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Journal of Physiology (2001), 533.1, pp. 83-89
© Copyright 2001 The Physiological Society
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ABSTRACT |
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Little spontaneous regeneration of function and structure occurs following spinal cord injury. A number of factors are thought to contribute to this lack of recovery, including glial scarring (for review see Fawcett & Asher, 1999), myelin inhibition (for review see Huber & Schwab, 2000), cell death (for review see Beattie et al. 2000), insufficient growth factor support and the lack of permissive substrates for axonal regeneration. Spontaneous regeneration occurs, however, after peripheral nerve injury, supported by both the ordered structure of the peripheral nerve and a number of intrinsically supportive properties of Schwann cells (SCs). SCs actively phagocytose peripheral nerve debris, produce neurotrophic factors, and secrete extracellular matrix molecules that support axonal regrowth. Thus, a feasible strategy for augmenting spinal cord regeneration is to adopt some of the favourable properties exhibited in peripheral nerve after injury; namely, the provision of neurotrophic factors and the grafting of SCs to promote axonal regeneration in the CNS. The following sections will describe effects of delivering neurotrophic factors and SCs to sites of spinal cord injury. Recent studies have also used the olfactory ensheathing cell to establish a cellular bridge in the injured spinal cord. These studies have been extensively outlined in recent reviews (Ramon-Cueto, 2000; Franklin & Barnett, 2000) and will not be further discussed in this review. Gene therapy as a vehicle for delivery of large quantities of neurotrophic factors to specific sites of spinal cord injury will also be discussed.
Neurotrophic factors and gene therapy in the injured spinal cord
Neurotrophic factors are key nervous system regulatory proteins that modulate neuronal survival, axonal growth, synaptic plasticity and neurotransmission. Mounting evidence over the last few years indicates that neurotrophic factors are instrumental in eliciting renewed axonal growth following spinal cord injury. An increasing repertoire of experimental lesion protocols, axonal markers and axonal tracers has helped identify various responses of specific axonal populations to different neurotrophic factors.
Neurotrophic factor-mediated axonal responses to injury vary in extent and are determined by axonal phenotype, the production of receptors to growth factors, the type of lesion, and the local environment (see Table 1). The distance of the lesion from the neuronal cell bodies and the extent of the lesion are critical in cell survival, degree of cellular response to the injury, and ability for renewed axonal growth to find the appropriate target (for review see Steeves & Tetzlaf, 1998). Axonal regrowth is also dependent on the extent of spared grey matter and the deposition of putatively inhibitory extracellular matrix molecules (Grill et al. 1997; Jones et al. 2000).
The route of neurotrophic factor administration to the injured spinal cord influences the nature of axonal responses. Many neurotrophic factors elicit a chemotropic effect, directing the growth of axonal populations to regions of the highest growth factor concentration. Thus, to promote axonal growth specifically into a site of spinal cord injury, initially the highest concentrations of growth factors should be delivered specifically to the injury site. Once axons penetrate the lesion site, or a graft in the lesion site, then a means of diversifying axonal gradients to points distal to the injury would be desirable to promote axonal growth 'downstream' toward denervated targets. Traditionally, neurotrophic factors have been introduced to the injured spinal cord by direct injection (Schnell et al. 1994), continuous infusion (Bradbury et al. 1998), or placement of growth-factor saturated Gelfoam, (UpJohn, Kalamazoo, MI, USA) (Shibayama et al. 1998). However, these methods did not achieve long-term, localized, high dose neurotrophic factor delivery. An alternative approach that achieves long-term and site-specific delivery of neurotrophic factors to the injured spinal cord is ex vivo gene therapy. The latter approach has been useful for promoting localized, robust axonal growth after spinal cord injury (for review see Tuszynski, 1997). However, gene therapy has not yet achieved the goal of diversifying axonal growth after initial axonal growth, although such methods are under development (Blesch et al. 2000). Nonetheless, ex vivo gene delivery to sites of spinal cord injury has been useful in identifying patterns of axonal sensitivity to growth factors in the context of both acute and chronic spinal cord injury.
Ex vivo gene therapy involves removal of cells from the host organism, genetic modification of these cells in vitro, characterization of the extent of gene product expression by the modified cells in vitro, followed by transplantation of these modified cells back into the host. Thus, specific cell types for gene therapy and neural repair can be targeted, such as Schwann cells, fibroblasts, glia or stem cells. Cells are selected in vitro for successful incorporation of the transgene and are then analysed for transgene expression and biological activity. Cells with high transgene expression are expanded in culture and grafted into the host spinal cord. There is no risk of immunological rejection because cells from the same individual are used. Thus, this approach has the capability of providing a high concentration of long-term, localized growth factor delivery to a site of injury.
Recent studies have broadened our understanding of differential axonal responses to neurotrophic factors following spinal cord injury. Emphasis has been placed on the 'classic' neurotrophin family, consisting of nerve growth factor (NGF), brain-derived neurotrophic factor (BDNF), neurotrophin-3 (NT-3), and neurotrophin-4/5 (NT-4/5). Growth factors of other families, including the cytokine growth factors and the transforming factor-
family that includes glial cell-line derived neurotrophic factor (GDNF), have also been demonstrated to enhance axonal growth in models of spinal cord injury. The regulative pattern of these neurotrophic factors in the brain demonstrate both specificity and overlap suggesting that each one plays a distinct role in modulating neural function (for review see Tuszynski, 1999). Indeed, specific neurotrophic factors elicit regrowth in specific axonal populations after spinal cord injury, and future experiments will probably examine the provision of a combination of neurotrophic factors to recruit the growth of multiple injured axonal populations.
Nerve growth factor. NGF has proved to be a potent stimulus for sensory axon growth after injury. A continuous infusion of NGF in the dorsal spinal cord rostral to a peripheral nerve graft promotes growth of sensory axons from the graft into the dorsal column white matter (Oudega & Hagg, 1996). After peripheral dorsal root crush lesions, only 3 % of sensory axons re-enter the spinal cord through the dorsal root entry zone (Oudega & Hagg, 1996). This proportion is significantly increased to 37 % after 14 days of continuous NGF infusion through catheters placed in the dorsal spinal cord. These regenerating axons penetrate spinal cord white matter and grow up to 3 mm past the dorsal root entry zone. More extensive experiments confirmed this NGF-driven regrowth of injured sensory fibres into the spinal cord (Ramer et al. 2000) and identified the regenerating axons as positive for calcitonin gene related peptide (CGRPpositive), a marker for small diameter, unmyelinated peptidergic axons. This study confirmed with electrophysiology that postsynaptic potentials could be evoked in the dorsal horn of the spinal cord after peripheral stimulation of the injured axons. Functional improvements were also observed using noxious behavioural testing.
In further experiments, NGF was delivered to the injured spinal cord using grafts of primary fibroblasts that were genetically modified to produce and secrete NGF into sites of spinal cord injury. In addition to confirming robust sensitivity of injured CGRP-expressing sensory nociceptive axons to NGF, these experiments showed that an NGF source within the spinal cord also attracted the growth of coerulospinal axons (immunolabelled for tyrosine hydroxylase and dopamine beta-hydroxylase) and ventral motor axons (immunolabelled for choline acetyltransferase, ChAT; Tuszynski et al. 1996). Growth from corticospinal and raphaespinal (serotonergic) axons was not detected. Of note, whereas axons extensively penetrated these NGF-secreting grafts, these axons did not exit the grafts, presumably because of the high continuous levels of growth factors expressed within the graft or unfavourable, non-permissive conditions outside the graft (i.e. glial scarring or myelin inhibition). Not surprisingly, therefore, these grafts do not support functional recovery (Tuszynski et al. 1997). This highlights the need for a neurotrophin delivery system that can sequentially deliver high doses of neurotrophins to injury sites, then switch delivery to sites 'downstream' from the injury site.
Brain-derived neurotrophic factor. A continuous infusion of BDNF in the dorsal spinal cord rostral to a peripheral nerve graft was shown to promote regeneration of sensory axons from the graft into the dorsal column white matter (Oudega & Hagg, 1999). Following a mid-thoracic contusion spinal cord lesion, continuous, high concentrations of BDNF, infused over a 4 week period into the intrathecal space, were found to enhance the growth of ChAT-positive motor axons at the injury epicentre (Jakeman et al. 1998). No growth-promoting effect was observed on raphaespinal serotonergic axons. Functional tests in these rats demonstrated that BDNF delivery stimulated hindlimb air-stepping, a co-ordinated movement thought to be regulated by interneuronal networks in the lumbar spinal cord referred to as central pattern generators. This suggests that BDNF could play a role in reducing the threshold for activation of existing, non-injured central pattern generators in the spinal cord below a lesion site, thereby generating a 'functional' improvement without necessarily promoting axonal regeneration. However, in a different study, fibroblasts were genetically modified to produce BDNF and were grafted directly into the spinal cord in a partial cervical hemisection lesion site. These grafts reportedly promoted regeneration of injured rubrospinal tract axons (Liu et al. 1999) into and distal to the lesion site, terminating in the spinal cord grey matter; there was increased use of the affected forelimb in these grafted animals. Long-distance growth reportedly occurred despite the lack of a known downstream growth factor source to promote axonal egress from the graft. Thus, the mechanism of this long-distance growth has not been established. Additional studies are required to investigate whether mechanisms other than regeneration of rubrospinal axons (i.e. BDNF effects on local plasticity) may generate functional recovery. Such experiments in all studies of regeneration should incorporate morphological, electrophysiological and behavioural analysis to determine better whether axonal regeneration directly contributes to functional recovery. Another study using genetically modified BDNF-secreting fibroblasts implanted into spinal cord lesion sites demonstrated significant growth of local motor axons, CGRP-labelled dorsal root sensory axons, and supraspinal coerulospinal axons (P. Lu, A. Blesch & M. H. Tuszynski, unpublished observations). The effects of BDNF on rubrospinal axons were not examined in this study. Previous studies involving infusion of BDNF into the intrathecal space failed to demonstrate effects on sensory axons (Bradbury et al. 1999; Ramer et al. 2000), in contrast to the gene delivery study mentioned above. This highlights the importance of achieving high-dose, localized growth factor delivery in order to elicit significant axonal growth.
Glial cell-line derived neurotrophic factor. Following dorsal root injury, GDNF proved to be the most effective neurotrophic factor in stimulating axonal growth across the dorsal root entry zone into the spinal cord white matter (Ramer et al. 2000). A robust regrowth of CGRP-positive sensory fibres was observed central to the transition zone following prolonged infusion of GDNF, with electrophysiology and noxious behavioural tests confirming successful regeneration. GDNF-transduced fibroblasts, grafted into a spinal cord dorsal hemi-lesion site, also greatly enhanced sensory axon growth (Blesch et al. 1998). In addition, GDNF elicited robust growth of lesioned local motor axons. However, GDNF gene delivery did not augment the growth of corticospinal, coerulospinal or raphaespinal axons.
Neurotrophin-3. To date, NT-3 is the only neurotrophic factor that has been identified to promote the growth of corticospinal axons after spinal cord injury (Schnell et al. 1994; Grill et al. 1997). When delivered as a single injection into the lesioned spinal cord, NT-3 promoted sprouting of corticospinal axons (Schnell et al. 1994). When delivered to the spinal cord using a continuous ex vivo gene delivery approach, NT-3 promoted the growth of corticospinal axons and achieved partial functional recovery (Grill et al. 1997). In the latter study, NT-3 gene delivery did promote corticospinal axon growth distal to the lesion site, perhaps as a result of a serendipitous event: corticospinal axons did not penetrate the NT-3-secreting cell grafts. Rather, corticospinal axons extended through remaining bridges of host grey matter surrounding the lesion site and extended for significant distances of up to 8 mm distal to the graft. Presumably, the milieu of the graft itself could not support corticospinal axon growth, or inhibitory molecules at the graft-host interface inhibited their penetration of the graft (Jones et al. 2000); thus, the corticospinal axons did not become enmeshed in the graft. Also confirming the finding that NT-3 supports corticospinal axon growth, Blesch et al. (1999) used an ex vivo gene delivery approach to deliver the cytokine growth factor, leukaemia inhibitory factor (LIF), to the spinal cord. It was found that LIF gene delivery upregulated NT-3 expression in the spinal cord, and that corticospinal axonal growth was significantly increased in grey matter at the lesion site. This study also highlighted the point that delivery of one growth factor to the CNS may augment the production of other growth factors, thereby sustaining neuronal growth or survival. In another study, a continuous infusion of NT-3 into the dorsal spinal cord was reported to promote the growth of sensory axons from a peripheral nerve graft into the dorsal column white matter (Oudega & Hagg, 1999). Also, infusion of NT-3 into the intrathecal space promoted regeneration of dorsal column sensory axons into and beyond a site of spinal cord injury (Bradbury et al. 1999).
Thus, a variety of axonal populations of the spinal cord respond to neurotrophic factors. The elucidation of specific patterns of sensitivity will allow the design of rational strategies for promoting more extensive axonal growth in the context of injury and degeneration.
Schwann cells promote axonal regeneration in the injured spinal cord
As early as 1913, Ramon y Cajal (1928) and his colleague, F. Tello, recognized the powerful regeneration-promoting properties of peripheral nerve, which contains SCs, the glia that form myelin around peripheral axons. Over two decades ago, Richard P. Bunge proposed the possibility of isolating and culturing SCs from a patient's peripheral nerve for autologous implantation into the injured spinal cord. Accordingly, large numbers of SCs can now be generated from both adult rat (Morrissey et al. 1991) and human (Levi et al. 1995; Rutkowski et al. 1995; Casella et al. 1996) peripheral nerve for transplantation studies or therapy, respectively.
The efficacy of SCs in promoting axonal regeneration in the injured adult spinal cord has been studied extensively (for review see Bunge, 1993, 2000; Bunge & Kleitman, 1997, 1999; see also Montgomery et al. 1996). Implantation in the completely transected adult rat thoracic spinal cord of a semipermeable polyacrylonitrile/ polyvinylchoride (PAN/PVC) polymer tube filled with SCs, isolated from adult rat sciatic nerve and mixed with Matrigel (Becton Dickinson Labware, Bedford, MA, USA) was shown to promote axonal growth across the gap (Xu et al. 1995b, 1997). The cord stumps at 1 month were united by a tissue cable that contained SCs, ensheathed unmyelinated and myelinated axons, blood vessels, fibroblasts, meningeal cells and usually some macrophages (Xu et al. 1995a,b, 1997; Chen et al. 1996; Oudega et al. 1997). Nerve fibres grew into the bridge from both cord stumps. An estimated 25 % of the total number of fibres within the bridge were myelinated by SCs. Myelinated axons were, in general, rare in Matrigel only bridges, demonstrating that SCs were key to axonal growth across the bridge.
Retrograde tracing experiments revealed that the majority of the axons in SC implants were derived from propriospinal neurons located as far rostral as the third cervical spinal cord level, and as far caudal as the fourth sacral level (Xu et al. 1997). The responding axons grew across the SC implant but did not grow into the opposite cord stump. Also, supraspinal fibres, which derive from brainstem nuclei and are involved in control of locomotion, were rare in these grafts; there was limited growth of serotonergic (raphe) and noradrenergic (locus coeruleus) fibres into the SC bridges. Important for future human application is that implantation of human SCs in the nude (T-cell deficient) rat spinal cord elicited an axonal growth response as well (Guest et al. 1997).
In the studies mentioned above, implantation of a SC-filled PAN/PVC tube in the transected spinal cord resulted in loss of tissue primarily in the stumps inserted into the tube (at the graft-host cord interfaces) (Xu et al. 1995a,b, 1997; Chen et al. 1996; Oudega et al. 1997). Loss of nervous tissue at the interfaces most likely limits the overall regenerative response. To reduce this tissue loss, the grafting of a SC-filled PAN/PVC tube was combined with administration of the neuroprotectant glucocorticosteroid, methylprednisolone (Chen et al. 1996). This combination treatment was shown to reduce tissue loss at the interfaces, to enhance the number of axons in the bridge, to promote supraspinal growth into the SC graft, and to enable modest growth of regenerated fibres from the graft into the cord (Chen et al. 1996; for review see Bunge & Kleitman, 1999). The observed preservation of nervous tissue and enhanced axonal growth may be related to the methylprednisolone-induced decrease in the number of activated microglia and/or macrophages near the graft-cord interfaces (Oudega et al. 1999).
Over the last two decades, the SC has been established as an important candidate for future surgical repair strategies aimed at functional recovery in humans with spinal cord injury. The implantation studies mentioned above, however, have clearly shown that grafting SCs alone in the injured adult rat spinal cord does not result in substantial regeneration of supraspinal axons or in fibre growth from the implant into the spinal cord (Xu et al. 1997). Additional interventions need to be combined with SC implantation strategies to stimulate additional axon populations to grow, and to achieve re-entry of regenerating fibres into the spinal cord, thus increasing the chances for functional recovery. Adding olfactory ensheathing glia to the SC bridge paradigm enabled growth of regenerated fibres from the graft into the cord. The use of these glia is therefore of potential utility in promoting axonal growth in spinal cord injury models (for further review see Plant et al. 2000; Kleitman & Bunge, 2000).
Schwann cells combined with neurotrophins improve axonal regeneration in the injured spinal cord
Among the interventions tested to improve the axonal regeneration achieved with SC bridges was the addition of neurotrophins. In one study (Xu et al. 1995a), BDNF and NT-3 were both infused into the space around the SC cable inside the PAN/PVC-channel for 14 days. After an additional 14 days, myelinated axon number in the graft and the number of propriospinal neurons that regenerated axons into the graft were both tripled compared with animals not receiving the neurotrophins. Also, neurons in the brainstem extended axons into the bridge in contrast to control animals in which there was little response because of the distance between the nerve cell bodies and the thoracic bridge. Responding neurons were predominantly located in vestibular nuclei. Thus, regeneration of some neuronal populations distant from the treatment was elicited by a combination of trophic factors and a favourable cellular substrate.
In another study (Menei et al. 1998), SCs were infected with a retroviral vector carrying the human prepro BDNF/cDNA before transplantation. These cells or untreated SCs were deposited in a 5 mm long trail extending into the distal cord from the complete transection site and into the transection site itself. The SC trails remained largely intact for at least a month, the duration of the experiment. When animals receiving the genetically modified SCs were compared with those receiving untreated SCs, more axons, including those from brainstem neurons, were present in the trail beyond the transection site. This finding was confirmed by the presence of tracer, injected at the distal end of the trail, in an increased number of brainstem neurons. Untreated SCs elicited a modest response from vestibular neurons, whereas the BDNF-secreting SCs engendered a larger response principally from reticular and raphe neurons (Menei et al. 1998). Neither brainstem axons nor labelled brainstem neurons were found rostral to the transection when SCs were not transplanted. Thus, transplantation of human BDNF-secreting SCs further improved the regenerative response across the transection site and into the thoracic cord the length of the SC trail.
Recently, it was tested whether adenoviral vector (Ad)-mediated delivery of BDNF and NT-3 distal to a SC graft in the completely transected adult rat thoracic spinal cord would promote axonal growth from the graft into the distal cord (Blits et al. 2000b). One week after implantation of the SC bridge, a mixture of Ad-BDNF and Ad-NT-3 (50:50; total: 2 
107 plaque-froming units) was injected into the cord 7 mm distal to the distal graft-cord interface. Control animals received Ad-LacZ or saline. Treatment with Ad-BDNF and Ad-NT-3 modestly but significantly improved hindlimb function compared with the control (Ad-LacZ) group. Using the axonal tracer, fast blue, it was reported that more axons extended beyond the SC graft in the neurotrophin group compared with the control groups. These results suggest that gene transfer techniques can be used to promote axonal regeneration from an intraspinal graft into the distal spinal cord.
To promote regeneration of corticospinal axons, Blits et al. (2000a) implanted in the hemisected adult rat thoracic spinal cord several intercostal nerves transduced with an Ad encoding for NT-3. Three months after implantation, neurofilament positive fibres were present in the grafts. Anterograde tracing with biotinylated dextran amine demonstrated that corticospinal axons avoided the grafts but grew in undamaged spinal cord tissue beyond the site of implantation. This implantation strategy reportedly improved hindlimb function (Blits et al. 2000a) using the Basso, Beattie & Bresnahan locomotor rating scale (BBB; Basso et al. 1995).
Conclusions
Although the problem of promoting regeneration after spinal cord injury remains formidable, progress has been made in enhancing the growth potential of injured axons. Neurotrophins can enhance extension of specific populations of injured axons. When combined with substances that constitute a permissive substrate for growth, such as Schwann cell bridges, growth is further enhanced. Some reports of limited functional recovery based on these approaches have begun to appear. Future studies will combine the provision of growth factor delivery and Schwann cells with additional strategies to promote axonal growth, including neutralization of myelin- and extracellular matrix-associated inhibitors.
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Acknowledgements
Leonard Jones is supported by the NIH (NINDS Grant NS 10927). Martin Oudega is a Werner Heumann Memorial International Scholar. Mary Bartlett Bunge is supported by the Christopher Reeve Paralysis Foundation, the NIH (NINDS Grant NS 09923) and the Miami Project to Cure Paralysis. Mark Tuszynski is supported by the NIH (NINDS Grant NS 37083) and the Veterans Administration.
Corresponding author
M. H. Tuszynski: Department of Neurosciences-0626, University of California, San Diego, 9500 Gilman Drive, La Jolla, CA 92093, USA.
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