Repeated Rapid Shear-Responsiveness of Peptide Hydrogels with

ment was normalized to mean residue ellipticity (θ), ex- pressed in units of deg ... applied strain had a 0.2% amplitude at a 1-rad/s frequency. Freq...
0 downloads 0 Views 157KB Size
Download PDF
Biomacromolecules 2005, 6, 1316-1321

1316

Repeated Rapid Shear-Responsiveness of Peptide Hydrogels with Tunable Shear Modulus Sivakumar Ramachandran,† Yiider Tseng,*,‡ and Y. Bruce Yu*,†,§ Department of Pharmaceutics and Pharmaceutical Chemistry and Department of Bioengineering, University of Utah, Salt Lake City, Utah, and Department of Chemical and Biomolecular Engineering, Johns Hopkins University, Baltimore, Maryland Received November 11, 2004; Revised Manuscript Received February 20, 2005

A pair of mutually attractive but self-repulsive decapeptides, with alternating charged/neutral amino acid sequence patterns, was found to co-assemble into a viscoelastic material upon mixing at a low total peptide concentration of 0.25 wt %. Circular dichroism spectroscopy of individual decapeptide solutions revealed their random coil conformation. Transmission electron microscopy images showed the nanofibrillar network structure of the hydrogel. Dynamic rheological characterization revealed its high elasticity and shear-thinning nature. Furthermore, the co-assembled hydrogel was capable of rapid recoveries from repeated shear-induced breakdowns, a property desirable for designing injectable biomaterials for controlled drug delivery and tissue engineering applications. A systematic variation of the neutral amino acids in the sequence revealed some of the design principles for this class of biomaterials. First, viscoelastic properties of the hydrogels can be tuned through adjusting the hydrophobicity of the neutral amino acids. Second, the β-sheet propensity of the neutral amino acid residue in the peptides is critical for hydrogelation. Introduction Hydrogels are a class of viscoelastic materials that possess numerous biomedical application potentials, for instance, as encapsulation matrixes for controlled drug release or as scaffolds for tissue engineering.1-4 Hydrogels are typically made of high molecular weight synthetic polymers (e.g., derivatives of acrylic acid, ethylene oxide and vinyl alcohol) or natural biopolymers (e.g., gelatin, collagen, fibrin and chitosan).3 Hydrogels obtained from natural sources are particularly appealing since they are more likely to be biodegradable, biocompatible and bioresorbable for in vivo applications.5 The gelation of protein- or polypeptide-based polymers that are synthesized either chemically6,7 or biologically 2,8 have been extensively studied. Recent studies have focused on oligopeptides9-12 or their derivatives13 that can form physical hydrogels through self-assembly (no chemical cross-linking). Compared with proteins or polypeptides, oligopeptides can be readily obtained through straightforward solid-phase synthesis with precise control over sequence (which is not restricted to natural amino acids), chain length, stereochemistry, and derivatization (e.g., N-, C-terminal blockage). Such precise control of chemical structures allows one to fine-tune the properties of materials formed by oligopeptides. A desirable property for hydrogels to have is the ability to respond rapidly to mechanical stresses,6,14 particularly shears, in the human body. Physical hydrogels have limited * Corresponding author. E-mail: [email protected] or [email protected]. † Department of Pharmaceutics and Pharmaceutical Chemistry, University of Utah. ‡ Johns Hopkins University. § Department of Bioengineering, University of Utah.

inherent mechanical strength due to the noncovalent nature of molecular entanglements and hence will break under shear stresses. However, if the hydrogel can recover quickly after the cessation of stresses, then the limitation in mechanical strength is circumvented. Rapidly recoverable viscoelastic materials can act as scaffolds for tissue engineering as it can accommodate cell growth while still maintaining its texture. Another potential application of recoverable viscoelastic materials is as injectable encapsulation matrixes for controlled drug release at specific in vivo sites. The key to rapid recovery is a fast and reversible assembly - disassembly - reassembly process. Previous design of peptide-based hydrogels rested on the principle of selfcomplementarity, i.e., spontaneous assembly of a single peptide chain complementary to itself.6,9,12,15 We chose a modular design strategy based on mutual attraction but selfrepulsion, analogous to the design of heterodimeric coiledcoils.16 This design principle was implemented through the following pair of decapeptide modules: Positively charged peptide module Negatively charged peptide module Acetyl-WK(VK)4-amide Acetyl-EW(EV)4-amide KVW10 EVW10

The alternating charge/apolar sequence pattern originates from the discovery that self-complementary oligopeptides with such a sequence pattern can form hydrogels.9,17-20 These peptide sequences are known to adopt β-sheet type structures.9,21 Valine was chosen as the default neutral amino acid since it is known to have high propensity to form β-sheets.22-24 Also, valine is found in natural elastic biopolymers such as elastin.25 To evaluate the effect of hydrophobicity and β-sheet propensity of the neutral residue on material properties, valine was subsequently replaced by alanine, serine, and proline.

10.1021/bm049284w CCC: $30.25 © 2005 American Chemical Society Published on Web 03/26/2005

Shear-Responsive Peptide Hydrogels

The N- and C-termini of the peptides were acetylated and amidated to eliminate terminal charges that will complicate the co-assembly process. This N- and C-termini modification might also enhance their resistance against exopeptidase degradation (which recognizes the free amino- and carboxyterminal) for future in vivo applications. The separation of positively (K) and negatively (E) charged side chains into two modules makes the peptide modules mutually attractive but self-repulsive. Such mutual attraction drives the co-assembly process upon mixing the two modules. High net charge density (50% of the side chains are charged in each peptide at neutral pH) should significantly enhance the long-range attraction between the two oppositely charged modules (which also enhances the solubility of these peptides). Further, due to the repetitiveness of each module, exact sequence matching is not necessary for association. These features should lead to faster assembly rate and hence quicker recovery. Rapid recovery should also benefit from rapid peptide diffusion in the solution due to their small sizes. Additionally, self-repulsion of each module prevents uncontrolled spontaneous self-assembly and allows one to control the initiation of gelation via mixing. Mixinginduced physical gelation can preserve the pH and ionic strength of the sample and hence will benefit in vivo applications. Experimental Section Sample Preparation. The decapeptides were synthesized using standard Fmoc Chemistry26 on Rink Amide MBHA resin (which gives a C-terminal amide). The N-terminal of each peptide was acetylated by acetic anhydride as the last step of solid-phase synthesis. Each peptide was then cleaved off the resin using 90% trifluoroacetic acid with 2.5% each of ethylene dithiol, tri-isopropyl silane, water (as scavengers) and dichloromethane. The cleavage products were rotavaped and washed with ether multiple times. The products were then dissolved in H2O/CH3CN (60:40 v/v) mixture and lyophilized. The lyophilized crude peptides were purified using reverse phase (ZORBAX 300SB-C18, 9.4 × 250 mm, 5 µm) and, if needed, ion-exchange (ZORBAX 300-SCX, 9.4 × 250 mm, 5 µm) liquid chromatography (Agilent Technologies, HP1100 chromatograph system, Wilmington, DE). The molecular weight and purity of each peptide was verified by MALDI mass spectrometry and analytical HPLC, respectively. All amino acids were purchased from Novabiochem in their protected forms and used directly for solidphase synthesis without further purification. A Trp (W) residue was incorporated in each peptide as a spectroscopic probe. Each peptide sample was dissolved in 50 mM phosphate or acetate buffer at appropriate pH and dialyzed at room temperature for 2-4 h using a dialysis membrane with a molecular weight cutoff of 100 Da. The concentration of each peptide sample was determined based on the UV absorption of the Trp residue in each peptide, using an extinction coefficient of 5690 M-1 cm-1 at 280 nm,27 with light scattering corrected.28 Circular Dichroism Spectroscopy. Circular dichroism (CD) spectra were obtained using an AVIV 62DS spectro-

Biomacromolecules, Vol. 6, No. 3, 2005 1317

polarimeter equipped with a water bath operated at 25 °C. Cylindrical cell of 0.1 mm path length was used for the measurements. The instrument was calibrated using 0.06% (w/v) ammonium d-10-camphor sulfonate before use and flushed with nitrogen during operation. Ellipticity measurement was normalized to mean residue ellipticity (θ), expressed in units of deg cm2 dmole-1. Transmission Electron Microscopy. Transmission electron microscopy (TEM) images were acquired from Hitachi H7100 electron microscope operated at 75KV accelerating voltage. Each sample (a thin layer of gel or solution) was placed on a 200 mesh copper grid and negatively stained with uranyl acetate for 1 h. The samples were left to dry in a desiccator before acquiring TEM images. Rheological Charecterization. Rheological studies of the hydrogel were conducted by loading the just mixed decapeptide-pair into a 50 mm cone-and-plate module of a straincontrolled, software-operated rheometer (ARES-100; TA instrument, Piscataway, NJ), followed immediately by a series of rheometrical tests at 25 °C. For the determination of viscoelastic properties, the hydrogel sample was sequentially subjected to an 8-hour of time sweep test, a frequency sweep test (frequency from 0.01 to 10 rad/s), and a series of stress relaxation tests with a defined step-strain (with amplitudes at 0.1%, 0.2%, 0.3%, 0.5%, 0.6%, 0.8%, 1%, 1.2%, 1.5%, 2%, 5%, 10%, 20%, 50%, and 100%, respectively). In time sweep tests, the applied strain had a 0.2% amplitude at a 1-rad/s frequency. Frequency sweep test was conducted at 0.2% amplitude and the data were acquired at a log mode with 4 data points per frequency decade. A shear modulus G versus strain profile was analyzed according to previously published procedures.29 G was acquired from the stress relaxation tests at elapsed time of 0.1, 1, 10, and 100 s, respectively. After which, another 8-hour of time sweep test at 0.2% strain and 1 rad/s frequency was conducted as a recovery test. To characterize the responsiveness of the hydrogel from repeated shear-induced breakdowns, the peptide mixture was allowed to gel for 4 h (under 0.2% amplitude strain and 1 rad/s frequency) and consequently subjected to a routine of recovery cycles for more than 15 h (Figure S1). Each cycle consists of a 2 min break period with a continuous 200% sine-wave strain, and a 30 min recovery period with a constant 0.2% strain at 1 rad/s frequency. The magnitude of the shear force (200% amplitude) and the duration of the break and recovery periods (2 and 30 min, respectively) were based on results from strain sweep and recovery tests. Congo Red Dye Binding Experiment. Concentration of Congo Red dye (CR) stock solution in 50 mM phosphate or ammonium acetate buffer at pH 6 was obtained using an extinction coefficient value of 59 300 M-1 cm-1 at 505 nm.30 Peptide samples dialyzed in appropriate buffers were mixed with CR stock solution such that in each experiment the total concentration of peptide was 0.25 wt % (∼1947 µM) and that of CR was 0.0071 wt % (25 µM). Results and Discussions Circular dichroism (CD) spectra are characteristic for different peptide/protein secondary structures:31 β-sheets are

1318

Biomacromolecules, Vol. 6, No. 3, 2005

Figure 1. Far UV circular dichroism spectra of 1 wt % peptide solutions (KSW10, ESW10, KPW10, EPW10) in 50 mM phosphate buffer, pH 6, 25 °C, and 1 wt % KVW10 and EVW10 peptide solutions in 30 mM ammonium acetate buffer pH 6, 25 °C. The solubility of the KVW10 peptide was not high enough at pH 7. When valine is replaced by alanine, serine, or proline, solubility at pH 7 was no longer an issue. For this solubility reason, physical characterizations of the peptides were conducted around pH 6.

characterized by a negative band near 215 nm and a strong positive band between 195 and 200 nm; R-helices are characterized by a double minima at 208 and 222 nm and a positive band at 190 nm; type I β-turn resembles R-helix with its positive band being weaker, whereas type II β-turn resembles β-sheet spectrum which is red shifted by 5-10 nm; the random coil conformation has a negative band around 195-200 nm, whereas the polyproline II conformation, normally found in some natural biopolymers such as collagen, has an additional weak positive band centered about 215-220 nm. Since the CD spectra (Figure 1) of individual peptide solutions have a minimum near 205 nm and lack characteristics of any standard secondary structures, it can be concluded that individual peptides are in the random coil conformation. However, the polyproline II type conformation32 cannot be ruled out for KSW10 due to the presence of a weak positive band at 220 nm. Mixing of oppositely charged peptide solutions induced the formation of a viscoelastic material. Transmission electron microscopy (TEM) was used to study the morphology of the co-assembled hydrogel. TEM images of hydrogel (both 0.25 wt % and 0.5 wt %) showed a network of nanofibrillar structures extending over several micrometers. On the other hand, individual peptide solutions did not have any fibrillar structures (Figure 2). A series of rheological measurements were conducted to evaluate the viscoelastic properties of the hydrogel formed by the decapeptide pair (Figure 3). First, a time sweep experiment conducted at 1 rad/s frequency and a strain of 0.2% amplitude indicates that the elastic modulus, G′, rose to ca. 750 dyn/cm2 within a few seconds and reached a plateau value of ca. 1700 dyn/cm2 after 3 h (Figure 3a). Kinetically, the elastic modulus profile was made of an elastic burst phase on the order of seconds followed by a slower elastic growth phase on the order of a few hours (Figure 3a). On the basis of previous mechanistic studies on the gelation process of proteins,29,33 a plausible interpretation of such a biphasic kinetic profile is that the initial burst phase

Ramachandran et al.

Figure 2. Transmission Electron microscopic images of peptide samples (solution and gel) prepared in pH 6 buffer. (a) 1 wt % KVW10 solution, (b) 1 wt % EVW10 solution, (c) 0.25 wt % KVW10:EVW10 gel, (d) 0.5 wt % KVW10:EVW10 gel. The scale bar in each figure represents 1 µm.

of elasticity might involve a rapid association of the peptide modules (due to long range electrostatic interactions) to form an uneven entangled nanofibrillar network, followed by the slow reorganization phase during which the fibrils distribute more evenly in the network. Over time, the degree of homogeneity increases in the hydrogel network and the elasticity reaches its plateau. A frequency sweep conducted at a strain of 0.2% amplitude revealed that G′ was relatively independent of frequency (ω) within the frequency range of 0.01-10 rad/ sec (Figure 3b), suggesting that the hydrogel had little mobility up to 600s (t ) 2π/ ω). While the plateau value of the elastic modulus G′ is in the 1000 dynes/cm2 range, the plateau value of the viscous modulus G′′ is in the 100 dynes/ cm2 range (Figure 3b), revealing an elastic, solidlike material at 0.25 wt % total peptide concentration. To test the resilience of the hydrogel, well-spanned strains were applied 10 min apart in a step-incremental manner to observe how the hydrogel relaxes strains.29 Shear modulus plotted against strain amplitude indicated the hydrogel cannot resist a strain of over 2% and beyond this yield strain it exhibited a shear-thinning property (Figure 3c). At 100% strain, G′ dropped to 4 dyn/cm2, suggesting complete disruption of the hydrogel network. To test the recoverability of the hydrogel from shearinduced breakdowns, the completely disrupted hydrogel (by a series of step-incremental strains up to 100% amplitude) was allowed to recover for 8 h. The elasticity profile of the recovery test was almost identical to that of the initial gelation process, indicating that the shear-induced breakdown is reversible (Figure 3a). Further, just like the initial gelation process, the recovery process had a burst phase in which the gel obtained ca. 50% of its mechanical strength within a few seconds and a slower growth phase in which the gel obtained the rest of its mechanical strength within a few hours. To monitor repeated recoveries of G′, the hydrogel underwent 30 cycles of break-and-recovery with a 2 min strain break period, followed by a 30 min recovery period. On the basis of the initial gelation and first recovery data

Shear-Responsive Peptide Hydrogels

Biomacromolecules, Vol. 6, No. 3, 2005 1319

Figure 3. Viscoelastic properties of a hydrogel assembled from the KVW10:EVW10 decapeptide pair. (a) Comparison of the original gelation curve (9) with the first recovery curve (0) after the hydrogel was subjected to 100% shear (pH 6). (b) Elastic (G′) and viscous (G′′) moduli vs frequency (ω) at an applied strain γ of 0.2% (pH 6). (c) Shear modulus G vs strain γ. The arrow points to the yielding strain at which the hydrogel started to breakdown (pH 6). (d) 12 cycles of hydrogel recovery from shear-induced breakdowns (pH 5.5). The dashed line denotes 90% of the original elasticity value (G0′) after 30 min of gelation. The total peptide concentration was 0.25 wt % for all the measurements. Note that recoverability was obtained at both pH 6.0 and 5.5 with no significant difference. This pH insensitivity makes such mixing-induced hydrogels applicable over a wider range of solution conditions.

Figure 4. (a) Time sweep measurement of different peptide pairs prepared in 50 mM phosphate buffer. Filled symbols represent G′, and open symbols represent G′′. 9/0 represent the KVW10:EVW10 pair (pH 6.0), [/] represent the KAW10:EAW10 pair (pH 7.0), 1/3 represent the KSW10:ESW10 pair (pH 7.0), and b/O represent the KPW10:EPW10 pair (pH 6.0). The total peptide concentration was 0.25 wt % for all the measurements. (b) Strain sweep measurement of different peptide pairs in 50mM phosphate buffersKVW10:EVW10 (0.25 wt %, pH 6.0, 9, γyield f 2%), KAW10:EAW10 (0.25 wt %, pH 7.0, [, γyield f 0.4%), KSW10:ESW10 (1 wt %, pH 7.0, 1, γyield f 0.2%), arrows point to the yield value (γyield) of the peptide pairs.

(Figure 3a), a 2 min break period was chosen to ensure complete disruption of the hydrogel network and a 30 min recovery period was chosen in order to reach the beginning of the plateau region (so that the kinetics of the recovery process can be observed). Figure 3d shows that G′ recovered back to 90% of its original value after 12 break-recovery cycles. Even after 30 cycles, G′ recovered to more than 70% of its original value (Figure S1). In all recovery processes, the elastic burst phase was completed on the order of seconds, indicating a fast recovery of the nanofibrillar network. Such a rapid responsiveness and shear-thinning nature would be particularly beneficial in designing injectable biomaterials for controlled drug release34 and tissue engineering applications.5,35 Cell seeding into these scaffolds for cellbased therapy or tissue engineering application could be performed at ambient conditions (while mixing the two peptide modules) which eliminates the exposure of cells to harsh conditions (like organic solvents, extreme temperatures, etc.) normally involved in the fabrication of other polymeric scaffolds for tissue engineering. In addition, high porosity of these scaffold (due to the low total peptide weight fraction

in the hydrogels) is highly desirable for tissue engineering as they facilitate nutrient diffusion, tissue ingrowth and cell attachment (by providing a larger internal surface area).36 Biomaterials which are dynamically responsive to the external strain might foster the growth of tissue scaffolds by increasing the cell motility and cell-cell interactions.37 It is interesting to note that for future tissue engineering applications, cell adhesive signal motifs (like RGD from integrin or YIGSR from laminin) can be incorporated into the peptide modules in a straightforward manner to enhance the biofunctionality of the hydrogel scaffolds. To further comprehend the design principles involved in this novel class of co-assembling biomaterials, the neutral amino acid residue in the sequence was varied systematically. Substitution of valine in the sequence with the less hydrophobic alanine or serine resulted in a weaker gel with lower G′ values (Figure 4a) and resilience (Figure 4b). Even though the G′ values for the valine and alanine pairs were not significantly different, the valine pair had higher yield value (2% at 0.25 wt % peptide concentration) than the alanine pair (0.4% at 0.25 wt % peptide concentration) and the serine

1320

Biomacromolecules, Vol. 6, No. 3, 2005

Ramachandran et al.

Conclusions A pair of mutually attractive but self-repulsive decapeptides were found to form viscoelastic materials when mixed with each other at very low concentrations (0.25 wt %). The material was able to rapidly regain its mechanical strength after repeated shear-induced breakdowns. This is the first time that repeated, rapid recoveries of elastic modulus after shear-induced breakdowns have been observed in peptide hydrogels. β-Sheet propensity, more than hydrophobicity, is critically important for the formation of this class of oligopeptide-based co-assembling viscoelastic materials. Acknowledgment. The authors thank Dr. Denis Wirtz for his thoughtful discussion. This work is supported in part by National Institute of Health under Grant EB004416. S.R. acknowledges the Graduate Research Fellowship from University of Utah and Stacey L. Carrier for her help with the solid-phase peptide synthesis and HPLC purification. Supporting Information Available. 30 cycles of repeated recoveries from shear-induced breakdowns (Figure S1). This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes

Figure 5. Congo Red (CR) dye binding experiment (a) KPW10: EPW10 decapeptide pair with CR in 50 mM phosphate buffer pH 6.0 and (b) KVW10:EVW10 decapeptide pair with CR in 50mM ammonium acetate buffer pH 6.0. The total peptide concentration was 0.25 wt % (∼1947 µM), and CR concentration was 0.0071 wt % (25 µM) for all the measurements. Hyperchromic shift of the CR absorption spectrum was seen in the KVW10:EVW10 pair.

pair (0.2% at 1 wt % peptide concentration). On the other hand, substitution of valine with proline, another important component of natural elastomer-elastin25 and a dominant component in collagen, did not form a hydrogel (Figure 4a). To investigate the basis of this behavior, Congo Red dye (CR) binding experiment was performed with the valine and proline decapeptide pairs. CR is a histological stain for β-sheet type amyloid aggregates.38,39 Binding of CR to the fibrillar aggregates causes a hyperchromic red shift in the absorption spectrum of CR.30,40 On the basis of the absorption spectra, individual peptide did not bind to CR (Figure 5), in agreement with their random coil conformation as seen in CD spectra. The proline decapeptide pair did not cause hyperchromic shift in the absorption spectrum of CR (Figure 5a), whereas the valine pair caused significant hyperchromic shift (Figure 5b). The increased light scattering at higher wavelengths (600-700 nm) in the valine pair is due to the increased size of the aggregates corresponding to hydrogel formation. The results indicate that even though proline has similar hydrophobicity like valine (proline has same number of saturated carbon atoms in its side chain as valine), the KPW10:EPW10 pair nonetheless was not able to form a hydrogel due to the low β-sheet propensity of proline.41,42 Hence, β-sheet propensity is more important than hydrophobicity when designing this class of biomaterials.

(1) Hubbell, J. A. Curr. Opin. Biotechnol. 2003, 14, 551-8. (2) Petka, W. A.; Harden, J. L.; McGrath, K. P.; Wirtz, D.; Tirrell, D. A. Science 1998, 281, 389-92. (3) Hoffman, A. S. AdV. Drug DeliV. ReV. 2002, 54, 3-12. (4) Drury, J. L.; Mooney, D. J. Biomaterials 2003, 24, 4337-51. (5) Holmes, T. C.; de Lacalle, S.; Su, X.; Liu, G.; Rich, A.; Zhang, S. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 6728-33. (6) Nowak, A. P.; Breedveld, V.; Pakstis, L.; Ozbas, B.; Pine, D. J.; Pochan, D.; Deming, T. J. Nature 2002, 417, 424-8. (7) Breedveld, V.; Nowak, A.; Sato, J.; Deming, T.; Pine, D. Macromolecules 2004, 37, 3943-3953. (8) Cappello, J.; Crissman, J. W.; Crissman, M.; Ferrari, F. A.; Textor, G.; Wallis, O.; Whitledge, J. R.; Zhou, X.; Burman, D.; Aukerman, L.; Stedronsky, E. R. J. Controlled Release 1998, 53, 105-17. (9) Zhang, S.; Holmes, T.; Lockshin, C.; Rich, A. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 3334-8. (10) Ozbas, B.; Kretsinger, J.; Rajagopal, K.; Schneider, J. P.; Pochan, D. J. Macromolecules 2004, 37, 7331-7337. (11) Pochan, D. J.; Schneider, J. P.; Kretsinger, J.; Ozbas, B.; Rajagopal, K.; Haines, L. J. Am. Chem. Soc. 2003, 125, 11802-3. (12) Schneider, J. P.; Pochan, D. J.; Ozbas, B.; Rajagopal, K.; Pakstis, L.; Kretsinger, J. J. Am. Chem. Soc. 2002, 124, 15030-7. (13) Hartgerink, J. D.; Beniash, E.; Stupp, S. I. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5133-8. (14) Kopecek, J. Nature 2002, 417, 388-9, 391. (15) Aggeli, A.; Bell, M.; Boden, N.; Keen, J. N.; Knowles, P. F.; McLeish, T. C.; Pitkeathly, M.; Radford, S. E. Nature 1997, 386, 259-62. (16) O’Shea, E. K.; Lumb, K. J.; Kim, P. S. Curr. Biol. 1993, 3, 65867. (17) Caplan, M. R.; Moore, P. N.; Zhang, S.; Kamm, R. D.; Lauffenburger, D. A. Biomacromolecules 2000, 1, 627-31. (18) Caplan, M. R.; Schwartzfarb, E. M.; Zhang, S.; Kamm, R. D.; Lauffenburger, D. A. J. Biomater. Sci. Polym. Ed. 2002, 13, 22536. (19) Caplan, M. R.; Schwartzfarb, E. M.; Zhang, S.; Kamm, R. D.; Lauffenburger, D. A. Biomaterials 2002, 23, 219-27. (20) Leon, E. J.; Verma, N.; Zhang, S.; Lauffenburger, D. A.; Kamm, R. D. J. Biomater. Sci. Polym. Ed. 1998, 9, 297-312. (21) Zhang, S.; Lockshin, C.; Cook, R.; Rich, A. Biopolymers 1994, 34, 663-72. (22) Street, A. G.; Mayo, S. L. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 9074-6. (23) Rossmeisl, J.; Kristensen, I.; Gregersen, M.; Jacobsen, K. W.; Norskov, J. K. J. Am. Chem. Soc. 2003, 125, 16383-6.

Shear-Responsive Peptide Hydrogels (24) Chou, P. Y.; Fasman, G. D. Biochemistry 1974, 13, 211-22. (25) Urry, D. W.; Hugel, T.; Seitz, M.; Gaub, H. E.; Sheiba, L.; Dea, J.; Xu, J.; Parker, T. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 2002, 357, 169-84. (26) Chan, W. C.; White, P. D. Fmoc Solid Phase Peptide Synthesis: A Practical Approach.; Oxford University Press: New York: 2000; pp 1-75. (27) Gill, S. C.; von Hippel, P. H. Anal. Biochem. 1989, 182, 319-26. (28) Winder, A. F.; Gent, W. L. Biopolymers 1971, 10, 1243-51. (29) Tseng, Y.; An, K. M.; Esue, O.; Wirtz, D. J. Biol. Chem. 2004, 279, 1819-26. (30) Klunk, W. E.; Jacob, R. F.; Mason, R. P. Anal. Biochem. 1999, 266, 66-76. (31) Fasman, G. D. Circular Dichroism and the Conformational Analysis of Biomolecules, 1st ed.; Plenum Press: New York: 1996; pp 2568. (32) Bochicchio, B.; Ait-Ali, A.; Tamburro, A. M.; Alix, A. J. Biopolymers 2004, 73, 484-93. (33) Tseng, Y.; Wirtz, D. Phys. ReV. Lett. 2004, 93, 258104. (34) Hatefi, A.; Amsden, B. J. Controlled Release 2002, 80, 9-28.

Biomacromolecules, Vol. 6, No. 3, 2005 1321 (35) Kisiday, J.; Jin, M.; Kurz, B.; Hung, H.; Semino, C.; Zhang, S.; Grodzinsky, A. J. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 999610001. (36) Lanza, R. P.; Langer, R.; Vacanti, J. Principles of Tissue Engineering, 2nd ed.; Academic Press: San Diego, 2000; p 251-262. (37) Van Tomme, S. R.; van Steenbergen, M. J.; De Smedt, S. C.; van Nostrum, C. F.; Hennink, W. E. Biomaterials 2005, 26, 2129-35. (38) Abruzzo, J. L.; Gross, A. F.; Christian, C. L. Br. J. Exp. Pathol. 1966, 47, 52-9. (39) Giovannelli, L.; Scali, C.; Faussone-Pellegrini, M. S.; Pepeu, G.; Casamenti, F. Neuroscience 1998, 87, 349-57. (40) Goeden-Wood, N.; Keasling, J.; Muller, S. Macromolecules 2003, 36, 2932-2938. (41) Li, S. C.; Goto, N. K.; Williams, K. A.; Deber, C. M. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 6676-81. (42) Wood, S. J.; Wetzel, R.; Martin, J. D.; Hurle, M. R. Biochemistry 1995, 34, 724-30.

BM049284W