Drug Deliv. and Transl. Res. (2012) 2:313–322 DOI 10.1007/s13346-012-0087-x

REVIEW ARTICLE

Electrospun collagen and its applications in regenerative medicine Matthew J. Fullana & Gary E. Wnek

# Controlled Release Society 2012

Abstract In recent years, electrospinning has increased in popularity as a processing technique for obtaining nanometerto-micron diameter polymer fibers collected to form a nonwoven scaffold. It possesses the ability to process collagen into nanofibrous scaffolds which have been used for a number of applications, such as artificial vascular grafts and for wound repair. This paper offers a review of some of the basic yet essential aspects of producing nanofibrous scaffolds of collagen by electrospinning. A primer to collagen structure, crosslinking techniques, and electrospinning principles is provided, along with some of the many applications of these unique materials.

antigenicity, promotion of cell attachment and growth, and is relatively non-immunogenic [3–5]. There are multiple varieties or “types” of collagen, all characterized by minor genetic variations. Over 20 genetically unique types exist, but type I remains the most prevalent in the body. Types I, II, III, V, and XI are within a special subclass of collagens known as fibrillar collagens, owing to their ability to form higher ordered collagen fibrils and represent over 90 % of the total collagen in humans. For the purposes of this review, we will be focused on type I collagen as it is most commonly used for the electrospinning process among other applications and accounts for up to 70– 90 % of the collagen found in the body [6].

Keywords Electrospinning . Collagen . Regenerative medicine . Tissue engineering . Nanofiber . Scaffolds . Composite . Cross-linking

Primary structure

Introduction to collagen structure As a tissue scaffold and drug release material, researchers continue to focus on collagen for a variety of reasons. The majority of human tissues contain collagen, making it the most abundant protein in mammalian extracellular matrix (ECM) [1] and constituting 30 % of all vertebrate body protein [2]. It serves as the major structural protein conferring strength to tissues such as dermis, bone, cartilage, tendon, ligament, and internal organs. In addition, it continues to be the preferred material for biomedical devices given its biocompatibility, low

All collagens consist of three protein chains. In type I, each chain is designated as an alpha chain (α-chain). Two of these chains are identical and are referred to as α1-chains, while the remaining third chain is referred to as the α2-chain [4]. Together, these three chains form a triple helical structure referred to as the collagen monomer or gamma structure [7]. The molecular conformation of the collagen triple helix possesses a peptide containing strings of five or six amino acid triplets of [Gly-X-Y]n, where X and Y are typically the amino acids proline and hydroxyproline, respectively, and n is 337–343 depending on the collagen type. In all fibrillar collagens, the α-chains are flanked by much shorter, nonhelical, terminal domains, about 20 residues in length [8]. Secondary structure

M. J. Fullana : G. E. Wnek (*) Department of Macromolecular Science and Engineering, Case Western Reserve University, 2100 Adelbert Road, Cleveland, OH 44106, USA e-mail: [email protected]

The central domain of the three α-chains forms a tight, right-handed α-helix with an axial residue-to-residue spacing of about 0.286 nm and an angular separation of 108°. The coiling of the helix is largely due to steric repulsion between the proline and hydroxyproline residues. The peptide

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bonds form the backbone of the α-helix, pushing the side chains of the α-helix to the outside of the chain. Given that the geometry of the α-helix results in roughly three residues per helical turn and since every third residue is a glycine, this generates a row of nearly superimposed glycine residues. This glycine row follows a slightly left-handed helix, possessing a pitch of approximately 8.6 nm which winds on the surface of the coiled molecule [9, 10]. Tertiary structure The glycine row creates a left-handed triple helix with a pitch of about 10.4 nm with all the glycine residues arranged inside the triple helix. This produces right-handed threads that form into a left-handed rope and confers stabilization to the collagen molecule [10]. Triple helix formation follows the C→N direction and is dependent upon the lack of steric hindrances provided by the glycine rows which constitutes the central axis of the molecule. As an aside, a point mutation resulting in a different residue appearing in place of glycine results in faulty molecular packing and is the cause of a variety of connective tissue disorders [9]. The formation of the triple helix gives rise to a long, rodlike structure which is ~1.5 nm wide and over 300 nm long and capped at both ends by globular domains. This stiff yet flexible form called procollagen is excreted into the extracellular space [8, 9]. Fibril formation During fibril formation, the terminal globular domains are cleaved by specific proteases. The remaining telopeptides contain some terminal non-helical fragments, but this entire reactive molecule is termed tropocollagen. By the process of spontaneous fibrillogenesis, it is able to self-assemble into large supramolecular structures. At this point, the organization of residues along the longitudinal axis forms a lateral intermolecular interaction with an axial stagger of roughly 234 amino acid residues or ~67 nm (or some integer multiple). Figure 1 illustrates the chemical structure of collagen with the stagger pattern depicted in I-C. Since the division of the total length of the molecule by the 234 amino acid residues does not result in an integer, a gap exists between the end of one molecule and the beginning of the next. This gap is responsible for the characteristic D-period of collagen fibrils, which occurs at ~67-nm intervals, and banding patterns can be seen using electron microscopy methods (see Fig. 2). Beyond fibrils Once fibril formation has completed, the ordered fibrils can combine to form bundles, which compose the collagen-rich tissues such as tendons and ligaments. At this stage,

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collagen becomes more specific for a given tissue type. For example, within the gaps which produce the D-period, biomineralization occurs in collagen produced for use in the bone, teeth, and cartilage. Tiny mineral particles first appear within the gap, at the fibril surface, and within the fibril itself. These needle-like particles develop into thin plates ~4 nm thick and extend into the overlap zones. Similarly, proteoglycans can also occupy the interfibrillar space, interacting with the collagen via side chains. These collagen– proteoglycan interactions are especially important for joining collagen fibrils together which confers mechanical strength to the fibrils by distributing the mechanical stresses throughout the whole tissue in the case of tendon or ligament. In cornea, a higher concentration of proteoglycans exists and is seen as being relevant for its transparency [10].

Collagen processing The earliest instance of preparing fibers of collagen occurred with Furukawa et al. by producing collagen fibers from solubilized collagen by spinning a collagen solution through a spinneret into a coagulating bath comprised of an aqueous solution of an inorganic salt, such as sodium sulfate, sodium chloride, ammonium sulfate, magnesium chloride, or aluminate sulfate [11]. In 1999, Fofonoff and Bell patented a method for producing fibers from wet spinning using a 0.5 M acetic acid solution [12]. The extruded collagen gel fiber is directed into a coagulation solution of alkaline alginic acid/boric acid. A year later, Hirano et al. published work on the wet spinning of chitosan–collagen fibers using an aqueous acetic acid–methanol solution spun into an aqueous ammonia solution containing ammonium sulfate [13]. While all of these methods successfully produced fibers of collagen, they all require lengthy preparations involving numerous laborious tasks and in some instances, utilize harsh solvents for preparation. Additionally, the use of spinnerets and wet spinning generates fibers possessing diameters in the tens and hundreds of microns, whereas native fibrillar collagen fibers in the human body range in size from 50 to 80 nm, depending on several factors, such as tissue type, age, etc. [14]. In order to improve biomimicry of collagen fibers, a technique capable of generating fibers on the order of nanometers would need to be adapted. Introduction to electrospinning The concept of electrospinning, also known as electrostatic spinning, has been around for centuries. The earliest mention of using electrical charge to prepare a fiber occurred in 1929 by Hagiwaba et al.; however, even earlier instances of applying an electrical charge to fluids date back as far as 1745 [15].

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Fig. 1 Chemical structure of collagen type I [2] (Reprinted with permission from Elsevier)

While there are two major divisions of electrospinning, solution and melt, we will focus on solution electrospinning since melt electrospinning is not possible with collagen due to denaturation of the collagen molecule. Typical solution electrospinning usually begins with solution preparation. A proper solution of the desired polymer(s) (in this case the polymer is primarily collagen) is prepared

using a suitable solvent for electrospinning. Choosing the proper solvent for electrospinning is regarded as one of the most crucial steps in electrospinning and depends on four critical factors: the dielectric constant, volatility, surface tension, and solvent conductivity [16]. The solution is loaded into a syringe equipped with a metal needle or nozzle. It is essential for the needle to be metal or at least conductive as it serves as

Fig. 2 TEM identification of the type I collagen N,N-bipolar fibril. a A TEM image of a N,N-bipolar fibril dispersed from 12-day embryonic chick metatarsal tendon and negatively stained with 2 % uranyl acetate (pH 4. 2). Analysis of the stain pattern along the fibril allows the molecular polarity to be established. The molecules outside the boxed region are packed in a polarized manner with their N-termini pointing towards the fibril tips, as indicated. The boxed region, shown at higher

magnification in (b), is the polarity transition region where molecules occur in antiparallel arrangement. Analysis of the stain patterns in the transition regions shows the axial arrangement of antiparallel molecules corresponds to the C-termini of antiparallel molecules being in axial register, as shown schematically in (c). The axial extent of the transition region appears invariant at close to 8 D-periods [58] (Reprinted with permission from Elsevier)

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an electrode, where a high electric field ranging from 100 to 500 kV/m is applied with currents typically measured in microamperes. The counter electrode or ground is placed within 10– 25 cm of the electrified needle and serves as the collection point for the fibers. This counter electrode can take several forms, such as a static metal plate or a rotating metal drum. The collection point, or target, determines the final form of the electrospun scaffold (i.e., flat scaffold, tubular shape for vascular graft, etc.) [17]. As an electrical potential is applied to the solution, a charge imbalance occurs at a critical voltage which overcomes the surface tension of the solution, from which a jet is produced. The jet is directed at the grounded target, allowing the solvent to evaporate and depositing dry fibers on the collection point [4]. Figure 3 shows a typical electrospinning setup. While simplistic in design, there are several variables which can impact the characteristics of the collected fibers: applied voltage, air gap distance (refers to the distance between the two electrodes), solution flow rate through the needle, polymer properties such as molecular weight and polydispersity; solution properties such as solvent, concentration, viscosity, and surface tension; and environmental factors such as ambient temperature and humidity [1, 4, 15]. Electrospinning collagen The earliest instance of electrospinning collagen-based scaffolds was performed by Huang et al. in 2001 with the electrospinning of collagen–polyethylene oxide (PEO) nanofibers [5]. Initially, fibers could not be formed from a 1–2 wt% of pure collagen prepared in 10 mM HCl at pH 2.0; however, upon addition of PEO, fibers were visible. The solution was electrospun to generate highly uniform fibers with a diameter range of 100–150 nm. While this was the first preparation of collagen fibers in the nanometer range, the resultant scaffold was not composed of pure collagen. In 2002, Matthews et al. published work on the successful electrospinning of type I and type III collagen from 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP) [4]. The electrospun fibers possessed average diameters ranging from 100 to 730 nm. It was discovered that a linear relationship existed between polymer concentration and fiber diameter Fig. 3 Schematic representation of typical electrospinning setup utilizing a rotating mandrel

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where 0.03–0.10 g/mL resulted in diameters ranging from 100 nm to 5 μm for type I collagen. The same relationship exists for type III collagen where a concentration range of 20– 80 mg/mL corresponds to a diameter range of 115–612 nm. It was discovered that HFIP was a very suitable solvent for electrospinning, being used to produce nanofibrous scaffolds from a variety of biopolymers such as artificial spider silk [18], chitin [19], type I and type III collagen [4], collagen/poly(Llactic acid)-co-poly(ε-caprolactone) [P(LLA-CL)] [20], fibrinogen [21], gelatin (otherwise known as denatured collagen) [22], and silk fibroin/chitin [23]. The effect of polymer concentration on collagen fiber formation was more thoroughly detailed by Li et al. [22]. They found that a decrease in collagen concentration would decrease fiber diameter to as low as ~100 nm; however, as the concentration decreased below 5 %, an increase in the presence of beaded fibers occurred. They concluded that a collagen concentration above 5 % would yield smooth and uniform fibers. In addition, they also found the tensile modulus of collagen fibers to be 262±18 MPa. While a seemingly perfect solvent for electrospinning, possessing ideal properties to produce nanofibers of nearly any polymer, both synthetic and natural, HFIP remains a highly volatile and corrosive solvent that poses health risks to humans. It has also been argued that the use of HFIP for electrospinning may denature collagen to gelatin [6]. Examination of work by Telemeco et al. provides evidence that the HFIP-produced scaffolds are structurally distinct from that of gelatin scaffolds [24]. The distribution of fiber diameters and pore dimensions observed between collagen and gelatin scaffolds differs to a subtle degree, with collagen scaffolds possessing a pore size range of 2,000–6,000 nm2 and gelatin scaffolds exhibiting a range of 1,500–4,000 nm2. Additionally, electrophoretic analysis revealed that fibrils of electrospun gelatin are composed of a complex mixture of protein fragments, shown by a lack of clear banding in SDS gel lanes. Lastly, electrospun type I collagen fibrils exhibit a 67 nm repeat structure not found in gelatin. While the high volatility of HFIP and the nature of electrospinning (evaporation of solvent from the fibers) would reduce the consequences of HFIP being present in the collagen

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scaffolds which could then be used for any range of biomedical applications, there is still an inherent risk to laboratory workers and concern over environmental hazards stemming from its use. To overcome this and to address any lingering concerns regarding HFIP degrading collagen, Dong et al. published work on the electrospinning of type I collagen scaffolds from a benign solvent system comprised of ethanol and phosphatebuffered saline 20× (PBS 20×; Fig. 4) [25]. It was found that a 1:1 ratio of PBS 20× and ethanol is a suitable solvent for dissolving water-insoluble lyophilized foam powder consisting of bovine-derived tropocollagen. A 16 wt% concentration of collagen was adequate for electrospinning using 20 kV, a pump rate of 1 mL/h, and an air gap distance of 10 cm. A range of PBS salt concentrations was examined, and at lower concentrations (PBS 5×), the collagen would continue to be dissolved, however fail to form fibers. PBS 10× was adequate for producing fibers with a diameter of 540±210 nm; however, increasing the salt concentration to 20× yielded fibers 210± 60 nm in diameter, which is on the same order of magnitude as the collagen fibers spun from HFIP. It also showed that the collagen scaffolds produced from HFIP and the benign solvent system are nearly identical. While HFIP continues to be a solvent of choice for electrospinning collagen for many, Liu et al. described electrospinning type I collagen from 40 % acetic acid [26]. Circular dichroism experiments were performed on scaffolds prepared from both HFIP and 40 % acetic acid to determine the extent of degradation caused by both solvents. While the results showed more denaturation in the HFIP-derived scaffolds than the ones produced using acetic acid, the experiments fail to include fibers spun from the benign solvent mixture described by Dong et al. [25].

Collagen cross-linking Cross-linking of collagen scaffolds has become an important step for producing scaffolds with enhanced mechanical properties as well as a means of producing water-insoluble scaffolds

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[25]. There are four major approaches for cross-linking collagen which all possess distinct advantages and disadvantages. Please note that only a brief overview of some of the most commonly encountered cross-linking methods will be covered here. A depiction of the different collagen cross-links can be seen in Fig. 5. Covalent chemical cross-linking of neighboring fibrils This method utilizes chemical cross-linking agents which covalently couple neighboring collagen fibrils using reactive groups in the collagen fibrils and the cross-linking molecules. Glutaraldehyde, epoxy compounds, and isocyanates are some examples. A disadvantage to these compounds is they are included within the cross-link which can produce chemical species not native to biological tissues and may cause any number of unfavorable conditions, such as inflammation, foreign body response, immunogenic reactions, etc. Glutaraldehyde Glutaraldehyde is one of the most frequently encountered fixatives in the biomedical field, due to its low cost, availability, rapid reactivity, and solubility in aqueous solution which makes it ideal for biological applications. The cross-linking of collagen-based tissues using glutaraldehyde most notably reduces biodegradation, making them biocompatible and non-thrombogenic while preserving anatomic integrity, leaflet strength, and flexibility [27]. The interaction of glutaraldehyde with collagen and the formation of cross-links were first described in 1968 [28]. Today, it is still the predominant crosslinking agent; however, it has received criticism as a crosslinking agent for implants as it was found to promote calcification [29] and for releasing toxic molecules from the implant [30]. Glutaraldehyde contains two reactive aldehyde groups flanking a short three-carbon aliphatic chain. An aqueous solution of glutaraldehyde contains a mixture of free aldehydes, monomeric and polymeric cyclic derivatives, and unsaturated polymers (see Fig. 6). At neutral pH, glutaraldehyde and its derivatives react with primary amines. If these amines are the side groups on proteins, a cross-link will form. Additionally, glutaraldehyde will also react with carboxy and amido groups present in proteins; however, these reactions do not constitute the main reactive sites for primary cross-link formation. Specific to collagen fibrils, glutaraldehyde reacts with ε-amino groups of the lysine residues in the collagen molecule [31, 32]. Physiochemical cross-linking methods

Fig. 4 SEM image of the electrospun collagen fiber from PBS (20×)/ ethanol (1:1) [25] (Reprinted with permission from John Wiley and Sons)

Physiochemical methods consist of catalyzing the formation of covalent cross-links to join collagen fibrils using the existing

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Fig. 5 A schematic representation of the basic structural design of a collagen fiber. The fundamental structural design of a collagen fiber is a staggered array of tropocollagen molecules. A five-stranded microfibril was suggested to account for such a substructure, which can then continue to grow by lateral and end-toend aggregation. Intrahelical and interhelical cross-links within or between tropocollagen molecules and intermicrofibrillar cross-links between adjacent collagen microfibrils are also illustrated [59] (Reprinted with permission from John Wiley and Sons)

reactive amino acid side chains without the addition of exogenous agents into the scaffold. This includes photooxidation, microwave irradiation, dehydration, and dehydrothermal treatment. This method is favorable over covalent chemical crosslinking agents such as glutaraldehyde because no chemicals are required and any risk of producing a toxic species is decreased. One method utilizing UV irradiation exposes the collagen scaffold to 254 or 514 nm wavelength light. However, in the case of UV irradiation as with others, overexposure of the collagen may irreversibly damage the fibril structure, denaturing it to gelatin. Although UV treatment is capable of improving the mechanical integrity of the scaffolds, it has little effect on reducing the degradation times in vivo [26, 33]. Intermolecular cross-linking of amino acid side chains Initiating intermolecular cross-linking of the reactive amino acid side chains requires the addition of a catalyst, such as carbodiimide or azyl azide, but offers the advantage of not becoming part of the cross-link like glutaraldehyde, epoxies, Fig. 6 Glutaraldehyde and the forms it takes in aqueous media [32] (Reprinted with permission from Elsevier)

and isocyanates, which is referred to as a zero-length crosslinking procedure. The catalyst and/or its byproducts can then be easily washed out of the scaffold following crosslinking. Carbodiimide cross-linking While glutaraldehyde still remains a popular choice for cross-linking collagen scaffolds, concern over complications arising from its use in vivo remains and has resulted in exploration of compounds which only catalyze the natural cross-linking between collagen fibrils and do not form the actual cross-link itself. Carbodiimide cross-linking has risen in popularity for its increased biocompatibility over glutaraldehyde, with numerous studies examining and confirming it as a noncytotoxic method [34–36]. The most common carbodiimide compound, 1-ethyl-3(3-dimethylaminopropyl) carbodiimide, reacts with the carboxyl groups of aspartic and glutamic acid side chains in collagen to form the o-isoacylurea derivative. The derivative

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then reacts by nucleophilic attack with primary amines of lysine side chains on collagen, which generates the amide cross-link and liberates the urea derivative of the carbodiimide. The urea derivative can easily be rinsed from the scaffold. Supplementing the reaction with N-hydroxysuccinimide will minimize internal rearrangement of the activated isoacylurea derivative, thereby reducing the occurrence of reaction intermediates and forming a two-step zero-length cross-linking method. The resultant isopeptide bond mimics the native peptide bond present in proteins, but bonds neighboring peptides. The carbodiimide reaction was found to be optimized at pH 5 [30, 31, 37]. Polymerizing compounds Polymerizing compounds do not chemically react with the collagen fibrils but instead interact with the fibrils to reinforce it. While the collagen and polymer matrix remain separate, considerations must be made to determine any ill effect produced by the polymer matrix in vivo.

Applications of collagen scaffolds There are hundreds of reports on the use of electrospun collagen scaffolds as biomedical devices with the most common applications being in use for vascular grafts and wound dressings. A major problem with processed collagen, including electrospun collagen scaffolds, is the lack of mechanical strength upon hydration [38]. The mechanical properties of processed collagen are not robust enough to allow its use for any application where it will be exposed to extreme pressures (vascular grafts) [39] or require some degree of elasticity (wound dressing or skin replacement) [40], even with cross-linking in some cases. This has resulted in a number of different approaches in developing collagen composite materials, using both synthetic and natural materials to reinforce and improve the properties of collagen-based materials. A variety of approaches have been taken to address the mechanical instability of collagen scaffolds by creating composite materials containing synthetic of naturally derived materials to reinforce the scaffold. Vascular grafts Venugopal et al. conducted a comparative study in 2005 between collagen and polycaprolactone (PCL) nanofibers for the proliferation of human coronary artery smooth muscle cells (SMCs) [41]. They found that for vascular graft applications, SMCs preferred collagen-coated PCL and collagen scaffolds over the PCL scaffolds. However, given the high intraluminal physiological pressure, it is likely that the pure collagen scaffold would succumb to mechanical failure

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[42]. This is supported by Girton et al. who found the maximum pressure of a collagen graft to be approximately 225 mmHg, supporting the use of collagen-coated PCL scaffolds or alternative combination scaffolds for vascular graft usage [43]. To further overcome the poor mechanical properties of a collagen scaffold and the lack of cell recognition signals [44], Stitzel et al. [45] and He et al. [20] described the preparation of collagen–poly(lactic acid) (PLA)-layered scaffolds and collagen-blended poly(L-lactic acid)-co-poly(ε-caprolactone) scaffolds, respectively. For the collagen–PLA-layered scaffolds, the PLA fibers were collected on a novel winding apparatus with variable pitch to simulate the smooth muscle cell arrangement within an artery. Collagen fibers were then deposited on top of the PLA fibers, followed by a final layer of PLA fibers. The biodegradable PLA fibers would confer improved mechanical strength, and the proper alignment of the fibers, produced by adjusting the collecting mandrel pitch, would encourage SMC orientation in the direction of the principal stress directions. For the collagen-blended P(LLA-CL) scaffolds, P(LLACL) copolymer (70:30) was dissolved with type I collagen in HFIP at 5 % (w/w). The scaffolds were electrospun at a rate of 1.2 mL/h, a voltage of 12 kV, and an air gap distance of 12 cm, which produced a range of fiber diameters between 100 and 200 nm. The blended collagen–P(LLA-CL) scaffold was found to have a desirable ultimate strain value (66±22 %) which is much closer to that of a human coronary artery (45– 99 %). It should be noted that the tensile strength and the ultimate strain of the P(LLA-CL) scaffold decreased significantly upon addition of collagen. While the collagen-blended P(LLA-CL) scaffold offered similar mechanical properties to native blood vessels, Buttafoco et al. took a different approach and looked to create a more natural blood vessel by electrospinning two native proteins together: collagen and elastin [46]. The two proteins were spun out of aqueous 10 mM HCl instead of HFIP at varying concentrations and ratios. The scaffolds required the addition of 1 % w/v PEO (Mw 08×106) as a carrier polymer and 42.5 mM NaCl to produce homogeneous and continuous fibers with fiber diameters ranging from 220 to 600 nm; however, no mechanical testing results were provided. Prior to this, Boland et al. constructed a blend of 40 % type I collagen, 40 % type III collagen, and 20 % elastin into a scaffold by electrospinning from HFIP at a concentration of 0.083 g/mL [39]. A popular synthetic polymer, polycaprolactone (PCL), is frequently incorporated into collagen scaffolds. Lee et al. dissolved type I collagen and PCL in HFIP and electrospun the blend to yield a vascular graft [47]. The fiber diameters were approximately 520 nm, and the burst pressure of the composite grafts (4,912±155 mmHg) exceeded the burst pressure of native blood vessels as well as the pure PCL grafts (914±130 mmHg). Later work explored the usage of the

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composite grafts in a rabbit aortoiliac bypass model. The study showed that platelets do not adhere to the endothelialized grafts [48]. A final collagen composite scaffold was created using polydioxanone (PDO). PDO is currently used in wound closure sutures and is marketed in the USA by Ethicon Inc. It is unique from other synthetic materials examined here because it displays shape memory, low inflammatory response, slower absorption rate when compared to poly(lactic-co-glycolic acid) (PLGA) and PGA, is colorless, and highly crystalline. A blend of polydioxanone (PDO) and type I and type III collagen, both dissolved in HFIP, was electrospun for blends of 100:0, 90:10, 80:20, and 70:30 (PDO/collagen by weight percent). Fiber diameters for the blends ranged from 210 to 340 nm. Interestingly, as the amount of collagen increased, the mean fiber diameter remained fairly constant; however, the addition of collagen to PDO did decrease the fiber diameter compared to PDO alone [49]. Wound repair As the major protein component of the ECM as well as the main structural protein in humans, collagen has been highly regarded as the preferred matrix material for new-age bandages and dressing for wound repair. Ideally, a tissue substitute should replicate both form and function of the native extracellular matrix; both mechanical strength and cellular interactions and signaling should not be compromised. In addition, any implanted biomaterial carries significant risk. The device must be biocompatible and not induce adverse reactions in the body, such as a foreign body reaction or thrombosis. It must be nonimmunogenic as well as noninflammatory as these types of reactions are likely to exacerbate preexisting or underlying conditions in the recipient [50]. Given the prevalence of collagen in the body, it is understandable why considerable effort has been made to exploit it as an engineered tissue. A recent effort exploring the effect of electrospun collagen scaffolds on early-stage wound healing was conducted by Rho et al. [51]. Type I collagen from calfskin was electrospun from HFIP and cross-linked using glutaraldehyde vapor. The resultant scaffolds were then used in an in vitro study examining cell adhesion and spreading of normal human oral keratinocytes (NHOK) and normal human epidermal keratinocytes (NHEK) on the collagen scaffolds. The same study also used the scaffolds in an in vivo open wound healing test on Sprague–Dawley rats. The cell seeding results showed that a relatively low level of normal human keratinocytes was observed on collagen nanofibers without an ECM coating compared to a polystyrene surface, while a collagen scaffold coated with laminin promoted adhesion of NHOK and NHEK, and that both materials support cell spreading. The open wound healing tests showed that collagen nanofibercovered wounds yielded similar wound closure results as

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gauze-covered control wounds, yet showed early-stage healing in the collagen group was accelerated, while the late-stage healing was similar for both groups. The use of collagen scaffolds can be applied to any number of topical injuries. In severely burned patients, rapid closure of full thickness wounds is critical for survival [3]. One study compared the effects of bovine-derived freeze-dried collagen sponges to electrospun collagen scaffolds as the electrospun scaffolds offer greater homogeneity and homology to natural ECM than porous sponges. The scaffolds were spun from HFIP and populated with human dermal fibroblasts and epidermal keratinocytes. The results showed electrospun scaffolds were capable of producing similar cellular organization, proliferation, and maturation to the collagen sponges as well as an ability to reduce wound contraction. Chen et al. developed an acellular collagen composite scaffold loaded with vancomycin, gentamicin (both antibiotics), and lidocaine (local anesthetic) for addressing infected burns [52]. The scaffolds were electrospun from HFIP to create a PLGA/collagen base layer, followed by PLGA loaded with vancomycin, gentamicin, and lidocaine, and lastly a top layer of PLGA/collagen. In vitro release kinetics showed an initial burst release of all three drugs, followed by a slower sustained release. In the case of vancomycin, a second burst was seen at day 15. The nanofibers displayed no signs of cytotoxicity, but the assay suggests inferior cell viability in the nanofibrous drug-eluding membrane at 3 and 7 days, possibly due to the released antimicrobials having a harmful effect on viable tissue, thereby delaying healing. Local anesthetics may also cause adverse reactions. While initial studies on electrospun collagen for wound healing supported its use as a suitable tissue substitute, it failed to meet expectations. To improve upon cellular responses, collagen was incorporated into PCL fibers with fibronectin to develop a multifunctional nanofibrous scaffold appropriate for engineering hierarchical tissue [40]. In this case, PCL and collagen fibers were spun from HFIP to create scaffolds with both biological activity and mechanical stability. The results showed that dermal fibroblasts preferred the PCL/collagen fibers over the pure PCL fibers. The addition of fibronectin to the PCL fibers showed that the bioactivity of fibronectin was not affected by the electrospinning process as fibroblasts adhered and spread across the fibers, confirming findings in previous studies. In addition to wound healing and burn treatment, scaffolds have also been developed for use as drug delivery agents for cancer therapy [53]. A collagen/poly(N-isopropyl acrylamide) (PNIPAA)/chitosan mat containing 5-fluorouracil (anticancer agent) was produced via electrospinning. Collagen and chitosan were both dissolved in 2 % HCl, while PNIPAA was dissolved in water. The three polymeric components were combined with 5-FU and glutaraldehyde and electrospun. The prepared scaffolds showed an initial burst phase release

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with a cumulative percentage between 8 and 15 %, most likely due to drug molecules coating the fiber surfaces. The second phase of release is most likely due to swelling of the polymer network, facilitating release of the drug. The third and final phase of release is related to the dissolution of the polymer network that the drug was loaded in. By increasing the chitosan concentration, a slower rate of release could be obtained while the PNIPAA conferred antithrombogenic properties to the scaffold. Release of recombinant human periostin (rhPN) from collagen scaffolds prepared from HFIP was successfully demonstrated in a periostin knockout mouse experiment. The mice which received collagen and rhPN showed a significant increase in α-smooth muscle actin immunoreactivity than mice receiving collagen-only scaffolds [54]. Further work by Liu et al. focuses on the release of neurotrophin-3 (NT-3) and chondroitinase ABC (ChABC) from collagen scaffolds for spinal cord repair which remains a challenge due to rapid clearance of drugs from the site of injury [55]. Collagen scaffolds were prepared from 40 % acetic acid and soaked in a solution of NT-3 or ChABC (with and without heparin). The NT-3-soaked scaffolds showed an initial burst release (35.9±1.7 %) in day 1, followed by a sustained release for up to 28 days, with 72.5±3.1 % of NT3 released during the first 6 days. Sustained release of ChABC was obtained for at least 32 days, with enzymatic activity decreasing from 66.5±0.91 % on day 1 to 32.34±2.61 % on day 32. A recent paper by Lin et al. explores the novel use of zein to improve the electrospinnability of collagen [56]. Zein, which is a plant protein derived from corn, has been used to develop biodegradable films and matrices for tissue regeneration [57]. In addition to zein, an antibiotic (berberine) was loaded into the fibers to create a scaffold with antibiotic properties. By increasing the concentration of berberine in the fibers, the average fiber diameter increased. Furthermore, the drug release study showed an initial burst release followed by a slower, sustained release as seen in other work described here [56].

Conclusions From the evidence presented here, collagen has tremendous potential as a tissue engineering scaffold. As a material native to the human body, it offers a biocompatible option for addressing a wide range of disorders. We have demonstrated electrospinning to be an effective means of processing collagen into nanofibrous scaffolds for use as biomedical devices. Lastly, we have shown that electrospun collagen scaffolds can be functionalized in numerous ways, either through the addition of various pharmaceutical agents to treat infection or cancer or by blending collagen with other electrospun polymer fibers, whether naturally derived elastin or a synthetic polymer such as

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PCL. The fabrication of these composites, as well as any of the cross-linking methods, can produce mechanically stable and highly durable materials for just about any application. While we have given a very succinct overview of electrospun collagen, remember that the options are truly limitless as the materials are further advanced through new and different techniques.

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