ARTICLE pubs.acs.org/Biomac
Degradation Mechanism and Control of Silk Fibroin Qiang Lu,*,†,‡ Bing Zhang,† Mingzhong Li,† Baoqi Zuo,† David L. Kaplan,§ Yongli Huang,† and Hesun Zhu|| †
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National Engineering Laboratory for Modern Silk, College of Textile and Clothing Engineering, Soochow University, Suzhou 215123, People’s Republic of China ‡ Jiangsu Province Key Laboratory of Stem Cell Research, Soochow University, Suzhou 215006, People’s Republic of China § Department of Biomedical Engineering, Tufts University, Medford, Massachusetts 02155, United States Research Center of Materials Science, Beijing Institute of Technology, Beijing, 100081, People’s Republic of China
bS Supporting Information ABSTRACT: Controlling the degradation process of silk is an important and interesting subject in the field of biomaterials. In the present study, silk fibroin films with different secondary conformations and nanostructures were used to study degradation behavior in buffered protease XIV solution. Different from previous studies, silk fibroin films with highest β-sheet content achieved the highest degradation rate in our research. A new degradation mechanism revealed that degradation behavior of silk fibroin was related to not only crystal content but also hydrophilic interaction and then crystal-noncrystal alternate nanostructures. First, hydrophilic blocks of silk fibroin were degraded. Then, hydrophobic crystal blocks that were formerly surrounded and immobilized by hydrophilic blocks became free particles and moved into solution. Therefore, on the basis of the mechanism, which enables the process to be more controllable and flexible, controlling the degradation behavior of silk fibroin without affecting other performances such as its mechanical or hydrophilic properties becomes feasible, and this would greatly expand the applications of silk as a biomedical material.
1. INTRODUCTION The degradability is very important for biomaterials used in tissue engineering.1,2 When damaged or diseased tissues are incapable of self-repair, a substitute biomaterial is often required to aid the healing process. The self-repairing periods of different tissues such as bone, tendons, ligament, or vessels are different, meaning that biomaterials used as scaffolds should have corresponding degradation rates to facilitate the formation of new tissues. Therefore, the controlling of the degradation of different scaffolds is still the major goal of tissue engineering research. Silk fibroin has aroused more and more interest in the field of biomedicine because of its excellent environmental stability, biocompatibility, morphologic flexibility, and mechanical properties.3-9 It has been found that silk fibroin had a promising future in tissue engineering,10,11 drug release,12,13 and optical apparatus.14 To satisfy the various requirements of different applications, the researchers still strive to extend the capabilities of silk fibroin from hydrophobic material to hydrophilic material,15,16 from filament to film, sheet, and scaffolds,11,17-19 and even from soft material to super rigid material.8,20 Some of these exciting developments have greatly stimulated studies on the silk-forming mechanism and the relationship between conformations, microstructures, and properties. However, the degradation is a serious obstacle for the applications of silk-based materials from the beginning. Although the degradability of silk-based materials can be sometimes changed through using different methods or adding different enzymes, the degrading process of silk is still bewildering.21-25 For example, r 2011 American Chemical Society
the β-sheet structure (Silk II) was considered to be a critical factor that stabilized silk fibroin in aqueous environments, but silk films, treated by methanol annealing and stretching, respectively, showed totally different degradation properties, even though the films had similar silk II content. To control the degradation behavior, it is necessary to explore potential factors that affect the degradation of silk fibroin. Silk I, the other main crystal structure of silk fibroin, is a hydrated structure and is considered to be a necessary intermediate for the preorganization or prealignment of silk fibroin molecules. In the nature, silk I structure was transformed to silk II structure after spinning process. Differing from the general opinion that considers the stability of silk fibroin to result from silk II structure, water-stable silk films mainly composed of silk I structure were prepared without the increase in silk II structure in our previous research.26 Compared with other silk films, the silk films in our research degraded more quickly in PBS and enzyme solutions. The water-stable silk films had special microstructures and flexible properties, making them suitable research objects for exploring the relationship between structures and degradation behavior. In this study, we researched the degrading process of three different insoluble silk films in enzyme solutions. These silk films had different crystal content and nanostructures, which is very Received: November 28, 2010 Revised: February 5, 2011 Published: February 25, 2011 1080
dx.doi.org/10.1021/bm101422j | Biomacromolecules 2011, 12, 1080–1086
Biomacromolecules useful for gaining a clear understanding of the relationship between structures, processing, and degradability. Therefore, it will become more predictable to control the degradability of silkbased materials according to the requirements of various tissue regenerations.
2. MATERIALS AND METHODS 2.1. Preparation of Silk Solutions. B. mori silk fibroin solutions were prepared following our previously published procedures.27 Cocoons were boiled for 20 min in an aqueous solution of 0.02 M Na2CO3, and then rinsed thoroughly with water to extract sericin proteins. The extracted silk was dissolved in 9.3 M LiBr solution at 60 °C, yielding a 20 wt % solution. This solution was dialyzed in water using Slide-a-Lyzer dialysis cassettes (Pierce, MWCO 3500) for 72 h. The final concentration of aqueous silk solution was ∼7.5 wt %, determined by weighing the remaining solid after drying. 2.2. Film Formation. We cast 1.5 mL of silk solution on polystyrene Petri dishes (diameter 30 mm). A lid with nine holes was placed over the dish to control the drying rate. The area of each hole was 3.13 mm2, and the drying time was 3 days when the hood airflow was maintained at 0.20 m s-1. Prepared by slow drying, the water-insoluble silk fibroin films with high silk I structure were termed SD-SF (SD, slow dry). Water insoluble silk fibroin films were also formed by water annealing treatment.21 In brief, silk fibroin films prepared by casting process were placed in a water-filled desiccator with a 25 in. Hg vacuum for 4 h to produce water-insoluble films, which were termed WA-SF (WA, water annealed). The other water-insoluble silk fibroin films were prepared by adding glycerol as additive. The silk fibroin solution (1.5 mL) was mixed with glycerol at weight ratios of 20% (w/w) and then poured in a Petri dish (diameter 30 mm) and dried at room temperature in a flow hood overnight. Once the films were prepared, they were stretched to 250% of their original length through using an Instron 3366 testing frame (Instron, Norwood, MA), resulting in the formation of silk II structure. Then, the treated films were incubated in distilled water for 24 h at room temperature to remove glycerol.28 Prepared by stretching and then dried in the air overnight, these silk fibroin films were termed ST-SF (ST, stretched). Through adjusting the volume of silk fibroin solution, the three water-insoluble films had similar thickness (Figure S1 of the Supporting Information). 2.3. Silk Degradation. Silk fibroin films were incubated at 37 °C in 40 mL of PBS solution that contained 0.23 U/mL protease XIV at pH 7.4. Each solution contained an approximately equivalent mass (40 ( 5 mg) of silk films. Solutions were replenished with enzyme and collected daily. At appointed time points, groups of samples were rinsed in distilled water and prepared for mass balance. 2.4. Differential Scanning Calorimetry (DSC). Samples of ∼5 mg were encapsulated in Al pans and heated in a TA Instruments Q100 DSC (TA Instruments, New Castle, DE) under a dry nitrogen gas flow of 50 mL/min. Standard mode DSC measurements were performed at a heating rate of 2 K/min. Temperature-modulated differential scanning calorimetry (TMDSC) measurements were also performed using a TA Instruments Q100, equipped with a refrigerated cooling system. The samples were heated at 2 K/min with a modulation period of 60 s and a temperature amplitude of 0.318 K. 2.5. Fourier Transform Infrared Spectroscopy (FTIR). FTIR analysis was performed with a Bruker Equinox 55/S FTIR spectrometer, equipped with a deuterated triglycine sulfate detector, a multiplereflection, and a horizontal MIRacle ATR attachment (using a Ge crystal, from Pile Tech.). For each measurement, 32 scans were coded with resolution 4 cm-1, with the wavenumber ranging from 400 to 4000 cm-1. Fourier self-deconvolution (FSD) of the infrared spectra covering the amide I region (1595-1705 cm-1) was performed by Opus 5.0 software. Deconvolution was performed using Lorentzian line
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shape with a half-bandwidth of 25 cm-1 and a noise reduction factor of 0.3. FSD spectra were curve-fitted to measure the relative areas of the amide I region components.29 2.6. Scanning Electron Microscopy (SEM). The surface morphologies of different silk films were imaged with a Zeiss Supra 55 VP SEM (Oberkochen, Germany). Then, these films were fractured in liquid nitrogen and sputtered with platinum. The cross-section images were also investigated with a Zeiss Supra 55 VP SEM. 2.7. Atomic Force Microscopy (AFM). The morphology of degraded silk fibroin in enzyme solution was observed by AFM (Veeco, Nanoscope III) in air. A 225 μm long silicon cantilever with a spring constant of 3 N/m was used in tapping mode.
3. RESULTS AND DISCUSSION 3.1. Structure of Silk Fibroin Films. Changes in the structure of silk films prepared with various methods were determined by FTIR. The infrared spectral region within 1700-1500 cm-1 was assigned to absorption by the peptide backbones of amide I (1700-1600 cm-1) and amide II (1600-1500 cm-1), which were usually used for the analysis of different secondary structures of silk fibroin. The peaks at 1610-1630, 1695-1700, and 1510-1520 cm-1 were characteristic of silk II secondary structure, whereas the absorptions at 1648-1654 and 15351542 cm-1 were indicative of silk I conformation.29,30 As shown in Figure 1A, the amide I band for the SD-SF films showed one strong peak at 1651 cm-1, corresponding to silk I structure. In the ST-SF films, the amide I band showed one strong peak at 1624 cm-1, with a shoulder at 1651 cm-1, whereas in the WA-SF samples, one peak at 1651 cm-1 appeared, with a shoulder at 1624 cm-1. The same trend in structural change was also found in the amide II region. From the SD-SF to the WA-SF and the ST-SF samples, the peak at 1539 cm-1 (silk I) decreased, whereas the peak at 1515 cm-1 (silk II) increased. The results indicated that silk films with different crystal structures were achieved by changing the process. Structural changes in the silk films after different processes were confirmed by DSC (Figure 1B). In our previous studies, it was found that silk I crystal structure degraded at ∼250 °C and silk II degraded at ∼260 °C.26 The SD-SF samples showed a main degradation peak at 251 °C and a minor degradation peak at 258 °C, indicating the formation of the stable silk I structure. For the WA-SF samples, the strength of degradation peak at ∼250 °C decreased, and two minor peaks at ∼260 °C appeared, implying the decrease in silk I crystal structure and the increase in silk II structure. Only one degradation peak at 261 °C was found for the ST-SF samples, meaning that they were mainly composed of silk II structure. Interestingly, compared with that of the SDSF samples, the degradation peak of the WA-SF samples moved from 251 to 247 °C, indicating that unstable silk I structure formed in the WA-SF samples, which might due to deficient selfassembly in water annealing process. The DSC results are consistent with the FTIR results, confirming that the ST-SF samples had the highest silk II structure and thermal stability, whereas the SD-SF samples had the lowest silk II structure. In our previous studies, we reported the observation of a lower Tg(1), which provided information about the removal of bound water, indicating the interaction between silk and bound water and further hydrophilic interaction between silk fibroin molecules.26 As shown in Figure 1C, the ST-SF samples had no Tg(1), meaning that traces of water formed strong interactions with silk fibroin protein. Tg(1) was found in both the 1081
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Figure 1. (A) FTIR spectra of silk fibroin films prepared with different processes; (B) DSC data from silk fibroin films prepared with different processes; and (C) TMDSC data from silk fibroin films prepared with different processes: (a) silk fibroin film derived from slow drying process, SD-SF, (b) silk fibroin film derived from water-annealing process, WA-SF, and (c) silk fibroin film derived from stretching process, ST-SF.
WA-SF and the SD-SF samples, with a higher heat capacity at Tg(1) being achieved for the SD-SF samples. The results indicated that compared with the WA-SF samples, stronger hydrophilic interaction formed in SD-SF films. The nanostructure of samples was investigated by SEM. The special nanostructure of the SD-SF samples was studied in our previous studies.26 Although the surface was relatively flat, the inside of silk fibroin films formed special core-layer globules with about 200-1000 nm in diameter, in which the core of globules was composed of nanofilaments containing high crystal content and surrounded by random nanofilaments (Figure 2a-c). Likewise, the core-layer structure composed of nanofilaments was also formed inside the WA-SF films (Figure 2d-f), which was confirmed by the following SEM, FTIR, and DSC results of the degraded WA-SF samples (Figures 4-6). The nanostructure of the ST-SF samples was different from that of the SD-SF and the WA-SF samples (Figure 2g,h). Their main structures consist of nanoparticles and short nanofilaments rather than core-layer globular structure. Some short nanofilaments with size of tens of nanometers were surrounded by nanoparticles with
