Fibrous protein-based biomaterials (silk, keratin, elastin, and resilin proteins) for tissue regeneration and repair

Fibrous protein-based biomaterials (silk, keratin, elastin, and resilin proteins) for tissue regeneration and repair

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F. Costa*,a, R. Silva†,a, A.R. Boccaccini† i3S - Instituto de Investigação e Inovação em Saúde, University of Porto, Porto, Portugal, † University of Erlangen-Nuremberg, Erlangen, Germany

*

7.1 Introduction Tissue engineering aims to restore or regenerate damaged tissue by combining cells derived from a patient biopsy with engineered biomaterial scaffolds that provide a temporary extracellular matrix (ECM) for the cells to attach and to proliferate [1]. At the same time, these scaffolds may also serve as carriers for growth factors, other enzymes, or drugs [2]. To do so, different types of materials have been developed or used, including foams, microsphere scaffolds, hydrogels, fibrous structures, and ­polymer–bioceramic composites among many others. In recent years, fibrous protein-based hydrogels have become popular [3,4] due to their structural and mechanical similarity with the native ECM and their relatively simple processing ability under mild, cell-compatible conditions [4]. Hydrogels are attractive as a cell matrix and as connective tissue substitutes due to their ability to form mechanically stable, porous, hydrated 3D polymer networks that facilitate the transport of nutrients and cell metabolic waste products [1,4]. Moreover, hydrogels can be formed in vivo and are therefore compatible with minimally invasive surgery methods: a liquid precursor solution together with suspended cells can be injected at the site of interest, and therefore the hydrogel-forming polymerization process takes place in the body [5–7]. A great variety of structural building blocks for hydrogel fabrication is available that can be derived from mechanically stable proteins, including silk fibroin from spider webs; collagen from skin, bone, and tendons; keratin from wool or hair; elastin from elastic tissues; fibrin from blood clots; resilin from insect tendons. Each of these biological materials shows unique properties unmatched by known technical materials. The present chapter gives an overview of the protein structure, basic fabrication principles, and properties of fibrous protein-based hydrogels and their applications in tissue engineering. In particular silk fibroin, keratin, elastin, and resilin will be discussed in detail. However, we will not discuss collagen and fibrin here since these are the focus of a

Both authors contributed equally to the manuscript.

Peptides and Proteins as Biomaterials for Tissue Regeneration and Repair. https://doi.org/10.1016/B978-0-08-100803-4.00007-3 Copyright © 2018 Elsevier Ltd. All rights reserved.

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other chapters of this book. The present chapter reproduces sections of a previous review paper [8], but it has been updated and adapted to fit the scope of the present volume.

7.2 Biopolymer-gels based on fibrous proteins: General considerations A wide range of natural materials can form non-cytotoxic polymeric hydrogels [7]. These natural polymers can be classified into proteins (i.e., silk, collagen, gelatin, fibrinogen, elastin, keratin, actin, and myosin), polysaccharides (i.e., cellulose, amylose, alginate, dextran, chitin, and glycosaminoglycan’s), or polynucleotides (i.e., DNA and RNA) [6,9]. In particular, protein-based hydrogels can mimic features of the ECM of human tissues and thus have the potential to promote the migration, growth, and organization of cells during tissue regeneration and wound healing. Protein-based hydrogels are therefore also often suitable materials for cell encapsulation [3,7]. Fibrous proteins, such as collagens, elastins, silks, and keratins are characterized by highly repetitive amino acid sequences that give these proteins unique mechanical and architectural properties [3,10]. These repetitive amino acid sequences result in the formation of relatively homogeneous secondary structures (e.g., β-pleated sheets, coiled coils, or triple helices), which in turn promote the spontaneous polymerization of protein monomers that self-assemble into structurally interesting hierarchical materials [11]. Furthermore, fibrous proteins are attractive materials for designing bioactive scaffolds, because cells can recognize and bind to specific sites within proteins, as well as secrete enzymes that may degrade specific amino acid sequences [12]. In the next sections, some of the most investigated fibrous proteins in tissue engineering scaffolds are discussed.

7.3 Silk fibroin Silks are naturally occurring proteins that can be found in a wide variety of insects and spiders. The most widely used and characterized silks are from the domesticated silkworm (Bombyx mori) and from some spiders (Nephila clavipes and Araneus diadematus) [13,14]. Silk proteins are usually produced within specialized glands. After biosynthesis by epithelial cells that line the glands, proteins are secreted into the lumen of the gland, and then spun into fibers [15]. Spider silk is lightweight, extremely strong, and elastic, and exhibits mechanical properties even higher than certain polymers such as kevlar [16]. Spider silk is spun at near room temperature and ambient pressure using water as the solvent, which makes it environmentally safe and noncytotoxic [17].

7.3.1 Protein structure Silk obtained from silkworms is mainly composed of two classes of proteins, fibroin, and sericin. Fibroin filaments are organized as bundles of nanofibrils corresponding

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to 70%–75% of the silk fiber weight. The remaining mass is sericin (20–310 kDa), a water-soluble glue-like protein that binds the fibroin fibers together. The B. mori fibroin is typically a dimer composed of a light chain (≈26 kDa) and a heavy chain (≈390 kDa), which are present in a 1:1 ratio and are linked by a single disulfide bond [15,18]. Sericins consist mainly of glycine (44%), alanine (29%), and serine (11%) [18]. In general, the primary sequence of fibroin consists of highly repetitive motifs. For B. mori, the primary repetitive sequence is the hexapeptide GAGAGS, or other g­ lycine-X repeats, with X being alanine, serine, threonine, and valine [18] (see Fig. 7.1). Protein coat

Fibroin

Sericin

(A)

β-Sheet crystalline domain

Degumming

Amorphous domain

(B) N-terminus

Heavy chain (H-chain)

C-terminus

G A G A G S G A G A G Y G A G A G A G A G V G A

Hydrophobic repetitive domains

(C)

Hydrophilic repetitive domain

Fig. 7.1  A schematic illustration of silk fibers produced by silkworms: (A) the raw silk fiber is composed of two fibroin fibers held together with sericin covered with a protein coat. After degumming, the removal of sericin, the fibroin fibers are dissolved in solution; (B) the illustration of β-sheet crystallite embedded in the amorphous matrix of silk fibroin fibers; and (C) each silk fibroin heavy chain (H-chain) consists of hydrophobic and hydrophilic repetitive domains. Reprinted with permission from Jao D, Mou X, Hu X. Tissue regeneration: a silk road. J Funct Biomater 2016;7.

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These predominant hydrophobic blocks induce extensive hydrogen and hydrophobic interactions throughout the protein chains resulting in homogeneous secondary structure (β-sheet). When the methyl groups and the hydrogen groups from the opposing β-sheets are interacting, they form antiparallel β-sheet stacks. These interactions are responsible for the crystallinity of the protein, which in turn is responsible for increased environmental stability [19,20]. The protein conformation of silk fibroin in the solid state can assume two polymorphs, the glandular state prior to crystallization/ spinning (silk I), and the spin silk state with a secondary structure (silk II) [19]. When methyl and hydrogen moieties from opposing β-sheets are interacting, they form antiparallel β-sheet stacks (silk II). The silk I structure is water soluble and, upon exposure to heat, organic solvents, or physical treatments, can be easily converted to silk II structure, which is water insoluble but can be dissolved by several chaotropes [21]. Recent studies further revealed that the nanoscale confinement of β-sheet nanocrystals as well as other semi-amorphous structures in silks have a fundamental role in achieving relatively high stiffness, resilience, and fracture toughness on a macroscale [22].

7.3.2 Extraction and purification Fibroin has been widely used to produce materials for medical applications. Silk proteins can be extracted from silk glands or silkworm cocoons (see Fig. 7.2). Typically, the fibroin is extracted from the silkworm cocoon by removal of sericin [23] and is then purified. There are several methods to extract and purify silk fibroin protein. One of the most widely used procedures for the removal of sericin (“degumming”) is sodium carbonate boiling and/or autoclaving. The fibroin is then extracted from the degummed silk by dissolving it in a concentrated solution of lithium bromide [24–26] or sometimes in a ternary solvent system of calcium chloride/ethanol/water [27–29]. After evaporation of the solvent, the fibroin can be further purified by dissolving it in 1,1,1,3,3,3-hexafluoro-2-propanol [22,29,30] and formic acid [28,31]. The relative ease with which silk proteins can be processed in water or various solvents to form gels, fibers, or sponges with different chemical functionalization, together with their excellent biocompatibility, enzymatic degradability, and mechanical resilience, make these proteins interesting candidates for many biomedical applications [21].

7.3.3 Hydrogels formation Predominance of hydrophobic amino acids such as glycine, serine, and alanine in ­fibroin makes gelation possible without addition of any gelling agent [32,33]. However, this process has a very slow rate, challenging the preparation of silk gels. This l­ imitation can be surpassed by stimulating fibroin gel formation by changing temperature, pH, or ionic concentration by addition of salts such as CaCl2 or KCl, and by using several methods such as shearing, sonication, removal of bulk water by osmotic stresses, vortexing, heating, and exposure to solvents, gases and surfactants such as sodium dodecyl sulfate and sodium N-lauroyl sarcosinate [34,35]. Generally, gelation time decreases with an increase in fibroin concentration, temperature, and concentration of additives such as Ca2+, or with a decrease in pH [22,32,35]. The pore size of the hydrogel ­decreases

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A. mylitta (Non-mulberry silkworm)

larvae

Cocoons

larvae

Cocoons

Isolated silk glands

Cut pieces of cocoon

Isolated silk glands

Cut pieces of cocoon

Glands are pierced and proteins are squeezed out in buffer

Fibroin solubilized in 1%SDS or 9.3M LiBr followed by dialysis

Sericin is removed by boiling in 0.1M Na2CO3 solution or hot water

Degummed fibers dissolved in 9.3M LiBr and then dialyzed

Glands are pierced and proteins are squeezed out in buffer

Sericin is removed by boiling in 0.1M Na2CO3 solution or hot water

Fibroin protein is solubilized in 1%SDS followed by dialysis

Degummed fibers dissolved in 9.3M LiBr and then dialyzed

Fibroin solution Gelation is induced by treatments like change in pH, temperature, shear forces or chemical crosslinking

Regenerated fibroin solution in teflon mold

Fibroin hydrogel (prototype image)

Fig. 7.2  Schematic representation of hydrogel fabrication from silk fibroin protein: Cocoons and the silk glands are main sources of silk fibroin proteins. Larvae and cocoons of domesticated mulberry silkworm, B. mori, and non-mulberry tropical tasar silkworm, Antheraea mylitta are shown. Silk fibroin protein is isolated from middle silk glands and cocoons of different silkworms and is regenerated. The regenerated silk fibroin protein is processed through various treatments to obtain hydrogels. Reprinted with permission from Kapoor S, Kundu SC. Silk protein-based hydrogels: promising advanced materials for biomedical applications. Acta Biomater 2016;31:17–32.

and the mechanical strength and stiffness increase with higher fibroin concentration or gelation temperature [30,32,34]. Depending on the conditions applied, different mechanical properties are obtained, which means that is possible to tune the gelation process regarding the final application of the gel. For instance, for cell encapsulation it is preferable to use mild temperature and pH, therefore applying ultrasonication or vortexing may be a way to obtain cell homogeneous distribution [30,36,37]. For example, human bone marrow-derived mesenchymal stem cells (hMSCs) have been mixed into 4% fibroin solutions after sonication, followed by rapid gelation. Subsequently, the cells proliferated in the gels over 21 days [37].

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In addition, the stiffness of these preformed hydrogels can recover quickly following injection through a needle, allowing application of these hydrogels as injectable cell delivery scaffolds.

7.3.4 Applications in tissue repair and regeneration Mechanical strength, biocompatibility, stability to heat and humidity, high permeability to oxygen, and other molecules along with the possibility to tune the silk ­protein-based characteristics, are some of the features that explain the potential of silk fibroin for biomedical purposes [22,26,34]. Moreover, as it is a hydrogel that has the ability to entrap cells during fabrication (mild conditions), providing a porous matrix (in situ formed), facilitating ease of nutrient/wastes transportation, allowing intercellular communication, and slow degradation in vivo, it has particular interest for tissue regeneration strategies [26]. Today the most important applications of silk protein-based materials in bioengineering includes (i) controlled drug delivery, (ii) cell culture studies, and (iii) tissue engineering strategies. Regarding controlled drug delivery, it has been extensively studied both as bulk material used for the drug release, and as a coating for nanoparticles [38]. For example, Wang et al. produced Chinese oak tasar Antheraea pernyi silk fibroin (ApF) nanoparticles loaded with differently charged small molecule drugs, such as doxorubicin hydrochloride, ibuprofen, and ibuprofen-Na, by simple absorption based on electrostatic interactions [39]. The release behavior of the compounds from the nanoparticles demonstrated that positively charged molecules were released in a prolonged or sustained manner. Dong et al. used regenerated silk fibroins with different dissolving times to coat ibuprofen-loaded liposomes. The morphology, drug encapsulation efficiency, in vitro release, and in vitro corneal permeation of silk fibroin-coated liposomes were investigated in comparison with the conventional liposome. Coated liposomes showed sustained drug release and in vitro corneal permeation of ibuprofen as compared with drug solution and conventional liposome [40]. Regarding cell culture applications, numerous cell types have already been cultivated using silk-protein hydrogels, films, and 3D scaffolds, namely fibroblasts, vascular smooth muscle cells, keratinocytes, and mesenchymal stem cells (that were further differentiated to other cell lineages as cardiomyocytes, osteoblasts, and adipocytes) [34,41–44]. In bone tissue regeneration, particularly in the case of critical bone defects or severe bone injuries, external intervention is mandatory. Implants that provide mechanical strength, promote natural tissue growth, and degrade/resorb in time are ideal in such applications [45]. Silk-based materials have important advantages as they can have enough mechanical strength and flexibility, tunable biodegradability, good tissue integration, and low immune response activation [26]. For example, the repair of confined, critical-sized cancellous bone defects has been demonstrated in a rabbit model using fibroin hydrogels obtained by treating a fibroin water solution with glycerol (Glygel), or keeping the solution at 4°C (Thermgel). The data showed that the Glygel substrate promoted osteoblast proliferation, whereas the Thermgel substrate mainly favored osteoblast activity and differentiation [34].

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An osteoblast cell line (MG63) cultured in vitro in an injectable fibroin hydrogel matrix showed increased TGF β1 production, a cytokine that controls cell proliferation and differentiation [42]. The same injectable silk fibroin hydrogel matrix was also studied in vivo in a rabbit femur model and was shown to promote the healing of critical size defects of the trabecular bone through enhanced-bone remodeling and maturation [42]. To enhance further the osteogenic features of sonication-induced silk fibroin hydrogels, vascular endothelial growth factor (VEGF), and bone morphogenic ­protein-2 (BMP-2) have been added, which are key regulators of angiogenesis and osteogenesis during bone regeneration [43]. Such functionalized gels were shown to promote bone regeneration and height maintenance in a rabbit sinus floor elevation model [43]. Cartilage tissue regeneration is particularly difficult to achieve as the natural tissue has low regenerative capability as a result of the absence of blood vessels, nerve tissue, and lymphocytes, with low cell density, slow cell proliferation, and slow matrix turnover [46]. Once again, due to the silk hydrogels resemblance to cartilage matrix, they can be used to encapsulate cells and growth factors, contributing to stabilize the tissue during regeneration [41]. Aoki et al. prepared fibroin hydrogel and compared its performance with collagen gels. They inoculated chondrocytes isolated from 4-week-old Japanese white rabbits into fibroin hydrogel sponges, formed by phase separation of freezed fibroin solution and collagen gels [47]. The cells were cultured for 4 weeks. Although the cell density in the collagen gels was seen to be higher initially, it did not increase over time as in fibroin hydrogel. Similarly, the rate of increase of chondroitin sulfate in fibroin hydrogel was higher than that in collagen gels [47]. Positive histological staining for key cartilage ECM components (chondroitin sulfate and type II collagen) indicated formation of hyaline cartilage-like tissue in the pores of the fibroin hydrogel. Chondrocytes maintained round morphology and retained their differentiated phenotype within the fibroin hydrogel [47]. Cell microaggregates seeded in fibroin hydrogel closely mimic the initial stages of tissue formation and have been found to be very efficient in forming ECM for cartilage tissue regeneration. Reconstituted silk fibroin can also be blended with other biopolymers such as gelatin [48–51], chitosan [52,53], alginate [54,55], hyaluronic acid [56–58], and cellulose [59,60], to form hydrogels with a large range of material properties for improved tissue engineering applications. Similar approach has been performed with ceramics such as hydroxyapatite for improved mechanical properties [61,62].

7.4 Keratins Keratins that are the main constituents of skin, fur, hair, wool, claws, nails, hooves, horns, scales, beaks, and feathers [63]. In recent years, keratins from human hair have been recognized as potent naturally derived biomaterial owing to the fact that it is a human-derived source, has excellent biocompatibility (with immune system nonactivation), possesses cellular interaction sites, and shows good biodegradability [64].

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Intermediate filament

Medulla

Cortex

+

Micro-fibril

Cuticle

Coiled-coil of two α-helices

Micro-fibril

Cross-section of hair

α-Helix

α-Helix

Keratin molecule

Fig. 7.3  Structure of the α-keratin fiber. Reprinted with permission from Lee H, Noh K, Lee SC, Kwon IK, Han DW, Lee IS, et al. Human hair keratin and its-based biomaterials for biomedical applications. Tissue Eng Regen Med 2014;11:255–265.

Generally, keratin fibers consist of two major morphological parts: a cortex (the inner part of the fiber) and the cuticle outer layer. The cortex comprises spindle-shaped fibrils that are separated from each other by a membrane (Fig. 7.3), which consists of nonkeratinous proteins and lipids [64,65]. The cuticle layer comprises 10% of the total weight and is mainly composed of 5–12 layered β-keratins that function to protect the inner layers and hold moisture [65]. Keratin proteins self-assemble into fibers in the hair follicle to generate hair. Prior to extrusion through the skin, the keratin fiber is formed into a highly stable structure by covalent bonds, oxygen-catalyzed disulfide cross-links, and noncovalent interactions. Cross-linking can occur between separate polypeptide chains (intermolecular) but also between different points of the same polypeptide chain (intramolecular) [66].

7.4.1 Protein structure Keratins constitute two classes: designated as type I (acidic) keratin and type II (basic) keratin. Hard keratins (5% sulfur) found in hair, horns, feathers, nails, and tongue papillae [67,68] are classified as types Ia (acidic-hard) and IIa (basic-hard), whereas epidermal keratins such as the stratum corneum in skin, known as soft keratins (1% sulfur), are classified as Ib (acidic-soft) and IIb (basic-soft) [69]. The classification depends on amino acid composition, distribution, and functions. With regard to secondary structure, keratins can be subdivided into α, β, and γ-keratins [64]. In the fiber cortex is a bundle of protein strands mainly composed of α-keratins (50%–60%), γ-keratins (20%–30%), and melanin granules trapped inside the cortex strands, which accounts for 70%–95% of the total mass of the whole hair. α-Keratins have an average molecular mass in the range of 40–60 kDa, are low in sulfur, partly crystalline, and form an α-helical secondary structure [64]. α-Keratins

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self-assemble into long filamentous fibers that can be stretched considerably without rupturing [70]. γ-Keratins form the matrix in between the α-keratin filaments. They constitute approximately 25 wt% of the total protein of a fiber, have a low molecular mass of around 15 kDa, they are globular and are noted for their high content in cysteine, glycine, and tyrosine residues [64]. γ-Keratins function primarily as disulfide cross-linkers that hold the α-keratin fibers together and give rise to the high mechanical strength, inertness, and rigidity of the cortical superstructure of hair and wool [71]. Finally, as mentioned above, β-keratins are found in the cuticle. They protect the cortical filaments from physical and chemical damage and are difficult to extract [72].

7.4.2 Extraction and purification Over the last years, several methods to extract keratins have been reported [64]. The extraction process for hair keratin consists of four steps, including delipidization, solubilization, dialysis, and freeze-drying [64,73]. During the extraction process, α-keratin is extracted mainly from the cortex layers and the insoluble β-keratins embedded in the cuticle layer are removed by centrifugation. For efficient extraction of keratin from human hair, various techniques have been developed using combinations of various reagents and including the use of heat, mechanical force, and hydrolyzing buffer [64]. These processes are based on oxidative [74] or reductive [75] chemical reactions to break disulfide bonds between the cysteines, which converts the keratins into their non-cross-linked form [76]. The protein mixture obtained with oxidative solvents, which convert the cysteines to cysteic acid, are referred to as keratose, whereas the protein mixture obtained with reductive solvents, which leave the cysteines intact and thus ready to form new cross bridges, are referred to as kerateines [64]. The keratose obtained from oxidative extraction [74] with peracetic acid or hydrogen peroxide is hygroscopic, water-soluble, non-disulfide cross-linkable, and at extreme pH values is susceptible to hydrolytic degradation [77]. Biomaterials generated with keratose degrade relatively fast in vivo, for example, in days to weeks [77,78]. Conversely, biomaterials generated with kerateines from a reductive extraction procedure can persist in vivo for weeks to months. This is because the kerateines can be re-cross-linked through oxidative coupling of cysteine groups, and as a consequence are less soluble in water and more stable at extreme pH. The reductive extraction is usually performed with dithiothreitol (DTT), 2-mercaptoethanol [79], or with sodium disulfite [80,81].

7.4.3 Hydrogel formation Keratin-based biomaterials come in different morphologies, including films, sponges, and hydrogels [64,78]. The processing of keratin protein solutions into gels, films, and scaffolds has first been attempted in the early 1970s [82]. The use of human hairbased keratin hydrogels as wound healing promoters was patented by Blanchard and coworkers in 1999 [83]. One of the most extensively studied properties of keratin solutions, both at the microscale and the macroscale, is the ability to spontaneously self-assemble, resulting

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in a 3D fiber network with a reproducible fiber architecture and porosity [76,84,85]. Therefore, a hydrogel of pure keratin can be formed by disulfide and hydrogen bonds alone, with no need of additional cross-linker agents.

7.4.4 Applications in tissue repair and regeneration Keratin has received great attention in the field of biomedical engineering owing to its excellent biocompatibility, biodegradability, good cellular interactions between cellular integrins and specific amino acid sequences within the keratin molecule that are also found on other ECM proteins, and very low immune reactions upon implantation [64,78]. Keratin, in particular acidic α-keratin, contains cell-binding motifs, such as ­arginine–glycine–aspartic acid (RGD) and leucine-aspartic acid-valine (LDV). These motives are similar to those found in ECM proteins such as collagen or fibronectin, and also interact with integrins to support cellular attachment, proliferation, and migration (microvascular endothelial cells, keratinocytes, and fibroblasts) [86–88]. Keratin-based biomaterials are being used in a variety of biomedical applications, including nerve conduit filler for peripheral nerve regeneration, hydrogels, or films for wound healing, hemostatic agents, and more recently, as drug delivery vehicles and cell culture systems [64].

7.4.4.1 Nerve regeneration Sierpinski et al. [83] and Apel et al. [89] demonstrated that keratin-based hydrogels were neuroinductive and capable of facilitating regeneration in a peripheral nerve injury model in mice. They showed that a keratin gel derived from human hair enhanced the in vitro activity of Schwann cells by inducing cell proliferation and migration, and by upregulating the expression of specific genes required for important neuronal functions. When translated into a mouse tibial nerve injury model, keratin gel-filled conduits served as a neuroinductive provisional matrix that mediated axon regeneration and improved functional recovery compared with traditional nerve autografts [83]. In another study, the time course of peripheral nerve regeneration was evaluated with respect to neuromuscular recovery and nerve histomorphometry [89]. Again, ­keratin-based hydrogel scaffolds facilitated peripheral nerve regeneration and promoted neuromuscular recovery that was equivalent to the gold standard, sensory nerve autografts [89]. Later, to support the finding that keratin hydrogel fillers have the potential to be used clinically to improve nerve repair, the authors developed a rabbit peripheral nerve defect model and assessed the effectiveness of a keratin hydrogel conduit filler [90]. They found that the use of keratin resulted in a significant improvement of electrical conduction speed compared with both empty conduits and autografts, as well as a significant improvement in amplitude recovery compared with empty conduits. Furthermore, nerves in keratin-treated conduits had a significantly greater myelin thickness than nerves in empty conduits. More recently, Van Dyke and coworkers [91] showed that keratin hydrogels provide a permissive matrix that is well tolerated and rapidly infiltrated by cells of the peripheral nervous system. In a 1-cm sciatic nerve

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injury model in rats at an early stage of regeneration, they studied conduits filled with keratin hydrogels and compared the cell responses to those in conduits filled with Matrigel or saline solution. Keratin hydrogels facilitated earlier migration of dedifferentiated Schwann cells from the proximal nerve end, a faster Schwann cell differentiation, better myelin debris clearance, and decreased macrophage infiltration of the distal nerve tissue [91].

7.4.4.2 Wound dressing When keratin solution and keratin-based films were applied to the wound sites of an animal model as a wound dressing, accelerated onset of epithelialization and subsequently more rapid wound healing was reported in comparison to a polyurethane dressing [92,93]. Kirsner et al. also reported a highly improved wound healing effect in a patient with recessive dystrophic epidermolysis bullosa when their wounds were treated with a mixture of a keratin-based solution or hydrogel, and a gauze bandage wrap [94,95]. They also reported the cost-effectiveness, in addition to the improved wound healing, resulting from the use of the keratin-based wound dressing compared with the soft-silicone–based primary dressing and absorbent form of secondary dressing that are currently widely used in clinics. The wound healing capabilities of a keratin biomaterial hydrogel were studied in two pilot studies: one using a chemical burn model in mice and the other a thermal burn model in swine. In both studies, keratin was shown to prevent enlargement of the initial wound area and promote faster wound closure. Interestingly, treating thermally stressed dermal fibroblast in culture demonstrated that soluble keratin was able to maintain cell viability and promote proliferation. Separation of the α and γ fractions of the keratin biomaterial had differential effects, with the γ fraction producing more pronounced cell survival and recovery. These results suggest that the γ fraction, composed essentially of degraded α keratin proteins, may facilitate cell rescue after thermal injury [96]. Moreover, in an in vivo skin wound healing assay in rats, keratin treatment accelerated the epithelialization and maturation process [97].

7.4.4.3 Hemostatic agent Keratin hydrogels from human hair have been reported to act as a hemostatic agent in a rabbit model of lethal liver injury [98]. In comparison to other commonly used hemostats (QuickClot and HemCon bandages), the keratin hemostatic gel improved 24-h survival and performed as well, if not better, than conventional hemostats in terms of total blood loss and shock index. The keratin gel used in these experiments acted on the injury site by instigating thrombus formation and by forming a physical seal of the wound site that acted as a porous scaffold to allow cellular infiltration and granulose tissue formation [98].

7.4.4.4 Cartilage tissue engineering Xu et al. reported a 3D keratin scaffold fabricated from the highly cross-linked keratin extracted from chicken feathers, which was de-cross-linked and disentangled into

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linear and aligned molecules with preserved molecular weight [99]. The solution was readily electrospun into scaffolds with ultrafine keratin fibers oriented randomly in three dimensions. Due to their highly cross-linked molecular structures, keratin scaffolds showed intrinsic water stability. Adipose-derived mesenchymal stem cells were able to penetrate deeper, proliferate, and chondrogenically differentiate [99].

7.4.4.5 Controlled drug delivery system Cardamone et  al. performed a study involving keratin sponge/hydrogel formation from reduction versus oxidation hydrolysis of wool, and they found that the produced materials were distinctly different in appearance and behavior [100]. Those formed by reduction hydrolysis appeared smooth and homogeneous. Those formed by oxidation hydrolysis appeared rough and inhomogeneous. Regardless of oxidation or reduction hydrolysis, keratin sponge/hydrogels showed swelling in a simulated gastric fluid but maintained their structural integrity over time. This behavior can be exploited for drug release with a dual-control kinetics for delivering higher dosages immediately after immersion, and lower dosages over prolonged time periods [101]. The Van Dyke research group developed a drug-loaded keratin hydrogel by dissolving 200 mg/mL human hair keratin and 2 mg/mL ciprofloxacin–HCl in phosphate-buffered saline at 37°C overnight. Their results showed that the release of ciprofloxacin from keratin hydrogels could be sustained in vitro up to 3 weeks due to the electrostatic interactions between ciprofloxacin and keratin molecules, and that the released drug successfully inhibited growth of Staphylococcus aureus for 2 weeks when implanted subcutaneously, thus demonstrating the use of keratin hydrogels for antibiotic release to prevent acute infection in regenerative medicine [102]. Moreover, de Guzman et  al. cross-linked polyethylene glycol and keratin into a sponge-like scaffold to absorb test proteins with different isoelectric points (pI): albumin (~5), hemoglobin (~7), and lysozyme (~11). The protein release kinetics was influenced by charge at physiological pH 7.4 (albumin > hemoglobin > lysozyme). Under acidic conditions (pH 4), all proteins including keratins were positively charged and therefore release was determined by size (MW) diffusion: lysozyme (14 kDa) > hemoglobin (64 kDa) > albumin (66 kDa) [103]. More recently, Ham et al. described an approach to fabricate keratin hydrogels with tunable rates of erosion by mixing keratose and kerateine. The variation in the keratose–kerateine ratio led to tunable control over release rates of recombinant human insulin-like growth factor 1 [10]. Passipieri et al. evaluated the utility of keratin hydrogel formulations as a cell and/ or growth factor delivery vehicle for functional muscle regeneration in a surgically created volumetric muscle loss (VML) injury in the rat tibial anterior muscle. Keratin hydrogel implantation promoted statistically significant and physiologically relevant improvements in functional outcomes of post-VML injury to the rodent model [104].

7.4.4.6 Cell culture systems Wang et  al. described the development of a 3D hair keratin hydrogel, which allowed living cell encapsulation under near physiological conditions. This keratin hydrogel was comparable to collagen hydrogels in terms of supporting viability

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and proliferation of L929 murine fibroblasts. Notably, the keratin hydrogels contracted significantly less as compared with the collagen hydrogels, over a 16-day culture period. In addition, preliminary in vivo studies in immunocompetent animals showed a mild acute host tissue response to such materials [105]. Also, Rueda et al. prepared a 3D keratin scaffold, however using a novel keratin-based resin and printing methodology [106].

7.5 Elastin Elastin is a protein component of the ECM and is abundant in organs that need to stretch and recoil, such as blood vessels, elastic ligaments, lungs, and skin [107–109]. Its concentration varies between different tissues, depending on the tissue structure and the required elastic properties [109,110]. These elastic fibers display notable resilience and structural stability, with aortic isolates of elastin exhibiting a half-life of ~70 years [110]. For example, elastin in the elastic lamina of the arterial wall is mostly responsible for the elastic recoil after vessel expansion, and is therefore important for the regulation of blood flow and maintenance of blood pressure during the diastole [111]. In the lung, elastin is arranged as a lattice that supports the shape stability of the alveoli during tidal breathing [111]. In skin, elastin fibers are enriched in the dermis where they impart skin flexibility and extensibility [112].

7.5.1 Protein structure Elastin is a protein comprised of approximately 800 amino acid residues [113]. It is synthesized from a ≈ 72 kDa precursor, tropoelastin that is water soluble, non-­ glycosylated and highly hydrophobic [109,114]. Tropoelastin is a highly elastic protein; it is capable of extending to approximately eight times its resting length with no evident hysteresis [115]. The tropoelastin molecule consists of two types of domains encoded by separate exons: hydrophobic domains rich in nonpolar amino acids (glycine, valine, alanine, and proline residues, which often occur in repeats of tetra-, penta-, and hexapeptides such as GVGVP, GVPGV, and GVGVAP), and hydrophilic domains (mainly lysine and alanine residues, which are potentially involved in cross-linking domains of tropoelastin) [116]. Desmosine and isodesmosine are the two predominant cross-links of native elastin, each involving four lysine residues that are cross-linked by lysyl oxidase [116]. A spring-like coil adjacent to the N-terminus is primarily responsible for its elasticity. The molecule is entropically driven to recoil after stretching because the configuration of water changes when hydrophobic regions are exposed upon stretching [117], decreasing the number of possible structural conformations [118]. Further flexibility arises from a hinge region that contains key lysine residues that participate in cross-linking to form elastin [119,120], adding stiffness to elastin fibers as demonstrated by the difference in Young’s moduli between tropoelastin and natural mature elastin (~3 kPa and 300–600 kPa, respectively) [115] (Fig. 7.4).

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Peptides and Proteins as Biomaterials for Tissue Regeneration and Repair Desmosine

Isodesmosine

—OC — CH — NH— CO CH NH

(CH2)3 (CH2)2

(CH2)2

+ N

CO

CO

CH

CH

NH

NH

(CH2)4

—OC — CH — NH—

CO (CH2)2

+ N

(CH2)2

CH

NH

(CH2)3

CH

CO

(CH2)4

NH

—OC — CH — NH—

Fig. 7.4  Specific elastin intermolecular cross-links include the tetrafunctional desmosine and isodesmosine formed from four Lys residues from two different tropoelastin molecules. Reprinted with permission from Daamen WF, Veerkamp JH, van Hest JCM, van Kuppevelt TH. Elastin as a biomaterial for tissue engineering. Biomaterials 2007;28:4378–4398.

A number of biophysical properties are crucial for the biochemical/physiological role of elastin in the body, such as elasticity [121,122], glass transition temperature [123], and coacervation [124]. The elasticity of elastin is entropy-driven, thus stretching decreases the entropy of the system. Elastic recoil is induced by spontaneous return to the maximum level of entropy [125]. The glass transition temperature (Tg) of elastin is highly dependent on its water content [126]. When dehydrated, Tg is about 200°C, and at 30% of hydration, Tg is around 30°C [127]. Upon raising the temperature above Tg, elastin and elastin-based materials, unlike other proteins that become denatured, form ordered structures and undergo a second phase transition (coacervation and droplet formation). It has been suggested that the nonpolar domains of elastin are responsible for the coacervation process through an entropic mechanism of hydrophobic association due to the loss of entropy from the protein chains that is compensated by the release of water [128]. Thus, the coacervation temperature can be affected by protein concentration, hydrophobicity (amino acid composition and distribution), pH, and ionic strength of the solvent [128,129]. After overnight incubation above the coacervation temperature, the loose network becomes a compact aligned fibrillar structure [128].

7.5.2 Extraction and purification Elastin-based biomaterials can be fabricated from natural, recombinant, and synthetic sources. Natural elastin and tropoelastin are difficult to source as the expression of tropoelastin is largely repressed in adults [130], and is hard to isolate because it is insoluble in its native form [131]. More recently, Mithieux et al. were able to in vitro synthesize elastin fibers regardless of the age of the dermal fibroblast donor [132]. Nevertheless, the amino acid sequences of tropoelastins from various sources (human, chick, bovine, and rat) have been determined, and all were found to have a close homology at both DNA and amino acid levels [124]. Due to this interspecies conservation of the tropoelastin gene, animal-derived tropoelastin has become a widespread alternative to natural human tropoelastin [133]. Elastin isolation can be achieved by a

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number of methods, resulting in a range of end products. Solubilization with hot alkali, guanidine, or more recently, oxalic acid (produces α-elastin), and potassium hydroxide (produces κ-elastin) treatment can be used to hydrolyze elastic fiber components harvested from animal tissues [109,134]. To fabricate elastin-based biomaterials, the insoluble elastin from animal tissue must first be hydrolyzed to obtain a soluble elastin protein. Soluble elastin can be obtained by hydrolysis of insoluble elastin using oxalic acid or potassium hydroxide to break the peptide bonds [116]. Solubilized elastin retains many physicochemical properties of tropoelastin, including the ability to self-assemble. However, these methodologies do not give rise to intact tropoelastin monomers and often consist of heterogeneous products due to their harsh nature [135]. A second alternative to human elastin employs recombinant human tropoelastin DNA in E. coli systems [136]. Optimization of the genetic codons for protein synthesis in E. coli has allowed synthetic human tropoelastin to be more readily obtained through bacterial culture, yielding high purity recombinant human tropoelastin [109]. Alternatively, repeated elastin-like sequences can be produced by synthetic or recombinant methods to obtain elastin-like polypeptides (ELPs), which will be further addressed in Chapter 12.

7.5.3 Hydrogel formation Elastin hydrogels are typically cast by cross-linking tropoelastin solutions under physiological conditions. In the initial stages of this process tropoelastin monomers reversibly self-organize into spherical nanoparticles. By altering the concentration of tropoelastin, the size of these nanoparticles can be modulated up to a maximum of ~200 nm. Upon the addition of cross-linkers, these spheres coalesce to form interconnected beaded networks with distinct concentration-defined morphologies. Hydrogels formed at high tropoelastin concentrations (30–40 mg/mL) have a porous structure, and possess fiber diameters resembling those of in vivo elastic fibers [137]. Hydrogel cross-linking can be achieved through numerous enzymatic, chemical, and irradiation methods. The degree of cross-linking is influenced by the concentration of the cross-linking agent and the duration of incubation [138]. Several cross-linkers such as glutaraldehyde (GA) [139], disuccinimidyl glutarate (DSG) [139], bis(sulfosuccinimidyl) suberate (BS3) [140], copper sulfate and pyrroloquinoline quinone (PPQ) [141], ethylene glycol diglycidyl ether (EGDE) [142], hexamethylene diisocyanate (HMDI) [143], tris-succinimidyl aminotriacetate (TSAT) [144], disuccinimidyl suberate (DSS) [140], and β-[tris(hydroxymethyl) phosphino] propionic acid (THPP) [145] have been used to cross-link genetically engineered ELPs [139,140,143–145], tropoelastin [138], and α-elastin [142]. The assembly of tropoelastin into a polymeric matrix is facilitated by the ­elastin-binding protein (EBP) (67 kDa) that assembles tropoelastin into a microfibrillar network seed, which serves as a scaffold for additional tropoelastin deposition [109]. The lysine residues are then cross-linked by lysyl oxidase and stabilize the mature insoluble fiber. In the absence of lysyl oxidase, tropoelastin tends to associate with ­glycosaminoglycan (GAG, with negative charge) due to the presence of α-amino groups in the elastin lysine residues, which have a positive charge [109].

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7.5.4 Application in tissue repair and regeneration The structural stability, elastic resilience, and bioactivity of tropoelastin, combined with its capacity for self-assembly, make this protein a highly desirable candidate for the fabrication of biomaterials. The most well-characterized cell–tropoelastin interactions are mediated through cell receptors such as EBP [146], GAGs [147], and integrins [148]. EBP is a transmembrane protein [149] that is an inactive, alternatively spliced variant of β-­galactosidase capable of binding XGXXPG sequences in elastin such as VGVAPG [150]. Binding of EBP to extracellular elastin fragments, formed through injury, triggers cell activity such as myofibrillogenesis mediated by vascular smooth muscle cells [151] and chemotaxis of fibroblasts and monocytes [150]. The overarching cellular response to tropoelastin hydrogels is attractive. A wide range of cell types including fibroblasts, endothelial cells, epithelial cells, embryonic kidney, and fibrosarcoma cells of both human and animals origin remain adherent and adopt proliferative states following seeding on these hydrogels [152]. Differential cell migratory responses are seen on the hydrogel surfaces, characterized by pervasive cell infiltration on the more porous top surface, and cell monolayer formation on the casting surface. Implantation of the hydrogels in the dorsum of guinea pigs to assess immunogenic response has demonstrated that tropoelastin hydrogels are innocuous and invoke only a mild foreign-body response comparable to collagen [138]. The biocompatibility of tropoelastin scaffolds engineered with high porosity has been assessed in subcutaneous murine models [153]. Six-week-old implants demonstrate multilayer encapsulation by fibroblast cells and moderate scaffold remodeling and degradation. The main contributors to scaffold degradation are infiltrating fibroblast cells that reconstruct the local environment by depositing native ECM proteins such as collagen fibers. Immunogenic factors including neutrophils and monocytes are not observed, which indicates that the scaffolds are well tolerated by the host. Tropoelastin scaffolds have also found applications as stem cell delivery v­ ehicles, as they mechanically and biologically reflect a native ECM, which is important in regulating stem cell differentiation. Scaffolds laden with adipose-derived stem cells are non-immunogenic, increase the rate of wound closure, and enhance wound healing in vivo [154]. Electrospun scaffolds also serve as a viable treatment alternative to allogenic and xenogenic skin grafts for >6 million severe burn injuries occurring worldwide each year [155]. While acceptable for short-term use, foreign skin grafts have limited availability, are strongly rejected by the host immune system, and possess elevated infection risks [156]. Elastin-based scaffolds are therefore appealing as they mimic the dermal environment and avoid the cytotoxic leaching common in synthetic polymers such as poly(lactide-co-glycolide) and poly(ε-caprolactone). Additionally, they offer advantages over similar ECM-like scaffolds formed from collagen, which contract and lead to reduced patient mobility [157]. Weiss and coworkers described the production and properties of synthetic elastin formed by chemically cross-linking recombinant human tropoelastin with BS3, allowing for the construction of elastic sponges, sheets, and tubes [138]. In vitro, these

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materials support the growth and proliferation of different cell types (adherent epithelial cells including M-1 murine cells and human fibrosarcoma cells). In vivo, subcutaneously inserted implants in guinea pig were well tolerated [138]. Furthermore, EGDE was used to cross-link elastin- based materials from α-elastin [142]. These cross-linked materials supported vascular smooth muscle cell (porcine VSMC) adhesion but, compared with polystyrene controls, they exhibited a decreased proliferation rate [142]. In another study, lysine-containing elastin-like polypeptide hydrogels were formed in aqueous solution by cross-linking with THPP under physiological conditions in the presence of mouse NIH-3 T3 fibroblasts [145]. These cells survived the cross-linking process and were viable after in vitro culture for 3 days [145]. High-pressure CO2 has been used as a foaming agent to produce porous elastin hydrogels [158,159] or tropoelastin/elastin hydrogels followed by chemical cross-­ linking [152]. Higher pressures resulted in an increase in the porosity, improvement of mechanical properties, and swelling ratio. Additionally, due to the formation of large pores in the hydrogels, cell proliferation, and growth were substantially enhanced. Alternatively, it is also possible to prepare elastic biomaterials from tropoelastin without cross-linking [160]. Under alkaline conditions, tropoelastin proceeds through a sol–gel transition leading to the formation of a stable elastic hydrogel that has been shown to support the growth of human skin fibroblast. Further, in vivo studies with recombinant human elastin hydrogels implanted in female Sprague–Dawley rats revealed its potential as a cell support biomaterial [160].

7.6 Resilin Resilin is an elastomeric structural protein found in insect cuticles (locusts, dragonflies, cicadas, and cockroaches) [161]. More recently, resilin has been identified in the clamp sclerites of monogenean fish parasites [162] and the opal teeth of copepod crustaceans [163]. Resilin from dragonfly tendons has an elastic modulus of 600–700 kPa and can be stretched to three times its original length before breaking (resilience) [164]. Indeed, resilin highly resilience allows it to recover immediately its original shape even after being stretched for weeks. This protein is one of the most stretchable elastomeric proteins currently known [165].

7.6.1 Protein structure It has been difficult to identify the primary sequence and molecular structure of resilin due to the reduced stability during purification [166]. The predicted 620 amino acid sequence of the Drosophila gene product CG15920 (a precursor of resilin) has a tripartite structure: the first exon with 323 amino acids (exon I reported as “pro-­ resilin”) consists a so-called signal peptide sequence with 17 amino acids followed by 18 pentadecapeptide repeats (GGRPSDSYGAPGGGN) [167,168]; the second exon with 62 amino acids (exon II) contains a typical cuticular chitin-binding domain

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17

Exon 1 26

Exon 2 322 342

403 412

NH2

Exon 3 602 620

COOH GGRPSDSYGAPGGGN

GYSGGRPGGQDLG

Fig. 7.5  The putative resilin sequence from the Drosophila melanogaster CG15290 gene product. The sequence consists of a signal peptide and three different exons (exons 1–3). The signal peptide is removed before secretion into the extracellular space. Exons 1 and 3 include 18 repeats of GGRPSDSYGAPGGGN and 11 copies of GYSGGRPGGQDLG, respectively. The sequence in exon 2 is involved in binding of chitin. Reprinted with permission from Su RS, Kim Y, Liu JC. Resilin: protein-based elastomeric biomaterials. Acta Biomater 2014;10:1601–1611.

[168]; the third exon with 235 amino acids (exon III) contains 11 tridecapeptide repeats (GYSGGRPGGQDLG) [169] (Fig. 7.5). Both, exon I and exon III are rich in proline and glycine residues, which provide high flexibility to resilin [170]. They also have tyrosine residues, which form intermolecular cross-links through di- and tri-tyrosines connecting resilin polypeptides [170]. Resilin behaves as an entropic elastomer consisting of unordered chains linked through stable cross-links. Its restoring elastic force arises due to the loss in conformational entropy upon stretching [171]. Unlike elastin, the resilin sequence is dominated by hydrophilic residues, suggesting that hydrophobic interactions are minimal [172]. Exon I consists of more hydrophilic blocks and has a more flexible structure that promotes self-­aggregation to fibrillar structures in water, compared with exon III, which is composed of hydrophobic and hydrophilic regions that tend to form micelles in water [173]. Exon II, which is relatively hydrophobic, forms micelles of different sizes in water [173]. A different interesting property of natural resilin is that it undergoes fluorescence resulting from its dityrosine cross-links [174].

7.6.2 Protein extraction and purification As mentioned above it is very difficult to extract resilin from its natural sources, mostly because of its low stability during purification. Resilin-based polypeptides have been produced in E. coli, which is the most common host for expressing heterologous proteins due to its fast growth rate, cost-effectiveness, and low probability of post-translational modification [175]. Previous studies of resilin-based proteins have utilized the T5/T7 expression system using three different induction methods: isopropyl β-d-1-thiogalactopyranoside (IPTG) induction, autoinduction, and lactose induction [176,177]. The first genetically engineered resilin-based material that exhibited the high resilience of natural resilin was a cross-linked recombinant protein comprising exon I of the CG15920 gene (rec1-resilin) [167]. Rheological studies showed that cross-linked rec1-resilin hydrogels exhibit outstanding elasticity and a resilience (yield strain) of 92%, which exceeds that of most other polymer hydrogels [178].

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Recombinant resilin-like polypeptides (RLP) have been purified by three different methods: nickel–nitrilotriacetate (Ni–NTA) affinity chromatography, salting-out and heating, and temperature-induced coacervation. The Ni-NTA affinity chromatography method consists on the incorporation of affinity tags in recombinant proteins to facilitate purification procedures [179]. A polyhistidine tag (His-tag), which usually contains a minimum of six histidine residues, is widely used and has a number of advantages: it has small size, is less immunogenic than large tags and rarely affects protein characteristics such as solubility, structure, and biological activity. The Histag, which binds to Ni–NTA chromatography resins, is placed at the N-terminus of RLP, facilitating protein purification in native conditions without disrupting protein structure [179]. The salting-out and heating method to purify RLP is based on the heat stability of cross-linked resilin [174]. Purification is performed by removing undesired proteins at low salt concentrations, precipitating, and enriching the resilin proteins at higher salt concentrations, and precipitating undesired proteins at high temperatures [174,180]. The high purity of different resilin-based proteins is then confirmed by SDS–PAGE analysis. Due to its simplicity, the salting-out and heating method is faster and more cost-effective than affinity chromatography. The concentrations of ammonium sulfate used to precipitate resilin-based proteins range from 20% to 40% (w/v). The minimum salt concentration necessary to precipitate proteins as well as the temperature subsequently used, depends on the individual resilin-like proteins characteristics (e.g., hydrophobicity) [168,181]. Lastly, temperature-induced coacervation method starts with resilin-like protein enrichment by selective precipitation with ammonium sulfate and then purification is achieved by incubating the protein solutions at a cold temperature (i.e., 4°C) based on the upper critical solution temperature (UCST) behavior of resilin-based polypeptides [182]. The use of temperature-induced coacervation minimizes possible protein degradation and denaturation, which may be seen during heat treatment (e.g., 4 h at 95°C) [182].

7.6.3 Hydrogel formation In order to produce hydrogels, Qin et al. synthesized full-length resilin from D. melanogaster resilin genes that was cross-linked by forming dityrosine bridges in two ways: enzymatically, using horseradish peroxidase, and photo-Fenton reaction, using FeSO4 and H2O2 followed by ultraviolet (UV) exposure [181]. Since then other researchers have also produced resilin-based hydrogels using a variety of other cross-linkers, or through the cross-linking of specific functionalized resilin sequences. Charati et al. [172] modified the resilin repeat by replacing the tyrosine with phenylalanine, in order to facilitate a photochemical cross-linking of the polypeptide [183]. Lysine residues have also been included outside the putative resilin repeat as additional cross-linking sites, as well as the cell-adhesion ligand RGDSP, derived from the fibronectin subunit module FN-III10 [184].

7.6.4 Application on tissue repair and regeneration Resilin-based materials as other protein-based polypeptides have been engineered to promote cell adhesion [183,185], material degradation [186], growth factor delivery

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[187], and cell differentiation [188]. The potential applications of resilin-based biomaterials have been broadened by incorporating multiple functionalities [189] such as the cell-binding RGD sequence [190,191], a matrix metalloproteinase (MMP)-sensitive degradation sequence [190,191], a heparin-binding domain [190,191], a VEGFmimicking peptide [168,192], and the chitin-binding domain [168]. Furthermore, a MMP-sensitive sequence (GPQG↓IWGQ, derived from the human α1(I) collagen chain and readily cleaved by MMPs) was included to promote proteolytic degradation [193]. In addition, the heparin-binding domain (CKAAKRPKAAKDKQTK) was included in the sequence for the noncovalent immobilization of heparin to allow for the sequestration and controlled release of growth factors [194]. The combination of these approaches for the generation of a resilin-like hydrogel-induced cell adhesion and proliferation of mouse NIH 3 T3 fibroblasts. The stiffness of this hydrogel was tuned within the range of 500 Pa to 10 kPa by changing the polypeptide concentration and cross-link ratio [191]. Consensus sequences derived from Anopheles gambiae (mosquito) genes have also been identified, and it was found that proteins obtained from the mosquito sequences display similar properties to rec1-resilin [171,174]. Based on these studies, a new modular protein containing repeating motifs derived from A. gambiae and a cell-binding domain derived from fibronectin was designed [180]. When cross-linked with tris(hydroxymethyl)phosphine, the hydrogels had a complex modulus of 22 kPa, a yield strain of 63%, and a compression modulus of 2.4 MPa, which is on the same order of magnitude as human cartilage. Human mesenchymal stem cells (hMSCs) cultured on such resilin-based hydrogels showed good spreading behavior and had a viability of 95% after 3 days of culture [180]. Recently, new RLPs, containing 12 repeats of the putative resilin consensus sequence and an MMP-1-sensitive domain were produced [195]. These RLP-based polypeptides exhibit largely random-coil conformation, both in solution and in hydrogels cross-linked with tris(hydroxymethyl) phosphine. Primary hMSCs encapsulated in RLP hydrogels were viable over extended time periods [195]. Additionally, RLP– PEG hybrid hydrogels were investigated. These hydrogels are cross-linked through a Michael-type addition reaction between cysteine residues on the polypeptide and a vinyl sulfone-terminated PEG [190]. These RLP–PEG hydrogels form stable networks upon mixing of the two components. Human aortic adventitial fibroblasts were successfully encapsulated in such hydrogels [190].

7.7 Final remarks and future perspectives In this chapter, we reviewed the properties of biomaterials based on naturally occurring fibrous proteins. Their degradability, biocompatibility, availability, and similarity with ECM proteins make them attractive for numerous biomedical applications, in particular in tissue engineering and regenerative medicine. In addition, one of the major advantages of these proteins is that they can be specifically modified and enhanced by genetic engineering strategies to add functionality, for instance to facilitate cell

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adhesiveness and controlled degradability by proteases. These proteins self-assemble under specific conditions, which render them compatible with emerging technologies such as rapid prototyping and biofabrication approaches. The importance of biomaterials based on naturally occurring fibrous proteins is bound to grow substantially in the future.

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