Skip to main content
NCBI home page
International Wound Journal logoLink to International Wound Journal
. 2026 Mar 16;23(3):e70859. doi: 10.1111/iwj.70859

Negative‐Pressure Induces Epithelial‐Mesenchymal Transition via Thrombospondin‐1 Upregulation in Intact Diabetic Skin

Toshifumi Yamashiro 1,✉, Toshihiro Kushibiki 2, Yoshine Mayumi 2, Masato Tsuchiya 1, Miya Ishihara 2, Ryuichi Azuma 1
PMCID: PMC13098103  PMID: 41840929

ABSTRACT

Negative‐pressure wound therapy (NPWT) is widely used in clinical practise to enhance wound healing; however, its biological effects on intact skin remain poorly understood. Given the expanding applications of NPWT, understanding its impact beyond open wounds is increasingly important. This study aimed to evaluate the biological responses of intact skin to negative‐pressure, specifically focusing on epithelial‐mesenchymal transition (EMT). The effects of negative‐pressure loading were assessed using an in vitro model of non‐diabetic human keratinocytes and an in vivo model of intact diabetic mouse skin. Human keratinocytes exposed to negative‐pressure exhibited increased expression of thrombospondin‐1 (THBS1), transforming growth factor‐beta 1 (TGF‐β1), plasminogen activator inhibitor‐1 (PAI‐1), and hypoxia‐inducible factor 1‐alpha (HIF‐1α), alongside decreased epithelial markers and increased mesenchymal markers. These EMT‐related changes were mitigated by inhibiting the THBS1‐TGF‐β1 interaction. Similarly, in diabetic mice, intermittent negative‐pressure loading applied to intact dorsal skin significantly increased THBS1 and TGF‐β1 levels, resulting in epidermal and dermal thickening, and promoted hypoxic, prothrombotic and angiogenic responses, as evidenced by increased HIF‐1α, PAI‐1, fibrinogen and vascular endothelial growth factor expression. These findings suggest that negative‐pressure loading can induce EMT‐like responses and tissue remodelling in intact skin primarily via mechanisms involving the THBS1‐TGF‐β1 signalling axis. This study expands the understanding of the biological influence of NPWT beyond traditional wound treatment applications, potentially informing future therapeutic considerations and safety guidelines.

Keywords: diabetic skin, epithelial‐mesenchymal transition, keratinocytes, negative‐pressure wound therapy, thrombospondin‐1, wound remodelling

Key Points

  • Negative‐pressure loading induces EMT‐like molecular changes in intact diabetic skin.

  • THBS1 upregulation appears to mediate NP‐induced activation of the TGF‐β1 signaling axis.

  • NP exposure increases hypoxia‐related and prothrombotic markers, including HIF‐1α and PAI‐1.

  • Angiogenic responses are enhanced in intact skin under NP conditions.

  • These findings provide mechanistic insight into both the therapeutic and potential adverse effects of NPWT.

1. Introduction

Since its widespread clinical implementation following the landmark studies of Morykwas et al. [1], negative‐pressure wound therapy (NPWT) has been extensively applied to various types of wounds, including chronic wounds, traumatic injuries, infectious wounds and postoperative incisions [2, 3, 4, 5]. Its clinical efficacy is widely recognised across diverse fields, with proposed mechanisms of action including exudate management, edema reduction and stimulation of angiogenesis [6]. However, despite its broad clinical use, the molecular biological mechanisms underlying its effects, particularly regarding epidermal responses, remain poorly understood [7, 8, 9, 10].

In recent years, NPWT technology has expanded [11, 12, 13, 14, 15, 16], with novel adaptations such as closed incision negative‐pressure therapy (ciNPT) being increasingly applied not only to surgical incisions but also to intact skin adjacent to wound sites [5, 11, 17, 18]. These novel applications have broadened therapeutic indications but have simultaneously raised concerns regarding adverse effects, such as skin blistering occurring on non‐wounded skin subjected to negative‐pressure (NP) loading [5]. Issues such as inflammation and blistering at wound margins, particularly where foam contacts healthy skin, have long been recognised with conventional NPWT for open wounds. To mitigate these complications, clinicians have developed protective strategies, including the use of skin barriers and modified application techniques [19, 20]. Nonetheless, detailed investigations into the underlying biological mechanisms of these skin‐related adverse effects remain limited [20]. This lack of detailed knowledge highlights the need to better understand how NP affects intact skin, particularly in vulnerable populations such as individuals with diabetes.

Diabetic skin frequently represents a target for NPWT due to its inherently impaired healing capacity [21]. It exhibits unique biological characteristics, including chronic low‐grade inflammation, abnormal extracellular matrix (ECM) remodelling and increased susceptibility to fibrosis [22, 23, 24]. Collectively, these alterations result in enhanced skin fragility and compromised responses to mechanical stress or injury. Despite the known susceptibility of diabetic skin to NPWT‐related effects, the specific biological effects of NP loading on intact diabetic skin, independently of open wounds, remain largely unexplored.

Therefore, we investigated the biological effects of NP loading on intact diabetic skin using in vitro and in vivo experimental models. Initially, we conducted a protein screening using a membrane antibody array (Proteome Profiler; R&D Systems, Minneapolis, Minnesota, USA) to examine culture supernatants and cell lysates obtained from keratinocytes exposed to NP. This preliminary screening identified increased expression of plasminogen activator inhibitor‐1 (PAI‐1) and thrombospondin‐1 (THBS1), both known mediators of epithelial‐mesenchymal transition (EMT) [25, 26]. EMT is a biological process characterised by the loss of epithelial characteristics and the acquisition of mesenchymal traits, which is often associated with fibrosis and reduced epithelial integrity [27, 28].

Based on these preliminary findings, we hypothesised that NP loading might induce EMT‐like changes even in intact diabetic skin. To test this hypothesis, we conducted detailed protein expression analyses in cultured keratinocytes and diabetic mouse skin subjected to NP. Given the established role of transforming growth factor‐β1 (TGF‐β1) as a pivotal regulator of EMT [29, 30], the recognised function of THBS1 as a key activator of latent TGF‐β1 [26], and the known involvement of hypoxia‐inducible factor‐1α (HIF‐1α) in tissue remodelling, fibrosis and angiogenesis [31, 32], we specifically focused on these molecules in our analysis. Our study aims to clarify the molecular responses of intact diabetic skin to NP loading, especially considering the expanding clinical use of NPWT technologies such as ciNPT.

2. Materials and Methods

2.1. Ethical Approval

All animal experiments were approved by the Ethics Committee for Animal Experiments of the National Defence Medical College (approval number: 21065). All procedures were conducted in accordance with ARRIVE guidelines and relevant institutional regulations. Humane care was provided throughout the study. An electronic laboratory notebook platform was not used.

2.2. Cell Culture and Negative‐Pressure Incubation

PSVK1 human keratinocytes were obtained from the Japanese Collection of Research Bioresources Cell Bank (Osaka, Japan). Cells (6 × 105) were seeded into 35‐mm culture dishes and maintained in Keratinocyte‐SFM (Thermo Fisher Scientific; Waltham, Massachusetts, USA) supplemented with 5 ng/mL epidermal growth factor, 50 μg/mL bovine pituitary extract and 100 μg/mL streptomycin–penicillin for 24 h. Following medium replacement, cells were divided into two experimental groups: ambient pressure (AP) and NP, and incubated for an additional 24 h. The AP group was cultured under standard atmospheric conditions in a CO2 incubator. The NP group was cultured in an air‐tight container containing humidified water and equipped with a data logger for environmental monitoring, as described previously [33]. Continuous NP of 120 mmHg was applied using the RENASYS TOUCH device (Smith & Nephew; Watford, Hertfordshire, UK) via a standard accessory, RENASYS Softport (Figure S1). All experiments were conducted using biological replicates.

2.3. In Vitro Protein Analysis

Culture supernatants and cell lysates were collected from AP and NP groups after 24 h of treatment. Quantitative protein analysis was performed using enzyme‐linked immunosorbent assay (ELISA). Concentrations of THBS1, TGF‐β1 and E‐cadherin in supernatants, and PAI‐1, HIF‐1α, vimentin and alpha‐smooth muscle actin (αSMA) in cell lysates were measured using commercial ELISA kits according to the manufacturer's protocols (Table S1). Total protein concentrations in lysates were determined using the TaKaRa BCA Protein Assay Kit (Takara Bio, Shiga, Japan), and ELISA values for lysates were normalised to total protein (mg/mL).

2.4. Gene Expression Analysis in PSVK1 Cells

PSVK1 cells (6 × 105) were seeded into 35‐mm dishes and treated with either 100 μM Leucine‐Serine‐Lysine‐Leucine (LSKL) peptide (MedChemExpress, Monmouth Junction, New Jersey, USA) or 10 μL dimethyl sulfoxide (DMSO) as vehicle control in 2000 μL serum‐free medium. After 24 h of treatment, cells were further exposed to either AP or NP conditions for an additional 24 h. Total RNA was extracted using TRIzol Reagent (Thermo Fisher Scientific) and purified with the RNeasy MinElute Cleanup Kit (QIAGEN, Venlo, Netherlands). Complementary DNA was synthesised using the SuperScript IV VILO Master Mix (Thermo Fisher Scientific). Quantitative real‐time PCR was performed using the TaqMan Universal Master Mix II with UNG (Thermo Fisher Scientific) on a ViiA 7 Real‐Time PCR System (Thermo Fisher Scientific). Gene expression levels were calculated using the comparative Ct method (ΔΔCt), with 18S rRNA serving as the internal control and the vehicle + AP group used as the calibrator. TaqMan Probe IDs used in the analysis are listed in Table 1.

TABLE 1.

List of TaqMan Probe IDs used for gene analysis.

Gene TaqMan Probe ID Corresponding protein
THBS1 Hs00962908_ml THBS1
TGFB1 Hs00998133_ml TGF‐β1
CDH1 Hs01023894_ml E‐cadherin
VIM Hs00958111_ml Vimentin
ACTA2 Hs00426835_gl αSMA
18S Hs99999901_sl
Open in a new tab

Abbreviations: αSMA: alpha‐smooth muscle actin, THBS1: thrombospondin‐1, TGF‐β1: transforming growth factor‐1 beta.

2.5. Animals and Treatment

Female C57BLKS/J db/db mice, an established model for obesity and diabetes, were purchased from SLC Japan (Shizuoka, Japan). Experiments were initiated when the mice were 9 weeks old. Anaesthesia was induced using a combination of 0.75 mg/kg medetomidine hydrochloride, 4 mg/kg midazolam and 5 mg/kg butorphanol tartrate, and reversed using 0.75 mg/kg atipamezole hydrochloride. Eighteen mice were randomly divided into three groups (n = 6 per group): a control group and two treatment groups (Figure S2). The sample size was determined based on preliminary experiments and feasibility considerations, in line with prior studies using similar experimental designs. On the day before treatment (Day 0), the dorsal skin was circumferentially depilated. No surgical incision or skin defect was created throughout the experiment. On Day 1, two 2 × 3‐cm areas were bilaterally marked on the back of each mouse, with the left side designated as the non‐intervention side and the right as the intervention side. For the treatment groups, the intervention side was covered with RENASYS Granufoam (Smith & Nephew), a porous polyurethane dressing, and sealed with polyurethane film. In the NP group, negative pressure of 120 mmHg was applied continuously using the RENASYS TOUCH system (Smith & Nephew) during each 3‐h session, resulting in an intermittent loading protocol (3 h/day). This intermittent pressure duration was selected to elicit mechanobiological responses whilst minimising excessive mechanical burden on intact skin during prolonged application. The AP group underwent identical dressing procedures without aspiration. The control group received anaesthesia only and no dressing. During the non‐treatment hours, all mice were housed under standard conditions without any additional management, as the skin remained intact. Treatments were conducted daily for four consecutive days. At the conclusion of the final treatment, mice were euthanized, and skin specimens were harvested from the centre of the treated area for further analysis.

2.6. In Vivo Protein Analysis

Circular skin biopsies (1 cm in diameter) were homogenised in 1000 μL of cold phosphate‐buffered saline containing 1% Triton X‐100 (MP Biomedicals, Irvine, California, USA) and 0.1% protease inhibitor cocktail (Sigma‐Aldrich, Burlington, Massachusetts, USA) using a μT‐12 tissue homogeniser (TAITEC, Saitama, Japan) at 3200 rpm for 2 min. Homogenates were centrifuged at 15000 ×g for 5 min, and the resulting supernatants were collected for quantification of THBS1, TGF‐β1, PAI‐1, HIF‐1α and vascular endothelial growth factor (VEGF) using specific ELISA kits (Table S1). Protein concentrations were normalised per milligramme of protein. Final values were calculated as the difference in protein levels between the intervention and non‐intervention sides for each mouse.

2.7. Histological Analysis

Formalin‐fixed, paraffin‐embedded skin tissues were sectioned at 4 μm and stained with haematoxylin and eosin (HE) or processed for immunohistochemistry (IHC) using specific antibodies (Table S2). HE staining was used to assess general tissue architecture, including epidermal and dermal thicknesses. IHC was performed to evaluate the expression of THBS1 as a marker of epithelial migration; HIF‐1α, PAI‐1 and fibrinogen to assess hypoxia, fibrosis and thrombotic vascular occlusion, respectively [34]; and CD31 to evaluate angiogenesis. All images were captured at 200× magnification. Histological and immunohistochemical quantifications were performed in a blinded manner. Quantitative measurements were averaged from at least 10 randomly selected fields per sample. Image analysis was conducted using ImageJ FIJI Software (US NIH, Bethesda, Maryland, USA) [35], and results were expressed as the difference between the intervention and non‐intervention sides for each mouse.

2.8. Statistical Analysis

All statistical analyses were performed using GraphPad Prism v.9.5.1 (GraphPad, San Diego, California, USA). For comparisons between two groups, an unpaired two‐tailed t‐test was used. For comparisons amongst three or more groups, one‐way or two‐way ANOVA was conducted, followed by Tukey's honestly significant difference test for post hoc analysis. Data are presented as mean ± standard deviation, and a p‐value of < 0.05 was considered statistically significant.

3. Results

3.1. EMT‐Related Responses Induced by Negative‐Pressure in Vitro

Protein expression analysis of culture supernatants and cell lysates revealed significantly increased expression of EMT‐related proteins in the NP group compared to the AP group, including THBS1 (p = 0.008), TGF‐β1 (p = 0.008), PAI‐1 (p = 0.001) and HIF‐1α (p = 0.022) (Figure 1a). In parallel, EMT marker analysis showed significantly decreased expression of the epithelial marker E‐cadherin (p = 0.031) and increased expression of the mesenchymal marker αSMA (p = 0.031). Additionally, vimentin expression tended to be higher under NP conditions (p = 0.051) (Figure 1b).

FIGURE 1.

FIGURE 1

Open in a new tab

EMT‐related protein and marker expression induced by negative‐pressure in vitro. (a) Enzyme‐linked immunosorbent assay (ELISA) of culture supernatants and cell lysates revealed significantly increased protein levels of thrombospondin‐1 (THBS1) (p = 0.008), transforming growth factor‐beta 1 (TGF‐β1) (p = 0.008), plasminogen activator inhibitor‐1 (PAI‐1) (p = 0.001) and hypoxia‐inducible factor‐1 alpha (HIF‐1α) (p = 0.022) in keratinocytes cultured under negative‐pressure (NP) compared with ambient pressure (AP). (b) Epithelial‐mesenchymal transition (EMT) marker expression showed a significant decrease in E‐cadherin (p = 0.031) and an increase in alpha‐smooth muscle actin (αSMA) (p = 0.031) in the NP group; vimentin expression also tended to increase (p = 0.051). n = 12 for each group. Values are presented as means ± standard deviations. Statistical significance is indicated as *p < 0.05 and **p < 0.01. αSMA: Alpha‐smooth muscle actin, AP: ambient pressure, ELISA: enzyme‐linked immunosorbent assay, EMT: epithelial‐mesenchymal transition, HIF‐1α: hypoxia‐inducible factor‐1 alpha, NP: negative‐pressure, PAI‐1: plasminogen activator inhibitor‐1, TGF‐β1: transforming growth factor‐beta 1, THBS1: thrombospondin‐1.

3.2. THBS1–TGF‐β1 Blockade Suppresses EMT‐Associated Gene Expression

Two‐way ANOVA demonstrated significant main effects of treatment (vehicle vs. LSKL) and pressure conditions (AP vs. NP) on thrombospondin‐1 (THBS1) expression (treatment: p = 0.008; pressure: p = 0.006), with a significant interaction between factors (p = 0.004). Post hoc analysis indicated that THBS1 expression was significantly reduced in the NP group compared to the AP group under LSKL treatment (p < 0.001), and notably higher in the LSKL group compared to the vehicle group under AP conditions (p = 0.001).

Transforming growth factor beta 1 (TGFB1) expression showed a significant main effect of treatment, with vehicle treatment resulting in higher expression than LSKL (p = 0.004), and a trend towards lower expression in the LSKL group under AP conditions (p = 0.09). Cadherin 1 (CDH1) exhibited a significant effect of pressure (AP > NP, p < 0.001); post hoc tests confirmed significantly lower expression in the NP group under both vehicle (p = 0.007) and LSKL conditions (p = 0.03). Vimentin (VIM) demonstrated a significant treatment effect (vehicle > LSKL, p < 0.001) and a significant interaction between factors (p < 0.001); NP conditions induced higher VIM expression compared to AP under vehicle conditions (p = 0.02), whereas NP significantly decreased VIM expression under LSKL treatment (p < 0.001). Finally, actin alpha 2 (ACTA2) showed a significant treatment effect (vehicle > LSKL, p < 0.001), with notably lower expression in the LSKL group under AP and NP conditions (both p < 0.001) (Figure 2).

FIGURE 2.

FIGURE 2

Open in a new tab

Gene expression in PSVK1 keratinocytes exposed to negative‐pressure and THBS1 inhibition. Real‐time polymerase chain reaction (PCR) was performed on cultured human keratinocyte PSVK1 cells exposed to ambient pressure (AP) or negative‐pressure (NP), with or without treatment with Leucine‐Serine‐Lysine‐Leucine (LSKL), a thrombospondin‐1 (THBS1) inhibitor or dimethyl sulfoxide (DMSO), a vehicle control. TaqMan assays revealed that THBS1 expression was reduced under NP when treated with LSKL. Transforming growth factor‐beta 1 (TGFB1) expression also decreased with LSKL under AP. Cadherin 1 (CDH1), which encodes E‐cadherin, was downregulated under NP regardless of treatment. Vimentin (VIM) expression increased under NP with vehicle but showed a decreasing trend with LSKL. Actin alpha 2 (ACTA2), encoding αSMA, was consistently downregulated by LSKL treatment under both pressure conditions. Data represent eight replicates per group and were analysed using two‐way analysis of variance with Tukey's range test. Significance is indicated as *p < 0.05, **p < 0.01 and ***p < 0.001. ACTA2: actin alpha 2, αSMA: alpha‐smooth muscle actin, CDH1: Cadherin 1, DMSO: dimethyl sulfoxide, LSKL: Leucine‐Serine‐Lysine‐Leucine, PCR: polymerase chain reaction, THBS1: thrombospondin‐1, TGF‐β1: transforming growth factor‐beta 1, VIM: vimentin.

3.3. THBS1 And TGF‐β1 Upregulation in Negative‐Pressure‐Treated Diabetic Mouse Skin

IHC demonstrated significantly higher THBS1 expression in the epidermis of the NP‐treated group compared to both control and AP groups (p < 0.001) (Figure 3a). Enzyme‐linked immunosorbent assay (ELISA) of skin samples also revealed significantly increased protein levels of THBS1 (p = 0.012 vs. control, p = 0.011 vs. AP) and TGF‐β1 (p = 0.008 vs. control, p = 0.006 vs. AP) in the NP‐treated group (Figure 4).

FIGURE 3.

FIGURE 3

Open in a new tab

Histological and immunohistochemical changes in NP‐treated diabetic mouse skin. (a) Immunohistochemistry (IHC) staining was performed to evaluate the epithelial‐mesenchymal transition (EMT)‐related marker thrombospondin‐1 (THBS1). THBS1‐positive pixels in the epidermis were quantified. THBS1 positivity was significantly higher in the negative‐pressure (NP) group compared with the other two groups (p < 0.001). (b) Skin thickness was assessed in haematoxylin and eosin (HE)‐stained sections. Both epidermal and dermal thickness were significantly greater in the NP group than in the other two groups (p < 0.001). (c) IHC staining for hypoxia‐inducible factor‐1 alpha (HIF‐1α), plasminogen activator inhibitor‐1 (PAI‐1), and fibrinogen was performed to evaluate tissue hypoxia and microthrombosis induced by NP. (c1) PAI‐1 positivity in the epidermis was significantly higher in the NP group (p < 0.001). (c2) The number of HIF‐1α positive cells per mm2 in the epidermis was significantly increased in the NP group (p < 0.001). (c3) The number of fibrinogen‐positive vessels per mm2 in the dermis was significantly higher in the NP group (p < 0.001). Arrowheads indicate fibrinogen‐positive vessels. (d) Angiogenesis was evaluated using IHC for CD31. The number of CD31‐positive vessels per mm2 in the dermis was significantly higher in the NP group (p < 0.001). Arrowheads indicate CD31‐positive vessels. N = 6 each. Values are presented as means ± standard deviations. Statistical significance is indicated as ***p < 0.001. Bar: 50 μm. AP: ambient pressure, ELISA: enzyme‐linked immunosorbent assay, EMT: epithelial‐mesenchymal transition, HE: haematoxylin and eosin, HIF‐1α: hypoxia‐inducible factor‐1 alpha, IHC: immunohistochemistry, NP: negative‐pressure, PAI‐1: plasminogen activator inhibitor‐1, THBS1: thrombospondin‐1, TGF‐β1: transforming growth factor‐beta 1.

FIGURE 4.

FIGURE 4

Open in a new tab

Quantitative analysis of EMT‐ and angiogenesis‐related proteins in NP‐treated mouse skin. Enzyme‐linked immunosorbent assay (ELISA) of skin tissue samples revealed significantly increased levels of thrombospondin‐1 (THBS1), transforming growth factor‐beta 1 (TGF‐β1), plasminogen activator inhibitor‐1 (PAI‐1), hypoxia‐inducible factor‐1 alpha (HIF‐1α) and vascular endothelial growth factor (VEGF) in the negative‐pressure (NP) group compared with the control and ambient pressure (AP) groups. THBS1: p = 0.012 versus control, p = 0.011 versus AP; TGF‐β1: p = 0.008 versus control, p = 0.006 versus AP; PAI‐1: p < 0.001 versus both; HIF‐1α: p = 0.004 versus control, p = 0.001 versus AP; VEGF: p < 0.001 versus both. N = 6 per group. Values are presented as means ± standard deviations. Statistical significance is indicated as *p < 0.05, **p < 0.01 and ***p < 0.001. AP: ambient pressure, ELISA: enzyme‐linked immunosorbent assay, EMT: epithelial‐mesenchymal transition, HIF‐1α: hypoxia‐inducible factor‐1 alpha, NP: negative‐pressure, PAI‐1: plasminogen activator inhibitor‐1, THBS1: thrombospondin‐1, TGF‐β1: transforming growth factor‐beta 1, VEGF: vascular endothelial growth factor.

3.4. Epidermal and Dermal Thickening Under Negative‐Pressure

Histological analysis using HE staining revealed significantly greater epidermal and dermal thickness in NP‐treated skin compared to both control and AP‐treated skin (all p < 0.001) (Figure 3b).

3.5. Hypoxic and Prothrombotic Responses to Negative‐Pressure

IHC analysis showed significantly increased epidermal expression of PAI‐1 in the NP group compared to the control and AP groups (p < 0.001). The number of HIF‐1α‐positive cells per mm2 was also significantly higher under NP conditions (p < 0.001). Furthermore, the number of fibrinogen‐positive vessels per mm2 in the dermis significantly increased in the NP‐treated group (p < 0.001) (Figure 3c). Consistently, ELISA demonstrated significantly higher protein levels of HIF‐1α (p = 0.004 vs. control, p = 0.001 vs. AP) and PAI‐1 (p < 0.001) in the NP‐treated skin samples compared to control and AP groups (Figure 4).

3.6. Angiogenesis Enhancement Under Negative‐Pressure Conditions

IHC analysis demonstrated significantly increased density of CD31‐positive vessels per mm2 in the dermis of the NP‐treated group compared to the other groups (p < 0.001) (Figure 3d). In agreement, ELISA analysis showed significantly increased VEGF protein levels in the NP‐treated group (p < 0.001) (Figure 4).

4. Discussion

This study demonstrates that NP loading is associated with EMT‐like changes and angiogenic responses in intact diabetic skin. Our investigation began with preliminary protein screening using a membrane antibody array, which identified increased expression of THBS1 and PAI‐1 in keratinocyte cultures exposed to NP. Both proteins are associated with EMT, a process closely linked to tissue remodelling and fibrosis. Based on these initial findings, we focused our subsequent analyses on factors related to EMT and angiogenesis to better understand the biological effects of NPWT, particularly on intact skin without wounds. Notably, these molecular changes were observed not only in vivo but also in monocultured human keratinocytes. Our cellular model consisted of monolayer cultures exposed to low‐pressure conditions in liquid media without direct physical contact with foam [33]. This approach excluded confounding factors such as microdeformation and immune responses typically associated with standard NPWT. Such conditions strongly suggest that epidermal responses observed primarily reflected the effects of NP rather than secondary consequences of foreign‐body reactions. Amongst the proteins differentially expressed, THBS1 was of particular interest as it has recently been identified as a mechanotransduction molecule linking mechanical stimuli to biochemical responses [36, 37]. Its role in conditions such as aneurysm formation [38] and osteogenesis [26] has been reported previously. Our findings suggest it may similarly mediate mechanical stress during NPWT.

Previous research has primarily focused on the beneficial effects of NPWT, such as promoting granulation tissue formation, enhancing local blood flow and reducing bacterial loads [39]. However, the potential of NPWT to induce adverse epithelial changes has received comparatively limited attention. Nuutila et al. [40] investigated gene expression profiles at skin graft donor sites treated with NPWT in burn patients, reporting a unique epithelial gene expression pattern compared to conventionally treated wounds. Although NPWT accelerated early‐phase epithelialization, their data suggested possible impairment in long‐term epithelial recovery due to disrupted differentiation. Our findings potentially provide a mechanistic explanation for this observation, as EMT is typically characterised by compromised epithelial integrity and function [27, 41].

During EMT, epithelial cells acquire mesenchymal characteristics, playing essential roles in embryonic development, wound healing and cancer progression [42, 43]. In skin wound healing specifically, EMT contributes to re‐epithelialization from wound edges, scar formation and subsequent fibrosis [27, 28]. TGF‐β1, a crucial EMT inducer, is vital for wound repair and tissue regeneration [26, 29, 30]. THBS1, an ECM glycoprotein, is a primary physiological activator of TGF‐β1, regulating EMT through the TGF‐β1/SMAD pathway [26, 44]. In skin, THBS1‐positive keratinocytes facilitate epidermal wound healing [45]. Consistent with these established roles, our study demonstrated increased expression of EMT‐related proteins, including THBS1, TGF‐β1, PAI‐1 and HIF‐1α, following NP loading in vitro and in vivo (Figures 1a and 4). Furthermore, EMT marker analysis in human keratinocytes revealed decreased epithelial markers and increased mesenchymal markers upon NP treatment (Figures 1b and 2). These molecular changes were notably suppressed by the peptide inhibitor LSKL (Figure 2), which specifically blocks THBS1‐mediated activation of TGF‐β1 [46], indicating a central role for THBS1 in NP‐induced EMT. These findings align with previous studies reporting similar protein expression patterns in response to NP conditions [9].

Furthermore, the NP environment may mimic pathophysiological conditions observed in pressure ulcers (PU), where tissue ischemia and microthrombosis are critical factors. Kaneko et al. [34] demonstrated that the HIF‐1α‐PAI‐1 axis contributes significantly to thrombus formation and subsequent tissue damage in PU. Although PAI‐1 is involved in normal wound repair, excessive expression of this protein promotes scar proliferation, tissue fibrosis, and impairs keratinocyte migration [25, 47, 48]. In our study, NP loading resulted in increased expression of HIF‐1α and PAI‐1, formation of fibrinogen‐positive vessels, and thickening of skin layers (Figures 1a, 3b,c, 4), suggesting hypoxia driven tissue remodelling. Collectively, these results indicate that NPWT‐induced epithelial changes may result from multiple interrelated mechanisms, including EMT‐driven structural destabilisation and ischemia‐mediated microvascular injury, partially resembling aspects of PU pathogenesis.

One of the most established mechanisms of NPWT is its capacity to enhance local blood flow [49, 50]. This occurs due to the pressure difference between the internal and external areas of the dressing, which compresses the wound surface and subsequently induces angiogenesis via the HIF‐1α‐VEGF signalling pathway in response to tissue ischemia [31, 32]. This angiogenic effect is widely recognised as central to the therapeutic efficacy of NPWT [49, 50]. Building on this principle, a novel strategy termed preconditioning NPWT has emerged, whereby NP is applied to intact skin prior to flap surgery to improve subsequent flap perfusion [51, 52]. In contrast to traditional NPWT methods that target open wounds, this preconditioning approach, similar to ciNPT, specifically targets intact skin. In our experiments, NP loading significantly enhanced the expression of HIF‐1α and VEGF and promoted CD31‐positive vessel formation in diabetic mouse skin (Figures 1a, 3c,d, 4). Similar angiogenic responses were also observed in vitro using keratinocyte monolayer cultures, suggesting that NPWT can promote angiogenesis through the HIF‐1α‐VEGF axis even in intact skin. These findings reinforce the potential use of NPWT as a non‐surgical preconditioning technique to enhance flap viability [51, 52]. Nevertheless, applying NPWT directly to healthy skin, as in ciNPT or preconditioning methods, raises concerns regarding potential adverse effects, particularly epithelial disruption related to EMT processes. Given our previous findings that NPWT enhances keratinocyte migration [33], the current results provide additional mechanistic insight supporting the beneficial role of NPWT in skin graft stabilisation and flap survival, particularly in postoperative skin‐grafting scenarios [53].

Ranieri et al. [54] reported enhanced EMT in keratinocytes under simulated microgravity conditions. Our findings extend this observation, demonstrating that EMT‐like changes can occur even under the relatively mild mechanical stress of approximately 0.1–0.15 atm, as typically applied in clinical NPWT. This suggests that even low‐intensity mechanical cues may be sufficient to trigger epithelial plasticity. To our knowledge, this study is amongst the first to propose a potential link between NPWT and EMT, highlighting important implications for both therapeutic benefits and possible adverse effects of NPWT on intact skin.

A key limitation of our study is the mismatch between the in vitro and in vivo models: human keratinocytes from non‐diabetic donors were used for the in vitro experiments, whereas diabetic mice were employed in the in vivo studies. Nevertheless, consistent changes in EMT‐ and angiogenesis‐related markers were observed across both models, suggesting partial conservation of keratinocyte responses to mechanical stress. Systemic factors associated with diabetes, however, may still influence the magnitude and regulation of these responses. Diabetic skin exhibits chronic inflammation, increased fibrosis and impaired tissue repair [22, 23, 24], all of which potentially influence keratinocyte behaviour, including EMT, under mechanical stress conditions. Indeed, previous studies have demonstrated that hyperglycemic conditions can modulate EMT in keratinocytes [55]. Future studies should therefore incorporate diabetic keratinocytes in vitro and apply NPWT to non‐diabetic skin in vivo to comprehensively assess how disease status modifies the cellular response to NP.

Additionally, Nuutila et al. [40] suggested that keratinocyte behaviour under NPWT evolves over time. Further studies are thus required to clarify temporal aspects of NPWT‐induced EMT, particularly to distinguish beneficial epithelialization effects from potential epithelial damage arising from prolonged NPWT application. Such insights would be valuable for optimising NPWT protocols, enabling clinicians to maximise wound healing efficacy whilst minimising potential adverse effects.

Collectively, our findings reveal the dual biological effects of NPWT on intact diabetic skin, emphasising its capability to promote vascular remodelling and epithelial mobilisation, as well as its potential to induce pathological tissue remodelling through EMT. These insights can inform safer and more effective clinical applications of NPWT, including ciNPT, flap preconditioning and post‐grafting care. Furthermore, a deeper understanding of epithelial plasticity in response to mechanical stimuli will be essential for refining NPWT protocols. Achieving an optimal balance between regenerative benefits and the risk of epithelial disruption will ultimately ensure safer, more effective patient care across various clinical settings.

Funding

This study was partially supported by the JSPS KAKENHI (20K12725).

Ethics Statement

All animal experiments were approved by the Ethics Committee for Animal Experiments of the National Defence Medical College (approval number: 21065). All procedures were conducted in accordance with ARRIVE guidelines and relevant institutional regulations. Humane care was provided throughout the study.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Data S1: iwj70859‐sup‐0001‐Supinfo.pdf.

IWJ-23-e70859-s001.pdf (291KB, pdf)

Acknowledgements

We would like to thank Professor Takahiro Nakamura (Department of Mathematics, National Defence Medical College) for advice regarding statistical analysis, and Editage (www.editage.com) for English language editing. The authors created Figures S1 and S2.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

References

  • 1. Morykwas M. J., Argenta L. C., Shelton‐Brown E. I., and McGuirt W., “Vacuum‐Assisted Closure: A New Method for Wound Control and Treatment: Animal Studies and Basic Foundation,” Annals of Plastic Surgery 38 (1997): 553–562, 10.1097/00000637-199706000-00001. [DOI] [PubMed] [Google Scholar]
  • 2. Li W. and Zheng J., “Negative Pressure Wound Therapy for Chronic Wounds,” Annals of Plastic Surgery 93 (2024): S19–S26, 10.1097/sap.0000000000003891. [DOI] [PubMed] [Google Scholar]
  • 3. Milcheski D. A., Clivatti G. M., Santos Junior R. A., González C. V. S., A. A. Monteiro, Jr. , and Gemperli R., “Effectiveness of Negative‐Pressure Wound Therapy With Instillation Compared to Standard Negative‐Pressure Wound Therapy and Traditional Gauze Layer Dressing for the Treatment of Acute Traumatic Wounds: A Randomized Controlled Trial,” Journal of Plastic, Reconstructive & Aesthetic Surgery 100 (2025): 208–218, 10.1016/j.bjps.2024.11.005. [DOI] [PubMed] [Google Scholar]
  • 4. Steingrimsson S., Gottfredsson M., Gudmundsdottir I., Sjogren J., and Gudbjartsson T., “Negative‐Pressure Wound Therapy for Deep Sternal Wound Infections Reduces the Rate of Surgical Interventions for Early Re‐Infections,” Interactive Cardiovascular and Thoracic Surgery 15 (2012): 406–410, 10.1093/icvts/ivs254. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5. Norman G., Shi C., Goh E. L., et al., “Negative Pressure Wound Therapy for Surgical Wounds Healing by Primary Closure,” Cochrane Database of Systematic Reviews 2022, no. 4 (2022): CD009261, 10.1002/14651858.CD009261.pub7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Yamashiro T., Kushibiki T., Mayumi Y., Tsuchiya M., Ishihara M., and Azuma R., “Negative‐Pressure Wound Therapy: What We Know and What We Need to Know,” Advances in Experimental Medicine and Biology 1436 (2023): 131–152, 10.1007/5584_2023_773. [DOI] [PubMed] [Google Scholar]
  • 7. Hsu C. C., Tsai W. C., Chen C. P., Lu Y. M., and Wang J. S., “Effects of Negative Pressures on Epithelial Tight Junctions and Migration in Wound Healing,” American Journal of Physiology. Cell Physiology 299 (2010): C528–C534, 10.1152/ajpcell.00504.2009. [DOI] [PubMed] [Google Scholar]
  • 8. Hsu C. C., Chow S. E., Chen C. P., et al., “Negative Pressure Accelerated Monolayer Keratinocyte Healing Involves cdc42 Mediated Cell Podia Formation,” Journal of Dermatological Science 70 (2013): 196–203, 10.1016/j.jdermsci.2013.03.007. [DOI] [PubMed] [Google Scholar]
  • 9. Chow S. E., Chen C. P., Hsu C. C., Tsai W. C., Wang J. S., and Hsu N. C., “Quantifying Cell Behaviors in Negative‐Pressure Induced Monolayer Cell Movement,” Biomedical Journal 39 (2016): 50–59, 10.1016/j.bj.2015.08.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 10. Huang C. H., Hsu C. C., Chen C. P., et al., “Negative Pressure Induces p120‐Catenin‐Dependent Adherens Junction Disassembly in Keratinocytes During Wound Healing,” Biochimica et Biophysica Acta 1863 (2016): 2212–2220, 10.1016/j.bbamcr.2016.05.017. [DOI] [PubMed] [Google Scholar]
  • 11. Wolvos T., “The Use of Negative Pressure Wound Therapy With an Automated, Volumetric Fluid Administration: An Advancement in Wound Care,” Wounds 25 (2013): 75–83. [PubMed] [Google Scholar]
  • 12. Sharp E., “Single‐Use Npwt for the Treatment of Complex Orthopaedic Surgical and Trauma Wounds,” Journal of Wound Care 22 (2013): S5–S9, 10.12968/jowc.2013.22.Sup10.S5. [DOI] [PubMed] [Google Scholar]
  • 13. van den Bulck R., Siebers Y., Zimmer R., Acton C., Janzing H., and Lang W., “Initial Clinical Experiences With a New, Portable, Single‐Use Negative Pressure Wound Therapy Device,” International Wound Journal 10 (2013): 145–151, 10.1111/j.1742-481X.2012.00954.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14. Borgquist O., Ingemansson R., and Malmsjo M., “The Effect of Intermittent and Variable Negative Pressure Wound Therapy on Wound Edge Microvascular Blood Flow,” Ostomy/Wound Management 56 (2010): 60–67. [PubMed] [Google Scholar]
  • 15. Lee K. N., Ben‐Nakhi M., Park E. J., and Hong J. P., “Cyclic Negative Pressure Wound Therapy: An Alternative Mode to Intermittent System,” International Wound Journal 12 (2015): 686–692, 10.1111/iwj.12201. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16. Bertelsen C. A., Fabricius R., Kleif J., Kristensen B., and Gogenur I., “Outcome of Negative‐Pressure Wound Therapy for Open Abdomen Treatment After Nontraumatic Lower Gastrointestinal Surgery: Analysis of Factors Affecting Delayed Fascial Closure in 101 Patients,” World Journal of Surgery 38 (2014): 774–781, 10.1007/s00268-013-2360-7. [DOI] [PubMed] [Google Scholar]
  • 17. Scalise A., Calamita R., Tartaglione C., et al., “Improving Wound Healing and Preventing Surgical Site Complications of Closed Surgical Incisions: A Possible Role of Incisional Negative Pressure Wound Therapy. A Systematic Review of the Literature,” International Wound Journal 13 (2016): 1260–1281, 10.1111/iwj.12492. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18. Kirkham A. M., Candeliere J., McIsaac D., et al., “Efficacy of Strategies Intended to Prevent Surgical Site Infection After Lower Limb Revascularization Surgery: A Systematic Review and Meta‐Analysis of Randomized Controlled Trials,” Annals of Surgery 278 (2023): e447–e456, 10.1097/SLA.0000000000005867. [DOI] [PubMed] [Google Scholar]
  • 19. Rentea R. M., Somers K. K., Cassidy L., Enters J., and Arca M. J., “Negative Pressure Wound Therapy in Infants and Children: A Single‐Institution Experience,” Journal of Surgical Research 184 (2013): 658–664, 10.1016/j.jss.2013.05.056. [DOI] [PubMed] [Google Scholar]
  • 20. de Jesus L. E., Martins A. B., Oliveira P. B., Gomes F., Leve T., and Dekermacher S., “Negative Pressure Wound Therapy in Pediatric Surgery: How and When to Use,” Journal of Pediatric Surgery 53 (2018): 585–591, 10.1016/j.jpedsurg.2017.11.048. [DOI] [PubMed] [Google Scholar]
  • 21. Chen L., Zhang S., da J., et al., “A Systematic Review and Meta‐Analysis of Efficacy and Safety of Negative Pressure Wound Therapy in the Treatment of Diabetic Foot Ulcer,” Annals of Palliative Medicine 10 (2021): 10830–10839, 10.21037/apm-21-2476. [DOI] [PubMed] [Google Scholar]
  • 22. Huang Y. and Kyriakides T. R., “The Role of Extracellular Matrix in the Pathophysiology of Diabetic Wounds,” Matrix Biology Plus 6‐7 (2020): 100037, 10.1016/j.mbplus.2020.100037. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Bermudez D. M., Herdrich B. J., Xu J., et al., “Impaired Biomechanical Properties of Diabetic Skin,” American Journal of Pathology 178 (2011): 2215–2223, 10.1016/j.ajpath.2011.01.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24. Tuleta I. and Frangogiannis N. G., “Diabetic Fibrosis,” Biochimica et Biophysica Acta ‐ Molecular Basis of Disease 1867 (2021): 166044, 10.1016/j.bbadis.2020.166044. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25. Ghosh A. K. and Vaughan D. E., “Pai‐1 in tissue fibrosis,” Journal of Cellular Physiology 227 (2012): 493–507, 10.1002/jcp.22783. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26. Deng R., Li C., Wang X., et al., “Periosteal cd68(+) f4/80(+) Macrophages Are Mechanosensitive for Cortical Bone Formation by Secretion and Activation of Tgf‐beta1,” Advanced Science 9 (2022): e2103343, 10.1002/advs.202103343. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27. Haensel D. and Dai X., “Epithelial‐To‐Mesenchymal Transition in Cutaneous Wound Healing: Where We Are and Where We Are Heading,” Developmental Dynamics 247 (2018): 473–480, 10.1002/dvdy.24561. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28. Yuan F. L., Sun Z. L., Feng Y., et al., “Epithelial‐Mesenchymal Transition in the Formation of Hypertrophic Scars and Keloids,” Journal of Cellular Physiology 234 (2019): 21662–21669, 10.1002/jcp.28830. [DOI] [PubMed] [Google Scholar]
  • 29. Sistigu A., Di Modugno F., Manic G., and Nistico P., “Deciphering the Loop of Epithelial‐Mesenchymal Transition, Inflammatory Cytokines and Cancer Immunoediting,” Cytokine & Growth Factor Reviews 36 (2017): 67–77, 10.1016/j.cytogfr.2017.05.008. [DOI] [PubMed] [Google Scholar]
  • 30. Koike Y., Yozaki M., Utani A., and Murota H., “Fibroblast Growth Factor 2 Accelerates the Epithelial‐Mesenchymal Transition in Keratinocytes During Wound Healing Process,” Scientific Reports 10 (2020): 18545, 10.1038/s41598-020-75584-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Erba P., Ogawa R., Ackermann M., et al., “Angiogenesis in Wounds Treated by Microdeformational Wound Therapy,” Annals of Surgery 253 (2011): 402–409, 10.1097/SLA.0b013e31820563a8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32. Biermann N., Geissler E. K., Brix E., et al., “Oxygen Levels During Negative Pressure Wound Therapy,” Journal of Tissue Viability 28 (2019): 223–226, 10.1016/j.jtv.2019.09.001. [DOI] [PubMed] [Google Scholar]
  • 33. Yamashiro T., Kushibiki T., Mayumi Y., Tsuchiya M., Ishihara M., and Azuma R., “Novel Cell Culture System for Monitoring Cells During Continuous and Variable Negative‐Pressure Wound Therapy,” Skin Research and Technology 29 (2023): e13262, 10.1111/srt.13262. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34. Kaneko M., Minematsu T., Yoshida M., et al., “Compression‐Induced Hif‐1 Enhances Thrombosis and Pai‐1 Expression in Mouse Skin,” Wound Repair and Regeneration 23 (2015): 657–663, 10.1111/wrr.12312. [DOI] [PubMed] [Google Scholar]
  • 35. Schindelin J., Arganda‐Carreras I., Frise E., et al., “Fiji: An Open‐Source Platform for Biological‐Image Analysis,” Nature Methods 9 (2012): 676–682, 10.1038/nmeth.2019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36. Wang N., Butler J. P., and Ingber D. E., “Mechanotransduction Across the Cell Surface and Through the Cytoskeleton,” Science 260 (1993): 1124–1127, 10.1126/science.7684161. [DOI] [PubMed] [Google Scholar]
  • 37. Reichelt J., “Mechanotransduction of Keratinocytes in Culture and in the Epidermis,” European Journal of Cell Biology 86 (2007): 807–816, 10.1016/j.ejcb.2007.06.004. [DOI] [PubMed] [Google Scholar]
  • 38. Yamashiro Y., Thang B. Q., Ramirez K., et al., “Matrix Mechanotransduction Mediated by Thrombospondin‐1/Integrin/Yap in the Vascular Remodeling,” Proceedings of the National Academy of Sciences of the United States of America 117 (2020): 9896–9905, 10.1073/pnas.1919702117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39. Orgill D. P., Manders E. K., Sumpio B. E., et al., “The Mechanisms of Action of Vacuum Assisted Closure: More to Learn,” Surgery 146 (2009): 40–51, 10.1016/j.surg.2009.02.002. [DOI] [PubMed] [Google Scholar]
  • 40. Nuutila K., Siltanen A., Peura M., et al., “Gene Expression Profiling of Negative‐Pressure‐Treated Skin Graft Donor Site Wounds,” Burns 39 (2013): 687–693, 10.1016/j.burns.2012.09.014. [DOI] [PubMed] [Google Scholar]
  • 41. Kalluri R. and Weinberg R. A., “The Basics of Epithelial‐Mesenchymal Transition,” Journal of Clinical Investigation 119 (2009): 1420–1428, 10.1172/jci39104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42. Nieto M. A., Huang R. Y., Jackson R. A., and Thiery J. P., “Emt: 2016,” Cell 166 (2016): 21–45, 10.1016/j.cell.2016.06.028. [DOI] [PubMed] [Google Scholar]
  • 43. Dongre A. and Weinberg R. A., “New Insights Into the Mechanisms of Epithelial‐Mesenchymal Transition and Implications for Cancer,” Nature Reviews. Molecular Cell Biology 20 (2019): 69–84, 10.1038/s41580-018-0080-4. [DOI] [PubMed] [Google Scholar]
  • 44. Jinesh G. G. and Brohl A. S., “Classical Epithelial‐Mesenchymal Transition (Emt) and Alternative Cell Death Process‐Driven Blebbishield Metastatic‐Witch (Bmw) Pathways to Cancer Metastasis,” Signal Transduction and Targeted Therapy 7 (2022): 296, 10.1038/s41392-022-01132-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 45. Siriwach R., Ngo A. Q., Higuchi M., et al., “Single‐Cell RNA Sequencing Identifies a Migratory Keratinocyte Subpopulation Expressing thbs1 in Epidermal Wound Healing,” iScience 25 (2022): 104130, 10.1016/j.isci.2022.104130. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46. Kuroki H., Hayashi H., Nakagawa S., et al., “Effect of Lskl Peptide on Thrombospondin 1‐Mediated Transforming Growth Factor β Signal Activation and Liver Regeneration After Hepatectomy in an Experimental Model,” British Journal of Surgery 102 (2015): 813–825, 10.1002/bjs.9765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47. Providence K. M., Higgins S. P., Mullen A., et al., “Serpine1 (Pai‐1) is Deposited Into Keratinocyte Migration “Trails” and Required for Optimal Monolayer Wound Repair,” Archives of Dermatological Research 300 (2008): 303–310, 10.1007/s00403-008-0845-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48. Qi L., Higgins S. P., Lu Q., et al., “Serpine1 (Pai‐1) is a Prominent Member of the Early g0 –> g1 Transition “Wound Repair” Transcriptome in p53 Mutant Human Keratinocytes,” Journal of Investigative Dermatology 128, no. 3 (2008): 749–753, 10.1038/sj.jid.5701068. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49. Ma Z., Shou K., Li Z., Jian C., Qi B., and Yu A., “Negative Pressure Wound Therapy Promotes Vessel Destabilization and Maturation at Various Stages of Wound Healing and Thus Influences Wound Prognosis,” Experimental and Therapeutic Medicine 11 (2016): 1307–1317, 10.3892/etm.2016.3083. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 50. Ma Z., Li Z., Shou K., et al., “Negative Pressure Wound Therapy: Regulating Blood Flow Perfusion and Microvessel Maturation Through Microvascular Pericytes,” International Journal of Molecular Medicine 40 (2017): 1415–1425, 10.3892/ijmm.2017.3131. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51. Rhodius P., Haddad A., Matsumine H., et al., “Non‐Invasive Flap Preconditioning by Foam‐Mediated External Suction Improves the Survival of Fasciocutaneous Axial‐Pattern Flaps in a Type 2 Diabetic Murine Model,” Plastic and Reconstructive Surgery 142 (2018): 872e–883e, 10.1097/PRS.0000000000005038. [DOI] [PubMed] [Google Scholar]
  • 52. Hong Y. G., Kim S. C., and Koh K. S., “Flap Preconditioning With the Cyclic Mode (Triangular Waveform) of Pressure‐Controlled Cupping in a Rat Model: An Alternative Mode to the Continuous System,” Plastic and Reconstructive Surgery 143 (2019): 88e–98e, 10.1097/PRS.0000000000005104. [DOI] [PubMed] [Google Scholar]
  • 53. Tsuchiya M., Kushibiki T., Yamashiro T., Mayumi Y., Ishihara M., and Azuma R., “Continuous Negative‐Pressure Wound Therapy Improves the Survival Rate of Skin Grafts and Shortens the Time Required for Skin Graft Survival,” Skin Research and Technology 30 (2024): e13865, 10.1111/srt.13865. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54. Ranieri D., Proietti S., Dinicola S., et al., “Simulated Microgravity Triggers Epithelial Mesenchymal Transition in Human Keratinocytes,” Scientific Reports 7 (2017): 538, 10.1038/s41598-017-00602-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55. Tan M. W. Y., Tan W. R., Kong Z. Q., et al., “High Glucose Restraint of Acetylcholine‐Induced Keratinocyte Epithelial‐Mesenchymal Transition Is Mitigated by p38 Inhibition,” Journal of Investigative Dermatology 141 (2021): 1438–1449.e1439, 10.1016/j.jid.2020.10.026. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Data S1: iwj70859‐sup‐0001‐Supinfo.pdf.

IWJ-23-e70859-s001.pdf (291KB, pdf)

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Articles from International Wound Journal are provided here courtesy of Wiley

RESOURCES