins is presented in Figure 2A. We identified a total of four differentially expressed proteins between the db/db+PEG-loxe and untreated db/db groups: Yes1 and Itgb1bp2 were downregulated, while IL-5 and Adam23 were upregulated in the PEG-loxe-treated group (Figure 2B). Notably, glucagon levels were significantly elevated in the db/db group compared to the control group. This upregulation was attenuated by the administration of PEG-loxe, which is consistent with the drug's pharmacological mechanism of action in lowering glucagon levels (Figure 2C).

To gain deeper insights into the systematic inflammatory alteration, we compared the expression levels of inflammation-related proteins across the three groups (Figure 2C). The results demonstrated that levels of key inflammatory factors, such as CCL2 (MCP-1), CCL20 (MIP-3α), and ERBB4, were elevated in diabetic mice compared to the normal control group. This is aligned with previous findings in db/db mice, indicating dysregulation of the inflammatory response [22,23]. Notably, most pro-inflammatory proteins in the db/db group were upregulated, and were downregulated following PEG-loxe treatment, although some changes were not statistically significant.

These findings suggested that PEG-loxe may accelerate wound healing in diabetic mice, likely by promoting neovascularization and modulating the inflammatory response toward a more regulated state.

EPCs play a vital role in neovascularization and wound healing. To evaluate the effect of PEG-loxe on neovascularization, we conducted in vitro experiments to assess EPC function. To simulate the hyperglycemic and lipotoxic environment characteristic of diabetes, EPCs were cultured in high-glucose and high-fatty-acid conditions (HG/FA group). We assessed tube formation and migration using Matrigel and transwell assays, respectively. The results indicated that exposure to high glucose and high fatty acid levels significantly impaired tube formation and diminished migratory abilities compared to untreated control cells (Figure 3A–D). However, when treated with PEG-loxe (HG/FA+PEG-loxe group), both tube formation and migration capacities were partially restored, suggesting that PEG-loxe mitigates the effects of glucotoxicity and lipotoxicity, thereby preserving EPCs' neovascularization potential.

EPC dysfunction in diabetes is associated with reduced nitric oxide (NO) bioavailability, primarily attributed to endothelial nitric oxide synthase (eNOS) uncoupling [24]. Phosphorylation of eNOS at Ser1177 represents an important post-translational modification required for its enzymatic activation. In the present study, EPCs exposed to high-glucose and -fatty-acid (HG/FA) conditions exhibited significantly decreased NO levels in culture supernatants compared to the control group (Figure 3E), a finding consistent with eNOS uncoupling. Notably, treatment with PEG-loxe markedly restored NO production in HG/FA-stimulated EPCs. However, Western blot analysis revealed no statistically significant differences in the phosphorylated eNOS (Ser1177)/total eNOS ratio among experimental groups (Figure 3F,G). These data indicated that PEG-loxe ameliorates impaired NO bioavailability in diabetic EPCs under glucolipotoxic stress, potentially through mechanisms independent of Ser1177 phosphorylation-dependent eNOS activation.

The elevated ROS can lead to eNOS uncoupling, promoting a cycle of oxidative stress and endothelial damage [25]. Mitochondria serve as the primary source of cellular ROS, and an increase in ROS within mitochondria can lead to a loss of mitochondrial membrane potential. To explore the role of mitochondrial ROS in EPC dysfunction, we employed MitoSOX staining to assess mitochondrial superoxide production in EPCs. Our results demonstrated that exposure to high glucose and fatty acids (HG/FAs) significantly increased mitochondrial ROS production. In contrast, PEG-loxe treatment significantly reduced HG/FA-induced ROS accumulation (Figure 4A,C). Furthermore, we observed that the mitochondrial membrane potential was significantly diminished under HG/FA conditions, but it was restored upon PEG-loxe treatment, as assessed by JC-1 staining (Figure 4B,D).

A Seahorse assay was employed to evaluate whether PEG-loxe could prevent mitochondrial dysfunction in EPCs. The results indicated that mitochondrial respiration rates, including both basal and maximal respiration, were significantly reduced in the HG/FA group. However, treatment with PEG-loxe enhanced the mitochondrial oxygen consumption rate (OCR) in EPCs (Figure 4E–I). These findings, alongside the restoration of mitochondrial membrane potential, suggest that oxidative stress impairs mitochondrial respiratory function. Additionally, PEG-loxe treatment effectively preserved mitochondrial membrane potential and reversed the impairment of mitochondrial respiration.

Building on the results mentioned above, maintaining mitochondrial homeostasis is essential for the overall health of EPCs. Autophagy is a crucial cellular process that plays a significant role in regulating mitochondrial quality. Impaired mitochondria are tagged and then delivered to lysosomes for degradation, ensuring that the quantity and quality of mitochondria are preserved [26]. To evaluate the induction and progression of autophagy, we analyzed the levels of the proteins LC3 || and p62. In control EPCs, the baseline levels of LC3 || and p62 were low; however, these proteins were significantly increased in response to HG/FA treatment (Figure 5A,C,D), consistent with autophagic flux impairment. In comparison to the HG/FA group, PEG-loxe treatment resulted in a reduction in p62 expression and a tendency toward lower levels of TOM20, a mitochondrial membrane protein (Figure 5B,E). This indicates improved autophagic flux and mitochondrial turnover.

To further investigate the interaction between mitochondria and lysosomes in EPCs, we performed a colocalization assay using MitoTracker and LysoTracker. The yellow dots in the merged images indicate the fusion of red mitochondria with green lysosomes (Figure 5F,G). The results demonstrated impaired fusion of mitochondria and lysosomes under HG/FA conditions. In contrast, treatment with PEG-loxe led to an increase in the number of mitolysosomes, suggesting enhanced mitophagy. Western blot analysis confirmed that, in the HG/FA group, levels of LC3 || and p62 levels did not further accumulate when treated with bafilomycin A1 (Figure 5H), a lysosomal inhibitor. However, PEG-loxe treatment reversed this effect, indicating a restoration of autophagic activity.

Collectively, PEG-loxe treatment partially restored autophagic flux and improved mitochondrial quality control that were impaired by glucolipotoxicity.

Male 8-week-old db/db mice for a model of type 2 diabetes mellitus and age-matched male db/m mice for the control group were purchased from GemPharmatech Co., Ltd. (Nanjing, China). Mice were housed in a temperature-controlled environment (22–24 °C) with a 12 h light/dark cycle and ad libitum access to standard chow and water. The study was approved by the Laboratory Animal Welfare Ethics Committee of the Peking Union Medical College Hospital (XHDW-2023-145).

Insulin tolerance tests (ITTs) and oral glucose tolerance tests (OGTTs) were performed at 5 and 6 weeks after initiating PEG-loxenatide treatment, respectively. For ITTs, mice were fasted for 8 h, and then injected intraperitoneally with 1 IU/kg body weight insulin (i.p.).

Blood glucose levels were measured from the tail tip at 0, 15, 30, 60, and 120 min post-injection using a glucometer. For OGTTs, mice were fasted overnight for 16 h and then orally gavaged with 1 g/kg body weight of glucose dissolved in water. Blood glucose was measured at 0, 15, 60, and 120 min.

After one week of adaptation, mice were divided into three groups: (1) individuals of the db/m strain as a control group; (2) untreated db/db mice; and (3) db/db mice treated with PEG-loxe (0.03 mg/kg/week, intraperitoneal injection). The drug was administered for 12 weeks before establishing a wound model. On the day of wound creation, mice were anesthetized with 3% isoflurane, and their dorsal fur was shaved. A full-thickness, circular wound with a 6 mm diameter was generated on the dorsum using a sterile punch biopsy. Afterward, the mice were housed individually in separate cages.

Photographs of the wounds were taken every two days starting from the day the wounds were created. The wound area was quantified using ImageJ software (version 1.54g), and the wound closure rate was calculated as follows: wound closure rate (%) = (An − A0)/A0 × 100 (A0 = the wound area on the day of wound creation; An = the remaining wound area on the respective day).

Mice were euthanized on day 10 and the full-thickness wound tissues along with a rim of surrounding unwounded skin margins were excised. Tissue samples were quickly frozen in liquid nitrogen and then transferred to −80 °C for storage.

Blood samples were collected via cardiac puncture and centrifuged at 3000× g for 15 min to isolate serum. Serum proteins from control, diabetic, and PEG-loxe-treated diabetic groups were analyzed using the Olink Mouse Exploratory Panel (Biotree, Shanghai, China). Normalized protein expression (NPX) values were log2-transformed, and differentially expressed proteins were identified using the DEGseq package (version 1.61.0) with a significance threshold of p < 0.05.

Human peripheral blood-derived endothelial progenitor cells (EPCs) were purchased from FuHeng Biology Co., Ltd. (Shanghai, China). EPCs were cultured in EGM-2 BulletKit medium (Lonza, Basel, Switzerland) supplemented with recombinant human rhEGF, rhFGF-B, rhVEGF, rhIGF-1, ascorbic acid, heparin, 5% fetal bovine serum, and 1% penicillin/streptomycin. The cells were maintained at 37 °C in a humidified incubator with 5% CO₄. To mimic diabetic conditions, EPCs were exposed to high glucose (30 mM) and palmitic acid (100 μM) (HG/FA group) for 24 h. For intervention, cells were co-treated with 250 nM PEG-loxe (HG/FA+PEG-loxe group).

The concentration of nitrite, an indicator of nitric oxide (NO) production, was determined using the Griess reaction. The nitrite detection kit (Beyotime, Shanghai, China) was used according to the manufacturer's instructions. Briefly, 50 µL of supernatant or sodium nitrite (NaNO₂) standard was mixed with 50 µL of Griess Reagent I and 50 µL of Griess Reagent II in a 96-well plate. After a 10 min incubation in the dark, the NO concentration was quantified by measuring the absorbance at 540 nm using a microplate reader. The NO content was calculated based on the optical density (OD) values, with each group normalized to the control group.

EPCs were pretreated for 24 h before the migration assay. Cells were then collected and resuspended in serum-free EBM-2 medium. A total of 2 × 10⁴ EPCs were added to the upper chamber of a 24-well transwell plate (8.0 µm pore size; Corning, New York, NY, USA), and 500 μL of EGM-2 medium containing 5% FBS was added to the lower chamber. Cells were incubated at 37 °C for 8 h. After fixation with paraformaldehyde, the cells were stained with crystal violet solution (Beyotime, Shanghai, China). Images of cells that migrated to the lower chamber were captured using an inverted light microscope (Nikon, Tokyo, Japan), and the number of migrating cells was counted using ImageJ.

For the tube formation assay, 50 μL Matrigel (Corning, New York, NY, USA) was added to the wells of a 96-well plate and incubated at 37 °C for 30 min for polymerization. EPCs (1.5 × 10⁴ cells per well) were then seeded onto the Matrigel. After 6 h of incubation, images of tube-like structures were captured, and the length of the capillary-like structures was measured using ImageJ software.

Mitochondrial ROS levels were quantified using MitoSOX™ Red (Thermo Fisher Scientific, Inc., Waltham, MA, USA), a fluorescent dye specific for mitochondrial superoxide. EPCs were stained according to the manufacturer's instructions, and fluorescence images were acquired using a confocal laser scanning microscope (Nikon, Tokyo, Japan).

The mitochondrial membrane potential was examined using JC-1 dye (Beyotime, C2003S), which is sensitive to changes in mitochondrial membrane potential. EPCs were incubated with the JC-1 working solution at 37 °C for 20 min. After treatment, cells were examined by fluorescence microscopy (Nikon AXR inverted confocal microscope). For quantitative analysis, three random fields (1000× magnification) were selected for each group, and the ratio of red to green fluorescence (indicative of mitochondrial membrane potential) was measured and analyzed using ImageJ software.

The colocalization of mitochondria and lysosome assay was performed as previously described [61]. In short, EPCs were stained with MitoTracker Deep Red (Invitrogen, Carlsbad, CA, USA, M22426) at a final concentration of 250 nM and LysoTracker Green (Beyotime, Shanghai, China, C1047S) at a final concentration of 50 nM according to the manufacturer's instructions, and then photographed using a Nikon AXR confocal laser scanning microscope to assess the colocalization of mitochondria and lysosomes.

The oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were measured using an Seahorse XF96 Analyzer (Agilent, Santa Clara, CA, USA). The experiments were conducted at 37 °C and pH 7.4, using Seahorse XF DMEM medium supplemented with glucose, pyruvate, and glutamine as the detection solution, following the manufacturer's instructions. All respiratory parameters were corrected for non-mitochondrial respiration and background signals by adding 1.5 µM oligomycin, 2 µM FCCP, and 0.5 µM rotenone/antimycin A (Rot/AA).

Total protein was extracted from skin tissue or EPCs using RIPA buffer containing protease inhibitors, PMSF, and phosphatase inhibitors. Equal amounts of protein (15–20 µg) were loaded onto a 4–20% SDS-PAGE gel (Boyi, Changzhou, China) and then transferred to PVDF membranes. After blocking with protein-free rapid blocking solution (Servicebio, Beijing, China) for 15 min, the membranes were incubated overnight at 4 °C with primary antibodies against the target proteins HIF-1α (CST, Beverly, MA, USA, 1:1000 dilution), VEGFA (Abcam, Cambridge, UK, 1:1000 dilution), SDF1/CXCL12 (CST, 1:1000 dilution), eNOS (CST, 1:1000 dilution), phosphor-eNOS (CST, 1:1000 dilution), p62 (CST, 1:1000 dilution), LC3A/B (CST, 1:1000 dilution), and β-actin (CST, 1:1000 dilution). The membranes were then incubated for 1 h at room temperature with HRP-conjugated secondary antibodies (Easybio, Zhenjiang, China). Protein bands were detected using a chemiluminescence (ECL) method and band intensities were quantified using ImageJ.

Data were displayed as mean ± SEM. Comparisons between the groups were analyzed by unpaired Student's t tests. Comparisons among multiple groups were analyzed by one-way analysis of variance (ANOVA) followed by Tukey's post hoc test. The two-way ANOVA was used for analyzing wound closure followed by Dunnett's post hoc test. A p-value < 0.05 was considered statistically significant.