DOI: 10.5301/ijao.5000317

Int J Artif Organs 2014; 37 ( 4 ) : 277-291

REVIEW

Recent advances in nerve tissue engineering Bill G.X. Zhang1,2,4, Anita F. Quigley2,3, Damian E. Myers1,4, Gordon G. Wallace3, Robert M.I. Kapsa2,3, Peter F.M. Choong1,4 Departments of Surgery, The University of Melbourne, St Vincent’s Hospital (Melbourne), Fitzroy - Australia Department of Neurology and Clinical Neuroscience, St Vincent’s Hospital (Melbourne), Fitzroy - Australia 3 ARC Centre for Electromaterials Science, University of Wollongong, Intelligent Polymer Research Institute, Wollongong Australia 4 Department of Orthopaedics, St Vincent’s Hospital (Melbourne), Fitzroy - Australia 1 2

Nerve injury secondary to trauma, neurological disease or tumor excision presents a challenge for surgical reconstruction. Current practice for nerve repair involves autologous nerve transplantation, which is associated with significant donor-site morbidity and other complications. Previously artificial nerve conduits made from polycaprolactone, polyglycolic acid and collagen were approved by the FDA (USA) for nerve repair. More recently, there have been significant advances in nerve conduit design that better address the requirements of nerve regrowth. Innovations in materials science, nanotechnology, and biology open the way for the synthesis of new generation nerve repair conduits that address issues currently faced in nerve repair and regeneration. This review discusses recent innovations in this area, including the use of nanotechnology to improve the design of nerve conduits and to enhance nerve regeneration. Keywords: Nerve, Conduit, Regeneration, Nanotechnology, Polymer Accepted: February 14, 2014

INTRODUCTION Nerve injury after trauma or surgery can lead to significant morbidity and disability. Despite developments in microsurgery, the results of recovery from nerve injury can often be disappointing with less than 40% of cases reporting satisfactory motor recovery (1). The traditional “gold standard” approach to therapy for nerve defects is autologous nerve transplant. However, this is limited by significant donor site morbidity, restricted range of harvest sites and mismatch in size between the graft and the recipient nerve. To overcome these shortcomings, various artificial and biologically-based nerve conduits have been developed and have achieved varying levels of success. Nerve conduits work on the principle of intubulation by enclosing the proximal and distal stump of the injured nerve within a cylindrical closed space that provides an

environment conducive to nerve regeneration. In order to optimize nerve regrowth, conduits must satisfy certain basic principles of design. In the first instance, conduits must minimize immunogenicity and inflammatory reactions that compromise nerve regrowth. Secondly, it is generally desirable that they be biodegradable while being sufficiently strong to support surgical manipulation (e.g., suturing), to prevent collapse and withstand pressure imparted by surrounding tissues. Thirdly, the walls of nerve conduits should promote efficient exchange of metabolic nutrients and waste while preventing the incursion of cells and biological materials that potentially interfere with active neural regeneration. The material surfaces within the conduit must optimally facilitate cell-substrate signaling and guidance of nerve regeneration to facilitate recapitulation of the ordered inner structure of the nerve. The FDA has approved a small range of biodegradable

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and biocompatible nerve conduits for repair of short nerve defects in humans (2), however, these conduits only partially satisfy the design principles mentioned above. As a result, motor function recovery after nerve repair using these approved products can be incomplete and the maximum length of repair limited to less than 4 cm in humans (3-6). The nerve regeneration thus achieved can also be accompanied by significant fibrosis with a layer of myofibroblasts surrounding and constricting the nerve, thereby impeding recovery and necessitating further invasive surgery for construct removal (7, 8). Recent advances in materials science, nanotechnology, and cell biology have resulted in the design of a new range of nerve repair conduits that aim to address the shortcomings of current conduits and better satisfy the basic principles of nerve scaffold design (9-13). In particular, the use of multimodal materials and biofactors that promote multiple and highly specific pro-neurogenic biomimetic effects may provide significant improvements on current designs. This review will outline some recent trends in nerve tissue engineering, including the use and incorporation of nanoscale materials, new growth factor delivery techniques, stem cells, and electropolymers into nerve conduits. Also, this review will explore how modern technology has advanced the therapeutic potential of biosynthetic nerve scaffolds through improvements to the conduit wall, the lumen, growth factor and cell delivery systems.

CONTEMPORARY ASPECTS OF NERVE REPAIR CONDUIT DESIGN Current nerve conduit designs are unable to fully replicate the natural niche of a regenerating nerve. Recently, in efforts to improve conduit performance, there has been a trend to downscaling the physical dimensions and improving the surface properties of scaffolds and supplementing the conduits with cells and growth factor delivery systems that mimic the physiological environment.

Scaffold fiber dimension and orientation Advances in nanoscale fabrication technologies make it possible to synthesize neural scaffolds with sub-micron architecture that approaches the topographical dimensions found within nerve extracellular matrix (ECM). Owing to their greater surface area-to-volume ratio, nanoscale scaffolds enhance 278

cell attachment, differentiation and growth when compared to microscale scaffolds (14). One of the most commonly employed techniques of nanofabrication is electrospinning. This technique involves the deposition of nanoscale fibers in a random or orientated manner, as required. Nerve conduits made from electrospun poly (ε-caprolactone-co-ethyl ethylene phosphate) (PCL-EEP) nanofibers with diameters of 3.96 um to 5.08 um have been used to bridge a 15 mm gap rat sciatic nerve defect, resulting in regeneration of all treated nerves at three months. This compares with regeneration apparent in only 50% of nerves using plain PCL-EEP conduits with smooth surfaces (15). Furthermore, the efficacy of the nerve constructs was enhanced by introducing aligned fibers orientated towards the axis of the nerve conduit on the conduit walls. Recent studies have reported that neurite length and linear orientation are both enhanced in neurons grown on aligned fibers in vitro (16-18). These findings are also reflected in vivo with aligned nanofibers significantly enhancing axon and Schwann cell proliferation compared to randomly orientated fiber conduits in rat nerve defect models (19).

Material composition Current nerve conduits approved for clinical use by the FDA are mainly synthesized from either biodegradable aliphatic polymers such as polyglycolic acid (PGA) or polyDL-lactide-caprolactone (PLCL) or from biological materials such as collagen 1 (2). Although biologically-based conduits often contain surfaces that facilitate interaction with cells, they are generally difficult to process and modify. Early nerve conduits based on biological materials often lack mechanical strength and/or are prone to collapse (20). However, with recent advancement in material science, biologically-based scaffold can be further modified physically and chemically to display acceptable mechanical properties. The main drawback of this type of scaffold is that there is usually significant batch-to-batch variability in construct quality and properties, making standardization of treatment and experimentation difficult. Compared to biologically-based conduits, artificial biodegradable polymers have the advantage of being more amenable to modification and processing with a greater control on variations in quality and properties. However, they are often hydrophobic and exhibit limited surface biomimetics for controlled cell signaling to promote specific and appropriate cell responses (21). As a result, there is a trend towards producing bio-synthetic materials by

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combining ECM molecules with biodegradable polymers, thereby combining the advantages of both classes of material. ECM molecules impart a hydrophilic surface to biodegradable polymers without significantly affecting the mechanical properties of the polymers (22). However, the effect of ECM biomolecules on the synthetic scaffold reaches a plateau after the concentration of the molecule exceeds a critical value. In fact, it has been reported that polycaprolactone (PCL) fibers containing 10% laminin have the same effect on neurite growth as those containing 30% and 50% laminin (22). In light of this finding, further research to optimize the types and amounts of biomolecules and synthetic compounds that comprise nerve scaffolds is warranted to maximize nerve regeneration. The potential disadvantages of using native ECM molecules such as laminin is that such materials need to be derived from animal sources; they confer the risk of immune rejection; and they may inadvertently introduce infection to the recipient. More recently, peptides that mimic the active binding domains of various ECM molecules have been used to produce bio-synthetic scaffolds (23, 24). These molecules do not pose an infection risk and are biocompatible with minimal risk of immune rejection (25). Peptides mimicking sequences of active domains on fibronectin, laminin-1, laminin-2, laminin 5 and collagen have been used to enhance the surface properties of artificial scaffolds in vivo. One of the earliest and most thoroughly characterized peptide is the RGD peptide. It is a sequence from the tenth type III repeat on the main cell binding domain of fibronectin (26). It promotes cell attachment through activation of multiple integrins on the cell surface and upregulates secretion of NGF, NT3, NT4 in regeneration nerves (27, 28). When collagen nerve conduits functionalized with RGD peptides were used to repair a 10 mm defect in rat sciatic nerve there was enhanced nerve regrowth and functional recovery at 90 days after surgery compared to plain collagen nerve conduits. This was associated with raised integrin alpha 5 expression and Schwann cell proliferation (23). The effect of RGD peptide can further be enhanced by addition of a second peptide PHSRN. PHSRN represents another sequence in the ninth type III repeat of the cellbinding domain of fibronectin (29). PHSRN is incapable of promoting cell adhesion by itself. However, when PHSRN is co-presented to the cell with RGD in a specific spatial array it acts synergistically with RGD to enhance cell spreading and adhesion (30). The spatial relationship between

RGD and PHSRN during presentation to the cell must resemble the natural spacing between the two sequences on the fibronectin molecule (29). Currently, PHSRN has only been employed to enhance RGD in bone and wound healing, with no study that has examined its effect in nerve regeneration (31, 32). In addition to RGD, many studies have also experimented with peptide sequences of laminin 1, 2, 5 and type 1 collagen. Laminin-1 related peptides YIGSR and IKVAV promote cell adhesion, Schwann cell proliferation, and nerve regeneration by binding to integrin V receptor and upregulating neurogenic growth factors and cytokines, including nerve growth factor (NGF), neurotrophin-3 (NT-3), neurotrophin-4 (NT4), and interleukin-1 (IL-1) (23, 27). In animal studies, IKVAV and YIGSR improve nerve regeneration after adsorption onto chitosan/hydroxyapatite (HA) nerve conduits. More importantly, in these studies, it was demonstrated that there was no significant difference in regeneration between groups receiving conduits coupled to laminin versus laminin peptides, suggesting minimal loss of function by using truncated laminin peptides (33). The effect of these laminin-1 peptides can be further enhanced by adding glycine spacers or linkers between the peptide sequence and the substrate to be coated (34, 35). This is proposed to increase the level of conformational degrees of freedom of the molecule and enhance its interaction with Schwann cells and regenerating neurons. More recently, researchers have also uncovered active sequences in laminin 2 and 5 that can also be exploited for tissue engineering. Both DLTIDDSYWYRI motif (Ln2-P3) and RNIPPFEGCIWN (Ln2-LG3-P2) from Large globular 1 and 3 domain from human laminin alpha 2 chain, respectively, promote neural cell adhesion and spreading (36-38). Ln2-P3 acts through syndecan-1 (38). Ln2-LG3-P2 binds to integrin alpha 3 beta 1 (37). Both peptide pathways converge on tyrosine phosphorylation of protein kinase C delta (37, 38). Ln2-P3 increased the number of myelinated fibers in vivo when they are coupled to the surface of artificial nerve conduits and used to repair a 10 mm defect in rat sciatic nerve (36). Similar to Ln2-LG3-P2, the laminin-5 related peptide PPFLMLLKGSTR also induces cell adhesion through activation of integrin alpha3 beta 1 and phosphorylation of focal adhesion kinase (39). However, it has only been studied in vivo in the setting of wound healing and its effects in nerve regeneration remain to be seen (40, 41). Collagen-related peptide sequence GFOGER also promotes cell adhesion

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(42-44); but like laminin-5 peptides, no study has tested its effects on nerve growth in vivo.

Physical construction and features Conduit wall permeability is important for effective removal of waste, promotion of new vessel infiltration, and nutrient diffusion into the lumen. Nerves repaired with permeable conduits show more effective nerve regeneration compared to impermeable ones (45). The size and morphology of pores on the nerve conduit wall have an important effect in determining nerve regrowth. Macroporous nerve conduits (83.5% porosity) lead to increased infiltration of myofibroblasts and perineural fibrosis compared to scaffolds with intermediate porosity (73.6% porosity) (46). Large pores with diameters ranging from 10 um to 20 um can also lead to increased surface area of hydrolysis of the biodegradable conduit and result in excessive swelling and obstruction of the conduit lumen (47). Current advances in nanofabrication have made it possible to control the size of pores in the nerve scaffold wall at both the microscale and the nanoscale. These advances have facilitated the development of asymmetric conduit walls with a microporous inner layer and a macrovoid outer layer conferring selective bidirectional permeability with low rates of inflow and high rates of outflow. When Schwann cells and fibroblasts were seeded onto the inner microporous and outer macroporous layers, respectively, there was increased proliferation of Schwann cells and inhibition of fibroblast growth. When the same conduits were used to repair a 10 mm defect in rat sciatic nerve there was increased number of myelinated A and B fibers in the middle of the conduit and the distal nerve after six weeks. The proposed mechanism was related to the neurogenic properties of increased Schwann cells and mitigated fibrosis secondary to reduce fibroblasts infiltration (46).

CONDUIT LUMEN FILLERS Current nerve conduits used in clinical practice lack scaffolds in the lumen that could potentially improve nerve guidance during regrowth. Adding a growth guidance scaffold to the lumen of nerve conduits can improve the length of repair achievable and functional outcomes after nerve repair (11, 48). In one study, nerve conduits with biodegradable aligned polyamide fibers in the lumen extended 280

the maximum length of sciatic nerve repair in rats from 10 mm to 15 mm (49). More recently, there has been a trend towards using nanofibers as lumen fillers. Nanomeshes made from blended PCL-laminin electrospun fibers with diameters ranging from 100 nm to 200 nm have been successfully used as lumen fillers in nerve conduits to repair 10 mm defects in rat tibial nerve. Recovery was superior in rats receiving conduits with aligned nanofibers compared to those receiving randomly oriented fibers, further emphasizing the importance of scaffold alignment (22). The neurogenic effect of the aligned fibers in the lumen is likely due to stabilization of the matrix rather than contactinduced cell signaling as the fibers become walled off from the axons by fibroblasts (49). Another problem commonly encountered during nerve repair is loss of target specificity after reinnervation and polyinnervation of muscles. To reduce this axonal “dispersion,” multiple longitudinal microchannels can be added to the center of the nerve conduit. Application of this principle has been shown to significantly reduce the number of neurons with dual projections to muscles and thereby significantly improve reinnervation accuracy (50). In addition to providing growth guidance and matrix stabilization in nerve conduits, lumen fillers may also relay important mechanical signals to cells affecting their growth and behavior. The mechanical qualities of the scaffold, such as stiffness, modulate the tension exerted by the growth cones on the neurites. Increases in substrate stiffness enhance the traction of neurites and alter cell gene expression (51, 52). However, under normal physiological control mechanisms, this tension must reach a critical threshold before nerve growth is activated (53, 54). Much research has focused on finding the ideal “stiffness” that corresponds to the optimal tension on the neurites during nerve growth. A number of groups have modified the stiffness of scaffolds by controlling the level of crosslinker and polymer concentration, such as the amount of bisacrylamide in polyacrylamide gels and the amount of polyethylene glycol (PEG) diacrylate in PEG-based hydrogels, and observing their effects on neural growth (55, 56). Others have adjusted the stiffness of the substrate by changing the concentration of ECM molecules such as fibrin, collagen, and fibronectin in hydrogels (51, 53, 57). In general, these studies have concluded that neural cells do not grow well on stiff substrates and that there exists an optimal window of substrate stiffness that is most conducive to neural growth. However, due to differences in experimental

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conditions and measurement endpoints there is a lack of consensus on the definition of optimal substrate stiffness with modulus values ranging from 5 Pa to 50 Pa depending on the constituents of the scaffold material (57). However, little is known about effects of substrate stiffness on neural growth in vivo. More research into optimizing substrate stiffness for lumen fillers in nerve conduits in animal models is required.

GROWTH FACTORS Absence of sustained neurotrophic stimulation accounts for at least some of the limited capacity of current nerve conduits to restore larger defects. To address this deficiency, a number of neurotrophic factors have been employed in nerve repair including: nerve growth factor (NGF), brainderived neurotrophic factor (BDNF), glial cell-derived neurotrophic factor (GDNF), ciliary neurotrophic factor (CNTF), neurotrophin 3, 4, 5 (NT-3,-4,-5), leukemia inhibitory factor (LIF), mechano growth factor 1 (MGF), vascular endothelial growth factor (VEGF), basic fibroblast growth factor (FGF2), and survival motor neuron-derived factor (SMDF) (11, 58-63). Some of these cytokines/growth factors have been used to promote multi-modal neurotrophic effects through targeting of diverse biological pathways, including pathways required for nerve regeneration (11, 60, 64, 65). Sustained growth factor release results in better nerve regeneration compared to rapid bolus release (66); as a result, various mechanisms for the sustained delivery of neurotrophic factors have been devised to achieve this end (67, 68). These delivery techniques include the immobilization of growth factors in hydrogels and microspheres and the use of heparin containing particulate systems for sustained delivery of heparin binding growth factors (66, 67, 69, 70). More recently, the nerve scaffold itself has been utilized as a vehicle for growth factor delivery. Conduits made from nanofibers electrospun from a mixture of GDNF and PCLEEP (poly(ε-caprolactone-co-ethyl ethylene phosphate)) have been successfully used to span a defect in rat sciatic nerve. These experiments demonstrate superior nerve regeneration in rats implanted with GDNFloaded PCLEEP nanofibers than with plain PCLEEP fibers alone (15). Growth factor delivery in such scaffolds results from a combination of diffusion from the scaffold surface and from release as the scaffold degrades. However, the distribution of growth factors in embedded scaffolds is

generally homogenous and fails to recapitulate growth factor gradients present in growing cell environments, thereby compromising directional guidance of nerve growth through the conduit (71-73). Growth factor gradients have been engineered into nerve scaffolds to simulate the physiological environment. In such conduits the concentration of the embedded growth factor increases from one end of the scaffold to the other. In one study, GDNF-loaded silk microspheres were distributed within the substance of the conduit in such a way as to create a growth factor gradient. When these scaffolds were used to repair a 15 mm gap in rat sciatic nerve there was significantly greater density of nerve regrowth compared to scaffolds with homogenously distributed GDNF six weeks after surgery (74). The embedding of growth factors into the fabric of the conduit requires consideration of a number of issues such as the effects on bioactivity of the molecule or the potential for unpredictable release profiles (75, 76). To address such limitations, other techniques for control of growth factor delivery have received attention, namely “core-shell” nanofibers and genetically modified cells. “Core-shell” nanofibers comprise a growth factor-laden core and a shell made of biodegradable polymers. These “core-shell” fibers can be synthesized from a process known as co-axial electrospinning, which involves simultaneous electrospinning of two solutions. One solution constitutes the “core” while the other solution forms a “sheath” surrounding the core. Coaxially electrospun poly(lactic-co-glycolic acid/basic fibroblast growth factor (PLGA/bFGF) fibers show a more sustained growth factor release compared to PLGA nanofibers embedded with bFGF (75). Furthermore, nerve defects repaired with conduits made of coaxially electrospun poly(lactic acidcaprolactone/nerve growth factor (P(LLA-CL)/NGF) fibers improve functional recovery compared to plain P(LLA-CL) fibers conduits loaded with NGF in the lumen (77). Neurotrophin delivery has also been achieved through the use of genetically modified stem cells and glia. Both Schwann cells and neural stem cells constitutively expressing GDNF or BDNF have been successfully employed as vectors for sustained growth factor delivery in nerve conduits with improved functional results after nerve repair (78, 79). The majority of these studies have been carried out using viral-based vectors for gene delivery. However, there are a number of safety concerns with the use of viral vectors, including immunogenicity and target specificity (80). As an alternative approach to viral-based gene therapies, plasmid-based gene delivery has also been used

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to increase expression of pro-neural factors in vivo. FGF isoforms (81, 82) as well as BDNF and NT-3 expressing Schwann cells (83) have been developed for the delivery of neurotrophins to neurons. Plasmid-based gene delivery also suffers from a number of limitations, including low transfection efficiency (approx. 20-40%) (84) and shortterm gene expression (82). However, plasmids generally do not confer the same safety concerns as virally-mediated gene delivery. The long-term impact of introducing genetically modified cells in vivo and associated health risks remain largely unknown. However, the use of vectors for gene delivery (both viral and plasmid) is a rapidly expanding area, not limited to the area of nerve regeneration. Since improvements to safety and efficiency are constantly occurring, with a number of vectors achieving clinical trial status (85), this therapeutic option is a real possibility in the near future. Further discussion of improvement in safety is not the subject of this review, however details on this can be found in an excellent review by Wirth and colleagues (86).

SUPPORT CELLS After nerve injury, resident Schwann cells become activated and switch from a myelinating state to a proliferating state (87, 88). This is accompanied by an upregulation of multiple growth associated genes such as NTR, GFAP, GAP-43, and netrin-1 (87). The Schwann cells secrete neurotrophins and form bands of Bunger to act as scaffolds to guide nerve regrowth (89). However, if nerve supply to the distal nerve stump is not re-established in a timely manner, the Schwann cells gradually lose their neurogenic potential and become “switched off” (90, 91). This lack of activated glial cells is postulated as one of the main reasons underlying the poor recovery after nerve injury. As a result many studies have examined supplementing nerve conduits with glial cells. Conduits seeded with Schwann cells have been used successfully to repair 2 cm defects in rat sciatic nerve, with functional recovery equivalent to nerve autograft at 6 months post-surgery (92). However, autologous Schwann cell use is limited by source availability and requirement for donor nerve sample for cell harvesting and exogenous Schwann cell transplantation can lead to immune rejection (93). Autologous neural stem cells, mesenchymal stems cells (MSCs), and induced pluripotent stem cells (iPSC) provide 282

alternative sources of trophic cells to autologous Schwann cells for neural tissue engineering. These stem cell types have the advantage that they can differentiate into any neural cell type required and have the capacity for selfrenewal, thereby potentially providing an unlimited supply of cells for seeding into conduits (94-97). Neural stem cells have been shown to improve nerve myelination and regeneration when grafted into chitosan-based nerve scaffolds (98). However, neural stem cells are predominantly derived from dentate gyrus of hippocampus and the subventricular zone of the adult brain and require a highly invasive procedure to extract brain tissue for cell isolation (99), limiting their applicability. From this viewpoint, MSCs cells that can be harvested from bone marrow, muscle, fat, or skin provide a more appropriate source of stem cells for such applications. MSCs are capable of differentiating into cells that are beyond the mesodermal lineage, such as neural and glial cells (100, 101). MSCs can differentiate into glial-like cells and promote neurite extension in vitro (102, 103). When collagen nerve conduits seeded with MSCs were used to repair a 3 mm defect in mice sciatic nerve there was increased number of myelinated fibers and improved functional outcomes at six weeks after surgery (104). Similar results have been reported by other authors using different nerve conduits and animal models (103-110). The main mechanisms that MSCs promote nerve regeneration are 1) secretion of neurotrophic factors and 2) trans-differentiation of the cells into glial-like cells that are directly incorporated into the growing nerve. MSCs secrete neurotrophins such as NGF, BDNF, bFGF, ciliary neurotrophic factor (CDNF) in vitro (105, 111, 112). The importance of this paracrine mechanism in nerve regeneration is best illustrated by the low survival rates of transplanted undifferentiated MSCs despite genuine improvements in nerve regeneration (108, 113, 114). This indicates that the improvements after stem cell treatment cannot be fully explained by stem cell transdifferentiation and that the MSCs are likely releasing neurotrophins to aid growth. The fate of the undifferentiated MSCs after implantation is a topic of great debate. Some studies postulate that the MSC transdifferentiate into Schwann cells, pointing to the observation that the implanted cells are incorporated into the growing nerve. However, others question the hypothesis and speculate that the MSCs are mainly fusing with the existing support cells (115). This is supported by the very low rate of expression of glial markers by the transplanted

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cells in vivo, which is inconsistent with the widespread incorporation of the cells into the nerve (106, 108, 113, 114, 116, 117). It is likely that both mechanisms are present, but the degree that each takes place is subject to differences in animal models and the type of nerve conduits used to deliver the MSCs. The phenotype and level of differentiation of the MSCs before cell seeding into the neural scaffold can have an impact on the success of the final reconstruction. Initially it was believed that by transplanting the MSCs in a naïve state the proliferating capacity of the cells can be preserved, thus minimizing the cell load needed on seeding. Furthermore, the milieu of the regenerating nerve will “instruct” and “teach” the stem cell to trans-differentiate into the cell that is required. However, in practice it was discovered that only a very low proportion of the implanted precursor cells differentiated into Schwann cells (0-6%) (106, 108, 113). As a result other authors have attempted to pre-differentiate the BMSCs before transplantation. With the aid of a cocktail of cytokines, approximately 50% to 60% of MSCs will differentiate and express glial markers in vitro (118, 119). When pre-differentiated MSCs were transplanted into collagen nerve conduits and used to repair a 12 mm defect in rat sciatic nerve, there were significantly more regenerating nerve fibers in the pre-differentiated stem cell group compared to rats that received undifferentiated MSCs three months after surgery (103, 120). Similar results have also been found by other authors with concomitant increases noted in nerve myelination and accelerated functional recovery in animals treated with differentiated MSCs (108, 110, 120). More recently, induced pluripotent stem cells (iPSCs) have become the focus of much research in nerve tissue engineering. iPSCs are derived from skin fibroblasts and can potentially provide an abundant source of autologous stem cells that display similar activity profiles to pluripotent stem cells (121). iPSCs seeded onto the walls of poly L-lactide (PLA) and poly ε-caprolactone (PCL)-based nerve conduits have been shown to improve both motor and sensory recovery when used to repair rat sciatic nerve defects (122). However, iPSCs can potentially form teratomas and require further research before clinical trials are possible (123). In addition, the necessity to transform cells with retrovirus in the production of iPSCs is a further limitation of this methodology to clinical translation. However, recent developments in iPSC technology that eliminate the need for retroviral transduction may overcome this limitation in future studies (124).

ELECTRICAL STIMULATION AND CONDUCTING POLYMERS Electrical stimulation has long been known to exert various effects on neural systems and is currently used in therapeutic applications. At a cellular level, electrical stimulation can promote neural and glial cell proliferation (125) as well as axonal outgrowth in vitro (17, 126, 127) and in vivo (128, 129). In terms of design, the incorporation of electrical conducting materials into nerve conduits is problematic and challenging. However, recent reports have focused on the use of organic conducting polymers (OCPs) as nervecompliant conducting structures in conduits (130-132). OCPs are polymerized from monomers, such as pyrrole or thiopene and have chemical backbones that are laden with loosely held electrons. During the polymerization process, a negatively charged counterion or “dopant” is employed to facilitate the polymerization reaction and to balance the charge on the polymer backbone. OCPs can be modified and manipulated using various processing techniques but maintain a high degree of electrical conductivity (133-135). The most common conductive polymer studied and implemented for tissue engineering applications is polypyrrole. Polypyrrole exhibits excellent biocompatibility and has excellent characteristics as a structural and ‘active’ scaffold in promoting neurite growth under electrical stimulation in vitro (134) as well as in vivo (131). A potential limitation for the in vivo use of polypyrrole is that it is not biodegradable and like many other artificial polymers, lacks a clearly defined biologically active surface. To address these limitations polypyrrole has been incorporated with a range of biodegradable polymers and biologically active molecules including neurotrophins, peptides, and cell adhesion molecules (136-139). Various approaches have been used to promote the breakdown of pyrrole-based conductive polymers in vivo. One approach involves embedding polypyrrole nanoparticles into a biodegradable polymer (poly (D, l-lactide) (PDLLA)) membrane, resulting in a final construct that was then degradable in vitro. In these studies, incorporation of polypyrrole nanoparticles significantly improved the electrical conductivity of the composite material compared to plain PDLLA membranes (140). Polypyrrole can also be doped with biologically active molecules to enhance bioactivity. Primary cortical neurons grown on polypyrrole doped with laminin-related peptides CDPGYIGSR or RNIAEIIKDI show higher cell density and neurite length compared to polypyrrole doped with non-biological molecules

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such as poly (styrene sulphonate) (138). In addition, polypyrole can be doped with NT3, NGF, BDNF either singly or in combination and designed for controlled cytokine release via electrical stimulation to promote neurotrophic effects in vitro (137). Recently, polypyrrole/chitosan-based nerve conduits have been evaluated for delivering electrical stimulation to enhance nerve repair in vivo. Rats that received blended polypyrrole/chitosan nerve conduits combined with electrical stimulation displayed superior nerve regeneration across a 15 mm nerve gap compared to plain chitosan scaffolds that were electrically stimulated (141). The conductive polymer conduit effectively isolated the electrical stimulation to the site of the nerve defect, providing a high degree of target specificity. The electroconductivity of most conductive polymers tends to decline after prolonged stimulation due to dopant “depletion” (142). As a result, more stable electro-conductive materials have been explored as a means to introduce electrical stimulation to tissue engineering in vivo. One such material is carbon nanotubes. Carbon nanotubes (CNTs) are nanoscale tubes rolled from sheets of carbon atoms arranged in hexagonal and pentagonal arrays. CNTs are 100 times stronger than steel and 1 000 times more electrically conductive than copper, with high capacitance and low impedance, while occupying a fraction of normal metal weight (143). CNTs provide a stable and reliable substrate for electrical stimulation in tissue engineering. Like conductive polymers, carbon nanotubes can be modified/functionalized by addition of functional chemical moieties to their surface or blending the CNTs with biological molecules to improve the bioactivity and biocompatibility of the surface (144-146). Functionalization of CNTs with laminin and collagen has been shown to improve nerve cell attachment while facilitating effective electrical stimulation of cells for enhancement of neuron differentiation and growth (147, 148). However, some studies have reported toxic properties of CNTs (149-151). The exact mechanism of toxicity is unclear. One study linked the cell toxicity to oxidative stress secondary to metal impurities within the nanotubes, with resulting accumulation of intracellular peroxide (149, 151). Other studies have reported the inhibition of nuclear translocation of intracellular growth proteins such as SMAD and depletion of environmental nutrients after cells were cultured with CNT suspensions (150, 151). Most studies have shown that CNT-related toxicity is associated with loosely bound CNTs in suspensions (152). Other studies have shown that CNTs, firmly 284

bound to or within the substrate, reduce cytotoxicity to undetectable levels (153). Nevertheless, concerns about CNT biocompatibility have hampered efforts to introduce CNTs into nerve conduits in vivo. Further studies are required to address the in vivo safety of carbon nanotubes in longerterm experiments. In addition to conductive polymers such as polypyrrole and CNTs, a number of conductive hydrogels have also been recently developed for the specific purpose of in vivo stimulation, particularly neural stimulation. PEDOT (154) and poly (3,4 ethylene dioxythiophene) (155)-based hydrogels have been developed for this purpose. These polymers are synthesized by incorporation of conducting polymers directly into a hydrogel or by polymerization of the conducting polymer through an existing hydrogel. The improved mechanical properties of hydrogel materials over rigid polymers or metals will most likely confer better integration in host tissues, minimizing inflammatory response and glial scarring around the electrode site. From this perspective, conductive hydrogels present a promising approach for the electrical stimulation of regenerating neural tissues.

SUMMARY AND FUTURE PERSPECTIVES Neural tissue engineering based on the principle of intubation has advanced significantly since the first experiments using silicone tubes to guide neuronal growth. Convergent developments in nanofabrication, polymers, gene and growth factor delivery and stem cell technologies enable new features to be incorporated and combined as a multimodal approach for the design of nerve conduits (Fig. 1). Techniques such as electrospinning and biofactor immobilization can be used to incorporate nanoscale surface topography, selective permeability, improved biocompatibility and specified bioactivity into nerve conduits. Lumen fillers consisting of aligned micro- and/or nano-scale features and fibers with optimized neuronal and neurogenic biomimetic properties may facilitate spatially orientated and structured nerve regeneration. Also, stem cells and glial cells can be seeded into scaffolds to provide glial cell support and growth factor delivery. Neurotrophin release mechanisms and glial supportive cells can be incorporated to provide a sustained (as necessary) and reliable supply of neurotrophins to the regenerating nerve. Finally, developments in electro-conductive materials such as conductive polymers and carbon nanotubes introduce the possible

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B

A

C

Fig. 1 - Schematic diagram of a nerve conduit with its main components and some recent advances in conduit component design. A) Electron microscope (EM) image of electrospun PLA fibers creating nanoscale topography for neural growth (images produced by the authors). B) Lumen fillers consisting of aligned PLGA fibers in electrospun PLA conduits. Image adapted from (11). C) Electron microscopy (EM) image of asymmetric permeable conduit walls with microporous inner layer and macrovoid outer layer allowing for preferential outflow across the wall compared to inflow. This maximizes waste removal and maintains effective nutrient exchange. Image adapted from (46). D) Electrospun PLA conduits used to repair a sciatic nerve defect in a rat. Image adapted from (11).

D

incorporation of novel electrical modalities that may benefit and expedite nerve regeneration. The convergence of these various disciplines will enable design of conduits that incorporate multi-modal stimulation for improved nerve regeneration. Identification and fine-tuning of conduit features that promote and optimize multiple neuroregenerative responses will create more effective conduits for nerve repair. In particular, addressing the spatio-temporal requirements for effective neuroregenerative responses can be facilitated; with these emerging technologies it is now possible to initiate structured nerve repair patterned from inherent neurobiological processes. Knowledge of the basic scientific mechanisms underlying the neurogenic effects of various materials and structures that will undoubtedly emerge from this exciting sub-discipline of tissue engineering will provide additional cues towards building highly effective biosynthetic nerve repair conduits. Molecular targets that are still emerging from current studies may be integrated into nerve conduit designs. This in turn, will undoubtedly improve nerve regeneration and facilitate a more pro-active development of new materials for nerve tissue engineering. The exciting developments in bio-synthetic nerve repair conduits evidenced in recent initiatives embrace major advances

in neurobiology and the materials sciences. Additive fabrication technologies (e.g., 3D printing of materials (156), cells (157), and functionalization of scaffold substrates (158)) have emerged that may address the spatial distribution of multiple components within nerve conduits. These technologies have the potential to provide surgeons with the means by which to digitally capture the dimensions of a nerve injury for tailored “printing” of scaffolds for nerve repair. With further developments, down to the nano-scale level, additive fabrication aligned with bioengineering technologies offers the potential for the design and even tailored construction of effective biosynthetic neural conduits for nerve repair. Financial Support: This work was funded through National Health and Medical Research Council (NHMRC) postgraduate Scholarship Scheme (BG-XZ), NHMRC project (Project ID635243; AFQ) and Australian Research Council (CE140100012; GGW, RMIK) funding sources. Conflict of Interest: None. Address for correspondence: Professor Peter F. Choong, MBBS, MD FRACS, FAOrthA Department of Surgery St Vincent’s Hospital (Melbourne) 41 Victoria Parade Fitzroy VIC 3065, Australia [email protected]

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REFERENCES 1.

2.

3.

4. 5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

286

Moneim M, Omer G. Clinical outcome following acute nerve repair. In: Omeg G, Spinner M, Van Beek A, eds. Management of peripheral nerve problems. Vol. 414. Philadelphia: Saunders; 1998:414-419. Meek MF, Coert JH. US Food and Drug Administration/Conformit Europe-approved absorbable nerve conduits for clinical repair of peripheral and cranial nerves. Ann Plast Surg. 2008;60(1):110-116. Pabari A, Yang SY, Seifalian AM, Mosahebi A. Modern surgical management of peripheral nerve gap. J Plast Reconstr Aesthet Surg. 2010;63(12):1941-1948. Navissano M, Malan F, Carnino R, Battiston B. Neurotube for facial nerve repair. Microsurgery. 2005;25(4):268-271. Weber RA, Breidenbach WC, Brown RE, Jabaley ME, Mass DP. A randomized prospective study of polyglycolic acid conduits for digital nerve reconstruction in humans. Plast Reconstr Surg. 2000;106(5):1036-1045, discussion 1046-1048. Battiston B, Geuna S, Ferrero M, Tos P. Nerve repair by means of tubulization: literature review and personal clinical experience comparing biological and synthetic conduits for sensory nerve repair. Microsurgery. 2005;25(4):258-267. Chamberlain LJ, Yannas IV, Arrizabalaga A, Hsu HP, Norregaard TV, Spector M. Early peripheral nerve healing in collagen and silicone tube implants: myofibroblasts and the cellular response. Biomaterials. 1998;19(15):1393-1403. Merle M, Dellon AL, Campbell JN, Chang PS. Complications from silicon-polymer intubulation of nerves. Microsurgery. 1989;10(2):130-133. Zhan X, Gao M, Jiang Y, et al. Nanofiber scaffolds facilitate functional regeneration of peripheral nerve injury. Nanomedicine. 2013;9(3):305-315. Biazar E, Heidari Keshel S. A nanofibrous PHBV tube with Schwann cell as artificial nerve graft contributing to rat sciatic nerve regeneration across a 30-mm defect bridge. Cell Commun Adhes. 2013;20(1-2):41-49. Quigley AF, Bulluss KJ, Kyratzis IL, et al. Engineering a multimodal nerve conduit for repair of injured peripheral nerve. J Neural Eng. 2013;10(1):016008. Jin J, Park M, Rengarajan A, et al. Functional motor recovery after peripheral nerve repair with an aligned nanofiber tubular conduit in a rat model. Regen Med. 2012;7(6):799-806. Lotfi P, Garde K, Chouhan AK, Bengali E, Romero-Ortega MI. Modality-specific axonal regeneration: toward selective regenerative neural interfaces. Front Neuroeng. 2011;4:11. Yang F, Murugan R, Wang S, Ramakrishna S. Electrospinning of nano/micro scale poly(L-lactic acid) aligned fibers and their potential in neural tissue engineering. Biomaterials. 2005;26(15):2603-2610. Chew SY, Mi R, Hoke A, Leong KW. Aligned Protein-Polymer Composite Fibers Enhance Nerve Regeneration: A Potential Tissue-Engineering Platform. Adv Funct Mater. 2007;17(8): 1288-1296.

16. Yao L, O’Brien N, Windebank A, Pandit A. Orienting neurite growth in electrospun fibrous neural conduits. J Biomed Mater Res B Appl Biomater. 2009;90B(2):483-491. 17. Quigley AF, Razal JM, Thompson BC, et al. A ConductingPolymer Platform with Biodegradable Fibers for Stimulation and Guidance of Axonal Growth. Adv Mater. 2009;21(43): 4393-4397. 18. Liu X, Chen J, Gilmore KJ, Higgins MJ, Liu Y, Wallace GG. Guidance of neurite outgrowth on aligned electrospun polypyrrole/poly(styrene-beta-isobutylene-beta-styrene) fiber platforms. J Biomed Mater Res A. 2010;94(4):1004-1011. 19. Kim YT, Haftel VK, Kumar S, Bellamkonda RV. The role of aligned polymer fiber-based constructs in the bridging of long peripheral nerve gaps. Biomaterials. 2008;29(21):3117-3127. 20. Wang S, Cai L. Polymers for Fabricating Nerve Conduits. Int J Polym Sci. 2010;2010:Article ID 138686:1-20. 21. Prabhakaran MP, Venugopal JR, Chyan TT, et al. Electrospun biocomposite nanofibrous scaffolds for neural tissue engineering. Tissue Eng Part A. 2008;14(11):1787-1797. 22. Neal RA, Tholpady SS, Foley PL, Swami N, Ogle RC, Botchwey EA. Alignment and composition of laminin-polycaprolactone nanofiber blends enhance peripheral nerve regeneration. J Biomed Mater Res A. 2011. 23. Rafiuddin Ahmed M, Jayakumar R. Peripheral nerve regeneration in RGD peptide incorporated collagen tubes. Brain Res. 2003;993(1-2):208-216. 24. Seo SY, Min S-K, Bae HK, et al. A laminin-2-derived peptide promotes early-stage peripheral nerve regeneration in a dual-component artificial nerve graft. J Tissue Eng Regen Med. 2013;7(10):788-800. 25. Hersel U, Dahmen C, Kessler H. RGD modified polymers: biomaterials for stimulated cell adhesion and beyond. Biomaterials. 2003;24(24):4385-4415. 26. Ruoslahti E. RGD and other recognition sequences for integrins. Annu Rev Cell Dev Biol. 1996;12:697-715. 27. Ahmed MR, Basha SH, Gopinath D, Muthusamy R, Jayakumar R. Initial upregulation of growth factors and inflammatory mediators during nerve regeneration in the presence of cell adhesive peptide-incorporated collagen tubes. J Peripher Nerv Syst. 2005;10(1):17-30. 28. Pierschbacher MD, Ruoslahti E. Cell attachment activity of fibronectin can be duplicated by small synthetic fragments of the molecule. Nature. 1984;309(5963):30-33. 29. Aota S, Nomizu M, Yamada KM. The short amino acid sequence Pro-His-Ser-Arg-Asn in human fibronectin enhances cell-adhesive function. J Biol Chem. 1994;269(40): 24756-24761. 30. Hojo K, Susuki Y, Maeda M, et al. Amino acids and peptides. Part 39: a bivalent poly(ethylene glycol) hybrid containing an active site (RGD) and its synergistic site (PHSRN) of fibronectin. Bioorg Med Chem Lett. 2001;11(11):1429-1432. 31. Benoit DS, Anseth KS. The effect on osteoblast function of colocalized RGD and PHSRN epitopes on PEG surfaces. Biomaterials. 2005;26(25):5209-5220.

© 2014 Wichtig Publishing - ISSN 0391-3988

Zhang et al

32. Hattori A, Hozumi K, Ko JA, et al. Sequence specificity of the PHSRN peptide from fibronectin on corneal epithelial migration. Biochem Biophys Res Commun. 2009;379(2):346-350. 33. Itoh S, Yamaguchi I, Suzuki M, et al. Hydroxyapatite-coated tendon chitosan tubes with adsorbed laminin peptides facilitate nerve regeneration in vivo. Brain Res. 2003;993 (1-2):111-123. 34. Wang W, Itoh S, Matsuda A, et al. Enhanced nerve regeneration through a bilayered chitosan tube: the effect of introduction of glycine spacer into the CYIGSR sequence. J Biomed Mater Res A. 2008;85A(4):919-928. 35. Tong YW, Shoichet MS. Enhancing the neuronal interaction on fluoropolymer surfaces with mixed peptides or spacer group linkers. Biomaterials. 2001;22(10):1029-1034. 36. Seo SY, Min SK, Bae HK, et al. A laminin-2-derived peptide promotes early-stage peripheral nerve regeneration in a dual-component artificial nerve graft. J Tissue Eng Regen Med. 2013;7(10):788-800. 37. Jung SY, Kim JM, Min SK, et al. The potential of laminin-2-biomimetic short peptide to promote cell adhesion, spreading and migration by inducing membrane recruitment and phosphorylation of PKCδ. Biomaterials. 2012;33(15):3967-3979. 38. Jung SY, Kim JM, Kang HK, Jang H, Min BM. A biologically active sequence of the laminin alpha2 large globular 1 domain promotes cell adhesion through syndecan-1 by inducing phosphorylation and membrane localization of protein kinase Cdelta. J Biol Chem. 2009;284(46):31764-31775. 39. Kim JM, Park WH, Min BM. The PPFLMLLKGSTR motif in globular domain 3 of the human laminin-5 alpha3 chain is crucial for integrin alpha3beta1 binding and cell adhesion. Exp Cell Res. 2005;304(1):317-327. 40. Min SK, Lee SC, Hong SD, Chung CP, Park WH, Min BM. The effect of a laminin-5-derived peptide coated onto chitin microfibers on re-epithelialization in early-stage wound healing. Biomaterials. 2010;31(17):4725-4730. 41. Damodaran G, Tiong WH, Collighan R, Griffin M, Navsaria H, Pandit A. In vivo effects of tailored laminin-332 α3 conjugated scaffolds enhances wound healing: a histomorphometric analysis. J Biomed Mater Res A. 2013;101(10):2788-2795. 42. Reyes CD, García AJ. Engineering integrin-specific surfaces with a triple-helical collagen-mimetic peptide. J Biomed Mater Res A. 2003;65A(4):511-523. 43. Wojtowicz AM, Shekaran A, Oest ME, et al. Coating of biomaterial scaffolds with the collagen-mimetic peptide GFOGER for bone defect repair. Biomaterials. 2010;31(9):2574-2582. 44. Reyes CD, García AJ. Alpha2beta1 integrin-specific collagen-mimetic surfaces supporting osteoblastic differentiation. J Biomed Mater Res A. 2004;69A(4):591-600. 45. Rodríguez FJ, Gómez N, Perego G, Navarro X. Highly permeable polylactide-caprolactone nerve guides enhance peripheral nerve regeneration through long gaps. Biomaterials. 1999;20(16):1489-1500. 46. Chang CJ, Hsu SH, Yen HJ, Chang H, Hsu SK. Effects of unidirectional permeability in asymmetric poly(DL-lactic acid-

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

co-glycolic acid) conduits on peripheral nerve regeneration: an in vitro and in vivo study. J Biomed Mater Res B Appl Biomater. 2007;83B(1):206-215. Meek MF, Den Dunnen WF. Porosity of the wall of a Neurolac nerve conduit hampers nerve regeneration. Microsurgery. 2009;29(6):473-478. Lee JY, Giusti G, Friedrich PF, et al. The effect of collagen nerve conduits filled with collagen-glycosaminoglycan matrix on peripheral motor nerve regeneration in a rat model. J Bone Joint Surg Am. 2012;94(22):2084-2091. Arai T, Lundborg G, Dahlin LB. Bioartificial nerve graft for bridging extended nerve defects in rat sciatic nerve based on resorbable guiding filaments. Scand J Plast Reconstr Surg Hand Surg. 2000;34(2):101-108. Yao L, de Ruiter GC, Wang H, et al. Controlling dispersion of axonal regeneration using a multichannel collagen nerve conduit. Biomaterials. 2010;31(22):5789-5797. Man AJ, Davis HE, Itoh A, Leach JK, Bannerman P. Neurite outgrowth in fibrin gels is regulated by substrate stiffness. Tissue Eng Part A. 2011;17(23-24):2931-2942. Koch D, Rosoff WJ, Jiang J, Geller HM, Urbach JS. Strength in the periphery: growth cone biomechanics and substrate rigidity response in peripheral and central nervous system neurons. Biophys J. 2012;102(3):452-460. Leach JB, Brown XQ, Jacot JG, Dimilla PA, Wong JY. Neurite outgrowth and branching of PC12 cells on very soft substrates sharply decreases below a threshold of substrate rigidity. J Neural Eng. 2007;4(2):26-34. Lamoureux P, Zheng J, Buxbaum RE, Heidemann SR. A cytomechanical investigation of neurite growth on different culture surfaces. J Cell Biol. 1992;118(3):655-661. Gunn JW, Turner SD, Mann BK. Adhesive and mechanical properties of hydrogels influence neurite extension. J Biomed Mater Res A. 2005;72A(1):91-97. Flanagan LA, Ju YE, Marg B, Osterfield M, Janmey PA. Neurite branching on deformable substrates. Neuroreport. 2002;13(18):2411-2415. Willits RK, Skornia SL. Effect of collagen gel stiffness on neurite extension. J Biomater Sci Polym Ed. 2004;15(12): 1521-1531. Nectow AR, Marra KG, Kaplan DL. Biomaterials for the development of peripheral nerve guidance conduits. Tissue Eng Part B Rev. 2012;18(1):40-50. Huang J, Xiang J, Yan Q, Li S, Song L, Cai X. Dog tibial nerve regeneration across a 30-mm defect bridged by a PRGD/ PDLLA/β-TCP/NGF sustained-release conduit. J Reconstr Microsurg. 2013;29(2):77-87. Dai LG, Huang GS, Hsu SH. Sciatic Nerve Regeneration by Co-Cultured Schwann Cells and Stem Cells on Microporous Nerve Conduits. Cell Transplant. 2013:22(11):20292039. de Boer R, Borntraeger A, Knight AM, et al. Short- and long-term peripheral nerve regeneration using a polylactic-co-glycolic-acid scaffold containing nerve growth

© 2014 Wichtig Publishing - ISSN 0391-3988

287

Advances in nerve engineering

factor and glial cell line-derived neurotrophic factor releasing microspheres. J Biomed Mater Res A. 2012;100A(8): 2139-2146. 62. Cao J, Xiao Z, Jin W, et al. Induction of rat facial nerve regeneration by functional collagen scaffolds. Biomaterials. 2013;34(4):1302-1310. 63. Grothe C, Haastert K, Jungnickel J. Physiological function and putative therapeutic impact of the FGF-2 system in peripheral nerve regeneration--lessons from in vivo studies in mice and rats. Brain Res Brain Res Rev. 2006;51(2):293-299. 64. Deister C, Schmidt CE. Optimizing neurotrophic factor combinations for neurite outgrowth. J Neural Eng. 2006;3(2): 172-179. 65. Hobson MI, Green CJ, Terenghi G. VEGF enhances intraneural angiogenesis and improves nerve regeneration after axotomy. J Anat. 2000;197(4):591-605. 66. Xu X, Yee WC, Hwang PY, et al. Peripheral nerve regeneration with sustained release of poly(phosphoester) microencapsulated nerve growth factor within nerve guide conduits. Biomaterials. 2003;24(13):2405-2412. 67. Lee AC, Yu VM, Lowe JB III, et al. Controlled release of nerve growth factor enhances sciatic nerve regeneration. Exp Neurol. 2003;184(1):295-303. 68. Yang Y, De Laporte L, Rives CB, et al. Neurotrophin releasing single and multiple lumen nerve conduits. J Control Release. 2005;104(3):433-446. 69. Midha R, Munro CA, Dalton PD, Tator CH, Shoichet MS. Growth factor enhancement of peripheral nerve regeneration through a novel synthetic hydrogel tube. J Neurosurg. 2003;99(3):555-565. 70. Joung YK, Bae JW, Park KD. Controlled release of heparinbinding growth factors using heparin-containing particulate systems for tissue regeneration. Expert Opin Drug Deliv. 2008;5(11):1173-1184. 71. Kennedy TE, Wang H, Marshall W, Tessier-Lavigne M. Axon guidance by diffusible chemoattractants: a gradient of netrin protein in the developing spinal cord. J Neurosci. 2006;26(34):8866-8874. 72. Cao X, Shoichet MS. Investigating the synergistic effect of combined neurotrophic factor concentration gradients to guide axonal growth. Neuroscience. 2003;122(2):381-389. 73. Gundersen RW, Barrett JN. Neuronal chemotaxis: chick dorsal-root axons turn toward high concentrations of nerve growth factor. Science. 1979;206(4422):1079-1080. 74. Lin YC, Ramadan M, Hronik-Tupaj M, et al. Spatially controlled delivery of neurotrophic factors in silk fibroin-based nerve conduits for peripheral nerve repair. Ann Plast Surg. 2011;67(2):147-155. 75. Sahoo S, Ang LT, Goh JC, Toh SL. Growth factor delivery through electrospun nanofibers in scaffolds for tissue engineering applications. J Biomed Mater Res A. 2010;93(4): 1539-1550. 76. Kenawy R, Bowlin GL, Mansfield K, et al. Release of tetracycline hydrochloride from electrospun poly(ethylene-co-

288

77.

78.

79.

80.

81.

82.

83.

84.

85.

86. 87.

88. 89.

90. 91. 92.

vinylacetate), poly(lactic acid), and a blend. J Control Release. 2002;81(1-2):57-64. Liu JJ, Wang CY, Wang JG, Ruan HJ, Fan CY. Peripheral nerve regeneration using composite poly(lactic acid-caprolactone)/nerve growth factor conduits prepared by coaxial electrospinning. J Biomed Mater Res A. 2011;96A(1):13-20. Li Q, Ping P, Jiang H, Liu K. Nerve conduit filled with GDNF gene-modified Schwann cells enhances regeneration of the peripheral nerve. Microsurgery. 2006;26(2):116-121. Fu KY, Dai LG, Chiu IM, Chen JR, Hsu SH. Sciatic nerve regeneration by microporous nerve conduits seeded with glial cell line-derived neurotrophic factor or brain-derived neurotrophic factor gene transfected neural stem cells. Artif Organs. 2011;35(4):363-372. Madduri S, Gander B. Schwann cell delivery of neurotrophic factors for peripheral nerve regeneration. J Peripher Nerv Syst. 2010;15(2):93-103. Timmer M, Robben S, Müller-Ostermeyer F, Nikkhah G, Grothe C. Axonal regeneration across long gaps in silicone chambers filled with Schwann cells overexpressing high molecular weight FGF-2. Cell Transplant. 2003;12(3):265-277. Haastert K, Lipokatic E, Fischer M, Timmer M, Grothe C. Differentially promoted peripheral nerve regeneration by grafted Schwann cells over-expressing different FGF-2 isoforms. Neurobiol Dis. 2006;21(1):138-153. Pettingill LN, Minter RL, Shepherd RK. Schwann cells genetically modified to express neurotrophins promote spiral ganglion neuron survival in vitro. Neuroscience. 2008;152(3): 821-828. Haastert K, Mauritz C, Chaturvedi S, Grothe C. Human and rat adult Schwann cell cultures: fast and efficient enrichment and highly effective non-viral transfection protocol. Nat Protoc. 2007;2(1):99-104. Mueller C, Flotte TR. Clinical gene therapy using recombinant adeno-associated virus vectors. Gene Ther. 2008;15(11): 858-863. Wirth T, Parker N, Ylä-Herttuala S. History of gene therapy. Gene. 2013;525(2):162-169. Walsh S, Midha R. Practical considerations concerning the use of stem cells for peripheral nerve repair. Neurosurg Focus. 2009;26(2):E2. Mirsky R, Jessen KR. The neurobiology of Schwann cells. Brain Pathol. 1999;9(2):293-311. Navarro X, Vivó M, Valero-Cabré A. Neural plasticity after peripheral nerve injury and regeneration. Prog Neurobiol. 2007;82(4):163-201. Weinberg HJ, Spencer PS. The fate of Schwann cells isolated from axonal contact. J Neurocytol. 1978;7(5):555-569. Hall SM. The biology of chronically denervated Schwann cells. Ann N Y Acad Sci. 1999;883:215-233. Sinis N, Schaller HE, Schulte-Eversum C, et al. Nerve regeneration across a 2-cm gap in the rat median nerve using a resorbable nerve conduit filled with Schwann cells. J Neurosurg. 2005;103(6):1067-1076.

© 2014 Wichtig Publishing - ISSN 0391-3988

Zhang et al

93. Guénard V, Kleitman N, Morrissey TK, Bunge RP, Aebischer P. Syngeneic Schwann cells derived from adult nerves seeded in semipermeable guidance channels enhance peripheral nerve regeneration. J Neurosci. 1992;12(9):3310-3320. 94. Vescovi AL, Parati EA, Gritti A, et al. Isolation and cloning of multipotential stem cells from the embryonic human CNS and establishment of transplantable human neural stem cell lines by epigenetic stimulation. Exp Neurol. 1999;156(1): 71-83. 95. Jiang Y, Jahagirdar BN, Reinhardt RL, et al. Pluripotency of mesenchymal stem cells derived from adult marrow. Nature. 2002;418(6893):41-49. 96. Sarugaser R, Hanoun L, Keating A, Stanford WL, Davies JE. Human mesenchymal stem cells self-renew and differentiate according to a deterministic hierarchy. PLoS One. 2009; 4(8):e6498. 97. Emborg ME, Liu Y, Xi J, et al. Induced pluripotent stem cellderived neural cells survive and mature in the nonhuman primate brain. Cell Rep. 2013;3(3):646-650. 98. Guo BF, Dong MM. Application of neural stem cells in tissueengineered artificial nerve. Otolaryngol Head Neck Surg. 2009;140(2):159-164. 99. Bottai D, Fiocco R, Gelain F, et al. Neural stem cells in the adult nervous system. J Hematother Stem Cell Res. 2003; 12(6):655-670. 100. Dezawa M, Kanno H, Hoshino M, et al. Specific induction of neuronal cells from bone marrow stromal cells and application for autologous transplantation. J Clin Invest. 2004; 113(12):1701-1710. 101. Wislet-Gendebien S, Hans G, Leprince P, Rigo JM, Moonen G, Rogister B. Plasticity of cultured mesenchymal stem cells: switch from nestin-positive to excitable neuron-like phenotype. Stem Cells. 2005;23(3):392-402. 102. Brohlin M, Mahay D, Novikov LN, et al. Characterisation of human mesenchymal stem cells following differentiation into Schwann cell-like cells. Neurosci Res. 2009;64(1): 41-49. 103. Ladak A, Olson J, Tredget EE, Gordon T. Differentiation of mesenchymal stem cells to support peripheral nerve regeneration in a rat model. Exp Neurol. 2011;228(2):242-252. 104. Pereira Lopes FR, Camargo de Moura Campos L, Dias Corrêa J Jr, et al. Bone marrow stromal cells and resorbable collagen guidance tubes enhance sciatic nerve regeneration in mice. Exp Neurol. 2006;198(2):457-468 105. Chen CJ, Ou YC, Liao SL, et al. Transplantation of bone marrow stromal cells for peripheral nerve repair. Exp Neurol. 2007;204(1):443-453. 106. Cuevas P, Carceller F, Dujovny M, et al. Peripheral nerve regeneration by bone marrow stromal cells. Neurol Res. 2002;24(7):634-638. 107. Mohammadi R, Azizi S, Delirezh N, Hobbenaghi R, Amini K, Malekkhetabi P. The use of undifferentiated bone marrow stromal cells for sciatic nerve regeneration in rats. Int J Oral Maxillofac Surg. 2012;41(5):650-656.

108. Shimizu S, Kitada M, Ishikawa H, Itokazu Y, Wakao S, Dezawa M. Peripheral nerve regeneration by the in vitro differentiated-human bone marrow stromal cells with Schwann cell property. Biochem Biophys Res Commun. 2007;359(4): 915-920. 109. Oliveira JT, Almeida FM, Biancalana A, et al. Mesenchymal stem cells in a polycaprolactone conduit enhance mediannerve regeneration, prevent decrease of creatine phosphokinase levels in muscle, and improve functional recovery in mice. Neuroscience. 2010;170(4):1295-1303. 110. Gu JH, Ji YH, Dhong ES, Kim DH, Yoon ES. Transplantation of adipose derived stem cells for peripheral nerve regeneration in sciatic nerve defects of the rat. Curr Stem Cell Res Ther. 2012;7(5):347-355. 111. Crigler L, Robey RC, Asawachaicharn A, Gaupp D, Phinney DG. Human mesenchymal stem cell subpopulations express a variety of neuro-regulatory molecules and promote neuronal cell survival and neuritogenesis. Exp Neurol. 2006;198(1):54-64. 112. Gu Y, Wang J, Ding F, Hu N, Wang Y, Gu X. Neurotrophic actions of bone marrow stromal cells on primary culture of dorsal root ganglion tissues and neurons. J Mol Neurosci. 2010;40(3):332-341. 113. Pan HC, Cheng FC, Chen CJ, et al. Post-injury regeneration in rat sciatic nerve facilitated by neurotrophic factors secreted by amniotic fluid mesenchymal stem cells. J Clin Neurosci. 2007;14(11):1089-1098. 114. Heine W, Conant K, Griffin JW, Höke A. Transplanted neural stem cells promote axonal regeneration through chronically denervated peripheral nerves. Exp Neurol. 2004;189(2): 231-240. 115. Kuroda Y, Kitada M, Wakao S, Dezawa M. Bone marrow mesenchymal cells: how do they contribute to tissue repair and are they really stem cells? Arch Immunol Ther Exp (Warsz). 2011;59(5):369-378. 116. Dezawa M, Takahashi I, Esaki M, Takano M, Sawada H. Sciatic nerve regeneration in rats induced by transplantation of in vitro differentiated bone-marrow stromal cells. Eur J Neurosci. 2001;14(11):1771-1776. 117. Marchesi C, Pluderi M, Colleoni F, et al. Skin-derived stem cells transplanted into resorbable guides provide functional nerve regeneration after sciatic nerve resection. Glia. 2007;55(4):425-438. 118. Keilhoff G, Goihl A, Stang F, Wolf G, Fansa H. Peripheral nerve tissue engineering: autologous Schwann cells vs. transdifferentiated mesenchymal stem cells. Tissue Eng. 2006;12(6):1451-1465. 119. Caddick J, Kingham PJ, Gardiner NJ, Wiberg M, Terenghi G. Phenotypic and functional characteristics of mesenchymal stem cells differentiated along a Schwann cell lineage. Glia. 2006;54(8):840-849. 120. Wang X, Luo E, Li Y, Hu J. Schwann-like mesenchymal stem cells within vein graft facilitate facial nerve regeneration and remyelination. Brain Res. 2011;1383:71-80.

© 2014 Wichtig Publishing - ISSN 0391-3988

289

Advances in nerve engineering

121. Löhle M, Hermann A, Glass H, et al. Differentiation efficiency of induced pluripotent stem cells depends on the number of reprogramming factors. Stem Cells. 2012;30(3):570-579. 122. Uemura T, Takamatsu K, Ikeda M, et al. Transplantation of induced pluripotent stem cell-derived neurospheres for peripheral nerve repair. Biochem Biophys Res Commun. 2012;419(1):130-135. 123. Miura K, Okada Y, Aoi T, et al. Variation in the safety of induced pluripotent stem cell lines. Nat Biotechnol. 2009;27(8): 743-745. 124. Okita K, Nakagawa M, Hyenjong H, Ichisaka T, Yamanaka S. Generation of mouse induced pluripotent stem cells without viral vectors. Science. 2008;322(5903):949-953. 125. Huang J, Hu X, Lu L, Ye Z, Zhang Q, Luo Z. Electrical regulation of Schwann cells using conductive polypyrrole/chitosan polymers. J Biomed Mater Res A. 2010;93(1):164-174. 126. Kimura K, Yanagida Y, Haruyama T, Kobatake E, Aizawa M. Electrically induced neurite outgrowth of PC12 cells on the electrode surface. Med Biol Eng Comput. 1998;36(4): 493-498. 127. Liu X, Gilmore KJ, Moulton SE, Wallace GG. Electrical stimulation promotes nerve cell differentiation on polypyrrole/poly (2-methoxy-5 aniline sulfonic acid) composites. J Neural Eng. 2009;6(6):065002. 128. Gordon T, et al. Brief electrical stimulation accelerates axon regeneration in the peripheral nervous system and promotes sensory axon regeneration in the central nervous system. Motor Control. 2009;13(4):412-441. 129. Udina E, Furey M, Busch S, Silver J, Gordon T, Fouad K. Electrical stimulation of intact peripheral sensory axons in rats promotes outgrowth of their central projections. Exp Neurol. 2008;210(1):238-247. 130. George PM, Saigal R, Lawlor MW, et al. Three-dimensional conductive constructs for nerve regeneration. J Biomed Mater Res A. 2009;91A(2):519-527. 131. Durgam H, Sapp S, Deister C, et al. Novel degradable co-polymers of polypyrrole support cell proliferation and enhance neurite out-growth with electrical stimulation. J Biomater Sci Polym Ed. 2010;21(10):1265-1282. 132. Wang X, Gu X, Yuan C, et al. Evaluation of biocompatibility of polypyrrole in vitro and in vivo. J Biomed Mater Res A. 2004;68A(3):411-422. 133. Liu X, Yue Z, Higgins MJ, Wallace GG. Conducting polymers with immobilised fibrillar collagen for enhanced neural interfacing. Biomaterials. 2011;32(30):7309-7317. 134. Lee JY, Bashur CA, Goldstein AS, Schmidt CE. Polypyrrolecoated electrospun PLGA nanofibers for neural tissue applications. Biomaterials. 2009;30(26):4325-4335. 135. Aznar-Cervantes S, Roca MI, Martinez JG, et al. Fabrication of conductive electrospun silk fibroin scaffolds by coating with polypyrrole for biomedical applications. Bioelectrochemistry. 2012;85:36-43. 136. Gomez N, Schmidt CE. Nerve growth factor-immobilized polypyrrole: bioactive electrically conducting polymer for

290

enhanced neurite extension. J Biomed Mater Res A. 2007; 81A(1): 135-149. 137. Thompson BC, Richardson RT, Moulton SE, et al. Conducting polymers, dual neurotrophins and pulsed electrical stimulation—dramatic effects on neurite outgrowth. J Control Release. 2010;141(2):161-167. 138. Stauffer WR, Cui XT. Polypyrrole doped with 2 peptide sequences from laminin. Biomaterials. 2006;27(11):2405-2413. 139. Xie J, Macewan MR, Willerth SM, et al. Conductive CoreSheath Nanofibers and Their Potential Application in Neural Tissue Engineering. Adv Funct Mater. 2009;19(14):2312-2318. 140. Shi G, Rouabhia M, Wang Z, Dao LH, Zhang Z. A novel electrically conductive and biodegradable composite made of polypyrrole nanoparticles and polylactide. Biomaterials. 2004;25(13):2477-2488. 141. Huang J, Lu L, Zhang J, et al. Electrical stimulation to conductive scaffold promotes axonal regeneration and remyelination in a rat model of large nerve defect. PLoS One. 2012;7(6):e39526. 142. Green RA, Lovell NH, Wallace GG, Poole-Warren LA. Conducting polymers for neural interfaces: challenges in developing an effective long-term implant. Biomaterials. 2008; 29(24-25):3393-3399. 143. Thostenson ET, Ren Z, Chou T-W. Advances in the science and technology of carbon nanotubes and their composites: a review. Compos Sci Technol. 2001;61(13):1899-1912. 144. Ni Y, Hu H, Malarkey EB, et al. Chemically functionalized water soluble single-walled carbon nanotubes modulate neurite outgrowth. J Nanosci Nanotechnol. 2005;5(10):1707-1712. 145. Hu H, Ni Y, Mandal SK, et al. Polyethyleneimine functionalized single-walled carbon nanotubes as a substrate for neuronal growth. J Phys Chem B. 2005;109(10):4285-4289. 146. Sayes CM, Liang F, Hudson JL, et al. Functionalization density dependence of single-walled carbon nanotubes cytotoxicity in vitro. Toxicol Lett. 2006;161(2):135-142. 147. Cho Y, Borgens RB. The effect of an electrically conductive carbon nanotube/collagen composite on neurite outgrowth of PC12 cells. J Biomed Mater Res A. 2010;95A(2):510-517. 148. Kam NW, Jan E, Kotov NA. Electrical stimulation of neural stem cells mediated by humanized carbon nanotube composite made with extracellular matrix protein. Nano Lett. 2009; 9(1):273-278. 149. Shvedova AA, Castranova V, Kisin ER, et al. Exposure to carbon nanotube material: assessment of nanotube cytotoxicity using human keratinocyte cells. J Toxicol Environ Health A. 2003;66(20):1909-1926. 150. Mooney E, Dockery P, Greiser U, Murphy M, Barron V. Carbon nanotubes and mesenchymal stem cells: biocompatibility, proliferation and differentiation. Nano Lett. 2008; 8(8):2137-2143. 151. Liu D, Yi C, Zhang D, Zhang J, Yang M. Inhibition of proliferation and differentiation of mesenchymal stem cells by carboxylated carbon nanotubes. ACS Nano. 2010;4(4): 2185-2195.

© 2014 Wichtig Publishing - ISSN 0391-3988

Zhang et al

152. Edwards SL, Werkmeister JA, Ramshaw JA. Carbon nanotubes in scaffolds for tissue engineering. Expert Rev Med Devices. 2009;6(5):499-505. 153. Nayagam DAX, Williams RA, Chen J, et al. Biocompatibility of immobilized aligned carbon nanotubes. Small. 2011;7(8): 1035-1042. 154. Mario Cheong GL, Lim KS, Jakubowicz A, Martens PJ, Poole-Warren LA, Green RA. Conductive hydrogels with tailored bioactivity for implantable electrode coatings. Acta Biomater. 2014;10(3):1216-26. 155. Abidian MR, Daneshvar ED, Egeland BM, Kipke DR, Cederna PS, Urbanchek MG. Hybrid conducting polymer-hydrogel

conduits for axonal growth and neural tissue engineering. Adv Healthc Mater. 2012;1(6):762-767. 156. Weng B, Shepherd RL, Crowley K, Killard AJ, Wallace GG. Printing conducting polymers. Analyst (Lond). 2010; 135:2779-2789. 157. Ferris CJ, Gilmore KJ, Beirne S, McCallum D, Wallace GG, Pahuis M. Bio-ink for on-demand printing of living cells. Biomater Sci. 2013;1:224-230. 158. Koh HS, Yong T, Chan CK, Ramakrishna S. Enhancement of neurite outgrowth using nano-structured scaffolds coupled with laminin. Biomaterials. 2008;29(26):3574-3582.

© 2014 Wichtig Publishing - ISSN 0391-3988

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