An innovative ischemic diabetic flap model for chronic wound research

Abstract

Preclinical animal models that are simple, reproducible, and capable of partially mimicking the pathophysiological features of clinical diabetic foot ulcers (DFUs) are essential for advancing research on refractory wound healing. Compared to mice, Sprague-Dawley (SD) rats offer advantages such as reduced stress and greater ease of modeling. Additionally, type 2 diabetes induced in rats via a high-fat diet combined with intraperitoneal streptozotocin (STZ) injection is more cost-effective than genotyped rats. To simulate ulcers caused by diabetic vasculopathy, we established two bipedicle ischemic flaps of different widths (2 cm and 2.5 cm, both 11 cm long) on the backs of diabetic rats. Two full-thickness skin defects (6 mm in diameter) were created bilaterally at the flap midpoint to prevent fascial contraction that is a common issue in conventional models. As controls, we created bipedicle ischemic flaps (11 cm long, 2 cm wide) and corresponding wounds on the backs of normal SD rats. Additionally, classic full-thickness wounds (6 mm in diameter) were established at the same location on diabetic rats without flaps. We evaluated wound healing rates, epithelialization, neocollagenesis, angiogenesis, macrophage polarization, and the expression levels of pro-inflammatory factors and MMPs. The wounds within the 2.0 cm-wide ischemic flaps exhibited delayed epithelialization, reduced neocollagenesis, and an increased number of CD31-positive endothelial cells, with elevated VEGF-A expression but fewer mature blood vessels. Higher levels of IL-1β, TNF-α, IL-6 MMP2 and MMP9, and M1 macrophages were also observed. In conclusion, a bipedicle ischemic flap (11 cm long, 2 cm wide) combined with full-thickness skin defects on the backs of type 2 diabetic rats induced by a high-fat diet with STZ injection provides an effective model for studying DFU-associated vasculopathy and chronic inflammation.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12893-025-03237-5.

Introduction

The rising prevalence of diabetes mellitus and its chronic complications represents a significant global health challenge [1, 2]. Among these complications, DFUs affect approximately 18.6 million individuals annually [3]. Additionally, the recurrence rate of ulcers within 3–5 years post-treatment as high as 65%, with an amputation rate of 20% and a 5-year mortality rate of 50–70% [4]. Despite the availability of various therapeutic strategies exist, most focus on managing ulcer infections, and no single approach is universally recommended for the treatment of DFUs [5]. However, the lack of suitable models has contributed to a high failure rate−up to 90%−in translating preclinical therapies into clinical applications [6]. Given these challenges, the development of appropriate animal models for preclinical studies is essential to understanding the molecular mechanisms of DFUs and evaluating potential therapeutic approaches [7].

Pigs and primates, which closely resemble human skin, have been used to study wound healing and STZ-induced models of diabetes [8], but their high acquisition and maintenance costs limit their widespread use. Rodents, particularly mice and rats, are widely selected for animal studies, with the choice depending on the physiological, anatomical, and pharmacological characteristics relevant to specific research objectives [9]. Compared to mice, rats are 8–10 times heavier, easier to handle, and better suited for creating larger wounds—advantages particularly valuable in chronic wound research [9]. Additionally, rats exhibit lower stress responses, further enhancing their suitability as research models [10]. Currently, chronic wound models are often generated by inducing underlying diseases, followed by the creation of full-thickness excisional wounds. However, in diabetic rat models, these wounds primarily heal through contraction and fail to replicate critical features of chronic wounds such as ischemia and prolonged inflammation [7, 11–14]. Thus, an accurate rat model for chronic wounds induced by comorbidities remains underdeveloped, largely due to the complexity of wound formation and the physiological differences between rats and humans.

Inducing peripheral vascular lesions that lead to chronic wounds in wild-type rats remains challenging, due to their natural resistance to vascular conditions [14, 15]. One widely used technique for studying skin ischemia-reperfusion is skin flap modeling, in which wounds are created on ischemic, reperfused skin to mimic chronic wounds in humans [16–19]. To prevent basal revascularization and sustain ischemia, silicone sheets are placed beneath the flap, delaying wound healing for approximately 28 days, thereby simulating chronic wounds [20]. Existing studies have primarily focused on the impact of hyperglycemia on flap survival [21, 22], while the effects of open wounds on the viability of ischemic flaps remain poorly understood [23]. In this study, we induced diabetes in rats and created ischemic skin flap wounds of varying sizes on their backs. Silicone was placed beneath the flaps, leading to necrosis 9–12 days post-surgery. We then refined this approach to develop a modified bipedicle ischemic skin flap model, which exhibited delayed healing and partially reproduced the pathophysiological features of chronic wounds in diabetic patients.

Materials and methods

Animals

Male Sprague-Dawley rats (6 weeks old, 160–200 g) were obtained from Guizhou Huijiu Biotechnology Co., Ltd. (China; License number: SYXK-Qian-2021-0004). All animal procedures were approved by the Institutional Animal Care and Use Committee of the Zunyi Medical University (approval number: zyfy-an-2023-0055).

Diabetes induction and rat bipedicle ischemic skin wound models

Rats were housed in a specific-pathogen-free (SPF) facility maintained at 23 ± 2 °C with a 12:12-hour light–dark cycle, and were provided with food and water ad libitum. Fifty rats were fed a high-fat diet (Sichuan Chengsi Biotechnology Co., Ltd, China) for five weeks, followed by an intraperitoneal injection of STZ (35 mg/kg; Sigma-Aldrich) to induce type 2 diabetes. Random blood glucose levels exceeding 16.7 mmol/L at one week post-injection were considered indicative of successful diabetes induction. Body weight and blood glucose levels were monitored weekly; animals with significant deviations were excluded from the study (Supplementary Table 1).

At 13 weeks of age, a bipedicle ischemic skin wound model was established on the rats’ backs, with modifications based on previously published protocols [20]. Rats were randomly assigned to eight groups (Table 1; n = 5 per group). Following anesthesia induction with 2.5% isoflurane (in 100% O₂ at 1 L/min) and maintenance at 2%, dorsal fur was shaved, and a surgical marking pen was used to outline a flap centered along the spine between the scapular base and iliac crest (Fig. 1A). Based on the method described by Gould et al. [19, 20], the flap center was marked at 5.0 cm (for 11 cm flap) or 4.5 cm (for 10 cm flap) from the top. According to HE S et al. [24], two full-thickness skin defects (6 mm in diameter) were symmetrically created on both sides of the marker point using a sterile biopsy punch. A dorsal bipedicle ischemic flap was then cut deep to the panniculus carnosus muscle (Fig. 1B), and a sterilized silicone sheet (Shenzhen Inmeda Insulation Plastic Material Co., China) was placed beneath it (Fig. 1C-D). The flap and silicone sheet were sutured using 3 − 0 braided silk sutures, with stitch intervals of approximately 0.9 cm to prevent displacement (Fig. 1E). After surgery, wounds were covered with sterile gauze, and rats were housed individually with Elizabethan collars (Quike Biologicals Ltd., China) to prevent self-inflicted wound trauma. Food and water were provided ad libitum. Postoperative photographs were taken to monitor flap perfusion and wound healing. Notably, most flaps exhibited necrosis by postoperative days 9–12 (Fig. 1F). Body weight and blood glucose levels were continuously monitored.

Table 1.

Dimensions of length, width, and silicone thickness for the design of various bipedicle flaps

Flap length (cm)	Flap width (cm)	Silicone Length(cm)	Silicone width (cm)	Silicone thickness(mm)
11	2	10	1.9	0.3
11	2.5	10	2.4	0.3
11	3	10	2.9	0.3
10	3	9.1	2.9	0.3
11	2	10	1.9	0.1
11	2.5	10	2.4	0.1
11	3	10	2.9	0.1
10	3	9.1	2.9	0.1

Fig. 1.

Establishment and characterization of an ischemic trauma model in diabetic SD rats using a bipedicle flap with silicone gel implantation. A Schematic of flap design and trauma induction. Flap dimensions: length = 10–11 cm, width = 2.0, 2.5, or 3.0 cm (n = 5). B Full-thickness skin excision (6 mm diameter) followed by creation of the bipedicle flap. C Elevation of the flap from the underlying vasculature, followed by hemostasis. D Placement of a sterile silicone sheet (0.1–0.3 mm thickness) to prevent hemorrhagic adhesion. E Interrupted suturing with silicone material to secure the flap. F Progression of flap necrosis and infection observed 9–12 days post-surgery. G Changes in body weight in rats implanted with silicone gels of 0.3 mm and 0.1 mm thickness

After reviewing the relevant literature [18], we modified the procedure by removing the silicone sheet. We then acquired 45 six-week-old SD rats, Type 2 diabetes was induced in 35 of them, following the procedure described above. Diabetic rats meeting inclusion criteria were randomly divided into three groups (n = 10). Both bipedicle ischemic flap groups had a flap length of 11 cm, with widths of either 2 cm (DM-Ischemic-2.0) or 2.5 cm (DM-Ischemic-2.5). Two 6-mm-diameter skin defects were created in the DM-Acute group as classical controls. Additionally, to control for potential surgical effects, ten rats fed a low-fat diet underwent the same ischemic flap procedure as the DM-Ischemic-2.0 group, forming the NC-Ischemic-2.0 group. Body weight and blood glucose levels were monitored weekly throughout the study (Fig. 2A-G).

Fig. 2.

Construction of the ischemic trauma model and body weight statistics in diabetic SD rats. A Bipedicle ischemic skin flap (11 cm in length, 2.0–2.5 cm in width) on the dorsum of diabetic rats; a flap of identical dimensions was also established on healthy SD rats as a control. B Bilateral full-thickness wounds (6 mm diameter) introduced at the midpoint of the flap or two identical acute wounds created on the backs of diabetic rats. C Skin flap excision procedure. D Detachment of the flap from the underlying vasculature, followed by hemostasis. E Placement of interrupted sutures across the skin. F Immediate postoperative condition of the flap. (G) Changes in body weight of diabetic SD rats in the DM-Ischemic-2.0, DM-Ischemic-2.5, NC-Ischemic-2.0, and DM-Acute groups

Histology

On days 6 and 15 post-injury, animals under deep anesthesia (2.5% isoflurane) were euthanized by cervical spinal dislocation, followed by aseptic excision of the wound and with a 5-mm peripheral margin using a sterile scalpel. The wound tissue was then bisected longitudinally; one half was frozen at −80 °C for qPCR analysis, while the other was fixed in 10% neutral formalin for 48 h. After fixation, tissues were dehydrated, paraffin-embedded, and sectioned at 4-µm thickness. These sections were stained with HE and Masson’s trichrome stain to assess wound morphology. Immunohistochemical staining was performed on diabetic normal skin, as well as wound tissues and adjacent skin of ischemic flaps (11 cm long, 2 cm wide) at postoperative days 3, 6, and 15, to quantify vascular density and assess flap ischemia (CD31, 1: 500, ab182981, rabbit, Abcam). Immunofluorescence staining of 15-day tissues was conducted to assess macrophage polarization (CD206, 1: 6400, ab64693, rabbit, Abcam; iNOS, 1: 500, ab178945, rabbit, Abcam) and vascular regeneration (CD31, 1: 8000, ab182981, rabbit, Abcam). Images were acquired using an Olympus slide scanning system (Olympus Corporation, Japan). Five randomly selected fields per group were analyzed. on days 6 and 9. The number of inflammatory cells, percentage of collagen, and positive cell counts (CD206, iNOS, and CD31) per field were quantified using ImageJ. Five visual fields per group were selected for analysis.

Quantitative real‑time reverse transcriptase polymerase chain reaction (qRT‑PCR)

Total RNA was extracted from each group on day 15 using TRIzol™ reagent (Invitrogen, Thermo Fisher Scientific, USA) and reverse-transcribed into single-stranded complementary DNA (cDNA) using a reverse transcription kit (Thermo Scientific) following the manufacturer’s instructions. qPCR was conducted using ChamQ Universal SYBR qPCR Master Mix (Vazyme, Nanjing, China) on an ABI Prism thermal cycler (Applied Biosystems, CA, USA). The thermal cycling protocol included an initial denaturation at 94 °C for 3 min, followed by 35 cycles of denaturation at 94 °C for 30 s, annealing at 58 °C for 30 s, and extension at 72 °C for 45 s. Specificity of amplification was verified by melting curve analysis. Gene expression levels were normalized to β-actin, and relative gene expression was calculated using the 2^-ΔΔCT method. Primer sequences are provided in Table 2.

Table 2.

Primers for qPCR

GENE	Primer sequence FW	Primer sequence RV
IL-1β	AATCTCACAGCAGCATCTCGACAAG	TCCACGGGCAAGACATAGGTAGC
IL-6	ACTTCCAGCCAGTTGCCTTCTTG	TGGTCTGTTGTGGGTGGTATCCTC
TNF-α	CCGATTTGCCATTTCATACCAG	TCACAGAGCAATGACTCCAAAG
MMP-2	CAAGCCCAAGTGGGACAAGAATC	GTAGTGGAGTTACGTCGCTCCATAC
MMP-9	GACCTGAAAACCTCCAACCTCAC	GCTTCTGAAGCATCAGCAAAGC
VEGF-A	CTTCAAGCCGTCCTGTGT	GAAGCTCATCTCTCCTATGTGC
β-actin	TGTCACCAACTGGGACGATA	GGGGTGTTGAAGGTCTCAAA

Statistical analysis

Data with normal distribution and homogeneous variance are expressed as mean ± standard deviation (SD) and were analyzed using one-way analysis of variance (ANOVA). The Shapiro–Wilk test was applied to assess normality. Post hoc comparisons were conducted with Bonferroni correction. All statistical analyses were performed using SPSS version 29.0 (IBM, USA), with p-values < 0.05 considered statistically significant.

Results

Weight loss in diabetic mice after modeling

After successful induction of type 2 diabetes, bipedicle dorsal flaps measuring 10–11 cm in length and 2.0 cm, 2.5 cm, or 3.0 cm in width were designed (Fig. 1A). The flaps were incised, and two 6-mm full-thickness skin defects were created within each flap (Fig. 1B). To disrupt blood flow at the flap base, sterile silicone sheets of 0.1–0.3 mm thickness were inserted (Fig. 1C-D). The flaps, along with the silicone interface, were then sutured in full thickness, interrupted stitches (Fig. 1E). Nearly complete flap necrosis was observed between postoperative days 9 and 12 (Fig. 1F). Following STZ injection, the rats exhibited weight loss, which continued for two weeks after surgery. No significant difference in body weight changes was observed between diabetic rats treated with 0.1 mm and 0.3 mm sterile silica gel (46.22 ± 8.02 g vs. 49.70 ± 5.31 g). Overall, body weight decreased by approximately 20% from the start of surgical modeling (Fig. 1G).

Following protocol refinement, silicone sheets were omitted and established a bipedicle ischemic skin flap (11 cm in length and 2–2.5 cm in width) on the dorsum of diabetic rats (Fig. 2A). As a control, a flap of identical dimensions was created on healthy SD rats (Fig. 2A). In addition, bilateral 6-mm full-thickness wounds were created at the midpoint of each flap (Fig. 2B). For comparison, two identical acute wounds were generated at the corresponding sites on diabetic rats (Fig. 2B). The flap was surgically incised, and perfusion between the flap and underlying tissue was disrupted to simulate ischemia (Fig. 2C-D). The skin was subsequently closed using interrupted sutures (Fig. 2E-F). Among the improved surgical groups, the DM-Ischemic-2.0 group showed the most pronounced weight loss (50.25 ± 7.37 g), followed by the DM-Ischemic-2.5 group (45.75 ± 9.18 g) and the DM-Acute group (42.5 ± 7.85 g). By contrast, the NC-Ischemic-2.0 group exhibited weight gain (+ 19.95 ± 3.97 g) (Fig. 2G).

The 2 cm bipedicle flap wound exhibited significant ischemia

Immunohistochemical analysis revealed a mature vascular network characterized by large luminal structures in normal skin, whereas ischemic flaps (11 cm long, 2 cm wide) demonstrated a clear pathological sequence: at postoperative day 3, tissue necrosis accompanied by a complete lack of vascular staining were observed; this avascular state persisted until day 6; by day 15, extensive yet immature neovascularization was evident, featuring numerous vessels with narrow lumina (Supplementary Fig. 1A-E).

The 2 cm bipedicle flap delays wound healing

Wound healing analysis (Fig. 3A and B) showed that the DM-Acute group exhibited higher wound healing rates than all other groups on days 3, 6, 9, 12, and 15. By day 6, the wound healing rate in the DM-Acute group was significantly accelerated, and epithelialization was nearly complete by day 9. In contrast, the scab shedding still existed in the NC-Ischemic-2.0 and DM-Ischemic-2.5 groups by day 6, and the crust sloughed off by day 9. On days 12 and 15, the DM-Ischemic-2.0 group remained covered with a crust, and minimal contraction, while the other groups had largely completed epithelialization (P<0.05). HE staining showed that on both days 6 and 15 (Fig. 4A-D), the DM-Ischemic-2.0 group still had extensive skin defects covered by scab shells and infiltrated by numerous inflammatory cells, whereas the other groups had largely completed epithelialization. Inflammatory cell counts revealed statistically significant increases in all groups compared to the DM-Acute group on day 6, with the DM-Ischemic-2.0 group still exhibiting elevated levels on day 15 (P<0.05).

Fig. 3.

Wound healing and statistical analysis of the DM-Ischemic-2.0, DM-Ischemic-2.5, NC-Ischemic-2.0, and DM-Acute groups. A Gross images showing wound healing. B Statistical analysis of the healing rate in each group. * Denotes comparison between the DM-Ischemic-2.0 and DM-Acute groups; # denotes comparison between the NC-Ischemic-2.0 and DM-Acute groups; + denotes comparison between the DM-Ischemic-2.5 and DM-Acute groups; & denotes comparison between the DM-Ischemic-2.0 and NC-Ischemic-2.0 groups. Statistical comparisons were made using the Kruskal-Wallis test, with pairwise comparisons corrected by the Bonferroni method. *, #, +, & indicate P < 0.05; **, P < 0.01; ***, P < 0.001

Fig. 4.

HE staining of traumatic lesions in the DM-Ischemic-2.0, DM-Ischemic-2.5, NC-Ischemic-2.0, and DM-Acute groups at 6 and 15 days postoperatively. A HE staining at 6 days showing inflammatory cell infiltration at the traumatic site. B HE staining at 15 days showing epithelialization and continued inflammatory cell infiltration. C Inflammatory cell count at 6 days postoperatively. D Inflammatory cell infiltration at 15 days postoperatively. Comparisons between groups were performed using one-way ANOVA with pairwise comparisons corrected by the Bonferroni method. **, P < 0.01; ***, P < 0.001

The 2 cm bipedicle flap impairs collagen deposition

Masson staining at day 6 revealed significant collagen in the DM-Acute group but minimal collagen formation in the other groups (Fig. 5A and C) with significantly greater collagen content in the DM-Acute group (P<0.05). By day 15 (Fig. 5B and D), well-organized collagen fibers were evident in the DM-Acute group, whereas the DM-Ischemic-2.5 and NC-Ischemic-2.0 groups showed moderate collagen deposition with less intense staining. The DM-Ischemic-2.0 group exhibited reduced collagen synthesis (P<0.05). Additionally, qPCR analysis on day 15 revealed elevated expression of MMP2 and MMP9 in the DM-Ischemic-2.0 group compared to all other groups (Figs. 5E-F, P<0.05).

Fig. 5.

Collagen expression in wounds assessed using Masson staining at 6 and 15 days postoperatively in the DM-Ischemic-2.0, DM-Ischemic-2.5, NC-Ischemic-2.0, and DM-Acute groups. A Masson staining at 6 days. B Masson staining at 15 days. C Collagen percentage at 6 days. D Collagen percentage at 15 days. E MMP2 gene expression. F MMP9 gene expression. Comparisons between groups were conducted using one-way ANOVA, followed by Bonferroni-corrected pairwise comparisons. *, P < 0.05; **, P < 0.01; ***, P < 0.001

The wounds of 2 cm bipedicle flap slowed angiogenesis

Immunofluorescence on day 15 revealed a high density of CD31-positive cells in the DM-Ischemic-2.0 group, but few mature vascular lumens were observed. In contrast, the NC-Ischemic-2.0 group displayed the greatest number of mature vascular lumens, despite a lower total vessel count, while the DM-Acute group exhibited fewer but more dilated lumens. Additionally, a notable presence of immature vascular lumens was observed in the DM-Ischemic-2.5 group (Fig. 6A). Quantification of individual mature vessels confirmed significantly fewer mature vascular lumens in the DM-Ischemic-2.0 group (Fig. 6B, P<0.05). qPCR analysis revealed that VEGF-A expression was highest in the DM-Ischemic-2.0 group at day 15 (Fig. 6C, P<0.05).

Fig. 6.

Immunofluorescence detection of blood vessels and qPCR analysis of VEGF-A expression at 15 days postoperatively in the DM-Ischemic-2.0, DM-Ischemic-2.5, NC-Ischemic-2.0, and DM-Acute groups. A Immunofluorescence staining of CD31-positive blood vessels in each group. B Quantification of CD31-positive vessel lumen. C qPCR analysis of VEGF-A gene expression. Statistical comparisons between groups were performed using one-way ANOVA, with pairwise comparisons corrected by the Bonferroni method. **, P < 0.01; ***, P < 0.001

The wounds of 2 cm bipedicle flap sustains chronic inflammation

Immunofluorescence on day 15 revealed the lowest density of CD206-positive macrophages in the DM-Ischemic-2.0 group, and the DM-Acute group exhibited fewer CD206-positive macrophages compared to the NC-Ischemic-2.0 group (Fig. 7A and C, P<0.05). Conversely, the highest number of iNOS-positive macrophages was observed in the DM-Ischemic-2.0 group, whereas the lowest levels were seen in control groups, with elevated levels also detected in the DM-Ischemic-2.5 group relative to the NC-Ischemic-2.0 group (Fig. 7B and D, P<0.05). qPCR analysis on day 15 further demonstrated, the expression of pro-inflammatory genes IL-1β, IL-6, and TNF-α were significantly upregulated in the DM-Ischemic-2.0 group compared to the other groups (Figs. 7E-G, P<0.05), indicating the presence of a sustained pro-inflammatory microenvironment.

Fig. 7.

Immunofluorescence detection of iNOS-positive M1-type macrophages and qPCR analysis of IL-1β, IL-6, and TNF-α expression at 15 days postoperatively in the DM-Ischemic-2.0, DM-Ischemic-2.5, NC-Ischemic-2.0, and DM-Acute groups. A Staining of CD206-positive M2-type macrophages. B Staining of iNOS-positive M1-type macrophages. C Counts of CD206-positive M2 macrophages. D Counts of iNOS-positive M1 macrophages. E Expression levels of IL-1β. F Expression levels of IL-6. G Expression levels of TNF-α. Statistical comparisons between groups were performed using one-way ANOVA, with post-hoc pairwise comparisons corrected by the Bonferroni method. *, P < 0.05; **, P < 0.01; ***, P < 0.001

Discussion

The development of clinically relevant animal models remains challenging due to the inherent complexity of chronic wounds in humans, which differ substantially from their manifestations in animals. Although existing models offer valuable platforms for preclinical evaluation, none fully recapitulate the pathological features of human chronic wounds [23]. In the present study, we established a bipedicle ischemic skin flap model in diabetic rats, consisting of an 11 cm-long and 2 cm-wide flap incorporating two ischemic trabeculae on either side of the flap’s midpoint. Diabetes was induced via a high-fat diet combined with intraperitoneal injection of streptozotocin (STZ). The model is technically simple, highly reproducible, and effectively circumvents the rapid wound contraction and re-epithelialization typically observed in conventional rodent wound models. Postoperative imaging, histological assessment, and molecular analyses confirmed that, compared with the normal skin of diabetic rats, the flap exhibited marked ischemic changes by days 3 and 6 post-surgery. By day 15, substantial neovascularization was observed, suggesting a transition from ischemia to vascular regeneration. This delayed neovascularization is likely attributed to compensatory hemodynamic remodeling, which may facilitate revascularization of the injured and adjacent tissues. Compared with the conventional acute wound model in diabetic rats, our ischemic flap model exhibited significantly delayed wound healing by day 15. Notably, wounds in the DM-Ischemic-2.0 group were characterized by persistent eschar formation and extensive tissue loss. Collectively, these findings indicate that the newly developed ischemic flap model in diabetic SD rats partially mimics the pathological hallmarks of clinical diabetic foot ulcers, particularly their chronicity, ischemic burden, and impaired healing dynamics.

Disruption of the native vascular supply in the flap induced localized ischemia, establishing a suitable foundation for the creation of full-thickness skin defects that effectively mimic ischemic chronic wounds [18]. Ischemic wounds generally require 14–21 days for complete healing, whereas non-ischemic wounds in healthy rats typically re-epithelialize within 10–12 days [16, 23]. A previous study developed ischemic skin flaps measuring 10 cm in length and 3 cm in width on the dorsum of Zucker Diabetic Fatty (ZDF) rats for trauma-related research [24]. However, such genetically modified models are expensive and less feasible for large-scale experimental use [25–28], In contrast, diabetes induced via a high-fat diet combined with streptozotocin (STZ) administration offers a more cost-effective and scalable platform for ischemic wound studies [29, 30]. Initially, we also aimed to place a silicone gel sheet beneath the flap to facilitate basal revascularization, as previously described [20, 24]. However, despite modifying flap dimensions (reducing the length from 11 cm to 10 cm, increasing the width to 3 cm, and decreasing silicone thickness from 0.3 mm to 0.1 mm), complete necrosis still occurred within 9–12 days post-surgery, thereby precluding further analysis. We speculate that progressive weight loss in diabetic rats, while the volume of the implanted silicone remained constant, may have intensified ischemia in the flap. Moreover, immune responses triggered by the implanted foreign material could have further compromised flap viability [23]. In a study by Chen et al. [18], an ischemic skin flap was successfully established on the backs of healthy SD rats without the use of silicone, and chronic wounds were confirmed by postoperative day 14. Building on this approach, our model refines the procedure by narrowing the flap width to 2 cm, thereby improving flap survival while maintaining a robust ischemic environment.

Although basal blood flow partially revascularized the flap due to the absence of silicone isolation in our model, wound healing in ischemic flaps—both in diabetic and normal SD rats—remained markedly slower than in acute wounds of diabetic rats. At 15 days post-injury, the DM-Ischemic-2.0 group still displayed extensive inflammatory cell infiltration, and Masson’s trichrome staining revealed a sparse distribution of nascent collagen fibers. Immunofluorescence analysis demonstrated sustained expression of the M1 macrophage marker iNOS, alongside significantly reduced levels of the M2 marker CD206. qPCR analysis further confirmed elevated expression of VEGF-A, MMP-2, and MMP-9 in the DM-Ischemic-2.0 group, indicating that the wounds remained in an active inflammatory phase. Despite the upregulation of VEGF-A, CD31 immunofluorescence showed impaired angiogenesis, evidenced by the limited formation of mature vascular lumens [31]. Elevated MMP activity, a recognized hallmark of chronic wounds, interferes with extracellular matrix (ECM) remodeling and delays tissue regeneration [32]. In particular, MMP-9 has been shown to inhibit wound repair, while MMP-9 knockout models demonstrate enhanced healing outcomes [33]. Consistent with previous findings [34], the DM-Ischemic-2.0 group also exhibited significantly increased levels of IL-1β and TNF-α. Notably, TNF-α has been reported to upregulate MMP expression while suppressing tissue inhibitors of metalloproteinases (TIMPs), thereby disrupting ECM homeostasis and impairing wound healing [18]. Moreover, this group showed abundant M1 macrophage infiltration and accumulation of necrotic cells around the wound bed, as evidenced by HE staining. These findings suggest diminished phagocytic capacity of inflammatory macrophages, which may contribute to the failure to clear apoptotic cells [35]. The persistent presence of apoptotic debris likely promotes a pro-inflammatory microenvironment, amplifying cytokine production, elevating MMP levels, and suppressing ECM deposition—findings that are consistent with our Masson staining results. Collectively, these data indicate that unresolved inflammation and excessive proteolytic activity synergistically contribute to the pathogenesis of chronic wounds and impede effective tissue repair [32].

Cutaneous wound repair remains a major global healthcare challenge. While most wounds heal under standard care, patients with impaired angiogenesis are prone to chronic wounds complicated by infection, sepsis, or osteomyelitis [36]. Here, we established a reproducible ischemic wound model in diabetic rats that provides a scalable and clinically relevant platform to investigate chronic wound pathophysiology and evaluate therapeutic strategies. The model is simple, stable, and adaptable for preclinical studies with pharmacological or bioengineered interventions, offering important support for advancing therapies for diabetic foot ulcers. Several limitations should be noted. First, rodent–human anatomical and physiological differences prevent complete replication of human pathology. Second, unlike the progressive vascular insufficiency underlying human ulcers, our model employs acute surgical ischemia, which may not fully reflect clinical conditions. Third, the absence of a physical barrier at the flap base permitted partial revascularization, reducing the ability to sustain long-term ischemia [37]. Nevertheless, we observed persistent elevation of MMP2, MMP9, IL-6, and IL-1β in the DM-Ischemic-2.0 group 15 days post-modeling, indicating that abnormal MMP and cytokine expression contributes to a pathological microenvironment that impairs wound repair [38]. Thus, our model serves as a valuable in vivo platform for mechanistic studies and therapeutic evaluation in diabetic ischemic wounds. Future refinement with biocompatible isolation materials may further restrict neovascularization and enhance clinical relevance.

Conclusion

Our findings demonstrate that delayed wound healing occurs in diabetic SD rats following the establishment of a bipedicle ischemic flap (11 cm in length and 2 cm in width), combined with bilateral 6-mm full-thickness excisional wounds at the flap’s midpoint. This model, induced via a high-fat diet and intraperitoneal STZ administration, effectively prolongs the wound healing timeline in diabetic animals and addresses key limitations of prior models that rely on acute wounds to approximate chronic pathology. Importantly, histological and molecular analyses revealed hallmark features of diabetic chronic wounds—such as persistent inflammation, impaired angiogenesis, and delayed extracellular matrix remodeling—thereby supporting the model’s translational relevance. Together, these results establish our ischemic flap approach as a scalable, reproducible, and physiologically relevant platform for preclinical investigation of therapeutic strategies targeting chronic diabetic ulcers in humans.

Abbreviations

DFUs

Diabetic foot ulcers

SD

Sprague-Dawley

STZ

Streptozotocin

IL-1β

Interleukin-1 beta

TNF-α

Tumor necrosis factor-alpha

IL-6

Interleukin-6

MMP2

Matrix metalloproteinase 2

MMP9

Matrix metalloproteinase 9

VEGF-A

Vascular endothelial growth factor A

Authors’ contributions

Zhiyuan Liu designed and established the ischemic flap wound model, and drafted the manuscript. Ronghua Li performed histopathological analyses. Yanji Zhang conducted statistical analyses. Qi Fang and Guangchao Xu critically revised the manuscript. Mulan Qaharprepared schematic illustrations. Jing Liu executed qPCR experiments. Zairong Wei and C.D. Chengliang Deng conceived the study and supervised the experimental design. All authors reviewed and approved the final manuscript.

Funding

This study was supported by the Guizhou Provincial Department of Science and Technology project (Qiankehe Foundation-ZK [2024] General 314), the National Nature Science Foundation of China (82472567).

Data availability

Data is provided within the manuscript or supplementary information files.

Declarations

Ethics approval and consent to participate

All animal procedures were approved by the Institutional Animal Care and Use Committee of the Zunyi Medical University (approval number: zyfy-an-2023-0055).

Competing interests

The authors declare no competing interests.