Injectable glycosaminoglycan hydrogels for controlled release of human basic fibroblast growth factor

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Biomaterials 26 (2005) 6054–6067 www.elsevier.com/locate/biomaterials

Injectable glycosaminoglycan hydrogels for controlled release of human basic fibroblast growth factor Shenshen Cai1, Yanchun Liu1, Xiao Zheng Shu, Glenn D. Prestwich Department of Medicinal Chemistry, The University of Utah, 419 Wakara Way, Suite 205, Salt Lake City, Utah 84108-125, USA Received 29 December 2004; accepted 7 March 2005 Available online 22 April 2005

Abstract Synthetic hydrogel mimics of the extracellular matrix (ECM) were created by crosslinking a thiol-modified analog of heparin with thiol-modified hyaluronan (HA) or chondroitin sulfate (CS) with poly(ethylene glycol) diacrylate (PEGDA). The covalently bound heparin provided a crosslinkable analog of a heparan sulfate proteoglycan, thus providing a multivalent biomaterial capable of controlled release of basic fibroblast growth factor (bFGF). Hydrogels contained 497% water and formed rapidly in o10 min. With as little as 1% (w/w) covalently bound heparin (relative to total glycosaminoglycan content), the rate of release of bFGF in vitro was substantially reduced. Total bFGF released increased with lower percentages of heparin; essentially quantitative release of bFGF was observed from heparin-free hydrogels. Moreover, the hydrogel-released bFGF retained 55% of its biological activity for up to 28 days as determined by a cell proliferation assay. Finally, when these hydrogels were implanted into subcutaneous pockets in Balb/c mice, neovascularization increased dramatically with HA and CS hydrogels that contained both bFGF and crosslinked heparin. In contrast, hydrogels lacking bFGF or crosslinked heparin showed little increase in neovascularization. Thus, covalently linked, heparin-containing glycosaminoglycan hydrogels that can be injected and crosslinked in situ constitute highly promising new materials for controlled release of heparin-binding growth factors in vivo. r 2005 Elsevier Ltd. All rights reserved. Keywords: Heparin; Hyaluronan; Chondroitin sulfate; Wound healing; Neovascularization; Synthetic extracellular matrix

1. Introduction The development of new biomaterials for tissue engineering has accelerated with the increasing demand for tissue regeneration as a substitute for organ transplantation [1]. The newer biomaterials can be derived from natural or synthetic polymers, and often growth factors or cell-surface interactive peptides are added to stimulate cell attachment, migration, and proliferation. Many of the biopolymers derived from the extracellular matrix (ECM) have been modified and adapted for medical uses. The natural polymer is first modified to contain reactive functional groups capable Corresponding author. Tel.: 801 585 9051; fax: 801 585 9053. 1

E-mail address: [email protected] (G.D. Prestwich). Contributed equally to research project.

0142-9612/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.biomaterials.2005.03.012

of further covalent bond formation, or by crosslinking to form monolithic materials that can be fashioned into meshes, sponges, fibers, and hydrogels. A variety of new materials, engineered to be slowly degraded in vivo, have been synthesized from hyaluronan (HA) [2,3], chondroitin sulfate (CS) [4–6], collagen [7], and fibrin [8,9], among other biopolymers. HA, the only non-sulfated glycosaminoglycan (GAG) and a major component of the ECM, has been chemically modified using a variety of chemistries to make new therapeutic biomaterials [10,11] and drug delivery systems [12,13]. Among the many functional modifications are amides [2], hydrazides [14,15], and thiols [16–18]. Pendant functionalities can be modified to introduce anti-cancer drugs such as taxol [19] or crosslinked as biomaterials containing anti-proliferative drugs such as mitomycin C [20] for prevention of

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post-surgical adhesions [21]. Crosslinked HA-based hydrogels incorporating the cell attaching tripeptide sequence RGD [22] have been employed for improved wound healing and tissue regeneration. CS, another component in ECM, has also been studied for skin and cartilage wound healing due to its biocompatibility and propensity for enhancing wound re-epithelialization without scarring [4,5]. Many of the chemically modified biopolymers above have been studied for growth factor delivery in promotion of tissue repair and new tissue growth. In addition, other materials employed for growth factor delivery include gelatin [23–29], alginate [30,31], ethylene-vinyl acetate [31,32], poly(glycolic acid), poly(lactic-co-glycolic acid [33–35], poly(ethylene glycol [36,37], chitosan [38], dextran [39], and poly (2-hydroxyethylmethacrylate) [40]. We have selected chemically modified HA and CS, two abundant GAGs in the ECM, to provide the required microenvironment for cell growth. Chemically modified HA and CS have become important building blocks for making medical polymers [11,41]. Recently, an injectable, in situ-crosslinkable synthetic extracellular matrix (sECM) was developed in which thiol-modified HA and CS were crosslinked with poly(ethylene glycol) diacrylate (PEGDA), producing hydrogels within 10 min [18]. These new sECM hydrogels have proven exceedingly versatile and have promising clinical applications in wound repair, adhesion prevention [21], 3-D cell culture and new tissue growth [42]. To further improve the range of applications of the in situ crosslinkable sECM, we required a material that could slowly release trophic growth factors that are required in wound healing and tissue regeneration. We selected basic fibroblast growth factor (bFGF) as a model growth factor for this purpose. bFGF, a 17-kDa polypeptide [43], is a primary promoter of cell proliferation and has applications in the treatment of wounds and bone fractures. When delivered without stabilization, bFGF diffuses rapidly, undergoes proteolysis, and consequently loses bioactivity under normal physiological conditions [31]. Thus, it is necessary to sustain its bioactivity to increase the utility of bFGF in practical treatments by developing dependable controlled release systems. Heparin (HP) is a polysulfated and heterogeneous GAG produced by mast cells to regulate thrombosis and blood vessel formation and regression [44]. HP binds bFGF to form a stable complex that maintains bFGF biological activity [45] and can retard bFGF release [2,30–32,46]. However, the current controlled release systems that incorporate HP use a sufficiently large quantity of HP that can lead to unfavorable side effects due to hemorrhage result as the HP-containing materials are degraded in vivo. We speculated that bFGF release from a multivalent, crosslinked sECM hydrogel would require much lower

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quantities of HP, thereby producing a therapeutic biomaterial with greater clinical applicability. We report herein a new method to incorporate bFGF into a hydrogel prepared by co-crosslinking small percentage of thiol-modified HP with similarly thiolated HA and CS. The HP-containing sECM hydrogels were readily prepared under sterile conditions and bFGF release from the hydrogels was maintained for over 4 weeks. In vivo studies in mice demonstrated that controlled release of bFGF increased neovascularization. These encouraging data suggest that HP-containing sECMs can act as controlled release materials for HP-binding growth factors, and that these injectable biomaterials have considerable potential for a spectrum of in vivo wound healing and tissue regeneration applications.

2. Materials and methods 2.1. Materials HA (1.5 MDa) was purchased from Clear Solutions Biotech, Inc. (Stony Brook, NY). The reagents 1-ethyl3-[3-(dimethylamino)propyl]carbodiimide (EDCI), 3,30 dithiobis(propanoic acid)(DTP), and hydrazine hydrate were from Aldrich Chemical Co. (Milwaukee, WI). CSC (from shark cartilage), Dulbecco’s phosphate-buffered saline (DPBS), bovine serum albumin (BSA), fluorescein isothiocyanate labeled human serum albumin (FITC-HSA), bovine testicular hyaluronidase (HAse, 608 U/mg) and HP sodium salt from porcine mucosa (unfractionated, Mw 15 kDa) were from Sigma Chemical Co. (St. Louis, MO). Dithiothreitol (DTT) was from Diagnostic Chemicals Limited (Oxford, CT) and 5,50 -Dithiobis(2-nitrobenzoic acid) (DTNB) was from Acros (Houston, TX). PEGDA (Mw 3400, degree of substitution 97%) was from Nektar Polymers (San Carlos, CA). Human basic fibroblast growth factor (hbFGF) was from Peprotech, Inc. (Rocky Hill, NJ), and mouse anti-hbFGF, both biotinylated and nonbiotinylated, were from R&D Systems, Inc. (Minneapolis, MN). 2.2. Synthesis of GAG-DTPH derivatives (HP-DTPH, HA-DTPH and CS-DTPH) Heparin-DTPH (HP-DTPH) was prepared from 15 kDa HP by EDCI-mediated condensation with DTP followed by DTT reduction and dialysis, as previously described for HA and CS modification [16]. HA-DTPH and CS-DTPH were synthesized as reported [16]. Structures of HA-DTPH and CS-DTPH were confirmed by 1H-NMR, with the degree of modification percent estimated from integration of the DTP methylene protons relative to the N-acetyl methyl protons.

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This technique is not applicable for HP-DTPH, since HP contains a significant degree of N-sulfation of the Nacetyl groups. Thus, the percentage of carboxylate groups modified into free thiols in HP-DTPH was determined by a modified DTNB method [47]. The calculated modifications for materials employed in this study were 56% for HP-DTPH, 42% for HA-DTPH and 63% for CS-DTPH. All products were lyophilized and stored at 80 1C. 2.3. pKa determination of GAG-DTPH The pKa values of the thiol groups in GAG-DTPH were determined using UV absorption of thiolates as previously reported [16,48,49]. HA-DTPH, CS-DTPH and HP-DTPH (2.5 mg each) were dissolved in 50 ml of DPBS solutions. UV absorptions were measured in freshly prepared solutions using a CARY 400 Bio UVvisible spectrophotometer (Varian, Palo Alto, CA) scanning from 220 to 300 nm, as the pH was gradually increased from 5.0 to 13.0 by addition of 1.0 N NaOH. 2.4. Preparation of GAG-DTPH hydrogels To prepare a range of HP-containing HA hydrogels, six different ratios of HP-DTPH:HA-DTPH (w/w) were investigated: 0:100, 1:99, 5:95, 25:75, 50:50, and 75:25. In addition, two different PEGDA concentrations (8% and 5.3%, corresponding to thiol:acrylate molar ratios of 2:1 and 3:1) were examined. The total GAG-DTPH concentration was fixed at 2% (w/v), and thus a total of 12 hydrogels were prepared. Lyophilized HA-DTPH and HP-DTPH were separately dissolved in DPBS in 2.5% (w/v) stock solutions and adjusted to pH 7.4. Two stock solutions of PEGDA, 8% and 5.3% (w/v), were prepared in DPBS. Next, HP-DTPH and HA-DTPH solutions were mixed in volume ratios of 0:100, 1:99, 5:95, 25:75, 50:50 and 75:25. Finally, the blended GAGDTPH solutions were mixed with PEGDA solutions in a volume ratio of 4:1. For the HP-containing CS hydrogels, four different ratios of HP-DTPH:CS-DTPH (w/w) (0:100, 1:99, 5:95 and 25:75) were employed. (Higher percentages of HP retarded or prevented gelation and were not further investigated.) For these materials, we evaluated three PEGDA concentrations (8%, 6% and 4.8%, corresponding to thiol:acrylate molar ratios of 3:1, 4:1 and 5:1, respectively). To compensate for the lower MW of CS relative to HA, the CS-HP gels had a GAG concentration of 3% (w/v). Thus, CS-DTPH and HPDTPH were dissolved separately in DPBS to give 3.75% (w/v) stock solutions and adjusted to pH 7.4. Three PEGDA stock solutions, 8%, 6% and 4.8% (w/v) in DPBS, were prepared. The HP-DTPH and CS-DTPH solutions were mixed in volume ratios of 0:100, 1:99,

5:95, and 25:75, and the blended solutions were further mixed with PEGDA solutions in a volume ratio of 4:1. Each of the mixtures was vortexed gently for about 20 s immediately following addition of the crosslinker. The rate of gelation was determined for different GAGDTPH blends and for different molar ratios of GAGDTPH to PEGDA. 2.5. Swelling characterization and determination of unreacted acrylate groups Hydrogels (0.5 ml) were prepared as described, soaked in DPBS at 37 1C in an orbital shaker at 150 rpm for 24 h to equilibrate to a fully hydrated state, and weighed to obtain a wet weight. The swollen hydrogels were then washed with distilled water (  3), blotted dry, and lyophilized to give dry sponge-like materials. The swelling ratio (SR) was defined as the ratio of the wet weight of swollen gel to the weight of dry gel. The unreacted acrylate groups were determined by the total acrylate moieties added minus those that had reacted with thiol groups. The total number of free thiol groups before and after gelation was determined using 2-nitro-5-thiosulfobenzoate (NTSB) [18]. The difference between the number of thiols present before and after gelation equals the reacted thiol groups. 2.6. Protein incorporation and determination of release kinetics 2.6.1. Release of FITC-HSA Hydrogels containing 0%, 1%, 5% and 25% (w/w) HP-DTPH, and PEGDA concentrations of 5.3% (w/v) for HA-DTPH gels (2% gels) and 8% (w/v) for CSDTPH gels (3% gels) were selected to evaluate FITCHSA release. FITC-HSA was dissolved in HP-DTPH solution before gelation to obtain a final concentration of 0.1% (w/v) in each of the final hydrogels (0.25 ml). Each release condition was performed in quadruplicate. Each hydrogel was soaked in 1.25 ml of DPBS, pH 7.4, at 37 1C, 150 rpm, or in 1.25 ml of 300 U/ml HAse solution in DPBS, pH 7.4. No HAse treatment was used for the CS-DTPH-based hydrogels. An aliquot of release medium (100 ml) was removed and FITC-HSA release was determined at 489 nm in 96-well plates at certain time points. DPBS-only and HAse in DPBS were used as references. Fresh release solution was added at each sampling to maintain volume and the cumulative release of FITC-HSA was plotted. 2.6.2. Release of bFGF The same hydrogel compositions tested for FITCHSA release were also used for bFGF release. For HADTPH hydrogels, release was performed using 250 ng of bFGF per 0.25 ml hydrogel in 1.0 ml release medium, with 50 U/ml HAse or without HAse. For CS-DTPH

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hydrogels, the same formulation was used, but no HAse was employed. To retain bFGF activity, BSA, sucrose, and EDTA were mixed with HP-DTPH and the pH was adjusted to 7.4; this yielded final concentrations of 0.1% BSA, 5% sucrose and 0.01% EDTA in the hydrogels [35]. (Note: when the amount of HP-DTPH was 0%, the additives, including bFGF, were mixed with HA-DTPH or CS-DTPH solution first.) Next, bFGF solutions were added together with the HP-DTPH solution to give a final bFGF amount of 250 ng per gel (0.25 ml). Hydrogels were then formed by mixing the GAGDTPH solutions with the PEGDA crosslinker solution. Release medium (1.0 ml)(DPBS supplemented with 10 mg/ml HP, 1% BSA, and 1 mM EDTA, pH 7.4) was added to the gel in a 1.5-ml polypropylene conical tube [35,43]. Subsequently, HAse (50 U/ml) was added to another set of tubes where HA-DTPH was contained in those 0.25 ml hydrogels. Each release condition was performed in quadruplicate. The release continued in 37 1C, 150 rpm agitation. At days 1, 2, 3, 7, 14, 21, 28, and 35, the release medium was sampled, stored in 1.5 ml conical tubes and immediately frozen at 80 1C until measurement. An equal volume of fresh release medium was added to maintain the total volume. 2.6.3. bFGF measurement with sandwich enzyme-linked immunosorbent assay (ELISA) bFGF titers in release medium were measured using a 96-well microtiter plate assay modified from the R&D bFGF ELISA kit. Briefly, 100 ml/well of 2 mg/ml antibFGF was immobilized on a 96-well polystyrene plate (E&K Scientific, Campbell, CA) by incubation at room temperature overnight. The plate was then washed 3  with wash buffer (DPBS with 0.05% Tween-20, pH 7.4) using 200 ml/well. Next, the wells were blocked with 200 ml/well assay block buffer (DPBS with 1% BSA, 5% sucrose and 0.05% NaN3) for 1 h at room temperature and then washed 3  . The bFGF solutions in release media were thawed and 100 ml/well was added to microtiter plate and incubated at room temperature for 1 h. A dilution series of known concentrations of human bFGF in release medium was prepared, kept frozen, and used to generate a fresh calibration curve for each day’s assays. After three additional washes, 100 ml/ well of biotin-anti-bFGF was added and incubated for another 1 h, and washed 3  . Next, 100 ml/well of a 1:1000 dilution of streptavidin-horseradish peroxidase (SA-HRP, Sigma, St. Louis, MO) was added. After 1 h incubation at room temperature, the wells were emptied and washed 3  . Finally, 100 ml 3,30 ,5,50 -tetramethylbenzidine (Sigma) was added to each well, and a blue color gradually appeared. The color-forming reaction was stopped by the addition of 100 ml/well 1 M H2SO4 when the blue color became stable. The plate was then read at 450 nm to determine the bFGF level released, with the standard bFGF samples providing the data for

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calibration. The accumulated bFGF quantities during the different release time periods were then calculated. 2.7. Cell proliferation assay for released bFGF NIH 3T3 fibroblasts (10,000/well) suspended in complete Dulbecco’s modified Eagle’s medium (DMEM) containing 10% bovine calf serum, 50 mg/ml streptomycin, and 50 U/ml penicillin, were seeded (100 ml/well) on 96-well tissue culture plates (Fisher Scientific, Pittsburgh, PA) and allowed to attach (37 1C, 5% CO2, 24 h). Then, an additional 100 ml/well of fresh complete DMEM was added, followed by the addition of 10 ml/well of the released bFGF samples or the bFGF standards in the release medium. The plates were incubated at 37 1C, 5% CO2 for 24 h. Cell numbers were determined using a Cyquant Cell Proliferation Kit (Molecular Probes, Eugene, OR) with fluorescence plate reader according to the manufacturer’s instructions. The bFGF concentration was determined from the calibration curve. The bioactivity of released bFGF was defined as: bioactivity (%) ¼ (bFGF concentration determined from cell proliferation assay/bFGF concentration determined from ELISA)  100%. 2.8. In vivo angiogenesis with bFGF loaded HA-DTPH/ HP-DTPH and CS-DTPH/HP-DTPH hydrogels 2.8.1. Preparation of bFGF-loaded GAG-DTPH hydrogels for in vivo implantation HP-DTPH proportions of 0%, 1% and 5% relative to GAG-DTPH (either HA-DTPH or CS-DTPH) were selected for in vivo experiments in order to prepare HAHP and CS-HP blended hydrogels. All hydrogels were prepared as for the in vitro studies, with a 4 mm depth in 12-well sterile culture plates (Fisher). Each component solution was sterilized through a 0.45 mm filter (Millipore, Molsheim, France) prior to mixing. Aliquots of a sterilized bFGF solution of 10 mg/ml was added to HADTPH+HP-DTPH or CS-DTPH+HP-DTPH mixture immediately prior to addition of the PEGDA crosslinker solution to give 1 mg bFGF per milliliter of hydrogel. All hydrogels formed within 10 min after gentle agitation. The plates were allowed to remain in a sterilize hood overnight at ambient temperature to allow complete gelation. 2.8.2. Surgical and experimental procedures The ability of crosslinked GAG hydrogels (with and without bFGF) to stimulate angiogenesis was evaluated in a mouse model. All procedures and experiments were carried out with the approval of the University of Utah Institutional Animal Care and Use Committee. Male Balb/c mice aged 6–8 weeks were anesthetized with intraperitoneal cavity injection of ketamine (80 mg/kg) and xylazine (10 mg/kg). Once a deep general anesthetic

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plane had been reached, the middle back area was shaved, cleaned with alcohol and Betadine, and a 15 mm midline incision was made on the dorsum. Through this incision, one subcutaneous pocket on each side was made by blunt disassociation of tissues. For the gel implantation groups, a total of eight different PEGDAcrosslinked gels were applied: (i) HA-DTPH+1% HPDTPH alone, (ii) HA-DTPH+bFGF, (iii) HADTPH+1% HP-DTPH+bFGF, (iv) HA-DTPH+5% HP-DTPH+bFGF, (v) CS-DTPH+1% HP-DTPH alone, (vi) CS-DTPH+bFGF, (vii) CS-DTPH+1% HP-DTPH+bFGF, and (viii) CS-DTPH+5% HPDTPH+bFGF. Gel discs (about 4 mm thick) were obtained from the bulk hydrogel using a 6-mm diameter Biopsy Punch (Products Corporation, Buffalo, NY), and the gel discs were then implanted into the preformed subcutaneous pockets. Therefore, each bFGFcontaining gel disc had a volume of 0.1 ml, delivering a dose of 0.1 mg of bFGF. The incision was then closed with interrupted sutures. Each mouse received two implants. Four mice were assigned to each experimental group. In addition, there were two groups of control animals in which a pocket was created but no gel was implanted; one control group had the sham surgery only, while the other control pocket received an injection of 100 ml of 1 mg/ml bFGF. The implantation or injection sites on the surface of the skin were labeled with tissue dye (IMEB INC., San Marcos, CA). On day 14 post-surgery, the mice were sacrificed in a CO2 chamber and the labeled skin, together with the underlying subcutaneous tissues and gels were excised and fixed in formalin. The tissues were then embedded in paraffin, thin-sectioned in planes perpendicular to the plane of the skin and stained with hematoxylin and eosin (H&E). Microvessels were directly counted in each sample under a conventional light microscope at 200  within the labeled areas. Identification marks on each slide were covered during counting so that the observer was not aware of the treatment group to which a given slide belonged. At this level of magnification, a very large number of fields were potentially available for observation in each sample. Consequently, 10 locations per sample were selected at random for quantification from within the labeled area; locations were selected to have vessel density representative of that sample. 2.8.3. Data analysis A total of four animals received implants in each treatment case for each of the eight treatment groups. Microvessel density data are accordingly presented as mean7standard deviation. In addition, vessel density in any implanted sample could in principle have been influenced both by the number of vessels present prior to surgery and by non-specific new vessel growth secondary to surgical probing. To account for these effects, so that the influence of HA/CS/HP and/or growth factor

on new vessel formation would be identifiable per se, we defined a dimensionless neovascularization index (NI) as treatment  mean sham , (1) mean normal In Eq. (1), ‘‘treatment’’ refers to the vessel count from each treatment, ‘‘mean sham’’ represents the mean vessel count from sham group, and ‘‘mean normal’’ is the mean vessel count from skin in the same area that had not undergone surgery [50]. Thus defined, NI represents the number of additional vessels present post-implant in a treatment group, minus the additional number due to the surgical procedure alone, normalized by the mean normal skin vessel count. Statistical significance was determined using 2-way ANOVA and post-hoc Fisher’s PLSD analysis (Stat-View 5.0, SAS Institute Inc., Cary, NC), with significance taken at the level po0:05.

NI ¼

3. Results and discussion 3.1. Synthesis of thiol-containing GAGs The carboxylate groups in the disaccharide units of GAGs were reacted with DTP, a disulfide-containing hydrazide using the carbodiimide EDCI as the condensing agent [16]. The hydrazide formation reaction, previously employed to prepare thiol-containing hydrazide derivatives of HA, CS, and gelatin [17], was also extremely efficient in producing HP-DTPH from HP (Fig. 1A), despite the high density of strongly basic Oand N-sulfates present in this GAG. The initially formed disulfide-bonded GAG-DTP was reduced with excess DTT to give the free thiol-containing HP-DTPH. After dialysis and lyophilization, the structure of HP-DTPH was confirmed by 1H-NMR. New resonances for GAGDTPH appeared at d 2.72 and 2.58, corresponding to the two side chain methylene groups that had been introduced. In Fig. 1A, these methylenes are labeled 1 and 2 in the structure and in the NMR spectrum. The integration of the four methylene protons of DTPH relative to the three N-acetyl protons that was employed for HA and CS [17] cannot be used, due to the variable degree of N-sulfation in HP. Thus, to quantify the percent modification of total carboxylates by DTPH thiol groups, the DTNB method was applied [47]. Using this spectrophotometric method, the thiol substitution percentages were 56% for HP-DTPH, 42% for HADTPH, and 63% for CS-DTPH. 3.2. pKa determination of GAG-DTPH The pKa values of thiol groups in each GAG-DTPH provide important data in evaluating the reactivity of each GAG-DTPH in the conjugate addition reaction

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Similarly, 12 different CS-based hydrogels were prepared to test the same two parameters. Blended GAGDTPH hydrogels were formed by first mixing HADTPH or CS-DTPH solutions in DPBS with an HPDTPH solution, and then crosslinking with PEGDA (Fig. 1B). The gel formation time was defined as the time required to form a solid such that no fluidity was visually observed in 1 min when the vessel was inverted [16]. The conjugate addition reactions between the free thiol and acrylate groups led to gelation within 3–33 min. Gelation time was dependent on both the ratio of HA-DTPH or CS-DTPH to HP-DTPH and on the molar ratio of thiol to acrylate groups. Table 1 summarizes the gelation time dependence on these parameters for HA-DTPH and CS-DTPH-based hydrogels. In general, slower gelation is observed with increasing amounts of HP-DTPH in the blend. Gelation is also slower at the lower molar ratio of PEGDA. The hydrogels were transparent and moderately soft. Gels with high HP-DTPH or with lower PEGDA ratios were predictably less robust. The pH dependence of gel formation on HADTPH+25% HP-DTPH and CS-DTPH+25% HP-

with PEGDA. The pKa values were determined spectrophotometrically as described previously [16,48,49]. In DPBS buffer, the pKa values were 9.05 for HA-DTPH, 9.39 for CS-DTPH and 10.37 for HP-DTPH (data not shown). This ordering of pKa values indicates that the relative reactivity of the thiol groups toward conjugate addition would be HA-DTPH4CS-DTPH4HPDTPH. This result is consistent with the presence of the basic 4-or 6-O-sulfate esters in CS-DTPH, and with the much higher sulfation (2-O-sulfation, 3-O-sulfation, 6-O-sulfation and N-sulfation) of HP-DTPH. 3.3. Hydrogel formation by conjugation addition of GAG-DTPH and PEGDA The use of HA, a non-sulfated GAG, and CS, a mono-sulfated GAG, as the basic crosslinkable component of the hydrogel was tested experimentally to determine which would be most effective in combination with crosslinkable HP for controlled release of bFGF. Thus, 12 different HA-based hydrogel compositions were selected to test two key parameters: (i) the relative percentage of HP-DTPH and (ii) the crosslinking ratio.

SH 2 1.

OSO3 H

HO O

HO O H

H

O

H O

1

O

-

H2 NHN

H O3 S NH H

OH

H OSO

HO 3

-

S

S

O

NHNH2 O HO O

EDCI, pH 4.75

H

2. DTT

x

H

NH HN O

OSO3 H

H O H

H O3 S NH H O OH H HO x H OSO 3

HP

HP-DTPH

2

1

5.5

(A)

5.0

4.5

4.0

3.5

3.0

2.5

2.0

ppm

Fig. 1. Panel A: Chemical modification of heparin to HP-DTPH, and 1H-NMR spectrum of HP-DTPH. Panel B, crosslinking of GAG-DTPH mixtures by conjugate addition using PEGDA.

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6060 HS

SH O O

HN H

NH O

O HO

OH OSO3 HO

HO

H O H OH H NH H H H H x O

+

OSO3 H

NH HN O

2-

HO O

-

H O3 S

HO

H

H

NH

H OH

O

H OSO

HO

H y

3

HP-DTPH

CS-DTPH

O S

O

O z

S

O

O

O O

O

O z

O

O

HN H

O

2-

NH O

OH OSO3 HO

HO

HO O

H O H NH H OH H H H H m O

O HO

H

OSO3H H O3 S NH H O OH H HO HO H OSO n

NH HN O

H

3

CS-DTPH

HP-DTPH

HS SH

O O

HN H O HO

NH O

HO

HOH HO

H OH H H

HO O

H

NH H H x O

+

HO O

OSO3 H

NH HN O

H O3 S

H OH

O

HO

HH

H y

HP-DTPH O S

O

HO

H OSO3 -

HA-DTPH

O

NH

O

O S

O

O

O

O

O z

O

HN

z O

H O HO

NH O

HO

HOH HO

H OH H H

(B)

HO O

H H

HA-DTPH

O

NH H m

HO O

OSO3 H H O3 S NH H O OH H HO HO H OSO3n

NH HN O

H H

HP-DTPH

Fig. 1. (Continued)

DTPH gels (2% w/v) was measured. Table 2 shows that the gelation rate increases dramatically with increased pH. This result is expected, as the rate of conjugate addition increases as the nucleophilicity or nucleophile concentration increases. Thus, at higher pH, the relative amount of thiolate anion relative to neutral thiol

increases, and thus the conjugation addition rate increases. As the sulfation of the GAG increases, the thiol-thiolate equilibrium at a given pH will shift towards the thiol form, and this can alter gelation rate and the percentage of acrylate groups reacting. At physiological pH, the rate of crosslinking is sufficiently

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Table 1 Swelling ratio and gelation time for HA-DTPH+HP-DTPH hydrogels (2%, w/v) (A) and CS-DTPH+HP-DTPH hydrogels (3%, w/v) (B) at thiol:acrylate ratios indicated (n ¼ 3) A HP-DTPH:HA-DTPH(w/w)

Gel formation time (min)

Swelling ratio

2:1

3:1

2:1

3:1

A 0:100 25:75 50:50 75:25

3 7 11 23

4 9 14 33

34.070.3 33.570.4 34.670.6 34.970.3

36.870.2 37.370 37.970.5 40.071.1

B HP-DTPH:CS-DTPH(w/w)

Gel formation time (min)

0:100 25:75

Swelling ratio

3:1

5:1

3:1

5:1

4 5.25

6.75 10

32.470.9 34.671.2

50.372.1 51.672.1

Table 2 pH dependence of HA-DTPH+HP-DTPH and CS-DTPH+HP-DTPH hydrogel formation. Gels contained 25% HP-DTPH, and different thiol:acrylate ratios indicated were tested (n ¼ 3) pH

7.0 7.4 8.0 8.5

Gel formation time (min) HA-DTPH+HP-DTPH (2% w/v in gels)

CS-DTPH+HP-DTPH (3% w/v in gels)

2:1 17 7 3 1

3:1 12 5 3 1

3:1 24 9 6 3

4:1 15 7 3 1

5:1 37 10 6 3

The pH was adjusted in each of the solutions used prior to mixing.

rapid to be useful for in situ hydrogel formation in the presence of cells, sensitive tissues, and biological agents such as growth factors. 3.4. Hydrogel characterization The SR of the different GAG-DTPH hydrogel blends are summarized in Table 1A (for HA-DTPH hydrogels) and Table 1B (for CS-DTPH hydrogels). With increasing amounts of HP-DTPH, a modest increase in SR for the hydrogels was observed. There was also a significant difference (po0:01) between hydrogels with different thiol:acrylate ratios, with less crosslinked gels having greater swellability. All hydrogels had very high water content (defined as (SR-1)/SR) exceeding 97%, thereby facilitating the exchange of nutrients and small hydrophilic molecules between the hydrogels and the cells. In addition, we calculated the unreacted acrylates in each of the hydrogels by determining consumed thiols using NTSB method [18,51]. Conjugate addition proceeded to completion in HA-DTPH hydrogels, with no

remaining acrylates were detected. In CS-DTPH hydrogels, some unreacted acrylates remained, but this fraction decreased as the fraction of HP-DTPH level decreased. Thus, 15% and 4% unreacted acrylates were found, respectively, in the 5% and 1% HP-DTPHcontaining CS gels (data not shown). These results can be rationalized based on the rank order of thiol reactivity of GAG-DTPHs: HP-DTPHoCSDTPHoHA-DTPH. As shown in Section 3.2, this reactivity correlates inversely with the pKa values: HPDTPH4CS-DTPH4HA-DTPH. Using the lowest possible HP content would keep unreacted acrylate groups to a minimum, and would also reduce any possibility of hemorrhagic toxicity following in vivo implantation. 3.5. FITC-HSA release with hydrogel swelling The hydrogels selected above were tested for their ability to release FITC-HSA, a 66-kDa protein. Fig. 2A shows the release over a 5 day period for CS-DTPH

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80

60

95% CS-DTPH, 5% HP-DTPH 99% CS-DTPH, 1% HP-DTPH

40

100% CS-DTPH 20

0 0

20

40

(A)

60 80 time (hour)

100

120

140

100

95% HA-DTPH, 5% HP-DTPH

80

99% HA-DTPH, 1% HP-DTPH

60

100% HA-DTPH

40

95% HA-DTPH, 5% HP-DTPH, 300 U/m HAse 99% HA-DTPH, 1% HP-DTPH, 300 U/m HAse 100% HA-DTPH,3% U/ml HAse

20

0 0 (B)

20

40

60 80 time (hour)

100

120

140

Fig. 2. FITC-HSA release from (A) CS-DTPH+HP-DTPH and (B) HA-DTPH+HP-DTPH blended, PEGDA-crosslinked hydrogels in DPBS. For HA-based materials, release was measured in the absence (open symbols) and presence (filled symbols) of 300 U/ml HAse.

hydrogels with different percentages of HP-DTPH. Fig. 2B shows the analogous data for the HA-DTPH hydrogel blends. To simplify the presentation, the data for the 25% HP-DTPH hydrogels are not shown, but the curve for that data overlaps that for the 5% HPDTPH gel. No enzyme was used to facilitate gel degradation in CS-DTPH hydrogels. Within each group, plots of varying HP-DTPH amounts were all overlapping, indicating that the HP-DTPH amounts did not significantly affect the density of gel network or the release of FITC-HSA. The initial burst of FITC-HSA release likely corresponds to the rapid diffusion of molecules from the surface layer of the hydrogel. We added 300 U/ml bovine testicular HAse to HADTPH hydrogels to accelerate gel degradation. The initial burst of FITC-HSA release overlapped with the no-enzyme curve for the first 12 h, supporting the hypothesis that this release arises from an undegraded surface layer. The HAse action on crosslinked gels begins with a slow surface erosion, as slow enzymatic cleavage of the crosslinked HA chains is accomplished. Gradually, the pitting and gradual depolymerization of

the hydrogel network permitted a steady release of FITC-HSA. Finally, after 5 days, about 80% of the labeled HSA had been released under enzymatic conditions, while approximately 50% of the HSA was released in the absence of enzyme. This indicated that the hydrogels could swell and hydrolyze slowly even under non-enzymatic conditions to permit release of FITC-HSA. Again, no dependence on the amount of HP-DTPH was observed. Compared to the 3% w/v CSDTPH hydrogels, the 2% w/v HA-DTPH gels had a slower release rate; this could be attributed to higher MW of the HA and increased entanglement of polypeptide and polysaccharide chains as crosslinking occurred. 3.6. bFGF release from HP-containing GAG hydrogels The selected hydrogels above (HP-DTPH 25%, 5%, 1% and 0%) were next tested for the controlled release of bFGF in vitro. Recombinant human bFGF solution in DPBS at pH 7.4 was used. As described in the Materials and Methods, several additives were required

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in the release buffer to maintain bFGF activity and prevent surface adsorption during the experimental release period. In the hydrogels, 0.01% EDTA was used to prevent trace metal-induced disulfide exchange between the hydrogels and bFGF, 5% sucrose was added to maintain bFGF structure in solid state and control the viscosity of aqueous pores in the polymers. In addition, 0.1% BSA was added to prevent bFGF adsorption to plastic surfaces. In the release medium, 10 mg/ml HP was added to maintain and sequester bFGF activity after it was released, 1% BSA was added to prevent adsorption, and 1 mM EDTA was added as a chelator [35]. When release was carried out in 0.25-ml gels that contained 250 ng of bFGF, we observed that bFGF was released over a 35-day period by crosslinked HP-DTPH in both HA-DTPH and CS-DTPH hydrogels (Fig. 3 and Table 3). For HA-DTPH gels (Fig. 3A), the release burst was reduced by as little as 1% HP-DTPH. HAse digestion only accelerated bFGF release when HPDTPH was absent, which again supported that bFGF

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binding by HP is the primary factor controlling bFGF release from the hydrogels. The accumulated release with over 35 days approached 96% (with HAse digestion) and 76% (without HAse digestion) for 0% HP-DTPH, 65% for 1% HP-DTPH, 41% for 5% HPDTPH and 33% for 25% HP-DTPH hydrogels. Similar results were obtained using CS-DTPH+HPDTPH hydrogels (Fig. 3B); note that the data for 25% HP-DTPH overlapped with the data for 5% HP-DTPH Table 3 Percent bFGF released from 0.25 ml crosslinked GAG hydrogels HP-DTPH % (w/w)

25 5 1 0

Released percent of bFGF over 35 days HA-DTPH+HP-DTPH (%)

CS-DTPH+HP-DTPH (%)

33 41 65 76

39 43 71 92

120

95% HA-DTPH, 5% HP-DTPH

100

99% HA-DTPH, 1% HP-DTPH

80 100% HA-DTPH 60 40 20 0 0

10

(A)

20 time (day)

30

40

95% HA-DTPH, 5% HP-DTPH, 50 U/ml HAse 99% HA-DTPH, 1% HP-DTPH, 50 U/ml HAse 100% HA-DTPH, 50 U/ml HAse

100 80

60

95% CS-DTPH, 5% HP-DTPH

40

99% CS-DTPH, 1% HP-DTPH 100% CS-DTPH

20 0 0 (B)

10

20 time (day)

30

40

Fig. 3. bFGF release from (A) HA-DTPH+HP-DTPH and (B) CS-DTPH+HP-DTPH blended hydrogels crosslinked with PEGDA. Each experiment used 250 ng bFGF per 0.25 ml gel, release in 1.0 ml medium in 1.5 ml conical tubes, n ¼ 4. For HA-based materials, release was measured in the absence (open symbols) and presence (filled symbols) of 300 U/ml HAse.

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and is not shown. As little as 1% HP-DTPH substantially controlled bFGF release, in contrast to hydrogels without HP-DTPH. Additional amounts of HP-DTPH above 5% did not significantly slow the release of bFGF. The accumulated release percentage at 35 days was approximately 92% without HP-DTPH, 71% for 1% HP-DTPH, 43% for 5% HP-DTPH and 39% for 25% HP-DTPH hydrogels (Table 3). CS-DTPH hydrogels showed a slightly higher release than HA-DTPH hydrogels, a result that can be attributed to the lower MW of CS. However, these release data are consistent with other bFGF controlled release systems that have been reported [2,27,31,35]. The possibility was considered that the free thiols in the hydrogel could form disulfide linkages with the cysteine residues of bFGF, since there are two nondisulfide bonded cysteines contained in the bFGF structure [52,53]. To rule out the possibility that it was the unreacted thiols in HP-DTPH rather the covalently crosslinked HP that binds to bFGF, unmodified HP was added to HA-DTPH gels that lacked crosslinkable HPDTPH. The physical entrapment of HP within an HADTPH-PEGDA hydrogel resulted in a similar retention of bFGF (data not shown), indicating that it was HP itself, not the thiols of HP-DTPH that were responsible for the retention of bFGF. It was also possible that the bFGF Cys residues could become covalently crosslinked via the PEGDA conjugate addition reaction. Two observations appear to minimize this possibility. First, the thiol residues of the GAG-DTPH are in enormous excess over the amount of bFGF present, and effectively act as a buffer for consuming the reactive PEGDA. Second, the fact that bFGF is released in free, biologically active, and immunoreactive form in vitro and in vivo (see below) argues for the fact the majority of bFGF is not covalently bound in the hydrogel, but simply retained by the high affinity HP-bFGF interaction.

3.7. Bioactivity of released bFGF in cell proliferation assay To examine the biological activity of released bFGF, a cell proliferation assay was performed using NIH 3T3 fibroblasts. Released bFGF samples collected at days 3, 14 and 28 from 0.25 ml gels in 1.0 ml medium were used. Although some loss of bioactivity did occur during release, Fig. 4 demonstrates that over 50% of the bioactivity of bFGF was maintained throughout the release period for hydrogels for different components and release conditions. There was no real difference between CS-DTPH and HA-DTPH hydrogels in maintaining the biological activity of released bFGF using this assay. Moreover, similar results were achieved for release from HA-DTPH gel with and without HAse.

140 CS-DTPH + HP-DTPH 120

HA-DTPH + HP-DTPH HA-DTPH + HP-DTPH, 50 U/ml HAse

100 bioactivity (%)

6064

80 60 40 20 0 Day 2-3

Day 7-14

Day 21-28

Fig. 4. Bioactivity of bFGF samples released. Bioactivity (%) ¼ (concentration determined from cell proliferation assay/concentration determined from ELISA)  100%.

Regression analysis showed that the correlation between the ELISA and the NIH 3T3 cell proliferation assay was significant with R2 ¼ 0:778 (data not shown). This further supports the notion that bioactivity promoting cell proliferation was due to the bFGF released from the hydrogels. 3.8. Controlled release of bFGF from in vivo implanted hydrogels improved vascularization in wound healing Hydrogels of different compositions that contained or lacked bFGF were surgically implanted in subcutaneous pockets in the backs of Balb/c mice. The implanted hydrogels (HA-DTPH+HP-DTPH and CS-DTPH+ HP-DTPH) that contained bFGF showed greatly increased neovascularization as shown in Fig. 5, in which the black arrows indicating newly formed capillaries. As seen in the histology image in Fig. 5 and the data summary in Fig. 6, no new blood vessels were formed in the sham surgery (surgical pocket creation only) after 2 weeks, and very few new capillaries were formed in the experimental hydrogels lacking bFGF. Importantly, the injection of a bolus of bFGF equivalent to that contained within the hydrogel implants did not promote the neovascularization significantly. However, when HA-DTPH+HP-DTPH+ bFGF or CS-DTPH+HP-DTPH+bFGF hydrogels were implanted in vivo, the production of new blood vessels was dramatically increased (Fig. 6). The NI in HA-DTPH+HP-DTPH+bFGF group was in the rank order: 1% HP-DTPH45% HPDTPH40% HP-DTPH. This indicates that covalently crosslinked HP was necessary to enhance bFGF activity and promote neovascularization, but too high a level of crosslinked HP appeared to retain bFGF with too high an avidity. The NI data is consistent with our in vitro analysis of controlled release of bFGF.

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Fig. 5. H&E stained representative images of skin and subcutaneous tissue for different implant types, day 14 post-surgery: (a) sham surgery; (b) aqueous bFGF bolus only; (c) CS-DTPH-PEGDA hydrogel containing bFGF; (d) CS-DTPH+HP-DTPH (1%) hydrogel; (e) CS-DTPH+HPDTPH (5%) hydrogel with bFGF; and (f) HA-DTPH+HP-DTPH (1%) hydrogel with bFGF. Note the greatly increased microvessel density along with many extravasated red cells in the CS-bFGF-HP (5%) hydrogel and HA-bFGF-HP (1%) hydrogel treated skin. Key: H ¼ hair follicle Ca ¼ capillary. Scale bar: 100 mm.

FG +b

+b he % S+ 5 C

1%

he

pa

pa

rin

rin

S+ C C S+

F

F

G bF

pa he 1%

FG

F

rin

F +b C S+

H A+

5%

he

pa

rin he

pa

rin

+b

bF A+ 1%

FG

F

G

FG

F

rin H H A+

H A+

1%

he

pa

G F bF

Sh a

m

14 12 10 8 6 4 2 0

Treatment Fig. 6. Neovascularization index (NI) at day 14 post-surgery. Results are shown for surgery without implant placement (sham), aqueous bFGF, and GAG-DTPH-PEGDA films with or without bFGF as indicated. Values shown are mean7s.d., n ¼ 8. NI is defined in the text, Eq. (1).

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Interestingly, in CS-DTPH+HP-DTPH+bFGF group, the rank order for the NI increase was different: 5% HP-DTPH41% HP-DTPH40% HP-DTPH. Again, crosslinked HP maintains bFGF bioactivity throughout the 2 week in vivo experiment. Although the higher dose of released bFGF in vitro was not consistent with the lower NI in vivo, we inferred, that because of the much faster degradation of CS-DTPH hydrogels (more than 90% degraded within 2 weeks) than HA-DTPH hydrogels (about only 70% degraded) in vivo, the amount of bFGF released from CS-DTPH hydrogels may be greater than from HA-DTPH gels. Therefore, higher amount of HP (i.e. 5%) could better help maintain the bioactivity of released bFGF than lower amount of HP (i.e. 1%) in this case. This was consistent with the result that CS-DTPH hydrogels promoted neovascularization better than HA-DTPH hydrogels (Fig. 6), which was probably attributed to the faster degradation of CSDTPH gels in vivo and the following greater amount of released bFGF. As a whole, we regarded CS-DTPH with 5% HP-DTPH was the most efficient to stimulate in vivo neovascularization (po0:05).

4. Conclusions We describe herein the synthesis and evaluation of GAG-based hydrogels that slowly release bFGF as novel biomaterials for wound repair. Several different hydrogels were characterized in terms of SR, crosslinking density, FITC-HSA release rates, in vitro bFGF release rates, and in vitro biological activity of released immunoreactive bFGF. In addition, HA-DTPH or CSDTPH hydrogels supplemented with different doses of co-crosslinked HP-DTPH, and incorporating bFGF were surgically implanted into mice to correlate the bFGF release in vitro with promotion of neovascularization promotion in vivo. We found that bFGF dramatically stimulated blood vessel formation when it was released from a crosslinked GAG hydrogel. The optimal result in vivo was obtained from CS-DTPH with 5% HP-DTPH hydrogel, which provided a 10fold increase in the rate of neovascularization when compared to controls. These hydrogels degrade within 1 month and produced only a minor inflammatory response. Taken together, these results demonstrate considerable potential for use of bFGF-containing injectable GAG-based hydrogels for controlled release of bioactive growth factors during clinical tissue reconstruction.

Acknowledgement This project was supported by NIH R01 DC04663 to the late S.D. Gray and to G.D.P. Additional funding to

G.D.P was provided by the University of Utah, and by the Center for Therapeutic Biomaterials, a member of the Utah Centers of Excellence Program. We thank Drs. J. Shelby, R. Peattie, and K. R. Kirker for helpful discussions and consultation in experimental design. Reference [1] Palsson B, Hubbell J. Tissue engineering. CRC Press: Boca Raton; 2003. [2] Liu L, Ng C, Thompson A, Poser J, Spiro R. Hyaluronateheparin conjugate gels for the delivery of basic fibroblast growth factor (FGF-2). J Biomed Mater Res 2002;62:128–35. [3] Luo Y, Kirker K, Prestwich G. Cross-linked hyaluronic acid hydrogel films: new biomaterials for drug delivery. J Control Release 2000;69:169–84. [4] Kirker KR, Luo Y, Nielson JH, Shelby J, Prestwich GD. Glycosaminoglycan hydrogel films as bio-interactive dressings for wound healing. Biomaterials 2002;23:3661–71. [5] Kirker KR, Luo Y, Morris SE, Shelby J, Prestwich GD. Glycosaminoglycan hydrogel films as supplemental wound dressings for donor sites. J Burn Care Rehab 2004;25:276–86. [6] Gilbert ME, Kirker KR, Gray SD, Ward PD, Prestwich GD, Orlandi RR. Chrondroitin sulfate hydrogel and wound healing in rabbit maxillary sinus mucosa. Laryngoscope 2004;114:1406–9. [7] Yamauchi K, Takeuchi N, Kurimoto A, Tanabe T. Films of collagen crosslinked by S-S bonds: preparation and characterization. Biomaterials 2001;22:855–63. [8] Sakiyama-Elbert S, Hubbell J. Development of fibrin derivatives for controlled release of heparin-binding growth factors. J Control Release 2000;65:389–402. [9] Sakiyama-Elbert S, Hubbell J. Controlled release of nerve growth factor from a hepairn-containing fibrin-based cell ingrowth matrix. J Control Release 2000;69:149–58. [10] Prestwich GD, Luo Y, Kirker KR, Ziebell MR, Shelby J. Hyaluronan biomaterials for targeted drug delivery and wound healing. In: HA 2000. Abington, UK: Woodhead Publishing Ltd.; 2002. p. 277–84. [11] Shu XZ, Prestwich GD. Therapeutic biomaterials from chemically modified hyaluronan. In: Garg H, Hales C, editors. Chemistry and Biology of Hyaluronan. Oxford, UK: Elsevier Ltd.; 2004. p. 475–504. [12] Vercruysse KP, Prestwich GD. Hyaluronate derivatives in drug delivery. Crit Rev Ther Drug Carr Syst 1998;15:513–55. [13] Luo Y, Prestwich GD. Novel biomaterials for drug delivery. Exp Opin Ther Patents 2001;11:1395–410. [14] Prestwich GD, Marecak DM, Marecek JF, Vercruysse KP, Ziebell MR. Controlled chemical modification of hyaluronic acid: synthesis, applications, and biodegradation of hydrazide derivatives. J Control Release 1998;53:93–103. [15] Pouyani T, Prestwich GD. Functionalized derivatives of hyaluronic acid oligosaccharides: drug carriers and novel biomaterials. Bioconjug Chem 1994;5:339–47. [16] Shu XZ, Liu Y, Luo Y, Roberts MC, Prestwich GD. Disulfide cross-linked hyaluronan hydrogels. Biomacromolecules 2002;3:1304–11. [17] Shu XZ, Liu Y, Palumbo FS, Prestwich GD. Disulfide-crosslinked hyaluronan-gelatin hydrogel films: a covalent mimic of the extracellular matrix for in vitro cell growth. Biomaterials 2003;24:3825–34. [18] Shu XZ, Liu Y, Palumbo FS, Luo Y, Prestwich GD. In situ crosslinkable hyaluronan hydrogels for tissue engineering. Biomaterials 2004;25:1339–48. [19] Luo Y, Prestwich GD. Synthesis and cytotoxicity of a hyaluronic antitumor bioconjugate. Bioconjug Chem 1999;10:755–63.

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