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. 2025 Aug 27;10(35):40101–40110. doi: 10.1021/acsomega.5c04901

DFO-Loaded Tussah Silk Nanofibrous Membranes for Guided Bone Regeneration

Miaomiao He †, Yumao Chen ‡, Min Jin †, Weijian Song †,*, Minxuan Han †,*, Shijun Lu †,*
PMCID: PMC12423857  PMID: 40949287

Abstract

Biopolymer nanofibrous membranes have gained considerable attention as advanced biomaterial scaffolds for guided bone regeneration owing to their remarkable structural and functional characteristics. In this study, we developed a novel composite membrane by incorporating deferoxamine (DFO) into tussah silk nanofibrous (Tsn) membranes to synergistically enhance angiogenic and osteogenic capabilities. The physically sheared Tsn membranes predominantly exhibited β-sheet conformations, confirming the preservation of silk’s natural secondary structure. Comparative analysis between pristine Tsn and DFO-loaded (DFO-Tsn) membranes revealed that drug incorporation resulted in modest reductions in mechanical parameters, including breaking stress, breaking strain, and Young’s modulus. In vitro evaluations demonstrated a time-dependent increase in cell proliferation within the DFO-loaded group, suggesting that the composite membrane maintained excellent biocompatibility. Most notably, when implanted in a rat cranial defect model, the DFO-Tsn group showed substantially enhanced bone regeneration compared to Tsn controls, as evidenced by comprehensive micro-CT analysis and histomorphometric evaluation. These findings collectively demonstrate that DFO-loaded Tsn membranes show great potential for use in bone tissue regeneration.

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1. Introduction

Bone regeneration is a complex and multifaceted process that necessitates coordinated biological efforts. Upon damage to bone tissue, the body activates a series of signaling pathways in the affected region to preserve the internal balance of the bone tissue microenvironment, thereby facilitating the regenerative process. A key aspect of this mechanism is angiogenesis, the process of restoring blood supply to damaged tissues. Blood vessels play an essential role by delivering oxygen and nutrients to bone tissue while simultaneously removing metabolic waste, which is vital for the survival and functionality of bone cells. Furthermore, blood vessels regulate the interactions among osteoblasts, osteoclasts, and vascular cells, providing the necessary biological signals for bone formation. , Enhanced vascularization at bone injury sites can substantially improve the efficiency and quality of osseous regeneration, highlighting the critical interplay between angiogenesis and osteogenesis. , The hypoxia-inducible factor (HIF) is a transcription factor that binds to DNA and activates various hypoxia-related genes under low-oxygen conditions. It serves as a cellular oxygen sensor and is essential for maintaining bone homeostasis and promoting angiogenesis. − Vascular endothelial growth factor (VEGF) is a key regulator of angiogenesis, primarily responsible for stimulating blood vessel formation, ensuring vascular stability, and facilitating tissue repair and regeneration. Research in bone tissue engineering has demonstrated that HIF-1α enhances neovascularization by upregulating VEGF expression under hypoxic conditions. , These findings demonstrate that the strategic incorporation of hypoxia-inducing biomaterials into bone tissue engineering scaffolds represents a promising therapeutic strategy, effectively enhancing osteogenic potential while promoting vascularized bone regeneration.

Deferoxamine (DFO) is a hexadentate iron chelator frequently employed in the long term clinical management of conditions associated with excessive iron accumulation. As a recognized stabilizer of hypoxia-inducible factor 1α (HIF-1α), DFO effectively increases the levels of HIF-1α and its downstream angiogenic factors, thereby facilitating angiogenesis and supporting bone development. − Prolyl and asparaginyl hydroxylases are essential enzymes responsible for the degradation of HIF-1α, and the presence of iron ions is critical for their enzymatic activity. DFO chelates these iron ions, thereby inhibiting the activity of hydroxylases and preventing the hydroxylation and subsequent degradation of HIF-1α, which leads to its accumulation within cells. This accumulation activates downstream target genes of HIF, significantly enhancing local VEGF expression, which promotes angiogenesis by facilitating endothelial cell differentiation. , Furthermore, the expression of HIF-1α initiates a transcriptional cascade of various hypoxia-responsive genes that are involved in cell proliferation, migration, angiogenesis, and tissue regeneration, underscoring the significance of this pathway in bone regeneration. , Extensive research has confirmed that deferoxamine (DFO) administration effectively stimulates angiogenesis and significantly enhances bone regeneration. − The activation of the HIF-1α pathway, in conjunction with other factors, creates a favorable local environment for vascular development, indicating that DFO functions as a stabilizer of HIF-1α to promote tissue regeneration and address bone defects. , Therefore, the incorporation of DFO is anticipated to be a promising strategy for enhancing the vascularization and bone regeneration of scaffolds. ,

Guided bone regeneration (GBR) technology employs a barrier membrane to cover the site of a bone defect. This membrane functions as a mechanical barrier, inhibiting the ingress of nonosteogenic cells into the defect area and creating a relatively enclosed environment that is conducive to tissue growth during the healing process. − The barrier membrane is a critical component of GBR, and an ideal membrane should demonstrate exceptional biocompatibility, bioactivity, mechanical strength, and biodegradability. Membranes are generally classified into absorbable and nonabsorbable categories based on their degradability. Among the materials utilized for absorbable membranes, silk fibroin emerges as a promising biomedical alternative, exhibiting favorable biocompatibility and mechanical properties. Recent research has indicated that silk fibroin can enhance the proliferation and vascularization of endothelial cells, thereby improving outcomes in tissue regeneration.

In our previous work, we successfully fabricated tussah silk nanofiber (Tsn) membranes using high-speed physical shearing technology. These membranes serve as an effective barrier at the interface between soft tissue and the bone repair site, preventing the rapid invasion of connective tissue into the defect area while allowing osteoblasts to preferentially populate the region, thereby facilitating new bone formation.

Both in vitro and in vivo studies have demonstrated that Tsn membranes exhibit excellent mechanical properties, biocompatibility, and osteogenic potential. , First, the tightly stacked fiber structure of Tsn membranes enhances their tensile strength, Young’s modulus, and low strain, offering robust mechanical support for bone defects and facilitating osteogenesis. Second, the fabrication process avoids toxic solvents and minimizes damage to the silk fibroin’s molecular structure, ensuring superior biocompatibility. Moreover, tussah silk fibroin contains intrinsic Arg-Gly-Asp (RGD) tripeptide sequencesa key cell adhesion motif that regulates cell-material interactions, including cell attachment and angiogenesis, further enhancing osteogenic activity. , Besides, the biodegradable and absorbable nature of Tsn membranes eliminates the need for surgical removal, reducing the risks of secondary infection and delayed healing.

Building on these findings, we functionalized Tsn membranes with deferoxamine (DFO) through amidation reactions between amino groups of silk fibroin macromolecules and anhydride-activated DFO. This strategy introduced carbon–carbon double bonds to both components, creating a DFO-modified membrane designed to enhance local vascularization and subsequent osteogenesis. Conventional guided bone regeneration (GBR) membranes primarily serve as mechanical barriers but lack bioactive properties to actively promote tissue regeneration. In this study, we innovatively integrate the robust mechanical support of tussah silk nanofibers (Tsn) with the pro-angiogenic effects of deferoxamine (DFO), thereby overcoming the functional limitations of traditional GBR membranes that act solely as passive barriers. Notably, in vivo experiments using a rat calvarial defect model demonstrated that the 300DFO-Tsn group induced near-complete bone regeneration within 12 weeks. These findings not only validate the dual functionality of the composite membrane but also present a promising therapeutic strategy for clinical large bone defect repair.

2. Materials and Methods

2.1. Materials

Tussah silk (Jiangsu, China), anhydrous sodium carbonate (Na2CO3, China National Pharmaceutical Group Corporation), anhydrous ethanol (China National Pharmaceutical Group Corporation), Human Umbilical Vein Endothelial Cells (HUVEC), DMEM high-sugar medium (Gibco), fetal bovine serum (FBS, Procell), penicillin–streptomycin (Sigma), 0.25% trypsin digestive solution (Sigma), dimethyl sulfoxide (Sigma), paraformaldehyde (PFA) (Sigma), DFO (MCE), Calcein/PI cell viability and cytotoxicity assay kit (Shanghai Beyotime Biotechnology Co., Ltd.), and a CCK-8 kit (Shanghai Beyotime Biotechnology Co., Ltd.).

2.2. Preparation of Tsn and DFO-Tsn Membrane

The Tussah silk was subjected to treatment in a boiling solution of 0.5 wt % sodium carbonate to remove sericin, utilizing a bath ratio of 1:50. After a duration of 30 min, the silk was removed and thoroughly rinsed, with this process being repeated three times before drying in an oven at 60 °C. As shown in Figure , the degummed tussah silk was subsequently cut into 0.5 cm fragments and mixed with deionized water. This mixture was then processed in a high-speed shearing machine operating at a rotational speed of 32,000 rpm for 30 min to obtain Tsn solution. The Tsn solution was evenly distributed in a Petri dish and placed in an oven at 60 °C to evaporate the water, resulting in the formation of a Tsn membrane.

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Schematic illustration of the preparation and application of DFO-Tsn membrane. (Created in BioRender. He, M. (2025) https://BioRender.com/xa73u8u.)

In accordance with our previous report, a silk fibroin (SF) solution was prepared. Then, a 2 wt % SF aqueous solution was mixed with methacrylic anhydride at a ratio of 100 mL to 50 mg and stirred in a water bath at 60 °C for 4 h to aminate the SF macromolecules (SF@AM). Subsequently, three concentrations of DFO (250, 500, and 1000 μL/mL) were added to the SF@AM solution, and after thorough mixing, a 0.5 wt % photoinitiator LAP was added. Finally, the mixed solution was combined with Tsn and precross-linked under blue light irradiation at an intensity of 45 mW/cm2 for 2 h, resulting in DFO-loaded Tsn membranes. The resulting membranes were designated as Tsn membrane (corresponding to a DFO concentration of 0), 75DFO-Tsn membrane (corresponding to a DFO concentration of 75 μmol/L), 150DFO-Tsn membrane (corresponding to a DFO concentration of 150 μmol/L), and 300DFO-Tsn membrane (corresponding to a DFO concentration of 300 μmol/L).

2.3. Morphology and Structure

The Tsn and DFO-Tsn membranes were mounted onto the stage of an electron microscope and subsequently coated with a gold spray, achieving a thickness of 20 to 30 nm. The morphological characteristics of the samples were analyzed using a Hitachi S8100 scanning electron microscope operating at a voltage of 3 kV. The absorption peaks of the samples were recorded within the wavenumber range of 400 to 4000 cm–1 utilizing a Fourier transform infrared spectrometer (FTIR), with a resolution of 4 cm–1 and 32 scans.

2.4. Physical Properties

2.4.1. Mechanical Properties

For the mechanical testing, we utilized an Instron 5967 universal material testing machine (Boston, USA) in a controlled environment maintained at a temperature of 20 ± 2 °C and a relative humidity of 65 ± 2%. The Tsn and DFO-Tsn membranes were cut into zigzag samples measuring 5 cm, with five parallel samples prepared for each group. The cut samples were immersed in deionized water until they were fully saturated, and any excess water on the surface was removed prior to the mechanical tests.

To evaluate the mechanical properties, we employed an Instron 5967 universal material testing machine (Boston, USA) under controlled environmental conditions, with the temperature maintained at 20 ± 2 °C and relative humidity at 65 ± 2%. The Tsn and DFO-Tsn membranes were carefully excised into rectangular specimens measuring 50 mm in length and 10 mm in width, with a thickness ranging approximately between 100 to 120 μm. To ensure statistical reliability, five replicate specimens were prepared for each experimental group. Before conducting mechanical testing, the prepared samples were thoroughly saturated by immersion in deionized water, after which any excess surface moisture was gently blotted away to maintain uniformity and reproducibility in testing conditions.

2.4.2. Degradation Experiment

Tsn/DFO-Tsn membranes (n = 3/group) were precisely weighed (designated as initial mass M 0) and placed in a 24-well plate. Each sample was subsequently treated with 2 mL of either phosphate-buffered saline (PBS) or protease XIV solution (1 U/mL in PBS), followed by incubation at 37 °C under controlled conditions. At predetermined time points (12, 24, 48, 72, 96, 120, 144, and 168 h), the membranes were retrieved, thoroughly rinsed with deionized water, completely dried, and weighed to determine their residual mass (recorded as M 1). The degradation kinetics were then quantified by calculating the remaining mass percentage using the following formula.

Calculationformula:Weightremaining(%)=(1−M0−M1M0)×100%

2.4.3. Drug Release Experiment

The samples (n = 3/group) were loaded into 5000 Da dialysis bags and placed in 10 mL of PBS. At 0, 1, 2, 4, 6, 12, 24, 48, and 72 h, 1 mL of the buffer was collected and replaced with an equal volume of fresh buffer. The absorbance was then measured using a microplate reader (Synergy NEO, BioTek) to determine the drug release efficiency.

2.4.4. Swelling Ratio Test

The swelling ratio of Tsn/DFO-Tsn membranes was evaluated using the gravimetric method. The dry weight of the samples (n = 3/group) was measured and recorded as M 0. The samples were then immersed in PBS at 37 °C. After 12 and 24 h, the samples were removed, excess liquid on the surface was gently wiped off, and the wet weight was recorded as M 1. The swelling ratio was calculated using the following formula:

Calculationformula:Swellingratio(%)=M1−M0M0×100%

2.5. Biocompatibility

2.5.1. Cell Culture and Proliferation

The Tsn and DFO-Tsn membranes were sectioned into circular discs measuring 5 mm in diameter. Subsequently, these membrane discs underwent sterilization via high-temperature and high-pressure treatment. After sterilization, the discs were carefully transferred to a 48-well plate and immersed in basal culture medium. Following a 24-h incubation period, the membranes were removed, and human umbilical vein endothelial cells (HUVECs) were seeded into the conditioned medium at a density of 4 × 104 cells per well. The cell cultures were maintained in a humidified incubator at 37 °C with 5% CO2 atmosphere, with regular medium replacement every 48 h. To assess cell proliferation, the culture medium was aspirated at specific time points (1, 2, and 3 days postinoculation), and 250 μL of Cell Counting Kit-8 (CCK-8) solution was added to each well. After a 2-h incubation, 100 μL of the supernatant from each well was measured at 450 nm using a microplate reader.

2.5.2. Living-Dead Cell Staining

The Calcein-AM/PI dual-staining was conducted for cellular viability assessment. The working solution was prepared by mixing 10 μL of Calcein-AM (1 mmol/L) and 15 μL of PI (1.5 mmol/L) stock solution with 5 mL of phosphate-buffered saline (PBS), yielding final concentrations of 2 μmol/L for Calcein-AM and 4.5 μmol/L for PI, respectively. Prior to staining, the culture medium was carefully aspirated, and the cells were gently washed with PBS three times to completely remove residual medium and serum components. The cells were then incubated with the staining solution for 30 min at 37 °C, followed by an additional five washes with PBS to remove unbound dye molecules. After thorough removal of the staining solution, cellular imaging was performed using a laser scanning confocal microscope.

2.6. Vessel-like Formation Assay

The 96-well plate was prechilled on ice for 5 min before being uniformly coated with 50 μL of growth factor-reduced Matrigel per well, followed by incubation at 37 °C in a 5% CO2 humidified atmosphere for 30 min to facilitate complete polymerization. HUVECs were then seeded onto the prepared Matrigel substrate at a density of 2 × 104 cells/well. Following a 6-h incubation period with the test extraction medium, cellular responses were documented through phase-contrast microscopic imaging.

2.7. In Vivo Study

2.7.1. Surgical Procedure

Fifteen male Sprague–Dawley rats (230–260 g) were used to establish critical-sized calvarial defect models (8 mm in diameter) for evaluating guided bone regeneration efficacy. All animal experiments were approved by the Ethics Committee of Soochow University. The rats were anesthetized by intraperitoneal injection of 4% chloral hydrate at a dosage of 1 mL per 100 g body weight. Upon achieving complete anesthesia, surgical access to the cranium was established, and bilateral circular defects, each measuring 8 mm in diameter, were created. Subsequently, the Tsn and DFO-modified Tsn (DFO-Tsn) membranes were positioned to cover the defects. At predetermined intervals of four and 12 weeks postimplantation, the animals were euthanized, and their crania were carefully reopened for comprehensive analysis of bone regeneration.

2.7.2. Microcomputed Tomography Analysis

The morphologies and volumetric characteristics of the regenerated bone tissue were quantitatively assessed through microcomputed tomography (micro-CT) analysis (SkyScan 1176, Bruker Corp., Kontich, Belgium). The analysis specifically focused on determining the percentage of regenerated bone relative to the original defect area. The scanning parameters were established at a voltage of 65 kV, a current of 100 μA, an exposure time of 600 ms, and an aluminum filter of 1 mm.

2.7.3. Histological Staining

Following microcomputed tomography (micro-CT) analysis, the harvested tissue samples were subjected to histological examination to evaluate bone regeneration, tissue integration, and the host response. The fixed tissues were then embedded in paraffin and sectioned axially at a thickness of 5 μm. For comprehensive histological assessment, sections were stained with hematoxylin–eosin (H&E) and Masson’s trichrome stain. The stained sections were mounted on glass slides and examined under a light microscope (Axioveter 40 CFL, Zeiss, Oberkochen, Germany).

2.8. Statistical Analysis

All quantitative data are presented as mean ± standard deviation (SD). Statistical significance was assessed using Student’s t test and one-way ANOVA, with p < 0.05 considered statistically significant.

3. Results and Discussion

3.1. Morphology and Structure

The morphological characteristics of the Tsn and DFO-Tsn membranes are depicted in Figure A. The Tsn membranes exhibited a well-defined network structure composed of numerous interconnected nanofibers. These nanofibers were further bonded together through silk fibroin hydrogel to form the DFO-Tsn membranes. The Tsn were fabricated via a top-down fibrillation approach. Owing to inherent limitations of this technique, the resulting fibers exhibit broad diameter distributions, leading to heterogeneous Tsn membranes as depicted in Figure A­(a). Upon integration with DFO-loaded SF hydrogel, these membranes form composite materials through interfacial bonding between the fibrous phase and gel matrix, as shown in Figure A­(b–d). Crucially, both pure Tsn membranes and DFO-SF gel composites function as effective physical barriers for bone regeneration, preventing soft tissue infiltration by virtue of their densely packed nanofiber architectures. The structural and conformational properties of the Tsn and DFO-Tsn membranes were analyzed using infrared spectroscopy, as illustrated in Figure B. The spectra revealed conformation-sensitive bands at 1628 cm–1 (amide I, representing CO stretching), 1517 cm–1 (amide II, associated with N–H bending), 1240 cm–1 (amide III, corresponding to C–N stretching), and 965 cm–1 (amide IV, indicative of C–C twisting), all of which are consistent with the characteristic β-sheet conformation of silk fibroin. −

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Characterization of DFO-Tsn Membrane. (A) SEM image of the DFO-Tsn Membranes (a: Tsn; b: 75 DFO-Tsn; c: 150 DFO-Tsn; d: 300 DFO-Tsn). (B) FTIR spectra (C); Breaking stress; (D) Breaking strain; (E) Young’s modulus of Tsn/DFO-Tsn Membranes. (F,G) In vitro degradation diagram of Tsn/DFO-Tsn membrane. (F) PBS; (G) XIV protease. (H) Drug release curve of DFO-Tsn membrane. (I) Swelling properties diagram of Tsn/DFO-Tsn membrane. (* p < 0.05, **p < 0.01, ***p < 0.001).

3.2. Physical Properties

The mechanical properties of the membranes in a hydrated state were evaluated to align with their intended application conditions. The mechanical characteristics of the Tsn and DFO-Tsn membranes were presented in Figure (Figure C–E). The Tsn membrane exhibited a breaking stress, breaking strain, Young’s modulus of 6.24 ± 1.27 MPa, 16.40 ± 1.50%, and 0.32 ± 0.04 MPa, respectively. In contrast, the DFO-Tsn membrane demonstrated lower values for these parameters. Specifically, the 75 DFO-Tsn group displayed a breaking stress of 2.52 ± 0.37 MPa, breaking strain of 12.46 ± 2.55%, and Young’s modulus of 0.24 ± 0.04 MPa. The 150 DFO-Tsn group showed further reductions, with values of 1.98 ± 0.23 MPa (breaking stress), 13.25 ± 1.95% (breaking strain), and 0.20 ± 0.02 MPa (Young’s modulus). The 300 DFO-Tsn group exhibited the lowest mechanical performance, with a breaking stress of 1.19 ± 0.37 MPa, breaking strain of 14.13 ± 3.43%, and Young’s modulus of 0.23 ± 0.02 MPa. The reduction in mechanical properties can be primarily ascribed to the swelling of the membranes triggered by the introduction of an aqueous solution. This swelling phenomenon leads to both an increase in membrane thickness and a weakening of interfiber bonding strength. Moreover, the limited interfiber adhesion capability of the silk protein gel further exacerbates the deterioration of mechanical performance.

In addition, we systematically characterized the degradation profile of DFO-Tsn membranes (Figure F,G). Appropriate degradation kinetics are crucial for bone tissue engineering, as timely degradation provides necessary space for tissue regeneration while avoiding secondary trauma from surgical removal. Our results demonstrated comparable degradation rates between Tsn and DFO-Tsn groups without significant differences. Both groups exhibited rapid degradation within the initial 24 h in PBS and 1 U/mL protease XIV solutions, followed by sustained slow degradation. This biphasic degradation behavior offers dual advantages: the initial rapid phase facilitates effective drug release to promote tissue regeneration in bone defect areas, while the subsequent prolonged degradation maintains structural integrity to protect the defect site. Such protection prevents premature fibrous tissue invasion while allowing sufficient time for osteogenic cell migration and new bone formation.

Further drug release analysis revealed that DFO-Tsn membranes achieved approximately 50% drug release within 24 h, followed by sustained release kinetics (Figure H). This dual-phase release profile, featuring both rapid initial release and sustained long-term delivery, represents an optimal kinetic pattern for promoting continuous bone regeneration. Moreover, DFO-Tsn membranes demonstrated exceptional swelling capacity (>2500% after 24 h), which provides two key benefits: (i) significantly reducing mechanical irritation to surrounding tissues during early implantation, and (ii) better mimicking native extracellular matrix (ECM) to enhance cell adhesion, migration and proliferation (Figure I).

3.3. In Vitro Cell Biocompatibility

The viability of HUVECs cultured on Tsn and DFO-Tsn membranes was assessed at 1, 2, and 3 days using Calcein-AM/PI dual staining. The live/dead fluorescence assay demonstrated a time-dependent proliferation of viable cells, indicated by the escalating intensity of green fluorescence associated with live cells (Figure A). The proliferative capacity of HUVECs on nanofiber membranes was quantitatively evaluated using the CCK-8 assay (Figure B). Quantitative analysis revealed comparable growth kinetics between the Tsn membrane and DFO-Tsn membrane during the culture period, with all groups exhibiting a progressive increase in cell viability. Notably, the DFO-Tsn groups displayed enhanced cellular activity on day 1, showing significantly higher optical density (OD) values compared to the Tsn group. By day 2, divergent proliferation patterns emerged: the 75 DFO-Tsn group demonstrated superior proliferative activity relative to Tsn, whereas the 150 DFO-Tsn group exhibited a nonsignificant reduction in proliferation. In contrast, the 300 DFO-Tsn group showed statistically significant growth inhibition compared to the Tsn control (p < 0.05). On day 3, while the drug-loaded groups maintained lower proliferation rates than the Tsn group, no significant difference was observed across experimental cohorts.

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Biocompatibility of DFO-Tsn membrane. (A) Living-dead cell staining (cell culture for 24, 48, 72 h). (B) In vitro cytotoxicity. Proliferative activity tested by CCK-8. (C) Representative images of tube formation after 6 h treatment with Tsn/DFO-Tsn membrane extracts. Quantitative analysis of (D) tubular nodes and (E) total tube length. (* p < 0.05, **p < 0.01, ***p < 0.001).

Given the well-established coupling between angiogenesis and osteogenesis, we evaluated the vascularization potential of DFO-incorporated membranes through in vitro tube formation assays. HUVECs treated with Tsn/DFO-Tsn membrane extracts for 6 h exhibited dose-dependent enhancement of capillary-like network formation (Figure C), with 75DFO-Tsn and 300DFO-Tsn groups demonstrating significantly more developed vascular lumens compared to Tsn controls. Quantitative analysis revealed that the 300DFO-Tsn group exhibited an increase in nodal junctions (p < 0.01 vs Tsn; Figure D), while both DFO-loaded groups showed greater total tube lengths (p < 0.05; Figure E). These findings demonstrate that the sustained release of DFO from Tsn membranes creates a pro-angiogenic microenvironment that may facilitate subsequent bone regeneration through enhanced vascular invasion.

3.5. In Vivo Bone Regeneration

Figure A demonstrated the successful development of a rat calvarial defect model. Postoperatively, all experimental animals demonstrated stable physiological recovery, retaining normal body mass indices and showing no clinical evidence of graft rejection, systemic infection, or wound complications throughout the 12-week observation period. Micro-CT imaging analysis (Figure B) demonstrated a progressive reduction in bone defect dimensions from 4 to 12 weeks, indicating significant osteogenic activity at the defect margins. Postoperative evaluation at one month revealed significantly enhanced osteogenesis in the 300 DFO-Tsn group compared to both the Tsn group (p < 0.05) and the 75 DFO-Tsn group (p < 0.05). Quantitative analysis of bone volume to total volume (BV/TV) ratios demonstrated a clear dose-dependent response: 300 DFO-Tsn (19.17 ± 1.18%), 75 DFO-Tsn (16.91 ± 0.30%), and Tsn (14.52 ± 1.08%) (Figure C). This hierarchical pattern persisted through the 3-month end point, with BV/TV values of 31.43 ± 3.10% (300 DFO-Tsn), 22.54 ± 1.05% (75 DFO-Tsn), and 17.05 ± 2.86% (Tsn), respectively. Statistical analysis confirmed that the 300 DFO-Tsn group maintained superior osteogenic performance compared to both the Tsn group (p < 0.01) and the 75 DFO-Tsn group (p < 0.05), while the 75 DFO-Tsn group consistently outperformed the Tsn group (p < 0.05) across both time points.

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(A) Establishment of rat skull defect model (a: Removal of hair from the surgical area of the rat; b: exposure of the surgical site; c: creation of the skull defect; d coverage with the membrane). (B) Micro-CT reconstruction images of bone defects at 4 and 12 weeks after operation. (C) New bone formation analyzed by Micro-CT at 4 and 12 weeks after operation. The value of BV/TV proves the role of osteogenesis (BV: bone volume; TV: tissue volume; * P < 0.05).

3.6. Histological Evaluation

To assess the integrated morphologies of soft tissues, neoformed bone, and native bone across experimental groups, tissue specimens were stained with hematoxylin-eosin and Masson’s trichrome at postoperative weeks 4 and 8 (Figures and ). The results revealed a substantial reduction in fibroblast proliferation and soft tissue infiltration at the bone defect sites across all groups, affirming the nanofiber membrane’s efficacy in preventing soft tissue invasion, consistent with previous findings. Bone regeneration was evident across all experimental groups at both 4-week and 8-week intervals, with a progressive increase in new bone formation over time. Notably, the Tsn group exhibited restricted osteogenic capacity, with substantial defect persistence at the 12-week follow-up, aligning with our prior study. This observation implies that while the Tsn membrane effectively serves as a physical barrier to facilitate osteogenesis, it lacks the biological activity necessary to substantially enhance bone regeneration. The incorporation of DFO into the Tsn membrane markedly improved bone defect repair, with significantly greater osteogenic activity observed at both 4-week and 12-week time points compared to the Tsn membrane alone, corroborating with our CT findings (Figure ). Histological examination revealed pronounced angiogenesis in the DFO-Tsn group, which appears to facilitate cellular proliferation, collagen matrix deposition, and subsequent bone formation. Extensive research has demonstrated that deferoxamine (DFO) activates and stabilizes the hypoxia-inducible factor-1α (HIF-1α) pathway, leading to upregulated expression of key angiogenic growth factors including vascular endothelial growth factor (VEGF), platelet-derived growth factor (PDGF), and stromal cell-derived factor-1 (SDF-1), consequently enhancing neovascularization. − This angiogenic enhancement likely represents the principal mechanism underlying the observed improvement in bone regeneration.

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HE staining section of rat skull defect at 4 and 12 weeks. M: nanofiber membrane; NB: new bone; OB: old bone. Black arrows represent blood vessels, white arrows represent osteoblasts, and blue arrows represent osteoblasts.

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Masson staining section of rat skull defect at 4 and 12 weeks. M: nanofiber membrane; NB: new bone; OB: old bone. Black arrows represent blood vessels, white arrows represent osteocytes, blue arrows represent osteoblasts, and yellow arrows represent collagen fibers.

4. Conclusions

In summary, Tsn membranes containing DFO were successfully fabricated through physical shear and silk fibroin gel as a carrier. These membranes were then utilized for the guided bone tissue regeneration. The Tsn membranes possessed a biomimetic nanofibrous structure, and the integration of DFO-loaded silk fibroin gel with Tsn membranes led to the aggregation of nanofibers. The mechanical properties of Tsn membranes were compromised due to the intrinsically weak mechanical strength of silk fibroin gel and its swelling effect on the Tsn membranes. Cellular assays demonstrated that higher DFO concentrations negatively impacted cell proliferation. In an in vivo rat experiments, the DFO-Tsn group exhibited significantly superior bone volume regeneration compared to the Tsn group, with better outcomes observed at higher DFO concentrations. The micro-CT imaging and histological staining confirmed that the 300DFO-Tsn membrane achieved complete bone tissue regeneration within 12 weeks.

While the current study demonstrates the osteogenic potential of DFO-Tsn membranes, the precise molecular mechanisms remain to be fully elucidated due to technical limitations. Future investigations incorporating both in vitro and in vivo approaches will be essential to delineate the genetic and molecular pathways underlying their bone-promoting effects. Overall, the DFO-Tsn membrane presents a promising solution for bone tissue regeneration applications.

Acknowledgments

This work was supported financially by the Project of State Key Laboratory of Radiation Medicine and Protection, Soochow University (GZK12024041), the 2023 Jiangsu Provincial Traditional Chinese Medicine Science and Technology Development Plan (MS2023170), and the China University Industry-University-Research Innovation Fund - Special Project on Infection and Control (2024GR089).

M.H.: Data curation, methodology, investigation, writingoriginal draft. Y.C.: Investigation, formal analysis, writingoriginal draft. M.J.: Data curation, methodology, conceptualization. W.-J.S.: Writingreview and editing, resources, project administration. M.Han: Supervision, resources, project administration, funding acquisition. S.L.: Methodology, investigation, funding acquisition, writingreview and editing, supervision, resources. M.H., Y.C., and M.J. contributed equally to this work.

The authors declare no competing financial interest.

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