Recent advances in stem cell-mediated peripheral nerve repair.

Review Accepted after revision: October 28, 2014 Published online: March 21, 2015

Cells Tissues Organs DOI: 10.1159/000369450

Recent Advances in Stem Cell-Mediated Peripheral Nerve Repair Joey Grochmal Rajiv Midha Department of Clinical Neuroscience, Faculty of Medicine, University of Calgary, Hotchkiss Brain Institute, Calgary, Alta., Canada

Key Words Stem cell biology · Regeneration · Peripheral nervous system · Myelin · Skin-derived precursor cells · Rodent

Abstract A major advance in the field of peripheral nerve repair has been the advent of stem and progenitor cell use to supplement the regenerative environment in animal models of nerve injury. As Schwann cell replacements, stem cells may be even better suited to promoting regeneration in these scenarios. We review the recent literature detailing the search for the definitive Schwann cell replacement cell, including a look at genetic modification of transplanted cells for nerve injury repair. © 2015 S. Karger AG, Basel

postinjury with neurotrophic factors playing key roles in axonal growth and survival [Chen et al., 2007b]. On the other hand, the maintenance of a supportive regenerative environment is also fundamental for successful axonal regeneration. Schwann cells (SCs) seem to be key to creating and maintaining this permissive regenerative environment [Fu and Gordon, 1997]. Following injury, SCs quickly change their phenotype from a myelinating to a growth-supportive mode [Fu and Gordon, 1997; Mirsky and Jessen, 1999]. They also proliferate postinjury in the distal stump, and migrate to form basal lamina-lined columns, or bands of Büngner [Büngner, 1891]. This prolif-

Abbreviations used in this paper

A multitude of factors affect the ability of a nerve to regenerate following nerve injury. From an axonal standpoint, intrinsic responses are essential for the conversion of neurons from a signaling to a regenerative neuronal phenotype. Upregulation of growth-related genes occurs © 2015 S. Karger AG, Basel 1422–6405/15/0000–0000$39.50/0 E-Mail [email protected] www.karger.com/cto

adipose-derived stem cells brain-derived neurotrophic factor dorsal root ganglia glial cell-derived neurotrophic factor myelin basic protein nerve growth factor peripheral nervous system Schwann cell skin-derived precursor Schwann cells

Dr. Joey Grochmal, MD 3330 Hospital Drive NW HM-109 Calgary, AB T2N 4N1 (Canada) E-Mail joey.grochmal @ albertahealthservices.ca

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Introduction

ADSCs BDNF DRG GDNF MBP NGF PNS SC SKP-SCs

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PNS regeneration. Although studies to date have yet to pinpoint the mechanism of their benefit, initial work has demonstrated functional improvement after sciatic nerve injury by repair with graft constructs seeded with induced pluripotent stem cell spheres differentiated towards a glial lineage [Uemura et al., 2012; Ikeda et al., 2013]. The detailed literature regarding the beneficial effects of these and other cell types in peripheral nerve regeneration has been reviewed extensively elsewhere [Mizuno, 2009; Walsh and Midha, 2009a, b; Liu et al., 2011]. Most stem cells with regenerative SC-like potential exist initially in an undifferentiated state. It would seem that encouraging these cells towards an SC phenotype has a positive effect on their ability to assist in nerve regeneration, a phenomenon initially described by Dezawa et al. [2001] whereby differentiated bone marrow-derived mesenchymal stem cells encouraged a more robust regenerative response in sciatic nerve, as compared to the undifferentiated counterpart. The cocktail of mitogens used to induce differentiation is tailored to the cell type in question, but usually includes the use of multiple glial growth factors (bFGF, PDGF, forskolin and neuregulin, as examples). Supporting this notion, differentiated ADSCs in vitro will encourage a robust neurite response of cocultured motor neuron-like cells, similar to that produced by SCs, while undifferentiated ADSCs fail to do so [Kingham et al., 2007]. In coculture with dissociated dorsal root ganglia (DRG) neurons, undifferentiated ADSCs do not tightly associate with neurites or assume a spindletype morphology over time, while differentiated ADSCs will not only do so, but will also form myelin basic protein (MBP)-positive myelin segments [Tomita et al., 2012]. Recently, human differentiated ADSCs have also been shown to produce greater neurotrophin quantity than undifferentiated human ADSCs (fig. 1) [Tomita et al., 2013]. Skin-derived precursor SCs (SKP-SCs) similarly demonstrate enhanced neurotrophin production over undifferentiated SKP spheres in vitro [Walsh et al., 2009a]. Of special interest to our lab, SKPs are multipotent stem cells that reside in the dermal papillae of mammalian skin [Toma et al., 2001] and have successfully been harvested and cultured from both rodents [Toma et al., 2001] and humans [Toma et al., 2005]. These cells can take on the morphology of SCs when exposed to the appropriate mitogens and in this state are referred to as SKP-SCs. These cells can myelinate axons in culture and appear to myelinate both PNS axons in normal and shiverer mutants distal to crush injury [McKenzie et al., 2006]. When transplanted into an acellular nerve graft, these Grochmal/Midha


eration peaks at about 3 days postinjury for previously myelinating SCs [Clemence et al., 1989; Fu and Gordon, 1997]. SCs that have dedifferentiated to the growth-supportive phenotype [Walsh and Midha, 2009b] secrete neurotrophins and cytokines [Tofaris et al., 2002], while elaborating extracellular matrix proteins that are growth supportive [Bunge, 1994]. They can also play a role in phagocytosis of debris at the early time points after peripheral nervous system (PNS) injury, but their main role is indirect as they act to recruit macrophages to the nerve undergoing wallerian degeneration. However, studies have demonstrated that the capacity for endogenous SCs to maintain a growth-supportive phenotype decreases over time [You et al., 1997], which may be a major reason for the decreased capacity of chronically denervated nerves to support axonal regeneration. Unfortunately, the inability to rapidly expand donor autologous SCs for transplantation into an injured host is currently a limiting factor for the clinical translation of SC transplant therapy [Nishiura et al., 2004], while the invasive nature of SC procurement, essentially involving nerve biopsy and sacrifice, currently makes these cells nonideal for clinical therapeutics. Therefore, research has focused on finding alternative Schwann-like cell types for transplantation. A pragmatic definition of an ‘ideal’ stem cell for clinical use includes the following qualifiers: it should be easily obtainable through a low-morbidity procedure, while being amenable to rapid in vitro expansion, genetic modification and incorporation into the host tissue, without the need for immunosuppression upon transplant [Azizi et al., 1998; Walsh and Midha, 2009a]. To date, many different types of cells have been investigated to fit this role. Bone marrow stromal cells have been shown to produce neurotrophins and improve regeneration when transplanted into peripheral nerve in vivo [Chen et al., 2007a] and may also have a limited capacity to migrate (‘home’) to injured areas of the central nervous system when injected in a systemic fashion [Akiyama et al., 2002]. Neural progenitor cells have been used in a peripheral nerve gap model, where they may contribute to improved morphological parameters of regeneration [Murakami et al., 2003]. Adipose-derived stem cells (ADSCs) are also potential candidates in this regard, transdifferentiating to an SC-like phenotype when exposed to mitogens in vitro [Kingham et al., 2007] and have been extensively studied as an adjunct to nerve repair. Induced pluripotent stem cells can now be differentiated along a neural lineage [Okada et al., 2008] and are another potential source of cell supplementation therapy under investigation for

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cells secrete neurotrophins and appear to improve early nerve regeneration (as compared to media injection), as well as improve upon morphometric parameters of regeneration [Walsh et al., 2009a]. Furthermore, these cells may be capable of reinvigorating the regenerative environment of chronically denervated nerve [Walsh et al., 2009b]. Coupled with their ease of harvest and culture expansion, these cells of interest have demonstrated great promise as a therapeutic option for the treatment of nerve injury. Recently, our work with both SKP-SCs and genetically modified SCs, along with recent work from other laboratories, suggests that this definition should perhaps be expanded. We assert that an ideal stem or precursor cell for SC transplantation should perform at least as well, if not

better, than host SCs or cultured nerve-derived SCs. In PNS injury models, these performance metrics may include debris clearance, production of growth factors, myelin formation and graft survival, with the end result being improved regenerative outcomes. The rest of this review will focus on recent progress by our lab and others towards characterizing and developing this caliber of cell for nerve injury repair in the context of recent similar advances in the field of regenerative biology. Beyond the goal of SC replacement therapy, a fascinating prospect in the field of nerve regeneration is that stem/precursor cells may be more biologically effective than nerve-derived autologous SCs at promoting regenerative outcomes. Much of the early literature demonstrated at least an equivalence of stem cell- and SC-facil-

Recent Advances in Stem Cell-Mediated Peripheral Nerve Repair


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enhanced neurotrophic profile as compared to undifferentiated human ADSCs (uhADSCs). Concentrations of neurotrophins in conditioned media from 4 separate cell lines after 48 h of culture as measured by ELISA: BDNF (a), NGF (b) and GDNF (c). Data are expressed as the mean ± SEM (n = 3). * p < 0.05, ** p < 0.01, ++ p < 0.01 vs. all uhADSC groups, ## p < 0.01 vs. all dhADSC groups (one-way ANOVA followed by post hoc Tukey’s test). hSC = Human Schwann cells; P1, P2, P3 and P4 = donor patients 1-4. Adapted from Tomita et al. [2013].

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itated regeneration outcomes. di Summa et al. [2010] showed increased axonal regeneration into fibrin-conduit gaps seeded with both differentiated ADSCs and differentiated mesenchymal stem cells, on par with that seen in SC-seeded conduits. Our own lab has also shown an equivalence between SKP-SC and SC treatment at promoting early (4 weeks) axonal regeneration in a freezethawed autograft repair paradigm [Walsh et al., 2009a]. However, individual data points from many of these cell therapy manuscripts have suggested that the chosen cell replacement therapy may be further enhancing the regenerative environment. Recent studies using differentiated ADSCs have shown trends towards superiority over SC treatment in certain injury models. In a fibrin conduit regeneration model they improve many parameters of recovery in a manner more robust than differentiated mesenchymal stem cells or SCs, and close to that achieved with autograft repair [di Summa et al., 2011]. In chronic injury models, ADSCs injected into the reapproximated distal stump of the common peroneal nerve 8 weeks post-transection demonstrated significant improved recovery over the noninjected control group, with a trend towards superiority over the SC-injected group in multiple parameters of regeneration [Tomita et al., 2012], including the walking track-determined peroneal function index, terminal time point compound muscle action potential maxima, and tibialis anterior wet muscle weights. They also seem to improve acute recovery across a gap injury, though perhaps not better than gold-standard fresh autograft containing an endogenous SC population [Liu et al., 2011]. From our own lab, we have seen suggestions that SKP-SCs hold the potential to act as an ‘enhanced’ SC [Walsh et al., 2009a; Grochmal et al., 2014; 4

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Khuong et al., 2014]. The pertinent question remains whether or not these enhanced ‘observations’ can translate into any real recovery benefit for patients with nerve injury. To address this, our most recent study focused on the behavioral recovery of rodents undergoing SKP-SC therapy in several injury paradigms [Khuong et al., 2014]. In a 10-mm-gap freeze-thawed autograft repair model, SKPSC-treated animals demonstrated improved functional recovery (skilled locomotion as assessed with ladder rung and tapered beam) over SC-treated and nerve isograftrepaired animals by the study terminus of 17 weeks. Furthermore, they likewise improve behavioral recovery after both chronic and acute repair, as compared to mediatreated groups. In particular, an improvement on an acute nerve repair paradigm by adjunctive SKP-SC therapy demonstrates a step forward over the current standard treatment for microsurgical repair of nerve transection injury, and suggests that these cells may be supplementing the support provided by endogenous SCs to improve the regenerative milieu as early as 5 weeks into the recovery process (fig. 2).

Potential Mechanisms whereby Stem Cells May Be ‘Outworking’ SCs in the Injured PNS

Debris Clearance and Modification of an Inhibitory Regenerative Environment It is becoming increasingly apparent that clearing the early postinjury environment of myelin debris may play a key part in improving regenerative outcomes. In vivo inhibition of phagocytic myeloid cells after sciatic nerve Grochmal/Midha


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Fig. 2. SKP-SCs enhance skilled locomotion. Following acute nerve repair, distal tibial nerves received immediate injection of either SKP-SCs, dead SKP-SCs or media. Using the ladder rung test, the SKP-SCtreated group demonstrated significantly enhanced recovery at 5 weeks when compared to the nonrepair group, the dead SKP-SC group and the media group (acute repair equivalent group). The slip ratio is defined as the mean number of errors/step for the injured limb, with an error being classified as a total miss, a deep slip or a slight slip on an individual ladder rung. ** p < 0.01; *** p < 0.001. Adapted from Khuong et al. [2014].

Myelination by Stem Cells and Improvement of Endogenous Myelination The ability of a differentiated stem cell to form myelin is often used as convincing evidence that a particular cell of interest can function as an SC in a physiologic manner [McKenzie et al., 2006; Tomita et al., 2013; Grochmal et al., 2014]. Congruently, the ability of transplanted stem cells to engage and encourage the myelination capacity of endogenous SCs may be a major mechanism by which they encourage recovery from nerve injury [Grochmal et al., 2014]. One interesting finding by Khuong et al. [2014] was that SKP-SCs improved early myelination after acute nerve repair. Keeping in mind that the previously reported survival percentages for naïve SKP injection into acutely injured nerve (10.48 ± 2.9%) are relatively low [Walsh et al., 2011], this was likely a secondary effect by SKP-SCs on endogenous remyelination in that environment. Direct stem cell myelination, as well as indirect effects on endogenous remyelination, have been demonstrated for many cell types of interest to nerve regeneration. Differentiated bone marrow mesenchymal stem cells have been shown to express P0, PMP22 and MBP proteins in

coculture with DRG neurons [Mantovani et al., 2010]. These cells have shown convincing evidence of myelination in vivo (rodent sciatic nerve), as evidenced by immunoelectron microscopy of transplanted myelin-forming green fluorescent protein-labeled cells that also express myelin-associated glycoprotein on immunohistochemical assay [Dezawa et al., 2001]. Differentiated ADSCs also express the aforementioned myelin proteins in coculture [Mantovani et al., 2010], and there is ample evidence of their ability to directly myelinate axons [Tomita et al., 2012, 2013]. They have also been linked to improved endogenous remyelination responses after injury. Liu et al. [2011] demonstrated overall greater myelin thickness post-ADSC transplantation into acellular nerve grafts for sciatic nerve gap injury. This result was similar to that found using differentiated ADSCs seeded in fibrin conduits, whereby they encouraged a regenerated myelin area parameter better than SCs, and almost on par with an autograft [di Summa et al., 2011]. Initial evidence that SKP-SCs can myelinate PNS axons was first established by McKenzie et al. [2006], who demonstrated MBP-positive myelin profiles of SKP-SCs cocultured with shiverer DRG explants, and also injected postsciatic crush into shiverer mice. These findings have since been confirmed in nonmutant paradigms [Walsh et al., 2011]. To address the functionality of SKP-SC-derived myelin, and its relevance in regard to improving myelin repair postinjury, our lab has recently studied SKP-SC injection into a demyelination/remyelination doxorubicin injury paradigm [England et al., 1988]. Injection of low doses of doxorubicin into the tibial nerve results in a delayed demyelination injury [England et al., 1988] free from significant axonopathy [Grochmal et al., 2014], and constitutes a model whereby the effects of cell therapy (injected in a delayed fashion) can be studied free from the confounder of axonal regeneration. We found that not only did SKP-SCs myelinate axons in vivo, but they also formed mature nodes of Ranvier and resultantly improved the electrophysiological recovery of this injury (fig. 3). While this suggests that SKP-SC

9 postinjury demonstrated significant improvement in tibial nerve recovery dynamics over media-injected nerves, as evidenced by improved foot plantar compound muscle action potential responses (day 48, p = 0.03, paired t test ± SEM). b A spectral unmixed image of a teased nerve fiber from a DiI (cell tracker CM-DiI)-labeled SKP-SC-injected tibial nerve 14 days after cell injection, demonstrating mature SKP-SC myelination (colocalization of DiI

with the myelin dye Nile red), including NaV1.6 (voltage-gated sodium channel subtype 1.6) nodal positivity. c An axial cryosection of tibial nerve also stained with Nile red (myelin), demonstrating green fluorescent protein-positive SKP-SC myelination on cross-section. SKP-SCs promote an earlier return to baseline Gratios in this model (d) and myelinate a small but measurable percentage of axons 14 days after cell injection (e). * p > 0.05 (d). Adapted from Grochmal et al. [2013]. (For figure see next page.)



Fig. 3. a Adriamycin-injured nerves injected with SKP-SCs at day

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crush attenuates the axonal regeneration response, and thereby prevents effective functional recovery [Barrette et al., 2008]. Also, it has recently been suggested that there is both an age-associated decline in the ability of endogenous SCs to function as debris-clearance cells, as well as a likewise decline in their ability to signal an endogenous macrophage response [Painter et al., 2014]. It is likely that stem-cell therapy may be modulating the early regenerative environment, perhaps by encouraging a more effective macrophage response. For example, Khuong et al. [2014] were able to demonstrate that SKP-SC therapy improved clearance of myelin debris from the distal segment of an acutely repaired nerve both 4 and 8 weeks after the repair, compared to media or dead cell treatment groups. Considering the inhibitory nature of nondegenerated myelin [Bisby and Chen, 1990; Brown et al., 1992], this observation may help to explain the improved acute repair outcomes.

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Fig. 4. SKP-SCs secrete measurable levels of NGF, neurotrophin-3 and BDNF. In particular, they secrete higher levels of NGF and BDNF as compared to SCs. NTF = Neurotrophic factor. * p < 0.05; *** p < 0.001. Adapted from Walsh et al. [2009a].

myelin is indeed functional, the quantified percentage of SKP-SC-derived myelin reached only about 15% on average. Considering that the SKP-SC-injected nerves also displayed a significantly higher total average G-ratio than the SC-injected nerves, the data also suggest that SKP-SCs facilitate early augmentation of the host SC response. Improved Cell Survival Cell survival in PNS transplant models is an infrequently reported statistic and, of note, many pioneering works on peripheral nerve SC replacement therapy [Guenard et al., 1992; Gulati et al., 1995] were done before the widespread use of modern cell tracing methods [Mosahebi et al., 2001]. The very importance of cell survival is still debated, as researchers have demonstrated improvement in regenerative outcomes after cell therapy with no clear evidence of significant long-term cell transplant survival or integration into the host milieu [Prockop, 2007; Marcus et al., 2008]. In general, it seems that SC-like stem cell transplants have survival ranges between approximately 5 and 15% [McKenzie et al., 2006; Shimizu et al., 2007; Walsh et al., 2011], with the highest survival seen in acute nerve injury, and lower survival seen in chronic nerve injury models and also in noninjured nerve. One suggestion in these studies is that neurotrophic support from dedifferentiated endogenous SCs may be increasing graft surRecent Advances in Stem Cell-Mediated Peripheral Nerve Repair

vival. The ideal cell for transplant may therefore benefit from a concomitant boost of trophic support administered with the graft injection. Of note, post-transplant apoptosis rates of amniotic fluid-derived stem cells seem attenuated by granulocyte colony-stimulating factor administration [Pan et al., 2009]. Recently, engineered SCs with enhanced neurotrophin-secreting abilities have been shown to have better survival in spinal cord than nonenhanced SCs [Kanno et al., 2014]. Our own work suggests that heuregulin administration may decrease the apoptosis of transplanted SKP-SCs, a finding supported by better SKP-SC survival in acutely injured as opposed to intact nerve [Walsh et al., 2011]. On the other hand, SCs transplanted into the spinal cord have been shown to undergo significant survival loss in the first week, and the vast majority of the loss was secondary to necrosis rather than apoptosis. These findings suggest that toxic or anoxic environments present within the injury zone may also contribute to the survival of transplanted cells in a significant way [Hill et al., 2007]. Interestingly, conditioning of SC-like stem cells by growth on supportive extracellular matrices has been shown to increase their ability to survive apoptotic conditions in vitro and to also increase their ability to support DRG neurite extension in coculture [di Summa et al., 2013]. Transplanted graft cell survival has recently been positively correlated with recovery outcomes in manuscripts that address both spinal cord injury [Kanno et al., 2014] and Parkinson’s disease. These results should hopefully provide the impetus for similar quantified study of survival phenomena in the PNS. Enhanced Neurotrophic Action by SC-Like Precursor Cells Transplanted stem cells may be enhancing the regenerative environment of the injured nerve through secretion of bioactive neurotrophic molecules [Walsh et al., 2009a; Zhao et al., 2009; Lopatina et al., 2011; Sowa et al., 2012]. Indeed, they may be inherently better at this than SCs. Our own research has shown that SKP-SCs secrete brain-derived neurotrophic factor (BDNF; trend), nerve growth factor (NGF; p < 0.01) and neurotrophin-3 (p < 0.05) at higher levels than SCs in culture (fig. 4) [Walsh et al., 2009a]. ADSCs have shown similar enhancement, producing greater amounts of BDNF, glial growth factor, neuregulin-1, vascular endothelial growth factor, hepatocyte growth factor and insulin-like growth factor mRNA than SCs in culture [Sowa et al., 2012]. This enhanced profile may contribute to positive regenerative effects even beyond the zone of nerve injury. For example, transCells Tissues Organs DOI: 10.1159/000369450

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Fig. 5. Doxycycline-regulated GDNF-producing SC treatment for nerve injury repair (5-mm gap sciatic nerve repair with distal stump cell injection) a ELISA demonstrating SC production of GDNF at 3 and 11 weeks postsurgery. After receiving only 1 week of doxycycline postsurgery, the Dox+/– group demonstrated effective downregulation of GDNF production after 3 and 11 weeks as

compared to the constant administration group (Dox+/+). b The Dox+/– group also demonstrated improved neurofilament-positive regeneration (p < 0.05 over SC injection control), reflected in improved slip ratio measurements as seen after 6 and 9 weeks (c). * p < 0.05. Adapted from Shakhbazau et al. [2013].

planted ASCs mitigate DRG neuronal loss after injury, likely by preventing the activation of caspase-3 in a neurotrophin-dependent manner [Reid et al., 2011]. To promote regeneration after injury, the use of genetically modified donor cells with a robust growth factor elution profile is not a novel concept [Weidner et al., 1999; Hu et al., 2005]. Recently, Bunge [1994] transplanted transduced SCs capable of producing both D15A (a trkA/B-activating neurotrophin) and chondroitinase ABC into a thoracic spinal contusion injury, and found the combinatorial therapy was superior at integrating into the injury and also at improving sensorimotor hind limb function as compared to nonenhanced SC therapy [Kanno et al., 2014]. In the PNS, Shakhbazau et al. [2012a, b] have recently reinforced the importance of cell transplant as a source of neurotrophins through a series of manuscripts. While it has already been well established that NGF is upregulated after nerve injury [Meyer et al., 1992; Frostick et al., 1998], these experiments demonstrated the SCs,

engineered to overexpress NGF, could encourage improved early axonal regeneration across a short gap (5 mm) tube repair in a rat. The results suggest that endogenous levels of neurotrophins reached postinjury in the distal regenerative milieu may not be optimal for the maximum regenerative response, and that enhanced cell therapy could potentially improve on this deficit. One major concern with this approach, however, is the ‘candy-store’ effect [Eggers et al., 2008; Tannemaat et al., 2008]. This is a phenomena whereby local hypersecretion of a growth factor results in excessive axonal trapping and failure of distal regeneration. The effect coincides with having a point focus of growth factor secretion surrounding a transplanted cell graft, and can be profound. This may pertain especially to glial cell-derived neurotrophic factor (GDNF) oversecretion [Tannemaat et al., 2008]. To address the candy store issue, our lab has recently developed a doxycycline-inducible GDNF expression system for use in donor graft SCs [Shakhbazau et al., 2013]. These transplanted cells secrete significant

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amounts of GDNF in a tightly controlled, doxycyclinedependent manner. When transplanted in the distal regenerative environment, these cells can promote improved recovery most effectively when GDNF secretion is turned ‘on’ for a short period immediately postinjury [Shakhbazau et al., 2013] (fig. 5). Conversely, prolonged GDNF treatment does not result in significant improved recovery, and is in fact associated with increased axonal misdirection [Shakhbazau et al., 2013].

Conclusions

For nerve injury repair, recent advances in the field of SC replacement therapy have made one concept increasingly clear – the regenerative capacity of the SC can be improved upon. The pursuit of this goal may provide well-defined clinical benefits in the near future, and many groups are currently in pursuit of human applications for stem cell use for peripheral nerve regeneration [Tomita et al., 2013]. Considering the above, the ideal cell for this context in the future may be genetically customized to maximize regenerative potential, and will likely be of a stem-cell lineage.

References


differentiated bone-marrow stromal cells. Eur J Neurosci 14: 1771–1776. di Summa, P.G., D.F. Kalbermatten, E. Pralong, W. Raffoul, P.J. Kingham, G. Terenghi (2011) Long-term in vivo regeneration of peripheral nerves through bioengineered nerve grafts. Neuroscience 5: 278–291. di Summa, P.G., D.F. Kalbermatten, W. Raffoul, G. Terenghi, P.J. Kingham (2013) Extracellular matrix molecules enhance the neurotrophic effect of Schwann cell-like differentiated adipose-derived stem cells and increase cell survival under stress conditions. Tissue Eng 19(3–4): 368–379. di Summa, P.G., P.J. Kingham, W. Raffoul, M. Wiberg, G. Terenghi, D.F. Kalbermatten (2010) Adipose-derived stem cells enhance peripheral nerve regeneration. J Plast Reconstr Aesthet Surg 63: 1544–1552. Eggers, R., W.T. Hendriks, M.R. Tannemaat, J.J. van Heerikhuize, C.W. Pool, T.P. Carlstedt, A. Zaldumbide, R.C. Hoeben, G.J. Boer, J. Verhaagen (2008) Neuroregenerative effects of lentiviral vector-mediated GDNF expression in reimplanted ventral roots. Mol Cell Neurosci 39: 105–117. England, J.D., E.K. Rhee, G. Said, A.J. Sumner (1988) Schwann cell degeneration induced by doxorubicin (adriamycin). Brain 111: 901– 913. Frostick, S.P., Q. Yin, G.J. Kemp (1998) Schwann cells, neurotrophic factors, and peripheral nerve regeneration. Microsurgery 18: 397– 405. Fu, S.Y., T. Gordon (1997) The cellular and molecular basis of peripheral nerve regeneration. Mol Neurobiol 14: 67–116. Grochmal, J., S. Dhaliwal, P.K. Stys, J. van Minnen, R. Midha (2014) Skin derived precursor Schwann cell myelination capacity in focal tibial demyelination. Muscle Nerve 50: 262– 272. Guenard, V., N. Kleitman, T.K. Morrissey, R.P. Bunge, P. Aebischer (1992) Syngeneic Schwann cells derived from adult nerves seed-


ed in semipermeable guidance channels enhance peripheral nerve regeneration. J Neurosci 12: 3310–3320. Gulati, A.K., D.R. Rai, A.M. Ali (1995) The influence of cultured Schwann cells on regeneration through acellular basal lamina grafts. Brain Res 705: 118–124. Hill, C.E., A. Hurtado, B. Blits, B.A. Bahr, P.M. Wood, M.B. Bunge, M. Oudega (2007) Early necrosis and apoptosis of Schwann cells transplanted into the injured rat spinal cord. Eur J Neurosci 26: 1433–1445. Hu, Y., S.G. Leaver, G.W. Plant, W.T. Hendriks, S.P. Niclou, J. Verhaagen, A.R. Harvey, Q. Cui (2005) Lentiviral-mediated transfer of CNTF to Schwann cells within reconstructed peripheral nerve grafts enhances adult retinal ganglion cell survival and axonal regeneration. Mol Ther 11: 906–915. Ikeda, M., T. Uemura, K. Takamatsu, M. Okada, K. Kazuki, Y. Tabata, Y. Ikada, H. Nakamura (2013) Acceleration of peripheral nerve regeneration using nerve conduits in combination with induced pluripotent stem cell technology and a basic fibroblast growth factor drug delivery system. J Biomed Mater Res A 102: 1370–1378. Kanno, H., Y. Pressman, A. Moody, R. Berg, E.M. Muir, J.H. Rogers, H. Ozawa, E. Itoi, D.D. Pearse, M.B. Bunge (2014) Combination of engineered Schwann cell grafts to secrete neurotrophin and chondroitinase promotes axonal regeneration and locomotion after spinal cord injury. J Neurosci 34: 1838–1855. Khuong, H., R. Kumar, F. Senjaya, J. Grochmal, A. Ivanovic, A. Shakhbazau, J. Forden, A.A. Webb, J. Biernaskie, R. Midha (2014) Skin derived precursor Schwann cells improve behavioral recovery for acute and delayed nerve repair. Exp Neurol 254: 168–179. Kingham, P.J., D.F. Kalbermatten, D. Mahay, S.J. Armstrong, M. Wiberg, G. Terenghi (2007) Adipose-derived stem cells differentiate into a Schwann cell phenotype and promote neurite outgrowth in vitro. Exp Neurol 207: 267–274.

9


Akiyama, Y., C. Radtke, J.D. Kocsis (2002) Remyelination of the rat spinal cord by transplantation of identified bone marrow stromal cells. J Neurosci 22: 6623–6630. Azizi, S.A., D. Stokes, B.J. Augelli, C. Digirolamo, D.J. Prockop (1998) Engraftment and migration of human bone marrow stromal cells implanted in the brains of albino rats – similarities to astrocyte grafts. Proc Natl Acad Sci USA 95: 3908–3913. Barrette, B., M.A. Hebert, M. Filali, K. Lafortune, N. Vallieres, G. Gowing, J.P. Julien, C.L. Lacroix (2008) Requirement of myeloid cells for axonal regeneration. J Neurosci 28: 9363– 9376. Bisby, M.A., S. Chen (1990) Delayed Wallerian degeneration in sciatic nerve of C57BL/Ola mice is associated with impaired regeneration of sensory axons. Brain Res 530: 117–120. Brown, M.C., E.R. Lunn, V.H. Perry (1992) Consequences of slow Wallerian degeneration for regenerating motor and sensory axons. J Neurobiol 23: 521–536. Bunge, R.P. (1994) The role of the Schwann cell in trophic support and regeneration. J Neurol 242(suppl 1): S19–S21. Büngner, O.V. (1891) Über die Degenerationsund Regenerationsvorgänge an Nerven nach Verletzungen. Beitr Pathol Anat 10: 321–387. Chen, C.J., Y.C. Ou, S.L. Liao, W.Y. Chen, S.Y. Chen, C.W. Wu, C.C. Wang, W.Y. Wang, Y.S. Huang, S.H. Hsu (2007a) Transplantation of bone marrow stromal cells for peripheral nerve repair. Exp Neurol 204: 443–453. Chen, Z.L., W.M. Yu, S. Strickland (2007b) Peripheral regeneration. Ann Rev Neurosci 30: 209–233. Clemence, A., R. Mirsky, K.R. Jessen (1989) Non-myelin-forming Schwann cells proliferate rapidly during Wallerian degeneration in the rat sciatic nerve. J Neurocytol 18: 185– 192. Dezawa, M., I. Takahashi, M. Esaki, M. Takano, H. Sawada (2001) Sciatic nerve regeneration in rats induced by transplantation of in vitro

10


Barres, T. Omura, C.J. Woolf (2014) Diminished Schwann cell repair responses underlie age-associated impaired axonal regeneration. Neuron 83: 331–343. Pan, H.C., C.J. Chen, F.C. Cheng, S.P. Ho, M.J. Liu, S.M. Hwang, M.H. Chang, Y.C. Wang (2009) Combination of G-CSF administration and human amniotic fluid mesenchymal stem cell transplantation promotes peripheral nerve regeneration. Neurochem Res 34: 518– 527. Prockop, D.J. (2007) ‘Stemness’ does not explain the repair of many tissues by mesenchymal stem/multipotent stromal cells (MSCs). Clin Pharmacol Ther 82: 241–243. Reid, A.J., M. Sun, M. Wiberg, S. Downes, G. Terenghi, P.J. Kingham (2011) Nerve repair with adipose-derived stem cells protects dorsal root ganglia neurons from apoptosis. Neuroscience 199: 515–522. Shakhbazau, A., J. Kawasoe, S.A. Hoyng, R. Kumar, J. van Minnen, J. Verhaagen, R. Midha (2012a) Early regenerative effects of NGFtransduced Schwann cells in peripheral nerve repair. Mol Cell Neurosci 50: 103–112. Shakhbazau, A., J.A. Martinez, Q.G. Xu, J. Kawasoe, J. van Minnen, R. Midha (2012b) Evidence for a systemic regulation of neurotrophin synthesis in response to peripheral nerve injury. J Neurochem 2012: 501–511. Shakhbazau, A., C. Mohanty, D. Shcharbin, M. Bryszewska, A.M. Caminade, J.P. Majoral, J. Alant, R. Midha (2013) Doxycycline-regulated GDNF expression promotes axonal regeneration and functional recovery in transected peripheral nerve. J Control Release 172: 841– 851. Shimizu, S., M. Kitada, H. Ishikawa, Y. Itokazu, S. Wakao, M. Dezawa (2007) Peripheral nerve regeneration by the in vitro differentiated-human bone marrow stromal cells with Schwann cell property. Biochem Biophys Res Commun 359: 915–920. Sowa, Y., T. Imura, T. Numajiri, S. Fushiki (2012) Adipose-derived stem cells produce factors enhancing peripheral nerve regeneration: influence of age and anatomic site origin. Stem Cells Dev 21: 1852–1862. Tannemaat, M.R., R. Eggers, W.T. Hendriks, G.C. de Ruiter, J.J. van Heerikhuize, C.W. Pool, M.J. Malessy, G.J. Boer, J. Verhaagen (2008) Differential effects of lentiviral vector-mediated overexpression of nerve growth factor and glial cell line-derived neurotrophic factor on regenerating sensory and motor axons in the transected peripheral nerve. Eur J Neurosci 28: 1467–1479. Tofaris, G.K., P.H. Patterson, K.R. Jessen, R. Mirsky (2002) Denervated Schwann cells attract macrophages by secretion of leukemia inhibitory factor (LIF) and monocyte chemoattractant protein-1 in a process regulated by interleukin-6 and LIF. J Neurosci 22: 6696–6703.

Toma, J.G., M. Akhavan, K.J. Fernandes, F. Barnabe-Heider, A. Sadikot, D.R. Kaplan, F.D. Miller (2001) Isolation of multipotent adult stem cells from the dermis of mammalian skin. Nat Cell Biol 3: 778–784. Toma, J.G., I.A. McKenzie, D. Bagli, F.D. Miller (2005) Isolation and characterization of multipotent skin-derived precursors from human skin. Stem Cells 23: 727–737. Tomita, K., T. Madura, C. Mantovani, G. Terenghi (2012) Differentiated adipose-derived stem cells promote myelination and enhance functional recovery in a rat model of chronic denervation. J Neurosci Res 90: 1392–1402. Tomita, K., T. Madura, Y. Sakai, K. Yano, G. Terenghi, K. Hosokawa (2013) Glial differentiation of human adipose derived stem cells: implications for cell-based trasplantation therapy. Neuroscience 236: 55–65. Uemura, T., K. Takamatsu, M. Ikeda, M. Okada, K. Kazuki, Y. Ikada, H. Nakamura (2012) Transplantation of induced pluripotent stem cell-derived neurospheres for peripheral nerve repair. Biochem Biophys Res Commun 419: 130–135. Walsh, S., J. Biernaskie, S.W. Kemp, R. Midha (2009a) Supplementation of acellular nerve grafts with skin derived precursor cells promotes peripheral nerve regeneration. Neuroscience 164: 1097–1107. Walsh, S., R. Midha (2009a) Practical considerations concerning the use of stem cells for peripheral nerve repair. Neurosurg Focus 26: E2. Walsh, S., R. Midha (2009b) Use of stem cells to augment nerve injury repair. Neurosurgery 65(suppl): A80–A86. Walsh, S.K., T. Gordon, B.M. Addas, S.W. Kemp, R. Midha (2009b) Skin-derived precursor cells enhance peripheral nerve regeneration following chronic denervation. Exp Neurol 223: 221–228. Walsh, S.K., R. Kumar, J. Grochmal, S.W.P. Kemp, J. Forden, R. Midha (2011) Fate of stem cell transplants in peripheral nerves. Stem Cell Res 8: 226–238. Weidner, N., A. Blesch, R.J. Grill, M.H. Tuszynski (1999) Nerve growth factor-hypersecreting Schwann cell grafts augment and guide spinal cord axonal growth and remyelinate central nervous system axons in a phenotypically appropriate manner that correlates with expression of L1. J Comp Neurol 413: 495– 506. You, S., T. Petrov, P.H. Chung, T. Gordon (1997) The expression of the low affinity nerve growth factor receptor in long-term denervated Schwann cells. Glia 20: 87–100. Zhao, L., X. Wei, Z. Ma, D. Feng, P. Tu, B.H. Johnstone, K.L. March, Y. Du (2009) Adipose stromal cells-conditional medium protected glutamate-induced CGNs neuronal death by BDNF. Neurosci Lett 452: 238–240.

Grochmal/Midha


Liu, G., Y. Cheng, S. Guo, Y. Feng, Q. Li, H. Jia, Y. Wang, L. Tong, X. Tong (2011) Transplantation of adipose-derived stem cells for peripheral nerve repair. Int J Mol Med 28: 565–572. Lopatina, T., N. Kalinina, M. Karagyaur, D. Stambolsky, K. Rubina, A. Revischin, G. Pavlova, Y. Parfyonova, V. Tkachuk (2011) Adiposederived stem cells stimulate regeneration of peripheral nerves: BDNF secreted by these cells promotes nerve healing and axon growth de novo. PLoS One 6: e17889. Mantovani, C., D. Mahay, M. Kingham, G. Terenghi, S.G. Shawcross, M. Wiberg (2010) Bone marrow and adipose-derived stem cells show expression of myelin mRNAs and proteins. Regen Med 5: 403–410. Marcus, A.J., T.M. Coyne, I.B. Black, D. Woodbury (2008) Fate of amnion-derived stem cells transplanted to the fetal rat brain: migration, survival and differentiation. J Cell Mol Med 12: 1256–1264. McKenzie, I.A., J. Biernaskie, J.G. Toma, R. Midha, F.D. Miller (2006) Skin-derived precursors generate myelinating Schwann cells for the injured and dysmyelinated nervous system. J Neurosci 14: 6651–6660. Meyer, M., I. Matsuoka, C. Wetmore, L. Olson, H. Thoenen (1992) Enhanced synthesis of brainderived neurotrophic factor in the lesioned peripheral nerve: different mechanisms are responsible for the regulation of BDNF and NGF mRNA. J Cell Biol 119: 45–54. Mirsky, R., K.R. Jessen (1999) The neurobiology of Schwann cells. Brain Pathol 9: 293–311. Mizuno, H. (2009) Adipose-derived stem cells for tissue repair and regeneration: ten years of research and a literature review. J Nihon Med Sch 76: 56–66. Mosahebi, A., B. Woodward, M. Wiberg, R. Martin, G. Terenghi (2001) Retroviral labeling of Schwann cells: in vitro characterization and in vivo transplantation to improve peripheral nerve regeneration. Glia 34: 8–17. Murakami, T., Y. Fujimoto, Y. Yasunaga, O. Ishida, N. Tanaka, Y. Ikuta, M. Ochi (2003) Transplanted neuronal progenitor cells in a peripheral nerve gap promote nerve repair. Brain Res 974: 17–24. Nishiura, Y., J. Brandt, A. Nilsson, M. Kanje, L.B. Dahlin (2004) Addition of cultured Schwann cells to tendon autografts and freeze-thawed muscle grafts improves peripheral nerve regeneration. Tissue Eng 10: 157–164. Okada, Y., A. Matsumoto, T. Shimazaki, R. Enoki, A. Koizumi, S. Ishii, Y. Itoyama, G. Sobue, H. Okano (2008) Spatiotemporal recapitulation of central nervous system development by murine embryonic stem cell-derived neural stem/progenitor cells. Stem Cells 26: 3086– 3098. Painter, M.W., B.S. Lutz, Y. Cheng, A. Latremoliere, K. Duong, C. Miller, S. Posada, E.J. Cobos, A.X. Zhang, A.J. Wagers, L.A. Havton, B.

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