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. 2025 Sep 16;14(9):e70160. doi: 10.1002/jev2.70160

Epidermal Stem Cell‐Derived Extracellular Vesicles Induce Fibroblasts Mesenchymal‐Epidermal Transition to Alleviate Hypertrophic Scar Formation via miR‐200s Inhibition of ZEB1 and 2

Miao Zhen 1, Juntao Xie 2, Rui Yang 1, Lijuan Liu 1, Hengdeng Liu 1, Xuefeng He 1, Suyue Gao 1, Junyou Zhu 1, Jingting Li 3, Bin Shu 1,✉, Peng Wang 1,✉
PMCID: PMC12438467  PMID: 40955457

ABSTRACT

Hypertrophic scar (HS) is a prevalent yet unresolved wound healing complication characterized by persistent hyperactive and proliferative fibroblasts, leading to excessive extracellular matrix (ECM) synthesis and collagen contraction. Our previous studies have identified epidermal stem cells (ESCs) as critical for wound healing and HS remodelling, with its extracellular vesicles (EVs) playing a vital role. However, the specific mechanisms remain unclear. In this study, we first discovered that ESC‐EVs could effectively induce the mesenchymal‐epidermal transition (MET) of HS fibroblasts (HSFs) and inhibit their biological activity. Furthermore, by next‐generation sequencing and multiplexed CRISPR/Cas9 system, we elucidated that this therapeutic effect is mediated by the miR‐200 family (miR‐200s) encapsulated in ESC‐EVs, which targeted and inhibited ZEB1 and ZEB2 in HSFs. This vital role and mechanism have been thoroughly validated in both in vitro cell experiments and in vivo rat tail HS (RHS) models. These findings not only shed light on a previously unidentified mechanism of ESC‐EVs for HS, but also provide potential novel targets and strategies for its precise treatment.

Keywords: epidermal stem cell, extracellular vesicles, hypertrophic scar, mesenchymal‐epidermal transition, miR‐200s, ZEBs

Schematic illustration of ESC‐EVs induced HSFs' MET to alleviate HS via the miR‐200s/ZEBs axis. Epidermal stem cells (ESC) were cultured in cell factories to obtain sufficient extracellular vesicles (ESC‐EVs). EVs were isolated using a differential ultracentrifugation method. Both in vitro cell experiments and in vivo rat tail HS (RHS) models thoroughly validated that ESC‐EVs were internalized and effectively inducing the MET of HSFs and inhibited its activation to attenuate fibrosis via the miR‐200s/ZEBs axis.

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1. Introduction

Hypertrophic scar (HS) emerges as a frequent consequence following dermal insults such as burns, trauma, puncture wounds, acne, surgical incisions and vaccinations (Ogawa 2017). Clinically, HS presents patients with tremendous challenges by various symptoms, including pruritus, pain, paresthesia, erosion of underlying bone structure and restriction of joint mobility (Berman et al. 2017). However, despite the array of therapeutic approaches available, from corticosteroids and pressure therapy to laser treatments and surgical excision, the therapeutic effect of HS remains unsatisfactory (Rabello et al. 2014). Urgent efforts are warranted to identify precise intervention targets and develop novel strategies for HS.

Recent studies have demonstrated that epithelial‐mesenchymal transition (EMT) is crucial in tissue fibrosis and scar formation (Youssef and Nieto 2024). EMT represents a ubiquitous biological process where polarized epithelial cells undergo loss of cell polarity and intercellular adhesion and acquire a mesenchymal cell phenotype, which culminates in heightened deposition of extracellular matrix (ECM), augmented resistance to apoptosis, and intensified migratory and invasive potential (Boyer et al. 2020). Although EMT is necessary for proper re‐epithelialization in routine wound healing, the uncontrolled continued transition from epithelial cells to HS fibroblasts (HSFs) eventually lead to aberrant ECM deposition, collagen contraction and disrupted structural integrity (Machesney et al. 1998). Therefore, inhibiting epithelial cells’ EMT or even inducing HSFs mesenchymal‐epithelial transition (MET) should be highly efficient in alleviating HS.

Emerging evidence suggests that extracellular vesicles (EVs) regulate EMT or MET and mediate the development of various fibrosis diseases (Jiang et al. 2022). As a crucial subset of epithelial cells and progenitors of diverse epidermal cell lineages, epidermal stem cells (ESCs) possess the capacity to proliferate and differentiate into various epidermal cell types, usually showing impeccable efficiency for wound healing and the mitigation of scar formation than other kind of stem cells (Blanpain and Fuchs 2006; Moore and Lemischka 2006). Encompassing a diverse array of proteins, DNA and RNA that mirror the physiological profile of ESCs, EVs derived from ESC (ESC‐EVs) mimic the functional properties of ESCs and present considerable potential as a substitute for cellular therapy. Encouragingly, our recent studies have discovered that ESC‐EVs have an ESCs’ identical therapeutic effect in wound healing and scar prevention (Wang et al. 2017; Wang et al. 2019; Wang et al. 2022), and exhibited a potential to modulate EMT and MET. However, its crucial mechanism urgently needs to be further explored.

In this study, we established an efficient ESC‐EVs isolation and application system and thoroughly validated their effects on MET induction and biological function inhibition of HSFs in vitro and in vivo. Furthermore, using next‐generation sequencing and multiplexed CRISPR/Cas9 system, we delineated that ESC‐EVs induced HSFs' MET mainly through miR‐200s/ZEBs axis. This innovative mechanistic insight suggests a promising avenue for future therapeutic strategies targeting HS and other skin fibrosis diseases.

2. Materials and Methods

2.1. Human Skin Samples Collection

This study was approved by the Medical and Ethics Committees of the First Affiliated Hospital of Sun Yat‐sen University (2022026), and all donors signed an informed consent before enrolling in this study. Human skin tissue samples were collected from surgically discarded patient specimens, with prior ethical approval and informed consent obtained. This study included three normal skin samples, three HS samples 6–7 months post‐burn, and three mature scar samples 2–5 years post‐trauma or post‐surgery. Sample details are presented in Table 1. Full terms for abbreviations and acronyms are presented in Table 4.

TABLE 1.

The demographics and characteristics of sample donors.

Sample Sex Age Cause of Scars Scars’ age Location
Normal skin 1 M 10 — 0 Chest
Normal skin 2 M 18 — 0 Leg
Normal skin 3 F 36 — 0 Chest
Hypertrophic scar 1 M 8 Burn 6 months Leg
Hypertrophic scar 2 M 12 Burn 6 months Leg
Hypertrophic scar 3 F 18 Burn 7 months Leg
Mature Scar 1 M 25 Trauma 5 years Leg
Mature Scar 2 M 55 Surgery 2 years Leg
Mature Scar 3 F 35 Surgery 2 years Leg
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TABLE 4.

Full terms for abbreviations and acronyms.

Abbreviations and acronyms Full name
EDGS EpiLife Defined Growth Supplement
EDTA Ethylenediaminetetraacetic acid
SDS‐PAGE Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis
PVDF Polyvinylidene Difluoride
BSA Bovine Serum Albumin
RIPA Radioimmunoprecipitation Assay
mIHC Multiplex Immunohistochemistry
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2.2. Cell Isolation and Culture

ESCs and fibroblasts (FBs) were isolated from human foreskins (n = 10; aged 6–20 years). RESCs and RFBs were isolated from the foetal rat skin (n = 5). Hypertrophic scar fibroblasts (HSFs) were isolated from human HSs as described (Wang et al., 2022; Wang et al. 2024). Briefly, subcutaneous tissues and fat were removed before the skin was disinfected and incubated overnight with DispaseII (3 mg/mL; D4693‐1G, Sigma–Aldrich) in keratinocyte serum‐free medium (Epilife, MEPI500CA, Gibco) with EDGS (S0125, Gibco) at 4°C to separate the epidermis from the dermis. After that, the epidermis separated from the foreskins was digested with prewarmed Trypsin/EDTA (0.25%, Gibco) at 37°C for 10 min to obtain ESCs, which were resuspended in Epilife and seeded at a density of 105 cells/cm2 in 100 µg/mL type IV collagen‐coated culture dishes. Cells adhering within 10 min were selected according to the rapid attachment method (Kim et al. 2004). Similarly, the dermis was digested with Collagenase IV (3 mg/mL, C6745‐1ML, Corning) and Hyaluronidase (2 mg/mL, 1141GR001, Biofroxx) in Dulbecco's modified Eagle medium (DMEM, C11995500BT, Gibco) at 37°C for 2 h to obtain FBs and HSFs, which were cultured in DMEM supplemented 10% foetal bovine serum with 5% CO2 at 37°C. All cells were cultured in cell factories to obtain sufficient amounts of EVs. Cells at the 3–5 passages were used for this study.

2.3. EVs Isolation, Characterization and Treatment

EVs were isolated from the culture medium of cells using a differential ultracentrifugation method (Lin et al. 2020). For the collection of conditioned medium, 80% confluent ESCs or FBs were washed with PBS before being cultured with serum‐free media for 2 days. Briefly, the medium was centrifuged at 300 × g for 10 min and 2000 × g for 10 min at 4°C, followed by ultracentrifugation at 100,000 × g for 70 min at 4°C on an Optima XPN‐100 ultracentrifugation with an SW32Ti rotor (Beckman Coulter, USA). The isolated EVs were washed with PBS and subjected to secondary ultracentrifugation at 100,000 × g for 70 min at 4°C. EVs were stored in PBS at −80°C. The particle size distribution and concentration were measured with ZetaView (Particle Metrix, Germany), and the morphology was examined by electron microscopy (HITACHI HT‐7700, Japan). The EV protein was quantified using a BCA protein assay kit (Thermo Fisher Scientific, USA). Western blotting was performed to detect positive markers Alix (1:1000, RGAB100‐50, Rengenbio), CD9 (1:1000, ab92726, Abcam), CD81 (1:1000, RGAB105‐50, Rengenbio), CD63 (1:1000, ab134045, Abcam), TSG101 (1:1000, ab125011, Abcam) and negative markers GAPDH (1:50000, 60004‐1‐lg, Proteintech) of EVs. For TSG101, two bands were detected, appearing around ∼55 kDa, which is consistent with the manufacturer's datasheet and previous studies (Dibsy et al. 2023; Jiang et al. 2023).

2.4. Cellular Uptake Assay of EVs

To evaluate EVs delivery, PKH67 membrane dye (MINI67, Sigma–Aldrich) was added to PBS at a concentration of 1 µM and incubated with EVs (10 µg) for 20 min. The incubation was stopped with 0.5%BSA (A3311, Sigma–Aldrich) before washing repeatedly. The labelled EVs were then resuspended (10 µg/mL) and incubated with HSFs for 24 h. After that, the cells were fixed with 4% paraformaldehyde for Actin (2 drops/mL, R37112, Thermo Fisher) and DAPI staining and observed using a confocal laser scanning microscope (OLYMPUS FV3000, Japan).

2.5. Flow Cytometric Analysis (FCA)

To identify ESCs, about 106 cells were collected, prepared as single‐cell suspensions and incubated with 2.5 µg Fc block in 100 µL PBS. Surface marker anti‐CD49f‐FITC (1:100, 313605, Biolegend, USA) was used to stain cells at room temperature for 30 min. While for intracellular staining, cells were permeabilized and fixed with the intracellular staining kit (554714, BD Biosciences) and then washed and stained with antibodies anti‐cytokeratin 15 (1:100, ab52816, Abcam) at room temperature for 30 min. After washing, cells were incubated with second antibodies (1:500, 4408S, CST) and washed twice before resuspending. The stained cells were analysed using a CytoFlex flow cytometer (Beckman, USA). Data were processed by FlowJo Software v10.8.1.

2.6. Cell Viability Assays

Cells were plated onto 24‐wells at a density of 105 cells/cm2 in a 37°C and 5% CO2 incubator. After 24 h, living and dead cells were detected using the Calcein/PI Cell Viability/Cytotoxicity Assay Kit (C2015S, Beyotime, China) in accordance with the manufacturer's instructions. Then the cells were observed under an inverted fluorescence microscope (OLYMPUS IX83, Japan).

2.7. Collagen Gel Contraction Assay

According to previous articles (Ngo et al. 2006; Huang et al. 2022), HSFs were resuspended in 1.8 mg/mL collagen solution at a concentration of 2 × 105/mL. The mixture of cellular collagen was seeded in 24‐well plates (0.5 mL/well) and incubated at 37°C for 1 h to coagulate. Then, 500 µL of the conditioned medium was gently added to each well, and the gel was separated from the plates by gently running the sterile pipette tip along the gel edges.

2.8. Multiplex CRISPR/Cas9 System and Lentivirus Transfection

As previously described, we assembled customized lentiviral vectors expressing sgRNAs targeting hsa‐miR‐200b‐5p, hsa‐miR‐429, hsa‐miR‐200c‐5p and hsa‐miR‐141‐3p into a lentiviral vector (CV279, Genechem, China) that ultimately expressed active Cas9 by Golden Gate cloning (Kabadi et al. 2014; Yu et al. 2022). The ESCs were infected with lentivirus expressing multiplex CRISPR system. Stable clones were selected after 10 days using 1 µg/mL puromycin.

2.9. Western Blot Analysis

Protein samples were lysate using RIPA lysis buffer supplemented with protease inhibitors (Calbiochem, USA). Equal amounts of proteins were loaded and separated by 10% SDS‐PAGE and then transferred to polyvinylidene difluoride membranes (PVDF, Millipore, USA). The membrane was blocked with 5% BSA and then incubated with one of the primary antibodies: anti‐alpha smooth muscle (1:1000, GB111364, Servicebio), anti‐COL1A1 (1:1000, E3E1X, Cell Signalling Technology), anti‐N Cadherin (1:1000, A19083, Abclonal), anti‐cytokeratin 1 (1:500, PA5‐113746, Invitrogen), anti‐cytokeratin 15 (1:10000, ab52816, Abcam), anti‐E Cadherin (1:500, ab1416, Abcam), anti‐ZEB1 (1:500, ab181451, Abcam) and anti‐ZEB2 (1:1000, AF5278, Affinity Biosciences) overnight at 4°C. Afterward, the PVDF membranes were incubated with HRP‐conjugated secondary antibodies at room temperature for 1 h. The immunoreactive bands were visualized with the enhanced chemiluminescence kit (Sigma–Aldrich) and analysed by densitometry using ImageJ software.

2.10. Fluorescence In Situ Hybridization (FISH)

The paraffin sections were prepared and fixed as previously described (Kaufer 2013). SweAMI probes against miR‐200s were used (Servicebio, China). Staining was performed as written in the manufacturer's protocol. Briefly, slides were treated with proteinase K (20 µg/mL, G1234, Servicebio) for 20 min and pre‐hybridization for 1 h at 37°C before hybridizing with probes overnight at 37°C. Then, the slides were counterstained with DAPI and scanned afterward. FISH probes are presented in Table 3.

TABLE 3.

FISH probes.

Gene Probe
hsa‐miR‐200a‐5p TCCAGCACTGTCCGGTAAGATG
hsa‐miR‐200b‐3p TCATCATTACCAGGCAGTATTA
hsa‐miR‐200c‐3p TCCATCATTACCCGGCAGTATTA
hsa‐miR‐141‐5p CCAACACTGTACTGGAAGATG
hsa‐miR‐429 ACGGTTTTACCAGACAGTATTA
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2.11. RNA Extraction and Real‐Time PCR Assay

Total RNA from cells was extracted using Trizol reagent (Thermo Fisher) and reverse‐transcribed into cDNA using the PrimeScript RT reagent kit (Takara) according to the manufacturer's protocol. Quantitative real‐time PCR (qRT‐PCR) reaction was performed with the Roche 480 system (Roche) using the LightCycler 480 SYBR Green I Master Mix (Roche). In contrast, miRNA was extracted using Trizol reagent (Thermo Fisher), reverse‐transcribed and performed qRT‐PCR using All‐in‐One miRNA qRT‐PCR Detection Kit (QP115, GeneCopoeia). The relative mRNA expression levels of target genes were calculated using the 2−ΔΔCt method. Primers used in qRT‐PCR are presented in Table 2.

TABLE 2.

qRT‐PCR primers.

Gene Forward primer Reverse primer
hsa‐COL1A1 CCCCTGGAAAGAATGGAGATG AGCTGTTCCGGGCAATCCT
hsa‐α‐SMA GCGGATAGAGATGTCTGGAAGC CGGCTTCATCGTATTCCTGTT
hsa‐N‐Cadherin AAGAGGCAGAGACTTGCGAAAC TGGAGTCACACTGGCAAACCTT
hsa‐Krt1 ATGGACAACAACCGCAGTCT TGCAGCTCTTCATACTTGCTCT
hsa‐Krt15 ACCACGAAGAGGAGATGAAGGA TTGCTGGTCTGGATCATTTCTG
hsa‐E‐Cadherin GCTGGACCGAGAGAGTTTCC CGACGTTAGCCTCGTTCTCA
hsa‐ZEB1 CGCAGTCTGGGTGTAATCGTAA ATGTCTTGAGTCCTGTTCTTGGTC
hsa‐ZEB2 AGGAAGATGAAATAAGGGAGGGT TCACTGTACCATTGTTAATTGCGG
GAPDH GGAAGCTTGTCATCAATGGAAATC TGATGACCCTTTTGGCTCCC
hsa‐miR‐200a‐5p CATCTTACCGGACAGT TCCAGCAC
hsa‐miR‐200b‐3p TAATACTGCCTGGTAA TCATCATT
hsa‐miR‐200c‐3p TAATACTGCCGGGTAAT TCCATCAT
hsa‐miR‐141‐5p CATCTTCCAGTACAGT TCCAACAC
hsa‐miR‐429 TAATACTGTCTGGTAA ACGGTTTT
U6 CTCGCTTCGGCAGCACA AACGCTTCACGAATTTGCGT
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2.12. In Vitro Immunofluorescent Staining

A total of 104 HSFs were resuspended in 500 µL DMEM with 10 µg/mL ESC‐EVs, FB‐EVs, or PBS and added into one well of an eight‐well removable glass chamber slide (Nunc Lab‐TekIIChamber Slide System, Thermo Fisher Scientific). After 24 h, cells were fixed with 4% formaldehyde, permeated with 0.1% Triton X‐100 (Solarbio, China) for 15 min, and blocked with 2% BSA for 1 h. Next, the cells were stained with primary antibodies, including anti‐alpha smooth muscle (1:500, GB111364, Servicebio), anti‐COL1A1 (1:100, E3E1X, Cell Signalling Technology), anti‐N Cadherin (1:200, ab98952, Abcam), anti‐cytokeratin 1 (1:100, PA5‐113746, Invitrogen), anti‐cytokeratin 15 (1:50, ab52816, Abcam), anti‐E Cadherin (1:50, ab1416, Abcam), anti‐ZEB1 (1:200, ab181451, Abcam) and anti‐ZEB2 (1:100, AF5278, Affinity Biosciences) overnight at 4°C. Cells were washed and incubated with secondary fluorescence antibodies (1:2000, 4408S, 4409S, 4412S, 4413S, Cell signalling Technology) for 1 h and DAPI for 10 min at room temperature. Then the cells were observed under an inverted fluorescence microscope (OLYMPUS IX83, Japan). Quantitative analysis of fluorescence intensity was conducted on a single field of view from independent experiments.

2.13. miRNA Sequencing Analysis of EVs

After total RNA was extracted by TRIzol, the RNA molecules in a size range of 18–30 nt were enriched by polyacrylamide gel electrophoresis (PAGE). The transcripts per million reads were used for the calculation of gene expression, and genes with |log2 (fold change)| ≥ 1 and p < 0.05 were considered statistically significant. Target genes of each miRNA in EVs were predicted using publicly available algorithms multiMiR (Ru et al. 2014). Kyoto Encyclopaedia of Genes and Genomes analyses and Gene Ontology analyses were performed to elucidate the functions and enriched pathways of statistically significant genes.

2.14. In Vivo Model of HSs

Male Sprague–Dawley rats aged 8 weeks were purchased from the Ruige Biological Technology Corporation (Guangzhou, China). All experimental procedures were approved by the Ethics Committee Board of the First Affiliated Hospital of Sun Yat‐Sen University and performed following the NIH Guide for the Care and Use of Laboratory Animals. Briefly, a 6 × 6 mm full‐thickness skin defect was created on the dorsal side of the rat tail. Then the wound site was stretched by attaching a 2 cm stainless steel ring to the ventral side of the tail. After 3 weeks, RHS had been successfully epidermalised. Thereafter, 12 rats were randomly assigned into three groups (n = 4 per group): the RESC‐EVs group (200 µL, 10 µg per week), the RFB‐EVs group (200 µL, 10 µg per week) and the control group (PBS, 200 µL per week). We administered weekly four‐point subcutaneous injections to the scar. On Day 28, all rats were sacrificed and the scar tissues were surgically removed for experiments (Zhou et al. 2019).

2.15. Histological Examination

Skin tissue was fixed overnight in 4% paraformaldehyde at room temperature and embedded in paraffin. Afterward, 5‐µm‐thick slices were cut out. Some of them were stained with haematoxylin‐eosin (H&E) and Masson and analysed using a microscope (Kfbio, China). Others were subjected to multiplex immunohistochemistry (mIHC) to enable simultaneous detection of multiple targets. They were blocked with goat serum and then stained with primary antibodies, including anti‐alpha smooth muscle, anti‐COL1A2 (1:200, A5786, Abclonal), anti‐N Cadherin, anti‐cytokeratin 14 (1:200, A15069, Abclonal), anti‐E Cadherin, anti‐ZEB1 (1:200, ab181451, Abcam) and anti‐ZEB2 overnight at 4°C. Sections were washed and incubated with HRP‐conjugated secondary antibodies (1:2000, ab6728, ab6721, Abcam) for 1 h, fluorochrome (15X, ASOP520, ASOP570, Asbio) for 10 min and DAPI for 10 min at room temperature. Slides were scanned afterward (Pannoramic MIDI, 3DHISTECH, Hungary) and analysed using CaseViewer 2.4. Quantitative analysis of fluorescence intensity was conducted on a single field of view from independent experiments.

2.16. Scar Assessment

Scar severity was evaluated using the Vancouver Scar Scale (VSS), which assesses four parameters: vascularity, pigmentation, pliability and height. Each parameter was scored individually, and the total score reflected the overall severity of the scar. For histological evaluation, the Scar Elevation Index (SEI) was calculated based on H&E‐stained tissue sections. SEI was defined as the ratio of the total dermal area—including both the hypertrophied dermis and the underlying dermis—to the area of the underlying dermis alone. The height of the underlying dermis was determined from adjacent unwounded dermis within the same section. All measurements were confined to the wounded area, and the epithelial height was excluded from the calculation.

2.17. Statistical Analysis

All data are presented as the mean ± SD. The dots in the quantitative analysis represent independent experiments from different donor populations. Differences among experimental groups were conducted by Student's t‐test (for single comparisons) or one‐way ANOVA followed by Fisher's post hoc test (for multiple comparisons). The relationship between epithelial markers (K14 and E‐cadherin) and ZEB1/2 expression was analysed using Pearson correlation. Statistical analyses were performed with GraphPad Prism 8.3.0 software (GraphPad Software, USA); p < 0.05 was considered statistically significant.

3. Results

3.1. Isolation and Characterization of the ESC‐EVs and FB‐EVs

As depicted in Figure 1a, the primary ESCs and FB were isolated from human foreskin or foetal rat skin tissues, displaying high proliferative capacity and forming colonies (Figure 1b). Flow cytometry (FCM) analysis revealed the ESCs was positive for integrin CD49f (97.3%) and Krt15 (95.1%), while the FB was positive for Vimentin (98.8%) and negative for Pan‐Keratin (99.8%) (Figure 1c).

FIGURE 1.

FIGURE 1

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Isolation and characterization of the ESC‐EVs and FB‐EVs. (a) Schematic diagram of cell separation, culture and the EVs isolation. (b) Calcein and PI stains showed the high proliferative capacity of ESCs, FBs and HSFs. Scale bar, 100 µm. (c) Flow cytometry analysis of ESCs markers (cd49f and Krt15) and FBs markers (Pan‐Keratin and Vimentin). (d) Transmission electron microscopy (TEM) images of ESC‐EVs and FB‐EVs. Scale bar, 500 nm (down) and 200 nm (up). (e) Nanoparticle tracking analysis of ESC‐EVs and FB‐EVs from ZetaView. (f) Endosomal (Alix, TSG101) and tetraspanin (CD81, CD63) expression in FBs, ESCs, FB‐EVs and ESC‐EVs. GAPDH was used as a negative EVs marker.

Purified EVs were characterized according to morphology, size distribution, surface markers and surface charge (Figure 1d–f). ESC‐EVs and FB‐EVs displayed a round shape with a bilayer structure (Figure 1d). Besides, NTA analysis identified that the mean diameters of the ESC‐EVs were 127.9 nm, and the FB‐EVs were 138.7 nm (Figure 1e). The protein content in the solution of the ESC‐EVs was 0.48 µg protein/mL medium, and the FB‐EVs was 0.56 µg protein/mL medium. The ESC‐EVs concentration was 28.4 × 106 particles/mL medium, which was equivalent to 59.2 × 106 particles/mg protein, while the FB‐EVs was 33.6 × 106 mL/medium and identical to 60.0 × 106 particles/mg protein. In addition, immunoblotting was performed to confirm the presence of typical EV markers Alix, CD9, CD81, CD63 and TSG101 (Figure 1f). These data indicated that the ESC‐EVs and FB‐EVs we collected were highly purified and suitable for the subsequent experiments.

3.2. ESC‐EVs Were Internalized and Effectively Inducing the MET of HSFs

To determine whether HSFs are recipient cells for ESC‐EVs, the ESC‐EVs were labelled with PKH67 (green) and the HSF with Actin (red), respectively. After 12 h of incubation, confocal imaging revealed green fluorescent spots in recipient HSFs, indicating the labelled EVs had been successfully delivered to HSFs (Figures 2a and S1).

FIGURE 2.

FIGURE 2

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ESC‐EVs were internalized and effectively inducing the MET of HSFs. (a) Confocal imaging showed the internalized PKH67‐labeled ESC‐EVs (green) in ACTIN‐labelled HSFs (red). Scale bar, 25 µm. (b) Western blotting analysis of COLI, α‐SMA, N‐cad, K1, K15 and E‐cad in HSFs incubated with PBS, ESC‐EVs at concentrations of 1.25, 2.5, 5 or 10 µg/mL or FB‐EVs (10 µg/mL). (c) Quantification of the relative band density compared to GAPDH. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (d) RT‐qPCR analysis of the mesenchymal markers COLI, α‐SMA and N‐cad and epithelial markers K1, K15 and E‐cad in HSFs treated with ESC‐EVs, FB‐EVs or PBS. Graph represented the expression of the markers relative to that of GAPDH. Data are presented as mean values ± SD (n = 9). Statistical analysis was performed using one‐way ANOVA. (e) Representative images of COLI, α‐SMA, N‐cadherin (green) immunofluorescence staining in ACTIN‐labelled HSFs (red) incubated with ESC‐EVs or PBS for 24 h. Scale bar, 50 µm. (f) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. (g) Representative images of K1, K15 and E‐cadherin (green) immunofluorescence staining in ACTIN‐labelled HSFs (red) incubated with ESC‐EVs or PBS for 24 h. Scale bar, 50 µm. (h) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p < 0.0005 and ****p < 0.0001.

Then we co‐cultured HSFs with different concentrations of ESC‐EVs. HSFs stimulated with ESC‐EVs exhibited a morphological trend resembling tufted cobblestone‐like keratinocyte patterns (Figure S2a). To ensure rigorous characterization, we primarily relied on the expression of specific molecular markers rather than morphological observations. As shown in Figure 2b,c, ESC‐EVs significantly decreased the expression of mesenchymal markers in HSFs including collagen I (COLI), α‐smooth muscle Actin (α‐SMA) and N‐cadherin (N‐cad), while increasing the expressions of epithelial markers like Krt1 (K1), Krt15 (K15) and E‐cadherin (E‐cad), and the optimal effective concentration was 5 µg/mL. RT‐qPCR analysis (Figure 2d) and immunofluorescence staining showed the same results (Figures 2e–i, S3 and S9). These findings indicated that ESC‐EVs effectively induced the MET of HSFs and inhibited its activation to attenuate fibrosis (Ngo et al. 2006).

3.3. miR‐200s Were Enriched in ESC‐EVs and Targeting ZEBs in HSFs

To further elucidate the potential mechanisms of ESC‐EVs in HSFs’ MET, we detected the miRNA expression profiles between the ESC‐EVs and FB‐EVs using microarrays (Figure 3a,b). A total of 209 miRNAs were discovered to be significantly different, of which 173 were highly expressed in ESC‐EVs and 36 were lowly expressed. Among them, we found that the miR‐200 family (miR‐200s), including miR‐200a, miR‐200b, miR‐200c, miR‐141 and miR‐429 were all highly expressed in ESC‐EVs, and the differences were significant (LogFC > 1, p < 0.05 and FDR < 0.05). Then we used the clusterProfiler package for Gene ontology (GO) and Kyoto Encyclopaedia of Genes and Genomes (KEGG) pathway (Wu et al. 2021) annotations, and the ggplot2 package for illustration. As shown in Figure 3c,d, the ZEB1 was enriched in GO analysis, and the ZEB1 and ZEB2 were discovered by KEGG analysis(Huang et al. 2019; Bakir et al. 2020).

FIGURE 3.

FIGURE 3

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miR‐200s were enriched in ESC‐EVs and targeting ZEBs in HSFs. (a) Heatmap of the 20 miRNAs with the largest LogFC in the microarray analysis of the miRNA expression profiles between ESC‐EVs and FB‐EVs. (b) Volcano annotated with miR‐200s probes (ESC‐EVs vs. FB‐EVs). Type, High expression (red), Low expression (blue) and No significance (grey). (c) Chord diagram of GO enrichment terms. (d) Chord diagram of KEGG enrichment terms. (e) RT‐qPCR analysis of the expression of miR‐200s in HSFs treated with ESC‐EVs, FB‐EVs or PBS for 24 h. Graph represented the expression of the markers relative to that of U6. Data are presented as mean values ± SD (n = 9). Statistical analysis was performed using one‐way ANOVA. (f) RT‐qPCR analysis of the expression of ZEB1 and ZEB2 in HSFs relative to GAPDH. Data are presented as mean values ± SD (n = 9). Statistical analysis was performed using one‐way ANOVA. (g) Western blotting analysis of ZEB1 and ZEB2 in HSFs incubated with PBS, ESC‐EVs at concentrations of 1.25, 2.5, 5 or 10 µg/mL or FB‐EVs (10 µg/mL). (h) Quantification of the relative band density compared to GAPDH. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (i) Representative images of ZEB1 and ZEB2 (green) immunofluorescence staining in ACTIN‐labelled HSFs (red). Scale bar, 50 µm. (j) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 5). Statistical analysis was performed using one‐way ANOVA. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p<0.0005 and ****p<0.0001.

To investigate whether the miR‐200s were successfully transferred to HSFs via ESC‐EVs and inhibited ZEBs, we detected the expression of miR‐200s in HSFs incubated with PBS, ESC‐EVs and FB‐EVs, respectively. After 24 h of incubation, the expression of miR‐200s was remarkably increased in the ESC‐EVs group (Figure 3e), while the expression of ZEB1 and ZEB2 was significantly reduced (Figure 3f–j). Taken together, we speculated that ESC‐EVs target the inhibition of ZEBs in HSFs by delivering miR‐200s.

3.4. HS Exhibited miR‐200s Deficiency and ZEBs Accumulation

To explicit the expression of miR‐200s and ZEBs in human normal skin (NS), HS and mature scar (MS) tissues, we subsequently examined their expression by fluorescence in situ hybridization (FISH). Compared with the NS, the expression of miR‐200s in HS was significantly decreased, while it was restored in MS (Figure 4a,b). At the same time, mIHC also showed the expression of ZEB1 and ZEB2 in the HS was much higher than in NS and MS (Figures 4c,d and S4). These results demonstrated a vital role of the miR‐200s/ZEBs axis in the occurrence and development of HS.

FIGURE 4.

FIGURE 4

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HS exhibited miR‐200s deficiency and ZEBs accumulation. (a) FISH stains of miR‐200s in normal skin (NS), hyperplasia stage (HS) and mature stage of hypertrophic scar (MS). Scale bar, 20 µm. (b) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (c) mIHC stains of α‐SMA, ZEB1, ZEB2, COLI, α‐SMA, N‐Cad and K14, E‐Cad in normal skin, hyperplasia stage or mature stage of hypertrophic scar. Scale bar, 200 µm, 20 µm. (d) Quantification of the relative positive cells. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p < 0.0005 and ****p < 0.0001.

3.5. ESC‐EVs Induced MET of HSFs via miR‐200s/ZEBs Axis

To further investigate whether the effects of ESC‐EVs are dependent on the miR‐200s, we targeted all five members of the miR‐200s located across two different genomic loci using a multiplexed CRISPR/Cas9 system in a single lentiviral vector based on previous reports (Yu et al. 2022). Two DNA fragments encoding guidelines for miR‐200b, miR‐200a, miR‐429 genes on chromosome 1, and miR‐200c, miR‐141 on chromosome 12, were designed to be deleted in the presence of the four sgRNAs and active Cas9 nuclease (Figure 5a,b). Thus, all miR‐200 family members were simultaneously knocked down in ESC‐EVs (Figure 5c).

FIGURE 5.

FIGURE 5

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ESC‐EVs induced MET of HSFs via miR‐200s/ZEBs axis. (a) The multiplex CRISPR/Cas9 in a single lentiviral vector. (b) Five sgRNAs are indicated by scissors targeting two genomic loci of human miR‐200 family host genes. (c) RT‐qPCR analysis of EVs secreted from wild‐type ESC‐EVs (WT), ESCs infected with scramble vectors (Scramble), or miR‐200s knockdown vectors (KD). Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (d) RT‐qPCR analysis of COLI, α‐SMA, N‐cad, K1, K15 and E‐cad in HSFs. Graph represented the expression of the markers relative to that of GAPDH. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (e) Western blotting analysis of COLI, α‐SMA, N‐cad, K1, K15 and E‐cad in HSFs. (f) Quantification of the relative band density compared to α‐tubulin. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (g) Representative images of COLI, α‐SMA, N‐cad, K1, K15 and E‐cad (green) immunofluorescence staining in ACTIN‐labeled HSFs (red) incubated with WT‐EVs, Scramble‐EVs, KD‐EVs, or PBS for 24 h. Scale bar, 50 µm. (h) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. (i) Representative images of ZEB1 and ZEB2 (green) immunofluorescence staining in ACTIN‐labelled HSFs (red). Scale bar, 50 µm. (j) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. (k) RT‐qPCR analysis of the expression of ZEB1 and ZEB2 in HSFs relative to GAPDH. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. (l) Western blotting analysis of ZEB1 and ZEB2 in HSFs. (m) Quantification of the relative band density compared to α‐tubulin. Data are presented as mean values ± SD (n = 3). Statistical analysis was performed using one‐way ANOVA. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p < 0.0005 and ****p < 0.0001.

After being incubated for 24 h, HSFs incubated with ESC‐EVs infected with miR‐200s knockdown vectors (KD) did not show apparent morphological change, aligning with those cultured with the normal medium (N) (Figure S2b). Furthermore, the results from RT‐qPCR (Figure 5d), WB (Figure 5e,f) and IF (Figures 5g,h and S5) consistently showed that miR‐200s knockdown significantly increased the expression of mesenchymal markers and decreased the expression of epithelial markers. Meanwhile, the knockdown of miR‐200s restored the expression of both ZEB1 and ZEB2 suppressed by wild‐type ESC‐EVs in HSFs (Figure 5i–m). These findings revealed that ESC‐EVs induced MET of HSFs via the miR‐200s/ZEBs axis.

3.6. ESC‐EVs‐miR‐200s Suppressed the Migration and Contractile Ability of HSFs

To express the effect of ESC‐EVs on HSFs' function, we detected HSFs' proliferation, migration and contractile ability under different treatments. Although CCK8 and Ki67 staining experiments consistently showed that the growth of HSF was not significantly affected by ESC‐EVs (Figures 6a–c and S6), the scratch test showed that the migration of HSFs was considerably inhibited by ESC‐EVs treatments compared to the PBS and FB‐EVs groups (Figure 6d,e). In addition, the collagen gel contraction test also displayed that ESC‐EVs significantly inhibited the contractile ability of HSFs while FB‐EVs did not (Figure 6f,g).

FIGURE 6.

FIGURE 6

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ESC‐EVs inhibited the activation of HSFs via miR‐200s. (a) CCK8 assay to detect the impact of both ESC‐EVs, FB‐EVs, or PBS on HSFs cell viability. Data are presented as mean values ± SD (n = 6). Two‐sided unpaired t‐test. (b) Representative images of Ki67 (green) immunofluorescence staining in ACTIN‐labelled HSFs (red). Scale bar, 100 µm. (c) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. (d) Scratch tests showed the comparison of relative migration areas among ESC‐EVs, FB‐EVs or PBS at 0 and 24 h. Scale bar, 100 µm. (e) Quantification of the migration rate at 6, 12, 24 and 48 h. Data are presented as mean values ± SD (n =6). Statistical analysis was performed using Two‐sided unpaired t‐test. (f) Collagen gel contraction test illustrated the contraction of ESC‐EVs, FB‐EVs or PBS populated collagen gels at 0, 24 and 48 h. (g) Quantification of contraction surface areas at 24, 48 and 72 h. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using Two‐sided unpaired t‐test. (h) Scratch tests showed the comparison of relative migration areas among WT‐EVs, Scramble‐EVs, KD‐EVs or PBS at 0 and 24 h. Scale bar, 100 µm. (i) Quantification of the migration rate at 12 and 24 h. Data are presented as mean values ± SD (n = 6). Statistical analysis was performed using two‐sided unpaired t‐test. (j) Collagen gel contraction test illustrated the contraction of WT‐EVs, Scramble‐EVs, KD‐EVs or PBS populated collagen gels at 0 and 72 h. (k) Quantification of contraction surface areas at 24, 48 and 72 h. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using Two‐sided unpaired t‐test. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p < 0.0005 and ****p < 0.0001.

However, after the knockdown of the miR‐200s in ESC‐EVs, both the scratch test and collagen gel contraction test showed that the migration and contractile abilities of HSFs were significantly enhanced compared to those directly treated with ESC‐EVs, indicating that the loss of miR‐200s in ESC‐EVs resulted in the loss of the ability to inhibit HSF activation (Fig 6h‐j). These findings revealed that wild‐type ESC‐EVs induce MET via the miR‐200s/ZEBs axis in HSFs.

3.7. Application of ESC‐EVs Alleviated the HS of Rat Tail (RHS)

To confirm the anti‐fibrotic efficacy of ESC‐EVs in vivo, we established the HS of rat tail (RHS) model according to previously reported (Wang et al. 2024). As shown in Figure 7a, a 6 × 6 mm full‐thickness wound was created on the rat tail. Three weeks after the operation, the rat tail defect was replaced by HS.

FIGURE 7.

FIGURE 7

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Application of ESC‐EVs alleviated the HS of rat tail (RHS). (a) Schematic representation of animal model preparation and administration in RHS. (b) Morphology of RHS on post‐injury Days 0, 7, 14, 21 and 28. (c) Quantitative evaluation of the scars volume. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using Two‐sided unpaired t‐test. (d) Representative images of H&E and Masson's trichrome stating of RHS sections. Scale bar, 200 µm, 100 µm. (e) Quantitative evaluation of the scar score. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using Two‐sided unpaired t‐test. (f) Quantitative evaluation of the scar elevation index. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. (g) Quantitative evaluation of the collagen volume fraction. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p < 0.0005 and ****p < 0.0001.

ESC‐EVs, FB‐EVs or an equal volume of PBS was subcutaneously injected once weekly according to the different groups. No death or abnormality was observed in any animal during the postoperative period. Digital photographs displayed the RHS changes on Days 7, 14, 21 and 28 post‐wounding (Figure 7b). After two applications of ESC‐EVs, the RHS demonstrated a notable decrease in thickness, heightened softness, volume and a more superficial appearance, thereby contributing to enhanced clinical aesthetics (Figure 7c). Subsequently, H&E and Masson's trichrome staining revealed a significant reduction in scar elevation index (SEI) and collagen density in the ESC‐EVs treated group compared to both control and FB‐EVs treated groups, resulting in more organized collagen fibres compared to the FB‐EVs and control groups (Figure 7d–g).

3.8. ESC‐EVs Alleviated the RHS Through the miR‐200s/ZEBs Axis

To further investigate the mechanism of ESC‐EVs alleviated the RHS, we detected the expression of miR‐200s (Figures 8a,b and S7a,b), ZEBs, E‐Cad, N‐Cad, α‐SMA, Collagen I and KRT14 (Figures 8c,d and S7c). Consistently, the expression levels of α‐SMA, N‐Cadherin and Collagen I were significantly reduced, while the miR‐200s, Krt14 and E‐Cadherin were notably increased in RHS treated with ESC‐EVs. In addition, levels of mesenchymal markers exhibited a significant positive correlation, whereas epithelial markers showed a negative correlation with ZEB1/2 (Figure S8), consistent with our in vitro observations. Taken together, these findings revealed the efficacy of ESC‐EVs in attenuating HS by promoting MET of HSFs via the miR‐200s/ZEBs Figures 9.

FIGURE 8.

FIGURE 8

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ESC‐EVs alleviated the RHS through the miR‐200s/ZEBs axis. (a) FISH stains of miR‐200a (red), miR‐200b (green), miR‐200c (pink), miR‐141 (red), miR‐429 (green) in PBS, RESC‐EVs, RFB‐EVs treated RHS. Scale bar, 20 µm. (b) Quantification of the relative fluorescence ratio. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. (c) Representative images of COLI, α‐SMA, N‐cad, K1, K15, E‐cad, ZEB1 and ZEB2 immunofluorescence staining. Scale bar, 200 µm, 20 µm. (d) Quantification of the relative positive cells. Data are presented as mean values ± SD (n = 4). Statistical analysis was performed using one‐way ANOVA. Significance levels are indicated as *p < 0.05, **p < 0.005, ***p < 0.0005 and ****p < 0.0001.

FIGURE 9.

FIGURE 9

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Schematic illustration of ESC‐EVs induced HSFs’ MET to alleviate HS via the miR‐200s/ZEBs axis. Epidermal stem cells (ESC) were cultured in cell factories to obtain sufficient extracellular vesicles (ESC‐EVs). EVs were isolated using a differential ultracentrifugation method. Both in vitro cell experiments and in vivo rat tail HS (RHS) models thoroughly validated that ESC‐EVs were internalized and effectively inducing the MET of HSFs and inhibited its activation to attenuate fibrosis via the miR‐200s/ZEBs axis.

4. Discussion

The overproduction of hypertrophic scar fibroblasts (HSFs) caused by excessive EMT is one of the most critical factors leading to HS (Yuan et al. 2019). Effectively inhibiting EMT and even inducing MET is significant for treating HS. In the present study, we first found that ESC‐EVs could effectively induce the MET of HSFs and inhibit their biological activity. Then, we further elucidated that this therapeutic effect is mediated by the miR‐200s encapsulated in ESC‐EVs, which targeted and regulated ZEB1 and ZEB2 in HSFs. This vital role and mechanism have been fully validated in both in vitro cell experiments and in vivo RHS models. These findings highlight the anti‐fibrosis potential of ESC‐EVs and their role in treating skin fibrosis disease.

As a formidable fibrotic skin condition, HS poses an enduring challenge in clinical practice worldwide (Finnerty et al. 2016; Ogawa et al. 2021). Although conventional treatments—including pressure therapy, silicone gel, corticosteroids and 5‐FU—are available, their efficacy remains limited and is often accompanied by significant adverse effects (Ogawa et al. 2021). To develop novel, practical therapeutic approaches explicitly focusing on its pathogenesis is imperative.

ESCs represent a pluripotent stem cell population capable of proliferating into specialized keratinocytes and skin appendages (Blanpain and Fuchs 2006). Residing in the basal layer of the epidermis, ESCs interface intimately with the underlying basement membrane, thereby encountering a milieu of dermal cells (Fujiwara et al. 2018). Its biological behaviour plays a vital role in the differentiation of FBs during wound healing and scar formation (Wang et al. 2017). Our previous studies have validated its capacity to enhance impaired wound healing and relieve HS (Huang et al. 2020). However, as cellular therapy remains beset by challenges, including cellular immunogenicity, storage stability, ethical considerations and risks associated with embolism and tumorigenesis, the clinical research and application of ESCs have been greatly limited (Herberts et al. 2011).

As central cell‐cell communication mediators, EVs share similar functions with the maternal cells (Théry et al. 2018). Compared with stem cell transplantation, EVs therapy has the advantages of low immunogenicity, high safety, easy storage and management and mass production, which is an ideal choice for clinical application (Lener et al. 2015). Recent studies have spotlighted the promising potential of EVs derived stem cells, such as mesenchymal stem cells (Helissey et al. 2022; Zhao et al. 2022) and adipose stem cells (Yang et al. 2024; Zhou et al. 2024) in mitigating and treating fibrotic diseases (Yang et al. 2022) mainly by inhibiting TGF‐β/Smad or PI3K/Akt signalling. Nevertheless, as the most critical progenitor cell for skin regeneration, the vital role of EVs derived from the primary cultured ESCs has not been thoroughly investigated (Duan et al. 2020).

Herein, we established an efficient ESC‐EVs isolation and application system and found it significantly inhibited the biological activity of HSFs in vitro and in vivo. ESC‐EVs not only inhibited HSFs’ migration and contractility but also prompted a noteworthy MET phenotypic shift in HSFs, marked by suppressed COLI, α‐SMA and N‐cadherin expression, alongside enhanced K1, K15 and E‐cadherin expression. On the RHS models, ESC‐EVs treated groups consistently displayed significant MET tendency and fibrosis alleviation. Therefore, ESC‐EVs exhibited great potential in catalysing MET, curtailing HSF activation and ameliorating abnormal fibrosis within HS. These findings suggest that ESC‐EVs uniquely reprogram fibrotic FBs toward a regenerative epithelial‐like state, distinguishing them from other EV‐based or conventional therapies.

To investigate the mechanism of ESC‐EVs on HS suppression, we further sequenced ESC‐EVs and found 173 miRNAs highly expressed miRNAs compared to FB‐EVs. Notably, members of the miR‐200 family (miR‐200s)—miR‐200a, miR‐200b, miR‐200c, miR‐141 and miR‐429—ranked prominently among the top 20 differentially expressed miRNAs. These results align with previous studies that prompt the vital involvement of miR‐200s in attenuating fibrotic processes (Liu et al. 2012; Li et al. 2014). Then, we assessed the miR‐200s expression in normal human skin, HS and mature scars by fluorescence in situ hybridization (FISH) and found that miR‐200s levels in HS were significantly decreased compared to normal skin, while a deficit resolved during scar maturation. However, When the ESC‐EVs were applied to the HSF or RHS, notably elevated miR‐200s could be observed, which reminded us that ESC‐EVs‐miR‐200s play a vital role between the crosstalk of ESCs and HSFs.

To clarify the target of ESC‐EVs in HSFs, GO/KEGG analysis of upregulated genes showed highlighted enrichment in pathways involving ZEB1 and ZEB2, paralogs belonging to the zinc‐finger E‐box‐binding homeobox (ZEB) transcription factor family known for promoting EMT (Bakir et al. 2020). Our investigation consistently confirmed heightened ZEB1 and ZEB2 expression in HS compared to normal skin and mature scars. Since extensive literature supports miR‐200s' role in suppressing EMT by targeting ZEB1/2 (Burk et al. 2008; Diaz‐Riascos et al. 2019; Bhome et al. 2022), we highly speculated that ESC‐EVs targeted ZEBs through miR‐200s. Notably, applying ESC‐EVs effectively elevated miR‐200s and mitigated ZEB1 and ZEB2 expression in HSFs, ultimately inducing its MET and alleviating HS. However, when we knocked down all five members of the miR‐200s using a multiplexed CRISPR/Cas9 system, the effect of ESC‐EVs on HSFs was significantly weakened, demonstrating the effects of ESC‐EVs on HSFs' MET are mainly dependent on the miR‐200s/ZEBs axis.

Clinically, current standard treatments for HS focus primarily on suppressing inflammation or collagen deposition, failing to address the fundamental dysregulation of FB phenotypic plasticity. In contrast, ESC‐EVs target the root pathological mechanism by reprogramming overactivated FBs toward a less fibro genic, epithelial‐like state—offering the potential for true tissue regeneration and durable therapeutic effects. Nevertheless, several challenges must be overcome before clinical translation. Although the ESCs in this study were ethically sourced from discarded human skin with informed consent, widespread clinical adoption will require stringent regulatory oversight. Additionally, scalable EV production and standardized purification protocols must be established (Guo et al. 2021; Thouvenot et al. 2024; Huang et al. 2025). Translational efforts must also account for interspecies differences in skin architecture and immune responses when extrapolating rodent model data to human patients. From a delivery perspective, the current perilesional injection approach suffers from uneven EV distribution and lacks sustained release. Emerging technologies, such as microneedle patches, could enhance bioavailability, improve dosing precision and increase patient compliance—addressing key limitations of existing methods.

This study has its limitations. Our investigation primarily examined the effects of ESC‐EVs on HSF, potentially overlooking other cellular and molecular components that may contribute to their therapeutic efficacy. Several recent studies demonstrated ESC‐EVs could modulate keratinocyte behaviour (Than et al. 2019), angiogenesis (Duan et al. 2020) and macrophage polarization (Zhou et al. 2020; Wang et al. 2022) in the wound healing environment, indicating their broader impact on cellular crosstalk and tissue remodelling in HS warrants further investigation. However, the primary objective of this project was to provide evidence of novel ESC‐EVs that relieve HS, explore its most critical mechanisms, and identify avenues for future investigation. To our knowledge, this study innovatively revealed that ESC‐EVs can induce MET, attenuate HSF activation and mitigate fibrosis in vitro and in vivo by modulating ZEB1 and ZEB2 through miR‐200s. These findings provided a novel therapeutic strategy and elucidated the mechanism for using ESC‐EVs and miR‐200s as a clinical treatment of HS and other fibrotic disorders.

5. Conclusion

In summary, our study highlights a significant breakthrough in understanding the role of ESC‐EVs in alleviating HS and uncovers a previously unknown mechanism by which ESC‐EVs could induce HSFs’ MET via miR‐200s/ZEBs axis. As soon as up‐taken by HSF, the ESC‐EVs‐miR‐200s directly targeted to ZEB1/2 and induced the MET of HSF, ultimately attenuating HSFs’ biological functions and alleviating HS (shown as Figure 9). Our findings are expected to provide novel targets and strategies for the precise clinical treatment of HS and other skin fibrosis diseases.

Author Contributions

Miao Zhen: Investigation (equal), validation (equal), writing – original draft (lead). Juntao Xie: Conceptualization (equal), investigation (equal), validation (equal). Rui Yang: Data curation (equal), formal analysis (equal), validation (equal). Lijuan Liu: Data curation (supporting), formal analysis (equal), methodology (equal). Hengdeng Liu: Formal analysis (equal), visualization (supporting). Xuefeng He: Data curation (supporting), formal analysis (supporting). Suyue Gao: Methodology (equal), validation (supporting). Junyou Zhu: Formal analysis (supporting). Jingting Li: Data curation (equal), methodology (supporting). Bin Shu: Project administration (equal), supervision (equal), writing – review and editing (equal). Peng Wang: Conceptualization (lead), funding acquisition (lead), project administration (lead), supervision (equal), writing – review and editing (equal).

Conflicts of Interest

The authors declare no conflict of interest.

Supporting information

Supplementary Figures: jev270160‐sup‐0001‐FigureS1‐S9.docx

JEV2-14-e70160-s001.docx (6.1MB, docx)

Acknowledgements

This work was mainly supported by the National Natural Science Foundation of China (No. 82002043, 82172207, 82273561), Natural Science Foundation of Guangdong Province (No. 2023A1515010265, 2023A1515010146) and Guangzhou Municipal Science and Technology Project (No. 2025A04J4078).

Zhen, M. , Xie J., Yang R., et al. 2025. “Epidermal Stem Cell‐Derived Extracellular Vesicles Induce Fibroblasts Mesenchymal‐Epidermal Transition to Alleviate Hypertrophic Scar Formation via miR‐200s Inhibition of ZEB1 and 2.” Journal of Extracellular Vesicles 14, no. 9: e70160. 10.1002/jev2.70160

Miao Zhen, Juntao Xie and Rui Yang contributed equally to this work.

Funding: This study was funded by the National Natural Science Foundation of China (Grant number 82002043, 82172207, 82273561), Guangzhou Municipal Science and Technology Project (Grant number 2025A04J4078), Natural Science Foundation of Guangdong Province (Grant number 2021A1515011806, 2023A1515010146, 2023A1515010265).

Contributor Information

Bin Shu, Email: shubin@mail.sysu.edu.cn.

Peng Wang, Email: wangp276@mail.sysu.edu.cn.

Data Availability Statement

All data are available in the main text and/or the Supplementary materials, tables and figures. Datasets related to this article can be found in the National Centre for Biotechnology Information Gene Expression Omnibus database, under the study accession number GSE287895. The data that support the findings of this study are available from the corresponding author upon reasonable request.

References

  1. Bakir, B. , Chiarella A. M., Pitarresi J. R., and Rustgi A. K.. 2020. “EMT, MET, Plasticity, and Tumor Metastasis.” Trends in Cell Biology 30, no. 10: 764–776. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Berman, B. , Maderal A., and Raphael B.. 2017. “Keloids and Hypertrophic Scars: Pathophysiology, Classification, and Treatment.” Dermatologic Surgery 43, no. 1: S3–S18. [DOI] [PubMed] [Google Scholar]
  3. Bhome, R. , Emaduddin M., James V., et al. 2022. “Epithelial to Mesenchymal Transition Influences Fibroblast Phenotype in Colorectal Cancer by Altering miR‐200 Levels in Extracellular Vesicles.” Journal of Extracellular Vesicles 11, no. 5: e12226. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Blanpain, C. , and Fuchs E.. 2006. “Epidermal Stem Cells of the Skin.” Annual Review of Cell and Developmental Biology 22: 339–373. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Boyer, M. J. , Kimura Y., Akiyama T., et al. 2020. “Endothelial Cell‐Derived Extracellular Vesicles Alter Vascular Smooth Muscle Cell Phenotype Through High‐Mobility Group Box Proteins.” Journal of Extracellular Vesicles 9, no. 1: 1781427. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Burk, U. , Schubert J., Wellner U., et al. 2008. “A Reciprocal Repression Between ZEB1 and Members of the miR‐200 Family Promotes EMT and Invasion in Cancer Cells.” Embo Reports 9, no. 6: 582–589. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Diaz‐Riascos, Z. V. , Ginesta M. M., Fabregat J., et al. 2019. “Expression and Role of MicroRNAs From the miR‐200 Family in the Tumor Formation and Metastatic Propensity of Pancreatic Cancer.” Molecular Therapy‐Nucleic Acids 17: 491–503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Dibsy, R. , Bremaud E., Mak J., Favard C., and Muriaux D.. 2023. “HIV‐1 Diverts Cortical Actin for Particle Assembly and Release.” Nature Communications 14: 6945. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Duan, M. , Zhang Y., Zhang H., Meng Y., Qian M., and Zhang G.. 2020. “Epidermal Stem Cell‐Derived Exosomes Promote Skin Regeneration by Downregulating Transforming Growth Factor‐β1 in Wound Healing.” Stem Cell Research & Therapy 11, no. 1: 452. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Finnerty, C. C. , Jeschke M. G., Branski L. K., Barret J. P., Dziewulski P., and Herndon D. N.. 2016. “Hypertrophic Scarring: The Greatest Unmet Challenge After Burn Injury.” Lancet 388, no. 10052: 1427–1436. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Fujiwara, H. , Tsutsui K., and Morita R.. 2018. “Multi‐Tasking Epidermal Stem Cells: Beyond Epidermal Maintenance.” Development Growth and Differentiation 60, no. 9: 531–541. [DOI] [PubMed] [Google Scholar]
  12. Guo, S. , Debbi L., Zohar B., et al. 2021. “Stimulating Extracellular Vesicles Production From Engineered Tissues by Mechanical Forces.” Nano Letters 21: 2497–2504. [DOI] [PubMed] [Google Scholar]
  13. Helissey, C. , Guitard N., Théry H., et al. 2022. “Two New Potential Therapeutic Approaches in Radiation Cystitis Derived From Mesenchymal Stem Cells: Extracellular Vesicles and Conditioned Medium.” Biology (Basel) 11, no. 7: 980. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Herberts, C. A. , Kwa M. S. G., and Hermsen H. P. H.. 2011. “Risk Factors in the Development of Stem Cell Therapy.” Journal of Translational Medicine 9: 29. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Huang, G.‐L. , Sun J., Lu Y., et al. 2019. “MiR‐200 Family and Cancer: From a Meta‐Analysis View.” Molecular Aspects of Medicine 70: 57–71. [DOI] [PubMed] [Google Scholar]
  16. Huang, H. , Xiao L., Fang L., et al. 2025. “Static Topographical Cue Combined With Dynamic Fluid Stimulation Enhances the Macrophage Extracellular Vesicle Yield and Therapeutic Potential for Bone Defects.” ACS Nano 19: 8667–8691. [DOI] [PubMed] [Google Scholar]
  17. Huang, J. , Zhang H., Fan X., and Guo J.. 2022. “Inhibition of miR‐497 Attenuates Oral Submucous Fibrosis by Inhibiting Myofibroblast Transdifferentiation in Buccal Mucosal Fibroblasts.” Oral Health & Preventive Dentistry 20: b3276183. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Huang, S. , Hu Z., Wang P., et al. 2020. “Rat Epidermal Stem Cells Promote the Angiogenesis of Full‐Thickness Wounds.” Stem Cell Research & Therapy 11: 344. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Jiang, J. , Li J., Zhou X., Zhao X., Huang B., and Qin Y.. 2022. “Exosomes Regulate the Epithelial‐Mesenchymal Transition in Cancer.” Frontiers in Oncology 12: 864980. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Jiang, X. , Yang J., Lin Y., et al. 2023. “Extracellular Vesicles Derived From Human ESC‐MSCs Target Macrophage and Promote Anti‐Inflammation Process, Angiogenesis, and Functional Recovery in ACS‐Induced Severe Skeletal Muscle Injury.” Stem Cell Research & Therapy 14: 331. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kabadi, A. M. , Ousterout D. G., Hilton I. B., and Gersbach C. A.. 2014. “Multiplex CRISPR/Cas9‐Based Genome Engineering From a Single Lentiviral Vector.” Nucleic Acids Research 42, no. 19: e147. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Kaufer, B. B. 2013. “Detection of Integrated Herpesvirus Genomes by Fluorescence In Situ Hybridization (FISH).” Methods in Molecular Biology (Clifton, N.J.) 1064: 141–152. [DOI] [PubMed] [Google Scholar]
  23. Kim, D. S. , Cho H. J., Choi H. R., Kwon S. B., and Park K. C.. 2004. “Isolation of Human Epidermal Stem Cells by Adherence and the Reconstruction of Skin Equivalents.” Cellular and Molecular Life Sciences 61, no. 21: 2774–2781. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Lener, T. , Gimona M., Aigner L., et al. 2015. “Applying Extracellular Vesicles Based Therapeutics in Clinical Trials—An ISEV Position Paper.” Journal of Extracellular Vesicles 4, no. 1: 30087. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Li, P. , He Q. Y., and Luo C. Q.. 2014. “Overexpression of miR‐200b Inhibits the Cell Proliferation and Promotes Apoptosis of human Hypertrophic Scar Fibroblasts In Vitro.” Journal of Dermatology 41, no. 10: 903–911. [DOI] [PubMed] [Google Scholar]
  26. Lin, S. , Yu Z., Chen D., et al. 2020. “Progress in Microfluidics‐Based Exosome Separation and Detection Technologies for Diagnostic Applications.” Small 16, no. 9: 1903916. [DOI] [PubMed] [Google Scholar]
  27. Liu, Y. , Yang D., Xiao Z., and Zhang M.. 2012. “miRNA Expression Profiles in Keloid Tissue and Corresponding Normal Skin Tissue.” Aesthetic Plastic Surgery 36, no. 1: 193–201. [DOI] [PubMed] [Google Scholar]
  28. Machesney, M. , Tidman N., Waseem A., Kirby L., and Leigh I.. 1998. “Activated Keratinocytes in the Epidermis of Hypertrophic Scars.” American Journal of Pathology 152, no. 5: 1133. [PMC free article] [PubMed] [Google Scholar]
  29. Moore, K. A. , and Lemischka I. R.. 2006. “Stem Cells and Their Niches.” Science 311, no. 5769: 1880–1885. [DOI] [PubMed] [Google Scholar]
  30. Ngo, P. , Ramalingam P., Phillips J. A., and Furuta G. T.. 2006. “Collagen Gel Contraction Assay.” Methods in Molecular Biology 341: 103. [DOI] [PubMed] [Google Scholar]
  31. Ogawa, R. 2017. “Keloid and Hypertrophic Scars Are the Result of Chronic Inflammation in the Reticular Dermis.” International Journal of Molecular Sciences 18, no. 3: 606. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Ogawa, R. , Dohi T., Tosa M., Aoki M., and Akaishi S.. 2021. “The Latest Strategy for Keloid and Hypertrophic Scar Prevention and Treatment: The Nippon Medical School (NMS) Protocol.” Journal of Nippon Medical School = Nippon Ika Daigaku Zasshi 88, no. 1: 2–9. [DOI] [PubMed] [Google Scholar]
  33. Rabello, F. B. , Souza C. D., and J. A. Farina, Jr . 2014. “Update on Hypertrophic Scar Treatment.” Clinics 69, no. 8: 565–573. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Ru, Y. , Kechris K. J., Tabakoff B., et al. 2014. “The multiMiR R Package and Database: Integration of microRNA‐target Interactions Along With Their Disease and Drug Associations.” Nucleic Acids Research 42, no. 17: e133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Than, U. T. T. , Leavesley D. I., and Parker T. J.. 2019. “Characteristics and Roles of Extracellular Vesicles Released by Epidermal Keratinocytes.” Journal of the European Academy of Dermatology and Venereology: JEADV 33: 2264–2272. [DOI] [PubMed] [Google Scholar]
  36. Théry, C. , Witwer K. W., Aikawa E., Alcaraz M. J., et al. 2018. “Minimal Information for Studies of Extracellular Vesicles 2018 (MISEV2018): A Position Statement of the International Society for Extracellular Vesicles and Update of the MISEV2014 Guidelines.” Journal of Extracellular Vesicles 7, no. 1: 1535750. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Thouvenot, E. , Charnay L., Burshtein N., et al. 2024. “High‐Yield Bioproduction of Extracellular Vesicles From Stem Cell Spheroids via Millifluidic Vortex Transport.” Advanced Materials (Deerfield Beach, Fla.) 37: e2412498. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Wang, P. , Hu Z., Cao X., et al. 2019. “Fibronectin Precoating Wound Bed Enhances the Therapeutic Effects of Autologous Epidermal Basal Cell Suspension for Full‐Thickness Wounds by Improving Epidermal Stem Cells' Utilization.” Stem Cell Research & Therapy 10: 154. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Wang, P. , Peng Z., Yu L., et al. 2024. “Verteporfin‐Loaded Bioadhesive Nanoparticles for the Prevention of Hypertrophic Scar.” Small Methods 8: e2301295. [DOI] [PubMed] [Google Scholar]
  40. Wang, P. , Shu B., Xu Y., et al. 2017. “Basic Fibroblast Growth Factor Reduces Scar by Inhibiting the Differentiation of Epidermal Stem Cells to Myofibroblasts via the Notch1/Jagged1 Pathway.” Stem Cell Research & Therapy 8, no. 1: 114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Wang, P. , Theocharidis G., Vlachos I. S., et al. 2022. “Exosomes Derived From Epidermal Stem Cells Improve Diabetic Wound Healing.” Journal of Investigative Dermatology 142: 2508–2517.e13. [DOI] [PubMed] [Google Scholar]
  42. Wu, T. , Hu E., Xu S., et al. 2021. “clusterProfiler 4.0: A Universal Enrichment Tool for Interpreting Omics Data.” Innovation (Camb) 2, no. 3: 100141. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Yang, S. , Sun Y., and Yan C.. 2024. “Recent Advances in the Use of Extracellular Vesicles From Adipose‐Derived Stem Cells for Regenerative Medical Therapeutics.” Journal of Nanobiotechnology 22: 316. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Yang, Y. , Liu Y., Chai Y., et al. 2022. “Exosomes in Pathogenesis, Diagnosis, and Treatment of Pulmonary Fibrosis.” Frontiers in Pharmacology 13: 927653. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Youssef, K. K. , and Nieto M. A.. 2024. “Epithelial‐Mesenchymal Transition in Tissue Repair and Degeneration.” Nature Reviews. Molecular Cell Biology 25: 720–739. [DOI] [PubMed] [Google Scholar]
  46. Yu, L. , Cao C., Li X., et al. 2022. “Complete Loss of miR‐200 Family Induces EMT Associated Cellular Senescence in Gastric Cancer.” Oncogene 41, no. 1: 26–36. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Yuan, F.‐L. , Sun Z.‐L., Feng Y., et al. 2019. “Epithelial‐Mesenchymal Transition in the Formation of Hypertrophic Scars and Keloids.” Journal of Cellular Physiology 234: 21662–21669. [DOI] [PubMed] [Google Scholar]
  48. Zhao, W. , Zhang R., Zang C., et al. 2022. “Exosome Derived From Mesenchymal Stem Cells Alleviates Pathological Scars by Inhibiting the Proliferation, Migration and Protein Expression of Fibroblasts via Delivering miR‐138‐5p to Target SIRT1.” International Journal of Nanomedicine 17: 4023–4038. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Zhou, B. , Chen Q., Zhang Q., Tian W., Chen T., and Liu Z.. 2024. “Therapeutic Potential of Adipose‐Derived Stem Cell Extracellular Vesicles: From Inflammation Regulation to Tissue Repair.” Stem Cell Research & Therapy 15: 249. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Zhou, S. , Wang W., Zhou S., Zhang G., He J., and Li Q.. 2019. “A Novel Model for Cutaneous Wound Healing and Scarring in the Rat.” Plastic and Reconstructive Surgery 143, no. 2: 468–477. [DOI] [PubMed] [Google Scholar]
  51. Zhou, X. , Brown B. A., Siegel A. P., et al. 2020. “Exosome‐Mediated Crosstalk Between Keratinocytes and Macrophages in Cutaneous Wound Healing.” ACS Nano 14: 12732–12748. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Supplementary Figures: jev270160‐sup‐0001‐FigureS1‐S9.docx

JEV2-14-e70160-s001.docx (6.1MB, docx)

Data Availability Statement

All data are available in the main text and/or the Supplementary materials, tables and figures. Datasets related to this article can be found in the National Centre for Biotechnology Information Gene Expression Omnibus database, under the study accession number GSE287895. The data that support the findings of this study are available from the corresponding author upon reasonable request.

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