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DEVELOPMENTAL DYNAMICS 226:295–307, 2003
REVIEWS
A PEER REVIEWED FORUM
Urodele Spinal Cord Regeneration and
Related Processes
Ellen A.G. Chernoff,1* David L. Stocum,1 Holly L.D. Nye,2 and Jo Ann Cameron2
Urodele amphibians, newts and salamanders, can regenerate lesioned spinal cord at any stage of the life cycle and
are the only tetrapod vertebrates that regenerate spinal cord completely as adults. The ependymal cells play a key
role in this process in both gap replacement and caudal regeneration. The ependymal response helps to produce a
different response to neural injury compared with mammalian neural injury. The regenerating urodele cord produces
new neurons as well as supporting axonal regrowth. It is not yet clear to what extent urodele spinal cord regeneration
recapitulates embryonic anteroposterior and dorsoventral patterning gene expression to achieve functional
reconstruction. The source of axial patterning signals in regeneration would be substantially different from those in
developing tissue, perhaps with signals propagated from the stump tissue. Examination of the effects of fibroblast
growth factor and epidermal growth factor on ependymal cells in vivo and in vitro suggest a connection with neural
stem cell behavior as described in developing and mature mammalian central nervous system. This review
coordinates the urodele regeneration literature with axial patterning, stem cell, and neural injury literature from other
systems to describe our current understanding and assess the gaps in our knowledge about urodele spinal cord
regeneration. Developmental Dynamics 226:295–307, 2003. © 2003 Wiley-Liss, Inc.
Key words: axolotl; Ambystoma mexicanum; urodele; spinal cord development; regeneration; spinal cord regeneration; spinal
cord; neurulation; ependymal cells; ependymoglia; radial glia; axial pattern formation; newt; salamander
Received 21 October 2002; Accepted 5 November 2002
INTRODUCTION
Anuran amphibians, reptiles, birds,
and mammals can only regenerate
spinal cord as larvae or embryos.
The anuran amphibians and higher
vertebrates can regenerate limited
regions of the spinal cord spontaneously or can regenerate after specific types of lesions (reviewed in
Chernoff et al., 2002). Urodele amphibian spinal cord regeneration is a
unique experimental system in that
urodeles are the only tetrapod vertebrates that are strong regenera-
tors in adulthood. Retention of embryonic character is cited as a
property that supports this regeneration process, but it is not clear to
what extent embryonic processes involved in central nervous system
(CNS) development must be retained or re-expressed. Regenerating urodele spinal cord clearly does
not recapitulate the early events of
neurulation. However, the example
of limb regeneration suggests that
some embryonic patterning and differentiative processes would be re-
quired to reconstruct spinal cord
structure and restore function completely (Stocum, 1996; Brockes, 1997;
Chistensen et al., 2002).
Caudal regeneration studies have
shown that urodele ependymal cells
produce neuronal cells (Egar and
Singer, 1972; Nordlander and Singer,
1978; Arsanto et al., 1992; Benraiss et
al., 1996; Zhang et al., 2000). The
modulation of urodele ependymal
cell proliferation and differentiation
by fibroblast growth factor-2 (FGF-2)
and epidermal growth factor (EGF)
1
Indiana University-Purdue University Indianapolis, Department of Biology, and Indiana University Center for Regenerative Biology and
Medicine, Indianapolis, Indiana
2
University of Illinois Department of Cell and Structural Biology and College of Medicine, B107 Chemical and Life Sciences Laboratory,
Urbana, Illinois
Grant sponsor: NSF; Grant number: PFI EHR-0093092; Grant sponsor: Indiana 21st Century Research and Technology Fund; Grant number:
031500-0064.
*Correspondence to: Ellen Chernoff, Indiana University-Purdue University Indianapolis, Department of Biology, and Indiana University
Center for Regenerative Biology and Medicine, 723 W. Michigan St., Indianapolis, IN 46202. E-mail: [email protected]
DOI 10.1002/dvdy.10240
© 2003 Wiley-Liss, Inc.
296 CHERNOFF ET AL.
demonstrates properties reminiscent
of mammalian neural stem cells
(O’Hara and Chernoff, 1994; Zhang
et al., 2000; Temple, 2001). It is possible that, instead of retention of embryonic properties per se, regeneration capacity reflects expression of
neural stem cell properties in specialized cell populations.
The higher vertebrate neural injury
literature has defined inhibitory
pathways and cellular responses
(Fawcett and Asher, 1999; Kwon
and Tetzlaff, 2001; McDonald and
Sadowsky, 2002). These processes include excitotoxicity due to excessive calcium transport and mobilization, glial scars formed by astrocytes
or meningeal cells, and toxic myelin
breakdown products. The corresponding processes and cell behavior in urodele systems will be discussed and compared. In this review,
we will integrate the information
available from several urodele species and several areas of research to
provide a picture of the literature
that forms the background for current urodele spinal cord regeneration research.
URODELE INJURY MODELS
Several parameters must be considered when using a urodele spinal
cord injury and regeneration model
system. Choices must be made regarding phase of life cycle, neotenic or metamorphosing species,
region of the spinal cord lesioned,
and type of lesion. The spinal cord
regenerates in embryonic, larval, juvenile, and adult urodeles. The regeneration that occurs in embryonic
or larval animals involves the formation of substantial numbers of new
neurons and is qualitatively different
from adult regeneration (Detwiler,
1947; Holtzer, 1951, 1952; Butler and
Ward, 1965, 1967; Stensaas, 1983;
Davis et al., 1989). The time course of
regeneration is also more rapid in
larval and young juvenile urodeles
than in adults (Piatt, 1955). Different
aspects of spinal cord regeneration
can be examined at different points
in the life cycle, emphasizing neurogenesis or axonal regrowth.
The choice between metamorphosing and neotenic animals can
be a compromise among factors
such as ease of husbandry, animal
availability, and biological differences. It has been argued that the
persistent regenerative capacity of
the adult axolotl (Ambystoma mexicanum) derives from its neotenic
state and continuous growth, leading to retention of embryonic/immature tissue properties (Holder and
Clarke, 1988; Holder et al., 1991).
Neoteny does not necessarily decrease the desirability of a spinal
cord model system because metamorphosing amphibians undergo
extensive neural changes that affect spinal cord regeneration and
complicate studies of properties permissive to neural regeneration (Shi,
2000). Adult neotenic urodeles produce a strong regeneration response from a predominantly stable,
fully differentiated, fully functional
spinal cord. The use of axolotl specifically has the advantage of using a
captive-bred, purpose-bred animal,
relatively extensive spinal cord regeneration literature, and an emerging
molecular biology database (Indiana
University Axolotl Colony: http://www.
indiana.edu/⬃axolotl/; S. Randal
Voss, Salamander Genome Project:
http://www.lamar.colostate.edu/
⬃srvoss/SGP/ (Note: Web site will be
moving to the University of Kentucky)).
The progress of regeneration is
easiest to follow in complete transections. In transection models the
relationship of the regenerating axons with ependymal cell processes
and the pia mater is very clear (Egar
and Singer, 1981). In suction wounds,
the meninges and pia mater can be
spared, resulting in axonal regrowth
along the inner surface of the pia
(Stensaas, 1983). Although widely
used in mammalian injury models,
crush injuries are not widely used in
urodele regeneration studies (Steward et al., 1999). Crushed cord undergoes a more extensive period
of cell death, debris removal, and
tissue remodeling than cleanly transected cord (personal observation).
Recovery of locomotion has been
demonstrated after transection of
brachial, thoracic, lumbar, and tail
spinal cord (Steffaneli, 1950; Holtzer,
1951, 1952; Piatt, 1955; Butler and
Ward, 1967; Nordlander and Singer,
1978; Stensaas, 1983; Clarke et al.,
1988; Davis et al., 1989; O’Hara et al.,
1992). Most of these studies use axonal regrowth as the measure of regeneration. Some axonal regrowth
occurs in axolotls as early as 2 weeks
after transection (reviewed in Stensaas, 1983). In a systematic study of
the restoration of normal swimming
behavior after thoracic transection
lesions in adult Eastern Spotted
Newts (Notaphthalmus viridescens),
the earliest recovery of coordinated
swimming was 4 weeks after lesioning, with 50% of the animals showing
swimming with deficits at 6 weeks
and fully coordinated swimming at 8
weeks (Davis et al., 1990). Retrograde labeling studies show a strict
correlation of coordinated swimming with regrowth of the descending supraspinal axons to the level of
the lumbar enlargement (Davis et
al., 1990). Experiments following regeneration for periods of 6 months or
less routinely report that the spinal
cord does not return to its original
thickness and that some urodele spinal cord tracts do not regenerate
(Piatt, 1955; Stensaas, 1983; Holder et
al., 1989). However, the time required for complete replacement of
cord structures can be very long,
e.g., 23 months in adult axolotls
(Clarke et al., 1988).
Variation in injury type and other
experimental parameters can make
it difficult to compare the results of
different laboratories. For example,
experiments that describe failure of
regeneration in adult urodeles may
have produced gross lateral displacement of the cord due to severing of the vertebral column. This
type of disruption prevents regenerating ends of the cord from meeting
(Butler and Ward, 1967). The use of
any one injury type may be limited
to a single laboratory and performed at a time when no biochemical or molecular analysis was
possible. This finding makes direct
comparison of current experiments
with some of the older literature difficult. Instead of cataloguing differences and aberrations in the
literature, we will concentrate on
processes and mechanisms that are
significant in the context of current
knowledge.
URODELE SPINAL CORD REGENERATION 297
EMBRYONIC PROPERTIES OF THE
URODELE SPINAL CORD
Urodele neural development can
be divided into three steps: neuralization, the induction of the neural
plate; neurulation, the formation
and closure of the neural tube; and
neural differentiation, the formation
of neurons and glia (reviewed in Duprat, 1996). Not all of these processes are relevant to urodele spinal
cord regeneration: the neural plate
is not reconstructed, and the regenerated spinal cord does not develop
by the folding of a neural plate, but
there are mechanisms involved in
developmental patterning and differentiation that may be relevant in
regeneration competence and tissue reconstruction.
Is there any recapitulation of neurulation events in urodele spinal cord
regeneration? Traditionally neural
tube formation in amphibian embryos has been described as a primary neurulation process (folding,
not cavitation), and this is true for
urodeles (Jacobson, 1981). In some
anurans, a process of fusion and relumination occurs (Davidson and
Keller, 1999). In Xenopus, prospective dorsal neural tube cells undergo
medial cell movement and intercalation that integrates deep and superficial cells to form the roof of the
neural tube. These rearrangements
are followed by relumination of the
neural tube, a process that shares
some features with secondary neurulation processes in amniotes (Davidson and Keller, 1999). A relumination-like process could be part of
regeneration in urodele cord.
The reconstructed spinal cord innervates muscles and organs along
the anteroposterior (A-P) axis. Ascending and descending axons
must be targeted appropriately
along the dorsoventral (D-V) axis,
and the various types of spinal neurons must connect with each other.
This finding suggests that some aspects of embryonic A-P and D-V
patterning might be re-expressed.
The aspects of spinal cord development that may help in understanding regenerative mechanisms are
discussed here in conjunction with
the corresponding regenerative process.
What are the residual embryonic
qualities of the adult axolotl spinal
cord that could facilitate regeneration? Postembryonic neurogenesis
and the persistence of radial
ependymal or radial glial fibers are
two often-cited properties. Labeling
studies demonstrate that new neurons are produced fairly frequently in
intact axolotls smaller than 7.5 cm
(juveniles 6 –7 months of age; Holder
et al., 1991). In larger/older intact
animals, new spinal cord neurons
are formed infrequently, reflecting
the stability of the mature CNS tissue
(Holder et al., 1991). The axolotl
ependymal cell spans the spinal
cord from lumen to basal lamina, as
do the embryonic radial glia. The radial fibers of the ependymal cells
contain glial fibrillary acidic protein
(GFAP) in the outer (basal) segment,
a region that starts in the outer gray
matter and spans the white matter,
whereas the cytoplasm of the cell
body contains intermediate filaments composed of cytokeratins
(Holder et al., 1990). However, the
presence of transepithelial radial
ependymal (radial glial) processes
also occurs in the spinal cord of
adult frogs, animals that do not regenerate their spinal cords; and frog
ependymal cell processes also express GFAP in the outer segment of
the process (Miller and Liuzzi, 1986).
There is probably no correlation between intermediate filament content of the ependyma and regeneration capacity, as phylogenetic
studies have shown no systematic
differences that would correlate
with success or failure of regeneration (Bodega et al., 1994; Clarke and
Ferretti, 1998). So, transepithelial extension and regionalization of intermediate filament proteins are not
sufficient indicators of spinal cord regeneration capacity.
SPINAL CORD STRUCTURE
The mature urodele spinal cord consists of an ependymal zone lining the
central canal, gray matter, and
white matter. The boundary between white and gray matter is not
as distinct as in mammals. Within the
gray matter, the ventral horns can
be distinguished, but a dorsal horn
region is not well-developed. As in all
amphibians, the ependymal cells
maintain radial fibers (ependymal
processes) with pial endfeet that terminate on, and produce, the glia
limitans (basal extracellular matrix
surrounding the cord; Roots, 1986;
Miller and Liuzzi, 1986; Holder et al.,
1989). Thick layers of fibrillar collagen
(pia–arachnoid matrix) produced
by the leptomeningeal cells (neural
crest origin) surround the basal lamina. Numerous blood vessels terminate on the surface of the spinal
cord. The ependymal processes and
endfeet form the blood/spinal cord
barrier. In pigmented animals, melanocytes are found on the meningeal
collagen.
The developing urodele spinal
cord contains nine groups of spinal
cord neurons: (1) dorsal RohanBeard primary sensory neurons, (2)
giant dorsolateral commissural interneurons, (3) multipolar dorsolateral
commissural interneurons, (4) dorsolateral ascending interneurons, (5)
multipolar dorsolateral ascending interneurons, (6) unipolar mid-cord
commissural neurons with glycinelike immunoreactivity, (7) bipolar or
multipolar descending interneurons,
(8) ventrolateral motor neurons, and
(9) ␥-aminobutyric acid–like KolmerAgduhr cerebrospinal fluid-contacting neurons (Harper and Roberts,
1993; Roberts, 2000). These nine
groups of neurons are identifiable in
embryos of the smooth newt, Triturus
vulgaris, at a stage that can swim
when removed from their egg membranes (stages 32– 48, Harper and
Roberts, 1993; Gallien and Bidaud,
1959). The giant dorsolateral commissural neurons are also present in
axolotl embryos from stage 39 on
and in other Ambystomatid salamanders (Bordzilovskaya et al., 1989;
Harper and Roberts, 1993; Roberts,
2000). A significant gap in urodele
cord regeneration studies is analysis
of the fate of each of these types of
neurons.
SPINAL CORD REMODELING
AFTER INJURY
Regeneration of the urodele spinal
cord has been examined in tail amputation and lesions of the cervical,
thoracic, and lumbar regions, although not, unfortunately, all in the
298 CHERNOFF ET AL.
Fig. 1. Spinal cord regeneration: gap replacement. A: Cartoon of gap replacement in the urodele spinal cord. (1) A complete
transection results in retraction of the stumps. (2) Mesenchymal ependymal outgrowth from cranial and caudal stumps fills the lesion site.
(3) The ependymal cells reepithelialize, and axons regrow through ependymal cell processes. B: Early in the gap replacement process
in adult axolotl spinal cord, 2 weeks of regeneration, ependymal cells grow out as a solid mesenchymal mass. Shown in cross-section.
C: Cranial to the lesion site, the ependymal cells remain in epithelial form. Mitotic cells are prominent. Voids are visible in the damaged
tissue. D: At a later stage in the regeneration process, 4 weeks after lesioning, white and gray matter are reconstituted. Small voids are still
present in the tissue. Images taken from material supplied by M.W. Egar.
same species (reviewed in Piatt,
1955; Davis et al., 1989). The most
significant differences are those between regeneration of the spinal
cord in amputated tail (caudal regeneration) and in spinal cord
transected more cranially (gap replacement). The type of tissue remodeling that occurs and the inductive effects of the spinal cord on
surrounding tissue differ (Piatt, 1955;
Egar and Singer, 1972; Stensaas,
1983). Figure 1 shows a diagrammatic representation of the gap replacement process and representative sections through regenerating
adult axolotl cord.
Tail regeneration is much more
complicated than body spinal cord
regeneration in the sense that complex non-neural structures must also
be replaced. Contact between the
regenerating spinal cord and the tail
stump wound epithelium results in induction of an epimorphic regeneration process similar to limb regeneration (Godlewski, 1928; Efrimov,
1951; Polazhaev, 1972). The wound
epithelium is not necessary to cord
regeneration of the tail spinal cord
but plays a role in the regeneration
of muscle and connective tissue. Tail
regeneration is a bit like regenerating a limb with a spinal column in it
instead of long bones and autopodia. The response of each of the
principal spinal cord cell types in the
regeneration process will be considered.
Ependymoglia
One feature common to the animals
that regenerate their spinal cord
(fish, urodele amphibians, young
anuran tadpoles, lizard tails, fetal
birds, and mammals) is an ependymal response (described below) after lesioning (reviewed in Chernoff,
1996; Clarke and Ferretti, 1998). The
mechanism that initiates this response is unknown, and many of the
contributions of these ependymal
URODELE SPINAL CORD REGENERATION 299
cells to the regeneration process are
still under investigation.
The tissue reorganization involved
in the ependymal response has
been described in detail. During
caudal regrowth in the regenerating
tail, ependymal cells form a hollow
terminal vesicle continuous with the
central canal of the stump cord. The
ependymal outgrowth occurs in the
form of a tubular extension that does
not lose its apical/basal polarity. The
epithelial intercellular junctional
complexes and basal lamina are
maintained (Egar and Singer, 1972,
1981; Clarke and Ferretti, 1998). Cell
proliferation is associated with the
ependymal response (see section
on Mitotic Capacity In Vivo). An
ependymal tube extends as the
cells continue to proliferate.
The remodeling of lesioned body
spinal cord (above the cauda
equina) also starts with an ependymal response. After nontail spinal
cord is transected, the ependymal
cells rearrange to seal the cut ends
of the spinal cord, forming an
ependymal bulb. The ependymal
cells migrate into the lesion site from
both cranial and caudal stumps during the process of gap replacement
(Butler and Ward, 1967; Singer et al.,
1979; Stensaas, 1983). Early in the regeneration process, the cell population within the lesion site consists of
ependymal cells and infiltrating
macrophages. Cells within the
ependymal outgrowth proliferate,
migrate, and remove existing extracellular matrix material and debris
from dead cells (Simpson, 1968; Egar
and Singer, 1972; Singer et al., 1979;
Stensaas, 1983; Anderson et al.,
1986; Chernoff et al., 1990, 2000).
In gap replacement, ependymal
outgrowth involves reorganization of
the epithelial ependymal cells into a
mesenchyme with characteristic
changes in cell– cell adhesive junctions, intermediate filaments, and
extracellular matrix components
(Butler and Ward, 1967; Egar and
Singer, 1972; O’Hara et al., 1992). This
outgrowth is in the form of a solid
mass lacking apical/basal polarity.
In adult axolotls (⬎13 cm), there is
extensive outgrowth from cranial
and caudal stumps at 2 weeks after
lesioning. The mesenchymal ependymal outgrowths meet and join at
approximately 3– 4 weeks to form an
ependymal bridge. The ependymal
bridge reorganizes from a solid cord
of cells into an epithelial tube at 4 –5
weeks (O’Hara et al., 1992; Fig. 1).
There are differences in individuals
based in part on the extent to which
the spinal cord retracts when cut,
and the amount of trauma to the
cut ends. Neuropil and newly myelinated axons can be seen in the
ependymal bridges of some animals
as early as 4 weeks of regeneration
(O’Hara et al., 1992). The axons and
dendrites grow in the spaces between ependymal endfeet or between ependymal processes in
close contact with the basal lamina
(Egar and Singer, 1981). Studies that
rigorously examine restoration of coordinated movement are relatively
rare. There are two reasons for this.
Either lesions are made caudally to
avoid paralyzing the bladder and
thus minimizing motor deficits or axonal regrowth is monitored in lieu of
scoring coordinated movement.
The characterization of the disorganization of ependymal cells in
gap replacement/body cord as an
epithelial/mesenchymal transformation is based on characteristics
found in other epithelial–mesenchymal transformations in vitro and in
embryonic development (Hay and
Zuk, 1995). E-cadherin is lost, matrix
metalloproteinases are secreted,
and basal lamina is removed
(O’Hara et al., 1992; Chernoff et al.,
2000; Chernoff, unpublished observations). The apical intercellular
junctional complex is removed with
concomitant loss of the subapical
microfilament bands and loss of desmosome-associated tonofilaments
(O’Hara et al., 1992; Chernoff et al.,
1998).
et al., 1990; O’Hara et al., 1992). During mesenchymal outgrowth in the
earlier stages of gap replacement
regeneration, the ependymal cells
lose their GFAP and epithelial cytokeratins and produce vimentin
(O’Hara et al., 1992). This change in
intermediate filament content with
tissue reorganization is seen in culture as well as in vivo (O’Hara et al.,
1992).
Extracellular matrix and
adhesion molecules.
Ependymal cells in intact axolotl spinal cord produce laminin. During
mesenchymal outgrowth, the laminin is lost and fibronectin is produced (O’Hara et al., 1992). In vitro,
a fibronectin-coated substratum
and the presence of EGF in the culture medium maintain injury-reactive ependymal cells as mesenchyme (Chernoff et al., 1998).
Tenascin is observed around ependymal cells in urodele spinal cord
and in the axonal tracts in developing animals and may play a role in
channeling axonal regeneration. In
the intact adult, the tenascin level
remains high in the axonal tracts but
is low in the ependymal cells. The
tenascin distribution in regenerating
animals is like that in developing animals (Caubit et al., 1994). The neural
cell adhesion molecule (N-CAM)
shows an interesting distribution in
the urodele spinal cord. During development, the polysialylated form
of N-CAM (PSA-N-CAM) is present
around the ependymal cells and axonal tracts. N-CAM is also present in
the adult, but PSA-N-CAM is only detected in the ependymal zone. PSAN-CAM is up-regulated in the regenerating spinal cord (Caubit et al.,
1993).
Intermediate filament content.
The intermediate filament composition in ependymal cells of intact
adult or juvenile axolotl spinal cord
shows the regional localization characteristic of all amphibian ependymoglia. The outer (basal) segment of
the ependymal process contains
GFAP. The cytoplasm of the juxtaluminal cell body contains epithelial
cytokeratins 8 and 18, simple epithelial cytokeratins, and also reacts with
antibodies to tonofilaments (Holder
Matrix-degrading enzymes.
During spinal cord regeneration in
the axolotl, matrix remodeling includes removal of the basal lamina
(glia limitans) early in ependymal
outgrowth, and remodeling of a
substantial amount of fibrillar collagen in the meninges. Matrix remodeling involves the action of matrixdegrading enzymes such as the
matrix metalloproteinases (MMPs).
MMPs are characterized by a zinc-
300 CHERNOFF ET AL.
binding region required for protease
activity and calcium-binding required
for substrate binding (Birkedal-Hansen
et al., 1993; Nagase and Woessner,
1999). They are found in zymogen
(granule storage) form within cells
and are proteolytically processed
and secreted. MMPs and related enzymes are active in a variety of normal and abnormal processes in the
nervous system (Yong et al., 2001).
Intact adult axolotl spinal cord does
not produce levels of MMPs detectable by zymography (Chernoff et
al., 2000). Injury-reactive ependymal
cells, however, contribute to the
remodeling of lesioned urodele spinal cord through MMP production.
Ependymal outgrowth in vivo and
mesenchymal ependymal cells in
vitro produce MMP-1 (interstitial collagenase), MMP-2, and MMP-9 (gelatinase a and b). The amount of
MMP activity is high at 2 and 4 weeks
after lesioning and declines by 8
weeks of regeneration. The endogenous MMP inhibitor, tissue inhibitor of
metalloproteinase-1 (TIMP-1), is detected in the intact spinal cord but is
absent during mesenchymal ependymal outgrowth (when cells have
deepithelialized and are dividing
and migrating; Chernoff et al., 2000).
The action of other matrix-degrading enzymes has not been analyzed
in the urodele spinal cord.
Mitotic capacity in vivo.
In tail cord regeneration, it is generally agreed that ependymal cells
possess a high level of mitosis. Based
on distribution of mitotic figures, the
ependymal cells are highly proliferative just cranial to the amputation
site, more so than in the ependymal
bulb (Egar and Singer, 1972; Nordlander and Singer, 1978). Proliferating cell nuclear antigen (PCNA) labeling in Pleurodeles waltl (the
Spanish ribbed newt) at early postamputation stages shows that most
of the ependyma cells near the amputation plane are labeled. Lower
numbers are labeled within the terminal vesicle (Zhang et al., 2000).
Later, new neurons are detected by
using a beta-3 tubulin probe, and
ependymal cells are still proliferating
at this time. Few PCNA-labeled cells
exist in intact P. waltl spinal cord
(Zhang et al., 2000). By using bro-
modeoxyuridine (BrdU) labeling,
Benraiss et al. (1999) reported a high
level of mitosis in ependymal cells
inside of and rostral to the terminal
bulb. Within the regenerating
ependymal tube, there does not appear to be a defined growth zone;
mitotic figures are distributed along
the tube’s length, and new neurons
differentiate (Clarke and Ferretti,
1998; Benraiss et al., 1999). Directly
comparable studies in other urodele
species and in nontail spinal cord
regeneration have not been published.
Astrocytes
The identification of GFAP as a fibrous astrocyte marker in vertebrate
produces some confusion in interpretation of glial differentiation in
amphibians (Bignami et al., 1972;
Holder et al., 1990; Soula et al., 1990;
Cochard et al., 1995). In the developing amphibian spinal cord, the radial glia and, later, the ependymal
cells span the neural epithelium. As
described earlier, these cells contain
GFAP in the basal (outer) portion of
the cell processes, with cytokeratin
comprising the intermediate filaments within the cell body (Miller
and Liuzzi, 1986; Holder et al., 1990).
The radial glia/ependymoglia in amphibians have been identified as astrocytes in several studies (e.g., Cochard et al., 1995). Astrocyte
differentiation has been described
as precocious in amphibians because of the presence of GFAP in
radial glia (Pleurodeles waltl: Soula
et al., 1990; Xenopus: Szaro and
Gainer, 1988). There is disagreement
on the existence of true astrocytes in
adult amphibian CNS. Some laboratories have identified cells, present in
low numbers, that express only GFAP
in their intermediate filaments, but it
is clear that most of the GFAP in the
intact axolotl spinal cord is in the
outer segments of radial ependymal
fibers (Zamora and Mutin, 1988;
Holder et al., 1990; O’Hara et al.,
1992). Electron microscopic examination has suggested that there are
“astrocytes” that lack radial processes (Schonbach, 1969; Egar and
Singer, 1972). It remains to be determined whether these cells are truly
equivalent to fibrous astrocytes, or
are ependymal cells that have retracted their endfeet from the basal
surface. Although the status of
urodele astrocytes may be ambiguous, it is clear that the lesioned
urodele cord does not produce astrocytic scars formed by GFAP-positive cells, or fibroblastic scars formed
by invading meningeal cells that inhibit axonal regrowth in mammals
(Fawcett and Asher, 1999; Steward
et al., 1999).
Oligodendrocytes
In anuran amphibians, as in other
vertebrates, the notochord is required to induce formation of oligodendrocytes in the ventral spinal
cord during dorsoventral patterning
of the spinal cord (reviewed in Orentas and Miller, 1996; Maier and Miller,
1997). The alteration of white matter
that results from early surgical ablation of the notochord in salamanders suggests that urodele oligodendrocyte formation is similar (Triturus,
formerly Triton, Lehmann, 1928; axolotl, Kitchin, 1949). Oligodendrocytes
have been described in the periependymal stratum of the mature
urodele spinal cord and also more
basally (P. waltl, Zamora, 1978).
Oligodendrocytes can affect the
regeneration process positively by
remyelinating new axons or negatively through the production of
toxic or inhibitory myelin products after injury. In mammals, myelin components such as myelin-associated
glycoprotein (MAG) and Nogo
(NI35/250) are known to inhibit regeneration after injury (reviewed in
Ng and Tang, 2002). Other CNS
components known to inhibit axonal
outgrowth in mammals include
chondroitin sulfate proteoglycan
and tenascin-R (Becker et al., 1999;
Ng and Tang, 2002). Nogo (NI35/
250) has not been found in regeneration-competent lower vertebrate,
such as fish and larval Xenopus, but
the other components are present
(Wanner et al., 1995; Lang et al.,
1995; Becker et al., 1999). It has been
suggested that they are rapidly removed after injury. In anuran amphibian (Xenopus laevis), spinal cord
myelin becomes nonpermissive to
axonal outgrowth with metamorphosis and reacts with IN-1, the
URODELE SPINAL CORD REGENERATION 301
Nogo-neutralizing antibody (Lang et
al., 1995). The recent literature on
urodele oligodendroglia is notably
sparse. In salamander optic nerve,
inhibitory components such as MAG
and tenascin-R are present but are
rapidly removed after injury, providing an environment more supportive
to axonal outgrowth (Pleurodeles
waltl; Becker et al., 1999). Specific
information on urodele spinal cord
myelin is absent.
Neurons
In urodele tail regeneration, morphologic studies and markers of
neuronal differentiation show that
new peripheral nervous system (PNS)
and CNS neurons are produced
from the ependymal tube (Notophthalmus viridescens and Pleurodeles
waltl; Egar and Singer, 1972; Nordlander and Singer, 1978; Arsanto et
al., 1992; Benraiss et al., 1999; Zhang
et al., 2000). Neural crest cells are no
longer present, so the regenerated
peripheral nervous system is derived
from either the ventrolateral part of
the regenerated ependymal tube or
dorsal aspect of the terminal vesicle
(Arsanto et al., 1992; Nicolas et al.,
1996; Benraiss et al., 1996). Labeled,
tail regenerate-derived ependymal
cells form cell types normally of neural crest origin in culture as well: melanocytes and Schwann cells (Benraiss et al., 1996).
Pulse-chase experiments using
BrdU show that new CNS neurons
are produced in Pleurodeles waltl
tail cord, in situ. In these studies,
GFAP was used as a glial marker,
and neuron-specific enolase (NSE)
and neurofilament antibodies were
used as neuronal markers. Threeweek regenerates show the proliferative ependyma giving rise to new
neurons (Benraiss et al., 1999). Double labeling of long-term labeled
cells shows that NSE- and GFAP-positive cells both label with BrdU at the
rostral and median levels of the regenerating cord, indicating the formation of new neurons and glia.
Confocal microscopy resolved individual labeled cells, showing that
both glia and neurons are generated from the ependyma. The stronger labeling of the glial and neuronal progeny showed they are not as
mitotic as their ependymal parents
(Benraiss et al., 1999). When BrdUlabeled regenerating cord cells
were cultured, they gave rise to labeled neurons, showing directly that
new neurogenesis was indeed occurring.
Axonal regrowth.
Studies of urodele spinal cord regeneration have reported a range in the
extent of functional recovery. The
number of axons that regenerate,
the number of functional synapses
that occur, and the extent to which
coordinated locomotion is recovered varies (Davis et al., 1989, 1990).
In the newt Notophthalmus viridescens (then Triturus the Eastern redspotted newt), retrograde labeling
shows that all of the regions of the
brainstem that project to the lumbar
spinal cord can regenerate (Davis et
al., 1989). Regeneration can occur
across a gap of 10 mm. It is widely
observed that the regenerated
urodele spinal cord is thinner than
the intact cord, there are fewer axons, and not all of the connections
made by regenerating neurons are
correct after regeneration over a
period of weeks or a few months
(Stensaas, 1983; Davis et al., 1989).
Long-term regeneration studies in
the axolotl (up to 23 months) show
that with enough time, the number
of neurons in the nuclei of the medulla, midbrain, and diencephalon
with axons that reach regenerated
lumbar spinal cord attains control
levels (Clarke et al., 1988).
MODULATORS OF
REGENERATION
Calcium
The intrinsic properties and environmental cues that trigger the onset of
the ependymal response after injury
have yet to be determined. The involvement of calcium early in postinjury events in mammalian cord suggests that it should be considered in
initiation of the regeneration process. Excitotoxicity is among the
damaging events and toxic reactions that occur in the mammalian
spinal cord after serious injury (reviewed in McDonald and Sadowsky,
2002). Extracellular calcium levels
decrease, and intracellular calcium
levels increase (Happel et al., 1981;
Stokes et al., 1983). Calcium influxes
occur in association with glutamate
excitotoxicity that can kill neurons
and oligodendrocytes (Panter et al.,
1990). Examination of calcium activity in injured rat spinal cord suggests
that cells within the dorsal gray matter are strongly involved and the effects are then propagated into the
white matter (Moriya et al., 1994;
McDonald and Sadowsky, 2002). By
using an in vitro system, it was shown
that stimulation of increases in cytosolic calcium in individual astrocytes
is propagated to neighboring astrocytes and to adhering neurons, providing an additional path for calcium-mediated toxic events after CNS
injury (Nedergaard, 1994). This process has been shown to have consequences in vivo in mammalian
CNS (Araque et al., 2001). The meningeal cells can also participate in
transfer of calcium (Grafstein et al.,
2000).
Calcium and the urodele CNS.
One major difference in the urodele
and mammalian responses to spinal
cord injury may lie in the ability of
injury-reactive ependymal cells to
act as a buffer, possibly through calcium uptake and sequestration that
protects neurons and oligodendrocytes from the processes that would
trigger secondary cell death. Although some cell death would occur, the propagation of toxic effects
would be limited. This putative calcium uptake process could also
serve as the trigger for the onset of
the ependymal response itself
through second messenger signaling pathways. This mechanism remains theoretical, the subject of future investigations using cell culture
model systems.
What is known about calcium and
the urodele CNS is that modulation
of intracellular calcium levels is important throughout amphibian neural development. Calcium influx
plays a specific role in amphibian
neuralization (induction of the neural plate; Moreau et al., 1994; Duprat, 1996). There are species differences among amphibian embryos:
axolotl embryos exhibit “autoneuralization” in that their cells have much
302 CHERNOFF ET AL.
higher resting internal Ca⫹⫹ levels
than P. waltl and X. laevis, and there
is no incremental effect with calcium channel agonists (Duprat,
1996). During neurulation, the lifting
of the neural folds is calcium-dependent. This finding was shown in studies with Ambystomatid embryos
(axolotls and spotted salamander
Ambystoma maculatum; Moran,
1976; Moran and Rice, 1976). The developmental role of calcium that is
most relevant to regeneration may
be in neuronal differentiation. There
is a threshold level of calcium required for neuronal differentiation
that can be achieved by different
mechanisms in different species.
Ca⫹⫹ is known to be necessary for
differentiation of spinal neurons in
Xenopus laevis, and the Ca⫹⫹ transients associated with this process
occur spontaneously (Holliday and
Spitzer, 1990). One consequence of
the activation of L-type calcium
channels in neuronal differentiation
is the activation of immediate early
gene proteins such as c-fos or Jun-B,
demonstrated in tissue culture cell
lines, such as PC12 cells (Sheng and
Greenberg, 1990). During neurite
outgrowth in cultured Xenopus neurons, calcium is required in the culture medium, but the optimum calcium concentration for axonal
outgrowth is low compared with levels for neuralization (Bixby and
Spitzer, 1984). In general, calcium
homeostasis changes during development with progressively lower levels of intracellular calcium from neuralization to axonal outgrowth
(Duprat, 1996).
Retinoic Acid
To examine the effects of retinoic
acid on neurite outgrowth, explants
of axolotl spinal cord were grown in
collagen gels with a source of alltrans RA (Hunter et al., 1991). These
studies used juvenile animals between 4 and 8 cm in length, which
are still growing and adding new
CNS neurons (Holder et al., 1991).
Neurite outgrowth occurs in the
presence of RA with no other growth
factors present, although the outgrowth is somewhat sparse. In vivo
immunohistochemical studies show
that cytoplasmic retinol binding pro-
tein is present in the ependymal cell
(radial glial) bodies surrounding the
central canal in spinal cord and
hindbrain as well as in the glial processes of the ventral floor plate. Cytoplasmic retinoic acid binding protein shows a reciprocal antibody
binding pattern within the axons of
the spinal cord white matter (Hunter
et al., 1991). Endogenous retinol and
retinoic acid are detected in the spinal cord. Combined information
from the axolotl studies and embryonic chick and mouse studies show
that ependymal cells can take up
retinol, convert it to retinoic acid,
and secrete it. Neurons can then
take up retinoic acid, which promotes neurite outgrowth. The retinoic acid source elicits directional
axonal outgrowth in vitro, suggesting
that the retinoic acid–producing
ependymal cells could exert a chemoattractant effect on axons, but
an in vivo effect in spinal cord regeneration has not yet been identified (Hunter et al., 1991).
Fibroblast Growth Factor
In the intact urodele tail cord, FGF-2
is produced in ventrolateral neurons,
not ependymal cells. FGF-2 mRNA is
produced in increasingly higher levels in the blastema of the regenerating tail. In this case, production of
FGF-2 has been shown to be associated with the ependymal cells. The
FGF-2 levels are high as the ependymal tube is extending and being rebuilt (Zhang et al., 2000). Two-week
regenerates are actively expressing
FGF-2 and making new neurons.
FGF2 is also expressed in the regenerating spinal sensory ganglia. Exogenous FGF-2 increases the number
of proliferating cells in vivo in P. waltl
(Zhang et al., 2000).
In other systems, FGF-2 plays a role
in regulation of neural differentiation
(Temple, 2001). In adult mammalian
CNS, FGF-2 is expressed by fibrous
astrocytes and subsets of neurons. It
is up-regulated after injury to adult
mammalian CNS, where it and the
FGF-2 receptor probably play a role
in the astrocyte proliferation that
generates glial scars and in oligodendrocyte proliferation (Fawcett
and Asher, 1999). In neural stem cell
studies in vitro, exogenous FGF-2
keeps cells dividing and undifferentiated (Svendsen and Caldwell,
2000; Temple, 2001).
Epidermal Growth Factor
Injury-reactive adult axolotl ependymal cells show a differential response to a variety of growth factors
in vitro. The most important observation is that exogenous epidermal
growth factor (EGF) is essential for
stimulating axolotl ependymal proliferation from background levels
(10%) to high levels (50 –70%) in vitro
(O’Hara and Chernoff et al., 1994).
EGF is also required to maintain the
migration of the axolotl ependymal
cells in culture (O’Hara and
Chernoff, 1994). The source of EGF
(or its potential substitute, transforming growth factor-␣) in vivo remains
to be determined. Cultured axolotl
ependymal cells did not respond to
exogenous FGF as did the P. waltl tail
cord cell in situ (O’Hara and
Chernoff, 1994; Zhang et al., 2000).
Perhaps axolotl ependymal cells
make sufficient endogenous FGF.
EGF receptor expression appears
during development of the rat cerebral cortex in response to stimulation
by FGF-2 (Lillien and Raphael, 2000).
EGF is important in the behavior of
adult mammalian neural stem cells
as well as in development (Weiss et
al., 1996).
NEURAL STEM CELL PROPERTIES
The growth factor requirements and
endogenous growth factor production of regenerating urodele cord
ependymal cells fit the emerging
picture of vertebrate neural stem
cells. During development, FGF-2
maintains the progenitor cell properties of early embryonic mammalian
neural stem cells and maintains their
proliferation (Temple, 2001). EGF is
involved in supporting proliferation
of mammalian neural stem cells derived from later embryos and adult
tissue (Weiss et al., 1996; Burrows et
al., 1997; Lillien and Raphael, 2000;
Temple, 2001). There are regional differences in the mammalian CNS.
Adult murine spinal cord stem cells
respond to EGF plus FGF, whereas
brain lateral ventricle stem cells proliferate in the presence of EGF alone
URODELE SPINAL CORD REGENERATION 303
(Weiss et al., 1996). The stem cells
that respond to FGF and/or EGF
maintain pluripotency.
One important marker of stem cell
properties is activity of the Notch-1
signaling pathway. Notch-1 signaling maintains neural stem cell properties, but does not create them (Hitoshi et al., 2002). Musashi-1 is an
RNA-binding protein first identified in
Drosophila for its involvement in the
maintenance of Notch-1 signaling
(Nakamura et al., 1994). MouseMusashi-1 is highly enriched in the
CNS stem cells and increases
Notch-1 signal activity by posttranscriptional down-regulation of Numb
(Sakakibara et al., 1996; Imai et al.,
2001). Downstream of Notch-1 receptor activity is activation of E(spl)C/Hes genes (enhancer of splithairy). Hes gene activity represses
expression of proneural genes such
as Mash and neurogenin, maintaining stem cell or progenitor cell properties (Artavanis-Tsakonas et al.,
1999; Ohtsuka et al., 1999; Johansson et al., 1999).
Expression of these genes is being
evaluated in amphibian spinal cord.
One difference between amphibian
spinal cord and most mammalian
neural stem cell source tissues is that
the amphibian spinal cord stem cells
are ventricular rather than subventricular (Momma et al., 2000; Morshead and van der Kooy, 2001;
Alvarez-Buylla et al., 2002). In amphibians, spinal cord cells with stem
cell properties have been identified
from regeneration-competent Xenopus tadpole and adult axolotl spinal cord (Chernoff et al., 2001; unpublished results of the Chernoff
Laboratory). These cells express
nrp-1, the Xenopus homolog of
Musashi-1 (Richter et al., 1990). Nrp-1
expression is down-regulated as regenerative capacity is lost in Xenopus and is up-regulated during adult
axolotl spinal cord regeneration (unpublished results of the Chernoff
Laboratory).
PATTERNING OF THE
REGENERATING CORD
In amphibian limb regeneration, it is
clear that the patterning process recapitulates a substantial portion of
limb development. In the spinal
cord, we must look primarily to nonamphibian systems for molecular
mechanisms of axial patterning of
the developing cord, as there is limited information available from
urodeles or other amphibians. For
spinal cord regeneration to completely reproduce missing structures
and produce functional axonal targeting, it may be necessary to express or re-express A-P and D-V axis
cues.
A-P Patterning
Detailed studies of A-P and D-V patterning of the developing spinal
cord have been performed primarily
in amniote embryos (reviewed in Altmann and Brivanlou, 2001). A-P patterning of the embryonic spinal cord
starts with additive expression of
Hox-a and Hox-b cluster genes and
progresses to establishing the identities of subgroups of spinal motoneurons through expression of genes
such as engrailed (McGinnis and
Krumlauf, 1992; Pfaff and Kintner,
1998). Cross-inhibitory regulation between the bHLH transcription factors
neurogenin, Math1 and Mash1 helps
to determine progenitor cell domains and neuronal fate in the spinal cord of higher vertebrates
(Shirasaki and Pfaff, 2002).
Although A-P patterning of the spinal cord occurs early in CNS development, there are later A-P cues,
such as those that pattern subgroups of motor neurons (Pfaff and
Kintner, 1998). The persistent expression of patterning genes in adult tissue has been seen in other urodele
systems. HoxA11 and distal-less family genes are expressed in intact
adult newt limb and in limb regeneration (Beauchemin and Savard,
1992; Beauchemin et al., 1994).
There are similar patterning gene examples in spinal cord regeneration.
In Pleurodeles waltl, two Nkx3-related
genes (PwNkx3.3 and PwNkx3.2 ) exhibit graded expression in normal
adult spinal cord along the A-P axis
(Nicolas et al., 1999). The gradient is
more pronounced for PwNkx3.3,
which is stronger posteriorly. Reverse
transcriptase-polymerase chain reaction of mRNA from 11 regions (anterior to posterior along the body)
shows increased expression of both
genes at the level of the lumbar en-
largement. These genes are up-regulated in many tissues of the regenerating tail and may be particularly
important in skeletal regeneration.
Wnt growth factor genes provide a
second example of adult expression
of pattern regulating genes. In Pleurodeles waltl, Pwnt-10a and Pwnt-7a
are expressed in normal adult tissue
and up-regulated during tail regeneration (Caubit et al., 1997a,b). Several of the Wnt genes have also
been identified with A-P axis-associated expression during regeneration. Pwnt-5a, -5b, and -10a are expressed in an A-P gradient. No
identification has been made of Wnt
gene involvement in D-V axis regeneration comparable to that found in
dorsal neural tube differentiation
(Muroyama et al., 2002).
Patterning genes are associated
specifically with regeneration of
missing structures in tail regeneration. Dlx3 (PwDlx3), a distal-less
gene, appears in the tail cord
ependymal cells that re-form the
dorsal root ganglia (Nicolas et al.,
1996). This is a gene that is not expressed in normal adult tissue and
may not be associated with the embryonic origin of dorsal root ganglia
(Beauchemin and Savard, 1992;
Nicolas et al., 1996).
D-V Patterning
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), from the notochord and neural tube floor plate, is
key in the ventralizing process,
whereas the bone morphogenetic
proteins, from the ectoderm and
neural tube roof plate, are important for dorsalization (Altmann and
Brivanlou, 2001; Eggenschwiler et al.,
2001).
Specific information on urodele
patterning responses is sparse, and
the details of patterning in development or regeneration may have
species-specific variations. The Gli
genes Gli1 and Gli2 are expressed in
response to SHH in amniote embryos. One of the known targets of
Gli is the forkhead family winged-helix transcription factor HNF3-␤, which
304 CHERNOFF ET AL.
can activate floor plate marker production (Briscoe and Ericson, 2001).
In Xenopus the winged-helix forkhead transcription factor XFKH1 (Pintallavis, XFD-1) is expressed during
gastrulation in the dorsal lip of the
blastopore and, subsequently, in the
notochord and the floor plate of the
neural tube (Dirksen and Jamrich,
1992; Ruiz i Altaba and Jessel, 1992;
Knochel et al., 1992). The apparent
axolotl homolog AxFKH1 is expressed
only in the floor plate (Whitely et al.,
1997). It has been suggested that
the relationship between the notochord and floor plate is different in
Xenopus and axolotls. Alternatively,
it is suggested that another fork
head gene, such as Ax FKH2 is actually functionally homologous to
XFKH1 (Whitely et al., 1997).
There is currently no information
about expression and regulation of
the urodele homologues of the amniote embryo homeodomain genes
involved in production of the dorsal
and ventral spinal cord interneurons
and motoneurons (Briscoe and Ericson, 1999, 2001; Lee and Jessell,
1999; Briscoe et al., 2000; Patten and
Placzek, 2000; Altmann and Brivanlou, 2001; Shirasaki and Pfaff, 2002).
This involvement of these genes,
Pax, Nkx, Dbx, Irx, and LIM-homeodomain genes, in regeneration is
an area of active interest at this
time.
In regeneration, D-V signaling
might be required to establish the
identity and differentiation of new
neurons and for the targeting of regrowing axons. Possibly, stump tissues provide signals to adjacent
cells that induce the patterning of
the regenerate. Also unknown is the
relationship between D-V signaling
and the A-P patterning of the regenerate. There is a strong indication
that the patterning process exists
and is generated differently than it is
during development: in tail regeneration, PNS ganglia are produced
from the ependymal tube rather
than from neural crest cells (see the
axonal regrowth section above).
Studies on the expression of patterning genes in urodele spinal cord regeneration are ongoing in several
laboratories.
PERSPECTIVES
Urodele spinal cord regeneration
does not appear to be a recapitulation of spinal cord development.
There are fundamental, structural
reasons for the differences. The predominant difference is that the progenitor tissues no longer exist and
are not reconstructed during regeneration. The neural plate is not reconstructed; therefore, regeneration does not repeat the events of
neurulation. The notochord is buried
in the centrum of the vertebrae and
is no longer a source of ventralizing
signals. Presumptive skin ectoderm
no longer exists, but the skin ectoderm does produce a wound epithelium that can act as a source of
signals to the spinal cord in caudal
regrowth (Godlewski, 1928; Polezhaev, 1972). In caudal replacement, the PNS ganglia must regenerate from the CNS, as the neural
crest no longer exists (Egar and
Singer, 1972; Nordlander and Singer,
1978; Arsanto et al., 1992; Benraiss et
al., 1999; Zhang et al., 2000). The
structural differences suggest that at
least some of the expression of axial
patterning genes will also differ from
that in embryonic neurulation. This
has been shown in A-P patterning of
caudal regrowth (Beauchemin and
Savard, 1992; Nicolas et al., 1996;
Caubit et al., 1997a,b). Studies of
D-V patterning during regeneration
are just beginning. Without the notochord and skin epidermis as signaling centers, it is possible that the D-V
patterning cues are propagated in
a planar manner from the stump tissue. The identity of these signals and
their spatial and temporal patterns
of expression are areas of current
research interest.
The ependymal outgrowth and
sprouting axons would serve as the
target for A-P and D-V patterning
signals in reconstruction of the regenerating spinal cord. These processes could be coordinated by signals induced or up-regulated in
stump tissue. Differentiation of the
ependymal outgrowth could be coordinated with rearrangement of
cells in the remodeling stump tissues
and axonal regrowth from stump tissue neurons.
Experimental observations of FGF
and EGF responsiveness, along with
studies of neurogenesis in urodeles,
link the ependymal cell response in
cord regeneration with studies of
mammalian neural stem cells (Holder et al., 1991; O’Hara and Chernoff,
1994; Benraiss et al., 1999; Zhang et
al., 2000). The expression of the neural stem cell marker Musashi-1 and
other elements of the Notch-1 signaling pathway will allow us to examine the differentiated state of
ependymal cells throughout the
process of spinal cord growth and
regeneration. Understanding the
regulation of stem cell properties
of urodele ependymoglia will be
important in understanding the
urodele spinal cord regeneration
process.
ACKNOWLEDGMENTS
E.A.G.C. received funding from the
NSF, and D.L.S. was funded by an
Indiana 21st Century Research and
Technology Fund.
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