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Strategies to Promote Neural Repair and Regeneration After Spinal Cord Injury

Kwon, Brian K. MD, PhD, FRCSC; Fisher, Charles G. MD, MPH, FRCSC; Dvorak, Marcel F. MD, FRCSC; Tetzlaff, Wolfram MD, PhD

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doi: 10.1097/01.brs.0000175186.17923.87
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The permanence and irreversibility of the paralysis associated with spinal cord injury have been recognized and accepted for thousands of years,1 with hope emerging in only the latter part of the 20th century that advances in our scientific understanding of the neurobiology of spinal cord injury might produce therapies for this devastating condition.2 The sequelae of spinal cord paralysis are most profoundly manifested in the loss of voluntary motor, sensory, urologic, and sexual function, but significant abnormalities of the respiratory, cardiovascular, gastrointestinal, and musculoskeletal systems are common.3 In adult patients, mechanical failure of the spinal column in association with nonpenetrating spinal cord injury can lead to progressive kyphotic, and less commonly, scoliotic deformity.4 In skeletally immature patients, spinal cord injuries are associated with a high incidence of progressive deformity, with chronic spinal column instability, asymmetric growth arrest, and neuromuscular imbalance contributing to the paralytic collapse of the spine. The role of neuromuscular imbalance is demonstrated by the near 100% incidence of scoliosis in children paralyzed before the age of 10 years or before the adolescent growth spurt,5,6 a statistic that in itself represents the relevance of emerging therapies for spinal cord injury in the discussion of spinal deformity and its management.

Current Areas of Focus in Spinal Cord Research

The mechanical forces imparted to the spinal cord during fractures and dislocations of the spine disrupt the cord’s local neuroglial architecture (the “primary” damage) and initiate a complex pattern of acute pathophysiologic processes that are thought to exacerbate the derangement at and around the epicenter of injury (the “secondary” damage) (reviewed by Kwon et al7). It is well recognized that such nonpenetrating injuries to the spinal column rarely result in complete transection of the spinal cord. Even in individuals who are deemed to have functionally “complete” spinal cord injuries graded as “A” by the American Spinal Injury Association (ASIA), the chronically injured spinal cord remains anatomically in-continuity and is often characterized by a peripheral rim of intact tissue encircling a cystic cavity,8,9 a gross morphology that can be reasonably reproduced in animal models of a dorsal contusion spinal cord injury (Figure 1).

Figure 1
Figure 1:
Gross histology of the spinal cord after contusion injury. This figure depicts the damage to a rat spinal cord (Sprague-Dawley) 12 weeks after a dorsal contusion injury (Ohio State University Impactor, 1.2 mm injury) at the injury epicenter (C), 3 mm away (B), and 6 mm away (A) (myelin-stained with Luxol Fast Blue, counterstained with hematoxylin and eosin). Note how the injury epicenter (C) is characterized by a peripheral rim tissue encircling a cystic cavity, and that 3 mm away from this (B) the cord still remain grossly disrupted, with little gray-white matter distinction, before assuming a more normal appearance (A). Neuroprotective interventions are aimed at minimizing the extent of damage seen in B and C. The appearance of the injury epicenter at C represents the difficult environment that must be traversed by growing axons in order for functional reconnections to occur across the injury site.

The implications of this observation of the cord histomorphology are numerous. One is that if the entire spinal cord at the site of impact does not completely succumb to the initial mechanical injury and subsequent secondary pathophysiologic processes, then an intervention that attenuates these processes and maximizes the extent of this spared tissue could potentially result in improved neurologic function. Cautious optimism for such a strategy is derived from the observation in both humans9,10 and animals11–13 that some distal cord function can in some cases be mediated by surprisingly small amounts of spared spinal cord at the injury site, documented at less than 10% in some of these studies (Figure 2). Such is the rationale for neuroprotective strategies after acute injury, of which methylprednisolone is the most widely recognized, administered, and contentiously debated pharmacologic agent. Such pharmacologic agents will be further discussed.

Figure 2
Figure 2:
The MRI evolution of cord injury and sparing at injury site. This 48-year-old man was struck by a motor vehicle while riding his bicycle, suffering a C3–C4 fracture-dislocation and severe spinal cord injury requiring immediate intubation for respiratory failure. Note the severe edema and hemorrhage within his spinal cord on the T2-weighted MRI sagittal (A) and axial (B) views taken 18 hours after injury. Six weeks after posterior decompression and fusion, his imaging was repeated (C and D), having surprisingly regained Grade 4 out of 5 strength in his left quadriceps, ankle dorsiflexors, and plantarflexors. Note the peripheral sparing of spinal cord tissue at the injury site along the left side of the cord (arrow) and the cystic cavity that has already encompassed the remainder of his cord. The fact that this extent of spinal cord tissue can mediate near-normal strength in the lower extremity provides a compelling rationale for neuroprotective therapies that maximize this spared tissue.

The second implication is that, while some tissue may remain spared at the injury site, the spinal cord environment that is established over time, to even the untrained eye (Figure 1), appears extremely nonpermissive to axonal growth. This environment embodies a major challenge for neural regeneration, and as such is the focus of much neuroscientific study that aims to characterize the molecular obstacles and inhibitors to axonal regeneration and develop means of circumventing them. For example, the strategy of transplanting cellular substrates such as stem cells into the injured spinal cord aims to provide a more permissive and encouraging environment for axonal growth. Research into the challenges of axonal regeneration has identified not only external obstacles to axonal growth but also the low “intrinsic” regenerative competence of central nervous system (CNS) neurons that contributes to the failure of neural repair after spinal cord injury. These extrinsic and intrinsic impediments to axonal regeneration will be discussed further as well.

Finally, if some cord tissue is typically spared, even after the secondary damage induced by the pathophysiologic processes initiated acutely after injury, then a substrate exists for strategies that might enhance this remaining tissue’s ability to mediate distal function. For example, axons that traverse the injury site but have been stripped of their myelin sheath are the target of potassium channel blocking agents such as fampridine (Acorda Therapeutics, Hawthorne, NY) which, by altering the voltage potential across the axonal membrane, facilitates impulse conduction. Transplantation of cells that could remyelinate axons also has the potential to improve signal transmission across the injury site. Furthermore, if these precious few axons that survive and traverse the injury site could be induced to extend collateral sprouts that might reinnervate distal or even local neural circuitry, some functional recovery could be achieved. Such “plasticity” is postulated to be one of the principal effects of physical rehabilitation. It may indeed also be chiefly responsible for the functional recovery that is observed in animal models with experimental axonal regeneration therapies (although it is, of course, hoped that such therapies would produce long-distance axonal regeneration from axons that are interrupted at the injury site).

Within this framework of neural repair challenges, it is important that realistic expectations are maintained in the research and clinical communities vis-à-vis the achievability of a “cure” for paralysis. Clearly, we are at an exciting time for repair strategies, as a number of international clinical trials have been initiated; however, even the most promising axonal regeneration and neuroprotective strategies alone or in combination are almost certainly not going to restore a complete quadriplegic individual to normal physical function. Nevertheless, the process must begin somewhere, and even short distances of axonal regrowth or small victories in neuroprotection have the potential to make an enormous impact on the quality of life for patients with spinal cord injury. With this in mind, we first discuss the acute pathophysiologic responses to spinal cord injury and neuroprotective strategies, past, present, and future, that act to attenuate them. We then discuss the obstacles to axonal regeneration after spinal cord injury and how these may be overcome with therapies such as neurotrophic factors and cellular transplantation.

Pathophysiologic Processes That Contribute to Secondary Injury

Laboratory research in spinal cord injury uses a large number of animal models in which the spinal cord is subjected to either a blunt or sharp injury. Examples of the former include contusion injuries induced by a dorsally applied force (weight drop14–16 or impactor17,18), clip compression,19,20 and more recently, lateral spinal dislocation.21 Because of their conceptual resemblance to human nonpenetrating injuries, these models are most useful for studying pathophysiologic processes and neuroprotective strategies after spinal cord injury.22

These models have depicted an increasingly complex array of secondary pathophysiologic processes that are rapidly initiated after the mechanical injury to the spinal cord. Intraparenchymal hemorrhage, disruption of the blood–brain barrier, thrombosis, vasospasm, and the loss of pressure autoregulation are vascular aberrations that are intimately linked to the development of local ischemia, a fundamental component of secondary damage in the injury penumbra.23 Concomitantly, an influx of inflammatory cells, including neutrophils and macrophages, the activation of microglia, and the expression of cytokines such as IL-1β, TNFα, IL-6, LIF, and a myriad of matrix metalloproteinases contribute to the secondary injury process.24–28 Free radical generation and lipid peroxidation can propagate the local destruction of cellular membranes, and the oxidative stress can fatally disrupt intracellular molecular and biochemical homeostasis of neurons and glial cells.29,30 Such cells are also subjected to excitotoxicity caused by the extracellular release and accumulation of glutamate and the activation of N-methyl-D-aspartate (NMDA) and non-NMDA receptors.31 These components of the secondary injury cascade, i.e., microvascular ischemia, inflammation, oxidative stress, and excitotoxicity, are not mutually independent but rather can synergistically act to promote further destruction through the necrotic and apoptotic demise of neurons and glial cells. They are mentioned only briefly here, as all have been recognized not only as important components of the secondary injury response, but also potential targets for intervention.

Neuroprotective Interventions

The concept of neuroprotection is based on the premise that attenuating these aforementioned pathophysiologic processes will improve the ultimate neurologic outcome. A substantial amount of data from animal studies supports this concept, but effective neuroprotection in human spinal cord injury has been exceedingly difficult to demonstrate. Improvements over the past 40 years in protocols for trauma stabilization, immobilization, and resuscitation are thought to be neuroprotective insofar as the proportion of complete spinal cord injuries appears to have decreased with a concomitant increase in the proportion of incomplete injuries over that period of time.32 Early surgical decompression has for many years been postulated to improve neurologic outcome after traumatic spinal cord injury, although human data to this point have been contradictory and a large-scale prospective multicenter trial to demonstrate the effect of early decompression has only recently begun. Data from this trial, entitled the “Surgical Treatment of Acute Spinal Cord Injury Trial” (STASCIS), are as yet unavailable but will hopefully provide insight on this important question.

Methylprednisolone and Other Previously Tested Pharmacologic Agents

The publishing of the second National Acute Spinal Cord Injury Study (NASCIS 2)33 in 1990 was heralded as the first convincing demonstration of a pharmacologic neuroprotection in humans, but more recent critiques34–36 of the execution and interpretation of this and the subsequent NASCIS 337 have led many prominent orthopedic, neurosurgical, and emergency medicine societies to speak out against methylprednisolone as a standard of care for individuals with acute spinal cord injuries.38,39 Additional pharmacologic agents that have been trialed in human spinal cord injury but have not demonstrated significant clinical efficacy include GM-1 ganglioside (Sygen),40 naloxone,33 thyrotropin releasing hormone,41 nimodipine,42 tirilazad mesylate,37 and gacyclidine (as yet unpublished negative results of a 280-patient prospective randomized trial43). The failure to demonstrate convincing efficacy in these human spinal cord injury trials stands in resounding contrast to the very promising results reported for each of these agents in preclinical animal studies (and for GM-1 ganglioside, even a small clinical study44). This fact serves as a sobering reminder of the important differences that exist between the human and animal condition, and should temper the enthusiasm for making strong predictions about the efficacy of neuroprotective agents based on animal results alone.

The debate over the merits of methylprednisolone has certainly pointed out the need for the development of more effective neurotherapeutic agents. While many compounds are evaluated in the laboratory setting, of particular interest to clinicians are those agents that are currently in widespread use for other human conditions that, almost surreptitiously, have been found to have neuroprotective properties. Human experience with such agents would therefore overcome many important safety and tolerance issues that would be understandably enormous for a trial on, for example, viral-mediated gene therapy. Leading the list of such agents are the antibiotic minocycline and the hormone erythropoietin.


Minocycline is a tetracycline derivative that is currently in common clinical use for the treatment of acne and chronic periodontitis. In addition to its broad spectrum of antibacterial activity, minocycline has been found to have a number of neuroprotective properties,45 including the inhibition of matrix metalloproteinases,46 the inhibition of microglial activation47–49 (both considered to be important aspects of neuroinflammation), and the prevention of programmed cell death.50–53 These neuroprotective properties have been demonstrated in promising results from animal models of ischemic stroke,54,55 Parkinson’s disease,56,57 Huntington disease,58,59 and amyotrophic lateral sclerosis.60–62 Such promising results have prompted the initiation of minocycline clinical trials for patients with Parkinson’s disease,63 Huntington disease,64 and amyotrophic lateral sclerosis.65 One should be aware, however, that not all studies of minocycline in this regard have been positive, and that negative or even deleterious effects have also been reported in animal studies of these neurologic disorders,66–69 illustrating the need for caution in the development of minocycline as a neuroprotective agent.70

Despite these conflicting reports, a number of laboratories have independently reported beneficial effects of minocycline in acute spinal cord injury. The first demonstration of this came from Wells et al in 2003, who observed that minocycline administration 1 hour after a spinal cord compression injury resulted in increased axonal sparing at the injury,71 results similar to those found by Lee et al after a contusion spinal cord injury.72 Stirling et al reported that minocycline treatment after a dorsal column transection reduced the apoptotic demise of oligodendrocytes and the activation of microglia/macrophages, and diminished axonal dieback from the injury site.53 The reduction in oligodendrocyte apoptosis was also reported by Teng et al, who demonstrated that minocycline inhibits cytochrome c release at the injury site.73 Importantly, all four of these studies reported that minocycline administration was associated with improved behavioral outcomes on locomotor and other functional testing. It is worth pointing out that the reproducibility of the minocycline effect by independent laboratories using different animal models of spinal cord injury, different dosing regimens, and different outcome measures is rather unique in the neuroscientific study of therapeutic agents, and lends credence to the potential efficacy of minocycline in this context. A pilot study to evaluate minocycline in patients with acute spinal cord injuries has been initiated in Calgary, Alberta, Canada.


Erythropoietin, a hormone produced mainly by the kidney in response to hypoxia, has long been recognized for its role in regulating erythropoiesis and is currently in widespread clinical use for the treatment of anemia related to such scenarios as chemotherapy, chronic renal failure, and autologous blood donation. It has also been subsequently discovered to have tissue-protective properties in animal models of stroke,74,75 multiple sclerosis,76 Parkinson’s disease,77 myocardial infarction,78,79 and spinal cord injury. In an ischemia model of spinal cord injury (induced by aortic occlusion), erythropoietin was found to prevent motoneuron apoptosis and to promote functional recovery.80 Neuroprotection after traumatic spinal cord injury was first demonstrated by Gorio et al, who found that after either a clip compression or a contusion injury to the rat spinal cord, the administration of erythropoietin immediately after injury promoted improved locomotor scores and tissue sparing at the injury site.81 Kaptanoglu et al subsequently demonstrated a dose-dependent reduction in lipid peroxidation and improved histologic appearance of the injury site with erythropoietin after contusive spinal cord injury.82 It is of interest that, while the inhibition of lipid peroxidation has been proposed as the major mechanism of action for methylprednisolone after spinal cord injury, in this study, the reduction of lipid peroxidation was significantly better in animals treated with high-dose erythropoietin than with the NASCIS 2 bolus dose of methylprednisolone (30 mg/kg).

A potential problem with the use of erythropoietin as a neuroprotective agent is the possibility that the stimulation of erythropoiesis (particularly in a nonanemic patient) could lead to an undesirable erythrocytosis and a prothrombotic state. In practical terms, this may not be a substantial problem for a short-term application of erythropoietin (such as acutely after spinal cord injury). Within this context, however, it has been recently demonstrated that erythropoietin may be modified in such a manner as to relinquish its erythropoietic properties but yet maintain its tissue-protective characteristics.83 Further study has demonstrated that the tissue-protective properties of erythropoietin may be mediated by different receptor interactions and intracellular signaling pathways than the erythropoietic properties.84 Most promisingly, a small prospective randomized double-blinded study of human patients with acute strokes revealed that a once daily injection of erythropoietin for 3 days was well tolerated and led to improved neurologic function and infarct size at an early time point.85 These animal and human results would strongly suggest that a clinical trial for spinal cord injuries is warranted.

Axonal Regeneration Therapies

The poor axonal regeneration and functional recovery after blunt spinal cord injury stand in stark contrast to the fairly robust axonal regeneration and functional recovery observed in even transected peripheral nerves after primary repair. The biologic factors that account for these differences are the subject of much research. In very simple conceptual terms, the success of neurons within the peripheral nervous system to regenerate is influenced by their ability to upregulate genetic programs necessary for axonal growth, and by the permissive growth environment provided by the Schwann cells within the injured peripheral nerve. Conversely, neurons within the CNS often fail to upregulate or sustain the expression of such genes as a reflection of their “intrinsic” growth incompetence, and must extend new axons into the inhibitory CNS environment.86 Therapies to promote axonal regeneration within the CNS therefore focus on either augmenting the neuron’s intrinsic ability to regenerate, or attenuating the inhibitory CNS environment into which they must regrow.

Augmenting Intrinsic Growth Propensity With Neurotrophic Factors

A commonly used approach for stimulating axonal growth within the injured CNS is the administration of neurotrophic factors. Neurotrophic factors are proteins that exert considerable influence on a wide spectrum of processes within the developing and mature nervous system, including neuronal survival, axonal growth, synaptic plasticity, and neurotransmission (reviewed by Tuszynski87) Since the identification of the first neurotrophic factor, nerve growth factor (NGF) by Levi-Montalcini and Hamburger over 5 decades ago,88,89 dozens of such factors have been uncovered. It is important to recognize that the extreme diversity of neurotrophic factors and their functions, in addition to the complex cytoarchitecture of the spinal cord, make it unrealistic to expect that the administration of a single trophic factor will, by itself, elicit the comprehensive regenerative response in all relevant neuronal and glial cell populations necessary for full recovery after spinal cord injury. While the induction of ectopic bone formation to facilitate spinal fusion appears to be achievable with the exogenous administration of single growth factors of the bone morphogenetic protein family (e.g., the commercially available forms of BMP-2 or BMP-7), the situation within the injured spinal cord is clearly more complex.

Like bone morphogenetic proteins, however, the biologic activity of neurotrophic factors depends on the target tissue possessing the appropriate receptors, and the presence of such receptors is an important consideration for the therapeutic application of exogenous growth factors. This has been well demonstrated in the rat rubrospinal system (a motor control system) after acute and chronic spinal cord injury. Following a cervical spinal cord injury in which the rubrospinal tract is cut, the immediate administration of brain derived neurotrophic factor (BDNF) into the spinal cord injury site promotes significant rubrospinal axonal regeneration and prevents axotomy-induced atrophy and/or death of rubrospinal neurons.90,91 This indicates that the acutely injured rubrospinal axons at the spinal cord injury site are responsive to BDNF and thereby conduct the appropriate intracellular signaling pathways to augment the intrinsic growth propensity of these CNS neurons. However, such a regenerative response is not elicited in the rubrospinal neurons if the administration of BDNF into the injury site is delayed by 6 to 8 weeks after injury.92,93 This loss of effectiveness in the “chronic” setting can be explained by the observation that the injured rubrospinal axons at the spinal cord injury site lose, over time, the full-length trkB receptors necessary to mediate BDNF activity (such receptors are present on uninjured rubrospinal axons and are thus presumed to mediate the effect of BDNF applied immediately after injury).94 Conversely, BDNF applied directly to the rubrospinal cell bodies within the brainstem either acutely95 or even 1 year after injury96 elicits a reversal of axotomy-induced neuronal atrophy, an upregulation of genes important for regeneration, and the promotion of rubrospinal axonal growth. Consistent with this, full-length trkB receptors are maintained on the cell bodies of rubrospinal neurons even 1 year after cervical injury.96 While this is but one example of one neurotrophic factor (BDNF) and its receptor (trkB) within one neuronal system (rubrospinal system), it illustrates the principle that the development of effective therapeutic strategies that use the administration of neurotrophic factors will require an understanding of the biologic responsiveness of the target tissue, responsiveness that may in fact change with time and thus differ between the acute and chronic injury settings.

Augmenting Intrinsic Growth Propensity by Elevating Intracellular Cyclic AMP

With the demonstrated ability of neurotrophic factors to augment the growth potential of CNS neurons after injury, it is obviously of great interest to elucidate the intracellular mechanisms that mediate this effect. One mechanism that has stimulated substantial interest over the last 5 years is the elevation of intracellular cyclic adenosine monophosphate (cAMP). Neurite outgrowth from cerebellar neurons is typically inhibited when cultured in vitro on nonpermissive substrates such as CNS myelin or myelin-associated glycoprotein (MAG). Filbin et al demonstrated, however, that this inhibition could be overcome and neurite growth could be facilitated by “priming” such neurons with an overnight exposure to the neurotrophic factors BDNF or glial-derived neurotrophic factor before plating them on the nonpermissive substrates.97 This promotion of the neuron’s ability to overcome the nonpermissive environment was mediated by a rise in the intracellular cAMP levels and could be reproduced by the exogenous application of dibutyryl cAMP (db cAMP), a cAMP analogue.

Subsequent in vivo experiments revealed that the injection of db cAMP into the dorsal root ganglions promoted the regeneration of the centrally projecting axons within the spinal cord after a dorsal column spinal cord injury, again demonstrating that the elevated cAMP levels within the neurons allowed them to grow axons over even long distances within the inhibitory CNS environment.98,99 Three recent in vivo studies in which the administration of cAMP (or alternatively, the inhibition of its hydrolytic breakdown) was combined with the of Schwann cells,100 bone marrow stromal cells,101 or embryonic spinal tissue102 into the injured spinal cord have demonstrated that the elevation of cAMP is an important determinant of the neuronal growth propensity and may facilitate both axonal regeneration and possibly even functional recovery after spinal cord injury. These studies demonstrate, in principle, the potential efficacy of using a “combinatorial” therapeutic strategy of integrating different treatment paradigms. An example of this would be elevating intracellular cAMP to augment the growth ability of the neuron, plus cell transplantation to provide a better growth environment for regenerating axons. It is widely thought that effective therapies for spinal cord injury in the future will require a combinatorial approach such as this.

Nonpermissive Environment of the Injured Spinal Cord: Myelin and the Glial Scar

As alluded to previously, the other conceptual obstacle to axonal regeneration after spinal cord injury is the inhibitory environment into which new axons must grow. A tremendous amount of research over the past two decades has gone into identifying the inhibitory CNS molecules that make the spinal cord less permissive to axonal regeneration than peripheral nerves. The two major impediments to axonal regeneration in the injured spinal cord are CNS myelin and the glial scar that is established at the injury site.

The full complement of inhibitory elements that reside within CNS myelin and obstruct axonal regeneration is currently unknown. The three best characterized inhibitory constituents include the protein called Nogo,101,103–105 MAG,106 and oligodendrocyte-myelin glycoprotein (OMgp)107 (reviewed by Grados-Munro and Fournier108). The discovery of the receptor for Nogo in 2001109 was followed by the somewhat surprising finding that this receptor also interacts with MAG and OMgp to mediate some of their inhibitory properties as well.110 This has prompted much interest in the Nogo receptor and its subsequent intracellular signaling as a potential target for therapeutic interventions, interventions that, by altering this single pathway, might attenuate the axonal growth-repulsive properties of multiple inhibitory molecules (i.e., Nogo, MAG, and OMgp).111 As proof of this concept, Strittmatter et al demonstrated that the intrathecal administration of a portion of the Nogo receptor that competitively binds to and neutralizes Nogo, MAG, and OMgp promoted axonal sprouting and functional recovery after a partial spinal cord injury in rats.112 It is worth noting, however, that despite increasing interest in Nogo and mounting enthusiasm for developing human therapies that target this molecule, studies on Nogo-deficient transgenic mice performed by three independent laboratories have come to contradictory conclusions regarding the in vivo importance of Nogo as an inhibitor of axonal growth.113–115 Furthermore, a very recent study of transgenic mice lacking the Nogo receptor demonstrated that neurite outgrowth of certain populations of cells was not enhanced, as one would expect in the absence of this receptor.116 Clearly, these contradictory observations emphasize the complexity of this biologic system and provide a compelling rationale for further investigation to better delineate the role of Nogo in spinal cord injuries before human translation.

The glial scar and cyst that become established at the injury site over time are thought to be the other major impediments to axonal regeneration after spinal cord injury. Some evidence exists to suggest that in the absence of glial scarring within the CNS, axonal regeneration can occur within CNS myelin,117 a finding that stirs some controversy as to whether the glial scar or CNS myelin is more responsible for inhibiting axonal regeneration after spinal cord injury (and is therefore the most attractive target for intervention). The predominant cellular constituent of the glial scar are astrocytes, which, in addition to ultimately forming a physical barrier to growing axons, also express a number of chondroitin sulfate proteoglycans (CSPGs), which are inhibitory to axonal growth (reviewed by Silver and Miller118).

A potentially promising therapeutic intervention to address the CSPG inhibition to axonal growth is to enzymatically degrade the proteoglycans, and in this regard, the enzyme chondroitinase ABC has received increasing attention. This enzyme removes a large proportion of the glycosaminoglycan chains from CSPGs, and the remaining protein core does not possess the same inhibitory properties as the intact macromolecule. The intrathecal infusion of chondroitinase ABC after a partial spinal cord lesion was reported to promote growth of corticospinal axons and result in functional recovery.119 The application of this strategy with cell transplantation therapies in order to attenuate the glial scar establishment at the host-transplant interface and thus facilitate the passage of axons through the graft and back into the spinal cord has also been successfully demonstrated.120

Cellular Transplantation

On observing the cystic cavity that often typifies the chronic spinal cord injury site, it is only intuitive to postulate that axonal regeneration could be facilitated by surgically filling that cavity with some form of growth-permissive cellular substrate. In this regard, a wide variety of cells have been investigated as therapeutic candidates for spinal cord injury, some of which have more recently been trialed in human patients. Primary candidates for such transplantation strategies include stem cells, bone marrow stromal cells, fetal tissue, Schwann cells, and olfactory ensheathing cells. It is worth noting that not all cell transplantation strategies are necessarily aimed at achieving the same thing. The undifferentiated nature of stem cells, for example, makes it possible for them to mature down neuronal or glial lineages, raising the potential for them to either form neurons to relay synaptic information across the injury, or to form glial cells that might, for example, remyelinate growing axons. Alternatively, the transplant may serve mainly to provide a favorable growth substrate for new axons and to subsequently remyelinate them to restore efficient conduction of action potentials (e.g., Schwann cells and olfactory ensheathing cells). In addition to their native properties, cells may be genetically modified ex vivo in order to confer specific properties that might be beneficial when transplanted, such as the augmented expression of neurotrophic factors.121 Because of recent developments and human experience with cell transplantation for spinal cord injury, this review will focus on only two of the many cellular substrates: Schwann cells and olfactory ensheathing cells.

Schwann Cells and the PNS-CNS Barrier

Schwann cells have long been recognized as the key cellular constituent of peripheral nerves that facilitates axonal regeneration. Indeed, seminal studies in the field of spinal cord research used this permissive growth environment, either as free-ending peripheral nerve transplants122 or as intercalary grafts bridging a completely transected spinal cord injury,123 to demonstrate axonal regeneration of CNS neurons after spinal cord injury. Further studies have shown that, in principle, this facilitation of axonal growth can be augmented by administering neurotrophic factors124 or by genetically manipulating Schwann cells to increase their neurotrophic factor production,125–127 again reflecting the potential efficacy of combinatorial therapeutic strategies.

A case report of the utilization of peripheral nerve transplantation for human chronic spinal cord injury was recently published by Cheng et al.128 The lead author, well known for his work with peripheral nerve transplants in acute123 and chronic129 rat spinal cord injury, used a similar approach by taking grafts from intercostal nerves and transplanting them into the spinal cord injury site of a 24-year-old man who, at the age of 20 years, had had a left-sided spinal cord hemisection in a stabbing incident. The authors reported an improvement in ASIA motor scores and motor-evoked potentials 2 years post-transplantation in this individual, although it is difficult to know how much of the recovery was attributable to the transplant versus the detethering and cyst decompression of his cord. A similar study in which autologous sural nerves segments have been used to bridge the spinal cord injury of 8 patients with complete spinal cord injuries from gunshot wounds has been performed by Dr. Tarcisio Barros at the University of Sao Paulo. Five-year follow-up results have suggested no improvement in neurologic function or electrophysiologic parameters.130

While many potential explanations exist for the limited success with peripheral nerve transplantation in these human reports, much animal literature has demonstrated that CNS axons that regenerate into such grafts are typically reluctant to leave the permissive PNS conditions and reenter the host, inhibitory CNS environment.131 This “PNS-CNS barrier” is likely related to the establishment of the glial scar and the presence of inhibitory myelin elements (discussed above) at the graft-host interface. Consistent with this, a recent study that combined Schwann cell transplants with chondroitinase ABC to attenuate this gliotic scar demonstrated that axons growing through the graft could indeed regenerate farther into the host CNS environment.120 This notwithstanding, the PNS-CNS barrier that inherently encumbers peripheral nerve and Schwann cell transplants has directed a great deal of attention toward a cell that appears capable of accompanying regenerating axons across this obstacle: the olfactory ensheathing cell.

Olfactory Ensheathing Cells

Sensory neurons within the olfactory epithelium (considered to be in the peripheral nervous system) are unique in that throughout adult life, they undergo a constant turnover and thus regenerate new axons that must extend through the cribriform plate and reconnect with second-order neurons in the olfactory bulb (considered to be in the inhibitory CNS). Olfactory ensheathing cells (OECs) are distinct glial cells that escort these growing axons across this PNS-CNS interface, and this property was identified as a potential solution for the problem encountered with Schwann cell and peripheral nerve transplants. The possibility that such cells could be harvested from an individual with a spinal cord injury, cultured in large quantities, and then transplanted into the same individual without risk of immunologic rejection gives them similar conceptual appeal as Schwann cells.

An important paper by Ramon-Cueto et al in 2000 accelerated international interest in what was already an expanding line of research into the therapeutic properties of OECs for CNS repair.132 These investigators reported that the transplantation of OECs into a complete spinal cord transection facilitated long-distance regeneration of corticospinal, noradrenergic, and serotonergic fibers across the lesion and into the distal spinal cord, observations that were associated with significant functional recovery. A large body of literature now exists on the use of OECs in spinal cord injury (reviewed by Barnett and Riddell133). While much of it is promising, the work from different laboratories using different techniques of cell acquisition, expansion, purification, and implantation into different animal models at different time points after injury has left many important questions to be answered.

Nonetheless, the human application of putative OECs has already begun in a number of centers around the world. These include Beijing, China, Lisbon, Portugal, and Brisbane, Australia, under the direction of Drs. Huang, Lima, and Mackay-Sim, respectively. For a more complete discussion of these and other clinical trials for spinal cord injuries, please see Steeves et al.130 It is important to note that only the project of Dr. Mackay-Sim in Australia has been designed as an epidemiologically sound Phase I clinical trial to evaluate the safety of purified OEC transplantation with four OEC and four placebo-treated patients and regularly scheduled assessments with blinded evaluators. Dr. Huang’s approach employs olfactory cells obtained from human fetuses aborted at 12 to 16 weeks’ gestation and, given the lack of strict inclusion criteria, placebo controls, nonblinded assessments, and standardized follow-up evaluations is more akin to an experimental treatment than a clinical trial. By February 2004, he had performed this transplantation on more than 300 patients with spinal cord injuries, and throughout the year has received increasing worldwide attention. Follow-up of these patients, and in particular, a reporting of safety and adverse events, has not been systematically and comprehensively documented. Dr. Lima’s trial in Portugal evaluated autologous transplants of olfactory mucosa in motor complete patients (ASIA A or B) and again lacks placebo controls and blinded assessment. Formal peer-reviewed reports of these human trials are anxiously awaited.


An accelerating pace of scientific discovery over the past three decades that has generated much hope for patients, scientists, and clinicians that a “cure for paralysis” will someday become a reality. While laboratory research in animal models of spinal cord injury has generated a number of promising experimental therapies, it has also revealed a daunting complexity of the neurobiologic challenges that impede neural repair after injury. A greater understanding of these obstacles will be necessary for the further development of therapeutic strategies. This review describes many encouraging reports of novel neuroprotective agents such as minocycline and erythropoietin, and a number of promising strategies for promoting axonal regeneration. These serve as the foundation for the cautious optimism that exists for the future, that in our generation we will see the emergence of truly effective therapies for patients with spinal cord injuries. The initiation of human trials for experimental therapies that have been extensively studied in the laboratory setting represents the forward movement of this knowledge but also speaks loudly for the need for these technologies to now be subject to epidemiologically sound and rigorous clinical evaluation. As the list of therapies available for human testing grows, equipoise will be necessary in this field of “spinal cord regeneration” that has traditionally been viewed as the realm of laboratory researchers but now demands leadership from clinicians with interest and expertise in clinical research methodology.

Key Points

  • An increased understanding of the acute pathophysiologic processes initiated after spinal cord injury is essential to the development of neuroprotective therapies.
  • While methylprednisolone remains a treatment option for acute spinal cord injury, a number of promising neuroprotective agents are being developed, many of which are already in clinical use for other indications.
  • Axonal regeneration of neurons within the central nervous system is limited by both their limited intrinsic capacity to regenerate and the inhibitory environment within the injured spinal cord.
  • Many experimental therapies are emerging from the laboratory as potential candidates for pilot clinical trials; spinal physicians will have an increasingly important leadership role in the design and execution of such trials.


1.Hughes JT. The Edwin Smith Surgical Papyrus: an analysis of the first case reports of spinal cord injuries. Paraplegia 1988;26:71–82.
2.Adams M, Cavanagh JF. International Campaign for Cures of Spinal Cord Injury Paralysis (ICCP): another step forward for spinal cord injury research. Spinal Cord 2004;42:273–80.
3.Sie I, Waters RL. Outcomes following spinal cord injury. In: Lin VW, ed. Spinal Cord Medicine. New York: Demos, 2003:87–103.
4.Vaccaro AR, Silber JS. Post-traumatic spinal deformity. Spine 2001;26(suppl):111–8.
5.Lancourt JE, Dickson JH, Carter RE. Paralytic spinal deformity following traumatic spinal-cord injury in children and adolescents. J Bone Joint Surg Am 1981;63:47–53.
6.Dearolf WW III, Betz RR, Vogel LC, et al. Scoliosis in pediatric spinal cord-injured patients. J Pediatr Orthop 1990;10:214–8.
7.Kwon BK, Tetzlaff W, Grauer JN, et al. Pathophysiology and pharmacologic treatment of acute spinal cord injury. Spine J 2004;4:451–64.
8.Bunge RP, Puckett WR, Becerra JL, et al. Observations on the pathology of human spinal cord injury: a review and classification of 22 new cases with details from a case of chronic cord compression with extensive focal demyelination. Adv Neurol 1993;59:75–89.
9.Kakulas BA. The applied neuropathology of human spinal cord injury. Spinal Cord 1999;37:79–88.
10.Kaelan C, Jacobsen P, Morling P, et al. A quantitative study of motoneurons and cortico-spinal fibers related to function in human spinal cord injury (SCI). Paraplegia 1989;27:153.
11.Blight AR. Cellular morphology of chronic spinal cord injury in the cat: analysis of myelinated axons by line-sampling. Neuroscience 1983;10:521–43.
12.Eidelberg E, Straehley D, Erspamer R, et al. Relationship between residual hindlimb-assisted locomotion and surviving axons after incomplete spinal cord injuries. Exp Neurol 1977;56:312–22.
13.Fehlings MG, Tator CH. The relationships among the severity of spinal cord injury, residual neurological function, axon counts, and counts of retrogradely labeled neurons after experimental spinal cord injury. Exp Neurol 1995;132:220–8.
14.Noble LJ, Wrathall JR. Spinal cord contusion in the rat: morphometric analyses of alterations in the spinal cord. Exp Neurol 1985;88:135–49.
15.Gruner JA. A monitored contusion model of spinal cord injury in the rat. J Neurotrauma 1992;9:123–6.
16.Basso DM, Beattie MS, Bresnahan JC. Graded histological and locomotor outcomes after spinal cord contusion using the NYU weight-drop device versus transection. Exp Neurol 1996;139:244–56.
17.Noyes DH. Electromechanical impactor for producing experimental spinal cord injury in animals. Med Biol Eng Comput 1987;25:335–40.
18.Stokes BT. Experimental spinal cord injury: a dynamic and verifiable injury device. J Neurotrauma 1992;9:129–31.
19.Rivlin AS, Tator CH. Effect of duration of acute spinal cord compression in a new acute cord injury model in the rat. Surg Neurol 1978;10:38–43.
20.Joshi M, Fehlings MG. Development and characterization of a novel, graded model of clip compressive spinal cord injury in the mouse: Part 1. Clip design, behavioral outcomes, and histopathology. J Neurotrauma 2002;19:175–90.
21.Fiford RJ, Bilston LE, Waite P, et al. A vertebral dislocation model of spinal cord injury in rats. J Neurotrauma 2004;21:451–8.
22.Kwon BK, Oxland TR, Tetzlaff W. Animal models used in spinal cord regeneration research. Spine 2002;27:1504–10.
23.Amar AP, Levy ML. Pathogenesis and pharmacological strategies for mitigating secondary damage in acute spinal cord injury. Neurosurgery 1999;44:1027–39.
24.Popovich PG, Wei P, Stokes BT. Cellular inflammatory response after spinal cord injury in Sprague-Dawley and Lewis rats. J Comp Neurol 1997;377:443–64.
25.Dusart I, Schwab ME. Secondary cell death and the inflammatory reaction after dorsal hemisection of the rat spinal cord. Eur J Neurosci 1994;6: 712–24.
26.Bartholdi D, Schwab ME. Expression of pro-inflammatory cytokine and chemokine mRNA upon experimental spinal cord injury in mouse: an in situ hybridization study. Eur J Neurosci 1997;9:1422–38.
27.Klusman I, Schwab ME. Effects of pro-inflammatory cytokines in experimental spinal cord injury. Brain Res 1997;762:173–84.
28.Kerr BJ, Patterson PH. Potent pro-inflammatory actions of leukemia inhibitory factor in the spinal cord of the adult mouse. Exp Neurol 2004;188:391–407.
29.Hall E. Free radicals in central nervous system injury. In: Rice-Evans CA, Burdon R, eds. Free Radical Damage and Its Control. New York: Elsevier Science, 1994:217–38.
30.Cuzzocrea S, Riley DP, Caputi AP, et al. Antioxidant therapy: a new pharmacological approach in shock, inflammation, and ischemia/reperfusion injury. Pharmacol Rev 2001;53:135–59.
31.Wrathall JR, Teng YD, Choiniere D. Amelioration of functional deficits from spinal cord trauma with systemically administered NBQX, an antagonist of non-N-methyl-D-aspartate receptors. Exp Neurol 1996;137:119–26.
32.Tator CH, Duncan EG, Edmonds VE, et al. Changes in epidemiology of acute spinal cord injury from 1947 to 1981. Surg Neurol 1993;40:207–15.
33.Bracken MB, Shepard MJ, Collins WF et al. A randomized, controlled trial of methylprednisolone or naloxone in the treatment of acute spinal-cord injury: results of the Second National Acute Spinal Cord Injury Study. N Engl J Med 1990;322:1405–11.
34.Hurlbert RJ. Methylprednisolone for acute spinal cord injury: an inappropriate standard of care. J Neurosurg 2000;93:1–7.
35.Coleman WP, Benzel D, Cahill DW, et al. A critical appraisal of the reporting of the National Acute Spinal Cord Injury Studies (II and III) of methylprednisolone in acute spinal cord injury. J Spinal Disord 2000;13:185–99.
36.Short DJ, El Masry WS, Jones PW. High dose methylprednisolone in the management of acute spinal cord injury: a systematic review from a clinical perspective. Spinal Cord 2000;38:273–86.
37.Bracken MB, Shepard MJ, Holford TR, et al. Administration of methylprednisolone for 24 or 48 hours or tirilazad mesylate for 48 hours in the treatment of acute spinal cord injury: results of the Third National Acute Spinal Cord Injury Randomized Controlled Trial. National Acute Spinal Cord Injury Study. JAMA 1997;277:1597–604.
38.Hugenholtz H, Cass DE, Dvorak MF, et al. High-dose methylprednisolone for acute closed spinal cord injury: only a treatment option. Can J Neurol Sci 2002;29:227–35.
39.Pharmacological therapy after acute cervical spinal cord injury. Neurosurgery 2002;50(suppl):63–72.
40.Geisler FH, Coleman WP, Grieco G, et al. The Sygen multicenter acute spinal cord injury study. Spine 2001;26(suppl):87–98.
41.Pitts LH, Ross A, Chase GA, et al. Treatment with thyrotropin-releasing hormone (TRH) in patients with traumatic spinal cord injuries. J Neurotrauma 1995;12:235–43.
42.Pointillart V, Petitjean ME, Wiart L, et al. Pharmacological therapy of spinal cord injury during the acute phase. Spinal Cord 2000;38:71–6.
43.Lammertse DP. Update on pharmaceutical trials in acute spinal cord injury. J Spinal Cord Med 2004;27:319–25.
44.Geisler FH, Dorsey FC, Coleman WP. Recovery of motor function after spinal-cord injury: a randomized, placebo-controlled trial with GM-1 ganglioside. N Engl J Med 1991;324:1829–38.
45.Zemke D, Majid A. The potential of minocycline for neuroprotection in human neurologic disease. Clin Neuropharmacol 2004;27:293–8.
46.Brundula V, Rewcastle NB, Metz LM, et al. Targeting leukocyte MMPs and transmigration: minocycline as a potential therapy for multiple sclerosis. Brain 2002;125:1297–308.
47.Li WW, Setzu A, Zhao C, et al. Minocycline-mediated inhibition of microglia activation impairs oligodendrocyte progenitor cell responses and remyelination in a non-immune model of demyelination. J Neuroimmunol 2005;158:58–66.
48.Dommergues MA, Plaisant F, Verney C, et al. Early microglial activation following neonatal excitotoxic brain damage in mice: a potential target for neuroprotection. Neuroscience 2003;121:619–28.
49.Tikka T, Fiebich BL, Goldsteins G, et al. Minocycline, a tetracycline derivative, is neuroprotective against excitotoxicity by inhibiting activation and proliferation of microglia. J Neurosci 2001;21:2580–8.
50.Pi R, Li W, Lee NT, et al. Minocycline prevents glutamate-induced apoptosis of cerebellar granule neurons by differential regulation of p38 and Akt pathways. J Neurochem 2004;91:1219–30.
51.Lee SM, Yune TY, Kim SJ, et al. Minocycline inhibits apoptotic cell death via attenuation of TNF-alpha expression following iNOS/NO induction by lipopolysaccharide in neuron/glia co-cultures. J Neurochem 2004;91:568–78.
52.Wang J, Wei Q, Wang CY, et al. Minocycline up-regulates Bcl-2 and protects against cell death in mitochondria. J Biol Chem 2004;279:19948–54.
53.Stirling DP, Khodarahmi K, Liu J, et al. Minocycline treatment reduces delayed oligodendrocyte death, attenuates axonal dieback, and improves functional outcome after spinal cord injury. J Neurosci 2004;24:2182–90.
54.Arvin KL, Han BH, Du Y, et al. Minocycline markedly protects the neonatal brain against hypoxicischemic injury. Ann Neurol 2002;52:54–61.
55.Wang CX, Yang T, Noor R, et al. Delayed minocycline but not delayed mild hypothermia protects against embolic stroke. BMC Neurol 2002;2:2.
56.Wu DC, Jackson-Lewis V, Vila M, et al. Blockade of microglial activation is neuroprotective in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine mouse model of Parkinson disease. J Neurosci 2002;22:1763–71.
57.Du Y, Ma Z, Lin S, et al. Minocycline prevents nigrostriatal dopaminergic neurodegeneration in the MPTP model of Parkinson’s disease. Proc Natl Acad Sci USA 2001;98:14669–74.
58.Wang X, Zhu S, Drozda M, et al. Minocycline inhibits caspase-independent and -dependent mitochondrial cell death pathways in models of Huntington’s disease. Proc Natl Acad Sci USA 2003;100:10483–7.
59.Chen M, Ona VO, Li M, et al. Minocycline inhibits caspase-1 and caspase-3 expression and delays mortality in a transgenic mouse model of Huntington disease. Nat Med 2000;6:797–801.
60.Kriz J, Nguyen MD, Julien JP. Minocycline slows disease progression in a mouse model of amyotrophic lateral sclerosis. Neurobiol Dis 2002;10: 268–78.
61.Van Den BL, Tilkin P, Lemmens G, et al. Minocycline delays disease onset and mortality in a transgenic model of ALS. Neuroreport 2002;13: 1067–70.
62.Zhu S, Stavrovskaya IG, Drozda M, et al. Minocycline inhibits cytochrome c release and delays progression of amyotrophic lateral sclerosis in mice. Nature 2002;417:74–8.
63.Hart RG, Ravina BM. Randomized clinical trials of neuroprotective agents in Parkinson’s disease: the NINDS registry. Available at Accessed January 16, 2005. 7-18-2003. Ref Type: Report.
64.Thomas M, Ashizawa T, Jankovic J. Minocycline in Huntington’s disease: a pilot study. Mov Disord 2004;19:692–5.
65.Gordon PH, Moore DH, Gelinas DF, et al. Placebo-controlled phase I/II studies of minocycline in amyotrophic lateral sclerosis. Neurology 2004;62:1845–7.
66.Tsuji M, Wilson MA, Lange MS, et al. Minocycline worsens hypoxic-ischemic brain injury in a neonatal mouse model. Exp Neurol 2004;189:58–65.
67.Smith DL, Woodman B, Mahal A, et al. Minocycline and doxycycline are not beneficial in a model of Huntington’s disease. Ann Neurol 2003;54:186–96.
68.Diguet E, Fernagut PO, Wei X, et al. Deleterious effects of minocycline in animal models of Parkinson’s disease and Huntington’s disease. Eur J Neurosci 2004;19:3266–76.
69.Yang L, Sugama S, Chirichigno JW, et al. Minocycline enhances MPTP toxicity to dopaminergic neurons. J Neurosci Res 2003;74:278–85.
70.Diguet E, Gross CE, Tison F, et al. Rise and fall of minocycline in neuroprotection: need to promote publication of negative results. Exp Neurol 2004;189:1–4.
71.Wells JE, Hurlbert RJ, Fehlings MG, et al. Neuroprotection by minocycline facilitates significant recovery from spinal cord injury in mice. Brain 2003;126:1628–37.
72.Lee SM, Yune TY, Kim SJ, et al. Minocycline reduces cell death and improves functional recovery after traumatic spinal cord injury in the rat. J Neurotrauma 2003;20:1017–27.
73.Teng YD, Choi H, Onario RC, et al. Minocycline inhibits contusion-triggered mitochondrial cytochrome c release and mitigates functional deficits after spinal cord injury. Proc Natl Acad Sci USA 2004;101:3071–6.
74.Brines ML, Ghezzi P, Keenan S, et al. Erythropoietin crosses the blood-brain barrier to protect against experimental brain injury. Proc Natl Acad Sci USA 2000;97:10526–31.
75.Kumral A, Ozer E, Yilmaz O, et al. Neuroprotective effect of erythropoietin on hypoxic-ischemic brain injury in neonatal rats. Biol Neonate 2003;83:224–8.
76.Sattler MB, Merkler D, Maier K, et al. Neuroprotective effects and intracellular signaling pathways of erythropoietin in a rat model of multiple sclerosis. Cell Death Differ 2004;11(suppl 2):181–92.
77.Genc S, Kuralay F, Genc K, et al. Erythropoietin exerts neuroprotection in 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine-treated C57/BL mice via increasing nitric oxide production. Neurosci Lett 2001;298:139–41.
78.Moon C, Krawczyk M, Ahn D, et al. Erythropoietin reduces myocardial infarction and left ventricular functional decline after coronary artery ligation in rats. Proc Natl Acad Sci USA 2003;100:11612–7.
79.Calvillo L, Latini R, Kajstura J, et al. Recombinant human erythropoietin protects the myocardium from ischemia-reperfusion injury and promotes beneficial remodeling. Proc Natl Acad Sci USA 2003;100:4802–6.
80.Celik M, Gokmen N, Erbayraktar S, et al. Erythropoietin prevents motor neuron apoptosis and neurologic disability in experimental spinal cord ischemic injury. Proc Natl Acad Sci USA 2002;99:2258–63.
81.Gorio A, Gokmen N, Erbayraktar S, et al. Recombinant human erythropoietin counteracts secondary injury and markedly enhances neurological recovery from experimental spinal cord trauma. Proc Natl Acad Sci USA 2002;99:9450–5.
82.Kaptanoglu E, Solaroglu I, Okutan O, et al. Erythropoietin exerts neuroprotection after acute spinal cord injury in rats: effect on lipid peroxidation and early ultrastructural findings. Neurosurg Rev 2004;27:113–20.
83.Leist M, Ghezzi P, Grasso G, et al. Derivatives of erythropoietin that are tissue protective but not erythropoietic. Science 2004;305:239–42.
84.Brines M, Grasso G, Fiordaliso F, et al. Erythropoietin mediates tissue protection through an erythropoietin and common beta-subunit heteroreceptor. Proc Natl Acad Sci USA 2004;101:14907–12.
85.Ehrenreich H, Hasselblatt M, Dembowski C, et al. Erythropoietin therapy for acute stroke is both safe and beneficial. Mol Med 2002;8:495–505.
86.Plunet W, Kwon BK, Tetzlaff W. Promoting axonal regeneration in the central nervous system by enhancing the cell body response to axotomy. J Neurosci Res 2002;68:1–6.
87.Tuszynski MH. Neurotrophic factors. In: Tuszynski MH, Kordower JH, eds. CNS Regeneration: Basic Science and Clinical Advances. San Diego: Academic Press, 1999:109–58.
88.Levi-Montalcini R, Hamburger V. Selective growth stimulating effects of mouse sarcoma on the sensory and sympathetic nervous system of the chick embryo. J Exp Zool 1951;116:321–62.
89.Levi-Montalcini R, Hamburger V. A diffusible agent of mouse sarcoma, producing hyperplasia of sympathetic ganglia and hyperneurotization of viscera in the chick embryo. J Exp Zool 1953;123:233–88.
90.Liu Y, Kim D, Himes BT, et al. Transplants of fibroblasts genetically modified to express BDNF promote regeneration of adult rat rubrospinal axons and recovery of forelimb function. J Neurosci 1999;19:4370–87.
91.Liu Y, Himes BT, Murray M, et al. Grafts of BDNF-producing fibroblasts rescue axotomized rubrospinal neurons and prevent their atrophy. Exp Neurol 2002;178:150–64.
92.Shumsky JS, Tobias CA, Tumolo M, et al. Delayed transplantation of fibroblasts genetically modified to secrete BDNF and NT-3 into a spinal cord injury site is associated with limited recovery of function. Exp Neurol 2003;184:114–30.
93.Tobias CA, Shumsky JS, Shibata M, et al. Delayed grafting of BDNF and NT-3 producing fibroblasts into the injured spinal cord stimulates sprouting, partially rescues axotomized red nucleus neurons from loss and atrophy, and provides limited regeneration. Exp Neurol 2003;184:97–113.
94.Kwon BK, Liu J, Oschipok L, et al. Rubrospinal neurons fail to respond to brain-derived neurotrophic factor applied to the spinal cord injury site 2 months after cervical axotomy. Exp Neurol 2004;189:45–57.
95.Kobayashi NR, Fan DP, Giehl KM, et al. BDNF and NT-4/5 prevent atrophy of rat rubrospinal neurons after cervical axotomy, stimulate GAP-43 and Talpha1-tubulin mRNA expression, and promote axonal regeneration. J Neurosci 1997;17:9583–95.
96.Kwon BK, Liu J, Messerer C, et al. Survival and regeneration of rubrospinal neurons 1 year after spinal cord injury. Proc Natl Acad Sci USA 2002;99:3246–51.
97.Cai D, Shen Y, De Bellard M, et al. Prior exposure to neurotrophins blocks inhibition of axonal regeneration by MAG and myelin via a cAMP-dependent mechanism. Neuron 1999;22:89–101.
98.Qiu J, Cai D, Dai H, et al. Spinal axon regeneration induced by elevation of cyclic AMP. Neuron 2002;34:895–903.
99.Neumann S, Bradke F, Tessier-Lavigne M, et al. Regeneration of sensory axons within the injured spinal cord induced by intraganglionic cAMP elevation. Neuron 2002;34:885–93.
100.Pearse DD, Pereira FC, Marcillo AE, et al. cAMP and Schwann cells promote axonal growth and functional recovery after spinal cord injury. Nat Med 2004;10:610–6.
101.Lu P, Yang H, Jones LL, et al. Combinatorial therapy with neurotrophins and cAMP promotes axonal regeneration beyond sites of spinal cord injury. J Neurosci 2004;24:6402–9.
102.Nikulina E, Tidwell JL, Dai HN, et al. The phosphodiesterase inhibitor rolipram delivered after a spinal cord lesion promotes axonal regeneration and functional recovery. Proc Natl Acad Sci USA 2004;101:8786–90.
103.Chen MS, Huber AB, van der Haar ME, et al. Nogo-A is a myelin-associated neurite outgrowth inhibitor and an antigen for monoclonal antibody IN-1 [see comments]. Nature 2000;403:434–9.
104.GrandPre T, Nakamura F, Vartanian T, et al. Identification of the Nogo inhibitor of axon regeneration as a Reticulon protein. Nature 2000;403:439–44.
105.Prinjha R, Moore SE, Vinson M, et al. Inhibitor of neurite outgrowth in humans. Nature 2000;403:383–4.
106.McKerracher L, David S, Jackson DL, et al. Identification of myelin-associated glycoprotein as a major myelin-derived inhibitor of neurite growth. Neuron 1994;13:805–11.
107.Wang KC, Koprivica V, Kim JA, et al. Oligodendrocyte-myelin glycoprotein is a Nogo receptor ligand that inhibits neurite outgrowth. Nature 2002;417:941–4.
108.Grados-Munro EM, Fournier AE. Myelin-associated inhibitors of axon regeneration. J Neurosci Res 2003;74:479–85.
109.Fournier AE, GrandPre T, Strittmatter SM. Identification of a receptor mediating Nogo-66 inhibition of axonal regeneration. Nature 2001;409:341–6.
110.McGee AW, Strittmatter SM. The Nogo-66 receptor: focusing myelin inhibition of axon regeneration. Trends Neurosci 2003;26:193–8.
111.Kwon BK, Borisoff JF, Tetzlaff W. Molecular targets for intervention in spinal cord injury. Mol Interventions 2002;2:244–58.
112.Li S, Liu BP, Budel S, et al. Blockade of Nogo-66, myelin-associated glycoprotein, and oligodendrocyte myelin glycoprotein by soluble Nogo-66 receptor promotes axonal sprouting and recovery after spinal injury. J Neurosci 2004;24:10511–20.
113.Zheng B, Ho C, Li S, et al. Lack of enhanced spinal regeneration in Nogo-deficient mice. Neuron 2003;38:213–24.
114.Simonen M, Pedersen V, Weinmann O, et al. Systemic deletion of the myelin-associated outgrowth inhibitor Nogo-A improves regenerative and plastic responses after spinal cord injury. Neuron 2003;38:201–11.
115.Kim JE, Li S, GrandPre T, et al. Axon regeneration in young adult mice lacking Nogo-A/B. Neuron 2003;38:187–99.
116.Zheng B, Atwal J, Ho C, et al. Genetic deletion of the Nogo receptor does not reduce neurite inhibition in vitro or promote corticospinal tract regeneration in vivo. Proc Natl Acad Sci USA 2005;102:1205–10.
117.Davies SJ, Fitch MT, Memberg SP, et al. Regeneration of adult axons in white matter tracts of the central nervous system. Nature 1997;390:680–3.
118.Silver J, Miller JH. Regeneration beyond the glial scar. Nat Rev Neurosci 2004;5:146–56.
119.Bradbury EJ, Moon LD, Popat RJ, et al. Chondroitinase ABC promotes functional recovery after spinal cord injury. Nature 2002;416:636–40.
120.Chau CH, Shum DK, Li H, et al. Chondroitinase ABC enhances axonal regrowth through Schwann cell-seeded guidance channels after spinal cord injury. FASEB J 2004;18:194–6.
121.Murray M, Kim D, Liu Y, et al. Transplantation of genetically modified cells contributes to repair and recovery from spinal injury. Brain Res Brain Res Rev 2002;40:292–300.
122.Richardson PM, McGuinness UM, Aguayo AJ. Axons from CNS neurons regenerate into PNS grafts. Nature 1980;284:264–5.
123.Cheng H, Cao Y, Olson L. Spinal cord repair in adult paraplegic rats: partial restoration of hind limb function [see comments]. Science 1996;273:510–3.
124.Xu XM, Guenard V, Kleitman N, et al. A combination of BDNF and NT-3 promotes supraspinal axonal regeneration into Schwann cell grafts in adult rat thoracic spinal cord. Exp Neurol 1995;134:261–72.
125.Menei P, Montero-Menei C, Whittemore SR, et al. Schwann cells genetically modified to secrete human BDNF promote enhanced axonal regrowth across transected adult rat spinal cord. Eur J Neurosci 1998;10:607–21.
126.Tuszynski MH, Weidner N, McCormack M, et al. Grafts of genetically modified Schwann cells to the spinal cord: survival, axon growth, and myelination. Cell Transplant 1998;7:187–96.
127.Weidner N, Blesch A, Grill RJ, et al. 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 1999;413:495–506.
128.Cheng H, Liao KK, Liao SF, et al. Spinal cord repair with acidic fibroblast growth factor as a treatment for a patient with chronic paraplegia. Spine 2004;29:E284–8.
129.Fraidakis MJ, Spenger C, Olson L. Partial recovery after treatment of chronic paraplegia in rat. Exp Neurol 2004;188:33–42.
130.Steeves J, Fawcett J, Tuszynski M. Report of international clinical trials workshop on spinal cord injury February 20–21, 2004, Vancouver, Canada. Spinal Cord 2004;42:591–7.
131.Xu XM, Chen A, Guenard V, et al. Bridging Schwann cell transplants promote axonal regeneration from both the rostral and caudal stumps of transected adult rat spinal cord. J Neurocytol 1997;26:1–16.
132.Ramon-Cueto A, Cordero MI, Santos-Benito FF, et al. Functional recovery of paraplegic rats and motor axon regeneration in their spinal cords by olfactory ensheathing glia. Neuron 2000;25:425–35.
133.Barnett SC, Riddell JS. Olfactory ensheathing cells (OECs) and the treatment of CNS injury: advantages and possible caveats. J Anat 2004;204:57–67.

spinal cord injury; neuroprotection; axonal regeneration; minocycline; erythropoietin; clinical trials; neurotrophic factors; olfactory ensheathing cells

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