Bioactive eggshell membrane-integrated nanofiber dressing with mesoporous polydopamine-mediated sustained dexamethasone delivery for enhanced wound regeneration
Lu Han, Hao Feng, Zhengchao Yuan, Muhammad Shafiq, Shuqi Lou, Mohamed EL-Newehy, Meera Moydeen Abdulhameed, Yan Xiong, Xiaojing Zhao, Xiumei Mo, Jiafei Chen
TL;DR
A new wound dressing combines eggshell membrane and nanofibers to reduce inflammation and speed up healing by delivering dexamethasone over time.
Contribution
A biodegradable nanofiber dressing integrating eggshell membrane and mesoporous polydopamine for sustained dexamethasone delivery is developed.
Findings
The dressing promotes fibroblast and endothelial cell migration and reduces oxidative stress in vitro.
In rats, the dressing accelerates wound closure and collagen deposition while reducing inflammation markers.
Transcriptomic analysis shows activation of antioxidant and extracellular matrix remodeling pathways.
Abstract
Wound healing is often impeded by excessive inflammation and oxidative stress, necessitating multifunctional dressings with therapeutic and regenerative properties. Here, a biodegradable nanofiber dressing (PE@MD) composed of poly(L-lactide-co-ε-caprolactone) (PLCL), water-soluble eggshell membrane (ESM), and dexamethasone (DEX)-loaded mesoporous polydopamine nanoparticles (MPDA) was developed via electrospinning technology. Proteomic and metabolomic analyses revealed that ESM contains abundant proteins and metabolites associated with cytoskeletal organization, antioxidation, and modulation of inflammation, providing intrinsic bioactivity to the composite. The incorporation of MPDA enabled sustained and controlled DEX release while enhancing the ROS scavenging capacity. The optimized PE@MD nanofibers exhibited good flexibility, biocompatibility, and degradation properties. In vitro, the…
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Figure 11- —Ongoing Research Funding program
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TopicsWound Healing and Treatments · Electrospun Nanofibers in Biomedical Applications · Tissue Engineering and Regenerative Medicine
Introduction
Acute or chronic wounds caused by persistent infection, burns, or scalding often exhibit delayed healing and are associated with multiple clinical complications [1] Approximately 1–2% of the global population suffers from acute or chronic wounds, which result in a healthcare burden amounting to tens of billions of dollars annually [2, 3]. Wound healing is a complex process comprising overlapping phases, including hemostasis (coagulation), inflammation, cell proliferation (granulation and angiogenesis), and tissue remodeling (maturation) [4]. Excessive accumulation of reactive oxygen species (ROS) during the inflammation phase, together with persistent production of pro-inflammatory factors such as Tumor Necrosis Factor alpha (TNF-α) and Interleukin-6 (IL-6), may induce oxidative stress, thereby delaying wound healing [5–7]. Such pathological conditions hinder the typical healing cascade, highlighting the need for advanced wound dressings that can actively modulate the wound microenvironment. Therefore, it is crucial to design multifunctional dressings with anti-inflammatory, antioxidant, and tissue-regenerative properties [8].
Electrospinning has attracted extensive attention due to its capacity to generate fibrous networks with nanoscale architecture, high porosity, and morphological features reminiscent of the extracellular matrix (ECM) [9]. Poly(L-lactide-co-ɛ-caprolactone) (PLCL) is a mechano-elastic and biodegradable polymer, which is extensively used as a scaffold material for regenerative medicine and tissue engineering (TE) [10]. Moreover, the mechanical properties, architecture, and degradability of PLCL copolymers can be tuned by varying individual monomers’ molecular weight and composition [11, 12]. Despite these advantages, conventional PLCL-based wound dressings are largely passive, lacking the capacity to regulate the wound microenvironment or suppress excessive inflammation actively, and their limited functionality hinders broader clinical translation [13]. To address these limitations, natural polymers such as collagen and chitosan have been incorporated to enhance cellular affinity. However, challenges persist, including high raw material costs, suboptimal mechanical stability, and the inability to provide antioxidant and anti-inflammatory effects concurrently. Moreover, most current approaches rely on inorganic additives (e.g., silver nanoparticles) to impart antibacterial activity, while the potential of agricultural by-products as sustainable bioactive resources has been largely underexplored.
Eggshell Membrane (ESM), a by-product of the egg processing industry, has emerged as a focal study point in biomaterials [5]. ESM is composed of 80–85% proteins, including collagen type 1 (Col I) and elastin, glycosaminoglycans (GAGs), and bioactive molecules such as natural antioxidants (e.g., squalene, coenzyme Q10, etc.) [14]. Besides, ESM possesses a porous fibrous structure and manifests biocompatibility, biodegradability, and mechanical robustness [15, 16]. Our group and others have explored the use of ESM for wound healing owing to its ability to guide cell migration, an ability guided by its porous structure and inherent bioactivity, coupled with its antioxidative properties [17, 18]. Nonetheless, ESM faces several challenges, including slow degradability, risk of inducing inflammatory responses, and the absence of an optimal application form, which collectively limit its translational potential in tissue engineering.
Mesoporous polydopamine nanoparticles (MPDA NPs) have recently emerged as an up-and-coming platform for advanced wound healing applications, owing to their unique multifunctional capabilities [19]. Derived from the self-polymerization of dopamine, MPDA combines the inherent advantages of polydopamine-including excellent biocompatibility, strong tissue adhesion, and intrinsic antioxidant activity-with the structural benefits of mesoporous materials [20]. The mesoporous architecture provides a high specific surface area and well-defined pores, enabling efficient loading and sustained release of therapeutic agents such as antimicrobial peptides, growth factors, and anti-inflammatory drugs, with drug loading capacities superior to those of non-porous polydopamine nanoparticles [21, 22]. We incorporated dexamethasone (DEX) into fibrous dressings owing to its potent anti-inflammatory activity and its ability to suppress pro-inflammatory cytokines and chemokines (e.g., IL-6, TNF-α) via inhibition of the Nuclear Factor kappa-light-chain-enhancer of activated B cells pathway (NF-κB pathway) during wound healing [23]. In addition, DEX exerts effects through glucocorticoid receptor-mediated transcriptional regulation [24]. However, direct application of DEX often results in rapid release at the wound site, potentially impairing tissue repair and increasing infection risk. At the same time, systemic exposure—particularly with long-term or high-dose use—may lead to side effects such as adrenal suppression and hyperglycemia [25]. On the other hand, encapsulation of DEX into MPDA NPs may enable sustained and controlled release of DEX, thereby prolonging its anti-inflammatory effect while mitigating potential side effects [26, 27].
Fig. 1. An overview of experimental design. (a) The preparation of MPDA, water-soluble ESM, and PE@MD dressings. (b) Illustration of the potential mechanism of PE@MD dressings for wound healing
This study aimed to fabricate multifunctional nanofiber dressings (PE@MD) by incorporating ESM and DEX-loaded MPDA NPs (MD) into PLCL fibers (Fig. 1a) with a focus on valorization of waste-derived resources, rational multifunctional material design, and multi-omics-driven mechanistic exploration, aiming to provide a comprehensive strategy for wound healing. The active components and biological functions of water-soluble ESM were delineated through proteomics and metabolomics, integrated with network pharmacology. The dressings were systematically evaluated via in vitro cytocompatibility, hemocompatibility, and cell migration assays. Moreover, whole transcriptome sequencing of L929 fibroblasts treated with PE@MD dressings was performed to elucidate the underlying wound healing mechanisms. Finally, a rat full-thickness excisional wound model was employed to evaluate in vivo skin regeneration capacity (Fig. 1b).
Experimental sections
Materials
Fresh hen eggs were obtained locally (Shanghai, China). DEX was purchased from Sahn Chemical Technology Co., Ltd (Shanghai, China). Dopamine hydrochloride (PDA), 1,3,5-trimethylbenzene (TMB), and polyether (Pluronic F127) were purchased from Aladdin Reagent Co., Ltd (Shanghai, China). Poly (L-lactide-co-ε-caprolactone) (PLCL, LA: CL = 50:50, inherent viscosity 2.9 dL/g) was purchased from Shenzhen Match Biomaterials Co., Ltd. Gelatin (type B) from porcine was purchased from Sigma-Aldrich Corporation (St. Louis, MO, USA). L929 fibroblasts, human umbilical vein endothelial cells (HUVECs), and RAW 264.7 macrophages were obtained from the Shanghai Institute of Biochemistry and Cell Biology, Chinese Academy of Sciences (Shanghai, China).
Preparation of water-soluble ESM
To reduce variability, eggs from the same breed and similar laying period were used, and all ESM samples were processed following a standardized protocol. Eggshells were soaked in water at 37 °C for 10 min, rinsed, treated with 0.5% (w/v) sodium bicarbonate solution for 15 min, washed with deionized water (×3), and dried at 40 °C for 2 h. The dried eggshells were then treated with acetic acid for 24 h, with the solution refreshed every 8 h, to remove the calcified components. Eggshell membranes were separated, washed with phosphate-buffered saline (PBS, ×3), crushed, and dissolved in 1.5 M sodium sulfide (Na₂S) at 55 °C for 4 h. The solution was dialyzed (MWCO 3,500 Da) against deionized water for 72 h to remove residual salts and Na₂S. The dialyzed soluble ESM solution was then analyzed by inductively coupled plasma mass spectrometry (ICP-MS) to confirm the effective removal of residual sulfide ions. Finally, the dialyzed solution was pre-cooled at -80 °C and lyophilized for 48 h to obtain ESM powder. Compared with traditional manual peeling, this chemical-assisted extraction method allows for more complete and uniform separation of the membrane from the shell, minimizes mechanical damage, and improves reproducibility and purity. Detailed protocols are provided in the Supplementary Information.
Compositional analysis of water-soluble ESM
The compositional analysis of water-soluble ESM was performed using proteomic, untargeted metabolomics, and network pharmacology. Detailed protocols are provided in the Supplementary Information.
Preparation of DEX-Loaded MPDA (MD) nanoparticles
Mesoporous polydopamine (MPDA) nanoparticles were synthesized using the triblock copolymer Pluronic F127 and π-electron-rich 1,3,5-trimethylbenzene (TMB) as organic templates. The particles were formed through organic self-assembly and π–π stacking interactions at the water/TMB interface, during which emulsion droplets were oriented for co-assembly. Dexamethasone (DEX) was subsequently loaded into the MPDA nanoparticles. Detailed protocols are provided in the Supplementary Information.
Preparation of dressings
A representative procedure for scaffold fabrication was as follows: 10 mg DEX-loaded MPDA nanoparticles (MD) were dispersed in 10 mL hexafluoroisopropanol (HFIP) using an ultrasonic bath (Fisher Scientific, USA). Subsequently, 0.9 g PLCL was added to the suspension and stirred for 5 h. Finally, 0.1 g ESM powder was introduced into the mixture and stirred overnight to obtain the electrospinning solution. PE@MD nanofibers were fabricated by electrospinning (SS-3556 H, Leye Technology, China. The electrospinning parameters were as follows: needle gauge, 20 G, applied voltage, 13 kV, flow rate, 2 mL/h, and spinneret-to-collector distance, 13 cm. PG (PLCL/Gelatin) was selected as a clinically established fibrous reference, while the introduction of ESM provides a low-cost, waste-derived, and sustainable alternative with comparable or improved biological performance, thereby enabling direct assessment of the functional advantages conferred by ESM relative to conventional gelatin-based dressings. Accordingly, for comparison, control scaffolds were prepared under identical electrospinning conditions, including the PG group (PLCL: Gelatin = 9:1, w/w) and the PE group (PLCL: ESM = 9:1, w/w). Detailed protocols are provided in the Supplementary Information.
Physicochemical analysis
The morphology and microstructure of scaffolds were examined by scanning electron microscopy (SEM) and transmission electron microscopy (TEM). Functional groups were analyzed using Fourier-transform infrared spectroscopy (FTIR). Mechanical performance was assessed by tensile testing. Surface wettability was measured by water contact angle (WCA). The release kinetics of DEX from nanofibers were determined. Detailed experimental conditions are provided in the Supplementary Information.
Hemocompatibility
Hemolysis assays were performed using fresh blood collected from New Zealand rabbits and anticoagulated with sodium citrate. Red blood cells (RBCs) were isolated by centrifugation at 3,000 rpm for 15 min, washed three times with saline, and diluted to 2% suspension. Scaffold samples (10 mm × 10 mm) were incubated at 37 ℃ for 30 min in 2 mL centrifuge tubes containing 200 µL of diluted RBC suspension, followed by incubation at 37 ℃ for 1 h. After centrifugation (2,500 rpm, 5 min), the absorbance of the supernatant was measured at 540 nm using a microplate reader (Multiskan MK3, Thermo Fisher Scientific, USA). Physiological saline (0.9%) and deionized water were negative and positive controls, respectively. The hemolysis rate was calculated according to Eq. (1):
\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$\:Hemolysis\:rate\left(\%\right)=\left[\frac{\left({A}_{s}-{A}_{n}\right)}{\left({A}_{p}-{A}_{n}\right)}\right]\times100\%$$\end{document}where As, An, and Ap represent the absorbance of the sample, negative Control, and positive Control, respectively.
Cytocompatibility
Scaffolds (diameter 10 mm) from PG, PE, and PE@MD groups were sterilized with 75% ethanol for 1 h, followed by UV irradiation for 12 h on each side. L929 fibroblasts and HUVECs were seeded on scaffolds in 48-well plates at a density of 2 × 10⁴ cells/well and cultured at 37 °C, 95% humidity, and 5% CO₂ for 1, 3, and 5 days. Medium was refreshed every other day. Cell proliferation was assessed using the Cell Counting Kit-8 (CCK-8, Beyotime Biotech, Shanghai, China). Live/Dead staining was performed with calcein-AM and propidium iodide (PI), and cells were imaged with a fluorescence microscope (DM300, Leica Microsystems, Germany).
Cell migration
The wound scratch assay and Transwell migration assay were employed as well-established in vitro models to evaluate cell migratory behavior driven primarily by soluble chemotactic cues [28, 29]. In the present study, these assays were specifically designed to probe the chemotactic effects of bioactive components released from the scaffolds, including MPDA-mediated dexamethasone and ESM-derived soluble factors. Therefore, scaffold extracts were used in order to decouple compositional bioactivity from topographical guidance effects associated with the fibrous architecture.
For scratch assays, L929 fibroblasts were seeded in 24-well plates (4 × 10⁴ cells/well) and cultured until 75% confluence. A scratch was introduced with a 200 µL sterile pipette tip, and debris was removed by PBS washing. Cells were cultured with serum-free medium (Control) or scaffold extracts from PG, PE, and PE@MD membranes. Images were captured at 0 h and 24 h. Migration rate was calculated by Eq. (2):
\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$\:Migration\:rate\left(\%\right)=\left[\left({A}_{0}-{A}_{t}\right)/{A}_{0}\right]\times100\%$$\end{document}where A0 and At are the scratch areas at 0 h and 24 h, respectively.
For transwell migration assays, cells were seeded in the upper chamber using a serum-free medium, while the lower chamber contained a serum-supplemented medium. After 12 h incubation, non-migrated cells were removed from the upper side, and migrated cells were fixed with 4% paraformaldehyde and stained with crystal violet. Migrated cell morphology was observed using a fluorescence microscope (DM300i, Leica Microsystems, Germany), and cell numbers were quantified.
Transcriptome sequencing
Transcriptome sequencing of L929 fibroblasts treated with different scaffolds was performed by OBiO Technology Corp., Ltd. (Shanghai, China). Library preparation, sequencing, and bioinformatics analyses were conducted according to standard protocols. Detailed methods are provided in the Supplementary Information.
Anti-inflammatory and antioxidant properties
Scaffold extracts (6 cm²/mL) were prepared by incubating membranes in serum-free medium at 37 °C for 24 h, followed by filtration through a 0.22 μm filter. RAW264.7 macrophages (1 × 10⁶ cells/well) were pretreated with scaffold extracts for 4 h and subsequently stimulated with LPS (100 ng/mL, 24 h, light-protected). Cells were stained with fluorescent anti-CD86 and anti-CD206 antibodies and analyzed by flow cytometry (BD Biosciences). Data were processed with FlowJo v11. For ROS quantification, cells were incubated with DCFH-DA for 30 min, washed, and analyzed by flow cytometry. For gene expression analysis, total RNA was extracted from treated cells, reverse-transcribed into cDNA, and subjected to quantitative real-time PCR (qRT-PCR). Relative expression levels of target genes were calculated using the 2^−ΔΔCt^ method with Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as the internal reference (n = 3). Primer sequences and detailed methods are provided in the Supplementary Information.
Animal experiments
All animal procedures were approved by the Welfare and Ethics Committee of the Department of Laboratory Animal Science, Donghua University (No. DWSY202505090156). A full-thickness excisional wound model was established in Sprague–Dawley (SD) rats. Rats were anesthetized by intraperitoneal injection of sodium pentobarbital (45 mg/kg), and dorsal skin was sterilized with iodophor. Under aseptic conditions, circular full-thickness wounds (10 mm in diameter) were created using a biopsy punch. For quantitative analyses, each wound was considered as an independent experimental unit (n = 3). Animals were randomly assigned to different treatment groups prior to surgery. Wounds were treated with gauze (Control), a commercial dressing (P-Control), PG, PE, or PE@MD. All dressings were standardized in size and thickness and replaced every three days. Post-operative analgesia was provided following full-thickness skin excision in accordance with institutional animal care guidelines [30]. Wounds were photographed at days 0, 3, 7, 10, and 14. Tissue samples from day 7 and day 14 were collected for histological evaluation (Hematoxylin & Eosin, Masson’s trichrome and Picrosirius Red staining) and immunohistochemical analysis for inflammatory markers (COX-2, IL-1β, MPO, and NF-κB p65). For histological and immunohistochemical quantification, three randomly selected, non-overlapping representative fields were analyzed per section. ImageJ was used for threshold-based segmentation with identical parameters applied across all groups. Quantification was performed in a blinded manner by two independent investigators. Wound closure rate was calculated according to Eq. (3):
\documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$\:Wound\:closure\:rate\left(\%\right)=\left(1-{A}_{t}/{A}_{0}\right)\times\:100\%$$\end{document}where A0 and At represent the wound area at day 0 and at the indicated time point, respectively.
Statistical analysis
All experiments were conducted with at least three independent samples. Quantitative data are presented as mean ± standard deviation (SD). Statistical analysis was performed using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test. Statistical significance was defined as *p < 0.05, **p < 0.01, and ***p < 0.001.
Results
Preparation and compositional analysis
Water-soluble eggshell membrane (ESM) powder was obtained through acetic acid treatment, Na₂S-mediated disulfide bond cleavage, dialysis, and lyophilization (Fig. 2a). In this process, Na₂S acts as a thiol-based reducing agent that selectively reduces disulfide crosslinks to facilitate ESM solubilization, without directly hydrolyzing peptide bonds or inducing nonspecific degradation of the polypeptide backbone [31–34]. ICP-MS analysis revealed that the \documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$\:{\mathrm{S}}^{2-}$$\end{document} concentration in the dialyzed ESM solution was 0.064 ± 0.005 mg/L, indicating that residual \documentclass[12pt]{minimal} \usepackage{amsmath} \usepackage{wasysym} \usepackage{amsfonts} \usepackage{amssymb} \usepackage{amsbsy} \usepackage{mathrsfs} \usepackage{upgreek} \setlength{\oddsidemargin}{-69pt} \begin{document}$$\:{\mathrm{S}}^{2-}$$\end{document} were effectively removed [35]. SEM analysis revealed irregular flaky particles with interconnected pores (Fig. 2b), a morphology that provides a favorable architecture for the encapsulation and sustained release of bioactive molecules [15].
Fig. 2. Characterization of Lyophilized Eggshell Membrane. (a) Schematic diagram of the preparation process for freeze-dried egg membrane. (b) Macroscopic images and scanning electron microscopy (SEM) images during the preparation of the egg membrane. (c) The top 10 protein structures identified via proteomic analysis. pLDDT is a per-residue measure of local confidence
Proteomic profiling identified 72 high-confidence proteins (FDR < 1%), including 21 with resolved three-dimensional structures in the PDB database. The top ten abundant proteins (Fig. 2c, Table S2), such as lysozyme (LYZ), ovostatin (SERPINB14), GAPDH, annexin A1 (ANXA1), actins (ACTB, ACTG1, ACTC1, ACTA1), and tubulins (TUBB4B, TUBA1A), represent the functional core of the ESM proteome. These proteins are implicated in cytoskeletal remodeling, inflammation regulation, oxidative stress defense, and apoptosis—processes tightly coupled to wound healing. For instance, actins and tubulins provide structural support for cell migration and granulation tissue formation; LYZ exhibits antibacterial activity; ANXA1 regulates inflammation resolution; cathepsins D and L contribute to immune responses; while antioxidant enzymes such as catalase (CAT) and peroxiredoxin-1 (PRDX1) mitigate oxidative stress.
Functional enrichment analyses further supported these findings. KEGG pathways (Fig. S1a, Table S3) were significantly enriched in apoptosis, necroptosis, phagosome formation, cell junction assembly, focal adhesion, and autophagy, indicating that ESM proteins may participate in the resolution of inflammation, barrier reconstruction, and cellular homeostasis. GO enrichment analysis (Fig. S1b) showed convergence from three perspectives: Molecular Function (ATP binding, hydrolase activity, actin filament binding), Cellular Component (extracellular region, keratin and actin filaments, focal adhesions), and Biological Process (keratinization, filament organization, regulation of apoptosis, microtubule-associated processes). These results demonstrate that water-soluble ESM provides a reservoir of structural and bioactive proteins directly relevant to multiple stages of wound healing.
Fig. 3. Untargeted metabolomics and network pharmacology analysis of freeze-dried ESM powder. (a) Chemical structures of the top 10 metabolites in ESM analyzed by non-targeted metabolomics. Network pharmacology analysis of the intersection between top 10 active components in eggshell membrane and wound healing targets: (b) Venn diagram, (c) GO enrichment analysis diagram, (d) KEGG pathway analysis
To characterize the small-molecule composition of water-soluble ESM, non-targeted LC-MS/MS metabolomics was performed. A total of 146 metabolites were identified, with the top 10 including fatty acids (palmitic acid, arachidonic acid, linoleic acid, myristic acid), amino acids and derivatives (5-aminovaleric acid, L-proline, L-arginine), carbohydrates (sucrose), and tricarboxylic acid (TCA) intermediates (citric acid, oxoglutaric acid) (Fig. 3a). These metabolites are primarily composed of long-chain fatty acids and bioactive amines, which may contribute to wound healing through roles in energy metabolism, membrane construction, inflammation regulation, collagen synthesis, and antioxidant defense.
All identified metabolites were further classified into 16 categories of primary metabolites (Fig. S2a) and 11 categories of secondary metabolites (Fig. S2b). Primary metabolites were dominated by fatty acids, sterols, and nitrogen-containing compounds that underpin fundamental cellular functions. In contrast, secondary metabolites—mainly benzenoids, indoles, and sterol derivatives—were associated with stress adaptation and defense responses. Together, these findings highlight the metabolic basis by which ESM may supports tissue repair.
A network pharmacology approach was employed further to assess the potential biological activities of these metabolites. Active compound–target prediction yielded 191 targets, accounting for 2.8% of the total (Fig. 3b). Among these, core nodes included EGFR, TP53, SRC, AKT1, CCND1, NFKB1, MMP9, ESR1, PTGS2, and PPARG, which formed highly connected hubs in protein–protein interaction networks (Fig. S3). These targets were functionally clustered into modules associated with inflammation regulation (NFKB1, PTGS2), cell proliferation (EGFR, CCND1, AKT1), and angiogenesis (SRC, MMP9). GO enrichment analysis linked the targets to EGFR-mediated signaling, suggesting roles in apoptosis inhibition and receptor regulation (Fig. 3d). KEGG pathway analysis revealed enrichment in VEGF signaling, MAPK signaling, and arachidonic acid metabolism, pathways potentially involved in angiogenesis, cell proliferation, and inflammatory homeostasis (Fig. 3c).
Together, these multi-omics and network pharmacology analyses provide exploratory, hypothesis-generating insights into the structural and metabolic features of water-soluble ESM, highlighting candidate pathways such as the EGFR/VEGFA–MAPK and IL6–STAT3 axes that could be further investigated in future mechanistic studies of wound healing.
To evaluate the influence of ESM content on the spinnability and functional performance of PLCL/ESM (P/E) nanofibers, P/E spinning solutions were prepared at different weight ratios (10/0, 9/1, 8/2, 7/3, 6/4, and 5/5, abbreviated as PE) (Fig. S4a).
SEM imaging revealed that fiber morphology was highly dependent on ESM proportion. The fibers exhibited uniform, continuous, and bead-free morphology at a ratio of P/E = 9/1 (w/w), whereas higher ESM contents (≥ 7/3) led to increasingly irregular and defective fiber structures, which can be attributed to the limited solubility and poor dispersion of ESM in the PLCL–HFIP system (Fig. S4b). Such structural instability at elevated ESM loadings is known to adversely affect the processability and mechanical integrity of electrospun matrices. Mechanical testing further demonstrated the composition-dependent performance of the nanofibrous membranes (Fig. S4c-f). Although pure PLCL (P/E = 10/0, w/w) exhibited the highest tensile strength and elongation, the absence of bioactive ESM limits its relevance for biofunctional wound applications. Among the ESM-containing groups, the membrane prepared at P/E = 9/1 (w/w) displayed a favorable mechanical profile, combining relatively high tensile strength (5.24 ± 0.68 MPa), high elongation at break (330.51 ± 62.69%), and a low Young’s modulus (0.033 ± 0.002 MPa), which are beneficial for maintaining mechanical robustness while accommodating dynamic skin deformation. In contrast, higher ESM contents (P/E ≥ 7/3) resulted in reduced ductility and increased stiffness, indicating compromised structural continuity and less efficient stress transfer within the fiber network, in agreement with the observed morphological deterioration. Surface wettability measurements showed that the incorporation of ESM progressively increased the hydrophilicity of the nanofibrous membranes (Fig. S4g). The water contact angle (WCA) decreased from 122.5 ± 3.02° for pure PLCL (P/E = 10/0, w/w) to 84.43 ± 0.50° at P/E = 9/1 (w/w), and further to 40.9 ± 1.51° at P/E = 5/5 (w/w), reflecting the introduction of hydrophilic functional groups from ESM on the fiber surface. While enhanced hydrophilicity is generally favorable for cell adhesion and exudate management, excessively low contact angles may compromise the barrier function and structural stability of wound dressings. On this basis, the ratio of P/E = 9/1 (w/w) was selected as optimal for subsequent experiments [36–38].
Preparation and characterization of composite nanofiber dressing
DEX was incorporated into mesoporous polydopamine nanoparticles (MPDA NPs) to endow the nanofiber membranes with anti-inflammatory activity and generate DEX-loaded MPDA (MD NPs). MPDA was synthesized by an emulsion templating method, in which dopamine polymerized around the TMB/Pluronic F127 emulsion domains under alkaline conditions. Subsequent removal of TMB and Pluronic F127 generated interconnected mesopores within the PDA framework. Nitrogen adsorption–desorption analysis of MPDA exhibited a type IV isotherm with a distinct hysteresis loop, characteristic of mesoporous materials (Fig. S5a). BET analysis revealed a specific surface area of 109.41 m²/g and a C constant of 38.9, with a high correlation coefficient (r = 0.9997), indicating reliable adsorption behavior. The corresponding pore size distribution calculated by the BJH method showed a predominant pore size within the mesoporous range (Fig. S5b), confirming the successful formation of a mesoporous structure [39, 40]. DEX was successfully encapsulated into the pores of MPDA via π–π stacking and hydrogen bonding interactions. TEM imaging revealed that both MPDA and MD displayed uniform spherical morphologies with dark contrast (Fig. 4b). Dynamic light scattering (DLS) confirmed an average hydrodynamic diameter of 193.8 ± 125.1 nm for MPDA and 215.8 ± 125.4 nm for MD (Fig. 4c). FTIR spectra further validated the incorporation of DEX, as characteristic peaks corresponding to C–H stretching, C–O stretching, and C–F bending vibrations of DEX were observed in MD, in addition to the PDA-specific bands (Fig. 4d) [41]. The DEX-loaded nanoparticles exhibited an encapsulation efficiency of 67.67 ± 1.08% and a drug loading of 40.36 ± 0.39%, further confirming efficient DEX incorporation. These results collectively confirmed the successful synthesis of MPDA and efficient DEX loading.
Figure 4a presents a schematic illustration of the fabrication of PE@MD nanofiber dressing, while TEM characterization further verified the successful incorporation of MD into PE fibers (Fig. S6). MD NPs were subsequently incorporated into PLCL/ESM (PE) nanofibers by electrospinning to generate PE@MD membranes. SEM revealed densely packed fibers with smooth and uniform morphology across all groups (PG, PE, and PE@MD), without observable nanoparticle aggregation (Fig. 4e). The average fiber diameters were comparable among PG (1253.3 ± 243.0 nm), PE (1216.5 ± 235.9 nm), and PE@MD (1209.0 ± 237.0 nm) (Fig. 4f). Capillary flow analysis (CFA) showed that pore size distribution was smallest in PG (0.95–1.95 μm) and largest in PE@MD (1.5–3.5 μm), with PE in between (1.1–3.3 μm) (Fig. 4g). The increase in pore size upon ESM addition, and further with MD incorporation, suggested that water-soluble components contributed to enhanced porosity.
Degradation studies in PBS revealed progressive mass loss over 14 days, with PE exhibiting the fastest degradation (25.94 ± 1.30%), followed by PE@MD (21.89 ± 1.24%) and PG (18.84 ± 1.37%) (Fig. 4h). The relatively high degradation rate of PE was attributed to the protein-rich ESM component. At the same time, the slight reduction upon MD incorporation suggested structural reinforcement by nanoparticles. Time-dependent SEM confirmed gradual fiber disruption in PE@MD (Fig. S7a). Thermal stability analysis further corroborated these findings: PE@MD exhibited a multistage decomposition profile with main degradation between 250 and 450 °C and distinct peaks at 340 °C and 400 °C (Fig. S7b), indicating a composite material with heterogeneous components.
Fig. 4. Characterization of nanofiber dressings. (a) Schematic illustration of the fabrication of composite fibers. (b) photographs and transmission electron microscopy (TEM) images of MPDA and MD, and (c) quantitative analysis of particle size distribution. Scale bar, 200 nm. (d) FTIR spectra of PDA, MPDA, DEX, and MD. (e) Macroscopic visual recording images and SEM images of PG, PE, and PE@MD dressings, (f) quantitative analysis of the distribution of average fiber diameter of different types of membranes, (g) quantitative analysis of pore size distribution, (h) quantitative analysis for in vitro degradation of membranes, and (i) representative stress-strain curves. Scale bar, 20 μm. (j) Percentage of DEX release over 14 days in an in vitro drug release assay. (n = 3)
In vitro drug release studies demonstrated stark differences between direct DEX-loaded fibers (PE@D) and nanoparticle-mediated release (PE@MD). PE@D exhibited a burst release of 95.37 ± 1.34% within 24 h, whereas PE@MD released only 30.69 ± 1.06% during the same period (Fig. S8a-b). By day 14, PE@MD achieved a cumulative release of 64.2 ± 2.48% (Fig. 4j), confirming that MPDA incorporation enabled prolonged and sustained DEX release.
Fig. 5. Biocompatibility of dressings in vitro. (a) Live/Dead staining of HUVECs seeded on PG, PE, and PE@MD dressings. (b) Live/Dead staining of L929 fibroblasts seeded on PG, PE, and PE@MD dressings. Scale bar, 200 μm. (c) Macroscopic photographs of hemolysis assay for the positive Control (water), negative Control (normal saline), PG, PE, and PE@MD dressings. (d) Quantitative analysis of hemolysis assay. Quantitative analysis of (e) HUVECs and (f) L929 fibroblasts using CCK-8 assay. Data are presented as mean ± SD (n = 3). Statistical analysis was performed with ANOVA with Tukey’s post hoc test, and * indicates p < 0.05, ** indicates p < 0.01, *** indicates p < 0.001
Mechanical testing showed group-dependent variations in tensile strength, elasticity, and strain. PG exhibited the highest tensile strength (Ultimate Tensile Strength, UTS 3.90 ± 0.31 MPa) but lower strain (214.6 ± 42.3%), while PE was softer and more elastic (UTS 2.72 ± 0.39 MPa; strain 313.5 ± 22.4%). Incorporation of MD reduced both UTS (1.54 ± 0.31 MPa) and strain (197.1 ± 15.4%), indicating easier deformation but lower resistance to mechanical stress (Fig. S9a–c). These properties suggest that PE-based membranes are sufficiently flexible to accommodate wound movements, while PE@MD maintains structural stability despite reduced elasticity [7].
Water contact angle (WCA) measurements revealed distinct differences in surface wettability. PG membranes were relatively hydrophilic (102.6 ± 2.3°), PE exhibited higher hydrophobicity (125.0 ± 3.1°), while PE@MD showed an intermediate value (109.7 ± 1.0°) (Fig. S10). Upon MD incorporation, this modulation of surface wettability suggested that PE@MD possessed balanced hydrophilic–hydrophobic characteristics, favorable for wound dressing applications.
Biocompatibility and biological functions in vitro
Further evaluated the in vitro biocompatibility of dressings. Hemolysis assay revealed no significant differences among groups; all membranes exhibited hemolysis ratios below 2%, within the international safety threshold of 5% (Fig. 5c–d) [42]. L929 fibroblasts and HUVECs were cultured on PG, PE, and PE@MD dressings for cytocompatibility for up to 5 days. Live/Dead staining confirmed high cell viability with minimal apoptosis and sufficient spreading across all groups (Fig. 5a–b). Consistently, CCK-8 assays demonstrated continuous proliferation of both cell types, with no significant cytotoxic effects observed (Fig. 5e–f).
Fig. 6. Biological functions of dressings in vitro. (a) Cell compatibility and anti-inflammatory properties of composite fibrous dressings in vitro. (b) Wound healing assay using L929 fibroblasts in Control, PG, PE, and PE@MD groups. (c) Transwell migration assay of L929 fibroblasts and HUVECs in Control, PG, PE, and PE@MD groups. Scale bar, 200 μm. (d) Quantitative analysis of scratch wound healing assay. Quantitative analysis of Transwell migration assay for (e) L929 fibroblasts and (f) HUVECs. Quantitative analysis of (g) relative ROS expression, (h) proportions of CD206⁺ and CD86⁺ cells. qRT-PCR analysis of inflammatory cytokine expression in RAW264.7 macrophages: (i) IL-4, (j) IL-10, (k) IL-6, and (l) TNF-α mRNA expression levels. Gene expression levels were normalized to GAPDH. Data are presented as mean ± SD (n = 3). Statistical analysis was performed with ANOVA with Tukey’s post hoc test, and * indicates p < 0.05, ** indicates p < 0.01, *** indicates p < 0.001
Figure 6a presents a schematic illustration of the in vitro cytocompatibility and anti-inflammatory properties of the composite nanofiber dressings. Cell migration capacity was further assessed using transwell and wound healing assays. In the transwell assay, the number of migrated L929 fibroblasts was 26.0 ± 8.54, 61.7 ± 4.51, 90.7 ± 16.2, and 145.7 ± 9.6 for Control, PG, PE, and PE@MD groups, respectively; for HUVECs, values were 17.7 ± 7.1, 47.3 ± 11.1, 52.7 ± 11.6, and 109.0 ± 8.2, respectively (Fig. 6c and e–f). PE@MD exhibited the highest cell migration. Similarly, wound healing assay showed superior wound closure in the PE@MD group (L929 fibroblasts: Control, 16.1 ± 9.0%; PG, 23.3 ± 5.7%; PE, 60.9 ± 3.9%; PE@MD, 82.4 ± 8.0%) (Fig. 6b and d). Enhanced migration in PE@MD may be attributed to the hydrophilic amine groups in ESM and MD, which can activate cell migration-related pathways Intracellular Reactive Oxygen Species (ROS) levels were measured under LPS induction to evaluate oxidative stress. Relative ROS levels for Control, LPS, PG, PE, and PE@MD groups were 1.00 ± 0.01, 2.22 ± 0.02, 2.03 ± 0.01, 2.02 ± 0.01, and 1.64 ± 0.02, respectively (Fig. 6g, Fig. S11b). Elevated ROS levels in LPS, PG, PE, and PE@MD confirmed successful induction of inflammation. Compared with LPS, all dressings reduced ROS production, with PE@MD showing the most pronounced reduction. These effects can be attributed to ESM and DEX’s combined antioxidant and anti-inflammatory actions. Specifically, ESM contains bioactive components such as glycosaminoglycans (GAGs) and antimicrobial peptides [43]. PDA scavenges ROS directly, while DEX suppresses inflammatory responses and further reduces ROS. Moreover, the mesoporous structure of MD enables sustained DEX release, maintaining anti-inflammatory activity and mitigating oxidative stress [19].
Flow cytometry was used to assess macrophage polarization. Compared with the LPS group, PE@MD membranes significantly increased CD206⁺ macrophages and reduced CD86⁺ macrophages (Fig. S11a). Quantitative analysis revealed a marked increase in the CD206⁺/CD86⁺ ratio in PE@MD (5.53 ± 0.53), compared with Control (1.14 ± 0.11), LPS (0.42 ± 0.03), PG (1.42 ± 0.23), and PE (2.84 ± 0.31) (Fig. 6h). These results demonstrated that PE@MD enhanced M2 macrophage polarization, indicating strong anti-inflammatory properties. Mechanistically, mesoporous MD can scavenge ROS and inhibit the NF-κB pathway, suppressing M1 polarization [44]. Sustained DEX release inhibits NF-κB while activating PPAR-γ, promoting M2 macrophages [45]. In parallel, collagen in ESM may further has been reported in prior literature to be associated with STAT6-dependent M2 polarization [46, 47].
The mRNA expression levels of the anti-inflammatory cytokines IL-4 and IL-10 and the pro-inflammatory cytokines IL-6 and TNF-α were analyzed by qRT-PCR (Fig. 6i-l). The results demonstrated that material treatment markedly modulated the inflammatory gene expression profile of macrophages. Compared with the control group, the PE and PE@MD groups significantly upregulated IL-4 and IL-10 expression, with the PE@MD group exhibiting the most pronounced increases (IL-4: 2.15 ± 0.10; IL-10: 3.82 ± 0.18), exceeding those of the PG and PE groups, indicating the strong immunoregulatory effects of ESM and MD. Meanwhile, the expression of pro-inflammatory factors IL-6 and TNF-α was significantly downregulated in the PE and PE@MD groups, particularly in the PE@MD group, which decreased to 0.26 ± 0.02 and 0.16 ± 0.003, respectively, suggesting that the composite dressing effectively suppresses inflammatory amplification. These gene expression trends further confirm that PE@MD promotes macrophage polarization toward a reparative phenotype by alleviating oxidative stress and regulating inflammatory cytokine expression, thereby establishing a more favorable immune microenvironment for wound healing [48–50].
Finally, RNA sequencing was performed to explore the regulatory mechanisms of dressings on L929 fibroblasts. All samples displayed high sequencing quality (Q30 > 95%; PE@MD Q30 = 96.81%), clean read alignment > 91%, and strong biological reproducibility (Pearson correlation > 0.95) (Table S4). PCA analysis showed distinct clustering between treatment groups (PE@MD, PE) and Control, indicating robust data quality. Compared with Control, PE@MD induced 2,206 differentially expressed genes (1,553 upregulated, 653 downregulated) (Fig. S12a–b). Clustering and volcano plots confirmed greater transcriptional changes in PE@MD vs. Control and PE@MD vs. PE groups, whereas PE vs. Control showed limited differences. Notably, aldehyde dehydrogenase family genes were strongly upregulated, suggesting enhanced antioxidant responses [51]. Tsc22d3, a key regulator in glucocorticoid signaling, was also elevated, consistent with DEX-mediated inflammation resolution (Fig. S13a, Fig. 7a) [52].
Fig. 7. Whole-transcriptome RNA sequencing results. (a) Volcano plots, (b) GO enrichment analysis of upregulated genes, (c) GO enrichment analysis of downregulated genes, and (d) KEGG pathway analysis for the comparative groups: PE vs. Control, PE@MD vs. Control, and PE@MD vs. PE
Venn analysis revealed that PE@MD substantially altered wound healing-related genes, whereas PE had minimal effects (Fig. S13b). GO enrichment indicated differential genes were mainly associated with extracellular space and DNA replication, including Pola1 and Mcm2, suggesting enhanced proliferation and ECM remodeling. KEGG analysis revealed activation of p53 signaling (e.g., Cdkn1a) and upregulation of ECM-receptor interaction pathways (e.g., Col1a1), implicating enhanced regulation of cell cycle and adhesion (Fig. 7b–d). These findings demonstrated that PE@MD enhanced fibroblast proliferation and adaptability by activating antioxidant (Aldh1a7), anti-inflammatory (Tsc22d3), and ECM remodeling pathways, elucidating molecular mechanisms underlying wound healing activity.
These transcriptomic results provide exploratory insights into the molecular responses of fibroblasts to PE@MD treatment, highlighting potential pathways for further mechanistic investigation.
Wound healing in vivo
A full-thickness excisional wound model in rats was established to evaluate the therapeutic efficacy of the dressings (Fig. 8a). Digital images of wounds were captured on days 0, 3, 7, 10, and 14, and the wound areas were quantified using ImageJ (Fig. 8b–d). By day 3, all wounds had formed scabs, indicating the initiation of the healing process. Progressive wound contraction was observed in all groups, while the PE@MD group exhibited markedly accelerated closure compared with both the Control and commercial P-Control groups. Quantitative analysis showed wound closure rates of 71.7 ± 3.7%, 70.7 ± 3.8%, 72.8 ± 4.3%, 83.1 ± 1.0%, and 98.0 ± 1.7% for the Control, P-Control, PG, PE, and PE@MD groups, respectively, on day 14 (Fig. 8e), confirming the superior wound contraction capacity of PE@MD.
To further elucidate tissue regeneration, inflammation, and extracellular matrix (ECM) remodeling during healing, wound tissues were harvested on days 7 and 14 for histological and immunohistochemical analyses, including hematoxylin–eosin (H&E), Masson’s trichrome (MT), and Sirius Red staining (Fig. 9a–b, Fig. S14a-b). Immunohistochemistry for COX-2, IL-1β, MPO, and NF-κB p65 was performed on day 14 to probe the underlying anti-inflammatory mechanisms (Fig. 10).
Fig. 8. The ability of dressings to induce skin regeneration in vivo. (a) Evaluation of dressings for wound healing using a full-thickness excisional defect model in SD rats. (b) Schematic diagram of wound treatment over 14 days. (c) Wound change model (Image J analysis mapping). Quantitative analysis of (d) the wound area and (e) the wound closure rate for the five groups of dressings: Control, P-Control, PG, PE, and PE@MD. Data are presented as mean ± SD (n = 3). Statistical analysis was performed with ANOVA with Tukey’s post hoc test, and * indicates p < 0.05, ** indicates p < 0.01, *** indicates p < 0.001
Fig. 9. Results of H&E staining and MT staining. (a) H&E and (b) Masson’s trichrome (MT) stained wound sections on day 7 and day 14. Scale bar, 200 μm. Quantitative analysis of (c) wound length, (d) number of inflammatory cells, (e) granulation tissue thickness, (f) re-epithelialization rate, and (g) percentage of collagen deposition area for the five groups of dressings on day 14. Data are presented as mean ± SD (n = 3). Statistical analysis was performed with ANOVA with Tukey’s post hoc test, and * indicates p < 0.05, ** indicates p < 0.01, *** indicates p < 0.001
H&E staining revealed that the PE@MD group exhibited the most pronounced anti-inflammatory effect, with inflammatory cell infiltration reduced to 27.00 ± 4.58 cells/high-power field compared with 79.33 ± 7.09 cells/high-power field in the Control group (Fig. 9d). The wound length in the PE@MD group (1.56 ± 0.18 mm) was significantly shorter than that in the Control group (4.10 ± 0.42 mm), indicating enhanced wound contraction (Fig. 9c). Granulation tissue analysis showed that the PE group presented the most significant granulation thickness (1.67 ± 0.08 mm). In contrast, the PE@MD group exhibited moderate thickness (1.39 ± 0.11 mm) accompanied by complete re-epithelialization (100%) (Fig. 9e–f), suggesting an earlier transition into the remodeling phase. MT staining demonstrated markedly increased collagen deposition in the PE@MD group, reaching 64.41 ± 9.07% on day 14, approximately 2.2-fold higher than that of the Control group (29.18 ± 5.48%) (Fig. 9g). Sirius Red staining confirmed this trend (Fig. S14), showing the most intense and uniformly distributed red collagen fibers in PE@MD-treated wounds. These results indicate that the DEX-loaded composite dressing effectively promotes fibroblast activity, accelerates ECM maturation, and enhances the mechanical integrity of regenerated tissue.
Immunohistochemical quantification further revealed the molecular basis of the anti-inflammatory effect (Fig. 10). Representative staining images of COX-2, IL-1β, MPO, and NF-κB p65 are shown in Fig. 10a, with scale bars of 50 μm. At day 7, the PE@MD group markedly suppressed all four inflammatory markers compared with Control, P-Control, PG, and PE groups. Specifically, COX-2 decreased to 2.04 ± 0.002% (Control: 14.59 ± 0.004%), IL-1β to 21.10 ± 0.030% (Control: 52.12 ± 0.065%), MPO to 27.87 ± 0.018% (Control: 58.52 ± 0.037%), and NF-κB p65 to 23.08 ± 0.031% (Control: 58.04 ± 0.015%) (Fig. 10b–e). By day 14, similar trends were observed, with PE@MD further reducing NF-κB p65 to 18.17 ± 0.02% (Control: 25.81 ± 0.01%) and downstream mediators COX-2, IL-1β, and MPO to 0.02 ± 0.0001%, 6.72 ± 0.002%, and 7.81 ± 0.006%, respectively (Fig. 10f–i). These findings indicate that PE@MD attenuates inflammatory responses by inhibiting NF-κB pathway activation and suppressing multiple downstream effectors, thereby establishing a pro-regenerative microenvironment consistent with reduced neutrophil infiltration observed in H&E staining.
Fig. 10. Results of anti-inflammatory effects in wound tissues on day 7 and day 14. (a) Representative immunohistochemical staining images of COX-2, IL-1β, MPO, and NF-κB p6. Scale bar, 50 μm. Quantitative analysis of COX-2, IL-1β, MPO, and NF-κB p65 expression in wound tissues at (b-e) day 7 and (f-i) day 14, based on positive area percentage. Data are presented as mean ± SD (n = 3). Statistical analysis was performed with ANOVA with Tukey’s post hoc test, and * indicates p < 0.05, ** indicates p < 0.01, *** indicates p < 0.001
In vivo results indicate that the PE@MD composite dressing accelerates wound healing through coordinated modulation of inflammation and tissue remodeling. The observed suppression of NF-κB signaling, together with enhanced collagen deposition and accelerated re-epithelialization, suggests that PE@MD establishes a microenvironment conducive to effective tissue repair.
Discussion
In this study, a multifunctional composite nanofiber dressing (PE@MD) was successfully fabricated by integrating biodegradable PLCL, bioactive water-soluble eggshell membrane (ESM), and dexamethasone (DEX)-loaded mesoporous polydopamine nanoparticles (MPDA NPs) (Fig. 1a). The dressing demonstrated superior physicochemical properties (Fig. 4e–i, S7, S9, S10), biocompatibility (Fig. 5), and in vivo regenerative performance through complementary effects on oxidative stress (Fig. 6g, S11b), inflammation (Figs. 6h-l and 10, S11a), and extracellular matrix (ECM) remodeling (Fig. 9g, S14), which may include synergistic interactions. Incorporating ESM, a sustainable biomaterial derived from food industry by-products, endowed the scaffold with intrinsic bioactivity. At the same time, MPDA enabled sustained and controlled DEX release (Fig. 4j, S8), achieving both therapeutic efficacy and environmental sustainability. The comparison between PE and PE@MD provides clear evidence for the functional contribution of MPDA/DEX within the ESM-containing fibrous system, establishing the effectiveness of the current material design. Further inclusion of component-specific controls in future studies will enable deeper quantitative resolution of individual contributions.
Proteomic and metabolomic analyses revealed that ESM contains a broad spectrum of structural and functional biomolecules, including collagens, actins, lysozyme, annexin A1, and antioxidant enzymes such as catalase and peroxiredoxin-1 (Figs. 2c and 3, Table S2). These components are closely associated with cytoskeletal organization, inflammation resolution, and redox regulation—key processes in wound healing. The enrichment of fatty acids, amino acids, and TCA intermediates further suggests that ESM may participate in energy metabolism, collagen synthesis, and antioxidant defense during the healing process (Fig. 3a). Such compositional complexity aligns with previous reports showing that natural ESM facilitates fibroblast adhesion and proliferation while modulating cytokine expression and macrophage activation through its intrinsic peptides and glycoproteins [53, 54]. Therefore, using water-soluble ESM improves the biological functionality of synthetic polymers and offers an eco-friendly approach for advanced wound dressings. Collectively, these multi-omics analyses provide hypothesis‑generating insights into potential molecular pathways involved in the biological effects of PE@MD, offering preliminary guidance for future mechanistic studies.
The complementary effects of ESM and MPDA on oxidative stress, which may include synergistic interactions, were crucial in regulating inflammation and oxidative stress. MPDA contains abundant catechol and quinone groups, capable of scavenging reactive oxygen species and inhibiting free-radical-induced damage (Fig. 6g, S11b) [55]. A dual-mode anti-inflammatory effect was achieved with sustained DEX release from the mesoporous framework. DEX exerts its function via glucocorticoid receptor-mediated transcriptional regulation, suppressing pro-inflammatory cytokines such as IL-6, TNF-α by inhibiting the NF-κB signaling pathway (Fig. 10) [56]. Results showed that PE@MD reduced ROS levels as indicated by DCFH‑DA staining (Fig. 6g–h), accompanied by decreased COX‑2 and NF‑κB p65 expression (Figs. 6k-l and 10) and promotion of M2 macrophage polarization (Fig. 6h, S11a). These findings reliably indicate an overall antioxidant effect of the dressing, offering a foundation for further mechanistic studies on oxidative stress modulation [57, 58]. This controlled inflammatory modulation is critical, as excessive inflammation often leads to delayed epithelialization and tissue fibrosis [59].
At the cellular level, PE@MD effectively enhanced fibroblast proliferation and migration (Fig. 6b–f), processes fundamental for granulation tissue formation and ECM remodeling. In this full-thickness wound model, the electrospun dressing remains largely present at the wound site during the early healing phase (up to 14 days), interacting with blood components and wound exudates to form a provisional wound interface rather than undergoing rapid degradation (Fig. 4h). Accordingly, the observed modulation of cellular behavior is primarily attributed to the sustained release of bioactive components from the dressing rather than to structural guidance arising from fiber degradation, which is consistent with prevailing concepts in wound dressing function where biochemical cues dominate early cellular responses [60, 61]. The extract-based migration assays employed here effectively delineate the chemotactic contribution of released bioactive components, while future studies will integrate direct cell–fiber interaction models to assess combined biochemical and structural influences of the dressing. Whole-transcriptome sequencing of fibroblasts treated with dressing extracts revealed upregulation of genes related to antioxidant defense (Aldh1a7), glucocorticoid signaling (Tsc22d3), and ECM organization (Col1a1, Mcm2) (Fig. 7, S12, S13), highlighting potential effects on fibroblast function in vitro and providing a reference for future studies exploring wound healing mechanisms in vivo. Enriched pathways such as p53 signaling and ECM–receptor interaction indicate that the dressing supports cell cycle regulation and structural reconstruction. Similar mechanisms in other bioactive composite scaffolds have been observed that activate ECM remodeling and antioxidant pathways to accelerate skin regeneration [62, 63]. These findings demonstrate that PE@MD is a physical support for cell adhesion and actively participates in molecular events driving tissue repair.
In vivo experiments confirmed that PE@MD accelerated wound closure (Fig. 8b–e) and promoted re-epithelialization and collagen deposition (Fig. 9, S14) while significantly reducing inflammatory cell infiltration (Fig. 9d). The enhanced collagen density observed in Masson’s trichrome and Sirius Red staining implies improved matrix organization and mechanical strength of regenerated tissue, consistent with the transition from the proliferative to the remodeling phase. Moreover, immunohistochemical analysis showed that PE@MD markedly suppressed the expression of NF-κB p65, COX-2, IL-1β, and MPO (Fig. 10), confirming that the dressing effectively interrupted inflammatory signaling cascades and established a regenerative microenvironment. These results agree with recent reports that PDA-based or glucocorticoid-modified nanofibrous scaffolds can activate MAPK and VEGF pathways, promoting angiogenesis and collagen synthesis during tissue regeneration [64, 65].
Although sustained DEX delivery is beneficial for inflammation control, prolonged local glucocorticoid exposure may carry risks including delayed epithelial turnover and local immunosuppression [66]. The DEX dose used in this study was selected to remain within a low therapeutic window, and no adverse effects were observed during the experimental period. Long-term safety evaluation and dose optimization remain important objectives for future studies.
As a naturally derived biomaterial, eggshell membrane (ESM) is subject to inherent batch-to-batch variability. To minimize this effect in the present study, all raw materials were sourced from the same supplier and processed under identical standardized protocols. Future studies will further incorporate systematic batch characterization and quality-control metrics to strengthen reproducibility and translational reliability.
Collectively, this work proposes a novel therapeutic paradigm for wound healing that marries bioresource valorization with advanced nanostructural design and robust mechanistic validation. By harnessing the intrinsic bioactivity of ESM and the multifunctionality of MPDA, the PE@MD dressing exhibits sustained anti-inflammatory activity, balanced degradation, and favorable tissue compatibility. Although the present study demonstrated significant healing efficacy in an acute wound model within 14 days, its applicability to chronic wound settings requires further validation. Long-term evaluation of immune responses and remodeling outcomes in chronic wound models (e.g., diabetic or ischemic conditions) is warranted to confirm its safety and therapeutic potential for complex, non‑healing wounds. Future optimization may incorporate additional bioactive cues, such as growth factors or antibacterial peptides, to enhance vascularization and prevent infection, ultimately translating this sustainable biomaterial platform into clinical use.
Conclusion
This study developed a biodegradable composite nanofiber dressing (PE@MD) by integrating poly(L-lactide-co-ε-caprolactone) (PLCL), bioactive water-soluble eggshell membrane (ESM), and dexamethasone (DEX)-loaded mesoporous polydopamine nanoparticles (MPDA). The dressing exhibited favorable mechanical flexibility, sustained drug release, and excellent biocompatibility. ESM provided intrinsic antioxidant and regenerative bioactivity, while MPDA enabled controlled DEX delivery and ROS scavenging. In vitro and in vivo studies demonstrated that PE@MD effectively suppressed inflammation and oxidative stress via inhibiting the NF-κB pathway, promoted fibroblast proliferation and M2 macrophage polarization, and enhanced collagen deposition and re-epithelialization. These results highlight the complementary and potentially synergistic anti-inflammatory and pro-regenerative functions of ESM and MPDA, offering a sustainable and multifunctional strategy for efficient wound healing.
Supplementary Information
Below is the link to the electronic supplementary material.
Supplementary Material 1
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