HAp/SF composite fibers with 10% HAp were selected as drug carriers to prepare drug-loaded fibers with a certain amount of NAC. The blending method was used to disperse NAC evenly in the spinning solution. Drug-loaded fibers were prepared by electrospinning. A solution was prepared by dissolving the required weight of NAC in 10 μl of water and adding it to the spinning solution, followed by stirring, and mixing. 0.4 g of HAp was added to a stirring bottle containing 2 ml of methane acid. The HAp particles were evenly dispersed by ultrasonication for 30 min 0.4 g of SF and 0.0082 g of polyethylene oxide (PEO) were then added to the solution and stirred and dissolved. After stirring for 1 h, 10 μL of an aqueous solution containing NAC (0.78 mg) was added, and the NAC/HAp/SF solution was then prepared by magnetic stirring for 5 h at room temperature. The electrospinning was carried out at a spinning voltage of 7–8 kV, an injection rate of 0.4 ml/h, a receiving distance of 15 cm, at room temperature, and an ambient humidity of 30–40%.

The contact angles of the different drug-loaded fiber membranes were measured using a contact-angle tester (JCY-1, Shanghai Fangrui Instrument Co., Ltd.). 0.3 μL of deionized water was dropped on the sample. The morphologies of the water droplets on the fiber membrane at different times were imaged. The angle between the water droplet and fiber membrane was measured to obtain the contact angle.

Ultraviolet spectrophotometry (Thermo Scientific Evolution 300, United States) was used to analyze drug release. Scaffold samples (20 mg, alcohol gas treated for 1 h) were immersed in 2 ml of distilled deionized water. The samples were then mixed with a 4-chloro-7-nitrobenzofurazan (0.006 wt%) chromogenic agent in a ratio of 1:18. After 30 min of reaction, the NAC concentrations were calculated based on the absorbance at 423 nm.

The rat iPSC-MSCs was induced by the rat iPS cells (Sidansai Biotechnology, Shanghai, China, China) cultured in MSCs medium and consisted of Dulbecco’s Modified Eagle’s Medium (Hyclone, United States), 1% penicillin/streptomycin (Tianjin Haoyang Biological Products Technology Co. Ltd., China), l-glutamine (Gibco, United States), and 10% fetal bovine serum (Gibco, United States). The iPSC-MSCs were cultured at 37°C in a 5% CO₂ humidified incubator and the culture medium was changed every 2 days.

To prepare nanofiber scaffolds for cell culture, smooth glass sheets (diameter 15 mm) were cleaned, sterilized, and placed on an aluminum foil to collect the fibers. After electrospinning of the 2-ml solution, the prepared HAp/SF membranes were dried in a vacuum-drying chamber for at least 5 days to remove residual solvents. After the solvent was volatilized, the samples were placed in the corresponding cell-culture plate. Four-to-six samples for each structure were utilized, with blank cover slides placed into a plate as a control group. After 10 min of methanol treatment, the SF structure was transformed, methanol was removed, and sample scaffolds were irradiated by ultraviolet light for 2 days after methanol volatilization. The samples were soaked in 75% ethanol for 2 h, and then washed for 10 min with PBS (Phosphate Buffer Saline) three times. Each pore was added to the corresponding cell culture medium. The sample was soaked in an incubator overnight for preculture. The sample was prepared squarely and placed in a cell-culture plate. iPSC-MSCs were planted according to the experimental requirements.

Cell spreading on fibers was observed using SEM. To prepare the samples for SEM, the following procedure was used: 1) Cell–scaffold complexes were cultured for 1 and 7 days and then removed from the culture medium and washed with PBS three times; 2) 4% glutaraldehyde was added to the complexes for a period of 2.5 h or longer to immobilize the samples; 3) after the glutaraldehyde was removed, the complexes were washed by PBS three times; 4) samples were washed for 15 min in each of six concentrations of alcohol (10, 30, 50, 70, 90, and 100%) for dehydration; 4) treatment with hexamethyldisiloxane was carried out overnight. After drying, the structures were cut into fragments, sprayed with gold, and observed using SEM.

The effects of the scaffolds on cell proliferation were evaluated by using cell counting kit-8 (CCK-8). At set times (1, 4, and 7 days), an appropriate volume of CCK-8 solution was added to the plates and incubated for 4 h. 200 μl of the culture medium was then extracted and added to a 96-well plate, without bubbles. The absorbance at 450 nm was determined using a microplate reader.

After 14 days incubation, the expressions of osteogenesis-related genes (ALP, COL, OCN (osteocalcin), and OPN (osteopontin)) in iPSC-MSCs cultured in HAp/SF, 10-NAC/HAp/SF, 20-NAC/HAp/SF nanofiber membranes, and the TCP blank control were detected by reverse transcription polymerase chain reaction (RT-PCR). All primer sequences were listed in Table 1. The procedure for RT-PCR included three principles processes: ribonucleic acid (RNA) extraction, reverse transcription deoxyribonucleic acid (DNA) synthesis, and RT-PCR gene amplification detection. RNA was rapidly extracted using a Biozol RNA extraction kit. Reverse transcription DNA synthesis was undertaken using a 20 L reactor containing 20–50 ng of RNA. RNA was reverse-transcribed to form complementary DNA (cDNA) using the FastQuant RT Kit (with gDNase). RT-PCR was performed using the SuperReal PreMix Plus (SYBR Green) kit, with the housekeeping gene (GAPDH) as a control. After 40 cycles, the relative expression of each target gene was calculated using the 2^−ΔΔct method.

Dual-source CT (SOMATOM Definition, Siemens) was used to detect regeneration and quantify the mineral density of the newly formed bone within the cranial defects. Under general anesthesia, the mouse was fixed and scanned over the entire length at a voxel size of 12 l m and medium resolution of 80 kVp, 110 µA with a 0.5-mm Al filter. The regeneration of bone within the defects was visually identified and the manufacturer’s evaluation software was used to separate bone from non-bone and the mineral density of the new bone-like tissue was computed and calibrated to a HAp phantom.

All quantitative values are expressed as mean ± standard error of at least three replicate samples. An analysis of variance for all quantitative tests was carried out with the software Origin and Tukey’s honestly significant difference. Post hoc tests were used for pair-wise comparisons between groups. Statistical significance was defined by either *p < 0.05 or **p < 0.01.

Composite fibers were prepared via electrospinning. The morphologies of the different electrospun fibers were observed by SEM (Figure 1) and all fibers exhibited a good morphology. White substances appeared on the surface of the fiber because of the aggregation of Hap. The TEM images showed that HAps were encapsulated within the electrospun silk fibers and the diameter of the fibers decreased with the incorporation of NAC and NAC/HAp, which may increase the conductivity and viscosity of the SF fibers, consistent with the results of Zhu et al. (2019). Typical tensile stress-strain curves and tensile mechanical properties of composite fiber films are shown in Figure 2. The addition of HAp, NAC and NAC/HAp had a significant effect on the mechanical properties of SF fibers. The average Young’s modulus and tensile strength of SF fibers were 53.76 ± 11.67 and 2.54 ± 0.30 MPa, respectively. With the addition of HAp, tensile strength and Young’s modulus significantly increased. The incorporation of NAC reduced the Young’s modulus and the tensile strength to 36.43 ± 19.99 MPa and 0.89 ± 0.09 MPa, respectively. When NAC/HAp were added, the Young’s modulus and the tensile strength increases significantly. The rate of elongation rate decreased with the addition of HAp, NAC and NAC/HAp into SF fibers. Generally speaking, the mechanical properties of the fibers were improved by adding NAC/HAp.

The hydrophilic angles of the composite fiber membranes were measured to evaluate their hydrophilic properties. Contact angles below 90° are indicative of hydrophilicity, which favors the diffusion of liquid on the materials. In contrast, contact angles exceeding 90° correspond to hydrophobicity (Trinca et al., 2017). The contact angles of NAC/HAp/SF were smaller than those of the other scaffolds (Figure 3). Thus the addition of NAC and HAps decreased the contact angle of the material surface through the synergy between NAC, which contains amine and carboxylic functional groups, and HAps, which contain hydroxyl groups.

Figure 4 shows the cumulative release of NAC per Gram of the composite fiber scaffold. The NAC/SF group exhibited an initial burst of release; NAC then became exhausted after a period of time. In contrast, the release of NAC from the NAC/HAp/SF group exhibited a biphasic pattern, characterized by an initial burst of release followed by a long-term, gradual, and continuous release, during which high NAC concentrations were maintained sufficient to promote cell differentiation. In the first stage, the apparent fast release of NAC drugs may have occurred because some of the small NAC molecules dispersed on the surface of the fiber and the inner layer of the fiber epidermis during the electrospinning can be released quickly in the PBS buffer solution (Zheng et al., 2013). In the second stage, NAC inside the composite fiber was gradually released. This indicates that HAp exerted an adsorption effect on NAC and also a significant effect on the drug release behavior of NAC.

The CCK-8 assay was used to detect the proliferation of iPSC-MSCs cultured on composite fiber membranes (Figure 5). The cell content in all groups increased with the duration of incubation, which indicates that the cells were in a good state of proliferation. During the process of cell culture, the capacity for cell proliferation of all material groups was higher than that of the coverslip group. Cell proliferation activity was significantly higher in the NAC/SF and NAC/HAp/SF groups than in the control group. This suggests that the addition of NAC and HAps positively affected cell viability.

The proliferation and spreading of iPSC-MSCs on different composite fibers were further observed by SEM (Figure 6). After 4 days of culture, the number of adherent cells in the NAC/HAp/SF group was higher than those in the other groups. After 7 days of culture, the number of cells on the fiber increased and the spread of the cells were incorporated with the fiber scaffold. From the perspective of the number of fiber cells, this trend was consistent with the CCK8 results.

To evaluate the capacity for osteogenic differentiation of iPSC-MSCs on different nanofiber scaffolds, the marker genes of osteogenic differentiation (ALP, COL, OCN, and OPN) were detected by RT-PCR. After 14 days of culture, cells in the corresponding culture plate were collected and RNA was extracted for detection. The SF group was used as a control group in this experiment. The expression of ALP in the NAC/HAp/SF fibers increased significantly after 14 days (Figure 7). This may be explained by the fact that as the fibers promote the osteogenic differentiation of iPSC-MSCs and the cell differentiation process is in the middle and late stages, the content of the early expression product of ALP gradually decreases. With the addition of HAp and NAC, the expression levels of COL and OCN were significantly increased (p < 0.01). According to the gene expression level of OPN, the addition of HAp increased the level of OPN after 14 days of cell culture, while the addition of NAC did not affect the expression of OPN. The addition of NAC increased not only the level of expression of ALP, but also those of COL and OCN, the marker genes in the middle and late stages of osteogenesis but had no significant effect on the expression of OPN. However, compared to the control group, HAp and NAC complemented each other and promoted the expression of ALP, COL, OCN, and OPN genes, thus promoting the osteogenic differentiation of iPSC-MSCs.

iPSC-MSCs were planted on different composite fibers. After 14 days of culture, ALP staining and quantitative analysis of ALP secreted by cells, including the total amount of ALP and amount of ALP secreted by a single iPSC-MSC, were undertaken. The results are shown in Figure 8. The addition of HAp resulted in an increase in the expression of ALP in iPSC-MSCs. The addition of NAC also promoted the expression of ALP in iPSC-MSCs, which indicated that it significantly enhanced the osteogenic differentiation of iPSC-MSCs.

iPSC-MSCs were seeded on composite fibers. After 14 days of culture, collagen was quantitatively examined using a testing kit (Figure 9). The detection results were similar to the results of cellular ALP expression. that is, the loading of NAC promoted not only the expression of total cell COL, but also the expression of COL in a single cell. The total COL expression of all cells and expression of COL in single cells on NAC/HAp/SF fibers were significantly higher than those in the other groups. This indicates that the NAC/HAp/SF composite nanofibers had the highest ability to promote collagen production. Thus, the loading of NAC can effectively improve the osteogenic differentiation performance of the fibers.

To verify the capability of the biomimetic scaffold of NAC/HAp/SF to regenerate bone in vivo, SF, HAp/SF, NAC/SF, and NAC/HAp/SF scaffolds were transplanted into calvarial defects in SD rats. After 8 weeks of implantation, CT images demonstrated that the whole defect was almost fully repaired by the formation of bone-like tissues in the NAC/HAp/SF group (Figure 10). Moreover, the bone mineral density was significantly increased (p < 0.01), peaking at 2.51, 1.18, 1.15, and 1.14 times those of the blank SF, HAp/SF, NAC/SF, and NAC/HAp/SF groups, respectively (Figure 10F). Notably, the NAC/HAp/SF scaffold without cells exhibited the highest capacity for repair (Figures 10A–D) and highest bone mineral density compared to the other groups (Figure 10F). The bone mineral density was higher in the NAC/SF group than in the HAp/SF group, but not significantly. The NAC/HAp/SF led to the best outcomes in regenerating cranial bone defects in rats.