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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 ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b19204 • Publication Date (Web): 27 Feb 2018 Downloaded from http://pubs.acs.org on February 27, 2018

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ACS Applied Materials & Interfaces

Biomimetic Silk Scaffolds with an Amorphous Structure for Soft Tissue Engineering Yonghuan Sanga,b,#, Meirong Lic,#, Jiejie Liuc, Yuling Yaod, Zhaozhao Dinga, Lili Wanga, Liying Xiaoa,b, Qiang Lua,b,*, Xiaobing Fuc,*, David L Kaplane

a

National Engineering Laboratory for Modern Silk & Collaborative Innovation Center of Suzhou

Nano Science and Technology, Soochow University, Suzhou 215123, People’s Republic of China b

Key Laboratory of Stem Cells and Biomedical Materials of Jiangsu Province and Chinese Ministry

of Science and Technology, Soochow University, Suzhou 215123, People’s Republic of China c

Healing and Cell Biology Laboratory, Institute of Basic Medicine Science, Chinese PLA General

Hospital, Beijing 100853, People’s Republic of China d

School of Biology and Basic Medical Sciences, Soochow University, Suzhou 215123, People’s

Republic of China e

Department of Biomedical Engineering, Tufts University, Medford, Massachusetts 02155, United

States

# The authors have contributed equally to the first author

Corresponding author: Qiang Lu, Tel: (+86)-512-67061649; E-mail: [email protected]

Xiaobing Fu, E-mail: [email protected]

Keywords: Silk; Biomimetic; Soft Tissue Regeneration; Nanofibers; Mechanical Cues

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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.

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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 to tune cellular behavior, extracellular matrix production and tissue reconstruction.1,2 Soft tissues, including skin, muscle, and nerve, are easily damaged due to 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 strong demands remain for novel scaffolds with the above multiple cue capabilities and tunability.

Silk fibroin (SF) has been considered as a promising natural protein biomaterial due to the cytocompatibility, biodegradability, impressive mechanical properties and tunability.13-19 Different fabrication strategies including lyophilization, salt-leaching, and electrospinning processes have been developed to prepare three-dimensional (3D) 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 ECM-biomimetic 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. 3



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 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 composted of beta-sheet structure 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 lower crystal structure and softer features were generated. These new scaffolds exhibited highly tunable stiffness in the range of 1-5 kPa, 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

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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 minutes 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 (FA, 98%) with a volume ratio of 1 : 23.2 and incubated at 60oC for 4 h to dissolve the silk fibers. After dialysis with deionized water for 3 days at 4oC and centrifugation at 9,000 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 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 -20oC to -5oC for 24 h and lyophilized for 72 h. When the freezing temperature was above -9oC, the insoluble scaffolds were directly prepared after lyophilizaiton without further treatment. The insoluble scaffolds were termed as NSF-9, NSF-7 and NSF-5 according to their freezing temperature. As control group, amorphous silk nanofiber solution and traditional silk solution were frozen at -20oC 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

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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. 64 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 XRD (X Pert-Pro MPD, PANalytical BV, Almelo, Holland) using monochromated Cu Kα radiation (30 mA, 40 kV) with a scanning speed of 6o min-1. The 2θ range was from 5o to 45o.

The thermal properties of the scaffolds were evaluated in a Q600 Thermal Gravimetric/Differential Scanning Calorimery (TGA/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 2oC min-1 under nitrogen gas flow of 50 ml min-1.

2.4 Properties of the Scaffolds Silk scaffolds were cultured with PBS solution in 50 ml tubes. The weight ratio of scaffold/water was kept at 1:99. The samples were placed at 37oC for 1, 4, 7, 14, 21 and 28 days, and then dried at 60oC 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

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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 37oC for 0.5, 2, 4, 8, 12, and 24 h. After dried at 60oC, 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 25oC. The cross head speed kept 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 its application was approved by the animal ethics committee of Soochow University. Different scaffolds were cut into small disks with diameter of 8 mm and height of 2 mm, which is suitable for 96-well plates. After sterilization with

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Co γ-irradiation at the dose of 50 kGy, the disks were used to culture BMSCs.

Dulbecco's modified Eagle medium (DMEM, low glucose) supplemented with 10% fetal bovine serum (FBS), 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

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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 FITC-phalloidin solvent (Sigma-Aldrich, St. Louis, MO) for 1 h, washing with PBS, and incubating with DAPI (Sigma-Aldrich, St. Louis, MO) for 5-10 min. The stained samples were observed with a 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 100µm in increments of 10µm.

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 for overnight at 56oC 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 the DNA content of the samples was calculated based on the standard curve.

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To measure possible differentiation capacity of the scaffolds, quantitative real-time PCR (qRT-PCR) and immunofluorescence staining were used to investigate endothelial gene expression of the cells cultured on the scaffolds. Based on the protocol of manufactures, when the cellular RNA was extracted from the samples with Trizol reagent (Takara), a cDNA Synthesis Kit (Takara) was used to reverse transcribed the 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 Table 1. Sequences of primers used in real-time PCR (PR-PCR). Genes Product

Primer sequence (F, R, 5′-3′)

length (bp)

TGGGTGTGAACCACGAGAA GAPDH

143 GGCATGGACTGTGGTCATGA CGGAGAAATCTGCTCGCTAT

Flt-1

190 CTTGGAAGGGACGACACG

Endothelial differentiation behaviors of BMSCs were characterized with immunofluorescence staining of cells with von willebrand factor (VWF, endothelial cell marker).47 After fixed with 4% paraformaldehyde solution (Sigma-Aldrich, St. Louis, MO, USA), the samples were incubated with the desired primary antibodies against VWF (Abcam, Cambridge, MA, USA) and secondary antibodies successively according to the reported schedule.40 When cell nucleus was counter-stained with Hoechst dye (Sigma, St. Louis, MO, USA) for 5 min, the samples were measured and imaged with a CLSM (Olympus FV10 inverted microscope, Nagano, Japan).

The PCR products were confirmed by agarose gel electrophoresis. PCR products were analyzed by 9



agarose gel electrophoresis on a 1.0% agarose gel in TAE buffer. The agarose gel was stained by ethidium bromide (EB). 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 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 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 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 (H&E) (Sigma-Aldrich, St. Louis, MO, USA), and also probed by primary and secondary antibody 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 (AxioVert 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 previous method.49 Masson trichrome staining was used to indicate collagen deposition. After regular staining process,40 the stained sections were observed with an inverted microscope (AxioVert A1, Carl Zeiss, Germany).

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2.7 Statistical Methods SPSS v.16.0 software was used in all statistical analyses. One-way AVOVA were performed to compare the significance of the values. Data are presented as means ± standard deviations. Unless otherwise specified, P

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