Nanofibers Made of Globular Proteins - American Chemical Society

Sep 20, 2008 - strength, high surface area/volume ratios, and biocompatibility resemble those of ... microscale polymeric fibers is by electrospinning...
0 downloads 0 Views 11MB Size
Download PDF
Biomacromolecules 2008, 9, 2749–2754

2749

Nanofibers Made of Globular Proteins Yael Dror,† Tamar Ziv,‡ Vadim Makarov,† Hila Wolf,‡ Arie Admon,‡ and Eyal Zussman*,† Faculty of Mechanical Engineering and Faculty of Biology, Technion - Israel Institute of Technology, Haifa 32000, Israel Received May 13, 2008; Revised Manuscript Received July 31, 2008

Strong nanofibers composed entirely of a model globular protein, namely, bovine serum albumin (BSA), were produced by electrospinning directly from a BSA solution without the use of chemical cross-linkers. Control of the spinnability and the mechanical properties of the produced nanofibers was achieved by manipulating the protein conformation, protein aggregation, and intra/intermolecular disulfide bonds exchange. In this manner, a low-viscosity globular protein solution could be modified into a polymer-like spinnable solution and easily spun into fibers whose mechanical properties were as good as those of natural fibers made of fibrous protein. We demonstrate here that newly formed disulfide bonds (intra/intermolecular) have a dominant role in both the formation of the nanofibers and in providing them with superior mechanical properties. Our approach to engineer proteins into biocompatible fibrous structures may be used in a wide range of biomedical applications such as suturing, wound dressing, and wound closure.

1. Introduction Fibrous proteins serve diverse structural functions in living organisms, including forming the extracellular matrix (ECM) and cellular scaffolds, and, in general, they provide a stabilizing and protective environment for cells in a variety of tissues.1,2 Their superior mechanical properties have inspired attempts to engineer similar versatile protein-based fibers for use in varied biomedical applications. Artificial nanofibers, whose dimensions, strength, high surface area/volume ratios, and biocompatibility resemble those of natural ECM fibers, are highly desired. One of the attractive methods to fabricate artificial nano- and microscale polymeric fibers is by electrospinning.3 This method is relatively simple, allowing for the manipulation of many parameters, such as fiber diameter and morphology, and enabling the formation of nonwoven mats with a controlled density. For these reasons, electrospinning has gained significant interest during the past decade as an alternative methodology for the development of biodegradable polymers for possible biomedical applications such as scaffolds for tissue engineering4-7 and as carriers for drugs and bioactive molecules, including enzymes or even bacteria.8-13 Significant efforts have recently been made to mimic nature by fabricating fibers made solely from natural proteins, hoping that such nanofibers would be less prone to rejection by the host. Unfortunately, the electrospinning of natural molecules has proven to be somewhat of a challenge because they do not behave like classic polymers lacking the viscoelastic properties essential for stable electrospinning. Consequently, biomolecules are often blended and cospun with synthetic polymers.4,14,15 Silk, the best known natural-spun fiber with its striking mechanical properties, was among the first to be mimicked using the electrospinning approach. A number of groups reported attempts to electrospin fibers of silk fibroin directly from aqueous solutions,16,17 with and without additional polymers.18,19 Yet, the mechanical properties of these electrospun silk fibers were * To whom correspondence should be addressed. E-mail: meeyal@ tx.tehnion.ac.il. † Faculty of Mechanical Engineering. ‡ Faculty of Biology.

inferior when compared to natural silkworm fibers. Posttreatment with methanol improved the mechanical properties of the electrospun silk fibers to a slight degree, most likely due to the increased formation of β-sheets. More recently, natural spider silk20 and other recombinant proteins were successfully electrospun from a hexaflouroisopropanol (HFIP) solution,21,22 whereby the semicrystalline structure was better preserved. Other fibrous proteins, such as collagen, gelatin (denatured collagen), elastin, and fibrinogen have also been successfully electrospun.23-26 However, the mechanical properties of these artificial fibers were rather poor as well. Chemical cross-linking was needed to attain mechanical properties suitable to create scaffolds. Electrospinning of keratin nanofibers was achieved by Thomas et al.27 Wool keratin was separated into two fractions, but only the smaller fraction, having a larger molecular weight and a lower amount of cysteine, could be spun in its pristine form. The larger fraction, with a low molecular weight and having a higher concentration of cysteine, could only be spun with the assistance of a supporting polymer matrix. Electrospinning of nanofibers made of the globular protein hemoglobin were described by Barnes et al.28 However, the mechanical properties of the product were unsatisfactory even after cross-linking. Here, we present the first successful electrospinning of strong nanofibers made of a globular protein, namely bovine serum albumin (BSA). This protein was selected under the assumption that, being one of the most abundant proteins in the body, nanofibers made from serum albumin would be regarded as being “less foreign” to the body and, therefore, less likely to be rejected. Indeed, its stability, adhesivity, and coating properties,29 together with its low cost, makes it a very good raw material candidate for the electrospinning of nanofibers. Furthermore, serum albumin is rich in cysteine, participating in as many as 17 intramolecular disulfide bridges. Our procedure, involving the opening of these disulfides while concomitantly unfolding the protein in a controlled fashion and allowing the reformation of new, extended structures rich with strong interand intramolecular disulfide covalent bonds, proved very effective in producing superb nanofibers. The main physical challenge encountered involved turning the low-viscosity globu-

10.1021/bm8005243 CCC: $40.75  2008 American Chemical Society Published on Web 09/20/2008

2750

Biomacromolecules, Vol. 9, No. 10, 2008

Table 1. Composition of the Solutions from which the Fibers were Spun No.

composition

A A0 A1 A2

12% BSA TFE/H2O 9:1 (weight ratio) 10% BSA TFE/H2O 9:1 + 10 equiv/bond β-ME 10% BSA TFE/H2O 9:1 + 10 equiv/>bond β-ME+ IAA 10% BSA TFE/ammonium hydroxide 0.1 M 9:1 + 10 equiv/ bond β-ME + IAA, pH ∼ 9 A3 10% BSA TFE/ammonium hydroxide 0.1 M 9:1 + 10 equiv/ bond β-ME, pH ∼ 9 A11 10% BSA TFE/HCl 0.1 M 9:1 + 10 equiv/bond β-ME, pH ∼ 2

lar protein solution into a polymer-like spinnable solution and producing keratin or elastin-like fibers with enriched intermolecular disulfide bridges.

2. Materials and Methods 2.1. Materials and Solutions. Bovine albumin fraction V was supplied by MP Biomedicals (Israel). Trifluroethanol (TFE) and β-mercaptoethanol (β-ME) were purchased from Aldrich (U.S.A.). Urea, dithiothreitol (DTT), 4-vinylpyridine (4-VP), and iodoacetamide (IAA) were purchased from Sigma. Different compositions of the BSA solutions are listed in Table 1. 2.2. Electrospinning. The solutions underwent electrospinning from a 5 mL syringe with a hypodermic needle having an inner diameter of 0.5 mm. The flow rate was Q ) 0.2-0.5 mL/hour. A copper electrode was placed in the polymer solution and the suspension was spun onto the edge of a grounded collector disk (cf., ref 30). The electrostatic field was E ) 1.1 kV/cm and the distance between the electrode tip and the edge of the disk (or the horizontal disk) was 12 cm. The linear speed at the edge of the disk collector was V ) 8.8 m/s. All the experiments were performed at room temperature (about 24 °C) and a humidity of about 50%. The samples were then stored for 24 h in a desiccator (humidity of about 30%). 2.3. Imaging. Images of the fibers were obtained using a Leo Gemini high resolution scanning electron microscope (HRSEM) at an acceleration voltage of 2-4 kV and sample-to-detector distance of 2-5 mm. The specimens were coated with a thin gold film. To image a crosssection of the fibers, an oriented mat was collected on a rotating wheel following Theron et al.’s approach30 and then cut. 2.4. Mechanical Properties. For tensile measurements, the electrospun nanofibers were collected on a grounded rotating disk forming a converging electric field.30 In this manner, a strip of oriented fibers 2.5 cm wide was formed. The final product was dried under vacuum for 48 h before testing. The thickness of the strip was measured using a SEM by imaging its cross-section. Tensile tests were conducted on a Testmashine D’Essais, a uniaxial tension machine designed for small samples. Each result is an average of 12-15 specimens. The effective cross-section of the strip, which takes into account the voids in between the collected fibers in the strip, was determined by considering the strip porosity 1 - Fs/F0 where Fs is the strip density, and F0 is the albumin density. The density was approximated as 1.44 ((0.06) g/mL31 considering that the samples were dried. 2.5. X-ray Diffraction. Wide-angle X-ray scattering (WAXS) measurements were performed with a Bruker Nanostar KFF CU 2 K-90 small-angle diffractometer with Cu Ka radiation (0.1542 nm), pinhole collimation (that yielded a beam of 300 µm in diameter), and a 10 × 10 cm2 two-dimensional position-sensitive wire detector, positioned 7 cm behind the examined sample. To perform the WAXS measurements, the fibers were collected on a vertical rotating wheel having a tapered edge so as to form a thin rope of oriented fibers. This rope was then fixed on an aluminum frame, which was mounted in the diffractometer with the rope fibers perpendicular to the X-ray beam. 2.6. Protein Separation by SDS-PAGE. A total of 10 µg of BSA fibers in 8 M urea were either alkylated with 150 mM IAA (30 min RT) or reduced with 40 mM DTT (30 min at 60 °C), followed by

Dror et al. alkylation with IAA at 150 mM. The proteins were resolved by 4-20% denaturing SDS-PAGE. 2.7. Mass Spectrometry. The protein fibers were dissolved in 8 M urea and 200 mM ammonium bicarbonate modified with 35 mM IAA and proteolyzed with modified trypsin (Promega) at a 1:100 enzymeto-substrate ratio. The resulting peptides were reduced (120 mM DTT) and modified with 0.5 M 4-VP. The resulting IAA- and 4-VP-modified peptides were resolved by reversed-phase chromatography on 0.075 internal diameter 200 mm length fused silica capillaries (J&W) packed with Reprosil C18 reversed phase beads. The peptides were eluted with linear 75 min gradients, of 5-45%, and 15 min at 95% acetonitrile with 0.1% formic acid in water at a flow rate of 0.25 µL/min. Mass spectrometry was performed with the Orbitrap mass spectrometer (Thermo-Fisher, San Jose) in a positive mode using a repetitive full MS scan followed by collision-induced dissociation (CID) of the seven most dominant ions selected from the first MS scan. The mass spectrometry data was analyzed using the Bioworks Sequest 3.31 software (Thermo-Fisher, San Jose) searching against the bovine part of the NCBI-NR database with optional cysteine modifications of 57 Da (IAA modified) or 105 Da (4-VP modified).

3. Results and Discussion 3.1. Structural Characterizations and Morphology. BSA solutions were prepared in a mixture of 9:1 trifluoroethanol/ water (TFE/H2O) with additional modifiers as listed in Table 1. TFE is known to affect protein structure differently in accordance with its concentration. Most prominently TFE at low concentrations, but also HFIP and to some extent methanol and ethanol, stabilize the native structure of the proteins in aqueous solutions. However, at high TFE concentrations, the structure of the water and the associated hydrophobic interaction within the proteins are disrupted (Luo and Baldwin32 and refs therein). This bulk effect causes unfolding of the tertiary structure of the proteins, while concurrently strengthening some intramolecular hydrogen bonds involved with secondary structures, such as R-helixes. This combined effect results in the formation of what is called the “open-helical structure” in which the interaction between helix segments is weak, while the hydrophobic segments are mostly exposed to the solvent. We worked with the higher concentrations of TFE under the assumption that the electrospinning process would benefit from the open conformation of the protein. Indeed, aqueous and lowratio TFE solutions of BSA were totally unspinnable even at high protein concentrations. This was most likely due to the compact globular shape of the protein and, hence, the improper rheological properties of the solutions. In contrast, the rich alcoholic solutions were much more spinnable, however, the electrospinning process was still unstable with the resulting fibers highly fragmented (see Figure 1). Upon the addition of β-mercaptoethanol (β-ME), an effective reducing agent of disulfide bonds, a remarkable improvement in the spinning process was achieved. Opening of the intramolecular disulfide bridges, together with the partial denaturing created by the TFE environment, produced a pronounced expansion of the protein. This, in turn, affected both the rheological properties of the solution and its spinnability. The improvement of the process was reflected in the morphology, structure, and mechanical properties of the resulting fibers. Scanning electron microscopy (SEM) showed that fibers produced with β-ME are continuous and have a smooth surface morphology (Figure 2a). The nanofibers could be spun into thick mats (Figure 2b) that could be shaped macroscopically into desired shapes, such as the ribbon shown in Figure 2c. 3.2. Mechanical Properties. The tensile properties of the nanofibers electrospun in the presence of β-ME were remarkably

Nanofibers Made of Globular Proteins

Biomacromolecules, Vol. 9, No. 10, 2008

2751

Figure 1. Image of electrospun BSA solution A (12% BSA TFE/H2O 9:1 (weight ratio). Due to the low viscoelasticity, the electrospinning process was unstable and the as-spun fibers were relatively short with uneven diameter.

high. The enhanced properties were mainly attributed to the reduction of the original intramolecular disulfide bonds enabling the spontaneous reformation of new inter- and intramolecular bonds. This claim is supported by the opposing effect created by the introduction of a sulfhydryl blocking reagent IAA (Table 1, A1 and A2, and Figure 3) after the original disulfide bonds had been reduced with β-ME. The IAA reacted with the opened sulfhydryl groups and covalently blocked them, thereby preventing most of the spontaneous reformation of both intra- and intermolecular disulfide bonds. The stress-strain characteristics and stiffness of these fibers (A1, A2) were significantly inferior (Figure 3a,b), while the ductility (Figure 3c) was higher, probably due to the relative free movement of the molecules in the absence of S-S bridges. Maximum stress and elastic modulus ranging from 30-60 MPa and 1.5-2 GPa, respectively, were achieved in nanofibers reinforced by spontaneous covalent cross-linking of S-S bonds in the absence of IAA. The variation of the density, F0 was in the range of ((0.06) and this did not significantly affect the overall spread of the calculated stress/modulus. 3.3. Morphology and pH Effects. Further manipulation of the mechanical properties was achieved by using either a low or high pH electrospinning solution. The extreme pHs induced important effects on the conformation and aggregation properties of the protein, on top of their already discussed effect on the stability of S-S bonds. Basic pH solutions favor reduction of the disulfides bonds. Therefore, more sulfhydryl sites become available for the establishment of new S-S bridges resulting in improved mechanical properties. Furthermore, BSA undergoes reversible conformational transitions upon exposure to extreme pHs. In acidic conditions it prominently expands a conformation referred to as the E form33 and its tendency to aggregate is reduced due to its high positive charge. These opened, elongated, and more soluble proteins are well suited for the electrospinning process. Indeed, the minor improvement in the mechanical properties of the fibers at the low and high pHs (A11 and A3) can be attributed to the alignment of the molecules and to the high quality of the spinning. At a neutral pH (as is the case of A0), which is close to the isoelectric point of the protein (pH ) 5.4),34 the molecules tend to aggregate more easily in the solution and even more so as the TFE evaporates. This aggregation may have disturbed the electrospinning process and restricted additional orientation during the

Figure 2. Electrospun bovine serum albumin (BSA) fibers. (a) Scanning electron microscope (SEM) images of cross-section (of A3 fibers), (b) top view of electrospun nanofibers, and (c) optical image of electrospun ribbon made of as-spun BSA nanofibers.

stretching phase. The combination of these various influences, the degree of reduction of intramolecular S-S bonds, the more open conformation of the molecules, the propensity to aggregate and the reduced viscosity, and surface tension of the solution, had cumulative effects on the mechanical properties of the nanofibers. Interestingly, among the strongest fibers (A0, A3, and A11), the high tensile strength of A3 fibers was not accompanied by a drastic reduction in elongation (Figure 3d). A3 fibers were much more ductile and had higher toughness than the other two. In fact, the tensile curve of A3 resembles that of a typical stiff and tough polymer, which is characterized by a high yielding point and subsequent cold drawing and moderate strain hardening. This behavior is possibly related to additional orientation that takes place during stretching in the

2752

Biomacromolecules, Vol. 9, No. 10, 2008

Dror et al.

Figure 3. Mechanical properties of electrospun BSA fibers obtained by tensile testing a ribbon of aligned fibers (A0, A1, A3, A2, and A11). (a) The elastic modulus of the fibers, (b) the yield stress of the fibers, (c) the maximal elongation (the strain at failure), and (d) typical stress-strain curve results of tensile tests of as-spun A0, A3, and A11 BSA fibers.

necking area. Specimens that attain their maximum orientation right at the beginning of the stretching phase, as is the case for A11 and A0 fibers, cannot undergo this elongation; instead, they break right after the elastic linear region of the stress-strain curve as would a hard and brittle polymer. Ultimately, A3 fibers have a remarkable combination of desirable properties, attaining the stiffness of collagen and keratin, and the extensibility of dragline silk and elastin. These values are far higher than those reported for fibers electrospun with the globular protein hemoglobin28 and from fibrous proteins such as collagen, gelatin, elastin, and fibrinogen24,26 and are even comparable to those of natural fibers.35 3.4. Mass Spectrometry Analysis. Correlation between the presence of newly formed intermolecular disulfide bridges and the mechanical properties of the fibers was provided first by subjecting the proteins to sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) with and without preopening of the disulfide bridges (Figure 4). Intermolecular cross-linked molecules are detected as a series of dimers, trimers and even higher order bands resistant to the denaturing conditions of the SDS-PAGE sample-buffer, which contained both 8 M urea and SDS. Further reduction of the disulfide bridges with DTT broke most of these intermolecular bridges and eliminated most of the very high molecular weight multimers that had been too large to enter the gel (Figure 4, boxed area at the top). It is interesting to note that in the A3 fibers, these multimers were less abundant than in the A11 and A0 fibers (Figure 4). Thus, it seems that A3 is cross-linked to a somewhat lesser extent, possibly explaining its ductility. Further quantitative data about the degree of disulfide bond formation resulting from the different treatments was provided by a quantitative proteomic

Figure 4. SDS-PAGE showing different BSA nanofibers and the starting material used for their production, with the molecular size indicated on the left. Addition of the reducing agent, 2-ME, to the electrospinning solution and the subsequent reduction of the disulfide bridges with DTT in the different spun nanofibers is marked at the top. P, BSA powder; A0, fibers spun at pH 7; A3, fibers spun at pH 10; A11, fibers spun at pH 3; and A, fibers spun without prereduction with 2-ME.

analysis. This was carried out by first blocking the open sulfhydryls with IAA followed by reduction of the S-S bridges formed during the solution preparation and spinning with DTT, and then further modifying them with 4-VP. The relative mass

Nanofibers Made of Globular Proteins

Biomacromolecules, Vol. 9, No. 10, 2008

2753

Figure 5. Ratio of the optical absorbance at 260/280 nm of the tryptic peptides mixture modified with 4-VP after first blockage of sulfhydryls with IAA. The absorbance of 4-VP was detected in 260 nm, while total protein concentration was followed in 280 nm.

spectrometry signal intensity of each of the cysteine-containing tryptic peptides modified with IAA relative to the same peptides modified with 4-VP served as an indicator of the degree of participation of each cysteine in disulfide bond formation (see Figure 5). The data shows a higher degree of disulfide crosslinking in the A11 fibers and a somewhat lower degree of crosslinking in the A3 fibers. The A1 and A2 spun in the presence of IAA served as controls, in which the degree of cross-linking was much reduced by the blocking effect of IAA on the sulfhydryls. The high degree of cross-linking observed by higher 4-VP content is also apparent from the optical absorbance at 260 nm of the tryptic peptide mixture modified with 4-VP (after the initial blocking of the sulfhydryls with IAA) relative to its absorbance at 280 nm, which reflects the total amount of the peptides. The relative amount of inter- versus intra-molecular bridges is impossible to establish, yet it is clear from the pattern of migration of the multimers in the SDS-PAGE (Figure 4) that a significant proportion of the molecules are indeed intermolecularly cross-linked, appearing as higher order multimers in the gel. 3.5. X-ray Results. X-ray diffraction patterns of different fibers are shown in Figures 6 and 7. The distinct rings observed in the diffraction patterns indicate that the cast films as well as the nanofibers are semicrystalline. Moreover, the 1D patterns of the diffraction peaks’ positions indicate similar d-spacing for all the samples. These patterns are characterized by two Bragg peaks at 2θ ) 9.3 and 20°, corresponding to 9.5 (inner ring) and 4.4 Å (outer ring) d-spacing, respectively. The 4.4 Å d-spacing is suggested to be related to the backbone distance within the R-helixes, while the larger d-spacing is attributed to lateral R-helix segment packing. This interpretation is supported by previous reports on the X-ray diffraction of natural R-proteins36 and it also agrees with the recent structural investigation conducted by Wang et al. on zein, a group of alcohol-soluble proteins found in corn endosperm.37 It should be borne in mind that BSA is a globular protein rich in R-helix content, which behaves in rich TFE/water mixtures quite similarly to zein proteins.38 This result suggests that neither the dissolution of the protein in TFE, the breaking of disulfide bonds by β-ME, nor the electrospinning process itself alters the dimensions of the helixes and their eventual rearrangement in the solid state. Nonetheless, for fibers spun with β-ΜE (A0, A3, and A11), the subdomains of laterally packed R-helix segments are more oriented along the fiber axis, as can be seen by the equatorial arcs of the inner ring (Figure 6), while the outer ring remains isotropic. This difference between the two peaks further supports

Figure 6. X-ray diffraction patterns of electrospun fibers and cast films. 2D X-ray diffraction patterns of (i) cast film of BSA from a solution of 9:1 TFE/H2O and β-ME, (ii) A1 fibers, (iii) A3 fibers, (iv) A11 fibers, and (v) A0 fibers.

the claim that they can be attributed to a separated periodic order and are not simply two reflections of the same d-spacing with a different order of the Bragg law. In all the other samples, including the fibers containing IAA (A1), which prevented the reformation of disulfide bonds, there is no preferred orientation as indicated in Figure 7. Hence, the electrospinning process, concomitantly with the opening of the original intramolecular disulfide bonds, which enabled the formation of new disulfide bonds, all contribute to this preferred orientation. Furthermore, the crystallinity and alignment of the R-helix subdomains may also contribute to the enhanced mechanical properties. Indeed, silk fibers electrospun from aqueous solution and remained amorphous as was indicated by the lack the typical diffraction peak found in natural silk, and assigned to crystalline β-sheet,16,19 exhibited poor mechanical properties. On the other hand, silk fibers electrospun from a fluorinated alcohol solution, as was done here with BSA, exhibited a crystalline diffraction pattern.20 It can thus be concluded that the improved mechanical properties were achieved by the combined effects of the unfolding of the protein chains, an increased crystallinity, greater cross-linking, and the orientation of the ordered packing.

4. Conclusions In conclusion, fibers made solely of globular proteins, with controllable and improved mechanical properties, have been

2754

Biomacromolecules, Vol. 9, No. 10, 2008

Figure 7. X-ray diffraction patterns of electrospun fibers and cast films. (a) 1D X-ray diffraction patterns after radial averaging; and (b) 1D X-ray diffraction patterns of the azymutal intensity of the 10.2 d-spacing (inner) peak.

successfully electrospun. Their improved response to tensile stress is attributed to spontaneous cross-linking of S-S interdisulfide bonds, increased crystallinity, and enhanced orientation of the helix-helix interactions. These improved characteristic were achieved by manipulating the protein chain conformation, altering their electrostatic charge, reducing the self-disulfide bonds and promoting the formation of new cross-links, all achieved through the various combinations of a denaturing TFE environment, a reducing agent, and extreme pHs. By manipulating these elements, desired combinations of mechanical properties of different natural fibrous proteins can be achieved, ranging from stiff and brittle to strong and ductile, each tailored according to the specific application Acknowledgment. The authors acknowledge the financial support from the Russel Berrie Nanotechnology Institute (RBNI) and the Elias Fund (both at the Technion). The technical assistance of S. Chervinski is appreciated.

References and Notes (1) Gosline, J.; Lillie, M.; Carrington, E.; Guerette, P.; Ortlepp, C.; Savage, K. Philos. Trans. R. Soc. London, Ser. B 2002, 357, 121–132. (2) Scheibel, T. Curr. Opin. Biotechnol. 2005, 16, 427–433. (3) Reneker, D. H.; Yarin, A. L.; Zussman, E.; Xu, H. AdV. Appl. Mech. 2007, 41, 43–195. (4) Bhattarai, N.; Li, Z. S.; Edmondson, D.; Zhang, M. Q. AdV. Mater. 2006, 18, 1463.

Dror et al. (5) Shin, H. J.; Lee, C. H.; Cho, I. H.; Kim, Y. J.; Lee, Y. J.; Kim, I. A.; Park, K. D.; Yui, N.; Shin, J. W. J. Biomater. Sci., Polym. Ed. 2006, 17, 103–119. (6) Sell, S.; Barnes, C.; Smith, M.; McClure, M.; Madurantakam, P.; Grant, J.; McManus, M.; Bowlin, G. Polym. Int. 2007, 56, 1349–1360. (7) Yang, F.; Murugan, R.; Wang, S.; Ramakrishna, S. Biomaterials 2005, 26, 2603–2610. (8) Dror, Y.; Salalha, W.; Avrahami, R.; Zussman, E.; Yarin, A. L.; Dersch, R.; Greiner, A.; Wendorff, J. H. Small 2007, 3, 1064–1073. (9) Jia, H. F.; Zhu, G. Y.; Vugrinovich, B.; Kataphinan, W.; Reneker, D. H.; Wang, P. Biotechnol. Prog. 2002, 18, 1027–1032. (10) Salalha, W.; Kuhn, J.; Dror, Y.; Zussman, E. Nanotechnology 2006, 17, 4675–4681. (11) Greiner, A.; Wendorff, J. H.; Yarin, A. L.; Zussman, E. Appl. Microbiol. Biotechnol. 2006, 71, 387–393. (12) Luu, Y. K.; Kim, K.; Hsiao, B. S.; Chu, B.; Hadjiargyrou, M. J. Controlled Release 2003, 89, 341–353. (13) Dror, Y.; Kuhn, J.; Avrahami, R.; Zussman, E. Macromolecules 2008, 41, 4187–4192. (14) Kwon, I. K.; Matsuda, T. Biomacromolrcules 2005, 6, 2096–2105. (15) Xie, J.; Hsieh, Y. L. J. Mater. Sci. 2003, 38, 2125–2133. (16) Wang, M.; Jin, H. J.; Kaplan, D. L.; Rutledge, G. C. Macromolecules 2004, 37, 6856–6864. (17) Putthanarat, S.; Eby, R. K.; Kataphinan, W.; Jones, S.; Naik, R. R.; Reneker, D. H.; Farmer, B. L. Polymer 2006, 47, 5630–5632. (18) Min, B. M.; Lee, G.; Kim, S. H.; Nam, Y. S.; Lee, T. S.; Park, W. H. Biomaterials 2004, 25, 1289–1297. (19) Chen, C.; Cao, C. B.; Ma, X. L.; Tang, Y.; Zhu, H. S. Polymer 2006, 47, 6322–6327. (20) Zarkoob, S.; Eby, R. K.; Reneker, D. H.; Hudson, S. D.; Ertly, D.; Adams, W. W. Polymer 2004, 45, 3973–3977. (21) Foo, C. W. P.; Patwardhan, S. V.; Belton, D. J.; Kitchel, B.; Anastasiades, D.; Huang, J.; Naik, R. R.; Perry, C. C.; Kaplan, D. L. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 9428–9433. (22) Stephens, J. S.; Fahnestock, S. R.; Farmer, R. S.; Kiick, K. L.; Chase, D. B.; Rabolt, J. F. Biomacromolecules 2005, 6, 1405–1413. (23) Wnek, G. E.; Carr, M. E.; Simpson, D. G.; Bowlin, G. L. Nano Lett. 2003, 3, 213–216. (24) Li, M. Y.; Mondrinos, M. J.; Gandhi, M. R.; Ko, F. K.; Weiss, A. S.; Lelkes, P. I. Biomaterials 2005, 26, 5999–6008. (25) Matthews, J. A.; Wnek, G. E.; Simpson, D. G.; Bowlin, G. L. Biomacromolecules 2002, 3, 232–238. (26) McManus, M. C.; Boland, E. D.; Koo, H. P.; Barnes, C. P.; Pawlowski, K. J.; Wnek, G. E.; Simpson, D. G.; Bowlin, G. L. Acta Biomaterialia 2006, 2, 19–28. (27) Thomas, E.; Heine, E.; Wollseifen, R.; Cimpeanu, C.; Moller, M. Int. NonwoVens J. 2005, 14, 12–18. (28) Barnes, C. P.; Smith, M. J.; Bowlin, G. L.; Sell, S. A.; Tang, T.; Matthews, J. A.; Simpson, D. G.; Nimtz, J. C. J. Eng. Fibers Fabr. 2006, 1, 16–28. (29) Peters, T. All About Albumin: Biochemistry, Genetics, and Medical Applications; Academic Press: New York, 1995. (30) Theron, A.; Zussman, E.; Yarin, A. L. Nanotechnology 2001, 12, 384– 390. (31) White, E. T.; Tan, W. H.; Ang, J. M.; Tait, S.; Litster, J. D. Powder Technol. 2007, 179, 55–58. (32) Luo, P.; Baldwin, R. L. Biochemistry 1997, 36, 8413–8421. (33) Carter, D. C.; Ho, J. X. AdV. Protein Chem. 1994, 45, 153–203. (34) Shi, Q. S.; Zhou, Y.; Sun, Y. Biotechnol. Prog. 2005, 21, 516–523. (35) Yamada, K. Strength of Biological Materials; Williams & Wilkins Company: Baltimore, 1970. (36) Arndt, U. W.; Riley, D. P. Philos. Trans. R. Soc. London, Ser. A 1955, 247, 409–439. (37) Wang, Y.; Lopes, F.; Geil, P.; Padua, G. W. Macromol. Biosci. 2005, 5, 1200–1208. (38) Lai, H. M.; Geil, P. H.; Padua, G. W. J. Appl. Polym. Sci. 1999, 71, 1267–1281.

BM8005243