Effects and the mechanism of pine pollen polysaccharides on diabetic wound healing in vitro and in vivo

Abstract

Introduction

Pine pollen polysaccharides (PPPS) has anti-inflammatory, immunomodulatory, anti-oxidant, hypoglycemic, and anti-bacterial properties. PPPS can accelerate wound healing in mouse cutaneous wounds, yet it is unclear whether PPPS can promote diabetic wound healing.

Methods

Fibroblasts, keratinocytes, and human umbilical vein endothelial cells (HUVECs) were stimulated with high glucose (HG) to mimic hyperglycemic environment. Cell viability, apoptosis, migration, and angiogenesis were assessed by cell counting, Western blot, transwell migration, and tube formation assays. Neutrophil extracellular traps (NETs) and macrophage polarization were analyzed by immunofluorescence and flow cytometry. Streptozotocin-induced diabetes mellitus (DM) mice were subjected to skin wounds and PPPS administration to validate the role of PPPS in diabetic wound healing.

Results

PPPS treatment impaired HG-induced viability reduction, apoptosis promotion, and migration repression of fibroblasts, keratinocytes, and HUVECs, accompanied by promotion of angiogenesis of HUVECs under HG stimulation. Specifically, PPPS treatment facilitated NET degradation and suppressed macrophage M1 polarization. Furthermore, PPPS accelerated diabetic wound healing in DM mice, along with decreased citrullinated H3 and PAD4 protein levels and elevated CD31 protein levels, suggesting that PPPS facilitated re-epithelialization and vascularization and reduced NETs.

Conclusions

PPPS accelerates diabetic wound healing via mediating NETs and the polarization of M1-type macrophages, providing new insights into the promoting role of PPPS in diabetic wound healing.

Highlights

• •PPPS mitigates HG-induced injury to keratinocytes, fibroblasts, and HUVECs.
• •PPPS facilitates NET degradation and represses macrophage M1 polarization.
• •PPPS fosters skin wound healing in DM mice.
• •PPPS enhances re-epithelialization and vascularization in wound location of DM mice.
• •PPPS decreases NETs in wound location of DM mice.

1. Introduction

Diabetes mellitus (DM) is a common chronic disease that affects an increasing number of people around the world. One serious chronic complication of DM is diabetic foot ulcers (DFUs), which may progress to soft tissue infection, gangrene, and even require amputation [1]. Currently, DFU wounds with infection and necrosis are treated clinically by removing necrotic tissues in combination with hypoglycemic agents, anti-microbial agents, negative pressure drainage, and traditional dressings. However, the 5-year mortality for DFU patients is about 30 % [2], which may be associated with the low efficacy of therapeutic measures in biofilm removal and wound healing promotion [3]. Thus, it is imperative to establish novel and effective treatments for diabetic wounds.

Epidermal integrity and dermal stability are the basis of skin protection against the invasion of external harmful substances. Dermis consists mainly of fibroblasts and microvascular endothelial cells, and epidermis consists largely of keratinocytes. Wound healing is a dynamic process in which various cells in the wound area undergo cellular interactions by secreting cytokines to enhance cellular function and propel wound healing [4]. Angiogenesis is effective in supporting wound closure, whereas the reduction and poor regeneration of neovascularization key elements in the difficulty of wound healing for diabetic patients [5]. Well-directed migration of fibroblasts is important for the acceleration of wound healing, where they can be transformed into myofibroblasts to enhance wound contractility and promote wound recovery [6]. Re-epithelialization, involved in migration and proliferation of keratinocytes, is responsible for the surface reconstruction of skin wounds [5]. In contrast, diabetic wounds often remain stalled in the inflammatory phase due to persistent hyperglycemia, oxidative stress, and immune dysfunction. These conditions impair fibroblast and keratinocyte function, delay macrophage transition from the M1 to M2 phenotype, and hinder neovascularization, all of which contribute to the chronic, non-healing nature of diabetic wounds [7]. Hyperglycemia has been shown to directly impair the biological functions of microvascular endothelial cells, fibroblasts, and keratinocytes. Thus, targeting hyperglycemia-induced cellular dysfunction is a key therapeutic strategy for improving diabetic wound healing.

As the first defensive line of intrinsic immunity, neutrophils play a defensive role via secreting cytokines, degranulation, phagocytosis, and forming neutrophil extracellular traps (NETs). NETs are web-like filamentous extracellular structures of DNA, histones, and cytotoxic granule-derived proteins. The activation of neutrophils leads to the formation of NETs, which play a role in capturing and destroying pathogens [8]. Recent studies suggest that DM activates the production of NETs by neutrophils, and excessive or persistent NETs lead to delayed wound healing in DM patients [9]. Neutrophils do not fight microbes alone in the body but work with other immune cells to eliminate pathogens and reduce inflammation. Macrophages are essential for normal wound healing. At initial infiltration, the pro-inflammatory cytokines and growth factors are secreted by pro-inflammatory macrophages (M1-type), thus mobilizing more immune cells and promoting proliferation of keratinocytes, fibroblasts, and vascular endothelial cells [10]. During tissue neogenesis, the microenvironment triggers the conversion of macrophages to an anti-inflammatory phenotype (M2-type) and secretion of anti-inflammatory cytokines to promote extracellular matrix synthesis and wound contraction [10,11]. However, aberrant phenotypic conversion of macrophages from M1 to M2 types can disrupt the normal wound healing process, leading to difficulties in wound healing [11]. Therefore, seeking measures to interfere with the release of NETs and changes in macrophage phenotype can help to develop new therapeutic strategies for treating wound healing in DM.

Pine pollen polysaccharides (PPPS) are water-soluble polysaccharides with hydrophilic and viscous properties, with high medical value. Available evidence shows the anti-tumor, hypolipidemic, anti-inflammatory, immunomodulatory, anti-oxidant, hypoglycemic, anti-viral, and anti-bacterial properties of PPPS [12,13]. Wang et al. suggested that PPPS accelerate wound healing in mouse cutaneous wounds via promoting HaCaT cell proliferation [14]. Therefore, PPPS is a prospective natural agent in the therapeutic setting of wound healing, yet it is unclear whether PPPS can boost wound healing in DM.

Therefore, the present study sought to elucidate the efficacy of PPPS on wound healing in DM, uncovering the underlying therapeutic value of PPPS in diabetic wound healing.

2. Methods

2.1. Extraction, separation, and purity of PPPS

Broken Pinus pollen was provided by the Xi'an Tianguangyuan Biotechnology Co., Ltd (broken rate >95 %; Shanxi, China). Production of PPPS was carried out by the water extract-alcohol precipitation method [15]. In short, broken Pinus pollen was boiled in distilled water for 5 h at a ratio of 1:14. After collecting the supernatant, rotary evaporation was employed to concentrate it to 10 % of the original volume. Next, 60 % ethanol (3-fold volume) (Aladdin, Shanghai, China) was added and mixed overnight for precipitation at 4 °C. The white flocculent precipitates were collected, followed by the removal of proteins using trichloroacetic acid precipitation and purification with Sephacryl™ S-400HR (GE-Healthcare Bio Sciences Ab, Marlborough, MA, USA).

2.2. Cell culture

Human dermal fibroblasts (#CP-H103; Procell, Wuhan, China), human epidermal keratinocytes (#CP-H113; Procell), and human umbilical vein endothelial cells (HUVECs; #CP-H082; Procell) were cultured in Dulbecco's modified Eagle's medium (DMEM; Thermo, Grand Island, NY, USA) containing 10 % (v/v) fetal bovine serum (FBS, Thermo) and 1 % (v/v) penicillin/streptomycin (Beyotime, Shanghai, China) at 37 °C in an incubator with 5 % CO₂.

2.3. Cell viability analysis

Gradient concentrations (0, 10, 20, 50, 100, 200, 400, and 800 μg/ml) of PPPS were deployed to treat the above three types of cells for 24 h to assess the cytotoxicity of PPPS. To study the effect of PPPS on the viability of fibroblasts, keratinocytes, and HUVECs under hyperglycemia in vitro, fibroblasts, keratinocytes, and HUVECs (5 × 10³ cells/well) were seeded respectively into 96-well plates at normal glucose (NG; 5.5 mmol/L) or high glucose (HG; d-glucose, 33.3 mmol/L) (Sangon, Shanghai, China) media for overnight, followed by treatment with high (H; 50 μg/ml), medium (M; 20 μg/ml), and low (L; 10 μg/ml) concentrations of PPPS for 24 h. The cell counting kit-8 (CCK-8) reagent (10 μl; Solarbio, Beijing, China) was added to each well. One hour later, the absorbance at 450 nm was measured using a microplate reader (BioTek, Winooski, Vermont, USA).

2.4. Western blot

After incubation with HG or HG combined with PPPS, fibroblasts, keratinocytes, and HUVECs were collected. After lysis with chilled RIPA buffer containing protease and phosphatase inhibitors (Sigma, St Louis MO, USA) the cell supernatants were harvested, followed by determination of protein concentration with a BCA assay kit (Solarbio). Proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to polyvinylidene fluoride membranes. After blocking with 5 % non-fat milk (Sangon) in TBST for 1 h, the membranes were incubated with anti-Bcl-2 (#AF6285, Beyotime; 1:1000), anti-Bax (#AB026; Beyotime; 1:1000), anti-citrullinated H3 (Cit-H3) (#97272; CST, Danvers, MA, USA; 1:1000), anti-H3 (#9715; CST; 1:1000), anti-peptidylarginine deiminase 4 (PAD4) (#Ab214810; Abcam, Cambridge, MA, USA; 1:1000), and anti-β-actin (#AF0003; Beyotime; 1:1000) antibodies overnight at 4 °C. Protein bands were visualized with enhanced chemiluminescence reagent (Thermo) after incubation with appropriate secondary antibodies (1:5000, Abcam). Quantification of grey-scale values of the blots was performed using Image J software.

2.5. Cell migration analysis

Fibroblasts, keratinocytes, and HUVECs (1 × 10⁵ cells/well) were starved respectively for 1 h and then treated with or without PPPS in the presence or absence of HG on the upper region of transwell chambers (8-μm pore size, Corning, NY, USA). The lower chambers were supplemented with a complete culture medium containing 10 % FBS for 24 h. After stabilization with 4 % paraformaldehyde (Beyotime) for 15 min, the migrated cells were stained with 0.1 % crystal violet (Solarbio) for 10 min. The cell images were captured using an inverted microscope (Olympus IX51, Tokyo, Japan) and quantified using ImageJ software (NIH, Bethesda, MD, USA).

2.6. Tube formation assay

In brief, 24-well plates were pre-coated with Matrigel (300 μl/well; BD Biosciences, San Diego, CA, USA) for 1 h in an incubator at 37 °C. HUVECs (1 × 10⁵) pretreated with different approaches in FBS-free DMEM media were reseeded 24-well plates pre-coated with Matrigel. Visualization and capturing of capillary-like structures was done through a microscope and quantification of the number of branching points was achieved by ImageJ software.

2.7. Isolation of mouse neutrophils

Blood samples collected from the retro-orbital plexus of anesthetized mice were placed in sterile EDTA-anticoagulated tubes. Mouse neutrophils were isolated from collected blood using the Mouse Peripheral Blood Neutrophil Isolation Solution Kit (#P9201; Solarbio) following the manufacturer's instructions. Neutrophils were isolated by centrifugation (25 °C, 20 min, 1000 g). Erythrocytes mixed with neutrophils were lysed with red blood cell lysis. Then, the neutrophils were washed and collected for subsequent analysis.

2.8. Immunofluorescence (IF) analysis

A murine macrophage cell line RAW264.7 (ATCC, Manassas, VA, USA) was cultured in DMEM containing1% (v/v) penicillin/streptomycin and 10 % (v/v) FBS. In the co-culture system, RAW264.7 cells (1 × 10⁵) were incubated in a complete culture medium containing LPS plus IFN-γ (Sigma) and vehicle (PBS) or PPPS in the upper Transwell chamber (Corning). The isolated neutrophils (2 × 10⁴) were seeded on the sterile cell slides pre-coated with poly-l-lysine (Sigma) and put into the lower Transwell chamber. After incubation for 24 h, the generation of NETs from neutrophils was observed by IF analysis. In brief, neutrophils were blocked with 5 % BSA and then incubated with an anti-histone H3 (#ab5103, 1:1000; Abcam) antibody after immobilization for 10 min with 4 % paraformaldehyde. After incubation with a secondary antibody (1:200; Proteintech, China), neutrophils were counterstained with hoechst (1 g/ml, Thermo) and imaged using a fluorescence microscope (Olympus IX51).

2.9. Flow cytometry for M1 macrophage polarization

The polarization of macrophages co-cultured with neutrophils treated with LPS plus IFN-γ and vehicle (PBS) or PPPS were analyzed. In brief, RAW264.7 cells (2 × 10⁵) were incubated in a complete culture medium in the upper Transwell chamber (Corning). The isolated neutrophils (2 × 10⁴) were incubated in a complete culture medium containing LPS plus IFN-γ and vehicle (PBS) or PPPS in the lower Transwell chamber. After incubation for 24 h, RAW264.7 cells were stained with anti-CD80-FITC (eBioscience, San Diego, CA, USA) for 30 min. Cell surface expression was detected using a Beckman Cytoflex flow cytometry (Brea, CA, USA), and the data were analyzed with FlowJo software (NIH).

2.10. Animal grouping and wound formation

C57BL/6J mice (male; weight: 26–30 g; age: 8 weeks) (Huafukang Experimental Animal Co, Ltd., Beijing, China) were kept in the clean animal laboratory with constant humidity (60 % ± 10 %), temperature (22 ± 2 °C), and light (a 12/12-h light/dark cycle). All procedures in animal experiments were conducted following the animal care and rules of the Institute of Guizhou Medical University.

After 1 week of adaptive feeding, diabetic mice (n = 18) were constructed by intraperitoneal injection of streptozotocin (STZ, Sigma) at a dose of 50 mg/kg for 5 days. The control mice (n = 6) were injected intraperitoneally with an equal amount of sodium citrate buffer solution (0.1 mmol/L, pH = 4.5). Four weeks after STZ induction, mice with fasting blood glucose levels >16.7 mmol/L but <33.3 mmol/L were considered successful for modelling. For skin wound production, all mice were de-haired after anesthesia with 0.6 % sodium pentobarbital (10 ml/kg, intraperitoneal injection). A full-thickness (skin and panniculus carnosus) excisional incision was created on the back of each mouse using a sterile 8-mm skin punch biopsy (Acuderm, USA). Diabetic mice were divided into 3 groups according to a random number and received different treatments: PBS, PPPS (100 mg/kg), and PPPS (200 mg/kg). PBS and PPPS were injected subcutaneously into multiple sites (6 sites/wound) twice a week for two weeks, respectively. The control mice were administered equal amounts of PBS in the same way. Representative pictures of the wound area were captured on days 1, 3, 7, and 14 during the treatment period. Analysis of wound closure was performed using ImageJ software, and the wound closure (%) was calculated as: (A₀ -A_n)/A_n × 100 % (A₀ and A_n referred to the wound area on day 1 and day n, respectively (cm²).

2.11. Glucose metabolism tests

Before the end of the experiment, mice underwent a 12-h fast, followed by receiving an oral glucose tolerance test (OGTT). Each mouse was injected intraperitoneally with glucose solution (2 g/kg) prepared in normal saline (10 μl/g). Blood samples from the tail vein were collected to measure glucose concentrations at 0, 30, 60, 90, and 120 min post-injection.

An insulin tolerance test (ITT) was undertaken on mice with a fast of 5 h. Insulin (0.50 IU/kg) (Sunncell, Wuhan, China) was injected intraperitoneally into each mouse. Glucose concentrations were measured at 0, 30, 60, 90 and 120 min post-injection by collecting blood from the tail vein.

2.12. Histological analysis

Mice were killed at day 14 after treatment with PBS or PPPS, and the skin tissues at wound sites were collected and fixed with formalin, embedded with paraffined, and sliced into 5 μm tissue sections. The tissues were stained with hematoxylin and eosin (HE) after deparaffinization and rehydration. The histological wound healing score was computed by determining scar width in each mouse [16].

For immunohistochemical (IHC) staining, the sections were blocked with 5 % BSA and incubated with a primary antibody against CD31 (#ab182981; 1:100, Abcam) at 4 °C overnight. Then, the slides were covered with the goat anti-rabbit IgG H&L (HRP), followed by staining with DAB (Boster, Wuhan, China). The slides were imaged using under a microscope (Olympus IX51). Blood vessel loops were analyzed quantitatively by Image J software.

2.13. Statistical analysis

For statistical analysis and graphic presentation, GraphPad Prism 9 software (GraphPad, San Diego, CA, USA) was utilized. In the bar graphs, data are shown as means ± standard error of the mean (SEM). Statistical significances in more than two groups were analyzed by one-way and two-way ANOVA with Tukey's multiple comparison test, with a pre-defined significance level of p < 0.05.

3. Results

3.1. PPPS attenuates HG-induced injury in fibroblasts and keratinocytes

Pine pollen extract and PPPS have been published to boost wound healing [14,17], yet whether PPPS boosts wound healing in a hyperglycemic environment is unknown. Hyperglycemia tends to damage the function of effector cells for wound healing, including vascular endothelial cells, fibroblasts, and keratinocytes [18,19]. To investigate the role of PPPS in diabetic wound healing, we first analyzed the toxicity of PPPS on fibroblasts, keratinocytes, and HUVECs. Concentration gradients of PPPS (0, 10, 20, 50, 100, 200, 400, 800 μg/ml) were allowed to incubate the above three types of cells for 24 h, followed by measuring the cell activity. The results showed that 10 μg/ml of PPPS did not affect the viability of fibroblasts, keratinocytes and HUVECs. Nevertheless, the activities of these cells were boosted progressively with increasing concentrations of PPPS (20–200 μg/ml), and there was a gradual decrease in the promoting effect on the activities of these cells when the concentration was ≥400 μg/ml (Fig. S1A–C). Because the changes in the viability of these three cell types were not significant when the concentrations of PPPS were 10 μg/ml, 20 μg/ml, and 50 μg/ml, they were chosen for subsequent cell analyses as low, medium, and high concentrations (Fig. S1A–C).

Subsequently, we assessed the alterations in viability, apoptosis and migration of HG-induced fibroblasts and keratinocytes under PPPS treatment. PPPS treatment improved the inhibitory effect of HG on fibroblast viability in a concentration-dependent pattern (Fig. 1A). We also discovered that HG repressed Bcl-2 protein levels and boosted Bax protein levels, whereas PPPS treatment impaired the changes in Bcl-2 and Bax protein levels mediated by HG stimulation (Fig. 1B). Moreover, the inhibitory effect of HG on fibroblast migration was ameliorated following PPPS treatment in a dose-dependent manner (Fig. 1C). Similarly, PPPS ameliorated HG-mediated effects on keratinocyte viability, apoptosis, and migration, with better efficacy with a higher dose (Fig. 1D–F). These results demonstrated that PPPS treatment lessens HG-induced damage to fibroblasts and keratinocytes.

Fig. 1.

PPPS weakens HG-induced fibroblast and keratinocyte injury. (A and D) Analysis of the viability of fibroblasts and keratinocytes in different groups (NG, HG, HG + PPPS-L, HG + PPPS-M, and HG + PPPS-H) was carried out by CCK-8 assays (n = 3). One-way ANOVA and Tukey's multiple comparison test. (B and E) Western bot was executed to assess Bcl-2 and Bax protein levels (n = 3). Two-way ANOVA and Tukey's multiple comparison test. (C and F) The migration of fibroblasts and keratinocytes was detected by transwell migration assays (n = 3). Scale bar = 50 μm. One-way ANOVA and Tukey's multiple comparison test. Data are presented as means ± SEMs. ∗∗p < 0.01 and ∗∗∗p < 0.001 versus NG; ^#p < 0.05, ^##p < 0.01, and ^##p < 0.001 versus HG.

3.2. PPPS alleviates HG-mediated effects on viability, apoptosis, migration, and angiogenesis of HUVECs

Vascular reconstruction is essential for wound healing. The migration of vascular endothelial cells into the wound area provides metabolic support for tissue repair and accelerates wound healing by forming capillary structures [20]. Thus., we further analyzed the influence of PPPS on HG-induced HUVECs. The results showed that HG-driven reduction in the viability of HUVEC was relieved post PPPS treatment in a dose-dependent manner, accompanied by an improvement in HG-mediated down-regulation of Bcl-2 protein levels and up-regulation of Bax protein levels (Fig. 2A and B). With increasing concentrations of PPPS, the migratory capacity of HUVECs was also boosted under HG stimulation (Fig. 2C). In addition, PPPS enhanced the angiogenic capability of HG-induced HUVECs (Fig. 2D). Collectively, these outcomes suggested that PPPS mitigates HG-mediated effects on HUVEC viability, apoptosis, migration, and angiogenesis.

Fig. 2.

PPPS improves HG-mediated impairment in viability, apoptosis, migration, and angiogenesis of HUVECs. (A) The viability of HUVECs in different groups (NG, HG, HG + PPPS-L, HG + PPPS-M, and HG + PPPS-H) was determined by CCK-8 assays (n = 3). One-way ANOVA and Tukey's multiple comparison test. (B) Relative protein levels of Bcl-2 and Bax were evaluated by Western blot (n = 3). Two-way ANOVA and Tukey's multiple comparison test. (C and D) The migration and angiogenesis of HUVECs were analyzed by transwell migration and tube formation assays (n = 3). Scale bar = 50 μm. One-way ANOVA and Tukey's multiple comparison test. Data are presented as means ± SEMs. ∗∗∗p < 0.001 versus NG; ^#p < 0.05, ^##p < 0.01, and ^##p < 0.001 versus HG.

3.3. PPPS elevates NET degradation and represses macrophage M1 polarization

Neutrophils play a crucial role in the innate immune response during the early phases of wound healing. Upon activation by pathogens or inflammatory stimuli, neutrophils can undergo either apoptosis or NETosis. In the physiological setting, apoptotic neutrophils are promptly cleared by macrophages through efferocytosis, which facilitates their transition toward the M2 phenotype. M2 macrophages in turn secrete reparative mediators such as IL-10, Arg1, VEGF, and TGF-β, promoting angiogenesis, fibroblast activation, and extracellular matrix remodeling. In contrast, NETosis, a specialized form of neutrophil death characterized by the release of DNA webs decorated with histones and MPO, contributes to tissue damage, prolonged inflammation, and impaired vascular regeneration. Excessive or unresolved NET formation has been implicated in delayed wound healing, particularly in diabetic wounds. These contrasting outcomes are summarized in a conceptual schematic (Fig. 3A), which illustrates the divergent consequences of neutrophil apoptosis versus NETosis on macrophage polarization and subsequent tissue repair. The diagram underscores the critical balance between pro-resolving and pro-inflammatory pathways in orchestrating effective wound healing. To experimentally evaluate the effect of PPPS on the interaction between macrophages and neutrophils undergoing NETosis, we co-cultured mouse-derived neutrophils with PMA-induced THP-1 macrophages (M0 phenotype) in the presence of IFN-γ (20 ng/ml) and LPS (1 μg/ml), with or without PPPS treatment. IF assays were performed with anti-histone 3 antibody to identify NETs, and the results displayed that extensive NET formation under inflammatory stimulation, which was markedly reduced upon PPPS treatment (Fig. 3B). To verify that PPPS may degrade NETs by mediating macrophage polarization, we detected changes in M1-type macrophages under the same incubation conditions. Flow cytometry assays demonstrated that IFN-γ and LPS promoted M1 polarization of THP-1 macrophages, as indicated by increased CD80 expression. This M1 shift was significantly attenuated by PPPS treatment (Fig. 3C). These findings suggested that PPPS treatment reduces NET formation and suppresses M1 macrophage polarization under inflammatory conditions, which may support wound healing in diabetic conditions.

Fig. 3.

Administration of PPPS facilitates NET degradation and represses macrophage M1 polarization. (A) Schematic illustration of neutrophil fate and its impact on macrophage polarization. Apoptotic neutrophils are cleared by macrophages, promoting M2 polarization and tissue repair. In contrast, NETosis leads to the release of Cit-H3, MPO, and DNA, sustaining inflammation, enhancing M1 polarization, and impairing wound healing in diabetes. (B) Representative IF images of Histone 3 in mouse neutrophils and co-incubated them with PMA-induced THP-1 cells (M0) for 24 h in the presence of 1 μg/ml of LPS, 20 ng/ml of IFN-γ, and PPPS (L, M, and H) (n = 3). Scale bar = 10 μm. (C) The surface marker of M1 macrophages (CD80) was detected by flow cytometry and quantitative analysis of CD80-positive macrophages (n = 3). One-way ANOVA and Tukey's multiple comparison test. Data are presented as means ± SEMs. ∗∗∗p < 0.001 versus control; ^##p < 0.01 versus vehicle.

3.4. PPPS promotes skin wound healing in mice with DM

To further validate the role of PPPS in diabetic wound healing, wounds were created on the back of STZ-induced DM mice and treated with PPPS (Fig. 4A). After 1 week of STZ injection, successful DM mouse models were confirmed by measuring blood glucose levels (>16.7 mmol/L). Glucose tolerance and insulin sensitivity were evaluated by OGTT and ITT. Compared with the control group, DM mice and PPPS-treated DM mice demonstrated high GTT and ITT values, accompanied by elevated AUC values, as shown in Fig. 4B and C. However, the above parameters didn't differ significantly between DM mice and PPPS-treated DM mice (Fig. 4B and C). Representative wound pictures and wound healing rates at the indicated times are shown in Fig. 4D. Wound healing rates among different groups on the 1st and 3rd day did not significant differences. On days 7 and 14, DM mice exhibited low wound healing rates compared to control mice, whereas PPPS treatment boosted wound healing rates in DM mice. These outcomes suggested that treatment with PPPS boosts skin wound healing in DM mice.

Fig. 4.

PPPS treatment fosters skin wound healing in DM mice. (A) Experimental procedure of the construction of DM mice and administration of PPPS. (B and C) Glucose tolerance and insulin sensitivity were determined by OGTT and ITT prior to the ending of the animal experiments (n = 6) One-way/two-way ANOVA and Tukey's multiple comparison test. (D) Representative images of wounds and wound contraction in different groups on day 1, 3, 7, and 14 post treatment (n = 6). One-way/two-way ANOVA and Tukey's multiple comparison test. Data are presented as means ± SEMs. ∗∗∗p < 0.001 versus control; ^##p < 0.01 and ^###p < 0.001 versus DM.

3.5. PPPS enhances re-epithelialization and vascularization and attenuates NETosis in DM mice

To further investigate the role of PPPS on skin wound healing in DM mice, we performed H&E staining with wound surface tissues from mice on day 14. The results showed that the wound areas in control and high-dose PPPS-treated mice were filled fully with granulation tissues. Quantitative analysis showed that mice in the DM group possessed higher scar widths versus the control group, but PPPS treatment improved scar widths (Fig. 5A). IHC staining of wound tissues with an anti-CD31 (vascular epithelial cell marker) antibody displayed that the number of CD34-positive blood vessels in wound tissues of DM mice was significantly reduced in comparison to wound tissues of control mice, yet treatment with PPPS elevated the number of CD34-positive blood vessels in wound tissues of DM mice (Fig. 5B). Additionally, Cit-H3 and PAD4 protein levels were detected in wound tissues to observe changes in NETs. As expected, Cit-H3 and PAD4 protein levels were up-regulated in wound tissues of DM mice, while these elevated protein levels were impaired following PPPS treatment (Fig. 5C). Together, these results manifested that PPPS enhances re-epithelialization and vascularization and attenuates NETosis in DM mice.

Fig. 5.

PPPS administration enhances re-epithelialization and vascularization as well as attenuates NETosis in skin-injured DM mice. (A) Representative images of HE staining and quantitative analysis of scar width (n = 6). Scale bar = 1000 μm. One-way ANOVA and Tukey's multiple comparison test. (B) Representative images of IHC staining of CD31 and quantitative analysis of the number of CD34-positive blood vessels (n = 6). Scale bar = 100 μm. One-way ANOVA and Tukey's multiple comparison test. (C) The protein levels of Cit-H3 and PAD4 were detected by Western blot (n = 6). Two-way ANOVA and Tukey's multiple comparison test. Data are presented as means ± SEMs. ∗∗p < 0.01 and ∗∗∗p < 0.001 versus control; ^#p < 0.05 and ^###p < 0.001 versus DM.

4. Discussion

Current therapeutic measures for diabetic wound healing have limited effectiveness, especially for ulcers with high Wagner grading [21]. The present study elucidated the efficacy of PPPS on wound healing in DM, and the outcomes demonstrated that PPPS promotes diabetic wound healing by alleviating HG-induced impairment in fibroblasts, keratinocytes, and HUVECs, degrading NETs, and inhibiting M1 macrophage polarization.

Wound healing depends on processes involved in the formation of fresh granulation tissue and re-epithelialization, the destruction of peripheral blood vessels and the reduction of neovascularization and difficulties in regeneration are important factors that lead to difficulties in the healing of diabetic wounds [22]. Typically, angiogenesis is governed by the proliferation and migration of vascular endothelial cells, and intact blood vessels can produce granulation in granulation tissues. During the process of wound epithelialization, keratinocytes migrate to the wound location, proliferate, and differentiate into different structures to restore the structural and functional integrity of the epidermis [23]. Dermal repair is largely dependent on fibroblasts, where lower fibroblasts participate in the maintenance of the morphological structure of the dermis and provide a stable environment for immune clearance activities and angiogenesis, while upper fibroblasts regulate follicle growth and hair regeneration through the formation of hair papillae [24]. Wound-associated disturbances (like DM) in cell behaviors can lead to healing impairments and the formation of chronic and non-healing wounds [18,19]. Thus, improving hyperglycemia-mediated injury to fibroblasts, keratinocytes, and vascular endothelial cells is the target for treating wound healing in DM.

PPPS holds hypoglycemic, immuno-strengthening, and anti-bacterial properties. It was pointed out that PPPS facilitates the growth of blood vessels in chick embryo chorioallantoic membrane and accelerates skin wound healing in mice [14]. In the current study, PPPS treatment attenuated HG-induced reductions in cell viability and migration and suppressed apoptosis in fibroblasts, keratinocytes, and HUVECs. It also enhanced angiogenic capacity in HUVECs under high glucose conditions. In vivo, PPPS administration increased skin wound closure rate in diabetic mouse models, along with reduced scar width and elevated CD31-positive vessel density in the wound location. These findings align with the pathological characteristics of diabetic wounds, which are often trapped in the inflammatory phase and show impaired angiogenesis and delayed tissue remodeling. In contrast, non-diabetic wounds proceed through well-regulated stages of inflammation resolution, granulation tissue formation, re-epithelialization, and vascular regeneration. Our model faithfully recapitulated this delayed healing phenotype in diabetic conditions. The ability of PPPS to partially reverse these diabetic-specific impairments highlights its potential as a therapeutic agent targeting key obstacles in diabetic wound healing.

Recent studies demonstrated that DM activates neutrophils to undergo NETosis and that excess NETs cause a delay in wound healing [25]. NETosis involved in wound repair is mostly induced by PMA. When the nuclear membrane fuses with the cytoplasmic granule membrane, neutrophil elastase (NE) degrades histone H1 that acts as a linker in chromatin and PAD4 catalyzes the citrullination of histone H3, resulting in the depolymerization of chromatin and forming a mixture of DNA and antimicrobial proteins, eventually discharging to the outside of the cell to form a network-like structure with the rupture of the cytoplasmic membrane [26]. High concentrations of NE lead to delayed wound healing by degrading the matrix of the wound, and NETs and histone 3 proteins cause a direct damage to epithelial and endothelial cells [27]. Diabetic mice exhibit elevated citrullinated H3 and delayed wound healing compared to normal mice, and patients with intractable DFUs show a significant elevation of NETs [28]. Wild-type mice yield a large quantity of NETs from cutaneous wounds, yet PAD4^−/− mice do not produce NETs and wound healing is accelerated in PAD4^−/− mice compared to wild-type mice [25]. These results imply that modulation of NETs holds a great prospect for diabetic wound healing. In the current research, PPPS was discovered to degrade NETs and PPPS treatment accelerated wound healing in DM mice, along with decreased Cit-H3 and PAD4 protein levels in wound tissues, highlighting that PPPS promotes wound healing in DM mice by degrading NETs.

An impairment of macrophage conversion from M1 to M2 types has been found to be one of the main causes of delayed wound healing in diabetic mice [29]. The persistence of M1-type macrophages in diabetic wounds leads to a sustained release of pro-inflammatory cytokines such as TNF-α and IL-1β, which suppress angiogenesis, impair fibroblast migration, and delay transition to the proliferative and remodeling phases [30]. In addition, the decrease in M2-type macrophages compromise their ability to clear neutrophils undergoing NETosis, leading to the accumulation of NETs and further aggravating tissue injury [30]. In the present study, PPPS treatment significantly inhibited M1 polarization in a co-culture system under inflammatory stimulation. This may relieve the chronic inflammatory state in diabetic wounds and create a microenvironment favorable for tissue regeneration. It is noteworthy that this effect may be mediated directly by PPPS or indirectly via its inhibition of NET formation. Although the precise mechanistic link between NET suppression and macrophage reprogramming remains to be elucidated, our findings support the hypothesis that PPPS promotes wound healing through coordinated immunomodulation targeting both neutrophils and macrophages.

Although PPPS have been reported to exhibit hypoglycemic properties in other contexts, our study found that at the doses used (100 and 200 mg/kg), PPPS did not significantly improve systemic glucose metabolism in diabetic mice, as assessed by OGTT and ITT assays. This suggests that the beneficial effects of PPPS on wound healing are not mediated by improvements in glycemic control under these experimental conditions. Instead, our findings indicate that PPPS primarily exerts local effects on the wound microenvironment, including the suppression of NET formation, attenuation of M1 macrophage polarization, and enhancement of angiogenesis, as summarized in Fig. 6. These results support the conclusion that PPPS promotes diabetic wound healing through immunomodulation and tissue remodeling, rather than systemic metabolic regulation.

Fig. 6.

Proposed mechanism by which PPPS promotes diabetic wound healing. In diabetic wounds, elevated AGEs and ROS lead to excessive NET formation, sustained M1 macrophage polarization, impaired angiogenesis, and delayed tissue remodeling. PPPS treatment suppresses NETosis and oxidative stress, facilitates M2 polarization, and enhances endothelial cell and fibroblast function, thereby improving angiogenesis, collagen deposition, and tissue regeneration.

Fig. 6 summarizes the proposed mechanism by which PPPS promotes diabetic wound healing. Under diabetic conditions, elevated AGEs and ROS drive NET formation, M1 macrophage polarization, impaired angiogenesis, and delayed tissue remodeling. PPPS treatment reduces NETosis and oxidative stress, facilitates M2 polarization, and promotes the function of endothelial cells and fibroblasts, enhancing angiogenesis, collagen deposition, and fibroblast regeneration. These combined effects contribute to improved tissue repair and scarless wound healing in diabetic mice. Overall, our findings indicate that PPPS promotes diabetic wound healing by modulating local immune responses and tissue remodeling processes rather than altering systemic glucose metabolism. This multifaceted mechanism supports the potential application of PPPS as a promising therapeutic strategy for chronic diabetic wounds.

5. Conclusions

In conclusion, PPPS contributes to wound healing by degrading NETs and repressing M1 macrophage polarization in DM, providing evidence to support that PPPS may be a valuable natural pro-wound healing agent in the hyperglycemic environment. Unfortunately, we did not explore the molecular mechanisms by which PPPS mediates NETs and macrophage polarization, which is a key thrust for our investigation in the future.

Funding

This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

Declaration of competing interest

The authors declare no potential conflicts of interest.

Appendix A. Supplementary data

The following is the Supplementary data to this article.

Fig. S1.

The toxicity of PPPS on fibroblasts, keratinocytes, and HUVECs. (A-C) The viability of fibroblasts, keratinocytes, and HUVECs treated with concentration gradients of PPPS (0, 10, 20, 50, 100, 200, 400, 800 μg/ml) were measured by CCK-8 assays (n = 3). One-way ANOVA and Tukey's multiple comparison test. Data are presented as means ± SEMs. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001 versus 0 μg/ml.