Research Article pubs.acs.org/journal/ascecg
Facile Preparation of Biocompatible Silk Fibroin/Cellulose Nanocomposite Films with High Mechanical Performance Yanfei Feng,† Xiufang Li,†,‡ Mingzhong Li,‡ Dezhan Ye,† Qiang Zhang,† Renchuan You,*,† and Weilin Xu*,† †
State Key Laboratory for Hubei New Textile Materials and Advanced Processing Technologies, College of Textile Science and Engineering, Wuhan Textile University, No. 1 Yangguang Avenue, Jiangxia District, Wuhan 430200, China ‡ National Engineering Laboratory for Modern Silk, College of Textile and Clothing Engineering, Soochow University, No. 199 Ren’ai Road, Industrial Park, Suzhou 215123, China ABSTRACT: Although regenerated silk fibroin (SF), which has excellent biocompatibility, biodegradability, and a low inflammatory response in vivo, has promising applications in tissue engineering, the mechanical properties and biofunctionality must be further improved to satisfy tissue-engineering applications. Cellulose nanofibrils (CNFs) are promising candidates for bionanocomposite production due to their ultrahigh strength and excellent biocompatibility. In this study, CNFs were extracted directly from microcrystalline cellulose using an aqueous lithium bromide solution, a typical solvent for dissolving SF fibers. As a result, SF/cellulose nanocomposite films with improved tensile strength were fabricated using aqueous lithium bromide solution as a novel solvent system for the dissolution and blending of SF and cellulose. The extracted CNFs were homogeneously dispersed within the composite films through the rapid gelation of cellulose. The degradability of the composite films in a protease XIV solution was strongly dependent upon the SF component, which significantly promoted the degradation rate of composite films. Adhesion and proliferation results showed that SF/cellulose nanocomposite films promoted cell viability. Our work suggests a facile and effective approach for designing SF/cellulose nanocomposites that may have wide potential applications in tissue engineering. KEYWORDS: Silk fibroin, Cellulose, Nanofibrils, Lithium bromide, Mechanical property, Biocompatibility
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INTRODUCTION Silk fibroin (SF) is an abundant and sustainable natural protein material extracted from Bombyx mori silk and holds distinctive biological properties including outstanding biocompatibility, tunable degradability, and a low inflammatory response in vivo.1−3 Recently, SF has been applied in numerous biomedical fields involving drug delivery, tissue engineering, and implantable devices.4,5 Because of their extraordinary strength, degummed SF fibers have been investigated particularly for the purposes of ligament and tendon repair.6,7 However, fiber-only materials are limited in tissue-engineering applications. After dissolving sericin-free SF fibers to obtain regenerated SF solutions in concentrated aqueous salts, typically using aqueous lithium bromide solution,2,8 we can further reform the desired material forms such as films, porous sponges, and nanofibers. However, regenerated SF matrixes are either too brittle in the dry state or too weak in the wet state, neither of which are mechanically strong enough for some tissue-engineering applications.9,10 Therefore, the mechanical properties of regenerated SF scaffolds must be further improved to satisfy the requirements of clinical applications. To improve the mechanical properties and biofunctionality of tissue-engineering scaffold materials, the use of composites © 2017 American Chemical Society
has been widely explored. Polymer blending is an effective method for offering new, desirable biocomposites. Cellulose has received increasing interest in research because it is safe, stable, nontoxic, biocompatible, renewable, low cost, and abundant in nature.11 SF/cellulose blend materials have been prepared to enhance mechanical performance.10,12,13 The cellulose/SF blend films prepared by ionic liquids are able to support the growth and proliferation of fibroblast cells.10 Singh et al. prepared cellulose/SF blend compositions using ionic liquids as a common solvent and found that the blend compositions can support mesenchymal stem cell growth and promote chondrogenic differentiation.12 Moreover, the incorporation of bioactive nanomaterials into polymeric tissue-engineering matrixes is also regarded as a promising solution for enhancing the mechanical properties and biofunctionality. The use of nanoscale reinforcements such as hydroxyapatite,14 graphene oxide,15,16 gold nanoparticle,17 carbon nanotubes,18,19 and carbon nanofibers20 as inclusions in SF matrixes have been investigated to date to increase the Received: April 17, 2017 Revised: May 26, 2017 Published: May 30, 2017 6227
DOI: 10.1021/acssuschemeng.7b01161 ACS Sustainable Chem. Eng. 2017, 5, 6227−6236
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ACS Sustainable Chemistry & Engineering
Figure 1. Dissolution of cellulose in aqueous lithium bromide solution. (a) Cellulose suspension. (b) SEM image of lyophilized CNFs. Scale bars: 1 μm.
Figure 2. Schematic representation of a new strategy for SF/cellulose nanocomposite films.
high crystallinity and complex hierarchical structure.22 Another major challenge with the preparation of CNFs/polymer nanocomposites is that CNFs are highly hydrophilic due to the nature of cellulose. As such, it is difficult to disperse CNFs uniformly in most of the common polymer matrixes.28,29 Here, we report a facile and effective approach to prepare SF/cellulose composites containing CNFs. First, we used a new strategy to directly separate microcrystalline cellulose on the single-CNF level by a partial dissolution using aqueous lithium bromide, a typical solvent for dissolving silk fibers. Next, aqueous lithium bromide solution was used as a novel solvent system for the dissolution and blending of SF and microcrystalline cellulose, producing SF/cellulose nanocomposites with outstanding biocompatibility and high mechanical performance.
mechanical performance of the SF materials, which improves cell growth, proliferation, and differentiation. Cellulose nanofibrils (CNFs) derived from cellulose can potentially be an excellent nanofiller for high-performance polymer nanocomposite biomaterials because of their outstanding mechanical properties with an elastic modulus of 145.2 ± 31.3 GPa and a strength of 1.6−3 GPa.21,22 Moreover, nanofibers can provide guidance cues and mechanical support for cell attachment, spreading, and migration.23 Some attempts to reinforce CNFs within polymer scaffolds have been made, and their cytocompatibility was also investigated.24,25 The incorporation of CNFs into porous scaffolds can provide anchor points for cells and enhance cell attachment.25 CNFs can be extracted in the form of aqueous colloidal suspensions by subjecting slurries of cellulose fibers to strong mechanical disintegration processes.26 Several pretreatments of cellulose fibers such as enzymatic hydrolysis or chemical oxidation by a (2,2,6,6-tetramethylpiperidin-1-yl)oxyl (TEMPO)-mediated system are often required prior to the mechanical treatments to reduce energy consumption and make CNFs more uniform in size.26,27 However, the extraction processing is complex and time-consuming. Direct extraction of CNFs from natural cellulose remains a challenge due to the
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EXPERIMENTAL SECTION
Silk Fibroin Extraction. Bombyx mori raw silks were degummed by the established procedure described previously.30 Briefly, Bombyx mori raw silk fibers (Huzhou, China) were boiled three times in 0.05 wt % Na2CO3 for 30 min to remove sericin, rinsed thoroughly with deionized water, and then dried in an oven. Dissolution of Microcrystalline Cellulose. Figure 1 summarizes the dissolution process of microcrystalline cellulose and shows the 6228
DOI: 10.1021/acssuschemeng.7b01161 ACS Sustainable Chem. Eng. 2017, 5, 6227−6236
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ACS Sustainable Chemistry & Engineering morphology of the separated CNFs. The dried microcrystalline cellulose powder (435236, Sigma−Aldrich, U.S.A.) was dissolved in 9.3 mol/L LiBr solution (bath ratio = 1:80) at 110 °C for 1 h, 110 °C for 2 h, and 120 °C for 1 h. After the microcrystalline cellulose powder was completely dissolved, the solution was quickly poured into poly(methyl methacrylate) plates and then cooled down to a translucent gel. The cellulose hydrogel was washed thoroughly with deionized water to remove salt, and the residual of LiBr was detected with AgNO3 solution. The cellulose hydrogel derived from 110 °C for 1 h was dehydrated with ethanol and t-butyl alcohol, followed by freeze-drying for the observation of CNFs. The lyophilized matrixes were observed using field-emission scanning electron microscopy (SEM, S-4800, Hitachi, Japan) after being sputtered with gold. Mechanical Tests of Cellulose Films. To study the effect of the dissolution process on mechanical properties of the regenerated cellulose materials, the cellulose gel was dried in vacuum drying oven at 37 °C for 48 h to produce the cellulose films. The mechanical properties of various cellulose films were measured using an Instron 5967 mechanical testing instrument at 25 °C and 50 ± 5% RH for dried films and for wet films after water soaking. The films were cut as uniform strips with a size of 50 mm × 10 mm (n = 5 for each sample). The gauge length and the drawing speed were preset at 10 mm and 10 mm/min. Preparation and Characterization of Silk Fibroin/Cellulose Nanocomposite Films. Film Preparation. Figure 2 summarizes the preparation process of the SF/cellulose nanocomposite films. First, the degummed silk fibers (bath ratio = 1:10, at 60 °C for 55 min) and dried microcrystalline cellulose powder (bath ratio = 1:80, at 110 °C for 55 min) were separately dissolved in 9.3 mol/L LiBr solution. Then, the SF and cellulose solutions were mixed and magnetically stirred at 110 °C for 5 min with the desired ratios of cellulose to SF of 100/0, 75/25, 50/50, and 25/75. After thorough mixing, the solution was quickly poured into poly(methyl methacrylate) plates and then cooled down to a gel-like cake. The blend matrixes were marked as CE75SF25, CE50SF50, and CE25SF75. After the solution solidified, the gel was immersed in 75% ethanol for 2 h and washed thoroughly with deionized water to remove salts. The desalted gel was dried in a vacuum drying oven at 37 °C for 48 h to produce composite films. To visualize more clearly the size and dispersion of CNFs on the film surface, the gels were solvent-exchanged from ethanol to t-butyl alcohol. The traditionally prepared pure SF film was used as a control group. In brief, the extracted SF fibers were dissolved in 9.3 mol/L LiBr solution at 60 °C for 1 h. Then the solution was dialyzed against deionized water for 3 days with a 12−14 kDa molecular weight cutoff dialysis tube to remove the salt. The pure SF films were prepared using a regenerated SF solution cast upon a poly(methyl methacrylate) plate. After casting, the solution was dried for 24 h at 25 °C in a fume hood. The dried films were immersed in 75% ethanol for 2 h for waterinsoluble treatment. SEM Observation. The surface and cross section of the films were observed using S-4800 SEM after sputtering with gold. Tensile Testing. The mechanical properties of films were measured using a mechanical testing instrument in accordance with the abovementioned conditions. Attenuated Total Reflectance Fourier Transform Infrared Spectroscopy. The secondary structure of the films was analyzed by attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR) on a VERTEX 70 spectrometer (Bruker, Germany). All spectra were taken in the spectral range of 4000−500 cm−1 using an accumulation of 64 scans with a resolution of 4 cm−1. Contact Angle Test. The static water contact angle was measured to estimate the hydrophilicity of the films using an Optical Contact Angle Meter (JY-PHb, Puhui, China). The droplet of 5 μL of deionized water was deposited on the surface of the samples (n = 3 per sample) to measure the static water contact angle at 15 s. Enzymatic Degradation Experiment. The samples were weighed and incubated at 37 °C in 0.1 M phosphate-buffered saline (PBS; pH 7.4) solution containing 1.0 U/mL Protease XIV (Sigma− Aldrich). The samples (n = 3 per time point) were incubated in
enzyme solution (bath ratio 1:100) for 1, 3, 7, 14, and 21 days under slow shaking and in PBS under otherwise identical conditions as a control. The enzyme solution was replaced with a fresh solution every 3 days. At the designated time points, the remaining samples were rinsed and then dried at 105 °C to constant weight. The degradation rate was expressed as the percentage of weight loss relative to the initial dry weight. Cell Culture and Viability Assay. Human umbilical vein endothelial cells (HUVECs; ATCC, U.S.A.) were cultured in Dulbecco’s modified Eagle medium (DMEM, low glucose) supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% streptomycin-penicillin (Gibco). Cells were cultured in a humidified incubator at 37 °C and 5% CO2, and the culture medium was replaced every 3 days. The films were first punched into 15-mm-diameter samples and then placed into 24-well plates. The samples were sterilized by 75% ethanol for 30 min and then rinsed three times with sterilized PBS solution (0.1 M). Cells were seeded onto the films at a density of 1 × 105 per well. Cell-viability assessment was evaluated by a CCK-8 kit (Beyotime, China) following the procedure described previously.23 In brief, a 500μL volume of culture medium containing 50 μL of CCK-8 solution was added to each well and incubated for 3 h at 37 °C. The optical density (OD) value at 450 nm was measured by using a microplate reader (Bio-Tek Synergy HT, U.S.A.). The background absorbance of the culture medium (without cells) was subtracted from all the samples. The amount of the formazan dye, generated by the activities of dehydrogenase in cells, is directly proportional to the number of living cells. Immunofluorescence Labeling. At the designated time points, the cell-seeded samples were washed with PBS and fixed in 4% paraformaldehyde in PBS for 30 min at room temperature and then rinsed three times with PBS. The cells were permeabilized with 0.2% Triton X-100 in PBS for 5 min and then blocked with 2% BSA in PBS for 30 min. The F-actin were stained with fluorescein (FITC)phalloidin (5 μg/mL, Sigma−Aldrich) for 2 h at room temperature. After thoroughly rinsing with PBS, the cell nuclei were labeled with 4′,6-diamidino-2-phenylindole (DAPI; 5 μg/mL, Sigma−Aldrich) for 10 min and rinsed thoroughly with PBS. The fluorescence images for cell observation were obtained using a confocal laser scanning microscopy (CLSM; IX81/FV1000, Olympus, Japan).
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RESULTS AND DISCUSSION Microcrystalline Cellulose Dissolution and CNFs Separation. Cellulose is produced with highly crystalline nanofibrils consisting of fully extended and uniaxially aligned cellulose molecules.22 Therefore, cellulose is extremely difficult to dissolve in water and most conventional organic solvents due to the high crystallinity and complex hierarchical structure. The microfibrils of cellulose consist of elementary fibrils. Cellulose molecule chains connect with each other to form elementary fibrils through a large number of hydrogen bonds,31 making cellulose fibrils strongly resistant to aqueous salts. However, the connection regions between microfibrils have much weaker resistance to aqueous salts in comparison to these cellulose microfibrils. Kim and his co-workers reported that cellulose could be completely dissolved in a heated aqueous lithium bromide solution.32 Considering the weaker connections between microfibrils in the lateral direction, we proposed the feasibility of using an aqueous lithium bromide solution, which is the most frequently used solvent for SF dissolution, to separate cellulose microfibrils along the fiber direction by controlling the dissolution conditions. During the dissolution, the lithium bromide gradually permeated into cellulose and first destroyed the weak connections between the microfibrils, forming a CNFcontained cellulose solution (partial dissolution). As shown in Figure 1, homogeneous and transparent cellulose solutions 6229
DOI: 10.1021/acssuschemeng.7b01161 ACS Sustainable Chem. Eng. 2017, 5, 6227−6236
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ACS Sustainable Chemistry & Engineering formed at 110 °C for 1 h, in which microcrystalline cellulose powder has been completely dissolved. The dissolved solution formed a translucent gel-like cake after cooling to ∼70 °C. After freeze-drying the cellulose gel, we observed a large number of CNFs (see Figure 1b). However, excessive dissolution should be avoided for the extraction of nanofibrils, because extending the dissolution time and increasing the dissolution temperature may dissolve fibrils into cellulose molecules. On the other hand, the dissolved solution is a CNFs-containing cellulose solution. Increasing the heating temperature and dissolution time led to a considerable reduction in the polymerization degree of cellulose molecules.32 Severe dissolution conditions resulted in the chain scission of cellulose molecules, which would lead to negative influences on the mechanical performance of the regenerated cellulose materials. The color of the dissolved solution gradually changed to yellow with a prolonged dissolution time and elevated heating temperature (Figure 1), suggesting that the destruction of the molecular chain occurred during dissolving in aqueous lithium bromide solution. A tensile test was carried out to evaluate the mechanical properties of the regenerated cellulose films. The stress−strain curves are presented in Figure 3, and the results are listed in
ously increased along with the content of SF (see the inset of Figure 2). After the content increased to >75%, the gel-like cakes were fragile. Therefore, the blend ratios of CE75SF25 and CE50SF50 were chosen for the preparation of composite films. After being placed into a vacuum atmosphere, the gels turned to robust films. The transparency of the films continuously increased along with the content of SF (Figure 4A). As shown in Figure 5A, the pure SF films cast from the regenerated SF solution presented a smooth surface, whereas the exposed CNFs could be observed on the surface of pure cellulose films (Figure 5C) and CE50SF50 (Figure 5B). To visualize more clearly the size and dispersion of CNFs on the film surface, the hydrogels were solvent-exchanged from ethanol to t-butyl alcohol. The solvent-exchange method allowed us to remove the water from the hydrogel before drying.33 Consequently, the pore collapse of the nanofibril network during the solvent evaporation was minimized by employing weak polar solvents because lower surface tension forces were exerted on CNFs.33−35 Therefore, the preservation of fibrillar morphology on the film surface was achieved more effectively by the solvent-exchange drying. As shown in Figure 6, clearer observation of CNFs morphology on film surface was better achieved after drying. The mean width of the fibrils was a few tens of nanometers (Figure 6a-1), which is in the range of expected native CNFs.21,22 In comparison to pure cellulose film, the CNFs on SF/cellulose composite film appear more closely packed due to the filling and cover of SF. In Figure 5c, the cross section of the cellulose film is presented. Fragile fracture on a cross section of the composite films also shows a large number of exposed CNFs (Figure 5b). Homogenous dispersion is a key challenge for nanocomposite preparation using nanoscale reinforcements. In general, producing well-dispersed CNFs in a composite is difficult because the addition of CNFs often leads to phase separation between the fibrils and polymer matrix: the nanofibrils are more attracted to each other.36 In this work, the dissolved cellulose solution can rapidly form a hydrogel to lock in situ CNFs in the matrix. The images in Figure 5C and c show that the CNFs are homogeneously dispersed within the cellulose films, and homogeneous dispersion is also observed from the SF/cellulose composite films (Figure 5B and b). The clearer observation for CNFs distribution in films can be found in Figure 6. The homogeneous dispersion of CNFs in matrix has a positive influence on enhancing the mechanical properties of the nanocomposite films. Hydrophilicity. Surface wettability is an important characteristic known to affect the biological response to biomaterials. The static water contact angle was measured, and the contact angle values are shown in Figure 4. The samples for different films showed a similar contact angle value within 60−65°. The pure SF films showed a hydrophilic characteristic with a contact angle of 62.0 ± 1.0° due to the presence of hydrophilic hydroxyl, amino, and carboxyl groups. There are a number of hydrophilic hydroxyl groups in the cellulose molecules, forming a more hydrophilic surface. The pure cellulose films showed a small contact angle of 60.0° ± 0.9°. However, the nanocomposite films had a larger contact angle (64.0 ± 1.1°) than SF films, which may result from the rough morphology with nanofibril structure (Figure 5A−C).37 Suitable hydrophilicity is required for biomaterials to ensure cytocompatibility. Many studies have demonstrated that moderately hydrophilic surfaces tend to enhance cell adhesion
Figure 3. Influence of dissolution processing on the tensile properties of dried cellulose films.
Table 1. Tensile Tests of Cellulose Films Prepared by Different Dissolution Conditions in Dry State (n = 5) dissolution conditions
breaking stress (MPa)
breaking strain (%)
Young’s modulus (GPa)
110 °C, 1 h 120 °C, 1 h 110 °C, 2 h
109.3 ± 12.2 82.9 ± 5.9 44.3 ± 2.3
11.6 ± 1.6 9.5 ± 0.9 7.6 ± 1.3
3.8 ± 0.2 2.6 ± 0.3 1.5 ± 0.1
Table 1. It can be found that the breaking stress and breaking elongation of cellulose films showed considerable reduction with a prolonged dissolution time and elevated heating temperature. On the basis of the aim of this study to improve the mechanical performance of regenerated SF-based composite materials, the most mild dissolution condition (at 110 °C for 1 h) was chosen for subsequent experiments. Morphology of the Nanocomposite Films. The dissolved cellulose solution formed a translucent hydrogel after being cooled to ∼70 °C. Pure cellulose gel is semitranslucent, but the transparency of blend gels continu6230
DOI: 10.1021/acssuschemeng.7b01161 ACS Sustainable Chem. Eng. 2017, 5, 6227−6236
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Figure 4. (A) Optical pictures of obtained films. (B) Contact angles of different films.
Figure 5. SEM images of obtained films: (A−C) surface morphology, (a−c) cross-sectional morphology. Scale bars: (A−C) 1 μm, (a) 50 μm, (a1) 5 μm, (b, c) 1 μm.
Additionally, an absorption peak at 3280 cm−1 ascribed to N−H stretching of SF was also recorded.39 The spectrum of pure cellulose was characterized by an intense and broad band around 3358 cm−1, associated with O−H stretching vibration. Moreover, a weak adsorption peak at 1645 cm−1 and a strong adsorption peak at 1013 cm−1 could be observed, which corresponded to the C−O stretching vibration and CO carbonyl stretching vibration of cellulose, respectively.42,43 In the blends, the peak at 1624 cm−1 (amide I) and the peak at 1516 cm−1 of SF disappeared due to the existence of cellulose. The peak of cellulose at 1645 cm−1 could be observed, but the peak at 1013 cm−1 corresponding to the CO carbonyl stretching vibration of cellulose became weaker due to the existence of SF molecular chains. Moreover, a new weak adsorption peak at 1544 cm−1 appeared, suggesting the existence of strong interactions between SF and cellulose
and proliferation.38 In this study, the contact angle values of the films are moderate. Secondary Structure of SF and Cellulose in Composite Films. In the previous studies, the characterizations of SF/ cellulose films by FTIR, X-ray diffraction (XRD), thermogravimetric analysis (TGA), differental scanning calorimetry (DSC), and dynamic mechanical thermal analyses (DMA) showed the existence of strong interactions between SF and cellulose molecular chains,39−41 determining that the hydrogen bonds between cellulose chains are partly substituted by the hydrogen bonds between cellulose and SF chains.10 In this study, FTIR measurements were performed on the films to determine the secondary structure of SF and cellulose in composite films (Figure 7). The pure SF film showed strong absorption bands at 1624 cm−1 (amide I) and 1516 cm−1 (amide II), corresponding to the β-sheet structure of SF protein. 6231
DOI: 10.1021/acssuschemeng.7b01161 ACS Sustainable Chem. Eng. 2017, 5, 6227−6236
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Figure 6. SEM images of films after solvent-exchange treatment. (a, a1, b) Magnified images. Scale bars: (A, B) 2 μm, (a, b) 500 nm, (a-1) 200 nm.
groups in SF (3280 cm−1) disappeared, and the −OH stretching vibration of cellulose around 3358 cm−1 was broadened and shifted to a lower wavenumber around 3305 cm−1; the formation of the new intermolecular hydrogen bonds between SF and cellulose molecules is inferred.39 Mechanical Properties of the Nanocomposite Films. Tensile tests were carried out to evaluate the mechanical properties of SF/cellulose nanocomposite films, both in dry and wet states. The stress−strain curves of films are shown in Figure 8. The Young’s modulus, maximum tensile strength, and elongation at break are shown in Table 2. The cellulose films showed the highest Young’s modulus value of 3.8 ± 0.2 GPa, and the modulus is the least for pure SF films with 0.6 ± 0.1 GPa. Apparently, the modulus gradually increased with increasing cellulose content in the blends. It was found that dried pure SF films show the lowest tensile strength of 10.3 ± 3.7 MPa and the cellulose films have a high tensile strength of 109.3 ± 12.2 MPa. From Figure 8 and Table 2, it can be observed that the tensile strength of the blends improved with increasing cellulose content. Tensile strength and break elongation mainly depend on components in the film and intermolecular interactions. CNFs have outstanding mechanical properties with a strength of ∼1.6 GPa.22 The incorporation of
Figure 7. FTIR spectra of the films with various blend ratios.
molecular chains. The spectra of regenerated cellulose are characterized by the −OH of the intramolecular-hydrogenbonding interacting chains at 3600−3340 cm−1 and the −OH of the intermolecular-hydrogen-bonding interacting chains at 3320−3000 cm−1.44,45 The strong peak corresponding to N−H
Figure 8. Tensile properties of obtained dried films (A) and wet films (B). 6232
DOI: 10.1021/acssuschemeng.7b01161 ACS Sustainable Chem. Eng. 2017, 5, 6227−6236
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∼7.5% in PBS solution. The weight loss of the SF films and SF/ cellulose composite films when incubated in protease XIV solution increased with increasing exposure time, and the weight losses were 21.0% and 12.5% after 21 days, respectively. The results demonstrated that the profiles of enzymatic degradation of nanocomposite films were strongly dependent upon the SF component, because cellulose does not contain the enzyme-specific cleavage site. The degradation of cellulose in animal and human tissues is limited because of the absence of cellulase,51 whereas the extracellular matrixes of human tissues contain abundant protease that can degrade silk protein. The SF component significantly promoted the degradability of composite films, whereas the cellulose rendered the composites more stable to enzymatic degradation. Therefore, the degradation rate of composites can be regulated by changing the blend ratio of SF and cellulose. In Vitro Cell Culture of the Nanocomposite Films. To evaluate the biocompatibility of the obtained nanocomposite films, the morphology of HUVECs on the films were observed by CLSM (Figure 10). According to the CLSM observation,
Table 2. Tensile Tests of Composite Films in Dry and Wet States with Different Blend Ratios (n = 5) groups dry film
wet film
cellulose CE75SF25 CE50SF50 SF (ethanol treated) SF cellulose CE75SF25 CE50SF50 SF (ethanol treated)
breaking stress (MPa)
breaking strain (%)
Young’s modulus (GPa)
109.3 ± 12.2 73.2 ± 9.6 32.1 ± 4.3 22.2 ± 2.8
11.6 ± 1.6 8.8 ± 1.1 5.7 ± 0.3 4.7 ± 0.4
3.8 1.9 1.5 0.8
10.3 ± 3.7 45.0 ± 5.2 30.4 ± 2.1 15.2 ± 0.8 5.8 ± 0.3
3.2 ± 0.5 18.0 ± 3.2 15.0 ± 1.5 9.6 ± 0.5 6.0 ± 0.5
0.6 ± 0.1 0.44 ± 0.03 0.32 ± 0.03 0.20 ± 0.02 0.10 ± 0.01
± ± ± ±
0.2 0.2 0.1 0.1
CNFs into films significantly enhanced the mechanical properties of the composite films compared to pure SF films. However, the tensile strength cannot achieve the strength of single CNF, because the arrangement of CNFs is random and the slippage of fibrils is inevitable during tensile failure. On other hand, the enhanced mechanical properties of the composite films may be attributed to the strong interaction between SF protein and cellulose.10,39−41 The wet films showed a similar trend. Along with the decreasing cellulose proportion, the tensile strength of the composite film decreased, indicating that the reinforcement of cellulose still works in a wet state of the film. In Vitro Enzymatic Degradation. The absence of degradation in mammalian systems has proved to be a major impediment to tissue substitution. Although regenerated cellulose material can be regarded as a slowly degradable implantation material, the in vivo degradation is very slow.46 The degradability of regenerated SF materials has been demonstrated in numerous in vitro and in vivo models,47−50 and the degradation rate can be effectively regulated by controlling the processing of regenerated SF materials.2,50 Figure 9 shows the weight loss of the different films over the degradation time. After 21 days, the weight loss of the cellulose films incubated in both protease XIV solution and PBS was
