The Ethics Review Committee of the Affiliated Hospital of Youjiang Medical University for Nationalities approved all of the experiments involving animals (YYFY-LL-2023-084). The study is reported according to the Animal Research: Reporting In Vivo Experiments (ARRIVE) guidelines. All processes were conducted strictly complying with the relevant guidelines. Mice were housed in a specific pathogen-free facility at 22 $\pm$ 2 °C and 55 $\pm$ 10% humidity. All mice were exposed to a 12-h light/dark cycle and had ad libitum access to standard chow and water.

C57BL/6J mice (8–10 weeks old; 17–18 g; male) were provided by the Guangxi Medical University Laboratory Animal Centre (Nanning, China). For diabetes induction, mice were administered a high-fat diet for 4 weeks, followed by an intraperitoneal injection of streptozotocin (STZ; HY-13753, MCE, Monmouth Junction, NJ, USA,50 mg/kg) for five consecutive days [15]. When the fasting blood glucose levels were $>$300 mg/dL, mice were considered to have been successfully induced with diabetes (DM; n = 21 per group). Mice fed a standard diet were used as controls (Control; n = 9 per group). For the wound healing experiments, a full-thickness excisional wound (8 mm in diameter) was created using a sterile biopsy punch on the dorsal surface of each mouse (Miltex, Inc., York, PA, USA) under anesthesia (1% sodium pentobarbital; P3761, Sigma-Aldrich, Burlington, MA, USA, 50 mg/kg).

To investigate the involvement of NETs in diabetic wound healing, nine control and nine DM mice were used. The wounds were photographed on designated days (1, 3, 7, and 14) after wounding. Mice were euthanized by cervical dislocation on days 3, 7, and 14 after wounding, and the wound tissues were harvested for subsequent analysis.

To explore the potential of Ro 106-9920 in improving wound healing in diabetic mice, DM mice were further divided into two groups, which received dimethyl sulfoxide (DMSO) (DM+vehicle; n = 6 per group) and Ro 106-9920 (5 mg/kg) (MCE, Monmouth Junction, NJ, USA) [16] (DM+Ro 106-9920; n = 6 per group), respectively. The subcutaneous injections were administered at multiple sites around the wounds. At 1, 3, 7, and 14 d after wounding, the blood glucose levels and weight of each mouse were assessed, and the wounds were photographed. The mice were euthanized by cervical dislocation 14 d after wounding for harvesting the wound tissues. All photographed wounds were collected and subsequently analyzed using the ImageJ software (Version number 1.52m, NIH, Bethesda, MD, USA).

Mouse neutrophils were isolated as previously described [17]. The harvested neutrophils were purified using a discontinuous Percoll gradient (62%/81%) and resuspended in RPMI-1640 medium (Gibco, Waltham, MA, USA) for further experiments.

The purity of neutrophil samples was assessed using flow cytometry analysis. The samples were incubated with anti-CD11b (12-0112-82, Thermo Fisher Scientific, Waltham, MA, USA) and anti-Ly6G (11-5931-82, Thermo Fisher Scientific) antibodies for 30 min at 4 °C. Subsequently, cells were washed with phosphate-buffered saline (PBS) and fixed in 2% paraformaldehyde (PFA, HY-Y0333, MCE, Monmouth Junction, NJ, USA) for 15 min at 4 °C. Finally, cells were washed and resuspended in 150 µL PBS before analysis on a BD FACSCanto II (BD Biosciences, Franklin Lakes, NJ, USA). Data were processed using FlowJo (version 10.4.2; BD Biosciences). Singlets were selected from the Forward Scatter Height (FSC-H) versus Forward Scatter Area (FSC-A) dot plot in the debris exclusion gate (gate 1), while in the singlet gate, CD11b and Ly6G double-positive events were gated to detect neutrophils.

Mouse neutrophils (1 $\times$ 10⁶) were seeded in each well of six-well plates with RPMI-1640 medium (Gibco, Waltham, MA, USA). The neutrophils were induced with phorbol 12-myristate 13-acetate (PMA; Sigma-Aldrich, Burlington, MA, USA) at a dose of 100 ng/mL for 3–5 h to trigger NET formation. Mouse RAW264.7 macrophages were obtained from ATCC (TIB-71; Manassas, VA, USA) and seeded at a density of 2 $\times$ 10⁵ cells per well into Transwell inserts (0.4 µm pore size; Corning, Corning, NY, USA) containing Dulbecco’s Modified Eagle Medium (Gibco). The inserts were placed in the six-well plates containing the neutrophils, with or without PMA treatment, for a 12-h coculture at 37 °C. Following coculturing, the conditioned medium (CM) was collected for subsequent analyses. The CM harvested from the PMA-stimulated coculture system was defined as neutrophil-macrophage coculture model (NET-CM).

To investigate the effect of NET formation on the biological functions of keratinocytes and fibroblasts, cells were treated with NET-CM, whereas controls were treated with CM harvested from the non-PMA-stimulated coculture system.

To determine whether Ro 106-9920 can suppress NET formation (thereby attenuating NET-induced injury of fibroblasts, keratinocytes, and endothelial cells), the PMA-stimulated coculture system was further treated with Ro 106-9920 (2.5 µM) [16], the CM of which was defined as (NET+Ro 106-9920)-CM. Next, mouse fibroblasts (NIH/3T3) (CRL-1658; ATCC), keratinocytes (HaCaT) (T9145; Cell Lines Service, Eppelheim, Germany), and endothelial cells (HUVEC) (PCS-100-010; ATCC) were divided into three groups: control, NET-CM, and (NET+Ro 106-9920)-CM. Cells were treated with NET-CM or (NET+Ro 106-9920)-CM before functional analysis; those treated with CM harvested from the non-PMA-stimulated coculture system were used as the control.

All culture media contained 1% penicillin-streptomycin and 10% fetal bovine serum. All cells were incubated in a humidified incubator at 37 °C and 5% CO₂. All cell lines were validated through negative testing of mycoplasma and STR profiling.

Mouse neutrophils (1 $\times$ 10⁶) were seeded on coverslips before treatment. Treated neutrophils were fixed and permeabilized before being blocked with 5% bovine serum albumin for 1 h. Next, neutrophils were incubated overnight at 4 °C with a primary antibody against citrullinated histone H3 (CitH3; ab281584, Abcam, Cambridge, UK) or NF-$\kappa$B (AF1234, Beyotime, Shanghai, China). After washing thrice with PBS, the neutrophils were incubated with an Alexa Fluor 488-conjugated secondary antibody (A-11008, 1:1000 dilution; Invitrogen, Waltham, MA, USA) for 1 h at room temperature in the dark. Nuclei were counterstained with 4^′,6-diamidino-2-phenylindole (DAPI) for approximately 5 min. Finally, fluorescence images were obtained using a TCS SP8 confocal microscope (Leica Microsystems, Wetzlar, Germany). The mean fluorescence intensity of CitH3 and NF-$\kappa$B from each group was quantified using FIJI/ImageJ (1.52m, NIH, Bethesda, MD, USA).

ELISA was performed to quantify the levels of interleukin (IL)-1$\beta$ and IL-18 in the supernatants collected from the coculture of neutrophils and RAW264.7 macrophages using the Mouse IL-1$\beta$ ELISA Kit (PI301, Beyotime) and Mouse IL-18 ELISA Kit (PI553, Beyotime), respectively. To monitor mouse liver and kidney function, the levels of alanine transaminase (ALT), aspartate aminotransferase (AST), serum creatinine (Cre), and blood urea nitrogen (BUN) were assessed using ALT ELISA kit (ELK1921; ELK Biotechnology, Wuhan, China), AST ELISA kit (MBS164085; MyBioSource, San Diego, CA, USA), Creatinine Assay Kit (MBS2611108; MyBioSource), and BUN Assay Kit (MBS2611085; MyBioSource), respectively. Briefly, the diluted supernatants and standards were added to precoated 96-well plates. The absorbance was determined at 450 nm using a microplate reader (BK-9622; BioBASE, Shandong, China). A standard curve was constructed by plotting the absorbance values against the known concentrations of the standards. The resulting regression equation was then used to calculate the unknown sample concentrations from their measured absorbance.

Mouse fibroblasts, keratinocytes, and endothelial cells were seeded into 96-well plates. After overnight cultivation for adherence, the cells were treated with different types of CM. Cell viability was evaluated at 24, 48, and 72 h after treatment using a Cell Counting Kit-8 (CCK-8; Dojindo, Kumamoto, Japan) by measuring the absorbance at 450 nm with a microplate reader (BK-9622, BioBASE).

Cell migration was assessed using a transwell assay [18]. Treated fibroblasts, keratinocytes, and endothelial cells (5 $\times$ 10⁴) were plated in the upper chamber of 8.0 µm pore size transwell inserts (Corning). The lower chamber was filled with normal medium (control), NET-CM, or (NET+Ro 106–9920)-CM. After 24 h of incubation, the migrated cells were fixed and subsequently stained with 0.1% crystal violet solution. The migratory cells were recorded under an inverted microscope (IX-83, Olympus, Tokyo, Japan). For quantification, cells were counted on five random fields per membrane and their numbers were then averaged.

In vitro tube formation experiments were performed to assess the ability of endothelial cells to form branch points [19]. Endothelial cells (1 $\times$ 10⁴) were seeded onto each well of growth factor-reduced Matrigel-coated plates in endothelial basal medium (Lonza, Walkersville, MD, USA) supplemented with normal medium, NET-CM, or (NET+Ro 106–9920)-CM. The cells were cultivated for 6 h, after which images of tube structures were captured under an inverted microscope (Olympus). The number of branch points was defined as the junctions where three or more tubes intersected and was quantified manually using the ImageJ software (NIH).

For hematoxylin and eosin (HE) staining, the sections of the collected wound tissues, livers, and kidneys were stained with hematoxylin solution (Sigma-Aldrich) for 5 min, followed by rinsing for 10 min. Next, the hematoxylin-stained sections were differentiated in 1% acidic alcohol for 30 s and then treated with lithium carbonate solution for 1 min to develop the blue color of the nuclei. After rinsing, the sections were further counterstained with eosin solution for 2 min prior to dehydrating, clearing, and mounting. Three random sections per tissue sample were selected, and images of the wound center of each section were captured.

To assess the angiogenesis ability of endothelial cells immunohistochemically, the sections were rehydrated and subjected to antigen retrieval. After blocking with 5% normal goat serum, the sections were incubated with a primary antibody against platelet endothelial cell adhesion molecule-1 (CD31) (ab9498, 1:200 dilution; Abcam) overnight at 4 °C. The next day, the sections were incubated with a horseradish peroxidase (HRP)-conjugated secondary antibody (1:2000 dilution, A0208; Beyotime) for 1 h. The signal was developed using a 3,3^′-diaminobenzidine (DAB) substrate kit (Sigma-Aldrich), followed by counterstaining with hematoxylin. Stained sections were dehydrated, cleared, and mounted in a synthetic mounting medium. For this analysis, three random sections from each mouse were selected, and five randomly chosen fields from the images were examined.

Histological stained images were captured under a DM500 light microscope (Leica Microsystems).

Wound tissues collected on days 1, 3, 7, and 14 after wounding and cells collected after treatment were lysed to obtain total protein. The protein concentration of each sample was determined using a bicinchoninic acid (BCA) Protein Assay Kit (Thermo Fisher Scientific). Each protein (30 µg) was separated on 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels and transferred onto polyvinylidene difluoride (PVDF) membranes (Millipore, Burlington, MA, USA). The membranes were blocked with 5% skim milk, followed by overnight incubation at 4 °C with primary antibodies against the following proteins: histone H3 (ab1791, Abcam), CitH3, peptidyl arginine deiminase 4 (PAD4) (LS-C806278-10, LifeSpan BioSciences, Seattle, WA, USA), NLRP3 (AF2155, Beyotime), pro-IL-1$\beta$ (ab234437, Abcam), IL-1$\beta$ (ab283818, Abcam), apoptosis-associated speck-like protein containing a caspase recruitment domain (ASC) (AF6234, Beyotime), caspase-1 (AF1681, Beyotime), p-NF-$\kappa$B (AF5881, Beyotime), and NF-$\kappa$B. $\beta$-actin (AF5003, Beyotime) served as the internal reference. All samples were diluted in 5% skim milk at 1:1000 for use. After washing thrice, the membranes were incubated with an HRP-conjugated secondary antibody (1:5000 dilution, A0208; Beyotime) for 1 h at 25 °C. Protein bands were visualized using an enhanced chemiluminescence detection kit (P0018FS, Beyotime) and imaged using the ChemiDoc MP Imaging System (Bio-Rad, Hercules, CA, USA). Densitometric analysis was performed using the ImageJ software (NIH), with H3 or $\beta$-actin serving as the loading controls.

All data are expressed as the mean $\pm$ standard error of the mean. Statistical analyses were performed using GraphPad Prism software (version 9.0, GraphPad Software, San Diego, CA, USA). Comparisons between two groups were performed using the Student’s t-test. Comparisons among groups were performed using one- or two-way analysis of variance (ANOVA), followed by Tukey’s post hoc test. Statistical significance was set at p 0.05.

The experimental design and groups are shown in Fig. 1A. We observed the wound healing process for 14 d, and found that it was significantly delayed in the DM group compared with that in the control group (Fig. 1B) (F (3, 64) = 31.26, p 0.001). Regarding NET formation markers, we detected a marked increase in CitH3 expression in the DM group from day 7 onward compared with that in the control group (p 0.001). In addition, the PAD4 levels were elevated in the DM group as early as day 3 (p 0.01), suggesting the early initiation of NET formation under diabetic conditions (Fig. 1C). Furthermore, we found that the NLRP3 inflammasome and IL-1$\beta$/pro-IL-1$\beta$ axis were significantly upregulated in the DM group from days 7 (p 0.001) and 3 (p 0.001), respectively (Fig. 1D). Cleavage of the inactive precursor pro-IL-1$\beta$ releases the bioactive molecule IL-1$\beta$ that triggers identical biological responses, such as inflammation [20]. Our results suggested that hyperglycemia can enhance the processing of pro-IL-1$\beta$ to IL-1$\beta$, thereby increasing IL-1$\beta$ secretion in the lesions. Hence, sustained NET formation and prolonged NLRP3 inflammasome activation may contribute to the delayed wound healing observed in DM.

Fig. 1.

Sustained neutrophil extracellular trap (NET) formation and nucleotide-binding oligomerization domain-like receptor protein 3 (NLRP3) inflammasome activation in the skin wounds of diabetic mice. (A) After successfully inducing a diabetic mouse model through a high-fat diet combined with streptozotocin injection, an 8-mm full-thickness skin wound was created on the dorsal surface of the mice. (B) Photographs of the dorsal wounds were taken on days 1, 3, 7, and 14 and the changes in wound closure were recorded (n = 3 per group). Western blot analysis was conducted using the wound tissues of mice. (C) Citrullinated histone H3 (CitH3) and peptidyl arginine deiminase 4 (PAD4) expression represented NET formation (n = 3 per group). (D) NLRP3 inflammasome activation was evaluated by assessing the protein levels of NLRP3, pro-interleukin (IL)-1$\beta$, and IL-1$\beta$ (n = 3 per group). ns, no significance, **p 0.01, ***p 0.001. HFD, high fat diet; WB, Western Blot; STZ, streptozotocin; DM, diabetes mice.

Neutrophil identification is shown in Supplementary Fig. 1. The isolated neutrophils displayed a uniform shape and spherical structure (Supplementary Fig. 1A). Further validation of the purity of the isolated neutrophils is demonstrated in the flow cytometry analysis, where $>$95% of the cells are positive for both Ly6G and CD11b, indicating a highly specific and pure neutrophil population (Supplementary Fig. 1B). To investigate the effects of NETs on skin cell function, we treated purified mouse neutrophils with PMA to induce NET formation (Fig. 2A). After coculturing with RAW264.7 mouse macrophages for 12 h, immunofluorescence and western blot analyses revealed a significant increase in CitH3 expression in PMA-stimulated neutrophils compared with that in the untreated control (Fig. 2B,C; both p 0.001). This finding confirmed the successful induction of NETs. Subsequent ELISA analysis of the coculture supernatant demonstrated a notable elevation in the levels of proinflammatory cytokines IL-1$\beta$ and IL-18 in the PMA group, which coincided with elevated CitH3 levels (Fig. 2D,E; both p 0.001). Additionally, western blot analyses of macrophages revealed the upregulation of the expression of NLRP3, ASC, and caspase-1 in response to PMA treatment, indicating the activation of the NLRP3 inflammasome (Fig. 2F; all p 0.001).

Fig. 2.

NETs impair the proliferation and migration of fibroblasts and keratinocytes by inducing NLRP3 inflammasome activation in macrophages. (A) The experimental design and groups of the in vitro analysis. Purified mouse neutrophils were treated with phorbol 12-myristate 13-acetate (PMA) to induce NET formation, which was verified by assessing CitH3 expression in (B) immunofluorescence (n = 3 per group, scale bar = 20 µm) and (C) western blot analyses (n = 3 per group). After coculturing with mouse macrophage RAW264.7 cells for 12 h, the supernatant was collected for (D) IL-1$\beta$ and (E) IL-18 measurement via enzyme-linked immunosorbent assay (ELISA) and used as conditioned media (CM) in subsequent assays (n = 3 per group). (F) Western blot analysis of macrophages cocultured with NET-forming neutrophils revealed the protein levels of NLRP3, apoptosis-associated speck-like protein (ASC), and caspase-1 (n = 3 per group). Fibroblasts and keratinocytes were subsequently treated with CM for 72 h. (G,H) Cell viability at 24, 48, and 72 h after treatment, as demonstrated in Cell Counting Kit-8 (CCK-8) assays (n = 3 per group). After treatment, (I,J) the migration rate of cells was detected using the transwell assay (n = 3 per group). (K) Immunofluorescence (scale bar = 50 µm) and (L) western blot analyses were performed to demonstrate the activation of the nuclear factor kappa B (NF-$\kappa$B) signal pathway in neutrophils (n = 3 per group) following PMA treatment. **p 0.01, ***p 0.001. DAPI, 4^′,6-diamidino-2-phenylindole; NET-CM, neutrophil-macrophage coculture model; OD, Optical Density.

To further assess the functional effect of NETs on skin cells, including fibroblasts and keratinocytes, we applied CM from the coculture system to these cell types (Fig. 2A). The CCK-8 assay demonstrated a significant reduction in the viability of both fibroblasts and keratinocytes after 24, 48, and 72 h of NET-CM treatment compared with that in cells treated with normal medium (control) (Fig. 2G,H; both p 0.001). Moreover, we observed that the migration rates of fibroblasts (p 0.001) and keratinocytes (p 0.01) were markedly decreased in the presence of CM, suggesting impaired wound-healing capabilities (Fig. 2I,J). To explore the potential involvement of the NF-$\kappa$B signaling pathway in this process, we conducted further immunofluorescence and western blot analyses. We detected a significant increase in NF-$\kappa$B expression, as indicated by the elevated fluorescence (Fig. 2K; p 0.001), and protein p-NF-$\kappa$B/NF-$\kappa$B levels in neutrophils following PMA treatment (Fig. 2L; p 0.01). Collectively, these findings indicated that NET formation promotes NLRP3 inflammasome activation in macrophages, leading to the secretion of proinflammatory cytokines that impair the proliferation and migration of fibroblasts and keratinocytes. Therefore, we hypothesized that the activation of the NF-$\kappa$B pathway may be involved in this process.

We introduced Ro 106-9920 to explore its effect in NET formation. We observed a significant reduction in p-NF-$\kappa$B protein expression in PMA-induced neutrophils after Ro 106-9920 treatment (NET vs. NET+Ro 106-9920 groups, p 0.001; Fig. 3A). Additionally, Ro 106-9920 treatment reversed the PMA-induced elevated expression of CitH3 (NET vs. NET+Ro 106-9920 groups, p 0.001), effectively inhibiting NET formation (Fig. 3B). This inhibitory effect extended to the secretion of proinflammatory cytokines, as evidenced by the decreased levels of IL-1$\beta$ (p 0.001) and IL-18 (p 0.01) in the culture supernatant of the NET+Ro 106-9920 group compared with that of the NET group (Fig. 3C,D). Furthermore, we observed that the expression of NLRP3 (p 0.01), ASC (p 0.001), and caspase-1 (p 0.001) was markedly suppressed in macrophages in the NET+Ro 106-9920 group, suggesting that Ro 106-9920 effectively attenuates NET-induced NLRP3 inflammasome activation (Fig. 3E).

Fig. 3.

Suppression of the NF-$\kappa$B signaling pathway attenuates the deleterious effects of NETs on fibroblasts and keratinocytes. Mouse neutrophil-macrophage coculture models were treated with Ro 106-9920, an NF-$\kappa$B inhibitor. (A) Western blot analysis verified that NF-$\kappa$B activation was suppressed (n = 3 per group). (B) The relative expression of CitH3 was used as a marker of NET formation (n = 3 per group). ELISA analysis of the culture supernatant was conducted to assess the levels of proinflammatory cytokines (C) IL-1$\beta$ and (D) IL-18 after Ro 106-9920 treatment (n = 3 per group). (E) NLRP3 inflammasome activation was detected through the expression of NLRP3, ASC, and caspase-1 proteins via western blot analysis (n = 3 per group). The coculturing supernatant was used as conditioned media (CM), and added to fibroblasts and keratinocytes. The viability of (F) fibroblasts and (G) keratinocytes at 24, 48, and 72 h after treatment was measured using the CCK-8 assay (n = 3 per group). The migratory ability of (H) fibroblasts (scale bar = 50 µm) and (I) keratinocytes (scale bar = 100 µm) was measured using the transwell assay (n = 3 per group). **p 0.01, ***p 0.001.

Subsequent functional assays revealed that the viability of fibroblasts and keratinocytes was significantly higher in the (NET+Ro 106-9920)-CM group than in the NET-CM group after 72 h (Fig. 3F,G; both p 0.001). Moreover, we found that Ro 106-9920 treatment significantly improved the impaired migratory ability of fibroblasts and keratinocytes in the NET-CM group (Fig. 3H,I; both p 0.01). These findings indicated that suppression of the NF-$\kappa$B signaling pathway by Ro 106-9920 not only inhibits NET formation but also mitigates the associated inflammatory response, thereby protecting fibroblasts and keratinocytes from NET-induced damage.

Angiogenesis provides the blood and nutritional supply necessary to support wound tissue repair and regeneration [21]. To investigate whether Ro 106-9920 could mitigate the adverse effects of NETs on the angiogenic capacity of endothelial cells, we applied CM collected from cocultures with or without Ro 106-9920 treatment to endothelial cells for 72 h. Despite a general decline in endothelial cell viability following NET formation, the viability of cells treated with Ro 106-9920-CM was significantly higher than that of cells treated with NET-CM (Fig. 4A; p 0.001). Migration assays using the transwell system revealed a similar trend. Although NET formation led to a reduced migration rate of endothelial cells, the degree of inhibition was lower in the Ro 106-9920-CM group, indicating that Ro 106-9920 partially restores the NET-induced impairment in endothelial cell migration (Fig. 4B; p 0.01). Moreover, an analysis of angiogenic potential showed that the number of branch points was significantly higher in the (NET + Ro 106-9920)-CM group than that in the NET-CM group (Fig. 4C; p 0.001). These results indicated that Ro 106-9920 confers a protective effect on endothelial cells by mitigating the detrimental effects of NETs.

Fig. 4.

Suppression of the NF-$\kappa$B signaling pathway attenuates the deleterious effects of NETs on endothelial cells. CM collected from neutrophil-macrophage cocultures with or without Ro 106-9920 treatment was applied to endothelial cells for 72 h. (A) Endothelial cell viability was determined via the CCK-8 assay after 24, 48, and 72 h treatment (n = 3 per group). (B) The migration rate was evaluated using the transwell system (n = 3 per group, scale bar = 100 µm). (C) The number of branch points was calculated to analyze the angiogenic potential of endothelial cells (n = 3 per group, scale bar = 100 µm). **p 0.01, ***p 0.001.

To confirm the potential of Ro 106-9920 in promoting wound healing, we conducted an in vivo experiment using the diabetic mouse model (Fig. 5A). We performed subcutaneous injection of Ro 106-9920 or DMSO (vehicle) on multiple sites around the wounds of diabetic mice and monitored the dorsal wounds for 14 d. We observed a decrease in the wound area over time, with the DM+Ro 106-9920 group exhibiting significantly faster wound healing than the vehicle-treated group (Fig. 5B) (F (3, 40) = 53.93, p 0.001). The histological evaluation further corroborated these findings. The wound area in the DM+vehicle group displayed wider and more irregular scarring with evident dermal disruption, leading to increased scar width. In contrast, the DM+Ro 106-9920 group demonstrated narrower scars with a more organized tissue architecture, indicating enhanced wound healing and re-epithelialization (Fig. 5C; p 0.05). We conducted immunohistochemistry analysis for CD31 to assess angiogenesis in the wound tissues. The number of loops was significantly increased in the Ro 106-9920-treated wound compared with that in the vehicle-treated wound (Fig. 5D; p 0.001). This finding suggested that Ro 106-9920 not only accelerates wound closure but also promotes angiogenesis, which is critical for effective wound healing. Notably, Ro 106-9920 treatment had no effect on the blood glucose levels and weight of DM mice (Supplementary Fig. 2A,B), suggesting that its beneficial role in diabetic wound healing may be independent of its antihyperglycemic effects.

Fig. 5.

Ro 106-9920 accelerates cutaneous wound healing and promotes re-epithelialization in diabetic mice. (A) Diabetic mice were subcutaneously injected with Ro 106-9920 or dimethyl sulfoxide (DMSO) (vehicle) around the wound area, and wound healing was monitored for 14 d. (B) Images of wounds on days 1, 3, 7, and 14 after wound generation, when the closure rates were also calculated (n = 3 per group). (C) Histological evaluation of wounds via hematoxylin and eosin (HE) staining. Scar width was determined using the ImageJ software (n = 3 per group). Scale bar = 1000 µm. (D) Immunohistochemical analysis of platelet endothelial cell adhesion molecule-1 (CD31) was performed to assess angiogenesis, reflected by the number of loops (n = 3 per group, scale bar = 100 µm). *p 0.05, ***p 0.001. IHC, immunohistochemistry.

To further elucidate the mechanism by which Ro 106-9920 enhances wound healing in DM, we assessed the levels of key proteins involved in NET formation and inflammasome activation in the wound tissues of DM mice. Treatment with Ro 106-9920 markedly reduced the expression of CitH3 and PAD4 in wound tissues (Fig. 6A). Furthermore, we observed a marked reduction in the protein levels of NLRP3 and IL-1$\beta$/pro-IL-1$\beta$ in the Ro 106-9920-treated group compared with that in the Ro 106-9920-untreated control (Fig. 6B). Collectively, these findings suggested that Ro 106-9920 may accelerate wound healing in DM mice by suppressing NET formation and blocking NLRP3 inflammasome activation.

Fig. 6.

Ro 106-9920 suppresses NET formation and NLRP3 inflammasome activation in the skin wounds of diabetic mice. Wound tissues of mice treated with Ro 106-9920 or DMSO were collected for western blot analysis. (A) The expression levels of CitH3 and PAD4 in wound tissues indicated NET formation (n = 3 per group). (B) NLRP3 inflammasome activation was determined by the relative protein levels of NLRP3 and IL-1$\beta$/pro-IL-1$\beta$ ratios (n = 3 per group). *p 0.05, **p 0.01.

Finally, to observe whether Ro 106-9920 causes hepato- or nephro-toxicity, we conducted renal and liver biochemical and pathological evaluations during wound healing in diabetic mice. HE staining revealed no necrosis or injury in the liver sections of mice across all groups (Supplementary Fig. 3A). We also observed no changes in the serum ALT and AST levels across all groups of mice (Supplementary Fig. 3B,C). HE staining of kidneys showed an integrated architectural structure of healthy glomerular, with no necrosis in tubular cells (Supplementary Fig. 3D). This suggested that Ro 106-9920 has no nephrotoxicity during wound healing in diabetic mice, which was further corroborated through the evaluation of Cre and BUN levels (Supplementary Fig. 3E,F).