Spinal cord regeneration: intrinsic properties and ...

seminars in CELL & DEVELOPMENTAL BIOLOGY, Vol. 13, 2002: pp. 361–368 doi:10.1016/S1084–9521(02)00092-7, available online at http://www.idealibrary.com on

Spinal cord regeneration: intrinsic properties and emerging mechanisms Ellen A.G. Chernoff ∗ , Kazuna Sato, Angela Corn and Rachel E. Karcavich include fetal tissue, stem cell, Schwann cell, olfactory ensheathing cell, and peripheral nervous system transplants.1, 4 Another approach examines those cases in which vertebrate spinal cord demonstrates intrinsic regeneration capacity.5–7 In adult animals, spinal cord regeneration succeeds in teleost fish,8–10 urodele amphibians (newts and salamanders),11–13 and lizards (tails only).14–17 Among immature animals regeneration occurs in premetamorphic tadpoles of anuran amphibians (frogs and toads),18–21 embryonic birds, and fetal mammals22–24 . In most cases, adult mammalian central nervous system (CNS) fails to regenerate spontaneously following injury, but there are examples in which brain tissue is resistant to excitoxic injury and spinal cord regeneration can occur following specific types of lesions.1, 7 Understanding the mechanisms of spontaneous spinal cord regeneration will generate an additional set of tools applicable to human spinal cord injury.

Injured spinal cord regenerates in adult fish and urodele amphibians, young tadpoles of anuran amphibians, lizard tails, embryonic birds and mammals, and in adults of at least some strains of mice. The extent of this regeneration is described with respect to axonal regrowth, neurogenesis, glial responses, and maintenance of an ‘embryonic’ environment. The regeneration process in amphibian spinal cord demonstrates that gap replacement and caudal regeneration share some properties with developing spinal cord. This review considers the extent to which intrinsically regenerating spinal cord demonstrates neural stem cell behavior and to what extent anterior–posterior and dorsal–ventral patterning might be involved. Key words: spinal cord regeneration / amphibian regeneration / ependymal cell / neurogenesis / neural stem cell © 2002 Elsevier Science Ltd. All rights reserved.

Introduction

What processes occur?

Numerous mammalian spinal cord injury models are being used to produce approaches useful in achieving human spinal cord regeneration. Regimens protecting against excitotoxicity and other pathological influences are pursued to provide mechanisms for the prevention of secondary cell death. A wide range of experiments in mammals explores the role of toxic myelin breakdown products, the supportive effects of neurotrophic factors, and the inhibitory effects of glial and fibroblastic scars.1–3 Transplantation strategies to augment mammalian spinal cord regeneration

There are phylogenetic differences in the regeneration capacity of organisms and different types of regeneration responses are mounted in different species.6 Spinal cord regeneration experiments have been conducted using different techniques and addressing different questions, so the absence of a particular aspect of spinal cord regeneration in a given species, such as new neurogenesis, may simply not have been addressed. In fish, neurogenesis and an ependymal response have been described.8–10, 25 Urodele amphibian regeneration studies have described axonal sprouting (uninjured neurons), frank regeneration (axonal regrowth), neurogenesis, and an ependymal response.11–13, 20, 26–28 In higher vertebrates, the limited examples of spontaneous spinal cord regeneration in developing animals and recent results in an adult mouse injury model demonstrate frank regeneration, sprouting, and glial responses.7, 22, 23

From the Department of Biology and the Indiana University Center for Regenerative Biology and Medicine, School of Science, Indiana University–Purdue University Indianapolis (IUPUI), 723 W. Michigan St., Indianapolis, IN 46202-5132, USA. * Corresponding author. E-mail: [email protected] © 2002 Elsevier Science Ltd. All rights reserved. 1084–9521 / 02 / $– see front matter

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The intrinsic properties that permit regeneration in spinal cord are complex. There are subtle differences among species within a given regeneration-competent phylogenetic group, and there can be expression of seemingly favorable properties in animals that fail to regenerate. Four features of regenerating cord that have been explored across multiple experimental systems are: axonal regrowth, retention of neurogenic capacity, the capacity to mount a beneficial glial response, and retention or reconstruction of an ‘embryonic’ trophic environment.20, 29 Beyond the source of cells and their behavior in regenerating tissue are fundamental questions about their differentiative state and patterning of the regenerating tissue. Are the cells mediating spinal cord regeneration radial glia or neural stem cells? How much of the embryonic gene expression mediating anterior–posterior (A–P) and dorsal–ventral (D–V) patterning is required for regeneration versus wound repair in the CNS? The following sections address the nature and extent of spontaneous spinal cord regeneration.

ation fails when fully metamorphosed frogs and toads are lesioned. Regeneration fails earlier in some species and some axonal tracts than others. In the ranid frog Rana temporaria, regeneration of dorsal column axons (dorsal root ascending sensory column) fails before regeneration of axons in the ventral funiculus.19, 20 In the bullfrog Rana catesbeiana, it appears that the dorsal ascending axons fail to regenerate at all.18 In Xenopus laevis tadpoles (a pipid frog), brainstem-spinal axons regenerate across a spinal transection site at a later equivalent stage than that reported for ranid tadpoles.21 In higher vertebrates axonal regrowth can occur in lesioned embryonic spinal cord. When embryonic chick spinal cord is transected at embryonic day 5, nerve fibers crossed the transection 48 h later. Shimizu et al.22 showed that chick spinal cord regeneration could occur until approximately day 15 of development. The limit for regeneration of brainstem-spinal neurons projecting to the lumbar spinal cord was identified as day 13 in chick embryos.23 Replacement of gaps in 1-day-old neonatal rat spinal cord with spinal cord segments from 14- to 16-day embryonic cord elicited axonal regrowth from brain stem across the lesion site to the lumbar enlargement.24 Retrograde labeling experiments in dura mater-sparing lesions of mature mice showed substantial regeneration associated with the rubrospinal and vestibulospinal tracts.7

Axonal regrowth The degree of spinal cord regeneration is often assayed by the extent of success or failure of axonal regrowth. This can be regrowth from axotomized neurons or sprouting of axons from intact neurons cranial or caudal to the lesion. Among strong regenerators, like the urodele amphibians, functional recovery occurs with the regeneration of relatively few axons in a short period of weeks or a few months in both the newt Notopthalmus viridescens (formerly Triturus) and the axolotl (Ambystoma mexicanum).13, 30 Production of control levels of axons requires an extended period of time. Adult axolotls required 23 months for the number of axons from neurons in the nuclei of the medulla, midbrain, and diencephalon to reach control levels in regenerated lumbar spinal cord.31 In the anuran spinal cord, regeneration fails as the tadpoles approach the end of metamorphic climax. Direct comparison of the stage at which regeneration fails among anuran species is difficult due to heterochrony in development, differences between rapid and slow developers and different staging schemes. When regeneration does occur, the regenerated axons follow their normal pathway.19 Anuran tadpoles lesioned during the permissive period must progress through metamorphosis in order to achieve complete regeneration.18, 21 It is generally agreed that regener-

Neurogenesis Mitotic labeling studies and detection of cell-cycleassociated proteins show that neurogenesis following spinal cord injury is common in lower vertebrate spinal cord. In fish, the ependymal layer generates new neuroblasts which proliferate in vivo and in vitro and differentiate into new neurons.8, 10, 25 The process can be asymmetrical with the caudal stump producing more new cells that the rostral stump.10 In urodele amphibians that continue to grow throughout their lives (indeterminate growth), new neurons are produced in intact animals. Neurogenesis is more frequent in younger animals, such as axolotls (A. mexicanum) less than 7.5 cm (less than 6–7 months).29 In urodele tail regeneration, new CNS and peripheral nervous system (PNS) neurons are produced from an ependymal tube.11, 27, 28, 32, 33 High levels of endogenous fibroblast growth factor-2 (FGF-2) are associated with generation of new CNS and PNS neurons in salamander tail cord regeneration.27 362

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dies following injury. In rats, crush injuries can be performed in which the dura mater is preserved, preventing retraction of the cranial and caudal stumps.1 A syrinx lined with glial scar or connective tissue forms and increases in size with time. Semaphorin3A, from the meningeal ‘fibroblasts’, and EphB3, from the astrocytes, have been implicated in scar formation.41, 42 In mice the spinal cord glial response is different. Crush or complete transection injuries in mice usually produce a connective tissue scar with very little syrinx formation. With time the scar remodels, bringing the cranial and caudal stumps together. Transection injuries to C57Bl/6 mice in which dura mater is minimally damaged, and the dorsal vein is intact, result in functional recovery in many animals. This form of lesioning the cord minimizes the formation of a connective tissue scar. Glial fibrillary acidic protein (GFAP) is localized in bridges across white and gray matter associated with axonal regrowth.7 Ependymal cells close the stump end central canal, and may be more proliferative at and above the lesion, but they do not grow out of the lesion site as in amphibian cord (A. Seitz and E. Heber-Katz, unpublished). The tissue response to excitotoxic brain injury varies widely among mouse strains with some strains are far more resistant to excitotoxic cell death than others.1 Strain differences in response to spinal cord injury might also exist. The C57Bl/6 strain that exhibits spinal cord regeneration is one of the strains resistant to excitotoxic injury, but at least one other strain of mice showed the same result, suggesting that the dura-sparing lesion may be more significant.1, 7 (A. Seitz and E. Heber-Katz, unpublished)

Glial responses Glial proliferative response and tissue reorganization varies between regeneration-competent and -incompetent spinal cord. The behavior of the spinal cord ependymal cells is an integral part of regeneration of larval, juvenile, and adult urodele amphibian spinal cord. In nontail spinal cord, a crush or transection stimulates the ependymal cells lining the central canal to rearrange, sealing the damaged cord stumps. Ependymal cells migrate into the lesion site from cranial and caudal stumps in a process termed gap replacement.12, 13 The ependymal cells are joined by infiltrating lymphocytes. The injury-reactive ependymal cells proliferate, migrate, and remove existing extracellular matrix material and debris from dead cells.9, 11–13, 15, 34, 35 In gap replacement, the ependymal cells reorganize from an epithelial form into a mesenchyme and back again, with characteristic changes in intermediate filament and extracellular matrix components.5, 26 In tail amputation (caudal regrowth), the ependyma do not reorganize completely.6, 32 In both types of urodele cord regeneration, axons and dendrites grow in the spaces between ependymal endfeet or processes in close contact with the basal lamina.36 In regeneration-competent anuran amphibian cord, there is an ependymal response while in regenerationincompetent anuran cord there is not.19, 37 In regenerating tadpole spinal cord, regrowing axons are in contact with ependymal processes. The spinal cord in higher vertebrates regenerates most readily in embryonic or fetal animals with growing CNS, during embryonic or fetal life.38 One phenomenon that is shown or noted, but not currently pursued, by researchers is the occurrence of an ependymal response. Reactive gliosis and the formation of fibroblastic scars are absent. Transection of embryonic avian spinal cord produces disorganization of the ependymal zone with occlusion of the central canal in conjunction with axonal regrowth.22 In rats, transplantation of fetal spinal cord into transected spinal cord of neonatal rats produces reorganization of periluminal cells in the graft that causes the disappearance of the central canal, suggesting an ependymal response from the fetal tissue.24 In the adult rat, transection of the spinal cord can induce both astrocyte gliosis and connective tissue proliferation.39, 40 In rats, the scar in lesioned spinal cord takes the form of a complex of astrocytic and connective tissue that forms between neural tissue and a syrinx (a fluid-filled cavity) that forms as tissue

Embryonic environment It has been suggested that salamander and frog tadpole cord regenerate because the animals are still growing and adding new neurons.29 In the case of salamanders, a great deal of research has used neotenic (nonmetamorphosing) species such as the axolotl. It was proposed that continued growth throughout the life of the axolotl reflects the retention of embryonic properties that permit regeneration.29, 43 While the animals are still growing, the mechanism for nerve cell replacement and tissue reconstruction may still be active. In the frog R. temporaria, it has been suggested that the period of neural regeneration in the dorsal column coincides with the capacity for neurogenesis in the dorsal root ganglia (DRG); spinal cord regenerates in the areas still undergoing growth.43 In urodele amphibians, new neurogenesis occurs as 363


part of the regeneration process.27, 28 Many urodeles do reach a final adult size, however, and can still regenerate. Adult N. viridescens normally show no mitotic figures in intact spinal cord, but can regenerate spinal cord across a gap over a period of months.44 Tail amputation in fully grown adult N. viridescens and Pleurodeles waltl also results in cord regeneration with neurogenesis.31 The persistence of tissue-spanning radial processes from the ependymal cells in urodele spinal cord could represent retention of embryonic radial glial properties. The intermediate filament content of the outer (basal) segment of the ependymal process consists of GFAP while the luminal cell body contains cytokeratins.45, 46 The radial fiber terminates on the spinal cord basal lamina in a GFAP-positive pial endfoot. Frog ependymal cells also retain GFAP-containing radial cell processes, and these processes persist after spinal cord lesioning.44, 45 However, the mature frog spinal cord does not regenerate.18, 29 Existence of radial glial processes per se is not an indicator of regenerative capacity. There are changes in extracellular matrix components correlated with amphibian spinal cord regeneration. These changes are often regarded as the retention or reconstruction of a more embryonic spinal cord. The intact amphibian spinal cord produces laminin, but in gap replacement the mesenchymal ependymal outgrowth produces fibronectin in both the axolotl26 and in regeneration-competent Xenopus tadpoles (E. Chernoff et al., unpublished). Tenascin and polysialylated N-CAM expression in regenerating urodele spinal cord is like that in developing animals with expression around ependymal cells as well as in axonal tracts.47, 48

in in vitro model systems, such as neurosphere culture. Removal of these growth factors and addition of neurotrophic factors stimulates neuronal or glial differentiation.50, 51 Adult axolotl spinal cord ependymal cells require EGF to divide in culture.52 In P. waltl tail cord regeneration, ependymal cells both produce and respond to FGF-2 as they produce new neurons during regeneration.27 Preliminary experiments with Xenopus tadpole ependymal cells in culture show cell proliferation and formation of ‘neurospheres’ reminiscent of mammalian neural stem cell behavior (E. Chernoff et al., unpublished). The Xenopus tadpole ependymal cells also require EGF to proliferate. In vivo, regeneration-competent and regenerating Xenopus spinal cord ependymal cells express nrp-1, the Xenopus Musashi-1 homolog (Figure 1).35, 53 Musashi-1 is an RNA-binding protein first identified in Drosophila for its involvement in the maintenance of Notch-1 signaling.54 Mouse-Musashi-1 is highly enriched in the CNS stem cells and increases Notch-1 signal activity by posttranscriptional downregulation of Numb.53, 55 One downstream effect of Notch-1 signaling is activation of E(spl)-C/Hes genes (enhancer of split-hairy) which repress expression of proneural genes, such as Mash and neurogenin, and maintain progenitor cell properties.56–58 Expression of these genes is being evaluated in amphibian spinal cord. One difference in amphibian spinal cord is that the apparent neural stem cells are ventricular rather than subventricular as in most mammalian experimental systems.59, 60

Patterning The positional cues required for axonal regrowth to appropriate targets in mouse cord regeneration studies may differ depending on the model system. In the regenerating mouse cord example, distances are small and only short-range axonal migration cues may be required. Glial bridges and channels and surrounding axons could support or provide Ephrin/Eph interactions, CAMS, cadherin, extracellular matrix components, and neurotrophic factor influences. In more extensive rodent cord injury models, successful axonal regrowth across implanted bridge materials may not result in appropriate synaptic connections.61 In these situations, necessary patterning or positional information may be lacking. For complete regeneration it may be necessary to re-express A–P and D–V axis cues active during embryonic development. These patterning systems may be active in amphibian cord regeneration.

Stem cell properties Mitotic labeling of the CNS of the newt P. waltl during regeneration shows that the ependymal layer proliferates during regeneration. Labeling regenerating tail cord with specific antibodies or lineage probes shows that the ependyma generate not only glia, but neurons and melanocytes, making multiple cell types produced by neuroepithelium during embryogenesis.28, 49 Are the new neurons and glia formed in amphibian spinal cord regeneration produced by neural stem cells as defined in mammalian systems? Mammalian neural subventricular or ventricular zone cells require fibroblast growth factor (FGF) alone or EGF and FGF to maintaining progenitor cell properties 364


Figure 1. Expression of nrp-1 (Musashi-1) in the Xenopus laevis tadpole spinal cord before and during metamorphosis. (A) Cross-section at stage 50. Nrp-1 is strongly expressed in the ependymal cells (ventricular zone) when the spinal cord is regeneration-competent. (B) Cross-section at stage 62. During metamorphosis, in the regeneration-incompetent stage, the ependymal cells show weak expression of nrp-1 (Chernoff laboratory, unpublished).

Since the embryonic sources of signaling molecules and the original progenitor cells populations involved in A–P or D–V patterning are no longer present or active, the source of signals and the mechanism of activation may differ. The cranial and caudal stump tissue flanking the lesion is a strong candidate for the source of patterning information in the regenerate.

A–P patterning of the embryonic CNS starts with additive expression of Hox-a and Hox-b cluster genes and progresses to establishing the identities of subgroups of spinal motor neurons.62 No correlation has been made between A–P patterning in regeneration and the early steps of embryonic A–P patterning, but patterning genes are expressed. Studies of caudal 365


regeneration of urodele spinal cord show that two Nkx3-related genes establish positional information along the A–P axis.63 A distal-less gene dlx3 (Pwdlx3) is expressed in the lateral ependymal tube in association with remaking the DRG. This gene is not expressed in uninjured adult tissue and may not be part of embryonic production of DRG.64 D–V patterning of the embryonic spinal cord is also a complex process involving many different pathways and transcription factors. At the top of the patterning cascade, Sonic Hedgehog (Shh) is key in the ventralizing process while the Bone Morphogenetic Proteins (BMPs) are important for dorsalization.62, 65 Pax genes are important in regionalization of the spinal cord. Class I and II transcription factors are involved in establishing neuronal identity.66–68 Investigation of the D–V patterning process in amphibian spinal cord regeneration is in its infancy. In the intact regeneration-competent Xenopus spinal cord Shh and nrp-1 are coexpressed in the floorplate. In regenerating Xenopus spinal cord, Shh is expressed ventrally in the mesenchymal outgrowth but in a location complementary to nrp-1/Musashi-1 expression, suggesting differences in regulation of D–V patterning of the regenerating and embryonic spinal cord.37 In the field of CNS regeneration, the spontaneously regenerating animal model systems have been relatively neglected. Elucidation of signaling systems, transcription factors, and transcription factor targets will provide tools such as proteins of therapeutic value and pathways that can be pharmacologically manipulated. When understood, the mechanisms used in animals intrinsically able to regenerate their spinal cord will provide new therapeutic approaches to human spinal cord regeneration.

2. Caroni P, Schwab ME (1988) Antibody against myelin-associated inhibitor of neurite growth neutralizes nonpermissive substrate properties of CNS white matter. Neuron 1:85–96 3. Keirstead HS, Hasan SJ, Muir GD, Steeves JD (1992) Suppression of the onset of myelination extends the permissive period for the functional repair of embryonic spinal cord. Proc Natl Acad Sci USA 89:11664–11668 4. Kwon BK, Tetzlaff W (2001) Spinal cord regeneration: from gene to transplants. Spine 26(Suppl 24):S13–S22 5. Chernoff EAG (1996) Spinal cord regeneration: a phenomenon unique to urodeles? Int J Dev Biol 40:823–831 6. Clarke JDW, Ferretti P (1998) CNS regeneration in lower vertebrates, in Cellular and Molecular Basis of Regeneration (Ferretti P, Geraudie J, eds) pp. 255–269. Wiley, New York 7. Seitz A, Aglow E, Heber-Katz E (2002) Recovery from spinal cord injury: a new transection model in the C57Bl/6 mouse. J Neurosci Res 67:337–345 8. Anderson MJ, Waxman SG (1983) Caudal spinal cord of the teleost Sternarchus albifrons resembles regenerating cord. Anat Rec 205:85–92 9. Anderson MJ, Choy CY, Waxman SG (1986) Selforganization of ependyma in regenerating teleost spinal cord: evidence from serial section reconstructions. J Embryol Exp Morphol 96:1– 18 10. Yamada H, Miyake T, Kitamura T (1997) Proliferation and differentiation of ependymal cells after transection of the carp spinal cord. Zool Sci 14:331–338 11. Egar M, Singer M (1972) The role of ependyma in spinal cord regeneration in the urodele, Triturus. Exp Neurol 37:422– 430 12. Singer M, Nordlander RH, Egar M (1979) Axonal guidance during embryogenesis and regeneration in the spinal cord of newt: the blueprint hypothesis of neuronal pathway patterning. J Comp Neurol 185:1–22 13. Stensaas LJ (1983) Regeneration in the spinal cord of the newt Notopthalmus (Triturus) pyrrhogaster, in Spinal Cord Reconstruction. (Kao CC, Bunge RP, Reier PJ, eds) pp. 121–149. Raven Press, New York 14. Egar M, Simpson SB, Singer M (1970) The growth and differentiation of the regenerating spinal cord of the lizard, Anolis carolinensis. J Morphol 131:131–152 15. Simpson SB Jr (1968) Morphology of the regenerated spinal cord in the lizard, Anolis carolinensis. J Comp Neurol 134:193– 210 16. Duffy MT, Liebich DR, Garner LK, Hawrych A, Simpson SB Jr, Davis BM (1992) Axonal sprouting and frank regeneration in the lizard tail spinal cord: correlation between changes in synaptic circuitry and axonal growth. J Comp Neurol 316:363– 374 17. Alibardi L, Meyer-Rochow VB (1988) Ultrastructure of the neural component of the regenerating spinal cord in the tails of three species of New Zealand lizards. New Zealand J Zool 15:535–550 18. Forehand CJ, Farel PB (1982) Anatomical and behavioral recovery from the effects of spinal cord transection: dependence on metamorphosis in anuran larvae. J Neurosci 2:654–662 19. Clarke JD, Tonge DA, Holder NH (1986) Stage-dependent restoration of sensory dorsal columns following spinal cord transection in anuran tadpoles. Proc R Soc Lond B Biol Sci 227:67– 82 20. Holder N, Clarke JDW, Wilson S, Hunter K, Tonge DA (1989) Mechanisms controlling directed axon regeneration in the peripheral and central nervous systems of amphibians, in NATO Advanced Research Workshop on Recent Trends in

Acknowledgements Part of the work presented here and authors were supported by NSF Grant EHR-0093092, Eli Lilly and Company, and the Indiana 21st Century Research and Technology Fund. Axolotls were supplied by the Indiana University Axolotl Colony.

References 1. Steward O, Schauwecker PE, Guth L, Zhang Z, Fujika M, Inman D, Wrathall J, Kemperman G, Gage F, Saatman KE, Raghupathi R, McIntosh T (1999) Genetic approaches to neurotrauma research: opportunities and potential pitfalls of murine models. Exp Neurol 157:19–42

366


21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

Regeneration Research (1988: Athens, Greece) (Kiorstsis V, Koussoulakos S, Wallace H, eds) pp. 179–190. Plenum Press, New York Beattie MS, Bresnahan JC, Lopate G (1990) Metamorphosis alters the response to spinal cord transection in Xenopus laevis frogs. J Neurobiol 21:1108–1122 Shimizu I, Oppenheim RW, O’Brien M, Shneiderman A (1990) Anatomical and functional recovery following spinal cord transection in the chick embryo. J Neurobiol 21:918–937 Hasan SJ, Keirstead HS, Muir GD, Steeves JD (1993) Axonal regeneration contributes to repair of injured brainstem-spinal neurons in embryonic chick. J Neurosci 13:492–507 Iwashita Y, Kawaguchi S, Murata M (1994) Restoration of function by replacement of spinal cord segments in the rat. Nature 367:167–170 Anderson MJ, Rossetto DL, Lorenz LA (1994) Neuronal differentiation in vitro from precursor cells of regenerating spinal cord of the adult teleost Apteronotus albifrons. Cell Tissue Res 278:243–248 O’Hara CM, Egar MW, Chernoff EAG (1992) Reorganization of the ependyma during axolotl spinal cord regeneration: changes in intermediate filament and fibronectin expression. Dev Dyn 193:103–115 Zhang F, Clarke JDW, Ferretti P (2000) FGF-2 up-regulation and proliferation of neural progenitors in the regenerating amphibian spinal cord in vivo. Dev Bio 225:381–391 Benraiss A, Arsanto JP, Coulon J, Thouveny Y (1999) Neurogenesis during caudal spinal cord regeneration in adult newts. Dev Genes Evol 209:363–369 Holder N, Clarke JDW, Stephens N, Wilson SW, Orsi C, Bloomer T, Tonge DA (1991) Continuous growth of the motor system in the axolotl. J Comp Neurol 303:534–550 Davis BM, Duffy MT, Simpson SB Jr (1989) Bulbospinal and intraspinal connection in normal and regenerated salamander spinal cord. Exp Neurol 103:41–51 Clarke JDW, Alexander R, Holder N (1988) Regeneration of descending axons in the spinal cord of the axolotl. Neurosci Lett 89:1–6 Nordlander RH, Singer M (1978) The role of ependyma in regeneration of the spinal cord of the urodele amphibian tail. J Comp Neurol 180:349–374 Arsanto JP, Komorowski TE, Dupin F, Caubit X, Diano M, Geraudie J, Carlson BM, Thouveny Y (1992) Formation of the peripheral nervous system during tail regeneration in urodele amphibians: ultrastructural and immunohistochemical studies of the origin of the cells. J Exp Zool 264:273– 292 Chernoff EA, Henry LC, Spotts T (1998) An ependymal cell culture system for the study of spinal cord regeneration. Wound Repair Regen 6:403–412 Chernoff EAG, O’Hara CM, Bauerle D, Bowling M (2000) Matrix metalloproteinase production in regenerating axolotl spinal cord. Wound Repair Regen 8:282–291 Egar M, Singer M (1981) The role of ependyma in spinal cord regrowth, in Mechanisms of Growth Control (Becker RO, ed.) pp. 93–106. Charles Thomas Publisher, Springfield, IL Chernoff EAG, Sato K, Smith RC (2001) Nrp-1 (Musashi) and Shh expression in ependymal cells of regenerating and non-regenerating Xenopus spinal cord. Mol Biol Cell 12:366a Dow KE, Ethell DW, Steeves JD, Riopelle RJ (1994) The cellular correlates of spinal cord repair in the embryonic chick: heparan sulphate and chondroitin sulphate proteoglycans. Exp Neurol 128:233–238

39. Reier PJ, Stensaas LJ, Guth L (1983) The astrocytic scar as an impediment to regeneration in the central nervous system, in Spinal Cord Reconstitution (Kao CC, Bunge RP, Reier PJ, eds) pp. 163–195. Raven Press, New York 40. Reier PJ, Houle JD (1988) The glial scar: its bearing on axonal elongation and transplantation approaches to CNS repair. Adv Neurol 47:87–138 41. Pasterkamp RJ, Verhaagen J (2001) Emerging roles for semaphorins in neural regeneration. Brain Res Rev 35:36–54 42. Miranda JD, White LA, Marcillo AE, Willson CA, Jagid J, Whittemore SR (1999) Induction of Eph B3 after spinal cord injury. Exp Neurol 156:218–222 43. Holder N, Clarke JDW (1988) Is there a correlation between continuous neurogenesis and directed axon regeneration in the vertebrate nervous system? TINS 11:94–99 44. Butler EF, Ward MB (1967) Reconstitution of the spinal cord after ablation in adult Triturus. Dev Biol 15:464–486 45. Holder N, Clarke JDW, Kamalati T, Lane EB (1990) Heterogeneity in spinal radial glia demonstrated by intermediate filament expression and HRP labeling. J Neurocytol 19:915–928 46. Miller RH, Liuzzi FJ (1986) Regional specialization of the radial glial cells of the adult frog spinal cord. J Neurocytol 15:187– 196 47. Caubit X, Riou JF, Coulon J, Arsanto JP, Benraiss A, Boucaut JC, Thouveny Y (1994) Tenascin expression in developing, adult and regenerating caudal spinal cord in the urodele amphibians. Int J Dev Biol 38(4):661–672 48. Caubit X, Arsanto JP, Figarella-Branger D, Thouveny Y (1993) Expression of polysialylated neural cell adhesion molecule (PSA-N-CAM) in developing, adult and regenerating caudal spinal cord of the urodele amphibians. Int J Dev Biol 37(2):327– 336 49. Benraiss A, Caubit X, Arsanto JP, Coulon J, Nicolas S, Le Parco Y, Thouveny Y (1996) Clonal cell cultures from adult spinal cord of the amphibian urodele Pleurodeles waltl to study the identity and potentialities of cells during tail regeneration. Dev Dyn 205:135–149 50. Temple S (2001) The development of neural stem cells. Nature 414:112–117 51. Svendsen CN, Caldwell MA (2000) Neural stem cells in the developing central nervous system: implications for cell therapy through transplantation, in Progress in Brain Research (Dunnett SB, Bjorklund A, eds) pp. 13–34. Elsevier, New York 52. O’Hara CM, Chernoff EAG (1994) Growth factor modulation of injury-reactive ependymal cell proliferation and migration. Tissue Cell 26:599–611 53. Imai T, Tokunaga A, Yoshida T, Hashimoto M, Mikoshiba K, Weinmaster G, Nakafuku M, Okano H (2001) The neural RNA-binding protein Musashi1 translationally regulates mammalian numb gene expression by interacting with its mRNA. Mol Cell Biol 21:3888–3900 54. Nakamura M, Okano H, Blendy JA, Montell C (1994) Musashi, a neural RNA-binding protein required for Drosophila adult external sensory organ development. Neuron 13:67–81 55. Sakakibara S, Imai T, Hamaguchi K, Okabe M, Aruga J, Nakajima K, Yasutomi D, Nagata T, Kurihara Y, Uesugi S, Miyata T, Ogawa M, Mikoshiba K, Okano H (1996) Mouse-Musashi-1, a neural RNA-binding protein highly enriched in the mammalian CNS stem cell. Dev Biol 176:230–242 56. Artavanis-Tsakonas S, Rand MD, Lake RJ (1999) Notch signaling: cell fate control and signal integration in development. Science 284:70–76

367


57. Ohtsuka T, Ishibashi M, Gradwohl G, Nakanishi S, Guillemot F, Kageyama R (1999) Hes1 and Hes5 as notch effectors in mammalian neuronal differentiation. EMBO J 18:2196–2207 58. Johansson CB, Momma S, Clarke DL, Risling M, Lendahl U, Frisen J (1999) Identification of a neural stem cell in the adult mammalian central nervous system. Cell 96:25–34 59. Morshead C, van der Kooy D (2001) A new “spin” on neural stem cells? Curr Opin Neurobiol 11:59–65 60. Momma S, Johansson CB, Frisen J (2000) Get to know your stem cells. Curr Opin Neurobiol 10:45–49 61. Bamber NI, Li H, Lu X, Oudega M, Aebischer P, Xu XM (2001) Neurotrophins BDNF and NT-3 promote axonal re-entry into the distal host spinal cord through Schwann cell-seeded mini-channels. Eur J Neurosci 13:257–268 62. Altmann CR, Brivanlou AH (2001) Neural patterning in the vertebrate embryo. Int Rev Cytol 203:447–482 63. Nicolas S, Caubit X, Massacrier A, Cau P, Le Parco Y (1999) Two Nkx-3-related genes are expressed in the adult and regenerating

64.

65.

66. 67.

68.

368

central nervous system of the urodele Pleurodeles waltl. Dev Genet 24:319–328 Nicolas S, Massacrier A, Caubit X, Cau P, Le Parco Y (1996) A distal-less-like gene is induced in the regenerating central nervous system of the urodele Pleurodeles waltl. Mech Dev 56:209– 220 Eggenschwiler JT, Espinoza E, Anderson KV (2001) Rab23 is an essential negative regulator of the mouse Sonic hedgehog pathway. Nature 412:194–198 Patten I, Placzek M (2000) The role of sonic hedgehog in neural tube patterning. Cell Mol Life Sci 57:1695–1708 Briscoe J, Ericson J (1999) The specification of neuronal identity by graded sonic hedgehog signaling. Semin Cell Dev Biol 10:353–362 Briscoe J, Pierani A, Jessell TM, Ericson J (2000) A homeodomain protein code specifies progenitor cell identity and neuronal fate in the ventral neural tube. Cell 101:435– 445