Conjugate electrospinning dermal-adipose bilayered skin: structural integration promoting regenerative skin remodeling

Abstract

The skin is a three-dimensional organ composed of multilayered tissues, in which the epidermis, dermis, and subcutaneous adipose layer cooperate to maintain protection, thermoregulation, and repair. Although recent advances in tissue-engineered skin substitutes have improved cutaneous regeneration, strategies that simultaneously promote dermal and adipose restoration remain limited. Here, we developed a bi-layered tissue-engineered skin scaffold with dermal–adipose architecture fabricated by conjugate electrospinning of polycaprolactone solutions containing acellular dermal matrix (ADM) or decellularized adipose tissue (DAT). The construct exhibited dual bioactivity: stimulating fibroblast proliferation and collagen remodeling in the dermal layer, while promoting adipose-derived stem cell proliferation and adipogenesis in the adipose layer. In a full-thickness nude rat wound model, the scaffold enhanced vascularization, modulated inflammation, and accelerated regeneration of both dermal and adipose tissues. These findings demonstrate a versatile platform for multilayered skin tissue engineering and provide new insight into dermal–adipose synergistic regeneration.

Graphical abstract

Supplementary Information

The online version contains supplementary material available at 10.1186/s12951-026-04051-8.

Introduction

The skin, as a composite organ formed by multiple tissue layers, derives its self-renewal, barrier, and immunological functions from the intricate interplay of these stratified structures [1–3]. Serving as the primary physical barrier and immunological interface between the human body and external environment, full-thickness skin defects caused by trauma, neoplasms, or metabolic disorders frequently disrupt systemic homeostasis through multi-layer tissue loss [4, 5]. Conventional clinical approaches employing autologous skin grafts, skin flaps, or composite tissue flaps inevitably cause donor site morbidity [6], highlighting the urgent need for tissue-engineered skin substitutes that eliminate secondary defects.

Over the years, researchers have proposed the endogenous tissue regeneration theory as a goal for wound repair. This strategy relies on placing functional materials within damaged tissue, where the material-created environment directs the body’s cells to regenerate tissue [7, 8]. These cell-free matrix materials create a three-dimensional supportive system that facilitates proliferation, migration, differentiation, and functional reconstruction of host cells. Compared to skin grafts requiring external cell intervention, this technology offers significant advantages: complete reliance on cells for regeneration avoids risks. Furthermore, by eliminating in vitro cell culture requirements, it demonstrates greater feasibility in ethical review and regulatory approval processes [9]. Based on this principle, commercial products like Apligraf^® and Integra^® have entered production phases [10]. However, existing commercial biomaterials still exhibit substantial limitations. While their single-component composition and simplified structural designs facilitate industrial-scale production, these engineered advancements neglect a critical element of native skin physiology: the subcutaneous adipose layer, a three-dimensional, intricate tissue with essential metabolic functions. Recent discoveries in adipose biology have demonstrated that subcutaneous adipose tissue holds critical importance. Subcutaneous adipose tissue not only serves as a triglyceride reservoir but also functions as a dynamic endocrine organ, releasing fatty acids and immunomodulatory cytokines during wound healing [11, 12]. Current studies on multilayer skin lack for attention on adipose layers [13, 14]. Notably, engineered skin substitutes with adipose inductive components and architectures to achieve multi-layered skin and subcutaneous tissue regeneration remain unexplored.

The specific bioactivity of dermal and subcutaneous adipose tissue originates from the corresponding extracellular matrix (ECM) component [15]. The unique 3D architectural integration of structural proteins and dynamic signaling molecules of ECM endows it with inherent tissue-specific bioactivity [16]. These components orchestrate cellular adhesion and differentiation through spatially organized biochemical gradients and biomechanical cues [17]. Decellularized ECM (dECM) scaffolds, engineered from tissue or cell-derived matrices, preserve this bioactive repertoire while mitigating immunogenicity, enabling groundbreaking applications in tissue-specific regeneration [18, 19]. Despite the proven efficacy of single-source dECM scaffolds in tissue-specific regeneration, such as acellularized dermal matrix (ADM) facilitating dermal repair through type I collagen networks, their inability to replicate the multilayer and multi-tissue interactions of native skin architecture remains a critical limitation [20]. For instance, decellularized adipose tissue (DAT), characterized by elastin-rich microenvironments and adipokine-mediated metabolic regulation, exhibits complementary roles in adipogenesis and immunomodulation but has hitherto not been systematically integrated with ADM [21, 22]. Combining ADM and DAT provides a biologically rational strategy to reconstruct the dermal–hypodermal interface. ADM contributes a collagen-rich, mechanically stable framework supporting fibroblast infiltration and dermal remodeling [23], while DAT offers adipose-derived ECM cues that promote angiogenesis and adipogenesis [24]. Their integration enables reciprocal interactions between dermal and adipose lineages, achieving coordinated regeneration of both layers. This ECM–ECM synergy forms the conceptual and innovative basis of our ADM/DAT composite design. This unresolved challenge fundamentally constrains the development of stratified scaffolds capable of emulating a multilayered skin construct. Our study sought to address this unmet need by innovatively engineering a bilayer ADM/DAT composite skin, aiming to decode their synergistic mechanochemical interactions and validate their capacity to restore full-thickness skin defects through dermal and adipose regeneration.

Recognizing the therapeutic potential of adipose and dermal originated ECM, we emphasize the importance of selecting appropriate fabrication methodologies for 3D adipose and dermal integrative constructs. Among the current widely used manufacturing methods for 3D tissue-engineered skin, 3D printing holds a pivotal position. Using 3D printing technology, precise three-dimensional skin structures can be produced on demand, enabling the construction of multilayered skin containing capillary networks and skin appendages, thereby advancing personalized regenerative medicine [25]. However, existing 3D printing remains limited in reaching micro-nanoscale levels. In contrast, electrospinning has emerged as an essential technique for tissue engineering due to its capacity to generate nanofibrous matrices with high porosity, surface area, and ECM-mimicking topography that facilitates cellular adhesion and proliferation. Nevertheless, conventional electrospinning membranes typically form thin, densely packed layers unsuitable for constructing stratified 3D architectures. To overcome this limitation, conjugate electrospinning (Co-ES) technology—employing dual opposed polarity nozzles to deposit interpenetrating positively and negatively charged fibers—enables fabrication of scaffolds with a 3D porous structure while preserving nanofibrous features. By establishing interconnected pore networks through alternating fiber deposition, conjugate electrospinning scaffolds inherently support cellular infiltration and oxygen diffusion. Accordingly, this technique represents an optimal choice for creating multi-layered skin equivalents, coordinating dermal and adipose regeneration.

Herein, we present a conjugate electrospinning technique to fabricate bilayered skin combining ADM and DAT layers for coordinated dermal-adipose regeneration. The ADM Conjugate-Electrospinning (ADM Co-ES) scaffolds specifically enhance fibroblast-mediated collagen synthesis and deposition, whereas the DAT Conjugate-Electrospinning (DAT Co-ES) scaffolds facilitate adipogenesis and lipid droplet formation in vitro. In vivo evaluation using full-thickness dorsal wounds in nude rats confirmed accelerated wound closure, enhanced vascularization, reduced inflammation, and simultaneous regeneration of both dermal and adipose layers. Transcriptomic analysis prompted the possible mechanisms underlying this dual regenerative capacity. The engineering of a bilayered skin substitute, integrating structural biomimicry through conjugate electrospinning with functional bioactivity via tissue-specific ECM integration, establishes a versatile platform for developing products with 3D stratified structures for tissue regeneration.

Materials and methods

Materials

All reagents and materials used in this study were of analytical grade and used without further purification unless otherwise specified. Polycaprolactone (PCL, Suzhou HuanTong Technology Co., China) and hexafluoroisopropanol (HFIP, Biosharp, China) were used for electrospinning. Phosphate-buffered saline (PBS, HyClone, USA), 0.25% trypsin–EDTA (Biosharp, China), DNase I and RNase A (Solarbio, China), and pepsin (Sigma-Aldrich, USA) were employed during decellularization procedures. Paraformaldehyde solution, hematoxylin–eosin (H&E) staining kit, Masson trichrome staining kit, Sirius Red staining kit, and Oil Red O dye were obtained from Shanghai Yuanye Biotechnology Co. (China). DAPI staining solution was purchased from Beyotime Biotechnology (China). Isopropanol, ethanol, and xylene were supplied by Sinopharm Chemical Reagent Co. (China). For Western blot, RIPA lysis buffer, PMSF, phosphatase inhibitors, BCA protein assay kits, SDS–PAGE reagents, and ECL substrates were purchased from Wuhan Boster (China). For ELISA, Commercial rat ELISA kits were obtained from Yiqiao Biotechnology (China), and absorbance was measured using a microplate reader (SpectraMax 190; Molecular Devices, USA). For qPCR, total RNA was extracted using TriQuick reagent (Solarbio, China), reverse transcription was performed using Evo M-MLV RT Mix (Accurate Biotechnology, China), and qPCR was conducted using PerfectStart Green qPCR SuperMix (TransGen Biotech, China). PVDF membranes were obtained from Beyotime (China), and HRP-conjugated secondary antibodies from Jackson ImmunoResearch (USA) and Sanying (China). All culture media and supplements were purchased from Gibco (USA).

Major equipment included a constant temperature shaker (Saiwei Instrument Co., Wuhan, China), a freeze dryer (Boyikang Instrument Co., Beijing, China), an electrospinning apparatus (Foshan Qingzi Precision Instrument Co., China), a contact angle measuring device (Shending Precision Instrument Co., China), a universal testing machine (Shimadzu, Japan), and a confocal laser scanning microscope (Leica, Germany). Homogenizer, refrigerated centrifuge, vertical electrophoresis/blotting system (YEASEN, China), −80 °C freezer (Meiling, China), microvolume spectrophotometer (Shanghai Jiapeng, China), and real-time PCR system (QuantStudio 1; Thermo Fisher, USA). All reagents and instruments were used following the manufacturers’ standard protocols unless otherwise specified.

Preparation of human ADM

The human skin tissue was derived from the skin tissue excised by plastic surgery in Wuhan Union Hospital, which has been approved by the Ethics Committee of Union Hospital. The steps of decellularization are as follows: First, the skin and subcutaneous adipose layer were cleanly separated, and no obvious adipose residue was observed by the nude eye. The skin tissue was then washed 3–5 times with normal saline for 1–2 min each time, after which the skin was soaked in 1% iodophor overnight for disinfection. After overnight rinsing with normal saline, the skin tissue was soaked in PBS solution containing 1% Penicillin-Streptomycin for 30 min for re-disinfection. The skin tissue was flushed with PBS 3–5 times. The disinfected skin tissue was irradiated with gamma rays at a dose rate of 1000 Gy/h and a total dose of 25 kGy. After irradiation, soak in 0.05% pancreatic enzyme/EDTA solution for 24 h. PBS was used to flush skin tissue 3–5 times for 1–2 min each time. Skin tissue was soaked in PBS with a DNase concentration of 100 μg/mL and an RNase concentration of 100 μg/mL. The skin tissue was transferred to a constant temperature shaker at 37 °C overnight. After overnight incubation, the skin tissue was immersed in PBS solution containing 1% Penicillin-Streptomycin and treated in a constant temperature shaker for 24 h. Finally, the processed tissues were dehydrated, freeze-dried, and ground into powder.

Preparation of human DAT

Human adipose tissue samples were derived from plastic surgery procedures at Wuhan Union Hospital. The preparation process was as follows: First, the adipose tissue was separated and trimmed into approximately cubic pieces (~5 mm³; ~1.7 × 1.7 × 1.7 mm) to ensure a consistent surface area for uniform processing. After pruning, 25 mL of adipose tissue and 25 mL of PBS solution were placed in a 50 mL centrifuge tube, followed by 5 freeze-thaw cycles alternating between −80 °C and 37 °C. The 25 mL PBS solution was replaced with 25 mL pancreatic enzyme/EDTA solution, and the sample was placed on a constant temperature shaking table at 37 °C for 16 h. The pancreatic enzyme/EDTA solution was then decanted and replaced with 25 mL PBS solution, followed by shaking on the table for 15 min and rinsing three times. The PBS solution was then decanted and replaced with 25 mL isopropyl alcohol solution for degreasing treatment, and the sample was placed in a constant temperature shaking bed at 37 °C for 48 h, with the isopropyl alcohol replaced once at 24 h. The tissues after degreasing with isopropyl alcohol were immersed in PBS solution with DNase concentration of 100 μg/mL and RNase concentration of 100 μg/mL and then in a constant temperature shaking bed at 37 °C for 16 h. For further degreasing, the tissue was placed in the same amount of isopropyl alcohol in a shaker for 16 h. Next, the tissues were cleaned with PBS solution 3 times for 1–2 min each time. Finally, the processed tissues were dehydrated, freeze-dried, and ground into powder.

Conjugate electrospinning

First, ADM powder and PCL particles were weighed respectively, according to the ratios ADM:PCL=50 mg:950 mg, 100 mg:900 mg, 150 mg:850 mg, and 200 mg:800 mg. Specifically, the ADM and PCL mixed solutes were divided into four groups, labeled as 5%ADM, 10%ADM, 15%ADM, and 20%ADM, respectively. Similarly, DAT powder and PCL particles were mixed in the same proportions, and a total of four groups of DAT and PCL mixed solutes were labeled as 5% DAT, 10% DAT, 15% DAT, and 20% DAT, respectively. Another control group consisted of 1000 mg of PCL. Then, the control group and the 5% ADM to 20% ADM, 5% DAT to 20% DAT, and PCL groups were placed into centrifuge tubes with 10 mL of hexafluoro-isopropyl alcohol (HFIP) and incubated on a constant temperature shaking table at 37 °C for 16 h. Next, 5 mL syringes were filled with 5% ADM using an 18G electrospinning needle. The syringe was placed on the injection pump of the electrospinning machine, and the high-pressure joints of the positive and negative poles were connected to the electrospinning needles on both sides. A cylindrical receiver, covered with tinfoil and attached to the rotating cup, was used for collection. Electrospinning parameters were adjusted to a distance of 15 cm, a voltage of ±20kV, a speed of 2.5 mL/h, and a rotating speed of 200 r/min, with the time set for 2 h. The electrospun product deposited on the tinfoil was obtained. The electrospun product was removed with a blade and collected in a drying bag for storage. Similarly, all electrospun scaffolds from the PCL control group and the ADM and DAT groups were obtained. After the best biocompatibility of ADM and DAT was determined, ADM and DAT composite scaffolds were constructed using the same method.

Extraction and culture of adipose-derived stem cells (ADSCs)

All human tissue cells obtained in this experiment were procured with signed informed consent from the patients and were approved by the ethical review board of Huazhong University of Science and Technology. First, the fresh adipose tissue was immersed in a sterile PBS solution and washed 3 times for 5–8 min each time. The washed adipose tissue was placed in a sterile centrifuge tube. Then the complete medium solution containing 0.2% IV collagenase was evenly mixed with the same volume of adipose tissue and placed in a constant temperature shaking table at 37 degrees Celsius for digestion for 4 h. The digested adipose tissue was placed in a centrifuge at a set speed of 1500 rpm and centrifuged for 5 min. The precipitation after centrifugation was collected, cleaned, and centrifuged with PBS again, and the cell suspension was resuspended after centrifugation. The cell suspension was filtered, the filtrate collected, and centrifuged again at 1500 rpm for 5 min. The precipitate was re-suspended using a medium. The suspensions were inoculated in petri dishes, placed in cell incubators, and changed every 3 days. Once the cell confluence reached 80%, passaging was performed.

Extraction and culture of fibroblasts (FBs)

All human tissue cells obtained in this experiment were procured with signed informed consent from the patients and were approved by the ethical review board of Huazhong University of Science and Technology. Firstly, chloramphenicol, neutral protease II and collagenase NB4 were dissolved in PBS, respectively, and prepared with 0.25% (w/v) chloramphenicol solution, 0.25% (w/v) neutral protease II solution and 0.3% (w/v) collagenase NB4 solution. All solutions were then filtered using a disposable 0.22 μm filter to obtain a sterile solution for use in the following steps. At the same time, the sterilized surgical instruments, required reagents, and consumables were placed on a super-clean table and irradiated by UV for 1 h. The skin tissue was soaked in 0.25% (w/v) chloramphenicol solution for 3 times, for 5–10 min each time, and then washed with PBS solution 3 times. The cleaned skin was then placed in 0.25% (w/v) neutral protease solution for overnight digestion at 4 ℃. The epidermis was then removed with tweezers, and the dermis was cut, placed in 0.3% collagenase NB4, and digested overnight at room temperature. The digestive solution was centrifuged at 1500 rpm for 5 min, the supernatant and undigested tissue were discarded, and after cleaning and precipitation with PBS, the cells were centrifuged at 1500 rpm for 5 min, and the cells were re-suspended with complete medium and filtered with a 70 µm filter. 1.5×10⁶ cells were inoculated in every 10 mL medium, placed in an incubator, and the fluid was changed every 2–3 days.

Cell inoculation and cellular compatibility evaluation

First, the Petri dish was removed from the cell incubator and placed on the ventilated operating table (which had been subjected to UV irradiation for 30 min prior to the procedure), and the medium was aspirated using a pipette. Sterile PBS solution was slowly added, and then the PBS solution was discarded. Subsequently, 2–3 mL of pancreatic enzyme solution was added to each dish, and the dishes were placed back in the cell incubator for 5 min. After this incubation, the Petri dish was removed and observed under a microscope, at which point the cells were noted to have become rounded and detached from the dish surface. The dish was then transferred back to the cell incubator, and 4–6 mL of complete medium was added for neutralization to halt digestion. The cell suspension was transferred to a centrifuge tube, placed in a centrifuge, and centrifuged at 1500 rpm for 5 min. The supernatant was discarded, and the cell pellet was resuspended in 1 mL of medium, and the cell count was performed. After cell counting, cell viability was determined by trypan blue exclusion assay. Only fibroblast and ADSCs suspensions exhibiting > 95% viability were used for seeding onto electrospun scaffolds. The disinfected electrospinning scaffold was immersed in complete medium for 5 min, after which the medium was discarded. Ten microliters of cell suspension were then carefully dripped onto each electrospinning scaffold. The electrospinning scaffolds were placed in the cell incubator for 2 h, and then 1 mL of medium was added to each. The medium was changed once daily thereafter.

To comprehensively evaluate the biocompatibility of scaffold materials, the cell proliferation activity of implanted scaffold materials was detected, including cell survival and death staining, CCK-8 quantitative detection, and phalloidin staining. The selected time points were day 1, day 4, and day 7 post-cell implantation. All experimental procedures described in this abstract were performed under dark conditions. For cell survival and death staining, the cells were stained with Calcein-AM and PI dyes, diluted 500-fold and 1000-fold, respectively, and incubated for 30 min. Observation was conducted using a confocal laser microscope. Subsequently, cell proliferation was quantitatively detected using a CCK-8 kit. 10 µL of CCK-8 stock solution were diluted with 100 µL of medium and incubated in a cell culture incubator for 2 h. Absorbance per well (at 450 nm) was measured using an enzyme-linked immunosorbent assay (ELISA) reader. The absorbance data for each group were then analyzed. Finally, the cytoskeleton was stained with phalloidin dye. After three washes with sterile PBS, the electrospinning scaffold was fixed with paraformaldehyde for 30 min. The cell membranes were permeabilized with a 0.25% Triton X-100 PBS solution. A 1% bovine serum albumin (BSA) solution was prepared, and tetramethylrhodamine B isothiocyanate (TRITC)-conjugated phalloidin was diluted 500-fold with the BSA solution. The prepared phalloidin dye was applied to the electrospinning scaffold. After 2 h of incubation, the scaffold was washed three times with PBS, and observation was performed using a confocal microscope.

Lipid induction and detection

To test the lipid-inducing properties of DAT-containing electrospinning scaffolds, lipid differentiation was induced using an integrated lipid induction solution, and the content of the best proliferative activity was compared with that of the PCL scaffolds without DAT. First, the electrospinning scaffolds containing DAT were cultured by ADSCs for 7 days. Subsequently, adipogenic differentiation was induced using a regimen of three days in adipogenic induction medium followed by one day in adipogenic maintenance medium. This induction cycle was repeated one to four times. The lipid droplets formed were stained with BODIPY dye, and lipid induction was assessed after each cycle. The specific experimental steps were as follows: First, the observation time points for lipid formation were selected as day 4, day 8, day 12, and day 16. The electrospinning scaffolds were washed three times with PBS. Then, the scaffolds were fixed with 4% paraformaldehyde for 30 min, followed by three washes with PBS. The cell membranes were permeabilized using a 0.25% Triton X-100 solution for 5 min, followed by another three washes with PBS. BODIPY dye at a concentration of 5 μM was added to the well plate, and the scaffolds were incubated for 30 min. The scaffolds were then washed three times with PBS. Phalloidin staining was performed as described previously. Finally, the stained scaffolds were observed under a confocal laser microscope.

Secretion detection

This study also aimed to evaluate the capacity of ADM-containing electrospinning scaffolds to promote collagen secretion and myofibroblast differentiation by FBs. To assess collagen secretion, FBs were seeded at the same density as ADSCs on cell coverslips in a 24-well plate. ADM scaffolds with optimal biocompatibility and control PCL scaffolds were selected, with five samples per group being cut, disinfected (using the same method as the DAT group), and then co-cultured with the FBs-seeded coverslips in a well plate for 3 days. Following co-culture, the coverslips were fixed with 4% paraformaldehyde, washed with PBS, and then incubated with a primary Anti-Collagen I antibody. After washing, a secondary antibody working solution was added, followed by further incubation and washing steps. Finally, an enzyme conjugate working solution was added, and after incubation and washing, the samples were observed using laser confocal microscopy. To assess α-SMA expression, indicating myofibroblast differentiation, cells were blocked with 5% normal goat serum, incubated with a diluted primary antibody overnight at 4 °C, and then rewarmed to room temperature. After washing with TBST and TBS buffers, a secondary antibody was applied for 1 h in the dark. Nuclei were then stained with DAPI, followed by washing, and the samples were observed using laser confocal microscopy.

Surgical procedures

All animal experiments were approved by the Institutional Animal Care and Use Committee of Huazhong University of Science and Technology (approval date: December 2021). Male nude rats (7 weeks old, 150–200 g) were purchased and approved by the Animal Experiment Ethics Committee of Huazhong University of Science and Technology. The nude rats were numbered and placed in an SPF feeding room for 3 days to adapt to the environment and regain weight. The nude rats were weighed and anesthetized with sodium pentobarbital (40 mg/kg, i.p., China, Cat#10-0005) and xylazine hydrochloride (16 mg/kg, i.p., 98% purity; China; Cat# 182503). Once adequate anesthesia was achieved, the hair on their backs was removed. Next, a circular area with a diameter of 1 cm was marked at two locations on each side of the dorsal surface using gentian violet. The skin within the marked areas was then excised layer by layer using a sharp scalpel to create full-thickness cutaneous defects involving epidermis, dermis, and subcutaneous adipose tissue, without exposure of underlying muscle or bone, and sterile gauze was applied with pressure to ensure hemostasis. From left to right, from top to bottom, wounds were labeled as PCL group, blank control group, ADM/DAT group, and ADM group.

Healing and sampling analysis of nude rats

On days 0, day 3, day 7, day 10, and day 14, the wound healing process of each group was recorded. Image J software was used to quantitatively analyze the healing area and speed. At the 1 month and the 3 months, H&E and Masson staining were used for histological evaluation. Angiogenesis was identified by CD31 immunofluorescence, and the pro-inflammatory and anti-inflammatory macrophages were identified by CD86 and CD163 immunofluorescence staining.

RNA-seq and analysis of the data

Total RNA underwent mRNA enrichment using oligo-dT magnetic beads to capture polyadenylated mRNA. The purified RNA was fragmented using fragmentation buffer, followed by first-strand cDNA synthesis with random N6 primers and subsequent second-strand synthesis to generate double-stranded DNA. The resulting DNA underwent blunt-end repair with 5’-end phosphorylation and 3’-end adenylation. Bubble-shaped adapters containing a single 3’-T overhang were then ligated to the DNA fragments. Adapter-ligated products were amplified by PCR using specific primers. The PCR amplicons were heat-denatured into single-stranded DNA and circularized via bridge primers to construct single-stranded circular DNA libraries. Quality-controlled libraries were subjected to sequencing.

Quantitative reverse transcription‑polymerase chain reaction (qRT‑PCR) analysis

Total RNA was extracted from tissue samples using TRIzol reagent (Invitrogen, USA) following the manufacturer’s protocol. RNA concentration and purity were assessed using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, USA). Subsequently, 500 ng of RNA was reverse-transcribed into cDNA using the PrimeScript RT Reagent Kit (Takara, Japan). qRT-PCR was performed on a StepOnePlus Real-Time PCR System (Applied Biosystems, USA) using SYBR Premix Ex Taq II (Takara, Japan) in a 20 μL reaction system. The thermal cycling procedure consisted of an initial denaturation at 95 °C for 30 s, followed by 40 cycles of 95 °C for 5 s and 60 °C for 30 s. Melt-curve analysis was conducted to confirm amplification specificity. Relative gene expression levels were calculated using the 2⁻ΔΔCt method, with GAPDH serving as the internal control. Primer sequences used in this study are provided in Table S1.

Mechanical Testing

The mechanical properties of both electrospun scaffolds and regenerated skin tissues were evaluated using a universal testing machine (Shimadzu, Japan).

1. Tensile testing of electrospun scaffolds

Rectangular specimens (30 mm ×10 mm) were cut from each scaffold, and thickness was measured at three locations using a digital micrometer. Samples were mounted with a gauge length of 20 mm and subjected to uniaxial tension at a crosshead speed of 5 mm/min until failure. Stress–strain curves were recorded, and Young’s modulus was calculated from the linear elastic region (0–10% strain). Each group contained three independent samples (n = 3).

• (2)Tensile testing of regenerated skin

Full-thickness skin strips were harvested at 3 months post-treatment from each group (Blank, PCL, ADM, ADM/DAT), together with contralateral unwounded dorsal skin as the native control. Specimens were trimmed into standardized strips (30 mm ×5 mm), and subcutaneous tissue was carefully removed to ensure uniform thickness. Samples were mounted with a 15-mm gauge length and stretched at 5 mm/min until rupture. The Young’s modulus was determined from the initial linear slope of the stress–strain curve. All samples were measured under identical hydration and temperature conditions (n = 3 per group).

Micro-CT characterization of bilayer scaffold architecture

The three-dimensional structure of the bilayer ADM/DAT scaffold was evaluated using micro-computed tomography (μCT, Nano Voxel 1000). Samples were scanned at a voxel resolution of 1 μm under standardized acquisition parameters. Reconstructed cross-sections were used to determine the overall scaffold thickness by measuring the vertical distance between the upper and lower surfaces at three randomly selected positions.

Layer-specific porosity was quantified from μCT 3D volume data using CTAn software (Bruker). Scaffold regions corresponding to the ADM and DAT layers were manually segmented, and binarized images were generated using a global threshold. Porosity was calculated as the ratio of void volume to total scaffold volume within the selected region of interest (ROI).

Pore-size distribution analysis was performed using the sphere-fitting algorithm within the software, and representative 3D reconstructions were generated to visualize the distinct microarchitectures of the two layers.

Western blot analysis

Total protein from regenerated skin tissues was extracted under ice-cold conditions. Briefly, samples were minced and homogenized in RIPA lysis buffer (10× tissue volume) containing protease inhibitors using a pre-cooled bead mill. Homogenates were incubated on ice with intermittent vortexing, followed by centrifugation at 12,000 rpm for 10 min at 4 °C. The supernatant was collected, mixed with 5× reducing loading buffer (4:1, v/v), denatured at 100 °C for 10 min, cooled, and stored at −80 °C until analysis.

Equivalent protein amounts were subjected to SDS–PAGE using 10–12% polyacrylamide gels and electrophoresed at 75 V (stacking) and 100 V (resolving). Proteins were transferred onto methanol-activated PVDF membranes via wet transfer at 300 mA for 1.5 h. Membranes were blocked with 5% non-fat milk in TBST for 30 min at room temperature and incubated overnight at 4 °C with primary antibodies diluted in 5% milk/TBST. After three TBST washes, membranes were incubated with HRP-conjugated secondary antibodies (1:5000) for 30 min and washed again.

Protein bands were visualized using enhanced chemiluminescence (ECL) reagents and imaged using a digital chemiluminescence system. Exposure time was adjusted after preview scanning to obtain optimal signal intensity. Band densitometry was performed using AlphaView/ImageJ software, and target band intensities were normalized to internal controls.

ELISA analysis of ECM-derived bioactive molecules

Targeted ELISA assays were performed to quantify ECM-associated bioactive molecules in ADM and DAT samples. Tissue samples were homogenized and extracted according to kit recommendations. Briefly, tissue homogenates or cell lysates were prepared by centrifugation at 1,000–5,000 × g to remove debris, and supernatants were collected for analysis. All samples and reagents were equilibrated to room temperature before use. Standards were serially diluted in 1× dilution buffer to generate a seven-point standard curve. For each assay, 100 μL of standards or appropriately diluted samples were added to antibody-coated 96-well plates and incubated at 37 °C for 1 h. Wells were washed three times with 1× wash buffer, followed by incubation with 100 μL of biotin-conjugated detection antibody for 1 h at 37 °C. After washing, 100 μL of streptavidin-HRP working solution was added and incubated for 30 min at 37 °C. Wells were washed again, and 50 μL each of chromogenic solutions A and B was added sequentially. After 15 min of color development at 37 °C, the reaction was stopped with 50 μL of stop solution. Absorbance was measured at 450 nm using a microplate reader. Concentrations of IGF-1, VEGF, FN1, and TGF-β1 were calculated from standard curves and normalized to sample input. All measurements were performed in triplicate, and results from three independent ADM and DAT preparations were included in the analysis.

Protein mass spectrometry

Protein samples were reduced, alkylated, and digested with Trypsin (1:50, w/w) at 37 °C for 20 h. The resulting peptides were desalted, lyophilized, and reconstituted in 0.1% formic acid (FA) for storage at − 20 °C prior to analysis. The mobile phases consisted of 0.1% FA in water (Solution A) and 0.1% FA in 84% acetonitrile (Solution B). After equilibrating the analytical column with 95% Solution A, samples were loaded onto a trap column via autosampler. Mass spectrometric data were acquired in data-dependent acquisition (DDA) mode, with each full MS scan (m/z 350–1500) followed by MS/MS fragmentation of the top 20 precursor ions. Raw files were processed using Proteome Discoverer 1.4 software for database searching, yielding definitive protein identifications.

Statistical analysis

All quantitative data were expressed as mean ± standard deviation (SD). For comparisons between two groups, a two-tailed unpaired Student’s t-test was applied. For experiments involving more than two groups, one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test was performed to determine statistical significance.

For in vitro assays (cell proliferation, phalloidin fluorescence intensity, adipogenic quantification, collagen-I and α-SMA expression), each experiment was conducted with at least three independent replicates (n ≥ 3). For in vivo wound-healing analysis, wound area measurements at predefined time points (days 0, 3, 7, 10, and 14) were quantified using ImageJ based on standardized region-of-interest selection. Histological analyses (H&E and Masson) and immunofluorescence evaluations (CD31, CD86, CD163) were quantified across five random fields per section.

Western blot band intensities were normalized to internal reference proteins and quantified using ImageJ/AlphaView. Transcriptomic data were analyzed using standard bioinformatics pipelines, with differentially expressed genes identified using |log₂ Fold change| ≥ 1 and adjusted P < 0.05 as significance thresholds.

All statistical analyses were performed using SPSS 21.0 and GraphPad Prism 9.0. A P value < 0.05 was considered statistically significant for all tests.

Results

Structural and biochemical properties of DAT and ADM

Following decellularization, the DAT and ADM appeared as pale white and highly hydrated matrices. Scanning electron microscopy (SEM) imaging of freeze-dried ADM and DAT samples revealed porous fibrous microarchitectures characteristic of the native ECM, suggesting minimal microstructural disruption during production processing (Fig. 1A). The effective removal of highly immunogenic nucleated components was confirmed by H&E and 4,6-diamidino-2-phenylindole (DAPI) staining, while Masson’s trichrome staining demonstrated well-preserved collagen and reticular fibers in both materials. Sirius red staining indicated retention of type I collagen fibers in the decellularized skin and adipose tissues. Oil Red O staining revealed abundant lipid droplets in native adipose tissue, whereas no significant residual lipid droplets were observed in DAT post-decellularization (Fig. 1B). Quantitative DNA analysis showed significantly reduced residual DNA content in DAT (28.48 ± 3.35 ng) compared to native adipose tissue (418.56 ± 5.54 ng) (Fig. 1C). Similarly, ADM exhibited markedly lower DNA levels (32.07 ± 2.13 ng) than untreated dermal tissue (390.22 ± 22.71 ng) (Fig. 1D), meeting the established decellularization criterion (< 50 ng dsDNA/mg dry weight) [26]. Proteomic analysis identified collagen, hemoglobin, and albumin as predominant proteins in DAT, along with moderate levels of elastin, keratin, and fatty acid-binding proteins (Table 1). In ADM, collagen and keratin constituted major components, supplemented by histones and elastin (Table 2). In addition to the proteomic profiling dominated by structural ECM proteins, we quantified low-abundance ECM-derived bioactive molecules using targeted ELISA. IGF-1, VEGF, FN1, and TGF-β1 were selected as representative adipogenic, angiogenic, adhesion-related, and matrix-regulatory factors. Across three donors, DAT contained significantly higher IGF-1 (26–31 ng/mL) and VEGF (480–550 pg/mL), whereas ADM showed enriched FN1 (400–540 pg/mL) and TGF-β1 (960–1090 pg/mL). These results confirm that both ADM and DAT retain tissue-specific signaling molecules after decellularization, providing biochemical cues consistent with the functional behaviors described in subsequent sections.

Fig. 1.

Morphological and decellularization efficiency analysis of adipose and dermis. (A) Macrostructural and microstructural characterization of adipose, dermis, DAT, and ADM. Scale bars: General/Hydrated/Dehydrated views=1mm, SEM 500X= 20 μm, SEM 10000X=1 μm. (B). Histopathological evaluation of native and decellularized tissues. From left to right: H&E, DAPI, Masson, Sirius and Oil Red O were used to evaluate adipose, DAT, dermis and ADM’s characterization. Scale bars: DAPI = 1 mm, others = 50 μm. (C) DNA quantification in adipose versus DAT (n = 3). (D) DNA quantification in dermis versus ADM (n = 3). (E–H) ELISA-based quantification of representative ECM-derived bioactive molecules in ADM and DAT: (E) VEGF, (F) TGF-β1, (G) FN1, and (H) IGF-1. All data are shown as mean±SD. Statistical significance was determined using unpaired two-tailed t-tests. *P < 0.05, **P < 0.01, ***P < 0.001, ***P < 0.0001

Table 1.

Protein composition of DAT ranked by abundance

Table 2.

Protein composition of ADM ranked by abundance

Structural and functional characterization of conjugate electrospun scaffolds

Electrospun scaffolds incorporating DAT (5–20%) or ADM (5–20%) with polycaprolactone (PCL) were systematically characterized. Macroscopic observation showed white fibrous structures in DAT Co-ES scaffolds (Fig. 2A). The fiber diameter distribution graph exhibited two distinct populations: thicker fibers (1200–1600 nm) and thinner fibers (400–800 nm), hypothesized to correspond to DAT fibers and PCL fibers, respectively (Fig. 2C). Hydrophilicity progressively increased with increasing DAT content (Fig. 2E, G), and was accompanied by an enhanced Young’s modulus, suggesting greater tensile strength (Fig. 2H). Similarly, in ADM/PCL composite electrospun scaffolds, two fiber populations were hypothesized: one corresponding to ADM fibers and another to PCL fibers (Fig. 2B, D). Contact angle measurements indicated hydrophilicity enhancement in ADM blends, reaching approximately 90° at 20% ADM (Fig. 2F, I). Mechanical testing showed that ascending Young’s moduli positively correlated with ADM concentration (Fig. 2J). A bilayered skin construct was successfully fabricated through two steps of conjugate electrospinning using different bioinks, featuring an ADM--containing upper layer and DAT-containing lower layer. Macroscopic imaging confirmed an intact 3D architecture, while confocal microscopy visualization using FITC (DAT layer) and Rhodamine B (ADM layer) fluorescent markers validated the bilayer construct as we designed (Fig. 2K). To complement the fiber-level morphology, μCT quantification was performed. The bilayer scaffold exhibited an overall thickness of 1.158 ± 0.071 mm (n = 3). Layer-specific porosity measured by μCT was 8.56 ± 0.50% for the ADM layer and 20.93 ± 0.14% for the DAT layer. The pore-size distribution obtained from μCT likewise reflected this bilayer gradient, with the ADM layer showing a narrow, small-pore–dominant profile (primarily 1–3 μm) and the DAT layer exhibiting a broader distribution with appreciable larger-pore fractions extending into the 10–40 μm range, consistent with the respective compact and open microarchitectures of the two layers (Figure S4), demonstrating that the scaffold reproduces the native dermis-to-adipose structural transition.

Fig. 2.

Material characterizations of electrospun scaffolds. (A) Macrographs and SEM images of scaffolds containing 5%DAT, 10%DAT, 15%DAT and 20%DAT, scale bar=10 μm. (B) Macrographs and SEM images of scaffolds containing 5%ADM, 10%ADM, 15%ADM, and 20%ADM, scale bar=10 μm. (C) Nanofiber diameter distribution of scaffolds in (A). (D) Nanofiber diameter distribution of scaffolds in (B). (E) Water contact angle measurements of scaffolds in (A). (F) Water contact angle measurements of scaffolds in (B). (G) Statistical analysis of water contact angle data from (A) (n=3). (H) Young’s modulus analysis of scaffolds in (A) (n=3). (I) Statistical analysis of water contact angle data from (B) (n=3). (J) Young’s modulus analysis of scaffolds in (B) (n=3). (K) Schematic diagram, macrographs, and fluorescence staining images of bilayer conjugated electrospun tissue-engineered skin. The red fluorescence (Rho B) represents the ADM-containing layer, while the green fluorescence (FITC) indicates the DAT-containing layer, *P < 0.05, **P < 0.01, ***P < 0.001

Evaluation of the biocompatibility and tissue-specific regenerative functions of ADM/DAT Co-ES scaffolds in vitro and in vivo

Maintaining cellular viability, morphology, and functionality within tissue-engineered skin constructs is critical for promoting skin regeneration. To evaluate the biocompatibility of DAT Co-ES scaffolds (5–20%), ADSCs viability and proliferation were assessed via Live/Dead staining and Cell Counting Kit-8 (CCK-8) assays. ADSCs exhibited accelerated proliferation rates and spindle-shaped morphology in higher DAT concentration groups after 4 and 7 days of culture (Fig. 3A). Quantitative CCK-8 analysis revealed higher cell abundance in the 15% DAT content group at day 7 compared to other groups (Fig. 3C). Phalloidin staining demonstrated cytoskeletal spreading across all groups (Fig. 3E), with the 15% DAT group exhibiting maximal spreading area and cell density. Semi-quantitative fluorescence area analysis confirmed significantly enhanced cellular expansion in the 15% DAT formulation (Fig. 3G), suggesting this concentration as optimal for subsequent functional evaluations. Parallel biocompatibility assessments were conducted for ADM Co-ES scaffolds with ADM concentrations of 5–20% using FBs. Live/Dead staining and CCK-8 assays indicated progressive morphological maturation and proliferation of FBs cultured on ADM Co-ES scaffolds over time (Fig. 3B). The 15% ADM group demonstrated peak cell number at day 7 (Fig. 3D), with phalloidin staining revealing a spindle-shaped morphology and dense cellular distribution (Fig. 3F). Fluorescence quantification confirmed maximal spreading area in the 15% ADM group (Fig. 3H), designating this formulation for downstream applications. Collectively, these findings demonstrate favorable biocompatibility of both DAT and ADM Co-ES scaffolds, with 15% (m/m) optimally preserving cellular viability and phenotypic characteristics in their respective tissue layers.

Fig. 3.

Cytocompatibility evaluation of electrospun scaffolds. (A) Live/dead staining of ADSCs cultured on 5–20% DAT scaffolds at days 1/4/7. Green fluorescence (Calcein-AM) represents viable cells, red fluorescence (PI) represents dead cells, scale bar=100 μm. (B) Live/dead staining of FBs cultured on 5–20% ADM scaffolds at days 1/4/7. Fluorescent labeling consistent with (A), scale bar = 100 μm. (C) Proliferation kinetics of ADSCs on DAT scaffolds quantified by CCK-8 assay (n=5). (D) Proliferation profile of FBs on ADM scaffolds via CCK-8 assay (n=5). (E) Cytoskeletal organization of ADSCs on DAT scaffolds visualized by phalloidin/DAPI staining. Red fluorescence (phalloidin) represents F-actin; blue fluorescence (DAPI) represents nuclei, scale bar=100 μm. (F) Morphological adaptation of FBs on ADM scaffolds through phalloidin/DAPI co-staining. Staining protocol matches panel (E), scale bar = 100 μm. (G) Semi-quantitative analysis of phalloidin fluorescence intensity reflecting cytoskeletal organization in ADSCs cultured on 5–20% DAT scaffolds at day 7. (H) Semi-quantitative analysis of phalloidin fluorescence intensity reflecting cytoskeletal organization in FBs cultured on 5–20% ADM scaffolds at day 7. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001

Following a comprehensive characterization of the three-dimensional conjugate electrospun scaffolds containing DAT and ADM components, the functional impacts on progenitor cell differentiation were evaluated. Adipogenic induction was performed on 15% DAT Co-ES scaffolds (experimental group) versus PCL controls through four induction cycles. Bodipy staining revealed progressive lipid droplet formation across cycles, with the 15% group exhibiting higher lipid droplet formation than the PCL group in all cycles (Fig. 4A). Quantitative analysis showed similar adipogenesis between groups during the first cycle, but significantly enhanced lipid accumulation (p<0.05) in the 15% DAT group during subsequent cycles (Fig. 4C), confirming DAT’s adipogenic potential. The adipogenic induction assays confirmed that DAT component in Co-ES scaffolds significantly enhanced adipose regeneration. Building upon these findings, we further investigated whether ADM Co-ES scaffolds could modulate FBs secretory activity and differentiation potential to promote dermal regeneration. Type I collagen (Col-1) immunostaining and α-smooth muscle actin (α-SMA) labeling were employed to assess extracellular matrix remodeling and myofibroblast activation. While no significant differences in Col-1 deposition or α-SMA expression were observed between groups 1 day after culture, the 15% ADM scaffolds demonstrated higher collagen deposition and greater α-SMA-positive area than PCL controls after 3 days of culture (Fig. 4B). Quantitative fluorescence intensity analysis revealed higher collagen synthesis (p<0.001) and myofibroblast density (p<0.01) in the 15% ADM group relative to both tissue culture polystyrene (TCP) and PCL controls at day 3 (Fig. 4D, E). The above findings collectively validated the distinct roles of DAT and ADM, suggesting that the bilayered structure may confer corresponding dual functionalities.

Fig. 4.

Electrospun scaffolds facilitate cell-specific functional enhancement. (A) Adipogenic induction of ADSCs cultured on 15%DAT vs. PCL over four cycles. Green (BODIPY) represents intracellular lipid droplets, blue (DAPI) represents nuclei; red (phalloidin) represents cytoskeletal F-actin, scale bar=100 μm. (B) Secretory function evaluation of FBs on TCP, PCL, and 15%ADM scaffolds at days 1/3, red represents collagen type I (Col-I), green represents α-smooth muscle actin (α-SMA), blue represents nuclei (DAPI). Scale bar=100 μm. (C) Quantitative analysis of adipogenic differentiation efficiency in PCL vs. 15%DAT groups across four induction cycles, n=5. (D) Temporal profiling of collagen-I secretion on TCP/PCL/15%ADM scaffolds. (E) Quantitative analysis of α-SMA expression dynamics reflecting myofibroblast activation, ns = not significant, *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001

ADM/DAT Co-ES skin promote wound healing and dermal-adipose regeneration: effects and mechanisms

Given the demonstrated adipogenic, collagen-secretory, and myofibroblast-differentiation capabilities of the ADM/DAT Co-ES scaffold in vitro, we further investigated its therapeutic potential through standardized in vivo assessment. Immunocompromised nude rats were selected to eliminate immune responses, with four full-thickness skin defects (1 cm diameter) surgically created on each dorsum, allocated to treatment groups: PCL, ADM, blank, and ADM/DAT composite. Six nude rats (n = 6) were used per group at each time point. Wound healing progression was longitudinally monitored via serial digital imaging at postoperative days (PODs) 0, 3, 7, 10, and 14. Subsequent computer-assisted planimetric analysis was conducted using ImageJ software to assess the healing conditions (Fig. 5A, B). Healing curves demonstrated that the ADM/DAT composite group exhibited accelerated wound closure at POD7, suggesting the most rapid healing rate, while both PCL and blank groups maintained comparable efficacy throughout the observation period, with no significant improvement observed (Fig. 5G). To systematically evaluate histological characteristics during the regeneration process, hematoxylin-eosin (H&E) staining and Masson’s trichrome staining were analyzed at 1- and 3 months post-implantation. Histological analysis demonstrated complete wound healing with dermis formation across all groups at 1 month. However, the ADM/DAT composite group exhibited organized collagen architecture in Masson-stained sections, accompanied by distinct vacuolated adipocyte structures within the hypodermal layer, a phenomenon not observed in other groups. Given the time-dependent nature of adipose regeneration, histological evaluation at 3-months revealed substantial adipocyte vacuolization exclusively in the ADM/DAT composite group, confirming successful adipose regeneration (Fig. 5C, D). Uniaxial tensile testing showed a clear mechanical recovery trend (Blank≈ PCL < ADM < ADM/DAT < native skin). Notably, the near-native modulus achieved in the ADM/DAT group corresponded well with its more organized collagen architecture and robust regeneration of subcutaneous adipose tissue (Figure S3). To assess angiogenesis, we conducted CD31 immunofluorescence staining at 1-month post-implantation and observed superior vascularization in the ADM/DAT composite group compared to other groups, showing comparable low-level fluorescence intensity (Fig. 5E, H). Quantitative analysis of CD31⁺ vessel density further confirmed significantly increased neovascularization in the ADM/DAT group compared with all other groups (Fig. 5K). Concurrent evaluation of inflammatory regulation through CD86 (pro-inflammatory M1 macrophage marker) and CD163 (anti-inflammatory M2 macrophage marker) staining revealed distinct immunomodulatory patterns. The ADM/DAT group exhibited minimal CD86 expression alongside maximal CD163 fluorescence intensity, whereas Blank and PCL groups demonstrated inverse polarization profiles (Fig. 5F, I, J). Correspondingly, quantitative assessment of CD86⁺ and CD163⁺ macrophage populations demonstrated a significantly reduced M1/M2 ratio in the ADM/DAT group, supporting its enhanced immunomodulatory capacity (Fig. 5L). Western blot analysis further confirmed enhanced M2 polarization in the ADM/DAT group, with sustained upregulation of CD206, Arg-1, TGF-β1, and IL-10 at both 1 and 3 months (Figure S2). These findings collectively suggest enhanced angiogenesis coupled with anti-inflammatory macrophage polarization are potential mechanisms contributing to the observed superior therapeutic effects of the ADM/DAT composite construct in wound regeneration.

Fig. 5.

In vivo evaluation of full-thickness skin regeneration in nude rat models. (A) Sequential wound closure documentation at days 0, 3, 7, 10 and 14. (B) Schematic reconstruction of dynamic healing trajectories across groups. (C) Histological evaluation of regenerated skin at 1 and 3 months post-implantation (H&E staining). (D) Collagen deposition analysis via Masson staining. (E) Neovascularization assessment at 1 month via CD31/DAPI co-staining, scale bar=50 μm. (F) Macrophage polarization analysis at 1 month through CD86(M1)/CD163(M2)/DAPI multiplex staining, scale bar=20 μm. (G) Wound closure rate quantification. (H) CD31⁺ vascular density at 1 month. (I) CD86⁺ M1 macrophage infiltration at 1 month. (J) CD163⁺ M2 macrophage recruitment at 1 month. (K) Quantitative analysis of CD31⁺ vessel density (vessels/mm²) in Blank, PCL, ADM, and ADM/DAT groups at 1 month post-implantation. (L) Quantitative analysis of macrophage polarization expressed as M1/M2 ratio (CD86⁺/CD163⁺) at 1 month. ns=not significant, *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001

To investigate the underlying mechanisms through which the ADM/DAT composite group facilitates dermal and adipose regeneration, we conducted sample collection and transcriptomic analysis comparing the ADM/DAT composite group with the blank group at 1- and 3-months post-implantation intervals. At 1 month, significant differences in gene expression were observed between the ADM/DAT composite group and the blank group, as demonstrated by volcano plots and bar graphs, with 1,146 genes upregulated and 665 genes downregulated (Fig. 6A, B). Gene Ontology (GO) analysis of differentially expressed genes, encompassing cellular components (CC), molecular functions (MF), and biological processes (BP), indicated that the bilayered skin construct enhanced ECM remodeling and assembly, cellular renewal, and immunomodulation (Fig. 6C, D, E). Furthermore, Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis identified alterations in ECM-receptor interaction pathways, as well as MAPK, cAMP, PI3K-Akt, and VEGF signaling pathways, which are potentially associated with accelerated cell proliferation, immune regulation, and vascularization (Fig. 6F). To explore the long-term tissue regeneration mechanisms, a transcriptomic analysis was similarly performed on samples collected at three months. Differential expression analysis identified 501 upregulated and 445 downregulated genes (Fig. 7A, B). GO analysis highlighted significant changes in adipose-related activities and metabolism, including lipase activity, lipid-binding protein activity, brown adipocyte differentiation, and tricarboxylic acid metabolism (Fig 7C, D, E). KEGG pathway analysis revealed activation of lipogenesis, adipokine-related signaling pathways, fatty acid elongation, and triglyceride synthesis pathways. Besides, key pathways such as cGMP-PKG and FoxO were implicated in adipose regeneration (Fig. 7F). To validate the sequencing results, qRT-PCR was conducted to examine the expression of VEGF, TGF-β1, C/EBPα and PPARγ in regenerated tissues. At 1 month, the ADM/DAT group showed a significant increase in TGF-β1 and VEGF expression compared with Blank (****P < 0.0001), while C/EBPα and PPARγ exhibited no significant change (ns). At 3 months, the expression pattern shifted, with C/EBPα and PPARγ markedly up-regulated relative to Blank (****P < 0.0001), whereas VEGF and TGF-β1 presented only moderate elevation or no difference (Fig. S1).

Fig. 6.

Transcriptomic Profiling of ADM/DAT vs. Blank Groups at 1 Month. (A) Volcano plot of differentially expressed genes (DEGs) between ADM/DAT and Blank groups. (B) Bar graph of upregulated/downregulated DEG counts. (C–E) Gene Ontology (GO) enrichment analysis of DEGs: (C) Cellular component. (D) Molecular function. (E) Biological process. (F) KEGG pathway enrichment map of DEGs

Fig. 7.

Transcriptomic Profiling of ADM+DAT vs. Blank Groups at 3 Month. (A) Volcano plot of differentially expressed genes (DEGs) between ADM/DAT and Blank groups. (B) Bar graph of upregulated/downregulated DEG counts. (C-E) Gene Ontology (GO) enrichment analysis of DEGs: (C) Cellular component. (D) Molecular function. (E) Biological process. (F) KEGG pathway enrichment map of DEGs

Discussion

The regenerative capacity of human skin involves coordinated interactions among its three layers: the keratinocyte-stratified epidermis, which facilitates barrier formation [27, 28], the fibroblast-populated dermis which produces structural proteins like collagen, elastin, and regulatory factors such as FGF, and the adipocyte-rich hypodermis, now recognized as not only an energy accumulator but also a multifunctional repair modulator [29–31]. The regenerative capacity of skin tissue involves coordinated interactions among these three layers [32]. Importantly, emerging evidence identifies hypodermal adipocytes as metabolic supporters through lipolysis-derived FAs, immunomodulators via macrophage polarization, and promoters of granulation tissue formation [33]. Besides, experimental models have confirmed the necessity of adipose tissue, as its ablation disrupts vascular regeneration and epithelial closure [34]. This study focused on cutaneous full-thickness defects limited to the skin and subcutaneous layers. Current tissue engineering strategies have predominantly focused on dermal reconstruction, largely neglecting adipose-mediated repair processes. Through ECM preserved bioactive molecules and conjugate electrospinning-created biomimetic nanostructure, our ADM/DAT Co-ES bilayer skin enhanced therapeutic efficacy through dermal-adipose regeneration, achieving accelerated wound healing, vascular regeneration, and M2 macrophage polarization. Western blot analysis further supported the immunoregulatory profile observed in vivo. Compared with Blank and PCL, the ADM/DAT group showed higher expression levels of CD206, Arg-1, TGF-β1 and IL-10 at both 1 and 3 months, whereas ADM alone presented an intermediate increase (Fig. S2). The persistence of these M2-associated markers indicates a stable pro-repair macrophage state rather than a transient shift, which corresponds to the reduced inflammatory response and progressive matrix remodeling seen in the regenerated tissue. This sustained polarization may contribute to the transition from early vascular formation to subsequent adipose reconstruction during late-stage healing. The mechanical recovery of regenerated skin exhibited a graded trend (Blank≈PCL < ADM < ADM/DAT < native), and this improvement closely paralleled histological remodeling. In particular, the ADM/DAT group achieved near-native stiffness, consistent with its well-aligned collagen fibers and reconstructed subcutaneous adipose tissue.

The development of multilayer tissue-engineered skin promoting dermal-adipose regeneration requires appropriate material selection, balancing bioactivity, and structural functionality. As decellularized biological scaffolds retaining collagen, growth factors, and adhesion proteins while eliminating immunogenic components, the ECM provides optimal biocompatibility and pro-regenerative signaling [35]. Its critical role in tissue regeneration, stems from enabling dynamic cell-ECM interactions that drive vascularization and immunomodulation [36]. Our experiments confirmed that the ADM/DAT Co-ES bilayer skin accelerated wound healing and promoted regeneration of dermal and adipose tissues by enhancing angiogenesis and inducing M2 macrophage polarization. Our proteomic analysis revealed type I collagen as the primary components of DAT. Previous studies have demonstrated that these collagens provide structural support for ADSCs while actively promoting their differentiation into adipogenic precursor cells and mature adipocytes, facilitating adipose tissue regeneration [37]. Proteomic characterization revealed that ADM is primarily composed of collagen, consistent with its extracellular matrix origin. Keratin was detected at relatively low abundance but may remain biologically active, potentially contributing to wound healing. Specifically, keratin provides scaffolding for epithelial cells and tissues, ensuring structural integrity under mechanical stress while facilitating cell adhesion, migration, and differentiation. Notably, keratin participates in intracellular signaling pathways, such as protecting tissues from stress and apoptosis, and wound healing processes [38].

While ECM effectively supports host tissue regeneration, inherent limitations in mechanical strength and three-dimensional structural integrity necessitate complementary approaches [39]. Although ECM alone exhibits limited mechanical robustness, our results showed that higher ECM concentrations within the composite were associated with increased tensile strength. This enhancement may be attributed to improved microstructural integration between ECM components and electrospun fibers, which facilitates load transfer and supports more cohesive fiber–matrix interactions. These findings indicate that, when incorporated into an organized fibrous scaffold, ECM can contribute to mechanical reinforcement in addition to providing biochemical cues. Synthetic polymers like PCL offer tunable physicochemical properties and enhanced durability through techniques such as electrospinning, with PCL nanofibrous scaffolds demonstrating sustained mechanical support due to slower degradation rates compared to PGA or PLGA [40–42]. Hybridization strategies combining ECM’s biological advantages with synthetic polymers’ structural reliability address these limitations, exemplified by PCL/gelatin composites optimizing both cellular compatibility and mechanical performance [43–45]. Our conjugate electrospinning technology successfully fabricated a multilayer construct mimicking native skin architecture by integrating ADM and PCL into a dermal layer as well as DAT and PCL into a subcutaneous fat layer (Fig 8). Experimental validation confirmed enhanced fibroblasts and adipose-derived stem cells proliferation, accelerated wound closure, and coordinated dermal-adipose regeneration in full-thickness defect models.

Fig. 8.

Preparation of Dermal-Adipose Bilayered Skin and its application in skin defects of nude rats

Beyond material composition, the macro-microstructural design critically influences cellular proliferation and functionality in tissue-engineered skin. Effective wound healing requires scaffolds mimicking the porous nanoarchitecture of native ECM to facilitate cell-matrix interactions. Electrospinning remains a fundamental technique for creating skin substitutes with high surface area through micro/nanofibrous matrices that enable cell adhesion and tissue regeneration [46, 47]. However, conventional electrospinning typically produces 2D thin membranes (5–10 μm thick) inadequate for volumetric tissue restoration [48]. Three-dimensional scaffolds with interconnected microporosity enhance host cell infiltration, wound integration, and granulation matrix deposition. Conjugate electrospinning overcomes dimensional constraints by generating 3D scaffolds that preserve nanofibrous architecture while providing physiologically relevant thickness and increased porosity for enhanced regeneration. Traditional hemostatic sponges demonstrate limited efficacy in cases involving deep penetrating wound hemorrhage. In contrast, novel 3D hemostatic sponges fabricated via conjugate electrospinning exhibit superior hemostatic performance. The high porosity, high surface area and hygroscopic property of conjugate electrospinning hemostatic sponges are beneficial to trigger the coagulation pathway [49]; Conjugate electrospinning can also be utilized to fabricate scaffolds with excellent mechanical properties and biocompatibility, thereby promoting bone tissue regeneration. Structural versatility, biomimetic design, and multifunctional integration of conjugate electrospinning scaffolds significantly enhance bone regeneration through optimized mechanical support, controlled bioactive release, and cellular microenvironment regulation [50]. Conjugate electrospinning creates 3D scaffolds with tiny fibers and interconnected pores, which help cells grow and nutrients move freely. These scaffolds can hold healing agents, adapt to different tissue shapes, and create ideal conditions for natural regeneration processes. PCL is known for its slow degradation under physiological conditions. In this study, the scaffold was fabricated by conjugate electrospinning of PCL with ADM and DAT, which substantially increased hydrophilicity and porosity. These features are expected to promote faster hydrolytic and enzymatic degradation compared with pure PCL. During the 3-month in vivo evaluation, partial scaffold remodeling was observed, suggesting that degradation occurred in parallel with tissue integration. The composite maintained adequate mechanical support in the early stage while allowing gradual load transfer to the regenerated tissue, ensuring long-term stability. These applications and our finding suggest that three-dimensional scaffolds produced via conjugate electrospinning hold distinct advantages in fabricating volume-equivalent materials, effectively balancing structural complexity with manufacturing reliability.

The transcriptomic analyses revealed multifaceted regenerative mechanisms of the ADM/DAT Co-ES bilayer skin. At one month, upregulated immune regulatory genes indicated microenvironment modulation through M2 macrophage polarization, evidenced by increased secretion of pro-regenerative factors (VEGF, TGF-β) that synergistically enhanced angiogenesis and ECM remodeling [51]. Simultaneously, activation of MAPK and PI3K-Akt pathways correlated with enhanced keratinocyte proliferation and survival, while VEGF pathway activation confirmed critical angiogenic stimulation. These mechanisms are consistent with physiological dermal repair processes [52, 53]. At three months, metabolic reprogramming emerged as the predominant feature, with significant upregulation of lipid biosynthesis pathways (fatty acid elongation, triglyceride synthesis) and adipogenic differentiation markers (PPARγ, C/EBPα) [54, 55]. Together with the RNA-seq findings, the qRT-PCR data—showing early upregulation of VEGF/TGF-β1 and late elevation of PPARγ/C/EBPα—support that the ADM/DAT scaffold drives a staged regenerative process characterized by initial immunoregulation/angiogenesis followed by adipogenic maturation (Figure S1). The coordinated activation of CGMP-PKG and FoxO pathways further substantiated mature adipocyte formation, indicative of physiological adipose development [56] (Fig. 7). Notably, sustained ECM receptor interaction pathway activation throughout the healing course demonstrated the bioactivity of ECM in guiding cellular adhesion/migration [57]. This temporal gene expression pattern, from initial immunomodulation and angiogenesis to subsequent adipogenesis, confirms the composite’s capacity to sequentially recreate dermal-hypodermal regenerative signaling networks through coordinated molecular network activation. Furthermore, considering future translation, donor-associated ECM variability (e.g., age/BMI/tissue site) was minimized through standardized donor selection and unified decellularization workflows, and proteomic profiling confirmed biochemical stability across batches. In addition, γ-irradiation preserved ECM bioactivity, and the implemented QC framework (DNA residual testing, lipid removal, microstructure, and cytocompatibility) aligns with GMP-compatible workflows and regulatory precedents for clinically used ADM materials.

While this study demonstrated the efficacy of the ADM/DAT Co-ES bilayer scaffold in promoting dermal–adipose regeneration, several limitations should be acknowledged. The ADM and DAT used were derived from human tissues, representing xenogeneic ECM materials. Although decellularization efficiently removed immunogenic components and nude rats were used to minimize immune interference, subtle innate immune responses cannot be entirely excluded. The absence of abnormal inflammation and the predominance of M2 macrophage polarization suggest good host tolerance, yet future large-animal studies employing autologous or allogeneic ECMs are required to further assess long-term biocompatibility. Future large-animal experiments will therefore incorporate these commercial benchmarks to enable direct structural and functional comparisons. Long-term follow-up and epidermal reconstruction studies are also needed to achieve anatomically and functionally complete skin regeneration.

Conclusions

In summary, we pioneered the fabrication of a bilayered tissue-engineered skin with structural and functional biomimicry using ECM conjugate electrospinning technology, featuring an ADM upper layer and DAT lower layer. In vitro evaluations demonstrated enhanced secretory activity and differentiation of FBs, coupled with promoted adipogenic differentiation of ADSCs. Subsequent in vivo studies confirmed the construct’s robust capacity for coordinated dermal and adipose regeneration, validated through transcriptomic analyses, which revealed the modulation of immune responses, angiogenesis, ECM remodeling, and lipid metabolism. This development of a stratified skin equivalent establishes a foundational platform for engineering anatomically precise, multifunctional tissue scaffolds that recapitulate native tissue layering for complex regenerative applications.

Author contributions

Zhuolun Hao: Writing – original draft, Validation, Software, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Yaqi Guo: Visualization, Validation, Resources, Methodology, Investigation, Formal analysis. Junjin Jie: Software, Resources, Investigation, Formal analysis. Wenbin Jiang: Software, Resources, Investigation, Formal analysis. Zhenxing Wang: Writing – review & editing, Supervision. Jiaming Sun: Writing – review & editing, Supervision. Nengqiang Guo: Writing – review & editing, Writing – original draft, Supervision, Project administration, Funding acquisition, Conceptualization. Muran Zhou: Writing – review & editing, Supervision, Resources, Project administration, Funding acquisition, Conceptualization.

Funding

This work was financially supported through grants from the National Natural Science Foundation of China (Grant No. 82102351), Hubei Provincial Natural Science Foundation (Grant No. 2019CFB561).

Data availability

The datasets generated and analyzed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

This study was approved by the Animal Care and Use Committee of Huazhong University of Science and Technology (HUST). The experiments were performed in compliance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. For the experiments involving human tissue samples, written informed consents were provided by participants and the protocol was approved by the Ethics Committee of HUST. Copies of the ethical approval documents have been uploaded.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.