Biomimetic Silk Scaffolds with an Amorphous Structure for Soft Tissue

Research Article Cite This: ACS Appl. Mater. Interfaces 2018, 10, 9290−9300

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Biomimetic Silk Scaffolds with an Amorphous Structure for Soft Tissue Engineering Yonghuan Sang,†,‡ Meirong Li,§ Jiejie Liu,§ Yuling Yao,∥ Zhaozhao Ding,† Lili Wang,† Liying Xiao,†,‡ Qiang Lu,*,†,‡ Xiaobing Fu,*,§ and David L. Kaplan⊥ †

National Engineering Laboratory for Modern Silk & Collaborative Innovation Center of Suzhou Nano Science and Technology, Key Laboratory of Stem Cells and Biomedical Materials of Jiangsu Province and Chinese Ministry of Science and Technology, and ∥ School of Biology and Basic Medical Sciences, Soochow University, Suzhou 215123, People’s Republic of China § Healing and Cell Biology Laboratory, Institute of Basic Medicine Science, Chinese PLA General Hospital, Beijing 100853, People’s Republic of China ⊥ Department of Biomedical Engineering, Tufts University, Medford, Massachusetts 02155, United States ‡

S Supporting Information *

ABSTRACT: Fine tuning physical cues of silk fibroin (SF) biomaterials to match specific requirements for different soft tissues would be advantageous. Here, amorphous SF nanofibers were used to fabricate scaffolds with better hierarchical extracellular matrix (ECM) mimetic microstructures than previous silk scaffolds. Kinetic control was introduced into the scaffold forming process, resulting in the direct production of water-stable scaffolds with tunable secondary structures and thus mechanical properties. These biomaterials remained with amorphous structures, offering softer properties than prior scaffolds. The fine mechanical tunability of these systems provides a feasible way to optimize physical cues for improved cell proliferation and enhanced neovascularization in vivo. Multiple physical cues, such as partly ECM mimetic structures and optimized stiffness, provided suitable microenvironments for tissue ingrowth, suggesting the possibility of actively designing bioactive SF biomaterials. These systems suggest a promising strategy to develop novel SF biomaterials for soft tissue repair and regenerative medicine. KEYWORDS: silk, biomimetic, soft tissue regeneration, nanofibers, mechanical cues ity.13−19 Different fabrication strategies including lyophilization, salt-leaching, and electrospinning processes have been developed to prepare three-dimensional matrices used in tissue regeneration.20−25 Although many studies have confirmed the feasibility of SF matrices for repairing soft tissues such as skin, muscle, or neural tissues,26−32 the present strategies face challenges in fabricating scaffolds with appropriate microstructures and mechanical properties. The scaffolds often show stiffer mechanical properties for soft tissues and also lack ECMbiomimetic nanofibrous−microporous structures.33 These features are important in order for SF scaffolds to actively direct cell fate to achieve optimized regeneration in soft tissue engineering. Recently, we developed a self-assembly mechanism to regulate the conformations and nanostructures of SF.34 The optimization of SF conformational compositions led to the

1. INTRODUCTION Biomaterial scaffolds are faced with increasingly stringent requirements in tissue engineering and regenerative medicine. They not only must serve a role as matrices for tissue regeneration but also provide suitable microenvironments with controllable biophysical and biochemical cues for tuning cellular behavior, extracellular matrix (ECM) production, and tissue reconstruction.1,2 Soft tissues, including skin, muscle, and nerve, are easily damaged because of traumatic injuries and tumor resection.3 Unlike hard tissues, the regeneration of soft tissues requires biomaterials with pliable properties to match the defect.4 Interconnected porous structures, ECM-like nanofibrous morphologies, as well as vascularization capacity, are also preferred for better tissue formation.5−9 Although various biomaterials have been used in soft tissue regeneration,10−12 there is still a strong demand for novel scaffolds with the abovementioned multiple cue capabilities and tunability. Silk fibroin (SF) has been considered as a promising natural protein biomaterial because of the cytocompatibility, biodegradability, impressive mechanical properties, and tunabil© 2018 American Chemical Society

Received: December 18, 2017 Accepted: February 27, 2018 Published: February 27, 2018 9290

DOI: 10.1021/acsami.7b19204 ACS Appl. Mater. Interfaces 2018, 10, 9290−9300

Research Article

ACS Applied Materials & Interfaces

Cu Kα radiation (30 mA, 40 kV) with a scanning speed of 6° min−1. The 2θ range was from 5° to 45°. The thermal properties of the scaffolds were evaluated in a Q600 thermal gravimetric/differential scanning calorimetry (DSC) instrument (TA Company, New Castle, DE). Before investigation, indium was used to calibrate heat flow and temperature of the instrument while the heat capacity was calibrated with aluminum and sapphire reference standards. Standard mode DSC curves were obtained at a heating rate of 2 °C min−1 under a nitrogen gas flow of 50 mL min−1. 2.4. Properties of the Scaffolds. Silk scaffolds were cultured with phosphate-buffered saline (PBS) solution in 50 mL tubes. The weight ratio of scaffold/water was kept at 1:99. The samples were placed at 37 °C for 1, 4, 7, 14, 21, and 28 days and then dried at 60 °C and weighed. The residual weight was obtained and divided by the initial weight to achieve residual mass (%). Five samples were supplied for each measurement. The scaffolds were further immersed in protease XIV solution (1 U mL−1) in PBS to assess degradation. At a scaffold/ solution weight ratio of 1:99, samples were soaked in protease XIV solution at 37 °C for 0.5, 2, 4, 8, 12, and 24 h. After being dried at 60 °C, the samples were weighted. The residual weight was obtained and divided by the initial weight to achieve the residual mass (%). Each group had five samples. The mechanical properties of the scaffolds were evaluated in hydrated conditions according to previous studies.40 Before measurement, the samples (10 mm in diameter and 5 mm in height) were hydrated in distilled water for 4 h. Then, an Instron 3366 testing frame (Instron, Norwood, MA) was used to compress the samples with a 10 N loading cell at 25 °C. The cross head speed was maintained at 2 mm min−1 until the compression deformation of the samples was above 30%. The linear elastic region was cut from the stress−strain curves to calculate the compressive modulus of the samples. Five scaffolds were measured for each group. 2.5. In Vitro Cytocompatibility of Scaffolds. The in vitro cytocompatibility of the samples was assessed with bone marrow mesenchymal stem cells (BMSCs). The BMSCs were extracted from Sprague−Dawley (SD) rats, and their application was approved by the animal ethics committee of Soochow University. Different scaffolds were cut into small disks with a diameter of 8 mm and a height of 2 mm, which is suitable for 96-well plates. After sterilization with 60Co γirradiation at the dose of 50 kGy, the disks were used to culture BMSCs. Dulbecco’s modified Eagle medium (low glucose) supplemented with 10% fetal bovine serum and 1% IU mL−1 streptomycin− penicillin (Invitrogen, Carlsbad, CA) was used to culture BMSCs in Petri dish. When the cell confluence was 80−90%, the cells (density 1.0 × 105/well) were detached and seeded into the 96-well plates containing scaffolds. Confocal microscopy was used to investigate the morphology of the cells on the scaffolds. When cultured for 1, 6, and 12 days, the scaffolds seeded with the cells were fixed by washing three times with PBS, immersing in 4% paraformaldehyde (Sigma-Aldrich, St. Louis, MO) for 30 min, and finally washing three times with PBS again. After permeabilized with 0.1% Triton X-100 for 30 min, the scaffolds seeded with the cells were stained through successively incubating in FITCphalloidin solvent (Sigma-Aldrich, St. Louis, MO) for 1 h, washing with PBS, and incubating with 4′,6-diamidino-2-phenylindole (DAPI, Sigma-Aldrich, St. Louis, MO) for 5−10 min. The stained samples were observed with confocal laser scanning microscopy (CLSM, Olympus FV10 inverted microscope, Nagano, Japan). The excitation/ emission wavelengths were 358/462 nm and 494/518 nm, respectively. Representative fluorescence images were obtained through scanning the samples from the surface to a depth of 100 μm in increments of 10 μm. The cell proliferation behaviors on the scaffolds were evaluated by the PicoGreen DNA assay (Invitrogen, Carlsbad, CA). At predetermined time points (1, 3, 6, 9, and 12 d), the samples were gathered and incubated in proteinase K buffer solution overnight at 56 °C to degrade the scaffolds.46 The samples (n = 5) were investigated by a BioTeK Synergy 4 spectrofluorometer (BioTeK, Winooski, VT) with excitation/emission at 480/530 nm, respectively. λ-Phage DNA was diluted to gradient solution to obtain the standard curve. Then,

formation of water-insoluble scaffolds with softer mechanical properties.11,22,35 The softer scaffolds exhibited vascularization capacity, suggesting the possibility of designing bioactive SF scaffolds without the addition of growth factors. SF nanofibers, mainly composed of β-sheet structures, were also prepared and used to fabricate scaffolds and hydrogels with hierarchical ECM biomimetic microstructures, further improving the biocompatibility of SF materials.36 However, further improvement is desirable for the SF materials used in soft tissue engineering. Recently, we fabricated amorphous SF nanofibers in aqueous solution.37 The nanofibers are composed of metastable intermediate conformations of the protein, endowing the possibility of forming tunable and softer scaffolds. Here, using the amorphous nanofibers as motifs, a mild strategy was developed to control secondary structures of SF scaffolds. Through tuning the freezing temperature of the nanofiber solution to control the kinetics of higher order structure formation from these metastable solutions, water-insoluble silk matrices with a lower crystal structure and softer features were generated. These new scaffolds exhibited highly tunable stiffness in the range of 1−5 kPa, which are significantly softer than SF scaffolds reported previously.19,38−43 Hierarchical nanofibrous−microporous structures provided useful microenvironments for cell proliferation and tissue regeneration. This kinetic-induced process provides options to generate tunable SF scaffold systems for soft tissue needs.

2. EXPERIMENTAL SECTION 2.1. Amorphous Silk Nanofiber Solution. The aqueous silk solutions composed of amorphous nanofibers were prepared according to our previous work.37 Bombyx mori silk fibers were boiled for 30 min in 0.02 M Na2CO3 aqueous solution and rinsed three times with water to remove sericin proteins. The degummed silk was added into a mixture solvent of lithium bromide (LiBr, 8.0 M) and formic acid (98%) with a volume ratio of 1:23.2 and incubated at 60 °C for 4 h to dissolve the silk fibers. After dialysis with deionized water for 3 days at 4 °C and centrifugation at 9000 rpm for 20 min, the amorphous silk nanofiber solution (0.8 wt %) was prepared. This solution was then concentrated to 2 wt % for further use. As a control, traditional silk solution was also prepared through procedures reported previously.44 2.2. Amorphous Silk Porous Scaffolds. The silk scaffolds mainly composed of the amorphous state were prepared through tuning the rate of silk nanofiber assembly during the lyophilization process. Amorphous silk nanofiber solution was frozen at various temperatures from −20 to −5 °C for 24 h and lyophilized for 72 h. When the freezing temperature was above −9 °C, the insoluble scaffolds were directly prepared after lyophilization without further treatment. The insoluble scaffolds were termed as NSF-9, NSF-7, and NSF-5 according to their freezing temperature. As the control group, amorphous silk nanofiber solution and traditional silk solution were frozen at −20 °C for 24 h and freeze-dried for 72 h to prepare soluble scaffolds. The scaffolds were immersed in methanol for 1 h to achieve insolubility and termed SF-MA and NSF-MA, respectively. 2.3. Characterization of the Scaffolds. The microstructure of the scaffolds was measured with scanning electron microscopy (SEM, Hitachi S-4800, Hitachi, Tokyo, Japan) at 3 kV. Before investigation, the samples were treated with platinum. The secondary structures of the scaffolds were analyzed with Fourier transform infrared spectroscopy (FTIR, Nicolet FTIR 5700, Thermo Scientific, FL, USA). The FTIR spectra were obtained in the wavenumber range of 400−4000 cm−1. Sixty-four scans were used under a resolution of 4 cm−1. PeakFit software was used to achieve Fourier self-deconvolution (FSD) of the amide I region (1595−1705 cm−1) for quantitative analysis of secondary structures of the scaffolds.45 The crystalline structures of the scaffolds were determined with X-ray diffraction (XRD) (X’PertPro MPD, PANalytical B.V., Almelo, Holland) using monochromated 9291


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ACS Applied Materials & Interfaces the DNA content of the samples was calculated based on the standard curve. To measure possible differentiation capacity of the scaffolds, quantitative real-time polymerase chain reaction (RT-PCR) and immunofluorescence staining were used to investigate endothelial gene expression of the cells cultured on the scaffolds. On the basis of the protocol of manufacturers, when the cellular RNA was extracted from the samples with TRIzol reagent (Takara), a cDNA Synthesis Kit (Takara) was used to reverse-transcribe RNA into cDNA. The targeted gene primers shown in Table 1 were used, and the results were obtained according to the same processes reported in our previous study.40

3. RESULTS AND DISCUSSION 3.1. Scaffold Structure Characterization. The selfassembly mechanism for material formation provides an effective strategy to tune SF nanostructures and conformations.50,51 The development of different silk materials, such as amorphous but water-insoluble scaffolds, is changing traditional opinions about SF materials, opening ways to match the SF structure to specific application. Amorphous and metastable SF nanofibers fabricated in aqueous solution widened the functional tunability of SF biomaterials (Figure S1).52 These waterinsoluble SF scaffolds with biomimetic morphologies were achieved by controlling assembly kinetics from amorphous nanofiber building blocks during the scaffold forming process. The freezing temperature was used as the variable to control the kinetics of amorphous SF nanofiber assembly, within the range of −20 to −5 °C (Scheme 1). When the freezing

Table 1. Sequences of Primers Used in RT-PCR genes product

primer sequence (F, R, 5′-3′)

length (bp)

GAPDH

TGGGTGTGAACCACGAGAA GGCATGGACTGTGGTCATGA CGGAGAAATCTGCTCGCTAT CTTGGAAGGGACGACACG

143

Flt-1

190

Scheme 1. Process of Formation of Water-Insoluble SF Scaffolds with Mainly Amorphous (Noncrystalline) States

Endothelial differentiation behaviors of BMSCs were characterized with immunofluorescence staining of cells with the von willebrand factor (VWF, endothelial cell marker).47 After being fixed with 4% paraformaldehyde solution (Sigma-Aldrich, St. Louis, MO, USA), the samples were incubated with the desired primary antibodies against the VWF (Abcam, Cambridge, MA, USA) and secondary antibodies successively according to the reported schedule.40 When the cell nucleus was counter-stained with Hoechst dye (Sigma, St. Louis, MO, USA) for 5 min, the samples were measured and imaged with CLSM (Olympus FV10 inverted microscope, Nagano, Japan). The PCR products were confirmed by agarose gel electrophoresis. PCR products were analyzed by agarose gel electrophoresis on a 1.0% agarose gel in TAE buffer. The agarose gel was stained by ethidium bromide. Electrophoresis was performed at 100 V for 50 min. 2.6. In Vivo Biocompatibility of the Scaffolds. The in vivo biocompatibility of the scaffolds was evaluated with a proven subcutaneous implantation model in our group.40 The use of SD rats was approved by the animal ethics committee of Soochow University. After sterilization and immersion in PBS for hours, the scaffolds with the same size of 10 × 10 × 3 mm were implanted into lateral incisions on the dorsal region. Animals were euthanized after implantation for 1, 2, 3, and 4 weeks. The specimens along with the adjacent tissues were gathered for the following investigation. The specimen sections (5−6 μm thickness) were prepared via regular fixation, embedment and section processes.40 The sections were deparaffinized and stained with hematoxylin and eosin (SigmaAldrich, St. Louis, MO, USA) and also probed by primary and secondary antibodies against the endothelial cell marker, CD34 (1:100 dilution, Abcam, MA, USA). After counterstained with hematoxylin (Histostain-SP kit, Invitrogen, MA, USA), the sections were visualized with an inverted microscope (Axio Vert A1, Carl Zeiss, Germany) to characterize lumen formation.48 The formed vessels inside the scaffolds were evaluated by counting above six view fields (under 20× magnification) randomly of the various sections from three individual rats. The vessel density was calculated according to the previous method.49 Masson trichrome staining was used to indicate collagen deposition. After the regular staining process,40 the stained sections were observed with an inverted microscope (Axio Vert A1, Carl Zeiss, Germany). 2.7. Statistical Methods. SPSS v.16.0 software was used in all statistical analyses. One-way analysis of variance was performed to compare the significance of the values. Data are presented as means ± standard deviations. Unless otherwise specified, P < 0.05 was considered significant.

temperature was above −9 °C, amorphous but water insoluble SF scaffolds were prepared via traditional lyophilization processes (Figure S2). Figure 1 shows the typical microstructures of the scaffolds prepared under the various freezing temperatures. Unlike the scaffolds derived from traditional SF solution (Figure 1, SF-MA), all of the SF nanofiber-based scaffolds showed interconnected porous structures and maintained nanofibrous topography on the pore surface, indicating the formation of hierarchical nanofibrous−microporous structures (Figure 1). Because of various rates of ice crystal growth under different temperatures, thicker porous layers appeared for the scaffolds prepared at higher temperatures (Figure 1b). Similar repeated morphological changes were achieved in our study, suggesting the reproducibility of the scaffolds. A number of studies have confirmed the favorable influence of similar ECM nanofibrous structures on cellular performance.53−56 The formation of better hierarchical ECM mimetic structures for the scaffolds could supply preferred cues for adhesion and growth of the cells. Structural changes of the SF scaffolds were measured with FTIR. The secondary structures of SF are usually assessed based on the absorption of amide I regions where the peaks of silk II conformation appear at 1610−1630 cm−1, while the peaks at 1635−1652 cm−1 belong to amorphous states.57−60 As shown in Figure 2a, the methanol-treated scaffolds (SF-MA and NSF-MA) showed a main peak at 1625 cm−1, while all the temperature-induced scaffolds (NSF-9, NSF-7, and NSF-5) had 9292


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to the FTIR results, the degradation peak at higher temperatures was achieved for the scaffolds prepared at higher freezing temperatures, further suggesting the conformational tunability for the scaffolds. The FTIR, XRD, and DSC results indicated the formation of water-insoluble SF scaffolds with tunable amorphous structures. 3.2. Scaffold Properties. Various degradation behaviors were achieved through changing the secondary compositions. Compared to the MA scaffolds with higher β-sheet content, all the temperature-induced scaffolds with amorphous states showed faster degradation after exposure to PBS and protease XIV solutions (Figure 3a,b). The tunable degradation behavior also appeared for the temperature-induced scaffolds, where gradually slower degradation rates were found for the scaffolds prepared at higher freezing temperatures. For example, when cultured in protease XIV solution for 24 h, the MA scaffolds remained above 85% of their original weight, while the temperature-induced scaffolds (NSF-9, NSF-7, and NSF-5) lost about 80, 70, and 50% of the original weight (Figure 3b), respectively. As expected, the temperature-induced scaffolds also showed tunable and lower stiffness because of the amorphous structures. The compressive modulus was 3.5, 2.7, and 1.5 kPa for NSF-5, NSF-7, and NSF-9 scaffolds, respectively (Figure 3c), which are significantly lower than those of MA scaffolds and acid-assisted amorphous scaffolds.40 SF scaffolds with vascularization capacity have been prepared through tuning the mechanical cues of the scaffolds in the acid-assisted lyophilization process. However, fine regulation of stiffness in the range of 1−7 kPa is impracticable for the process. Here, temperature-induced lyophilization was used to regulate the assembly rate of amorphous SF nanofibers, achieving subtle adjustments of the modulus between 1 and 4 kPa. Therefore, the present method introduces new ECM mimetic microstructures into silk scaffolds, while also enriching the mechanical features of the scaffolds for soft tissue engineering. The elastic properties of these scaffolds were then investigated, and they showed better elasticity than traditional methanoltreated scaffolds (Figure S4 and Table S2). The extension at break was 53.1, 62.7, and 69.1% in the wet state for NSF-5, NSF-7, and NSF-9 scaffolds, respectively, but decreased to 35.1% for methanol-treated scaffolds, suggesting their suitability in soft tissue regeneration. 3.3. In Vitro Cytocompatibility. In vitro cytocompatibility of the temperature-induced scaffolds was evaluated using BMSCs. The BMSCs showed persistent proliferation on all the SF scaffolds without reaching a plateau in the culture period of 12 days, suggesting cell compatibility (Figures 4 and S5). Various microstructures and conformations of the scaffolds resulted in various cell behaviors.8,61−63 Compared to methanol-treated scaffolds with few nanofibrous structures, BMSCs grew significantly better on methanol-treated scaffolds composed of nanofibers. The results confirmed that the nanofibrous structures provided favorable physical cues for cell growth. The cytocompatibility of SF scaffolds was also influenced by secondary structures. When the scaffolds were prepared from the same amorphous SF nanofiber solution, DNA content suggested that the cells proliferated significantly better on the scaffolds composed of amorphous states than β sheet-rich structures (NSF-MA). Further improvement of cell proliferation could be achieved through tuning the amorphous structures. Although all of the scaffolds mainly composed of amorphous states showed better cytocompatibility than those

Figure 1. SEM morphologies of SF scaffolds: (a) microporous structures of the scaffolds; (b) thickness of the porous wall for the different scaffolds; (c) high magnification of the topography of the pore wall at the nanoscale. The samples were as follows: SF-MA, methanol-treated scaffolds derived from traditional silk solution; NSFMA, methanol-treated scaffolds derived from amorphous silk nanofiber solution; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C.

a chief peak at 1640−1650 cm−1. The results indicated that all the water-insoluble temperature-induced scaffolds mainly composed of amorphous states, which is similar to the acidassisted scaffolds reported previously.40 Deconvolution (amide I region) was then used to further analyze the secondary structure (Figure S3 and Table S1). The silk II content of SFMA scaffolds was 43%, which decreased to below 30% for all of the temperature-induced scaffolds. Following the freezing temperature increases from −9 to −5 °C, the β-sheet content increased slightly from 24.9 to 25.2% and 27.8%, suggesting the tunability of the conformations for SF nanofiber scaffolds. XRD curves also suggested the amorphous states in the temperatureinduced scaffolds (Figure 2b). Unlike the SF-MA and NSF-MA scaffolds, the temperature-induced scaffolds showed broader peaks between 10° and 40°, confirming the lower crystalline structure. The conformational changes of the temperatureinduced scaffolds were also evaluated by DSC (Figure 2c). All the samples had an endothermic peak at 50−100 °C and a degradation peak at 250−280 °C without the nonisothermal crystallization peak at 200−230 °C that is usually found in previous amorphous SF scaffolds.34 The results suggested that more stable amorphous structures formed for the temperatureinduced scaffolds, which could restrain the crystallization transformation at high temperatures. Compared to SF-MA and NSF-MA scaffolds, the degradation peak appeared at lower temperatures for all the temperature-induced scaffolds, confirming the existence of the less β-sheet structure. Similar 9293


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Figure 2. FTIR (a), XRD (b), and DSC (c) curves of different SF scaffolds. The samples were as follows: SF-MA, methanol-treated scaffolds derived from traditional silk solution; NSF-MA, methanol-treated scaffolds derived from amorphous silk nanofiber solution; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C.

Figure 3. The degradation behaviors and modulus of the different SF scaffolds: (a) degradation behaviors in PBS solution, (b) degradation behaviors in protease XIV solution (1 U mL−1), and (c) compressive modulus of the scaffolds in the wet state. The samples were as follows: SF-MA, methanoltreated scaffolds derived from traditional silk solution; NSF-MA, methanol-treated scaffolds derived from amorphous silk nanofiber solution; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C. * P ≤ 0.05,** P ≤ 0.01, *** P ≤ 0.001.

with rich β-sheet structures, the best cell proliferation appeared on the NSF-5 scaffolds that contained the highest β-sheet content among the amorphous scaffolds. Several previous studies have suggested that more amorphous states of SF

scaffolds facilitate cell proliferation.38,40,65 Considering that the SF scaffolds prepared previously had significantly higher content of β-sheet structures, our present results indicated that suitable secondary structures rather than lower β-sheet 9294


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Figure 4. BMSC proliferation on various SF scaffolds: (a) fluorescence microscopy images of BMSCs cultured on the scaffolds at days 1, 6, and 12; nuclei and silk scaffolds were stained blue (DAPI); F-actin was stained green (FITC-labeled phalloidin); (b) BMSC proliferation on the samples measured with DNA analysis. The samples were as follows: SF-MA, SF scaffold was treated by methanol; NSF-MA, amorphous SF nanofiber scaffold was treated by methanol; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C. * P < 0.05,** P < 0.01, *** P < 0.001.

optimize suitable mechanical factors for inducing endothelial differentiation on the silk scaffolds, the present system provided an appropriate material platform to evaluate the function of mechanical cues on the cell behavior. 3.4. In Vivo Biocompatibility. A suitable niche is essential for better vascularization and tissue regeneration in vivo. Various chemical and physical cues including growth factors, nanostructures, and mechanical properties have been reported to promote vascularization and tissue regeneration.10,64,65 To further evaluate the effects of multiple physical cues on vascularization and tissue regeneration in vivo, the temperatureinduced scaffolds with ECM mimetic structures and different stiffness were implanted subcutaneously in rats. Two methanoltreated scaffolds were used as control. The histology of tissue sections and immunohistochemistry analysis for the implanted scaffolds are shown in Figures 6 and 7. Significantly better vasculature in the scaffolds appeared in the different temperature-induced scaffolds at 28 d after implantation but not in the methanol-treated scaffolds, suggesting accelerated neovascularization. The different temperature-induced scaffolds also exhibited different vascularization capacity where the most extensive vasculature was found in the NSF-5 scaffolds. Figure 7 shows the quantified number of vessels inside the scaffolds. After 4 weeks, about 39 ± 2.7 vessels per mm2 formed in the methanol-treated scaffolds, which increased to 49 ± 3.7, 52 ± 1.4, and 60 ± 5.6 respectively in NSF-9, NSF-7, and NSF-5, respectively, confirming better vascularization capacity for the

structures provided a preferable microenvironment for cell proliferation. Confocal microscopy results confirmed the better cell growth on the nanofibrous−microporous scaffolds with amorphous structures (Figure 4). Therefore, the cell studies in vitro suggested that the nanofibrous SF scaffolds provided multiple stimulating cues (secondary structures and microstructures) for better cell responses. Because previous studies suggested that matrix stiffness in the range of 1−7 kPa could induce the differentiation of stem cells into endothelial cells,66 the endothelial differentiation of BMSCs on the various scaffolds was evaluated with immunofluorescent staining for the VWF (Figure 5). The NSF-9 scaffolds were destroyed in the cell culture process because of their rapid degradation, resulting in the failure to obtain information on cell differentiation. Both the immunofluorescent staining and the agarose gel electrophoresis results of vascular endothelial growth factor receptor-1 (Flt-1) showed similar VWF expression for the cells cultured on temperatureinduced scaffolds (NSF-7 and NSF-5) and methanol-treated scaffolds (SF-MA and NSF-MA), indicating that the temperature-induced silk scaffolds did not achieve improved endothelial differentiation capacity. Considering that most of previous studies were obtained from hydrogels rather than dense scaffold states and failed to finely tune the stiffness of the materials,66−68 it is reasonable that the silk scaffolds with a stiffness of 1−4 kPa exhibit a differentiation capacity that is different than expected. Although further study is necessary to 9295


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Figure 5. Differentiation capacity into endothelial cells of BMSCs on various SF scaffolds: (a) expression of the VWF (red) in different samples at day 14 by confocal microscopy, blue (Hoechst) for nuclei and SF; (b) expression of Flt-1 at day 14 detected by agarose gel electrophoresis. The samples were as follows: 1, positive control for endothelial cell; 2, mesenchymal stem cells were induced by adding the vascular endothelial growth factor; SF-MA, SF scaffold was treated by methanol; NSF-MA, amorphous SF nanofiber scaffold was treated by methanol; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C. No significant difference of BMSC differentiation behaviors was found for all the scaffolds.

Figure 6. HE staining photography of sections of different sample implants at 7, 14, 21, and 28 days after implantation in vivo. The samples were as follows: SF-MA, SF scaffold was treated by methanol; NSF-MA, amorphous SF nanofiber scaffold was treated by methanol; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C. Better neovascularization and tissue ingrowth appeared in temperature-induced scaffolds (scale bar = 200 μm). The protoplasm was stained with picrosirius red, and nuclei were stained with blue. The arrows indicated the newly formed vessels.

The influence of vascularization on tissue regeneration was evaluated (Figures 6 and 8). Similar to the vasculature results, significantly better neogranulation tissue formation appeared inside the temperature-induced scaffolds. Twenty-eight days after implantation, only about 72% of the methanol-treated scaffolds was occupied by tissue ingrowth, while granulation tissue had overtaken about 95% of the spaces inside the

temperature-induced scaffolds. Because the temperatureinduced scaffolds had an endothelial differentiation capacity similar to the methanol-treated scaffolds, the improved vascularization in vivo is likely due to better cell proliferation and tissue ingrowth capacity of the scaffolds. Both in vitro and in vivo results indicated that the vascularization inside the silk scaffolds was affected by multiple factors. 9296


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Figure 7. (a) Immunohistochemistry staining images of different scaffold implants at 7, 14, 21, and 28 days after implantation in vivo (scale bar = 200 μm). The samples were as follows: SF-MA, SF scaffold was treated by methanol; NSF-MA, amorphous SF nanofiber scaffold was treated by methanol; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C. The microvessels were stained with dark gray positive for CD34, and nuclei were stained with blue. (b) Vessel density within the different scaffolds after implantation into the lateral incisions on the dorsal region. * P < 0.05, ** P < 0.01.

Figure 8. Masson trichrome staining images of sections of different scaffold implants at 7, 14, 21, and 28 days after implantation in vivo (scale bar = 200 μm). The samples were as follows: SF-MA, SF scaffold was treated by methanol; NSF-MA, amorphous SF nanofiber scaffold was treated by methanol; NSF-9, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −9 °C; NSF-7, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −7 °C; and NSF-5, insoluble scaffolds derived from amorphous silk nanofiber solution and frozen at −5 °C. Collagen fibers are stained blue, nuclei are stained dark red/purple, and cytoplasm is stained pink.

different temperature-induced scaffolds. The results confirmed that optimization of vascularization capacity for the SF scaffolds could promote tissue regeneration. Interestingly, the two methanol-treated scaffolds with similar stiffness and various microstructures exhibited similar vascularization but different

tissue regeneration. Significantly more granulation tissue was observed inside the NSF-MA scaffolds with nanofibrous structures, suggesting the capacity of microstructures in inducing neotissue formation. Various tissue regeneration behaviors also appeared on the different temperature-induced 9297


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ACS Applied Materials & Interfaces Author Contributions

scaffolds. Although most of the spaces inside all the scaffolds were occupied by neogranulation tissues after 28 d, best tissue ingrowth was achieved inside the NSF-9 scaffolds at 14 d and then appeared inside the NSF-5 scaffolds at 28 d. Considering that various degradation behaviors of the scaffolds, quicker degradation of the NSF-9 scaffolds facilitated the tissue ingrowth in the first 2 weeks but failed to provide better matrices after 4 weeks than that of the NSF-5 scaffolds. The collagen deposition behaviors inside the scaffolds also showed a similar tendency to tissue ingrowth. Although more collagen deposited on the NSF-9 and NSF-7 scaffolds at 14 d, best collagen deposition happened inside the NSF-5 scaffolds after 28 d. The results implied that the various degradation behaviors of the scaffolds had significant influence on tissue regeneration. More interestingly, unlike traditional silk scaffolds, the present scaffolds achieved tunable and faster degradation, as well as improved in vivo vascularization behaviors, implying their promising application in absorbable soft tissue replacement. Therefore, our findings indicated that the amorphous SF scaffolds provided multiple physical signals that optimized vascular ingrowth and tissue regeneration.

Y.S. and M.L. have contributed equally to the first author. Q.L., X.F., and D.L.K. designed the study. Y.S., M.L., J.L., Y.Y., and Z.D. performed the experiments. Q.L., L.W., and L.X. analyzed the data. Y.S., Q.L., M.L., and D.L.K. wrote the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the National Key Research and Development Program of China, (2016YFE0204400, 2017YFC110304, and 2017YFC1104701), the National Natural Science Foundation of China (81721092), the NIH (R01NS094218, R01AR070975), and the AFOSR. They also thank the second affiliated hospital of Soochow University preponderant clinic discipline group project funding (no. XKQ2015010) for support of this work.

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4. CONCLUSIONS SF scaffolds with tunable stiffness and ECM mimetic microstructures were developed through regulating the assembly rates of amorphous silk nanofibers in a lyophilization process. The scaffolds had hierarchical nanofibrous−microporous structures similar to the ECM and also showed tunable stiffness to provide optimized physical cues for vascularization and tissue regeneration. The optimization of cell proliferation capacity in vitro and the enhancement of neovascularization capacity and tissue regeneration in vivo suggest a promising future for soft tissue engineering. The kinetic control of the SF assembly process provides a mild and feasible strategy of actively designing scaffolds with multiple cues, while the mild all-aqueous processing further enriches the possibility of the incorporation of other bioactive molecules that are otherwise heat labile, increasing their adaptability in various biomedical applications.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.7b19204. SEM and FTIR of amorphous silk nanofiber solutions; the stability of the amorphous silk nanofiber scaffolds with various freezing temperatures before and after water immersion for 4 h at 37 °C; FSD of FTIR spectra was obtained through PeakFit software to quantitatively analyze secondary structures of the scaffolds; FTIR determination of secondary structures of different silk scaffolds through FSD of the amide I region; and the elasticity of the scaffolds in the dry and wet states (PDF)

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REFERENCES

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. Phone: (+86)-512-67061649 (Q.L.). *E-mail: [email protected] (X.F.). ORCID

Qiang Lu: 0000-0003-4889-5299 David L. Kaplan: 0000-0002-9245-7774 9298


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Biomimetic Silk Scaffolds with an Amorphous Structure for Soft Tissue

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