1
Title
Axonal regeneration in zebrafish
Authors
Thomas Becker, Catherina G. Becker
Centre for Neuroregeneration, School of Biomedical Sciences, The Chancellor’s
Building, 49 Little France Crescent, Edinburgh EH16 4SB, UK
For correspondence: catherina.becker@ed.ac.uk, thomas.becker@ed.ac.uk
Abstract
In contrast to mammals, fish and amphibia functionally regenerate axons
in the central nervous system (CNS). The strengths of the zebrafish model, i.e.
transgenics and mutant availability, ease of gene expression analysis and
manipulation and optical transparency of larvae lend themselves to the analysis
of successful axonal regeneration. Analyses in larval and adult zebrafish suggest
a high intrinsic capacity for axon regrowth, yet signalling pathways employed in
axonal growth and pathfinding are similar to those in mammals. However, the
lesioned CNS environment in zebrafish shows remarkably little scarring or
expression of inhibitory molecules and regenerating axons use molecular cues in
the environment to successfully navigate to their targets. Future zebrafish
research, including screening techniques, will complete our picture of the
mechanisms behind successful CNS axon regeneration in this vertebrate model
organism.
Introduction
2
The high capacity of fish and amphibians to regenerate organs,
appendages and CNS structures has long been recognized [1]. However, it is still
unresolved why amniotes, and in particular mammals, have lost the capacity for
CNS regeneration during evolution. There is hope that mechanisms of successful
regeneration can be gleaned from studying regenerating model vertebrates, such
as zebrafish, and translating knowledge to analyse non-regenerating species. The
zebrafish combines experimental and genetic accessibility with an enormous
regenerative capacity and is, therefore, advancing to be a widely used vertebrate
model of successful CNS regeneration. In the zebrafish CNS, entire neuronal
populations regenerate from progenitor cells, which has recently been expertly
reviewed elsewhere [2,3]. Here we focus on the events related to injury of axons
in the CNS and regrowth from those neurons that have been axotomized.
Extent of axon regrowth
In mammals, axon regrowth in the CNS is extremely limited and some
types of axotomized neurons, such as retinal ganglion cells, even perish [4]. In
contrast, anamniotes, including zebrafish, have an astonishing capacity to
successfully regrow long-range projection axons over distances that are much
greater than when these axons first made connections during development. For
example, after a crush or complete transection lesion of the optic nerve, retinal
ganglion cells (RGCs) survive [5,6] and their axons regrow to faithfully and
topographically re-innervate nine termination fields in the adult brain [7].
Regenerating RGC axons reach the tecum opticum, the largest termination field,
by 8 days post-lesion (dpl). During the entire regeneration process only a few
projection errors are made. A few axons erroneously grow ipsi-laterally into the
3
brain, and some optic axons from the dorsal retina grow erroneously through the
dorsal brachium of the optic tract and vice versa [7]. However, these pathfinding
errors do not lead to apparent errors in target innervation, as retinotopy of the
projection is reestablished by 42 dpl in a pattern that is indistinguishable from
unlesioned animals.
After a complete transection of the spinal cord, severed axons of
brainstem neurons with spinal projections cross into the distal part of the spinal
cord by 14 dpl and project at least 3.5 mm beyond the initial transection site by
42 dpl [8,9]. Interestingly, regenerating axons re-route through the central grey
matter, rather than growing through the peripheral white matter, thus
navigating novel pathways [10]. Remarkably, not all severed axons regenerate
equally well. Some brain nuclei with descending axons show poor axon regrowth
[11]. This includes the individually identifiable paired Mauthner neurons, unique
large brainstem neurons in aquatic vertebrates. Moreover, descending
monoaminergic axons manage to cross the lesion site, but penetrate only a few
micrometers into the distal spinal cord [12]. Severed dorsal root axons and
ascending axons of intraspinal neurons also show little detectable regrowth [13].
Thus the regrowth capacity for a number of axon types varies and is well
established in the adult zebrafish, providing a model to study differences in the
regenerative success of CNS axons.
Axon regeneration is functional
Despite minor targeting errors in the regenerated optic projection and
imperfect regeneration of spinal axons, functional recovery is spectacular. After
optic nerve injury the optokinetic and optomotor responses are recovered by 14
4
and 28-35 dpl, respectively, matching axon regrowth and re-establishment of
retinotopy. More complex visually guided behaviours, such as two fish chasing
each other, take longer to recover (3 months), perhaps related to long-term
synaptic rearrangements [5,14].
After spinal cord transection, fish are completely paralyzed caudal to the
lesion site, but within 42 dpl, most regain swimming activity and the ability to
maintain their position in a water flow, similar to uninjured control animals
[15,16]. Creating a physical barrier to axon regrowth in the spinal cord prevents
recovery [9] and re-transecting the spinal cord abolishes it [12], providing
evidence that regrowth of axons is indispensible for recovery. Hence, functional
recovery after axonal regeneration in the optic projection and spinal cord are
robust and can be assessed by a number of quantitative assays.
Extrinsic determinants of axon regrowth
In mammals, lack of axon regrowth is brought about in part by a hostile
growth environment in which astrocytes and other cells form the glial scar
containing growth-inhibitory extracellular matrix (ECM) components, such as
chondroitin sulfate proteoglycans (CSPGs). In addition, myelin and myelin debris
from degenerating fibers contain growth-inhibitory molecules [17]. After a lesion
of the optic nerve in zebrafish, there is no evidence of a CSPG expressing glial
scar [18]. In the spinal cord, ependymo-radial glial cells, which also have
astroglial functions, even form bridges that re-connect the severed spinal cord. In
transgenic larvae, expressing green fluorescent protein under the glial fibrillary
acidic protein promoter in ependymo-radial glial cells, rejoining of the spinal and
glial bridges can be observed by time-lapse microscopy [8]. These bridges have
5
been suggested to guide or support axon growth across the lesion site [8].
Ependymal progenitor cells in the mammalian spinal cord, which are similar to
ependymo-radial glial cells in zebrafish, generate scar cells [19]. Remarkably, in
zebrafish, but not in mammals, ependymo-radial cells generate neuronal cell
types after a lesion [20,21], perhaps in lieu of scar cells, which might be one of
the reasons for the absence of a detectable glial scar in zebrafish.
Myelin-associated inhibitory molecules, such as NogoA/RNT4 [22] and
MAG/siglec-4 [23] do exist in the zebrafish CNS. However, at least for zebrafish
NogoA it has been shown that the NogoA-specific N-terminal inhibitory domain
is missing, and the other protein domain that is inhibitory in mammals (Nogo66)
fails to elicit growth cone collapse of regenerating axons in vitro [22]. Moreover,
zebrafish oligodendrocytes, in contrast to mammalian oligodendrocytes,
increase expression of recognition molecules that may promote axon growth,
such as contactins [24,25], P0 [26] and L1-related molecules [27], after a CNS
lesion.
Receptors for inhibitory molecules, such as Nogo66 and CSPG receptor
NgR [28] and the CSPG receptors RPTP-sigma and LAR [29] are expressed in the
zebrafish CNS and at least for NgR there is evidence for expression on
regenerating axons [22]. However, the exact spatio-temporal regulation of
receptor expression during axon regeneration needs further investigation.
Similar to mammals, there is also a strong activation of
macrophages/microglial cells after an optic nerve or spinal lesion in zebrafish
[10,26,30,31]. Lysophosphatidic acid induced boosting of the immune response
after spinal injury negatively impacted neurite growth in a recent study [30].
However, the exact mechanisms how the immune response influences axonal
6
regrowth need to be determined. Overall, the cellular and molecular composition
of the adult lesioned CNS in zebrafish presents an environment that is
presumably more conducive to axon regrowth than the CNS environment in
mammals.
Neuron-intrinsic factors regulating axon regrowth
In general, severed CNS axons in zebrafish have a high capacity for
regrowth. For example, in retinal explant culture, RGC axons grow much more
vigorously than mammalian counterparts and they upregulate a number of well
known regeneration/growth-associated molecules, such as GAP-43 [32], L1related proteins [33] and alpha-1 tubulin [34]. Similar to mammalian neurons
with non-regenerating axons, L1-related proteins and GAP-43 are not
upregulated in those zebrafish brain nuclei that show a low regenerative
capacity [11]. The functional importance of these genes has been demonstrated
in vivo or explant culture by inserting a gelfoam pledget soaked with anti-sense
morpholino oligonucleotide into either spinal or optic nerve lesion site. The
morpholino is retrogradely and selectively transported to the somata of
axotomized neurons where it suppresses expression of target genes for weeks,
long enough to show effects on regeneration [33,34].
While the above-mentioned genes are also expressed during
developmental axon growth, there is evidence that they are differently regulated
during development and adult regeneration. Transgenic reporter lines, using
different fragments of the regulatory sequences of GAP-43 or alpha-1 tubulin to
drive reporter gene expression, have demonstrated the presence of
regeneration-specific mechanisms of gene regulation [32,35]. Indeed, expression
7
profiling of retinal ganglion cells undergoing axon regrowth revealed that some
genes are uniquely expressed during regeneration, such as the transcription
factor KLF6/7, which in turn regulates expression of alpha-1 tubulin [34,36,37].
The powerful combination of expression profiling with morpholino knock-down
has led to the identification of a number of additional genes that may play
specific roles in regeneration [38 and citations therein].
Interestingly, not all genes that are upregulated in neurons with
regenerating axons promote axon growth. For example, socs3, a strong neuronintrinsic inhibitor of axon regeneration in mammals, is upregulated in retina
ganglion cells after an optic nerve lesion in zebrafish, and attenuates axonal
regeneration [39]. This suggests that the molecular injury response in zebrafish
neurons may be more similar to that in mammals than previously thought.
Studying zebrafish neurons that do not regenerate axons and in which
upregulation of regeneration-associated genes fails, might be particularly
instructive. For example, the Mauthner neuron’s regenerative capacity is limited
already in larvae, which makes it possible to directly observe effects of
manipulations that augment axon regrowth, as in the PNS (see Box 1). Like in
mammalian neurons, increasing the levels of cAMP in the axotomized Mauthner
neuron leads to increased and directional axon regrowth, resulting in recovery of
function [40]. Moreover, the small size of the zebrafish larvae makes it ideally
suited for drug screening efforts and indeed, paradigms are being established for
semi-automated high-throughput laser-sectioning of the Mauthner axon, greatly
facilitating screens that aim to identify factors to promote regeneration [41].
Guidance of regrowing axons
8
It is perhaps the most astonishing property of adult axon regeneration in
zebrafish that axons reach their targets over long distances to make functional
reconnections. Indeed, it has been noted in mammalian systems that even when
regrowth of CNS axons is experimentally induced, axons frequently fail to
navigate correctly [42,43]. It could be hypothesized that regenerating axons in
the zebrafish CNS simply retrace their former pathways along degenerating
tracts due to physical constraints, similar to the PNS [44]. The observation that
most regenerating axons in the spinal cord re-route to the gray matter during
regeneration and not through the denervated white matter does not support this
idea [10]. Moreover, we tested this hypothesis experimentally in a mutant of the
robo2 recognition molecule. This mutation leads to the random and variable
appearance of ectopic tracts during development [45]. If degenerating tracts
guided optic axons, these should faithfully be re-used by regenerating axons.
This was, however, not the case, refuting the hypothesis that degenerating tracts
present the predominant guidance cue to regenerating axons [46].
What guides regenerating adult axons? CSPGs and other inhibitory
molecules have functions in developmental axon guidance, by repelling axons
from areas that are not to be innervated. Regenerating zebrafish optic axons are
sensitive to axon-repellent/inhibitory guidance molecules. For example, these
axons do not penetrate a substrate boundary of axon-repellent ECM molecules,
such as tenascin-R and CSPGs in vitro [18,47]. Indeed, in vivo, tenascin-R
surrounds the optic projection, consistent with repellent axon guidance.
Tenascin-R and CSPGs are particularly strongly expressed in the posterior
pretectal nucleus, a diencephalic nucleus that is engulfed by optic axons, but does
not receive primary visual input. Enzymatic removal of CSPGs in vivo allows
9
optic axons to partially invade this nucleus, suggesting that repellent ECM
molecules guide regenerating optic axons around this nucleus [18]. Adult
zebrafish also retain graded expression of axon repellent ephrins in the tectum
[7], which is important for correct retinotopic mapping of optic axons during
development. The presence of guidance cues in the adult optic system in
zebrafish may be related to the continuous addition of retinal ganglion cells in
adults, which need to find their way to the tectum. These cues are available also
to regenerating axons. In adult rats, in which no new axons are added to the optic
projection, ephrin gradients and appropriate receptor gradients in retinal
ganglion cells are down-regulated. However, this is reversed upon a lesion of the
adult optic nerve, indicating that the ephrin guidance system might also be
available in the lesioned CNS of mammals [48].
Conclusion
Successful CNS regeneration is a process for which a number of intrinsic
and extrinsic factors have to interact to allow functional reconnections. Analyses
of axon regrowth in the CNS of zebrafish show high intrinsic axon growth
capacity, minimal scar formation, low expression of growth inhibitors and
guidance of regenerating axons by molecular cues. It is striking that in zebrafish,
environmental and intrinsic factors are so well co-ordinated to allow for axonal
regeneration, whereas in mammals the opposite appears to be the case. However,
intrinsic and extrinsic factors might be functionally connected, such that
changing only a few parameters affects both axons and environment. For
example, stabilizing microtubuli in the lesioned spinal cord of mammals using
Taxol improves both intrinsic axon regrowth and environmental scarring [49].
10
Given the unique array of genetic, optic and screening tools available in the
zebrafish model, we expect zebrafish to contribute to elucidating the molecular
mechanisms underpinning axon regrowth and navigation leading to functional
reconnections in the vertebrate CNS.
Acknowledgements
We thank Drs. David Lyons and Dirk Sieger for critical reading.
11
TEXTBOX 1
The peripheral nervous system (PNS) of larvae is highly accessible to study
cell-cell interactions after laser-microsurgical lesions.
Here we review some recent studies on peripheral nerve regeneration in
larval zebrafish, because similar techniques may be used in the CNS. As in
mammals, the PNS of zebrafish shows axonal regrowth, also in translucent larvae.
Lesion studies of the larval posterior lateral line nerve, which grows along the
body to innervate sensory hair cells, indicate that debris clearance by immune
cells, presence of Schwann cells, and target derived factors [50,51] are important
to facilitate and guide successful nerve regeneration. Lesion studies of peripheral
sensory axons in the skin have shown that environmental inhibition of axon
regeneration also takes place in the PNS [52]. Laser-lesions of motor nerves
revealed that macrophages are attracted to injured nerves very early,
independently of Schwann cells and only enter the nerve upon fragmentation of
the axons [53]. Perineurial glia use notch signaling during initial motor nerve
development, but not during nerve regeneration, suggesting that nerve
regeneration is not simply a recapitulation of development [54]. The
combination of laser-lesions and time-lapse observations in the CNS has the
potential to elucidate cell-cell interactions during axon regrowth in vivo in
unprecedented detail.
END TEXTBOX 1
12
FIGURE
Fig. 1 Graphical summary of cellular and molecular events after lesions of
the optic nerve (above) and spinal cord (below) in adult zebrafish.
13
Commented literature:
1. Gilbert SF: Developmental Biology edn 10th. Sunderland, MA: Sinauer
Associates Inc; 2013.
2. Grandel H, Brand M: Comparative aspects of adult neural stem cell activity
in vertebrates. Dev Genes Evol 2013, 223:131-147.
3. Gemberling M, Bailey TJ, Hyde DR, Poss KD: The zebrafish as a model for
complex tissue regeneration. Trends Genet 2013.
4. Fischer D, Leibinger M: Promoting optic nerve regeneration. Prog Retin Eye
Res 2012, 31:688-701.
5. Zou S, Tian C, Ge S, Hu B: Neurogenesis of retinal ganglion cells is not
essential to visual functional recovery after optic nerve injury in
adult zebrafish. PLoS One 2013, 8:e57280.
6. Nagashima M, Fujikawa C, Mawatari K, Mori Y, Kato S: HSP70, the earliestinduced gene in the zebrafish retina during optic nerve
regeneration: its role in cell survival. Neurochem Int 2011, 58:888-895.
7. Becker CG, Meyer RL, Becker T: Gradients of ephrin-A2 and ephrin-A5b
mRNA during retinotopic regeneration of the optic projection in
adult zebrafish. J Comp Neurol 2000, 427:469-483.
*This paper demonstrates the astonishingly precise regeneration of the optic
projection
8. Goldshmit Y, Sztal TE, Jusuf PR, Hall TE, Nguyen-Chi M, Currie PD: Fgfdependent glial cell bridges facilitate spinal cord regeneration in
zebrafish. J Neurosci 2012, 32:7477-7492.
**The absence of a glial scar in the lesioned spinal cord and active bridging of the
lesion site by glial cells is demonstrated.
9. Becker T, Wullimann MF, Becker CG, Bernhardt RR, Schachner M: Axonal
regrowth after spinal cord transection in adult zebrafish. J. Comp.
Neurol. 1997, 377:577-595.
10. Becker T, Becker CG: Regenerating descending axons preferentially
reroute to the gray matter in the presence of a general
macrophage/microglial reaction caudal to a spinal transection in
adult zebrafish. J Comp Neurol 2001, 433:131-147.
11. Becker T, Bernhardt RR, Reinhard E, Wullimann MF, Tongiorgi E, Schachner
M: Readiness of zebrafish brain neurons to regenerate a spinal axon
correlates with differential expression of specific cell recognition
molecules. J Neurosci 1998, 18:5789-5803.
*Not all brainstem neurons with spinal projections initiate regeneration-related
gene expression, including the individually identifyable Mauthern cells.
12. Kuscha V, Barreiro-Iglesias A, Becker CG, Becker T: Plasticity of tyrosine
hydroxylase and serotonergic systems in the regenerating spinal
cord of adult zebrafish. J Comp Neurol 2012, 520:933-951.
13. Becker T, Lieberoth BC, Becker CG, Schachner M: Differences in the
regenerative response of neuronal cell populations and indications
14
for plasticity in intraspinal neurons after spinal cord transection in
adult zebrafish. Mol Cell Neurosci 2005, 30:265-278.
14. Kato S, Matsukawa T, Koriyama Y, Sugitani K, Ogai K: A molecular
mechanism of optic nerve regeneration in fish: The retinoid
signaling pathway. Prog Retin Eye Res 2013.
15. Dias TB, Yang YJ, Ogai K, Becker T, Becker CG: Notch signaling controls
generation of motor neurons in the lesioned spinal cord of adult
zebrafish. J Neurosci 2012, 32:3245-3252.
16. van Raamsdonk W, Maslam S, de Jong DH, Smit-Onel MJ, Velzing E: Long
term effects of spinal cord transection in zebrafish: swimming
performances, and metabolic properties of the neuromuscular
system. Acta Histochem 1998, 100:117-131.
17. Fawcett JW, Schwab ME, Montani L, Brazda N, Muller HW: Defeating
inhibition of regeneration by scar and myelin components. Handb
Clin Neurol 2012, 109:503-522.
18. Becker CG, Becker T: Repellent guidance of regenerating optic axons by
chondroitin sulfate glycosaminoglycans in zebrafish. J Neurosci 2002,
22:842-853.
** In vivo degradation of GSPGs by intraventricular injections of chondroitinase
demonstrates that regenerating optic axons are repelled by CSPGs in the
posterior pretectal nucleus during active pathfinding.
19. Meletis K, Barnabe-Heider F, Carlen M, Evergren E, Tomilin N, Shupliakov O,
Frisen J: Spinal cord injury reveals multilineage differentiation of
ependymal cells. PLoS Biol 2008, 6:e182.
20. Reimer MM, Sörensen I, Kuscha V, Frank RE, Liu C, Becker CG, Becker T:
Motor neuron regeneration in adult zebrafish. J Neurosci 2008,
28:8510-8516.
21. Kuscha V, Frazer SL, Dias TB, Hibi M, Becker T, Becker CG: Lesion-induced
generation of interneuron cell types in specific dorsoventral
domains in the spinal cord of adult zebrafish. J Comp Neurol 2012,
520:3604-3616.
22. Abdesselem H, Shypitsyna A, Solis GP, Bodrikov V, Stuermer CA: No Nogo66and NgR-mediated inhibition of regenerating axons in the zebrafish
optic nerve. J Neurosci 2009, 29:15489-15498.
**Molecular dissection of the function of NogoA shows how domains and signal
transduction of the protein differ in zebrafish to allow axon regrowth.
23. Lehmann F, Gathje H, Kelm S, Dietz F: Evolution of sialic acid-binding
proteins: molecular cloning and expression of fish siglec-4.
Glycobiology 2004, 14:959-968.
24. Schweitzer J, Gimnopoulos D, Lieberoth BC, Pogoda HM, Feldner J, Ebert A,
Schachner M, Becker T, Becker CG: Contactin1a expression is
associated with oligodendrocyte differentiation and axonal
regeneration in the central nervous system of zebrafish. Mol Cell
Neurosci 2007, 35:194-207.
25. Haenisch C, Diekmann H, Klinger M, Gennarini G, Kuwada JY, Stuermer CA:
The neuronal growth and regeneration associated Cntn1
(F3/F11/Contactin) gene is duplicated in fish: expression during
15
development and retinal axon regeneration. Mol Cell Neurosci 2005,
28:361-374.
26. Schweitzer J, Becker T, Becker CG, Schachner M: Expression of protein zero
is increased in lesioned axon pathways in the central nervous
system of adult zebrafish. Glia 2003, 41:301-317.
27. Bernhardt RR, Tongiorgi E, Anzini P, Schachner M: Increased expression of
specific recognition molecules by retinal ganglion cells and by the
optic pathway glia accompanies the successful regeneration of
retinal axons in adult zebrafish. J Comp Neurol 1996, 376:253-264.
28. Klinger M, Taylor JS, Oertle T, Schwab ME, Stuermer CA, Diekmann H:
Identification of Nogo-66 receptor (NgR) and homologous genes in
fish. Mol Biol Evol 2004, 21:76-85.
29. van der Sar A, Betist M, de Fockert J, Overvoorde J, Zivkovic D, den Hertog J:
Expression of receptor protein-tyrosine phosphatase alpha, sigma
and LAR during development of the zebrafish embryo. Mech Dev 2001,
109:423-426.
30. Goldshmit Y, Matteo R, Sztal T, Ellett F, Frisca F, Moreno K, Crombie D,
Lieschke GJ, Currie PD, Sabbadini RA, et al.: Blockage of
lysophosphatidic acid signaling improves spinal cord injury
outcomes. Am J Pathol 2012, 181:978-992.
31. Hui SP, Dutta A, Ghosh S: Cellular response after crush injury in adult
zebrafish spinal cord. Dev Dyn 2010, 239:2962-2979.
32. Kusik BW, Hammond DR, Udvadia AJ: Transcriptional regulatory regions
of gap43 needed in developing and regenerating retinal ganglion
cells. Dev Dyn 2010, 239:482-495.
33. Becker CG, Lieberoth BC, Morellini F, Feldner J, Becker T, Schachner M: L1.1
is involved in spinal cord regeneration in adult zebrafish. J Neurosci
2004, 24:7837-7842.
** This study establishes retrograde loading of axotomized projection neurons
with morpholino, a powerful technique to selectively reduce expression of
specific genes in neurons with regenerating axons.
34. Veldman MB, Bemben MA, Goldman D: Tuba1a gene expression is
regulated by KLF6/7 and is necessary for CNS development and
regeneration in zebrafish. Mol Cell Neurosci 2010, 43:370-383.
**Regeneration-specific regulation of axon growth related genes is demonstrated
by showing that the transcription factors KLF6a and 7a, which are only
induced during axon regrowth and not axon development, bind the
tuba1a promoter. This increases expression of tuba1a, which is necessary
for axon growth.
35. Veldman MB, Bemben MA, Thompson RC, Goldman D: Gene expression
analysis of zebrafish retinal ganglion cells during optic nerve
regeneration identifies KLF6a and KLF7a as important regulators of
axon regeneration. Dev Biol 2007, 312:596-612.
36. McCurley AT, Callard GV: Time Course Analysis of Gene Expression
Patterns in Zebrafish Eye During Optic Nerve Regeneration. J Exp
Neurosci 2010, 2010:17-33.
16
37. Saul KE, Koke JR, Garcia DM: Activating transcription factor 3 (ATF3)
expression in the neural retina and optic nerve of zebrafish during
optic nerve regeneration. Comp Biochem Physiol A Mol Integr Physiol
2010, 155:172-182.
38. Fang P, Pan HC, Lin SL, Zhang WQ, Rauvala H, Schachner M, Shen YQ: HMGB1
Contributes to Regeneration After Spinal Cord Injury in Adult
Zebrafish. Mol Neurobiol 2013.
39. Elsaeidi F, Bemben MA, Zhao XF, Goldman D: Jak/STAT signaling
stimulates zebrafish optic nerve regeneration and overcomes the
inhibitory actions of socs3 and sfpq. J Neurosci 2014, 34:2632-2644.
*This study provides evidence that after axonal lesion, during successful axon
regeneration zebrafish neurons upregulate signalling pathways that
inhibit axon regeneration in mammals.
40. Bhatt DH, Otto SJ, Depoister B, Fetcho JR: Cyclic AMP-induced repair of
zebrafish spinal circuits. Science 2004, 305:254-258.
**This is the first demonstration of the usefulness of the transparent larval
zebrafish for imaging of CNS regeneration, using the Mauthner neurons.
The Mauthner neurons have the advantage that they hardly regenerate,
even at larval stages and that cAMP improves their regrowth, which
makes them similar to mammalian projection neurons.
41. Pardo-Martin C, Chang TY, Koo BK, Gilleland CL, Wasserman SC, Yanik MF:
High-throughput in vivo vertebrate screening. Nat Methods 2010,
7:634-636.
*A forward-looking study showing how larval zebrafish can be used for largescale regeneration screening.
42. Pernet V, Joly S, Jordi N, Dalkara D, Guzik-Kornacka A, Flannery JG, Schwab
ME: Misguidance and modulation of axonal regeneration by Stat3
and Rho/ROCK signaling in the transparent optic nerve. Cell Death Dis
2013, 4:e734.
43. Luo X, Salgueiro Y, Beckerman SR, Lemmon VP, Tsoulfas P, Park KK: Threedimensional evaluation of retinal ganglion cell axon regeneration
and pathfinding in whole mouse tissue after injury. Exp Neurol 2013,
247:653-662.
44. Graciarena M, Dambly-Chaudiere C, Ghysen A: Dynamics of axonal
regeneration in adult and aging zebrafish reveal the promoting
effect of a first lesion. Proc Natl Acad Sci U S A 2014, 111:1610-1615.
45. Fricke C, Lee JS, Geiger-Rudolph S, Bonhoeffer F, Chien CB: Astray, a
zebrafish roundabout homolog required for retinal axon guidance.
Science 2001, 292:507-510.
46. Wyatt C, Ebert A, Reimer MM, Rasband K, Hardy M, Chien CB, Becker T,
Becker CG: Analysis of the astray/robo2 zebrafish mutant reveals
that degenerating tracts do not provide strong guidance cues for
regenerating optic axons. J Neurosci 2010, 30:13838-13849.
*Stochastically developing ectopic optic tracts are not repopulated during optic
nerve regeneration, demonstrating that degenerating optic tracts are not
a strong guidance cue per se.
17
47. Becker CG, Schweitzer J, Feldner J, Schachner M, Becker T: Tenascin-R as a
repellent guidance molecule for newly growing and regenerating
optic axons in adult zebrafish. Mol Cell Neurosci 2004, 26:376-389.
48. Symonds AC, King CE, Bartlett CA, Sauve Y, Lund RD, Beazley LD, Dunlop SA,
Rodger J: EphA5 and ephrin-A2 expression during optic nerve
regeneration: a 'two-edged sword'. Eur J Neurosci 2007, 25:744-752.
49. Hellal F, Hurtado A, Ruschel J, Flynn KC, Laskowski CJ, Umlauf M, Kapitein LC,
Strikis D, Lemmon V, Bixby J, et al.: Microtubule stabilization reduces
scarring and causes axon regeneration after spinal cord injury.
Science 2011, 331:928-931.
50. Schuster K, Dambly-Chaudiere C, Ghysen A: Glial cell line-derived
neurotrophic factor defines the path of developing and regenerating
axons in the lateral line system of zebrafish. Proc Natl Acad Sci U S A
2010, 107:19531-19536.
51. Villegas R, Martin SM, O'Donnell KC, Carrillo SA, Sagasti A, Allende ML:
Dynamics of degeneration and regeneration in developing zebrafish
peripheral axons reveals a requirement for extrinsic cell types.
Neural Dev 2012, 7:19.
52. Martin SM, O'Brien GS, Portera-Cailliau C, Sagasti A: Wallerian
degeneration of zebrafish trigeminal axons in the skin is required
for regeneration and developmental pruning. Development 2010,
137:3985-3994.
53. Rosenberg AF, Wolman MA, Franzini-Armstrong C, Granato M: In vivo
nerve-macrophage interactions following peripheral nerve injury. J
Neurosci 2012, 32:3898-3909.
*An excellent in vivo time-lapse study showing the interaction of different cell
types during motor nerve regeneration at high temporal resolution.
54. Binari LA, Lewis GM, Kucenas S: Perineurial glia require Notch signaling
during motor nerve development but not regeneration. J Neurosci
2013, 33:4241-4252.