The functional roles of PlexinD1 in chicken - ETH E

DISS. ETH No. 15930
The functional roles of PlexinD1 in chicken nervous system
during development
A dissertation submitted to the
SWISS FEDERAL INSTITUTE OF TECHNOLOGY ZURICH
for the degree of
Doctor of Sciences
presented by
JOËLLE GEMAYEL
DEA “Metabolism, Endocrinologie et Nutrition” Université Claude Bernard-Lyon
born May 27, 1974
citizen of Lebanon
accepted on the recommendation of
Prof. Martin Schwab, examiner
Prof. Lukas Sommer, co-examiner
Dr. Matthias Gesemann, external supervisor
2005
DISS. ETH No. 15930
The functional roles of PlexinD1 in chicken nervous system
during development
A dissertation submitted to the
SWISS FEDERAL INSTITUTE OF TECHNOLOGY ZURICH
for the degree of
Doctor of Sciences
presented by
JOËLLE GEMAYEL
DEA “Metabolism, Endocrinologie et Nutrition” Université Claude Bernard-Lyon
born May 27, 1974
citizen of Lebanon
accepted on the recommendation of
Prof. Martin Schwab, examiner
Prof. Lukas Sommer, co-examiner
Dr. Matthias Gesemann, external supervisor
2005
Summary
A functional nervous system results from the coordinated generation and assembly of billions
of neural cells into highly structured and well organized networks. Building blocks for these
networks are neurons, which arise from neural progenitors that are deployed from specialized
neuroepithelia. These neuronal precursors migrate along specific pathways populating
different areas within the developing brain, spinal cord and peripheral nervous system. Once
correctly positioned, differentiated neurons send out axons along highly stereotypical
pathways, specifically linking neurons within different parts of the nervous system,
generating a high number of functional neuronal networks. The use of appropriate migratory
routes as well as directed axonal outgrowth along specific predetermined pathways is
achieved by specific receptor-ligand interactions that are characteristic for each subpopulation
of migratory cells or growing axons.
Several families of cell surface receptors and axon guidance cues involved in directing cell
migration and/or axonal outgrowth have been described over the last decade. One of the
largest families of axon guidance receptors is the plexin family. Plexins and their semaphorin
ligands have been shown to be involved in several aspects of axonal targeting and cell
migration. However, numerous studies describe also functional roles for plexins in the
development outside the nervous system, notably in the development of the cardiovascular
system. In this respect, PlexinD1 (PD1) has been extensively studied in vasculogenesis, and
several groups document its implication in heart development as well as in vessel patterning.
However, PD1 is also expressed in several regions of the developing brain, but no available
data describe its potential role in the nervous system.
Experiments in the first part of this thesis demonstrate that PD1 transcripts are not only found
in mouse brain but also in chicken spinal motor neurons during the period motor axons sort in
the limb plexus. PD1 knock down, using in ovo RNAi, results in motor axon misguidance in
the dorsal as well as the ventral crural nerve trunk, suggesting PD1 involvement in motor
axon guidance and/or sorting of the crural nerve. Interestingly, PD1 loss of function
experiments showed also unexpected defects in dorsal sensory root formation and
abnormalities in the area of motoneuron exit points. In PD1 knock down animals, dorsal roots
are less compact, and several axon bundles grow erroneously into neighboring segments
whereas motor exit points display a severely altered morphology appearing much broader
than usual, and the ventral motor roots show axons with aberrant courses.
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Summary
While the defects in the crural nerve could be easily explained by the lack of PD1 expression
in motoneurons, phenotypes observed in the dorsal and ventral roots support a role for PD1 in
either neural crest migration or the existence of a tight relation between intersomitic vessel
formation and dorsal root development. In the first case, PD1 loss of function in a
subpopulation of neural crest cells would lead to erroneous positioning of Schwann cells and
/or boundary cap cells resulting in outgrowth and defasciculation defects within the ventral
and dorsal roots. Alternatively, PD1 knock down might affect endothelial cells surrounding
the spinal cord, in particular the intersomitic vessels, leading to aberrant formation of these
vessels. Intersomitic vessels could form in close contact with sensory growing axons and
could play a role for correct axonal outgrowth. Their aberrant formation could result,
therefore, in axon guidance defects observed in the present study.
Experiments described in the second and third chapter of this thesis analyze the presence and
expression of semaphorins as well as plexins in the developing chicken spinal cord. Chicken
plexins and semaphorins display very complex, often complementary, but also overlapping
expression patterns that are highly regulated developmentally. These results suggest
multifunctional roles for these proteins, most likely exerting their activities in large
complexes containing intricate combinations of several members.
Résumé
Le bon fonctionnement du système nerveux dépend du correct assemblage de milliards de
cellules neurales générées pendant le développement embryonnaire afin de pouvoir former
des réseaux complexes et structurés.
Les précurseurs des cellules neurales se forment au niveau d’une structure bien spécialisée
appelée l’épithélium neuronal. Une fois générées, ces cellules entament leur migration tout le
long de parcours bien définis afin de former les différentes structures du système nerveux
central et périphérique. Par la suite les cellules neuronales projettent leurs axones tout le long
de voies bien définies, souvent à de très longues distances de leur corps cellulaire pour
finalement de se connecter à leur cible finale.
L’achèvement d’une migration cellulaire correcte ainsi que la navigation précise des axones
jusqu’à leur destination finale dans un système embryonnaire nécessite l’intégrité de familles
de protéines parmi lesquelles les plexines.
Les plexines ont été récemment identifiées et caractérisées; elles constituent une très grande
famille de protéines contenant environ 9 membres et exercent leur action en se liant à leur
ligands, les semaphorines. Plusieurs études décrivent l’implication de ces protéines dans le
développement du système nerveux central mais aussi dans la formation d’autre tissues
embryonnaires.
PlexinD1 appartient à la famille des plexines et a été intensivement décrite comme un facteur
majeur dans le développement du système vasculaire in vitro et in vivo. Bien que l’expression
de cette protéine ait été identifiée dans le cerveau de souris pendant le développement
embryonnaire, aucune étude ne décrit le rôle de plexinD1 dans le développement du système
nerveux central.
Pendant ce travail de thèse, nous avons souhaité investir le rôle potentiel de plexinD1 dans le
développement de la moelle épinière dans l’embryon de poulet. Nous avons établi que cette
protéine est bien exprimée pendant le développement par les neurones moteurs de la moelle
épinière et cette expression correspond au moment où ces neurones s’arrangent dans le plexus
à la base du pied.
Les outils moléculaires visant à bloquer l’expression de plexinD1 en utilisant une technique
appelée in ovo RNAi, ont pu démontrer que l’absence de plexinD1 dans les motoneurones
induit la malformation de certaines branches du nerf crural. De plus, nous avons observé des
anomalies au sein des fibres sensorielles résultant en la fusion des fibres entre elles, bien que
plexinD1 ne soit pas exprimée par les neurones sensorielles mais par les cellules endothéliales
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Résumé
entourant la moelle épinière et les ganglions de la racine dorsale. Afin de pouvoir expliquer
ces aberrations dans la formation des fibres sensorielles, nous avons proposé deux modèles.
Notre première hypothèse suppose que plexinD1 n’est pas exclusivement exprimée par les
cellules endothéliales mais aussi dans les cellules de Schwann et les boundary cap cells. Une
migration erronée de ces cellules menant à leur mauvais positionnement peut expliquer les
anomalies observées. Une seconde explication est que la malformation des fibres sensorielles
peut résulter d’une anomalie dans le développement des vaisseaux sanguins exprimant
plexinD1, en particulier les vaisseaux intersomitiques, qui normalement assistent et guident
les fibres sensorielles pendant leur élongation.
Nous avons également montré dans une deuxième partie de notre travail, l’expression de
toutes les semaphorines et plexines dans la moelle épinière du poulet au cours des différents
stades du développement. Nos résultats démontrent que les ARNm étudiés sont exprimés
d’une façon très complexe et dynamique suggérant l’existence de multiples fonctions exercées
par ces protéines qui très souvent doivent agir en synergie plutôt que séparément.
Acknowledgements
I would like to thank Dr. Matthias Gesemann for giving me the opportunity to do my PhD
thesis in his laboratory and introducing me to the world of axon guidance molecules.
I am very grateful to Professor Martin Schwab, not only for supervising my thesis but also for
always taking time to discuss my data and giving me inputs about the future directions to
adopt in my projects and experiments.
I thank also Professor Lukas Sommer, for being always available for supervising my thesis
work and investing time to discuss my data and results.
This work would not have been possible without the collaboration as well as the precious help
of Professor Esther Stoeckli and all her team. I would like to thank her for giving me the
chance to learn a lot about chicken embryos and helping me all through the in vivo
experiments. All my gratitude goes also toward all the Stoeckli team, especially Rejina Sadhu
for teaching me all I know about chicken and Olivier Mauti, for their help and cooperation
and for making the work at their laboratory very enjoyable.
Many thanks go to my colleagues in the Gesemann groups (Regis, Esther, Pascal, Connie,
Daniele and Peter), for making hard moments possible to survive and for helping me at any
moment with all the energy they have.
Despite the hard moments, a researcher can pass through, the wonderful and international
environment the Hifo offers, make science more efficient and the work in the lab much easier.
Without the great and continuous help of all the neighboring labs, I could not have achieved
my thesis. I would like to thank specially the Schwab group for rendering the institute as a big
family and for helping me always as if I was a member of their team. My special thanks to
Franziska Christ, Lisa Schnell, Regula Schneider, Dana Dodd, Carri Duncan, Irin Maier,
Elisabeth Aloy, Barbara Niederoest and Florence Bareyre.
Special thanks to Roland Schoeb for his continuous support in arranging and printing my
pictures especially during the time I was writing my thesis.
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Acknowledgements
Many thanks also for the Neuhauss Group for their kind support and cooperation, especially,
Ronja Bahadori ,Oliver Rinner, Oliver Biehlmaier, David Belet and Yury Makhankov.
How cheap words can be when confronted to endless love and unlimited support.
This thesis is dedicated with all my love to my family and friends.
To my dad, for filling gaps life made, for teaching me how to be strong and fight for my
dreams, for giving me the great chance of flying away to build a life but mainly for giving me
endless love that one need at any time and any age.
To my brothers Gino and Tony, and my sister Jessy, for believing in me in all circumstances,
for all the laughers and the tears we shared and we will still share, for being four parts of one
entity and finally for your unlimited love that gives me always strength to advance.
This thesis is dedicated with all my love to Christian. I could have not achieved this work
without your daily support and continuous love. For standing by me no matter if the in situ
and cloning worked or failed, despite my long working hours in the lab even during week
ends, for pushing me to advance always forward and to push further my limits.
A mes meilleures amies, Chantal, Rozlaine et Rania pour avoir su être toujours présentes
malgré les distances et les frontières, pour m’avoir connu mieux que je ne me connais et pour
m’avoir aimé sincèrement pour ce que je suis.
How can I ever thank all of you in the Hifo and neighboring institutes who are for me more
than colleagues and for standing by my side during the hardest moment? For Zeina, Ronja,
Elisabeth, Carri, Rejina, and Florence, “thank you” seems meaningless!!! For wiping my tears
and making me laugh over endless coffee breaks, champagne drinking sessions and great
dinners, you are the best souvenir I will take with me from Zurich.
Je n’oublie pas de mentionner aussi tous mes amis bien loin de Zurich qui n’ont cessé de me
soutenir malgré la distance et le temps. Mes plus sincères pensées et remerciements vont vers:
Mme Jomain, Dr. Alain Geloen, Dr. Geneviève Barret et toute l’équipe du Laboratoire de
physiologie Lyon Nord pour avoir été comme une famille pour moi en France, avoir cru en
mes capacités et m’avoir aide à parvenir jusqu’aux bouts de mes ambitions.
Contents
Summary .................................................................................................................................... i
Résumé .....................................................................................................................................iii
Acknowledgements................................................................................................................... v
Contents................................................................................................................................... vii
Part I: Introduction.................................................................................................................. 1
1.1 Nervous system development........................................................................................... 2
1.1.1 Neural cell induction and migration.......................................................................... 2
1.1.2 Neural cell differentiation ......................................................................................... 4
1.1.3 Axon guidance......................................................................................................... 12
1.2 Axon guidance forces..................................................................................................... 17
1.3 Families of axon guidance molecules ............................................................................ 18
1.4 Plexins and Semaphorins ............................................................................................... 20
1.4.1 Semaphorin family .................................................................................................. 20
1.4.2 Semaphorin receptors and receptor complexes....................................................... 26
1.4.3 Semaphorin and Plexins beyond axon guidance ..................................................... 30
1.5 RNAi in chicken spinal cord .......................................................................................... 31
1.6 Goal of the present thesis: Role of PlexinD1 in nervous system development.............. 32
Part II: Paper 1....................................................................................................................... 35
2.1 Introduction .................................................................................................................... 37
2.2 Material and methods ..................................................................................................... 40
2.2.1 in ovo RNAi ............................................................................................................ 40
2.2.2 In situ hybridization ................................................................................................ 41
2.3 Results ............................................................................................................................ 43
2.3.1 Expression of PlexinD1 in chicken spinal cord during motor axon sorting in the
limb plexus ....................................................................................................................... 43
2.3.2 PD1 knock down specifically in chicken embryo spinal cord leads to motor axon
pathfinding errors at the hindlimb level ........................................................................... 44
2.3.3 PD1 knock down animals show fusion of the dorsal root entry zones ................... 46
2.3.4 PD1 knock down animals exhibit defects at the motor exit points of the ventral
roots.................................................................................................................................. 47
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Contents
2.4 Discussion ...................................................................................................................... 49
2.4.1 Chicken spinal motor neurons express PD1 mRNA ............................................... 49
2.4.2 PD1 involvement in the navigation of the crural nerve toward the limb ................ 49
2.4.3 PD1 knock down affects the dorsal roots entry zones and the motor exit points ... 51
2.4.4 PD1 potential binding partner(s) in chicken embryos during development............ 53
Part III: Paper 2 ..................................................................................................................... 55
3.1 Introduction .................................................................................................................... 57
3.2 Material and Methods..................................................................................................... 60
3.2.1 Assembly of chicken semaphorin cDNAs .............................................................. 60
3.2.2 Phylogenetic tree analysis ....................................................................................... 61
3.2.3 Cloning of semaphorin cDNA fragments................................................................ 61
3.2.4 In situ hybridization ................................................................................................ 62
3.3 Results ............................................................................................................................ 63
3.3.1 Identification of Sema3G, a novel member of the class III semaphorins ............... 63
3.3.2 The chicken genome has a reduced number of semaphorin genes ......................... 64
3.3.3 Class III semaphorins are highly expressed in developing motoneurons ............... 66
3.3.4 Floor plate cells expresses high levels of semaphorin V transcripts....................... 66
3.3.5 Class VI semaphorins are highly expressed in boundary cap cells..................Error!
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3.3.6 Semaphorin7A is expressed in endothelial cells and motoneurons ........................ 71
3.4 Discussion ...................................................................................................................... 72
3.4.1 The chicken genome has fewer semaphorin genes that the mammalian genome ... 72
3.4.2 Expression patterns of semaphorins are dynamically regulated during spinal cord
development ..................................................................................................................... 73
Part IV: Paper 3 ..................................................................................................................... 77
4.1 Introduction .................................................................................................................... 79
4.2 Material and methods ..................................................................................................... 81
4.2.1 Assembly of chicken plexin cDNAs ....................................................................... 81
4.2.2 Phylogenetic tree and domain identity analysis ...................................................... 82
4.2.3 In situ hybridization ................................................................................................ 82
4.3 Results ............................................................................................................................ 84
4.3.1 Plexin and Neuropilin genes in chicken.................................................................. 84
4.4 Discussion ...................................................................................................................... 93
Contents
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4.4.1 Mouse plexin genes located on the X sex chromosome are absent in chicken....... 93
4.4.2 Expression patterns of plexins and neuropilins are dynamically regulated during
spinal cord development................................................................................................... 94
4.4.3 PlexinBs and PD1 are only transiently expressed in neurons ................................. 95
4.4.4 PlexinC1 is not expressed in early stages of neuronal development....................... 96
Part V: Conclusion and Outlook........................................................................................... 97
Conclusion and Outlook ........................................................................................................ 98
References ............................................................................................................................. 103
List of Publications............................................................................................................... 117
Curriculum Vitae ................................................................................................................. 119
Part I:
Introduction
1
1 Introduction
1.1 Nervous system development
The nervous system of vertebrates is a very complex structure that coordinates a variety of
functions ranging from the integration of simple sensory and motor information to the
processing of very complex behavioral tasks such as learning. The proper functioning of an
adult nervous system depends mainly on the appropriate generation of different types and
numbers of neural cells, the well-defined migration of these cells to their specific final
position and the correct establishment of highly precise neuronal connections between the
very large numbers of generated neurons.
The first stage in nervous system development is the induction of the neuroectoderm to form a
columnar epithelium. This so-called neural plate is underlaid by axial along with paraxial
mesodermal cells and flanked by epidermal ectoderm. During a process called neurulation,
the neural plate first buckles at its midline to form the neural folds and the floor plate
resulting later, once the dorsal tips of the neural folds fuse, in a tube like structure called the
neural tube. At the dorsal midline of the neural tube, roof plate cells along with neural crest
cells are generated. All neural crest cells subsequently initiate migrations to populate different
area of the developing embryo. As the neural tube folds, cell division starts at its luminal side.
Dividing cells are influenced by several inductive or repressive factors secreted from the floor
plate, the roof plate and the surrounding tissue that contribute to the acquisition of diverse
neuronal cell fates. Neural induction and early regional fate of neural cells appear to be
linked, as it is impossible to separate the induction of neural properties from the acquisition of
anteroposterior regional identity.
1.1.1 Neural cell induction and migration
During early neural development, axial mesodermal cells provide inductive signals to initiate
neural tissue formation. The major pathway of neural induction is mediated by the inhibition
of bone morphogenic protein 4 (BMP4) signaling, which when blocked lead to the formation
of the anterior neural tube. Several structurally unrelated xenopus proteins (follistatin, noggin,
chordin) were shown to inhibit BMP4 activity (Godsave and Slack, 1989). The neural tissue
induced by follistatin, noggin and chordin exhibit an anterior character (Hemmati-Brivanlou
and Melton, 1994; Lamb et al., 1993), suggesting that distinct signaling pathways may be
required for inducing posterior neural tissue. Exposure of the ectoderm to fibroblast growth
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1.1 Nervous system development
3
factor members (FGFs), under conditions in which BMP4 signaling is reduced or eliminated,
leads to the generation of posterior neural plate tissue. Moreover, neural tissue characteristics
of intermediate levels of the neuraxis midbrain and hindbrain can be induced by exposure of
the ectoderm to both noggin and FGF (Lamb and Harland, 1995). Additionally, retinoids
belonging to another class of molecules appear to be involved in the generation of posterior
neural tissue: treatment of embryos with retinoic acid leads to an anterior-to-posterior
transformation in the regional character of the neural tube (Durston et al., 1989; Hill et al.,
1995).
However, in vivo studies demonstrate that BMP4 and follistatin are not the only factors
responsible for neural induction. Mutant animals lacking BMP4, follistatin or other factors
from the Hensen’s node do not exhibit any obvious defect in neural induction, implying that
these factors are not the only players required for neural plate formation (Matzuk et al., 1995;
Winnier et al., 1995). Furthermore, it now appears that neural induction begins prior to the
formation of the organizer region or the node. Thus, different factors expressed potentially in
other regions than the organizer or node such as the mesoderm and endoderm might act on the
neural induction of the ectoderm. These findings suggest that the suppression of BMP
signaling may maintain rather than initiate the process of neural differentiation.
Neuronal progenitors are generated from the rapid division of neural stem cells at the
germinal neuroepithelium of the newly formed neural tube. While symmetric cell divisions
along the luminal side of the neural tube give rise to two similar stem cells, asymmetric cell
divisions lead to the generation of differentiated neuronal and glia cell types migrating toward
the pial surface of the neural tube (Hollyday, 2001).
One hallmark of vertebrate nervous system development is long-range cell migration of
neuronal precursor cells. During the formation of the brain, the spinal cord and the peripheral
nervous system, neuronal precursor cells migrate extensively and undergo significant
rearrangements prior to differentiation into either neurons or glia. Two major patterns of
migratory movements can be distinguished during nervous system development. While
laminated structures such as the cerebellum, the hippocampus and the cerebral cortex are
formed largely due to cell migration along radial glia, other brain areas such as the hindbrain
and the thalamus are also formed by tangentially migrating neurons (Park et al., 2002; Hatten,
1999). Laminated structures built by radial migration exhibit a specific pattern of inside-out
formation where newborn neurons by pass their older siblings to populate more cortical
layers. This type of migration is based on cell-cell interactions between the moving neurons
and the radial glial cells, of which the latter form a migratory scaffold that extends from the
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Part I: Introduction
ventricular zone to the pial surface. This process implies the existence of signals on or near
the glia that can promote migration of neuroblasts in appropriate directions and arrest the
movement at appropriate locations. Identification of a key molecule in radial migration came
from the isolation of a new allele of a mouse mutant, reeler (Rice and Curran, 1999). In these
mutants, late migrating neurons fail to pass their older siblings, leading to a scrambling of the
normal inside-out relationship between birthdates and laminar position. Reelin is concentrated
on the superficial cortical laminae and seems to promote dissociation of neuroblasts from the
radial glia surface.
While glia guided migration is certainly a key component for laminated structure formation, it
has been shown lately that many cortical interneurons arise in subcortical areas rather than in
the cortical ventricular zone and migrate tangentially into the cortex (Hatten, 1999). Migration
of these neurons into the cortex occurs via a tangential pathway and is glia independent.
Tangentially migrating neurons develop a specialized transient structure that is called the
“leading process”. Although a lot remain to be discovered regarding structures and molecules
implicated in tangential cell migration, it is well understood that migrating cells use their
leading process to sense and probe environmental cues in the surrounding (Marin and
Rubenstein, 2003).
1.1.2 Neural cell differentiation
The allocation of cell fate in the central nervous system depends on two signaling systems
that are activated together with the more basic program of neural induction. These two
signaling systems intersect along the rostrocaudal and dorsoventral axes of the neural tube
establishing a grid-like set of positional cues. The initial position of the neuronal progenitor
cells along these two axes determines their exposure to different types and concentrations of
inductive signals and therefore influences directly their fate (Jessell, 2000).
Signaling along the rostrocaudal axis gives rise to the different subdivisions of the nervous
system: the forebrain, midbrain, hindbrain and spinal cord (Lumsden and Krumlauf, 1996).
Some of the factors playing a role in neural induction seem also to determine rostrocaudal
differentiation. Relatively few signaling factors (retinoid, TGF β signaling, sonic hedgehog
(Shh), FGFs and Wnts) account for many features of regional cell specialization within the
anterior neural tube (Tanabe and Jessell, 1996). All these factors are proposed to function in
different locations or developmental windows and their combinatorial actions in a single
region or cell can establish regional patterning and neuronal diversity. Moreover, some
1.1 Nervous system development
5
inductive signals such as Shh can act as a gradient signal to induce different subtypes of
neurons at different concentration (Poh et al., 2002).
While general cues along the dorsoventral axis lead to the diversity of cell types within each
rostrocaudal subdivision (Pituello, 1997), two primary signaling factors appear to induce
“ventralization” or “dorsalization” of the neural tube. As neural tube folding occurs, the
underlying mesodermal cells of the notochord initially provide a signal identified as Sonic
Hedgehog (Shh) that “ventralize” the developing neural tube. Shh initiates the differentiation
of a special set of neuroepithelial cells located immediately adjacent to the notochord called
“floor plate”. Floor plate cells are the first cells in the neural plate to show signs of overt
differentiation (Patten et al., 2003). Once induced, the floor plate cells themselves secrete Shh
and thereby provide patterning information for neural cell types. Shh seems to function as a
morphogene which can induce differentiation of distinct types of ventral cells at different
concentration thresholds (Ericson et al., 1997). The graded activity of Shh subsequently
activates different transcription factors in progenitor cells placed at various locations along
the ventro-dorsal axis of the neural tube and subdividing a previously uniform territory into
distinct domains. These transcription factors are expressed in a combinatorial manner in the
dividing multipotential progenitors of the ventricular zone. Importantly, these transcription
factors can repress each other in specific combinations leading to the translation of a transient
graded response to Shh into establishment of sharp boundary between two cell populations
(see Figure 1) (Briscoe et al., 2000).
Fig. 1: Three phases of Shh-mediated ventral neural patterning. a) Shh mediates the repression or the induction
of different transcription factors at variable threshold. Shh signaling defines five progenitor domains in the
ventral neural tube. c) The relation between neural progenitors (p) domains and the positions at which postmitotic neurons are generated along the dorsoventral axis of the ventral spinal cord (adapted from(Jessell,
2000)).
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Part I: Introduction
Concomitant with the ventral polarization of neural tissue, signals from the epidermal
ectoderm adjacent to the lateral edge of the neural plate seem to impose the dorsal pattern.
BMP4, BMP7 and Dorsalin-1 are expressed in the dorsal aspect of the neural tube and are
implicated in dorsalizing activity (Nguyen et al., 2000).
The region of the ventral neural tube that gives rise to motoneurons subsequently generates
also oligodendrocytes. Thus, the effect of Shh and BMPs extend beyond determining ventral
neurons identity as it also contributes to the location of the founder cells of the
oligodendrocyte lineage. Spinal cord oligodendrocytes arise from cells in the motor neuron
domain (Richardson et al., 1997; Miller, 2002), and this localization is dependent on the local
expression of Shh that is antagonized by BMPs signaling.
Interestingly, most of the neuronal subtypes generated within the spinal cord are represented
at all segmental levels, raising the issue of whether rostrocaudal positional information
contributes significantly to the establishment of neuronal subtype identity at the spinal cord.
1.1.2.1 Motor neuron differentiation and migration
While all spinal motoneurons neurons derive from a single ventral progenitor domain, they
acquire many distinct subtype identities. The identity of a motoneuron is defined by its
location in the CNS and its final synaptic connectivity in the PNS. Motor neurons
differentiate exclusively in the ventral spinal cord, but once they become post mitotic, they
may migrate to occupy more dorsal positions on the ipsilateral side of the spinal cord
(Hollyday, 2001). In higher vertebrates, motor neurons with common target projections are
aligned into longitudinally oriented columns (see Figure 2). These columns occupy distinct
and discontinuous domain along the rostrocaudal axis of the spinal cord. This columnar
organization is closely linked to specific formation of axon tracks and neuronal connections
(Jessell, 2000). Motor neurons in the medial motor column (MMC), innervating hypaxial
(back muscles) and epaxial (body wall muscles) muscles, are present along the entire
rostrocaudal axis. Whereas, neurons of the lateral motor column (LMC), which innervate the
limbs, are found only at brachial (forelimb) and lumbar (hind limb) levels and column of
Terni (CT) neurons project to the sympathetic ganglia (Landmesser, 1978a).
At a second level of organization, neurons within the same motor column are segregated into
medial and lateral division and project axons along different trajectories. Within the LMC,
motoneurons in the medial and the lateral divisions project to ventral and dorsal limb muscles
respectively (Landmesser, 1978b), whereas motor neurons from the medial division of MMC,
innervate the epaxial muscle and the lateral one target the body wall muscles. A final level of
1.1 Nervous system development
7
organization is reached when motor neurons within each division of the LMC segregate into
discrete pools to innervate specific muscles in the limb. Anatomically defined motor neuron
subclasses are also molecularly distinct, as defined by the restricted expression pattern of
specific transcription factors. The main different columnar subclasses of motor neurons can
be distinguished by the combinatorial expression of LIM homeodomain (LIM-HD) proteins
(Tsuchida et al., 1994), whereas individual motoneurons pools within the LMC can be
classified based on their expression of members of ETS proteins (Lin et al., 1998).
Fig. 2: Columnar organization of motor neuron subtypes in the chick spinal cord. The target specificity of motor
neuron subtypes are defined by distinct combinations of LIM-HD transcription factors. Axonal pathways of
motor neurons subtypes are represented in transverse sections at the brachial and thoracic levels. The expression
of LIM-HD transcription factors in individual motor neurons subtypes is shown by color-coding. bw, body wall;
dlb, dorsal limb bud musculature: dm, dermamytome; sg, neurons of the sympathetic ganglia; vlb, ventral limb
bud musculature. LMCL (blue), lateral half of lateral motor column; LMCM (green), median half of lateral motor
column; MMCL (orange), lateral half of medial motor column; MMCM (red), median half of medial motor
column (adapted from (Shirasaki and Pfaff, 2002)).
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Part I: Introduction
1.1.2.2 Interneurons differentiation and migration
Spinal interneurons constitute a large number of functionally distinct classes of cells present
at the dorsal as well as the ventral part of the neural tube. They appear as small groups of
neurons showing no particular distinguishing features or spatial organizations. However, they
comprise a large number of separate neuronal populations with widely varying functions
greatly outnumbering the amount of motor neurons. While some spinal interneurons process
sensory information, others modulate motoneuron activity or coordinate activity at different
spinal levels, and some relay sensory or proprioceptive information to the brain.
The distinguishing markers for dorsal neural tube progenitor cells seem to be bHLH
transcription factors rather than homeodomain transcription factors seen in the ventral spinal
domains. Interestingly at the dorsal spinal domain a combination of bHLH and HD domain
are likely to define progenitor domains.
Interneurons are induced by BMPs signaling factors secreted by the roof plate and/or
epidermal ectoderm. The down stream cascade is not very well understood but wnt1 and
wnt3a are likely to be candidates because wnt1 at least is induced by BMP signaling
(Panchision et al., 2001). Some transcription factors induced by BMP signaling seem to
modulate the timing of cell cycle exit of progenitor cells thus controlling the cell number.
Although BMPs have been considered the key signaling molecules originating from the dorsal
midline, other signaling molecules are expressed in this region during generation of dorsal
interneurons, such as members of FGF and Wnt families. It is still not clear if FGF play a role
in dorsal interneurons formation, however Wnts are implicated in patterning, proliferation and
cell determination (Wodarz and Nusse, 1998).
Dorsally located interneurons can be grouped into eight distinct subtypes defined by either the
expression of specific transcription factors, initial migration, dependence on roof plate
signals, or date of birth (Gross et al., 2002; Muller et al., 2002).
In mice, there are six early-born post mitotic dorsal interneuron populations called dI1-dI6,
which are generated between E10-E12.5 and two later-born post mitotic populations called
dILA and dILB produced between E11-E13. All populations are defined by the expression of
specific homeodomain transcription factors. These neurons can be further classified by their
dependence on roof plate signaling for formation: class A (dI1-dI3) are dependent on, and
class B (dI4-dI6, dILA/B) independent of, roof plate signals (Lee et al., 2000). While class A
neurons end up in deep dorsal horn layers to form relay interneurons that project
contralaterally to transmit sensory information to higher brain regions (Bermingham et al.,
2001), dILA/B neurons migrate to more superficial laminae of the dorsal horn and develop into
1.1 Nervous system development
9
association neurons, which serve to integrate sensory input and characteristically project
ipsilaterally (see Figure 3).
Fig. 3: Patterning of dorsal spinal cord showing the different classes of dorsal interneurons, the time they are
generated and the transcription factors implicated in their differentiation in addition to their migratory paths
(adapted from (Helms and Johnson, 2003)).
In contrast to dorsal interneurons, ventral interneurons are differentiated by graded Shh
induction and can be subdivided into four domains expressing differential transcription
factors. The V3 domain is located close to the floor plate below the differentiating motor
neurons whereas the V0, V1 and V2 are located dorsally to motor neurons and are separated
by sharp boundary depending on combinatorial expression of various transcription factors
(see Figure 1). Interestingly, the generation of certain sets of interneurons in the dorsal-most
region of the ventral neural tube is Shh independent. These interneurons subtypes can be
induced by a parallel signaling pathway that is mediated by retinoids derived from the
paraxial mesoderm and possibly from the neural plate cells (Pierani et al., 1999).
1.1.2.3 Neural crest differentiation and migration
A very special population of neural precursor cells, called neural crest cells, is found at the
dorsal site of the early neural tube. Neural crest cells form at the border between the neural
plate and the future epidermis and delaminate from the neuroepithelium in a rostro-caudal
wave. Neural crest cell precursors are multi-potent progenitors and can differentiate into a
very broad range of cell types. They form most of the peripheral nervous system including
10
Part I: Introduction
sensory, sympathetic and enteric neurons and glia. Additionally they give rise to melanocytes,
smooth muscle, dermis, connective tissue, cartilage and bone (Le Douarin and Dupin, 2003).
The embryological origin of crest cells is little understood. While it has been known for some
time that inductive interactions between the neural plate and the non-neural ectoderm underlie
the initial specification of neural crest cells at the neural plate border, the molecular nature of
these signals has been less clear. Earlier in vitro experiments in chicken suggested a role for
bone morphogenic proteins (BMPs) as endogenous neural crest inducers (Basler et al., 1993),
(Liem et al., 1995). Indeed, many BMPs can mimic the ability of epidermis to induce the
formation of crest cells from neural plate explants. However, recent studies in frog and
chicken provide strong evidence that Wnts are rather used as endogenous neural crest
inducers (Garcia-Castro et al., 2002). Both gain of function as well as loss of function
experiments in chicken show that Wnt6 plays a primary role as a neural crest cell inducer.
Furthermore, a novel extracellular glycoprotein called Noelin-1 has been implicated in neural
crest development. Noelin-1 is expressed at the lateral edges of the neural plate where it
appears to maintain the competence of neural epithelial cells to form neural crest cells.
Neural crest cells migrate along specific pathways to form a very diverse range of cells,
ranging from melanocytes in the skin to neurons in the sensory ganglia. Although very little is
known about the signals governing the differentiation of melanocytes, in vivo and in vitro
analysis demonstrates that the decision to differentiate into pigment cells or neural derivatives
is made early during migration. In zebrafish, medially located neural crest cells are induced
by local wnt-1 and wnt-3a signals to form pigment cells, whereas lateral cells form neurons
(Dorsky et al., 1998). In contrary, the autonomic nervous system induction seems to be BMP4 and BMP-7 dependent. These two factors are expressed in the dorsal aorta that is adjacent to
the sympathetic ganglia and seem to induce sympathetic neurogenesis (Reissmann et al.,
1996). Whereas sensory neurons are specified by the bHLH proteins Neurogenin-1 (ngn-1)
and -2 that are expressed in the neural crest precursor of sensory neurons (Ma et al., 1999),
several in vitro and in vivo studies implicate neuregulin, ErbB2 and ErbB3 as well as a
transient effect of the notch signaling in the specification of glial cells (Morrison et al., 2000).
Following their induction in the dorsolateral neural tube, neural crest cells undergo an
epithelial to mesenchymal transition and begin to migrate. Recent evidence suggests that in
addition to being implicated in neural crest induction, BMP signaling is also involved in
downstream aspects such as the onset of neural crest migration (Sela-Donenfeld and
Kalcheim, 1999). The earliest known response to this induction is the expression of two
transcription factors slug and snail (Nieto et al., 1994; Sefton et al., 1998). Following
1.1 Nervous system development
11
expression of snail in epithelial cell lines, the cells become mesenchymal and migratory
(Cano et al., 2000).
A number of molecules that either serve as substrates for migrating crest cells or delimit
migratory pathways by forming repulsive boundaries have been identified (Robinson et al.,
1997; Bronner-Fraser, 1993; Perris and Perissinotto, 2000). In avian trunk, neural crest cells
travel along two distinct pathways. Some cells emerging from the dorsal neural tube, adopt a
ventromedial path through the rostral but not caudal somitic sclerotome, whereas other crest
cells travels dorsolaterally in a uniform manner between the somites and overlying ectoderm.
While cells of the ventrolateral path aggregate bilaterally along the developing spinal cord to
form DRGs and sympathetic ganglia, migratory cells of the dorsolateral path contribute
mainly to the formation of pigment cells (see Figure 4) (Krull, 2001).
Fig. 4: Avian trunk neural crest cells travel on two distinct pathways after emigration from the neural tube.
Schematic diagram representing a longitudinal view at the trunk region of an avian embryo. Some trunk crest
cells (red) emerge from the dorsal neural tube and travel ventromedially, through the rostral but not caudal,
somitic sclerotome. Other neural crest cells (black) migrate dorsolaterally in a uniform manner between the
somites and overlying ectoderm. DM, dermamyotome; Scl, sclerotome; No, notochord; Ao, aorta, Ec, ectoderm;
NT, neural tube; R, rostral; C, caudal (adapted from (Krull, 2001)).
12
Part I: Introduction
1.1.3 Axon guidance
While cell migration is a prerequisite for the correct positioning of neurons within the
developing embryo, directed axon outgrowth is essential for the accurate wiring of the
nervous system during development. To form functional contacts with appropriate targets,
axons grow in a highly stereospecific manner over considerable distances. The precise wiring
of the nervous system occurs mainly by two types of mechanisms: early acting mechanisms
independent of neural activity (molecular mechanism) and later-acting activity based
mechanisms.
Studies performed over the last two decades in an attempt to understand the early acting
mechanisms have provided a detailed understanding of the cellular interactions between
axonal growth cones and their surroundings that direct pathfinding.
1.1.3.1 Pathfinding of motor axons
During development, motor neurons acquire distinct identities that are reflected in their
choice of specific axon pathways and their synaptic targets. Motor axon pathfinding occurs in
a stepwise manner and is dependent on the differential action of guidance cues, which are
serially deployed at discrete locations along the axonal pathway. Thus, a motor neuron
journey is divided into several stages: axonal exit from the CNS, growth along a shared
common pathway and navigation to and away from different choice points (Schneider and
Granato, 2003).
The first step in a motor axon’s pathway is to correctly exit the CNS and project its axon into
one of the segmental nerves, connecting the CNS to the periphery. Motor axons grow initially
away from the floor plate and penetrate the neuroepithelium at specific exit points. Co-culture
experiments in vitro demonstrate that all classes of motor axons are repelled when placed
adjacent to floor plate cells (Guthrie and Pini, 1995). Additionally, all spinal motor axons
choose a single ventral root within each somitic hemi-segment to leave the spinal cord. A
specific type of neural crest-derived cells, called the boundary cap cells, is located at these
exit points (Niederlander and Lumsden, 1996). These cells serve, most likely, as gatekeeper
between the CNS and the PNS and may play a role in inducing segmental nerve patterning
(Golding and Cohen, 1997). Support for this speculation comes from a recent studies which
demonstrates that boundary cap cells are not implicated in attracting motor axons to their exit
point but rather in keeping their cell body confined in the CNS (Vermeren et al., 2003).
The mesoderm adjacent to the spinal cord is divided into a series of segmented blocks, the
somites, which become partitioned into sclerotome and dermamyotome components. After
1.1 Nervous system development
13
motor axons emerge from the spinal cord at exit points, they traverse the sclerotome
component of the somite only within its rostral half. Repulsive and attractive activities,
derived from the caudal and rostral halves of the sclerotome respectively, impose the periodic
arrangement of motor nerves exiting from the spinal cord (Keynes and Stern, 1984). Despite
emerging evidence from in vitro studies proposing the involvement of diverse proteins in
spinal motor axon segmentation, to date no in vivo data exist that demonstrate the implication
of any axon guidance or cell adhesion molecules in this process. Thus, while caudal
sclerotome cells play an essential role in guiding the segmental exit of motor axons, the
identity of the cues by which they do so remains unclear.
Once motor axons have left the CNS, different classes of motor neurons innervate variable
muscle targets following predefined pathways. For example, motor neurons of the MMCL
send axons to innervate epaxial muscle and grow towards the dermamyotome, whereas other
motor axons of the MMCM avoid the dermamyotome and navigate ventrolaterally to innervate
hypaxial muscles (see Figure 2).
At the hindlimb level, spinal motor axons appear to grow along well-defined highways
making pathway changes at specific choice point regions. In chicken, multiple spinal nerves
converge to form the crural (segment L1-L3) and sciatic nerve trunk (segment L4-L8) (see
Figure 5). Motor axons in each of these nerve trunks grow to the base of the limb, called the
plexus area, where they pause for 24 hours before entering the limb bud. Apparently, axons
wait in the plexus for limb maturation to occur (Varela-Echavarria et al., 1997). Within the
plexus region, axon trajectories are highly individualistic with many abrupt turns, perhaps
reflecting a process of active sorting (Tosney and Landmesser, 1985a). This process appears
to be at least in part dependent on the differential expression of a set of cell adhesion
molecules by different motor axons.
Motor axons of the LMCM and LMCL further subdivide at the limb base to form distinct
ventral and dorsal nerve trunks. This step seems to be dependent on target-derived
chemoattractants such as HGF and guidance cues produced in the developing limb.
Additionally, some tissues act as barriers to axons as they navigate to the hindlimb. Motor
axons seem to avoid the perinotochordal mesenchyme and the pelvic girdle precursor tissue
(Tosney and Oakley, 1990; Oakley and Tosney, 1991).
Once motor axons are near their target muscle, they have to recognize and form synapses with
the appropriate muscle fiber. Motor neurons that innervate muscles are matched with their
target in such a way to generate a precise topographic map (Laskowski and Sanes, 1987).
14
Part I: Introduction
Fig. 5: In avian at the hindlimb level, multiple spinal nerves converge to form the crural (segment L1-L3) and
sciatic nerve trunk (segment L4-L8). Each nerve divides further into dorsal or ventral trunk targeting different
muscles of the hindlimb (adapted from (Landmesser, 2001)).
1.1.3.2 Interneurons and midline crossing
In the CNS of a wide variety of bilaterally symmetric organisms, different interneurons
project axons along specific trajectories, which are parallel or perpendicular to the midline.
The best-studied interneurons are the commissural interneurons, whose cell bodies are located
in the dorsolateral spinal cord extending axons in a circumferential path toward the floor
plate.
The earliest commissural axons or “pioneer” axons travel along the lateral edges of the spinal
cord until they reach the floor plate whereas the later projecting axons or the “followers”
extend along the same pathway. After reaching the midline, these commissural axons cross
1.1 Nervous system development
15
through the ventral most third of the floor plate, subsequently turning orthogonally at the
contralateral side of the floor plate (Bovolenta and Dodd, 1990). While another subtype of the
dorsally located commissural interneurons, known as association interneurons execute right
angle turns and extend parallel to the floor plate along the ipsilaterally-projecting lateral
funiculus (Colamarino and Tessier-Lavigne, 1995b). A more ventrally positioned population
of interneurons that develop in a region between the floor plate and motor neurons extend
their axons along the ipsilateral longitudinal pathway within the ventral funiculus (Yaginuma
et al., 1990).
Regardless of the fact whether specific interneuron populations cross or do not cross the
midline, all axonal interneurons are directed by the floor plate that affects noticeably the
behavior of a growth cone in its vicinity.
1.1.3.3 Projections of dorsal root ganglia
A subpopulation of multipotent neural crest cells migrates along stereotypic pathways and
coalesces at specific locations to form the spinal sensory ganglia also called the dorsal root
ganglia (DRG). Spinal sensory neurons comprise a morphologically and functionally
heterogeneous group of neurons, specialized in the transfer of different sensory modalities
(Farinas et al., 2002). Each DRG innervates a full array of targets in the periphery, including
skin, muscle, and viscera. Individual DRG neurons connect to specific types of sensory
receptors, conveying information about position in space (proprioception), pain (nociception),
distension, or touch (mechanoception) to the CNS. Neurotrophins play an essential role in the
maintenance of a normal complement of neurons since all sensory neurons require the
presence of at least one neurotrophin during development . Although the neurotrophic
hypothesis postulates that neurons become dependent on a particular neurotrophin when their
axons encounter their final targets there is evidence demonstrating that neurotrophins are
expressed during early development before axon-target recognition and are therefore also
implicated in gangliogenesis (Buchman and Davies, 1993; Farinas et al., 1996).
Anatomical and physiological data, document very well the early peripheral projection of
sensory neurons. Sensory fibers innervating the hindlimb are established in a precise orderly
manner (Honig, 1982). During normal development, sensory axons appear to grow on the
axons of adjacent motoneurons and always project to the same muscles as the neighboring
motor neurons (Tosney and Landmesser, 1985b). Therefore, the absence of motor neurons
causes also severe missprojections of sensory neurons. Interestingly sensory neurons
innervating skin or muscle in the periphery appear less rigidly specified than motoneurons
16
Part I: Introduction
and have more flexibility in their pathway and target choices. At the stages when innervations
are being established, cutaneous as well as muscle afferents, unlike motoneurons, may not yet
have acquired specified identities and the ability to recognize and respond selectively to their
appropriate targets (Adams and Scott, 1998).
Additionally the central projections of sensory neurons follow a strict spatio-temporal pattern
with different DRG neurons having central arborisations in the spinal cord that are specific for
the sensory modality. In chicken, cutaneous and muscle axons of sensory afferents reach the
spinal cord by stage 23, stalling there for 24 hours in the primordium of the dorsal funiculus
before extending axons rostrally as well as caudally. At around stage 28 central projections
begin to enter the gray matter of the spinal cord. While cutaneous afferents branch frequently
remaining in the dorsal horn (Mendelson et al., 1992), proprioceptive axons reach the vicinity
of motor neuron dendrites without branching and form functional contacts around stage 32
(Davis et al., 1989). The segregation of afferent inputs into laminar-specific projections is
dependent on diffusible factors, integral proteins and/or extra cellular matrix proteins (Ozaki
and Snider, 1997). Both types of sensory neuron projections (peripheral and central) are
established precisely and correctly from the outset, and neither cell death nor retraction of
axons plays a role in the development of appropriate connectivity. During the initial stage of
DRG axonal growth, surrounding “non target” tissues such as dermamyotome, the notochord,
and the ventral spinal cord release strong chemorepulsive signals that inhibit DRG axons
(Masuda et al., 2003) in vitro. However, despite many advances made in the identification of
axon guidance cues affecting sensory neurons outgrowth in vitro, our understanding about the
molecular mechanisms required in vivo for the axonal pathfinding of these neurons remain
fragmentary and incomplete.
1.1.3.4 Molecular mechanism of cell migration and axon guidance
Neurons extend axons to appropriate targets with the help of a very specialized structure at
the tip of the advancing axon called the “growth cone”. On the molecular level, our
understanding about how a growth cone recognizes which path it has to take and how it
reaches its precise synaptic target while encountering numerous potential, other targets on its
way, is still fragmentary. Nevertheless, many advances have been made toward a better
understanding of the molecular mechanisms of growth cone based on axon guidance. The
identification and characterization of different families of axon guidance molecules and the
use of functional tests to discover the biological roles of these guidance cues in vivo, has
greatly advanced our knowledge.
1.2 Axon guidance forces
17
To accomplish the complex task of reaching a distant target, the initial axon trajectory seems
to be broken into short segments that terminate at specialized cells forming intermediate
targets also called “choice points”. These intermediate targets provide guidance information
enabling axons to select and initiate growth along a particular segment. Growth cones that
approach an intermediate target reduce their speed and assume a more complex morphology
with more filopodia, presumably to better sample the environment. Therefore, two types of
cellular behaviors can characterize axon growth: a simple linear growth along “highways”,
punctuated by a more complex decision-making behaviors at choice points as axons switch
from one highway to another. In insects, some intermediate targets are made up of cluster of
“guidepost cells”, whose ablation results in misrouting of axons that normally contact them
(Raper et al., 1984; Bastiani et al., 1984). Additionally, the wiring process of the nervous
system occurs in a stepwise manner. While pioneer axons navigate through an axon free
environment when the embryo is still relatively small, later developing axons benefit from the
scaffold provided by earlier projecting axons allowing the later developing ones to grow
along pre-existing tracts or fascicules for at least some of their trajectory only switching from
one fascicule to another at specific choice points. This “selective fasciculation” strategy
simplifies the assembly of large nervous system network by allowing follower axons to use
predefined paths (Kolodkin et al., 1992).
Despite their intrinsic characteristics, growth cones are not endowed with a predestined and
autonomous ability to find their correct targets. For this, they must respond to and depend on
cues produced by their surroundings (O'Connor, 1992). Therefore, a growth cone must
express a specific set of receptor on its leading edge that reacts appropriately to the
surrounding molecules (Tessier-Lavigne and Goodman, 1996). Many proteins playing a role
in axon guidance have been identified in invertebrates and vertebrates and it appears that both
the mechanisms and the molecules are conserved among species. In fact, evolutionary
conservation of guidance molecule is so great that insights gained in invertebrates can be
immediately relevant to vertebrates, and vice-versa (Tessier-Lavigne and Goodman, 1996).
1.2 Axon guidance forces
Over a century ago, Cajal already proposed long-range chemoattraction as a mean to guide
axons to specific targets. Only in the late 1970s, Gunderson and Barrett showed that sensory
neuron growth cones can indeed respond to a gradient of a protein called nerve growth factor
(NGF). Although NGF is unlikely to be a long-range chemoattractant in the developing
organism, this study initiated a resurgence of interest in chemotropism. Later, experiments
18
Part I: Introduction
performed by Lumdsen and colleges (Lumsden and Davies, 1983), demonstrated that cocultures of neurons and their target areas, embedded in a collagen matrix produce stable
molecular gradients originating from these target tissues that are capable of attracting
extending axons. More recently, the finding that axons could also be repelled in vitro by
tissues that these axons normally avoid, provided strong evidence that guidance information is
not only attractive but also repulsive (Fitzgerald et al., 1993; Colamarino and TessierLavigne, 1995a). Axons can also be guided at short-range by contact-mediated mechanisms
involving non-diffusible cell surface or extracellular matrix (EMC) molecules. While the
process of contact attraction has been implicated in “selective fasciculation”, in which growth
cones confronted with several pre-existing axons fascicules select a specific pathway (Raper
et al., 1984), contact repulsion of axons has been extensively documented in the retinotectal
system.
Thus, the growth cone appears to be guided by four general mechanisms: contact and
chemoattraction as well as contact and chemorepulsion (Tessier-Lavigne and Goodman,
1996).
The presence of discrete classes of diffusible and non-diffusible factors, some attractive while
others repulsive are however not as strict as one might expect. Instead, axon guidance
molecules appear to act through conserved mechanisms activating or inhibiting oftenoverlapping signal transduction pathways. The same molecule may function as growth
inhibitor in one assay but as an attractant in another, suggesting that most of guidance
molecules are not exclusively attractive or repulsive but rather bifunctional, playing variable
roles in guiding different growth cones (Tessier-Lavigne and Goodman, 1996). Additionally it
was recently shown that a growth cone can exhibit attractive as well as repulsive responses to
the same guidance cue depending on the level of cytoplasmic cyclic AMP (cAMP) (Ming et
al., 1997; Song et al., 1997). Much of the current studies of axon guidance are directed toward
defining the precise complement of forces orchestrating particular guidance decisions. An
individual axon might be “pushed” from behind by a chemorepellent, “pulled” from afar by a
chemoattractant and “hemmed” in by attractive or repulsive local cues. These forces appear to
play in concert all together to ensure accurate and specific axon guidance.
1.3 Families of axon guidance molecules
In the early 1990s, the introduction of powerful in vitro assays to detect guidance activities in
the developing vertebrate nervous system and the growing interest in invertebrate axon
guidance led to the discovery of several conserved families of axon guidance molecules. The
1.3 Families of axon guidance molecules
19
most prominent axon guidance cue families are the netrins, slits, semaphorins, and ephrins.
These molecules are not the only known guidance molecules but are by far the best studied
ones.
Netrins are highly conserved proteins that while exerting a chemoattractant activity on
commissural axons (Culotti and Merz, 1998) are responsible for a chemorepellent effect
(Colamarino and Tessier-Lavigne, 1995a) on growth cones of trochlear motor axons. In
nematodes, Unc6/Netrin is known to mediate its activity through binding to unc-5 and unc-40
receptors (Hedgecock et al., 1990). Biochemical and genetic studies have confirmed that Unc40/DCC and Unc-5 receptors function as netrins receptors in several different species (Culotti
and Merz, 1998; Keleman and Dickson, 2001), guiding a variety of axons and cells in vivo as
well as in vitro (Winberg et al., 1998a; Yee et al., 1999).
Slit proteins represent another family of conserved axon guidance molecules Slits are large
secreted proteins acting as midline repellent whose function appears to be conserved along
evolution. Interestingly, slit was also purified as a factor that stimulates sensory axon
branching and elongation (Wang et al., 1999), suggesting that Slits like netrins may be
multifunctional. However, the best-studied functions of slits are its involvement in midline
guidance. In the Drosophila embryo, slit acts as a short-range repellent to prevent ipsilateral
projecting axons from crossing the midline and commissural projecting axons from recrossing
the midline (Battye et al., 1999). In vertebrates, slit has been shown to play a crucial role in
the formation of the optic chiasm and in axon guidance at the ventral midline guidance
through binding to its receptor Roundabout (Robo) preventing inaccurate crossing of
ipsilaterally as well as contralaterally projecting axons (Kidd et al., 1998; Plump et al., 2002;
Erskine et al., 2000).
One of the largest families of mainly repulsive axon guidance molecules are the ephrins.
Ephrins were initially characterized as membrane bound proteins, involved in guiding
vertebrate retinal axons to appropriate topographic locations in the tectum. Ephrins and Eph
receptors fall into two classes; ephrin-As, which are anchored to the membrane by a GPI
linkage that bind to EphA receptors and ephrins-Bs, which are transmembrane proteins
binding to EphB receptors. In the visual system, topographic mapping of retinal axons along
the anterior-posterior axis depends on ephrin-A mediated repulsion. Ephrin-A are expressed
in a gradient in the tectum and their receptors EphA are expressed in a complementary
gradient in the retina. Thus, retinal axons with successively higher EphA levels map to
successively lower points along the ephrin-A gradient (Wilkinson, 2001).
20
Part I: Introduction
However, emerging evidence nowadays suggest that ephrins and their receptors control axon
guidance in many other places in addition to their ability to signal bi-directionally and
mediate both attraction and repulsion.
1.4 Plexins and Semaphorins
1.4.1 Semaphorin family
Semaphorins belong to a large family of cell surface or secreted proteins and represent one of
the best-studied families of axon guidance molecules. Initially all semaphorin family
members were believed to mediate inhibitory actions on axon pathfinding, branching or
targeting, but nowadays there is increasing evidence that some semaphorin proteins also play
a role in chemoattraction and cell migration (Kolodkin et al., 1992; Luo et al., 1993; Matthes
et al., 1995; Messersmith et al., 1995).
Semaphorins are defined by a highly conserved 500 amino-acid extracellular domain, the
Sema domain, which contains 14 to 16 cysteines and some conserved N-linked glycosylation
sites. More than 20 different semaphorin members have been identified up to now. Based on
their degree of sequence similarity and domain organization semaphorins can be grouped into
seven different subclasses. While semaphorins belonging to subclass I and II are found
exclusively in invertebrates, semaphorins belonging to the subclasses III-VII are specific for
vertebrates. While in general the majority of semaphorin subtypes are transmembrane
proteins, semaphorin members belonging to the subclasses II, III, and VII, represent
exceptions, which are secreted or GPI anchored molecules respectively (see Figure 6)
(Tamagnone and Comoglio, 2000).
While the cytoplasmic domains of transmembrane semaphorins share no significant
homology to any known protein and have very little similarity to each other, the structural
conservation of the Sema domain implies that this part is essential for semaphoring binding
and function. Interestingly, all semaphorins form disulfide-linked homodimers, whose
oligomerization is crucial for semaphorin-signaling, and their biological activity seems to be
determined by a relatively short stretch of amino acids within the Sema domain (Koppel and
Raper, 1998; Klostermann et al., 1998; Koppel et al., 1997).
1.4.1.1 Semaphorin Class I & II
SemaI, the first semaphorin member to be identified, is a transmembrane protein expressed on
subsets of fasciculating axons and on stripes of epithelial cells in the grasshopper limb bud
(Kolodkin et al., 1992). Antibody perturbation experiments in grasshopper demonstrated that
1.4 Plexins and Semaphorins
21
SemaI plays a crucial role in steering a pair of sensory neurons to the limb bud by regulating
axon defasciculation and branching.
The other invertebrate semaphorin, SemaII is expressed transiently in a subset of motor
neurons and a single thoracic muscle during motor neuron outgrowth and synapse formation.
Gain-of-function analysis in Drosophila, show that ectopic SemaII expression in muscle cells
inhibits normal synaptic terminal arborisation of two different motor neurons subtypes
without affecting the growth cones of other motor axons (Matthes et al., 1995). The fact that
SemaII ectopic expression does not affect the oriented growth of axons toward these muscles
suggests that SemaII rather serves as a selective target-derived signal inhibiting the formation
of specific synaptic terminal arbores than influencing early aspects of axon guidance (Matthes
et al., 1995).
Fig. 6: Semaphorins are phylogenetically related proteins, sharing sema domains similar to the cystein rich Metrelated sequences (MRSs). Semaphorins are divided into 7 subgroups, Class 1 & 2 existent only in invertebrates
whereas Class 3-7 are exclusive for vertebrates. Class 2&3 are secreted semaphorins while all the other members
are membrane bound; Class 1, 4, 5 and 6 are transmembrane and Class 7 contains GPI linked members.
Abbreviations: G-P/ IPT motif, glycine-proline repeat/ immunoglobulin-like fold; GPI, glycosyl
phosphatidylinositol,; MRS, Met-related sequence (adapted from (Tamagnone and Comoglio, 2000).
1.4.1.2 Semaphorin Class III
Class III Semaphorin members share three structural motifs, the 500 amino acid sema
domain, a C-2 type immunoglobulin (Ig) domain, and a positively charged carboxy terminal
tail. Secreted semaphorins act as diffusible signals, although their diffusion distance might be
limited because they are tethered to the cell surface and extracellular matrix in vivo by the
charged sequence at the C-terminus of the protein (Bagnard et al., 2000).
So far, Sema3A is the most extensively studied semaphorin among all class III members. It
has been identified by two independent approaches. While Kolodkin and colleges identified
vertebrate Sema3A based on its homology to SemaI (Kolodkin et al., 1993), Raper’s group
22
Part I: Introduction
identified Sema3A as the major growth inhibitory protein for sensory axons(Luo et al., 1993).
Sema3A is widely expressed in the developing central and peripheral nervous system as well
as in several tissues surrounding the spinal cord where it is responsible for collapsing a broad
range of different axons. Sema3A is expressed, in the perinotochordal mesenchyme as well as
mesenchymal cells around the spinal cord and DRG with the exception of nerve exit points
(Puschel et al., 1995). Additionally Sema3A transcripts have been found in the dorsal aorta,
the connective tissue separating the developing muscles and in the caudal half of the
sclerotomes suggesting a potential involvement of Sema3A as a repulsive guidance molecule
in axon pathfinding of spinal neurons (Shepherd et al., 1996; Giger et al., 1996).
Despite its relatively broad expression in the developing embryo the effect of Sema3A is
highly specific as in dorsal root ganglia only NGF sensitive sensory axons are affected
(Messersmith et al., 1995). Nevertheless, it also collapses axons of sympathetic ganglia
neurons (Puschel et al., 1995), motor neurons (Shepherd et al., 1996; Varela-Echavarria et al.,
1997), sensory neurons from the trigeminal, facial and vagal cranial ganglia, in addition to
axons from olfactory (Kobayashi et al., 1997), pontine (Rabacchi et al., 1999), cortical
(Bagnard et al., 1998) and hippocampal neurons (Chedotal et al., 1998; Steup et al., 1999).
Interestingly, Sema3A cannot induce collapse of retinal ganglion growth cones,
demonstrating that Sema3A is specific to a subset of neuronal growth cones. Recent studies
show that Sema3A can also suppress the migration of avian trunk neural crest cells in vitro
(Eickholt et al., 1999).
The analyses of Sema3A function in vivo using gene disruption in mice gave rather surprising
results. While the nervous system appears normal with the exception of some minor defects in
sensory axon projections, mutant mice exhibit sever malformation of skeletal structures in
addition to pronounced and selective hypertrophy of the right ventricle of the heart resulting
in a high degree of mortality around birth (Behar et al., 1996). The fact that Sema3A is highly
expressed in rat heart, sclerotomes, ribs and pelvic girdle during development (Wright et al.,
1995) might account to explain such defects after Sema3A gene disruption. Intriguingly,
Sema3A mutant mice from a different laboratory are viable until adulthood, do not show any
heart defects and have normal dorsal root projections. However, they appear to have
disturbances in some peripheral nerve projections that are affected by Sema3A in vitro. Many
cranial nerves (trigeminal, facial, vagal, accessory and glossopharyngeal) as well as spinal
nerves are highly defasciculated and some nerves overshoot their targets (Taniguchi et al.,
1997). These data imply that Sema3A expressed in the surrounding tissues may drive axons
into fascicules and that its absence might provoke axons in the periphery of mutant animals to
1.4 Plexins and Semaphorins
23
enter regions that are normally strongly repulsive. Surprisingly, despite the wider aberrant
paths adopted by some nerves, the overall target recognition seems to be well achieved in the
knock out mice. The moderate defects of the nervous development in vivo observed in
Sema3A might be explained by the presence of six additional class III members with
overlapping distributions and potentially overlapping activities. To answer the question
whether Sema3A and other Class III semaphorin play combinatorial roles in nervous system
development, it will be essential to generate not only knock out animals for other Class III
members but also to disrupt the expression of several semaphorin members at the same time.
Other members of semaphorin class III have also been analyzed to a lesser extend than
Sema3A. To date, only Sema3C and Sema3F have been reasonably well characterized in vitro
and in vivo. In vitro Sema3C repels neurites from CA1 and medial septum but has no effect
on CA3, dentate gyrus and entorhinal axons. Interestingly Sema3C appear to exert a dual role
in vitro depending on the neuronal population studied. It acts as a repellent on sympathetic
axons, has no effect on DRG neurons but in contrary attracts axons of cortical explants
(Bagnard et al., 1998). As for Sema3A, Sema3C mutant mice show no obvious defects in the
development of the nervous system. The terminations of septal fibers in the inner and outer
molecular layers of the dentate gyrus are normal. However, mutant mice display severe
congenital cardiovascular defects and die soon after birth because of the interruption of the
aortic arch and improper septation of the cardiac outflow tract (Feiner et al., 2001). This
phenotype is consistent with the expression of Sema3C in the mesenchyme surrounding the
branchial arch arteries and in the myocardial cuff as well as the cardiac outflow tracts
suggesting a role for Sema3C in guiding migratory cardiac crest cells.
Sema 3F is another semaphorin member that appears to govern the pathfinding of certain
nerves in the CNS and PNS in vitro and in vivo. It is expressed in embryonic hippocampal
regions in mice at the time of axonal outgrowth (E15 to E17) and shows repulsive activity on
CA1, CA3 and dentate gyrus axons in vitro (Chedotal et al., 1998). In addition, it is also
capable of collapsing trochlear motor axons (Giger et al., 2000). Sema3F is also present in the
vomeronasal organ and in cells that flank the path of vomeronasal sensory neurons as well as
in the accessory olfactory bulb and the main olfactory bulb. Interestingly in vivo studies,
based on the generation of Sema3F mutant mice, demonstrate that Sema3F is crucial for axon
fasciculation and segregation but not for target recognition in the olfactory system (Cloutier et
al., 2004). Additional analyses of the CNS of Sema3F mutant mice reveal that this protein is
essential in the ventral forebrain for anterior commissure axons to fasciculate and decussate
normally at the CNS midline as well for the formation of the infrapyramidal tract (Sahay et
24
Part I: Introduction
al., 2003). Furthermore, Sema3F seems to be important for the proper organization of specific
cranial nerve projections in the PNS, as Sema3F mutants show severe defects in the trochlear
nerve where only few axons exit the hindbrain-midbrain junction. Moreover, the occulomotor
nerve is largely defasciculated but maintains its peripheral trajectory (Sahay et al., 2003).
The other members of Semaphorins class III have been less well studied and understood. In
vitro experiments demonstrate that Sema3B can repel sympathetic axons (Takahashi et al.,
1998) but no data were reported about the role of Sema3D and Sema3E in vitro or in vivo
(Koppel et al., 1997; Raper, 2000).
1.4.1.3 Membrane attached Semaphorins
In contrast to Semaphorins in subclasses III and IV, Sema5A and Sema5B belonging to the
subclass V lack the IgG domain and have instead seven carboxy-terminal thrombospondine
repeats followed by a short intracellular C-terminus that is unique for this subclass. The
thrombospondine repeats, which account for more than half of the protein sequence, are
components of different extracellular matrix proteins and have been shown to promote
potentially neurite outgrowth. Both semaphorins are expressed during early murine
embryogenesis and adult tissues in mutually exclusive domains. Sema5A is present in axial
and paraxial mesodermal tissues, limb bud, optic disc and nerve whereas Sema5B expression
is exclusively restricted to the neuroepithelium along the entire anterio-posterior axis (Adams
et al., 1996 117). While there is no available information concerning in vitro as well as in vivo
functions of Sema5B, data on Sema5A clearly suggest its functional involvement in axon
guidance.
The fact that Sema5A is present in the paraxial mesoderm prior to the arrival of growing
axons to the limb bud, suggests a potential role for this semaphorin not only in directing crest
cell migration and in sensory or motor axonal outgrowth but also in somitogenesis (Adams et
al., 1996 117). Additionally, Sema5A seems to be implicated in vitro in collapsing retinal
axon growth of rodents (Goldberg et al., 2004), which has been confirmed in vivo by the use
of function blocking antibody. Perturbation of Sema5A function leads to retinal axons
straying out of the optic nerve bundle indicating that Sema5A normally help ensheathing the
retinal pathway (Oster et al., 2003). This repulsion activity is mediated through the Sema
domain of the protein. The thrombospondine repeats do no show any repulsive or attractive
activity in vitro on axonal growth of retinal ganglion cells (Goldberg et al., 2004).
Semaphorins grouped into the subclass VI, display a relatively simple extracellular part, in
which only the highly conserved Sema domain is present. However, their intracellular stretch
1.4 Plexins and Semaphorins
25
is quite long comparing to other Semaphorin members, suggesting that this part in Class VI
semaphorin might have unique functions. Four different Class VI family members have been
identified so far (6A-6D), with Sema6A, being the best studied one.
In mice, several tissues express Sema6A during embryonic development; however, this
expression is strongly down-regulated perinataly. In the nervous system, Sema6A mRNA is
present in the spinal cord and the DRGs and in different regions of the Brain. While early
embryonic expression is restricted to the ventral spinal cord, at later stages expression is also
observed in the dorsal spinal cord in areas of lamina I and II. Although Sema6A is, absent
from all cervical and thoracic sympathetic ganglia, it is expressed in skeletal muscles near the
tissue encircling the ganglia as well as in glossopharyngeal and cochlear ganglia (Zhou et al.,
1997). Interestingly, in vitro Sema6A acts as a repellent on E8 chicken growth cone of
sympathetic ganglia and sensory NT-3 and NGF sensitive DRG neurons.
Surprisingly, abolishing Sema6A transcripts in vivo by gene trapping, gives mutant mice that
are viable and fertile without any behavioral or morphological defects. Although most of the
tracts in the CNS appear normal, the thalamocortical tract shows pathfinding defects at the
caudal level. While most rostral projections are similar to wild type, the caudal part fails to
turn through the internal capsule and instead projects towards the amygdala (Leighton et al.,
2001). Recent studies have demonstrated the existence a Sema6A isoform that contains a
longer cytoplasmic tail. This semaphorins 6 variant is capable of directly linking Ena/VASP
proteins, which are known to play a crucial role in actin filament dynamics (Klostermann et
al., 2000).
Sema6B is the least understood semaphorin member of all Class VI Semaphorins. Sema6B
expression appears early in development of dorsal root ganglia, somites and brain of rodents
(Kikuchi et al., 1997). In contrast to other semaphorins, Sema6B appear to be homogenously
expressed throughout the entire spinal cord. Additionally, Sema6B expression persists in
adulthood in many tissues such as brain, heart, and lungs. Interestingly, Sema6B in vitro binds
specifically the SH3 domain of the proto-oncogene c-src suggesting that it can trigger
intracellular signaling and act as a receptor (Eckhardt et al., 1997).
Sema6C is expressed in rat spinal cord as well as in dermamyotome, DRGs and the notochord
during development. Later on, Sema6C message can be found also in cranial motor ganglia,
the olfactory epithelium and the cerebellar plate. Postnataly, Sema6C expression is present in
different cerebellar layers, pontine and inferior olive nuclei as well as in adult skeletal muscle
tissue and many CNS structures (Kikuchi et al., 1999). Thus, temporal expression of Sema6C
in neurons and in their target areas during development suggests a potential role for this
26
Part I: Introduction
protein in axon guidance of motor and sensory neurons as well in directing commissural or
cerebellar neurons. Sema6C shows, growth cone collapse activity on DRGs in vitro and the
target regions of DRG neurons express Sema6C during development. Two isoforms of
Sema6C, derived from alternative splicing, were identified and their expression pattern is
regulated in tissue and age dependent manner (Kikuchi et al., 1999).
Sema6D, the last member of Class VI semaphorin to be characterized, exists in five different
splice variants whose expression patterns are tissue specific. In vitro Sema6D has been shown
to induce growth cone collapse of DRG and hippocampal neurons but had no effect on
cortical neurons (Qu et al., 2002). Sema6D knock down or over expression in mice or chicken
causes morphological abnormalities of the cardiac tube as well as of the neural tube,
suggesting that Sema6D is involved in cardiac morphogenesis and in the neural tube
formation (Toyofuku et al., 2004b; Toyofuku et al., 2004a). Such phenotypes are in
accordance with the observed expression pattern of Sema6D in normal mice where high
levels of Sema6D expression are observed in the developing heart and neural folds.
Semaphorins belonging to subclass IV and VII have been studied mainly in the immune
system where they exert immuno-modulatory effects. However increasing evidence, also
point to a potential implication of these proteins in the nervous system development. Sema4D,
the only semaphorin Class IV member studied in the nervous system development, has been
shown to stimulate outgrowth of embryonic DRG sensory neurons in vitro (Masuda et al.,
2004). Similarly, Sema7A is expressed in several structures of the rat embryonic brain and
promotes in vitro the growth of numerous central axons such as the vomeronasal epithelium,
the olfactory epithelium, the olfactory bulb and the cortex as well as the peripheral axons of
the dorsal root ganglia. However, the disruption of Sema7A gene in mice, leads only to minor
defects in the lateral olfactory tract whose axons fail to branch or to project to the most caudal
region of the olfactory cortex (Pasterkamp et al., 2003).
1.4.2 Semaphorin receptors and receptor complexes
Over the last couple of years, several Semaphorin-binding proteins potentially involved in
semaphorin signaling have been identified. The most prominent members can be grouped into
two different subclasses: the Neuropilins and the Plexins. While neuropilins seem to bind
different members of Class III semaphorin, plexins have been shown to interact directly with
all different subclasses of semaphorins.
1.4 Plexins and Semaphorins
27
Fig. 7: Plexins are subdivided into four subfamilies (plexins A to D). Their extracellular domains contain a sema
domain and MRS repeats. The extracellular domain of plexin B contains potential cleavage sites for furin-like
convertases. Neuropilin-1 and Neuropilin-2 are transmembrane proteins containing a very short cytoplasmic tail.
Abbreviations: G-P/ IPT motif, glycine-proline repeat/ immunoglobulin-like fold; SP domain, sex-plexin
domain; CUB domain, complement-homology domain; MAM domain, meprin/A5/mu-phosphatase homology
domain, MRS, Met-related sequence (adapted from (Tamagnone and Comoglio, 2000)).
1.4.2.1 Neuropilins
Neuropilins (NP) are transmembrane proteins lacking a signaling-competent cytoplasmic
domain. Two neuropilins genes have been identified in the genome of birds and mammalians
(NP-1 and NP-2); however, no neuropilin gene has been identified in invertebrates. The
extracellular domain of neuropilins contains two repeated complement-binding domains
(CUB domains a1/a2 domains), two coagulation-factor-homology domains (b1/b2 domains)
and a juxtamembrane meprin/A5/mu-phosphatase (MAM) homology domain (see Figure 7).
While the CUB a1/a2 and b1/b2 domains seem to be essential to define the profile of
semaphorin specificity, the MAM domain seems to be crucial for the functionally required NP
non-covalent oligomerization on the cell surface.
Neuropilins bind to the members of semaphorin class III with different affinities. While NP-1
binds with high affinity to Sema3A (He and Tessier-Lavigne, 1997; Kolodkin et al., 1997) but
not to Sema3F, NP-2 binds to Sema3F with higher affinity than Sema3A (Chen et al., 1997).
The fact that NPs aggregate into dimers and NP-1 with NP-2 can for heterodimers when co-
28
Part I: Introduction
expressed suggests a model whereby NP-1 homodimers confer responsivity to Sema3A; NP-2
homodimers are responsible for responsivity to Sema3F and a cooperation of both neuropilins
perhaps by heterodimerization confers responsivity to Sema3C (Chen et al., 1997; Takahashi
et al., 1999; Renzi et al., 1999).
NP-1 is expressed in particular classes of neurons, including most peripheral sensory neurons,
autonomic neurons of the sympathetic ganglia, motor neurons in the spinal cord and the
medulla, neurons in the hippocampal formation, retinal ganglion cells and olfactory receptors
and their target neurons in the olfactory bulb (Kawakami et al., 1996). The expression of NP1 in the nervous system is developmentally regulated in both peripheral and central nervous
systems. NP-1 appears first in newly differentiated neurons and persists throughout the period
of active axonal growth disappearing only after the frameworks of neuronal circuits have been
established.
Mice, carrying a null mutation for the NP-1 gene, are embryonic lethal and exhibit analogous
but somewhat stronger axon guidance defects to those observed in Sema3A knock out mice.
In addition to the abnormal defasciculation of cranial nerves, peripheral nerves in the trunk
are also defasciculated, DRG packages appear to be loose and sympathetic neurons are
displaced. Interestingly, sympathetic neuronal precursors are not accumulated at their initial
target sites around the dorsal aorta in NP-1 mutants (Kitsukawa et al., 1997), a defect also
observed in Sema3A knock out mice (Taniguchi et al., 1997). However, the migratory
pathways of sympathetic neuron progenitors within the rostral sclerotomes are normal
(Kawasaki et al., 2002), suggesting that NP-1-mediated Sema3A activity may play a
functional role in prohibiting incorrect migration of neural crest cells of sympathetic neuron
lineage, promote aggregation of sympathetic neurons and influence sympathetic neurons
fasciculation. NP-1 deficiency is also associated with altered vascularization in the brain and a
variety of defects in the large vessels of the heart outflow (Kawasaki et al., 1999). The
observed high degree of lethality may or may not be a result of interfering with semaphorin
signaling, since NP-1 also interacts with VEGF by increasing its affinity to its receptor. The
strongest phenotype observed in NP-1 mutants in comparison to Sema3A mice might reflect
the functional loss of more than one class III semaphorin and/or the loss of semaphorinindependent functions of NP-1.
NP-2 has an overall structure, which is similar to NP-1. The protein exists in six different
isoforms that are generated by alternative splicing (Chen et al., 1997). NP-2 is expressed in
multiple areas in the developing CNS and PNS as well as many non-neural tissues. The
expression pattern partially overlaps with NP-1 but is mostly complementary. In contrast to
1.4 Plexins and Semaphorins
29
NP-1, NP-2 expression is not detected in the heart or in capillaries but is only found in the
dorsal aorta. Two independent groups have recently reported the phenotype of knockout mice
for NP-2. These mice are generally viable until adulthood and exhibit defects of axon
fasciculation and targeting of selected cranial nerves and central projections (Chen et al.,
2000; Giger et al., 2000). The mutants also lack the trochlear nerve and showed irregular
trajectories of the occulomotor nerve but have no clear abnormalities in the projections and
trajectories of spinal nerves.
While neuropilins are clearly required for axon guidance of several neuronal populations, the
absence of an obvious intracellular domain to propagate signaling suggests the existence of
additional proteins to form functional signal-transducing complexes (Nakamura et al., 1998).
In addition, the absence of neuropilin genes in invertebrates and the lack of binding of nonClass III semaphorins to neuropilins support the existence of other semaphorin binding
proteins.
1.4.2.2 Plexin family
First evidences that plexins are Semaphorin-binding proteins were obtained not by studies in
the nervous system but by experiments carried out to identify a receptor for virally encoded
semaphorins. Using this approach a protein called VESPR was identified which was later
renamed PlexinC1. Subsequently nine different plexins have been identified in the
mammalian genome, which can be sub-grouped into four different classes (Tamagnone et al.,
1999). In contrast to neuropilins, plexins are also found in invertebrates, suggesting that they
represent functional receptors for Class I and II semaphorins (Winberg et al., 1998b).
All plexins are large integral membrane proteins with a highly conserved cytoplasmic tail.
Interestingly at their amino-terminus, they contain, as their putative ligands, a highly
conserved Sema domain, suggesting that an interaction between both proteins is mediated by
Sema-Sema interactions. In addition to the Sema domain, the plexin extracellular domain is
characterized by two or three Met-related sequence repeats (MRS). The large cytoplasmic
moiety of plexins contains a strikingly conserved sex-plexins (SP) domain, which is likely to
trigger novel signal-transduction pathways (Raper, 2000; Tamagnone and Comoglio, 2000)
(see Figure 7). The SP domain of plexins is unrelated to any other domain found so far,
though its primary sequence is highly conserved among family members and across
evolution, suggesting that plexins share common biochemical functions and signaltransduction pathways.
30
Part I: Introduction
Initially plexins were studied in Drosophila where PlexinA was found to bind directly to the
transmembrane Class I Semaphorins and controls important aspects of axon guidance
(Winberg et al., 1998b). In contrast to plexin-semaphorin interactions in invertebrates,
biochemical and cellular studies in vertebrates, suggest that members of the Plexin-A
subfamily form stable complexes with NP-1 or NP-2 rather than interacting directly with
semaphorins. Such complex formation does not depend on the presence of the ligand (Raper,
2000; Takahashi et al., 1999; Tamagnone et al., 1999; Rohm et al., 2000) nevertheless,
Plexins do not simply provide a signaling moiety to NPs but actively influence their binding
efficiency for the different subsets of secreted semaphorins (Takahashi et al., 1999; Rohm et
al., 2000). Moreover Plexins, have been shown to also interact directly with transmembrane
or GPI anchored semaphorins, influencing axon guidance and other developmental processes
in a neuropilin independent way (Tamagnone et al., 1999; Comeau et al., 1998; Winberg et
al., 1998a). In mice, PlexinA subfamily members are widely expressed in the central and
peripheral nervous system and are spatio-temporally regulated. PA-1 and PA-2 mRNA
expression is limited to some neurons in the DRG and absent from sympathetic ganglia
(Murakami et al., 2001) whereas PA-3 appears to be expressed in all peripheral ganglia,
including trigeminal and vagal ganglia, in addition to DRG and sympathetic ganglia.
Additionally, PlexinA3 is strongly expressed in the whole spinal cord and PlexinA2 is
expressed selectively in the dorsal spinal cord (Suto et al., 2003) whereas PlexinA4 is the
most abundant plexins in DRG.
Targeted disruption of PlexinA3 gene demonstrates that this plexin plays a role in
fasciculating the ophthalmic branch of the trigeminal nerve and regulates the development of
hippocampal projections in vivo (Cheng et al., 2001). However, PlexinD1 knock out animals
display cardiac defects but unrelated to cardiac crest migration. PlexinD1 seems unnecessary
for cardiac crest migration but appear implicated in outflow tract septation, development of
aortic arch artery and intersomitic vessels sprouting (Gitler et al., 2004).
Despite the generation of a large number of mutants for different semaphorin and plexin
genes, our understanding about the biological functions in vivo of most of these proteins
families in the nervous system development remain limited.
1.4.3 Semaphorin and Plexins beyond axon guidance
Semaphorins and plexins are highly expressed in several tissues outside the nervous system
where they play very important roles in normal and pathological situations, including
cardiovascular development, immune system formation, and tumor genesis. The
1.5 RNAi in chicken spinal cord
31
transmembrane Sema4D was shown to modulate the functions of T and B-lymphocytes (Hall
et al., 1996; Delaire et al., 1998), and two non-neurotropic viruses encode semaphorin like
molecules might interfere with the immune system of the host contributing to the immune
evasion, protecting the virus from being destroyed. Interestingly, Sema3C and 3E are also
over-expressed in invasive and metastasizing tumor cells, possibly mediating cell dissociation
and/or protection from apoptosis (Yamada et al., 1997; Christensen et al., 1998). In contrast to
this, Sema3A was reported to induce apoptosis of selected sensory and sympathetic neurons
(Gagliardini and Fankhauser, 1999; Shirvan et al., 1999).
Another function for Sema3A seems the inhibition of endothelial cell motility through
competition between Sema3A and vascular endothelial growth factor (VEGF) for NP-1
binding . Emerging evidence suggests that in pathological situation as diverse as nerve injury
and tumor progression, the specific expression of semaphorins is modulated. Semaphorins
seem to be a perfect example of a cell-cell communication code, exploited by a variety of
cells and in different instances from embryo development to adult pathology . Despite all the
advances over the last years, biological functions for the majority of semaphorins remain
incomplete representing important challenge for the future.
1.5 RNAi in chicken spinal cord
Plexins and semaphorins are broadly expressed in several different tissues outside the nervous
system making the investigation of their potential roles in the nervous system in vivo very
hard to tackle. So far, the generation of mutant mice for different semaphorin and plexin
genes gave little insights on the implication of these genes in axon guidance, cell migration,
or synapse formation during nervous development. Nevertheless, the broad and very often
complementary expression of semaphorins and their receptors in the central and peripheral
nervous system strongly argues for multiple yet undetected functional roles for these
molecules. To understand the potential roles of some of these proteins in spinal cord
development, the chicken embryo has proven to be a powerful model system (Pekarik et al.,
2003).
The chicken embryo has served as a classic model system for developmental studies due to its
easy access for surgical manipulations and a wealth of data about chicken embryogenesis.
Recently, the method of in ovo RNAi has led to a revival of the chicken system. Nevertheless,
due to the lack of appropriate genetic knock out possibilities, functional studies in chicken
embryos have been limited to antibody perturbation and dominant negative expression assays.
With the sequencing of the chicken genome near completion, this approach provides a
32
Part I: Introduction
powerful opportunity to examine the function of chicken genes. While in ovo electroporation
has been effectively used earlier for ectopic or over expression analyses, the injection of long
double stranded RNA (ds RNA) into the spinal central canal followed by electroporation
represents an excellent tool to study loss of function phenotypes. Interestingly, this method
allows the targeting of specific subpopulations of neural cells within the developing spinal
cord depending on embryonic age at the injection time. Ds RNA injected- around stage 14
(E2) targets, in addition to spinal neurons, also crest cell derivatives, whereas injections after
stage 16 (E2.5) specifically targets spinal cells, and injections later than stage 18 (E3) fails to
target motor neurons (Pekarik et al., 2003).
Thus, it seems appealing to use the RNAi technique in chicken embryo to knock down
different plexins or semaphorins in crest cells and/or spinal cells with the intention to
understand better their function in spinal cord development in vivo.
1.6 Goal of the present thesis: Role of PlexinD1 in nervous
system development
Despite the numerous studies describing the role and function of PlexinD1 in the vascular
system development in vivo, very little is known about the involvement of this protein in axon
guidance and/or cell migration in the nervous system formation (Gitler et al., 2004; TorresVazquez et al., 2004). However, several lines of evidence show the expression of PlexinD1 in
the central nervous system of mice and rats (van der Zwaag et al., 2002), (unpublished data).
Our expression analysis using two chicken EST clones matching the PlexinD1 sequence
revealed that PlexinD1 is not only expressed in endothelial cells but also in motor neurons.
Expression in chicken motor neurons is highly regulated developmentally, and the highest
expression levels correspond well with the timing motoneurons sort in the limb plexus and
project into different nerve trunks.
PlexinD1 knock down experiments lead to several defects in motor axon guidance and
surprisingly also to aberrant growth of sensory neurons at the dorsal root entry zones. Our
data strongly suggest that PlexinD1 besides influencing directly motor axon outgrowth also
affects directly or indirectly the sensory dorsal root afferents as well as the ventral motor
roots.
In an attempt to investigate whereas any member of the semaphorin family might correspond
to a potential binding partner(s) mediating PD1 signaling in vivo, we performed a detailed
spatio-temporal expression study to analyze the distribution of all semaphorins in the
1.6 Goal of the present thesis: Role of PlexinD1 in nervous system development
33
developing chicken spinal cord. Different plexin and semaphorin members display
complementary as well as overlapping expression patterns in chicken spinal cord, suggesting
the presence of large functional complexes between different compounds that are responsible
for mediating their biological activity in vivo.
34
Part II:
Paper 1
Functional knock down of PlexinD1 in chicken results in misguidance of
motor axons and alterations of dorsal and ventral roots organization
2 Dummyheading
35
Functional knock down of PlexinD1 in chicken results in misguidance of
motor axons and alterations of dorsal and ventral roots organization
1
Joelle Gemayel, 2Rejina Sadhu, 1Regis Babey, 2Esther T. Stoeckli, and 1Matthias Gesemann
1
Brain Research Institute, University of Zurich, and department of biology, ETH Zurich, and
2
Institute of Zoology, University of Zurich,
Winterthurerstrasse 190, 8057 Zurich, Switzerland
correspondence to:
[email protected]
phone:
+41 44 635 3283
fax:
+41 44 635 3303
key words:
36
chicken embryo, PlexinD1, motoneurons, vascular system, crest cells
Abstract
Plexins belong to a large family of molecules mainly implicated in directing axonal
outgrowth, tissue vascularization, and cell migration. PlexinD1 (PD1), a unique member of
the plexin super family, has been extensively studied in vascular system development.
However, very little is known about its potential function in nervous system formation. In this
paper, we show that PlexinD1 mRNA is expressed in chicken spinal motor neurons at a time
corresponding to the period of motor axon sorting in the limb plexus. PlexinD1 knock down
using in ovo RNAi lead to several defects in motor axons pathfinding of the crural nerve
trunk. Interestingly, we also observed abnormalities in axon sorting at the dorsal root entry
zones and the ventral motor root exit points, phenotypes that cannot simply be explained by
the knock down of PlexinD1 in motor neurons. Our data, combined with earlier reports
demonstrating the role of PlexinD1 in intersomitic vessels formation in mice, suggest a novel
role for PlexinD1 in the migration of a subpopulation of neural crest cell derivatives and/or a
tight relation between intersomitic vessels sprouting and spinal root formation.
2.1 Introduction
The formation of precise and accurate sensory and motor connections, are a prerequisite for
perceiving and integrating environmental information and for performing complex motor
tasks. The fundamental basis for the formation of sensory and motor nerves are neural crest
cells that delaminate from the neural crest shortly before or during neural tube closure, and
motor neurons that are formed by progenitor cells located in the ventral half of the spinal cord
(deLapeyriere and Henderson, 1997; Baker and Bronner-Fraser, 1997; Jessell, 2000).
Motor neurons located along the antero-posterior axis of the embryo sort into different motor
neuron pools, such as the lateral motor column (LMC), the medial motor column (MMC) or
the column of Terni (CT) (Landmesser, 1978b). Within the different subgroups, neurons can
be easily identified either based on their position within the developing spinal cord and/or
their axonal trajectory in the limb bud. Despite various studies attempting to explain
motoneuron pathfinding in the chicken embryo, our understanding of the molecular
mechanisms implicated in axonal targeting is still fragmentary. Initial axon outgrowth occurs
away from the floor plate towards a region called the motor exit point (Guthrie and Pini,
1995). After exiting the spinal cord, motor axons converge to reach the limb base where they
pause for 24 hours in the limb plexus, undergoing intensive sorting prior to dividing into a
dorsal and ventral nerve trunks (Varela-Echavarria et al., 1997). In the hindlimb, a lateral
37
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population of LMC motoneurons, called LMCL send axons through the dorsal nerve trunk to
innervate dorsal muscles, whereas axons of motor neurons located in the medial part of the
LMC (LMCM) navigate along a ventral path to innervate the ventrally developing limb
muscles (Tosney and Landmesser, 1985b). The crural nerve trunk, which innervates anterior
muscles, further subdivides into two major branches innervating the anterior dorsal thigh and
the anterior ventral thigh respectively. The final targeting towards the different peripheral
muscles is accomplished by additional branching events increasing the complexity of motor
axon guidance. For some parts of their path, motor axons intermingle with axons of sensory
neurons that have their cell bodies in the dorsal root ganglia (DRG).
Sensory neurons are generated from neural crest cells, which can give rise to a wide variety of
cells ranging from neurons, to smooth muscle cells, cardiac cells, melanocytes and a specific
type of glia located at the dorsal root entry zones (DREZs) and the motor exit points called
the boundary cap cells (Knecht and Bronner-Fraser, 2002; Golding and Cohen, 1997).
Precursors of sensory and sympathetic neurons populating the DRG and the sympathetic
ganglia the migrate between stage 14 (E2) and stage 18 (E3) along a ventrolateral pathway,
passing through the rostral half avoiding the caudal half. In contrast to this melanocyte
precursors migrate after stage 18 (E3) along a dorsal pathway (Krull, 2001). After completing
migration, dorsal root sensory neurons extend axons bi-directionally forming the dorsal roots
and together with motor axons the spinal nerve. Dorsal root afferent axons reache the
primordium of the dorsal funiculus (DF) at stage 23 (E4) entering the spinal cord via the
dorsal root entry zones (DREZs) (Davis et al., 1989). The dorsal roots acquire a highly
organized pattern during development in which adjacent dorsal roots are equally spaced
exhibiting a stereotypic segmental organization.
So far, little is known about molecules and mechanisms that are responsible for the formation
of highly stereotypic sensory and motor networks. While some axon guidance molecules
(ephrinA4) (Helmbacher et al., 2000) and transcription factors (lhx3 and lim-1) (Sharma et
al., 1998; Sharma et al., 2000) have been shown to direct the ventral or dorsal path of motor
axons, very little is known about molecules involved in axon sorting at the limb bud plexus.
The plant lectin peanut agglutinin (PNA)-binding glycoproteins, ephrins, netrins, Hepatocyte
growth factor (HGF), extracellular matrix molecules including F-spondin and tenascin
(Davies et al., 1990; Debby-Brafman et al., 1999; Faissner and Kruse, 1990; Flanagan and
Vanderhaeghen, 1998) have been shown to exert certain repulsive and attractive influences on
motor axons. Interestingly, Semaphorins have been shown to influence pathfinding and cell
migration of a broad range of neurons in both the central as well as peripheral nervous system
2.1 Introduction
39
(Shepherd et al., 1996; Puschel et al., 1995; Eickholt et al., 1999). Two receptor families have
been shown to bind semaphorins directly or indirectly: plexins and neuropilins (Puschel,
2002). While plexins are transmembrane proteins with an extracellular semaphorin (Sema)
domain, a cysteines-rich motif and a conserved intracellular plexin-specific sex-plexin (SP)
domain (Tamagnone and Comoglio, 2000), neuropilins have a highly conserved extracellular
MAM domain but only a very short cytoplasmic tail (Takahashi et al., 1999). Due to the
absence of an intracellular neuropilin domain implicated in signal transduction, it is believed
that semaphorin signaling is mainly transmitted through the activation of plexins (Raper,
2000). In vertebrates, nine different plexins have been identified so far, that based on their
structural similarities have been grouped into four subfamilies (A-D) (Tamagnone et al.,
1999).
Plexins have been reported to mediate multiple biological functions including axon guidance
and cell migration (Rohm et al., 2000; Winberg et al., 1998b; Cheng et al., 2001; Brown et al.,
2001; Gitler et al., 2004). PlexinA3 for example, mediates a semaphorin dependent
fasciculating activity in the hippocampal commissural pathway (Cheng et al., 2001), whereas
PlexinA2 is implicated in semaphorin dependent cardiac crest cell migration (Brown et al.,
2001). Interestingly, plexins and semaphorins are also expressed in many tissues outside the
nervous system, especially in endothelial cells and the heart, a finding that goes hand in hand
with recent studies demonstrating defects in vascular system development in plexin and
semaphorin knockouts. Disruption of the PlexinD1 (PD1) gene in mouse leads to severe
cardiovascular defects such as outflow tract septation, pharyngeal arch arteries malformation
and intersomitic vessels disorganization (Gitler et al., 2004). Similarly, PD1 knock down in
fish leads to angiogenesis defects of intersomitic vessels although heart and main vessels
formation are otherwise normal (Torres-Vazquez et al., 2004). While these two initial studies
suggested a role for a PD1-neuropilin-sema3A complex in intersomitic vessel formation, a
more recent study demonstrates that a neuropilin independent Sema3E-PD1 interaction is
required for accurate vascularization of the intersomitic vessels (Gu et al., 2004).
Surprisingly, although PlexinD1 message has been shown to be expressed not only in the
vascular system but also in several regions of the developing nervous system (van der Zwaag
et al., 2002 161), no potential role for this protein in nervous system development has been
documented.
In this paper, we now show that in the chicken embryo, PlexinD1 is expressed in spinal motor
neurons around the time motor axons reach and initiate sorting in the limb plexus. PlexinD1
knock down using in ovo RNAi leads to motor axons misguidance and disorganization of the
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dorsal and ventral spinal roots. This suggests that PlexinD1 besides its important role in
vascularization is implicated in multiple new functions during nervous system development.
2.2 Material and methods
2.2.1 in ovo RNAi
2.2.1.1 Production of long double stranded RNA (ds RNA)
Double stranded RNA for targeting PlexinD1 was produced from the following chicken EST
clones ChEST697F11 and ChEST867E24. ESTs were linearized using NotI or EcoRI and.
Two µg of linearized plasmid were mixed with a final concentration of 4mM dNTPs (Roche),
two µl T3 or T7 RNA polymerase (15U/ µl; Roche), 4 µl of 5X transcription buffer and 0.5 µl
RNasin (30 U; Promega) in a total volume of 20 µl. After completion of transcription (37°C
for 4 hours), DNAse I (Roche) was added and the RNA was extracted using once acidic
phenol-chloroform (25:24:1 vol/vol/vol phenol/chloroform/isoamyl alcohol) and once
chloroform/isoamyl alcohol (24:1 vol/vol). Following precipitation with ethanol, the RNA
was dissolved in 20 µl PBS. Subsequently, equal amounts of sense and anti-sense RNA were
mixed, heated to 95°C for 5 minutes, and double stranded RNAs were allowed to form by
slow cooling of the reaction for several hours.
2.2.1.2 Electroporation of long ds RNA
Electroporation of double stranded RNA was performed as earlier described by Perrin and
Stoeckli (Perrin and Stoeckli, 2000). In summary, 0.1-0.5µl phosphate-buffered saline (PBS),
containing either a mixture of, PlexinD1 ds RNA (200-500 ng/µl) and YFP plasmid (under
the control of β-actin promoter), or plasmids encoding YFP alone were injected into the
central canal of the chicken spinal cord. Before and after electroporation a few drops of sterile
PBS were added to cool the embryo. Platinum electrodes (BTX, Genotronics) of 4 mm length
with a distance of 4 mm between anode and cathode were used. The number of pulses and the
voltage were chosen dependent on the embryo age. After two to three days, embryos were
sacrificed and processed for immunofluorescence analysis.
2.2.1.3 Whole mount immunostaining
Embryos for whole mount immunostaining were taken between stages 24-26. Internal organs
and non-required body parts were removed and the clean embryo was fixed in 4% PFA-PBS
for 1 hour. After several washes with PBS, the embryos were permeabilized at ambient
temperature for 1 hour with constant but gentle agitation, using PBS containing 1 % Triton.
2.2 Material and methods
41
Following two washes with PBS, 20 mM lysine in 0.1 M sodium phosphate (pH 7.3 – 7.4)
was added for 1 hour and was subsequently replaced by PBS containing 10% goat serum
(blocking buffer). After two hours, the first antibody mixture containing a mouse monoclonal
anti-neurofilament antibody at a dilution of 1:1500 (RMO-270; Zymed laboratories) and a
rabbit anti-YFP polyclonal antibody (ab 290; Abcam) in a 1:500 dilution was added and
incubated at 4°C for at least 48 hours. Several PBS washes were performed before incubating
the embryos in blocking buffer at 4°C overnight. Embryos were than incubated with the
secondary antibodies, a Cy3-conjugated goat anti-mouse IgG (Jackson ImmunoResearch) and
an Alexa-conjugated goat anti-rabbit IgG. Following an overnight incubation at 4°C, embryos
were washed several times with PBS (5 times for at least 1 hour), and subsequently
dehydrated in methanol and immersed in BBDA. Specimen, were analyzed using indirect
Immunofluorescence on a Zeiss axiovert.
2.2.1.4 Immunofluorescence staining
Embryos were dissected at the designed stages in PBS and fixed for 1 hour in 4% PFA-PBS.
After 30 minutes washing in PBS, tissues were embedded in OCT and quickly frozen in
isopentane on dry ice. Cryostat sections, cut at 25-µm thickness were washed for 3 minutes in
PBS at 37°C and subsequently transferred for 30 minutes to 20 mM Lysine. Sections were
than washed with PBS and incubated for 1 hour at room temperature in PBS containing 10%
goat serum (blocking buffer). The primary antibody mixture (a rabbit anti-YFP polyclonal
antibody (ab 290; Abcam) 1:500 mixed with either a mouse monoclonal anti-neurofilament
antibody 1:1500 (RMO-270; Zymed laboratories), or a mouse monoclonal anti-HNK-1
antibody (sigma) 1:500 or mouse monoclonal anti-1E8 were added overnight at 4°C. Slides
were than washed in three changes of PBS and incubated for 1 hour in blocking buffer. 1:250
diluted fluorochrome-conjugates secondary antibodies (Cy3-conjugated goat anti-mouse IgG
(Jackson ImmunoResearch laboratories) and Alexa 488-conjugated goat anti-rabbit IgG
(molecular probes) were added subsequently for 2 hour at room temperature. Finally, the
sections were washed 5 times in PBS and mounted in Fluoromount before being examined by
indirect Immunofluorescence on a Zeiss axiovert.
2.2.2 In situ hybridization
2.2.2.1 cRNA probe labeling
The Chicken DNA plasmid derived from the two EST clone (ChEST697F11 and
ChEST867E24), found by data base search, were linearized using restriction endonucleases
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(NotI or EcoRI; Roche). The linearized PD1 plasmid was DIG labeled by incubating 2 µg of
DNA with 2 µl digoxigenin (DIG) labeling mix (Roche), 2 µl of T3 or T7 RNA polymerase
(Roche), 2 µl of 10 X transcription buffer (Roche), and H2O added to a final volume of 20 µl,
at 37°C for 2 hours. After incubation, 2 units of Rnase free DNaseI (Roche, 10U/µl) was
added to the mix, and incubated at 37°C for 30 min, after which 2 µl of 0.2 M EDTA, pH 8.0,
was added to stop the nuclease treatment. The cRNA probe was ethanol-precipitated and
dissolved in 50 µl of Rnase-free H2O.
2.2.2.2 RNA in Situ Hybridization
Chick embryos at the designated stages were dissected in PBS and fixed in 4% PFA-PBS for
1 hour. After 30 minutes washing in PBS, tissues were embedded in OCT and quickly frozen
in isopentane on dry ice. Sections of 25-µm thick were cut, collected on Super frost Plus
(Fisher Scientific) microscope slides, dried at room temperature and stored at -20°C until use.
Alternatively, embryonic tissue from different stages were collected after dissection in PBS
and immediately embedded in OCT prior to quick freezing in isopentane on dry ice.
Tissue sections were post-fixed half an hour in 4% PFA-DEPC PBS before a single 5 minutes
wash in PBS followed by a 5 minutes wash in DEPC water were carried out. Sections were
subsequently acetylated for 10 minutes, washed for 5 minutes once in PBS and once in 2X
SSC-DEPC and subsequently incubated with the prehybridization buffer containing 40%
formamid , 5X SSC-DEPC, 5X denhardts’ solution, 0.5 mg/ml yeast tRNA, 0.5 mg/ml
salmon sperm DNA at 54°C for 3 hours.
cRNA probes, diluted in prehybridization buffer at final concentration of 3 ng/µl, were added
to the slides and incubated over night at 54°C. The next morning, slides were washed as
following: 5 minutes in 5X SSC, 5 minutes in 2X SSC, 5 minutes in 0.2X SSC, 20 minutes in
0.2X SSC containing 40% formamid at 54°C followed by one wash for 5 minutes in 2X SSC
at room temperature. All the following steps were carried out at ambient temperature. Slides
were afterwards washed twice for 10 minutes in detection buffer (0.1 M Tris-base, 15 mM
NaCl, pH 7.5) before incubation in blocking buffer (3% milk in detection buffer) to block non
specific binding. The anti-DIG phosphatase-conjugated antibody diluted in blocking buffer at
1:2000, was added to slides and left for 1 hour at room temperature prior to washing twice in
detection buffer and one wash in alkaline phosphatase buffer (0.1 M Tris-Base, pH 9.5, 0.1 M
NaCl and 50 mM MgCl2) for 5 minutes each wash. The bound probe was detected by adding
NBT/BCIP substrate (Roche). For each ml of alkaline phosphatase buffer, 4.5 µl NBT and
2.3 Results
43
3.5µl BCIP were added and the mixture was added to tissue sections and developed in the
dark over night at 4°C. Images were recorded on a Zeiss Axioskop.
2.3 Results
2.3.1 Expression of PlexinD1 in chicken spinal cord during motor axon
sorting in the limb plexus
In embryonic mice and zebra fish, PlexinD1 is expressed in endothelial cells where it is
implicated in intersomitic vessels formation and sprouting (Torres-Vazquez et al., 2004),
(Gitler et al., 2004). However, earlier reports have shown that PlexinD1 is also expressed in
the developing nervous system (van der Zwaag et al., 2002), suggesting that it might have
additional functions than directing vascularization. Therefore, we analyzed PlexinD1
expression in the chicken embryo between stage 17 (E3) and stage 37 (E12). Similarly, to
mice and fish, PlexinD1 mRNA is present in endothelial cells. First, transcript can be detected
around stage 17 (Fig. 1A, arrow) in cells of the developing perineural vascular plexus that
sprouts and extends around the ventral half of the neural tube. By stage 20 PlexinD1
expression, has spread to the perineural vascular plexus surrounding the spinal cord (Fig. 1B)
and DRGs (Fig. 1C, arrows). PD1 expression in this region persists until stage 36 (data not
shown). Around stage 24, PD1 transcripts are detected additionally in endothelial cells of
many developing and sprouting vessels throughout the entire embryo (Fig. 1E and 1F,
arrowheads) and specifically in endothelial cells sprouting from the perineural vascular plexus
to vascularize the ventral part of the spinal cord (Fig. 1E and 1F, arrows).
Interestingly, between stage 23 and 26, PlexinD1 expression can also be detected in the spinal
cord, displaying a very specific and restricted spatio-temporal pattern (Fig. 1C, 1D, and 1E).
Expression of PlexinD1 in the spinal cord is limited to neurons of the lateral motor column
(LMC) that send axons towards different areas of the developing limb bud. The onset and
offset of PD1 expression in motor neurons correlates very well with the period of motor axon
sorting in the limb plexus, suggesting that PlexinD1 might be important for nerve segregation.
Interestingly PlexinD1 is not expressed equally throughout the entire rostrocaudal axis of the
spinal cord, suggesting that PlexinD1 might be involved only in motor axons sorting at the
hindlimb plexus.
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Fig. 1: Chicken PlexinD1 mRNA is expressed in spinal motor neurons and endothelial cells. In situ
hybridizations of transverse section through the developing chicken neural tube (nt) between stage 17 and stage
25 using a Dig labeled PlexinD1 anti-sense probe are shown. At stage 17 (A; arrows) PlexinD1 mRNA is
expressed in endothelial cells of the perineural vascular plexus, surrounding the spinal cord and the developing
DRGs (B and C; arrows). At stage 24 and stage 25 (E and F), PlexinD1 transcripts are also seen in endothelial
cells inside the spinal cord and in sprouting vessels in throughout the entire embryo. Between stage 23 and stage
25 (D, E and F) the time motor axon sort in the limb plexus, PlexinD1 is transiently expressed in motor neurons
and this expression persist until stage 26 (data not shown). Abbreviations: nt, neural tube; mc, motor column;
drg, dorsal root ganglia. Scale bars are equivalent to 200 µm.
2.3.2 PD1 knock down specifically in chicken embryo spinal cord leads to
motor axon pathfinding errors at the hindlimb level
To test the functional role of PlexinD1 in motor axon pathfinding, we performed RNA knock
down experiments in the chicken embryonic spinal cord using in ovo RNAi. Injection and
subsequent electroporation of PD1 long double stranded RNA into the central canal of stage
14 to 18 chicken embryos resulted in specific targeting of cells in one half of the spinal cord,
allowing the comparison of lumbosacral nerve patterns at the right side and the left side of the
same embryo (Fig. 2). While in control-injected embryos the crural nerve displays a highly
reproducible branching pattern, PlexinD1 knock down embryos reveal several axon
pathfinding defects in several subdivisions of the crural nerve. At stage 25 (Fig. 2C, arrow)
axons of the dorsal crural trunk begin to arrange and organize in a specific and highly
structured manner. The nerve starts to defasciculate and forms small branches and each newly
formed branch elongates and defasciculates later to give rise to the highly organized structures
we observe at stage 26 (Fig. 2D, arrow). At stage 25 this highly structured arrangement of
fibers appears strikingly disturbed in one side of PD1 injected embryos (Fig. 2C, arrowhead)
2.3 Results
45
where bundles of axons emerge highly defasciculated and disordered compared the control
side of the embryo. At stage 26, we can clearly observe that different bundles of the main
nerve fail to correctly defasciculate and branch, in the PD1 dsRNA electroporated side (Fig.
2D, arrowhead). Additionally, in embryos lacking PlexinD1 in the spinal cord, the ventral
branch that emerges from the crural nerve and forms the branch innervating the anterior
ventral thigh, fails to enter the limb bud in many embryos. Motor axons of the ventral crural
nerve trunk at the injected side seem to stall completely in the plexus region or only partially
leave the plexus to enter the limb. These defects are also observed when we compare the right
limb innervations in PD1 ds RNA injected embryos to control injected embryos (Fig. 2
control/dorsal crural trunk and control/ventral crural trunk, arrows; compared to arrowheads
in PlexinD1 knock down embryos).
Fig. 2: PD1 loss of function in chicken spinal motor neurons, results in aberrant axonal outgrowth of crural nerve
branches. Whole mount anti-neurofilament immunostaining of chicken embryos co-injected with long ds RNA
and YFP at lumbosacral levels, are shown. Within the same embryo, PlexinD1 is knocked down at the right half
of the spinal cord (arrowheads) whereas the left side is unaffected serving as a control (arrows). The injection
side is confirmed by immunostaining using an antibody against YFP (not shown). At stage 25 (A) and stage 26
(B) the ventral branch of the crural trunk at the right half of the embryo (arrowhead) fails to grow into the right
limb in contrary to the left side (arrow) where the same nerve navigate through the limb and form branches (A,
B). In addition, these embryos exhibit defects at one branch of the dorsal crural nerve trunk at stage 25 (C,
arrowhead) and stage 26 (D, arrowhead); the nerve fail to segregate, elongate and branch correctly (arrowheads
in C and D) compared to the control sides (arrows in C and D). These defects are also visible when we compare
the ventral crural nerve trunk at the left side of PlexinD1 knock down embryo (arrowheads in A and B) to
control injected embryos (arrow in control/ventral crural trunk). The same abnormalities are observed comparing
the dorsal crural nerve trunk of PlexinD1 knock down embryos (arrowheads in C and D) and control injected
embryos (arrow in control/dorsal crural trunk). Scale bars are equivalent to 200 µm.
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These data suggest that the lack of PlexinD1 expression in motor neurons is responsible for
the complete or partial retention of ventral crural trunk axons at the plexus level and for the
erroneous segregation of dorsal crural trunk axons in the limb bud.
2.3.3 PD1 knock down animals show fusion of the dorsal root entry zones
The electroporation of PlexinD1 long double stranded RNA co-injected with an YFP
expression vector into the central canal of embryos around stage 14 (E2) results in addition to
single side targeting of cells within the spinal cord also in targeting of neural crest cells
destined to populate both sides of the developing neural tube. However, YFP expression in
migrating crest cells appears to be much stronger on the right side of the injected embryo,
suggesting that most crest cells are staying on the ipsilateral side.
Fig 3: DREZs in PlexinD1 knock down animals loose their stereotypic segmental pattern. Whole mount antineurofilament immunostainings of dorsal roots at the lumbosacral levels of chicken embryos co-injected with
long ds RNA and YFP plasmid are shown. At stage 24 (B, arrowheads), stage 25 (D, arrowheads) and stage 26
(F, arrowheads) PlexinD1 knock down animals exhibit imperfections in the organization of the dorsal roots. The
normal evenly spacing between adjacent dorsal root entry zones in control animals at stage 24 (A), stage 25 (C)
and stage 26 (E) is disturbed in treated animals. Scale bars are equivalent to 200 µm.
Unexpectedly, PlexinD1 knock down embryos exhibit defects at the dorsal root entry zones.
The segmental patterning of the dorsal roots and the equal spacing between neighboring
dorsal root entry zones, usually observed in control animals between stages 24 and 26 (Fig.
3A, 3C and 3E), is no longer visible in injected embryos. Several dorsal roots appear
completely or partially fused (Fig. 3B, arrowhead), (Fig. 3D, arrowhead) and (Fig. 3E,
2.3 Results
47
arrowhead). Interestingly, the aberrant DREZ formation in PD1 knock down embryos was
often detected on both side of the spinal cord, suggesting the involvement of migratory cells,
traveling bilaterally through the embryo. This phenotype strongly suggests a functional role
for PlexinD1 not only in motor axon guidance but also in a subpopulation of crest cells
derivatives, possibly Schwann cells, and/or boundary cap cells.
2.3.4 PD1 knock down animals exhibit defects at the motor exit points of the
ventral roots
Encouraged by the circumstantial evidence that PlexinD1 might be expressed by a
subpopulation of migratory crest cells, we started to analyze the integrity of the ventral roots
at the motor neuron exit points. Immunostaining on cross sections of chicken spinal cord at
hindlimb levels, using different markers such as anti-HNK-1, anti-neurofilament and anti-1E8
an antibody that stain Schwann cells and boundary cap cells (Fig. 4), demonstrates clear
abnormalities at the ventral root level in injected embryos lacking PD1.
When compared to control embryos, the ventral roots in PlexinD1 knock down animals
appear wider and less compact as seen by immunostaining using the HNK-1 antibody (Fig.
4A arrow, 4B arrowhead). In knock down animals at the motor exit point we can observe a
split appearance of the ventral roots that elongate away from the motor exit points in
apparently two separate bundles (Fig. 4B, arrowhead). This is in clear contrast to control
embryos where the ventral roots grow and extend as a single branch (Fig. 4A, arrow).
Additionally, some fiber bundles seem to navigate apart from the main motor ventral root or
exit away from the ventral exit point when visualized by anti-neurofilament immunostaining
(Fig. 4C arrow, 4D arrowheads). In contrast to the straight route adopted by motor fibers in
control animals, in injected embryos several axon bundles clearly follow an aberrant path
after exiting the spinal cord (Fig. 4D, arrowheads). Interestingly, the aberrant axonal
trajectory is apparent on both sides of the spinal cord at the ventral root level prior to the
fusion between the motor ventral root and the sensory root. Additionally, some motor fibers in
PlexinD1 knock down animals exit the spinal cord at erroneous locations away from the usual
normal motor exit point. Furthermore, in injected embryos aggregated boundary cap cells
stained by the anti-1E8 antibody seem to have an altered morphology where the ventral root
appears split and often defasciculated (Fig. 4F, arrowheads). Interestingly these defects are
also obvious on both sides (right and left) of the spinal cord again suggesting that an aberrant
positioning of neural crest derivatives is responsible of the observed sorting errors at the
ventral roots, rather than a knock down of PlexinD1 in motor neurons.
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Fig. 4: PlexinD1 knock down in chicken spinal cord resulting in abnormalities at the motor exit points. A
comparison of ventral motor root axons between PlexinD1 knock down embryos and control injected embryos is
shown by immunostaining performed on cross sections of chicken spinal cord at stage 26 using anti-HNK-1 (A,
B), anti-neurofilament (C, D) and anti-1E8 antibodies (E, F). Crest cells expressing HNK-1 at stage 26, appear
less compact at the ventral motor bundle in injected animals (arrowhead in B) when compared to control animals
(arrow in A). Similarly, motor axons of PlexinD1 knock down animals exhibit less compact bundles after exiting
the spinal cord (D, arrowheads) in comparison to controls (C, arrow). Additionally, boundary cap cells stained
by the 1E8 antibody appear disorganized in animals lacking PlexinD1 and axon bundles seem defasciculated (F,
arrowheads) on both sides of the spinal cord. Scale bars are equivalent to 200 µm.
2.4 Discussion
49
2.4 Discussion
2.4.1 Chicken spinal motor neurons express PD1 mRNA
Several studies demonstrate contradictory results concerning the expression of PD1. While
studies investigating the role of PD1 in the vascular system, report the exclusive expression of
PD1 in endothelial cells (Torres-Vazquez et al., 2004; Gu et al., 2004; Gitler et al., 2004),
another laboratory describes PD1 expression, in addition to endothelial cells, in several
structures of the mouse central nervous system (van der Zwaag et al., 2002). Neither group
reported expression of PD1 in developing spinal motor neurons of embryonic mice. However,
our data clearly demonstrate the expression of PlexinD1 mRNA in chicken spinal motor
neurons during a time window that corresponds to hindlimb innervation. This raises the
question whether the previously reported expression patterns in mice are incomplete or
whether motor neuron specific expression is avian specific. The fact that PD1 can indeed be
expressed by neurons in the developing mouse or rat brain(Gesemann et al., 2001; van der
Zwaag et al., 2002), suggest that a functional role for PD1 in nervous system development is
not restricted to the chicken embryo. However, only a careful analysis of PlexinD1 expression
in the spinal cord of other species will answer the question whether motor axon guidance in
other species is at least in part PlexinD1 dependent.
2.4.2 PD1 involvement in the navigation of the crural nerve toward the limb
Motor axons grow in a direct manner and commit few mistakes during their pathfinding
journey. Once they exit the spinal cord, motor axons reach the base of the limb where they
sort in the limb plexus. Axon arrangement is highly specific and determines subsequently the
diverse trajectory choices that nerves innervating distinct ventral or dorsal muscles will take.
Already at the plexus region, various axon bundles innervating the same muscle assemble and
occupy particular spatial locations (ventral, dorsal medial or lateral) within the proximal nerve
trunks (Lance-Jones and Landmesser, 1981; Tosney and Landmesser, 1985c). These
reorganization events are decisive not only for ensuing dorsal or ventral trajectory choices
distinct nerves make but also for recognition of appropriate choice points where a muscle
nerve branches away from the main nerve trunk before continuing the journey to its final
target location. Selective adhesion between axons belonging to a single motor neurons pool
may contribute to this sorting process. However, axons also group into specific spatial
locations within the nerve trunks, therefore probably responding to extrinsic guidance cues
located at the base of the limb (Tosney and Landmesser, 1984).
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Our in situ data demonstrate that PD1 mRNA is expressed by all motor neurons of the LMC.
Although we require whole mount in situ hybridization experiments to confirm the precise
spinal levels of PD1 expression, we noticed on cross sections that PlexinD1 mRNA is
restricted to the most rostral lumbar and the most caudal thoracic levels while being
completely absent from all cervical and sacral motor neurons. Interestingly, PlexinD1 knock
down in chicken embryos specifically affect the normal patterning of those branches of the
crural nerve which emerge from motor neurons of the LMC at lumbar spinal levels (L1-L3)
(Landmesser, 1978a). In PlexinD1 knock down embryos, a part of the axons in the dorsal
crural nerve trunk motor axons cross the limb base to enter the limb mass, but assemble in a
disordered manner and without respecting the highly organized normal structure, whereas
axons in the ventral nerve trunk fail completely or partially to navigate correctly within the
limb bud.
Thus, the differential expression of PD1 mRNA at different spinal levels and its restricted
temporal expression between stage 23 and stage 26 in combination with the motor axon
defects noticed in PlexinD1 knock down embryos at specifically two subdivisions of the
crural nerve strongly suggest the involvement of PlexinD1 in motor axon guidance of the
dorsal and ventral crural trunk.
Our in situ data only give information about the presence of PlexinD1 mRNA, and we can
only speculate about the subcellular localization of PlexinD1 protein. As observed for other
axon guidance receptors, it seems likely that PD1 is present on advancing growth cones and
/or elongating axons. The absence of this protein in the dorsal and ventral crural nerve trunk
of knock down embryos may reduce the capability of normally PlexinD1 positive-growth
cones to react to specific cues in the surrounding tissues and subsequently results in
inaccuracy of the axonal outgrowth. Alternatively, PD1 protein could be responsible for axon
sorting in the limb plexus. At stage 23, motor axons of the LMCL and LMCM defasciculate at
the plexus level and rearrange spatially into the proximal nerve tracts before extending further
dorsally or ventrally in the limb muscle mass. The lack of PlexinD1 protein could prevent
correct defasciculation between lateral and medial lateral motor column axons, leading to
aberrant arrangements of nerve bundles at the plexus level that can subsequently position
axons at irregular locations within the limb bud. Thus, the defects observed in the course of
the dorsal and the ventral crural trunk of PlexinD1 knock down animals might be a
consequence of abnormal axon sorting at the plexus. The growth cones of abnormally
positioned axons might not be able to interpret or react to environmental cues present in their
unusual location and commit various pathfinding errors. Our in situ data combined with the
2.4 Discussion
51
aberrant projections observed in the dorsal and ventral proximal crural trunks of PlexinD1
deficient embryos support a potential role for PlexinD1 in axon guidance or alternatively in
axon sorting (involving fasciculation and defasciculation events) of the crural trunk nerves at
the limb plexus and /or proximal nerve trunks.
2.4.3 PD1 knock down affects the dorsal roots entry zones and the motor
exit points
In addition to the aberrant outgrowth of the crural trunk, PD1 knock down in chicken spinal
cord shows unexpected defects at the dorsal root entry zones (DREZs) and the motor exit
points. Surprisingly, the abolished segmental pattern noticed at the dorsal root entry zones and
the abnormalities observed at the motor ventral roots are perceived on both sides of the spinal
cord. PlexinD1 mRNA expression was never detected in dorsal root sensory neurons in
chicken embryos, eliminating a possible direct involvement of PlexinD1 in axon guidance of
sensory neurons to the dorsal root entry zones. Moreover, defects observed at the motor exit
points are unlikely due to PlexinD1 knock down in motor neurons, since changes in axon
guidance are observed on both sides of the injected embryo, while electroporation targets only
motor neurons located at one side of the spinal cord. Conversely, a Schwann cell marker
(1E8) and a neural crest marker (HNK-1) demonstrate a disorganization of boundary cap cells
in the region of the motor exit points and a defasciculation of ventral roots.
Based on these intriguing data, we can propose two models to explain the defects observed in
PlexinD1 knock down animals. The first hypothesis, suggests that the observed alterations in
spinal root formation in PlexinD1 knock down animals are based on migration defects of a
subpopulation of neural crest cell derivatives, notably Schwann cell precursors and/or
boundary cap cell precursors. These crest derivatives might migrate erroneously and would
fail to reach their appropriate final locations. The abnormal positioning of Schwann cells
and/or the boundary cap cells in PlexinD1 deficient embryos might be responsible for the
fusions observed at the dorsal root entry zones as well as the defects seen at the motor exit
points. Certainly, additional experiments are required to show an expression of PlexinD1 in
crest cell derivatives such as double in situ hybridization for PlexinD1 and a neural crest cell
marker (sox 10, slug, HNK, etc).
Nevertheless, to date all studies (in mice and zebra fish) related to PD1 expression reported its
presence exclusively in endothelial cells and excluded a role for PlexinD1 in neural crest cell
migration (Gitler et al., 2004; Torres-Vazquez et al., 2004). While, Gilter and colleges
claimed to have eliminate a possible involvement of cardiac neural crest cells for the defects
52
Part II: Paper 1
observed in PD1 mutant animals on E10.5 frontal sections. Our data clearly suggest a
potential involvement of crest cells in axon guidance. This discrepancy between our data and
earlier published reports can be due to species differences as well as to divergence in the
analyzed spinal levels. Our study mainly focuses on lumbosacral level of the spinal cord,
whereas the other study describes a more rostral region at the pharyngeal arch level in the
mouse embryo. In addition, while we analyzed axonal outgrowth at the DREZ using whole
mount immunostaining, the study by Gilter and al exclusively relies on the use of tissue
sections, which make the defects at the DREZ quite difficult to assess.
Alternatively, another hypothesis to explain our present findings derives from the potential
interaction between growing axons and blood vessels. PlexinD1 has been shown to play a
crucial role in intersomitic vessels sprouting and formation in vivo. Our neural defects can
therefore be seen as secondary effects resulting from PlexinD1 knock down in endothelial
cells that might support or guide growing axons. This would be consistent with a recent and
very elegant study in xenopus where Levine and colleges demonstrated that intersomitic
arteries form in tight relation to spinal nerves during development (Levine et al., 2003). In our
case, the electroporation of PlexinD1 long double stranded RNA in chicken spinal cord at
stage 14 might as well knock down PlexinD1 in the intersomitic vessels precursors leading to
defects in the patterning of these vessels as earlier reported in the zebra fish and mice (Gitler
et al., 2004; Torres-Vazquez et al., 2004). Subsequently, outgrowth of axons in the area of the
dorsal root is perturbed by abnormally formed intersomitic vessels leading to aberrant
arrangement of the DREZs. According to this model, defects in vascular patterning would
lead to axon misguidance.
Although this model appears interesting and eliminates contradiction between our current
study and earlier published work, it certainly requires many additional experiments to
determine its validity. To date we do not have evidence about the possibility of targeting nonneural cells with our electroporation technique. In addition, we did not examine the eventual
disturbances of the vascular system especially in intersomitic vessels in PD1 knock down
animals. Importantly, we also could not observe any YFP positive cells in the peripheryoutside the spinal cord and the DRG- in double injected (ds RNA and YFP vector) or control
animals (injected only with YFP vectors). This argues for the specificity in targeting only
neural cells. However, it is essential to mention that DNA vector and long double stranded
RNA do neither have the same size and nor the same charges. Thus, the possibility of a wider
diffusion of long double stranded RNA comparing to the YFP-vector cannot be absolutely
excluded.
2.4 Discussion
53
Despite the enormous advances achieved in the field of axon guidance, our knowledge about
pathfinding choices taken by different neurons in vivo remains very fragmentary.
2.4.4 PD1 potential binding partner(s) in chicken embryos during
development
PD1 knock down in spinal cord leads to aberrant motor axon pathfinding as well as defects in
the dorsal root entry zones and ventral motor roots formation.
Our current study does not give indications about the potential binding partner(s) of PlexinD1
that might be responsible for mediating its biological functions in vivo. Earlier reports,
demonstrating a biological function for PD1 in vivo (Gitler et al., 2004; Gu et al., 2004),
suggest the involvement of different signaling pathways using different ligands or ligand
complexes.
In the first report, PlexinD1 was found to co-immunoprecipitates with NP-1 and NP-2
proteins that bind with high affinity to Sema3A and Sema3C respectively (Gitler et al., 2004).
In contrast, Gu and colleagues confirm that PlexinD1 and Sema3E display complementary
expression patterns in mice as well as similar defects in PlexinD1 and Sema3E knock out
animals. Interestingly, Sema3E-PD1 signaling did not require neuropilins (Gu et al., 2004),
presumed obligate co-receptor for Semaphorins Class III members (Tamagnone et al., 1999;
Raper, 2000).
In a parallel work, we investigated the spatio-temporal expression pattern of chicken
semaphorins and plexins during development (chapter 3 and 4). Our study shows, that
between stage 22 and stage 26, the period chicken spinal motor neurons express PlexinD1
messages, neuropilin-1, neuropilin-2, Sema3A, Sema3C, and Sema3E messages are also
detected in chicken motor neurons (chapter 3 and chapter 4). It is tempting to speculate that
these semaphorin individually or combined in a complex, might act also in chicken as
potential ligands to PlexinD1. In this case, secreted semaphorin in combination with
neuropilins or independently of it, can bind PlexinD1 driving defasciculation events at the
crural nerve trunk.
Surprisingly, at stage 22 and stage 25 PlexinC1 (PC1) is detected in endothelial cells outside
the spinal cord, however to a lesser extend in comparison to PlexinD1 expression (chapter 4),
suggesting a possible binding between these plexins to mediate signaling in endothelial cells.
However, our current data are not sufficient to conclude whether the same binding partners
described earlier in mouse, activate also PlexinD1 signaling in chicken embryo.
Supplementary experiments, using biochemical assays are essential to elucidate the potential
54
Part II: Paper 1
binding of chicken PlexinD1 to one or more of the different semaphorin class III and
neuropilins members. Additionally, functional knock down of sema3A, 3C and 3E and
neuropilins genes in the chicken spinal cord, can give insights about the possible interactions
in vivo between PlexinD1 and these different semaphorins.
Part III:
Paper 2
Developmental regulation of semaphorin expression in the chicken spinal
cord and peripheral nervous system suggests different functions and
interactions
3 Dummyheading
55
Developmental regulation of semaphorins in spinal cord and peripheral
nervous system suggests different semaphorin functions and interactions in
the chicken embryo
1
Joelle Gemayel, 2Rejina Sadhu, 2Olivier Mauti, 2Esther T. Stoeckli, and 1Matthias Gesemann
1
Brain Research Institute, University of Zurich, and department of biology, ETH Zurich, and
2
Institute of Zoology, University of Zurich,
Winterthurerstrasse 190, 8057 Zurich, Switzerland
correspondence to:
[email protected]
phone:
+41 44 635 3283
fax:
+41 44 635 3303
[email protected]
phone:
+41 44 635 4840
fax:
+41 44 635 6879
key words:
56
chicken embryo, semaphorin, motoneurons, interneurons, DRG
Abstract
Semaphorins are axon guidance molecules with mainly repulsive activities that exist in a
variety of different subclasses. While functions and expression patterns of members in the
secreted subclass III have been described in detail, far less is known about expression and
function of molecules in the subclasses IV, V, VI and VII and few data are available about the
expression of these molecules in the developing chicken embryo. We have now performed a
combined EST and genomic database search to identify chicken semaphorins and to analyze
their expression patterns in the developing chicken spinal cord. In contrast to mouse, human
and rat, the chicken genome contains a reduced number of semaphorins, as no homologues
have been found for the class IV semaphorins Sema4A, Sema4C and Sema4F. In addition, no
counterpart could be identified for Sema6C. Interestingly, the chicken genome contains a
class IV semaphorin that cannot be assigned to class IV semaphorins found in other species.
Our databank search also revealed a yet non-described novel semaphorin Sema3G that can be
found in chicken as well as other vertebrate species. Expression analysis of the different
chicken semaphorins in the spinal cord revealed that while semaphorins of the subclass III
seemed to have conserved expression patterns across species, expression domains for class V
and VI semaphorins seemed to be altered compared to mice or rats and that class IV
semaphorins are not expressed in the developing chicken spinal cord. Taken together, our
results demonstrate that the chicken genome shows a slightly altered composition of
semaphorins compared to other species, resulting in a changed expression pattern for several
members that might most likely reflect changes in their binding partners and functions.
3.1 Introduction
Directed cell migration, accurate axonal navigation, and correct target recognition are crucial
for the proper functioning of the nervous system. Migration and axon outgrowth occur along
specific and highly stereotypic pathways that are determined by interactions of specific cell
surface receptors with environmental cues that confer positional and guidance information to
the migrating neuron or advancing axon (Tessier-Lavigne and Goodman, 1996). Therefore,
defining mechanisms as well as identifying molecules by which migrating neurons and
growing axons select their pathways, maintain a directed growth along them, and later
recognize and innervate the appropriate target areas, are fundamental issues in neurobiology.
Several different families of axon guidance molecules have been identified within the last
decade(Dickson, 2002; Huot, 2004; Zou, 2004). One of the largest families of axon guidance
57
58
Part III: Paper 2
cues, being mainly involved in redirecting axonal outgrowth by selectively collapsing the
axon growth cone, is the semaphorin family (Raper, 2000). Today, more than 20 different
semaphorins are known which based on their structural similarities can be grouped into seven
different subclasses. While Semaphorins falling into the subclasses I and II are exclusively
found in insects, semaphorins belonging to the higher subclasses are found in many different
vertebrate species ranging from fish to avian to men (Fiore and Puschel, 2003; Kolodkin et
al., 1992; Matthes et al., 1995). Initial functional experiments suggested that Semaphorins are
mainly repulsive axon guidance molecules. However, it has become increasingly clear that
semaphorins serve a far wider range of functions such as chemoattraction, directing cell
migration, being involved in immune responses and in the formation of the vascular system
(Fujisawa, 2004; Pasterkamp et al., 2003; Tamagnone and Comoglio, 2004). Nevertheless, the
most prominent function of semaphorin family members remains their role in conferring
guidance information for growing axons.
In the spinal cord, a great number of precise connections must be established to enable proper
integration of sensory information from and accurate delivery of motor impulses to the
periphery. While nociceptive and thermoceptive sensory neurons establish connections with
the dorsal-most layer of the spinal cord, proprioceptive neurons form synapses with
interneurons and motoneurons. Survival and axonal outgrowth of different functional
subtypes of sensory neurons critically depend on the presence of different neurotrophins, with
nociceptive and thermoceptive neurons requiring NGF/TrkA signaling (Patel et al., 2000),
whereas proprioceptive sensory neurons clearly depend on NT-3/TrkC activation (Patel et al.,
2003). The majority of sensory input is relayed to the brain via spinal interneurons. While
many different subtypes of interneurons exist in the spinal cord, the best-studied neuronal
subtype are commissural interneurons, which are located in the dorsal part of the spinal cord
(Yaginuma et al., 1994). This particular subtype of interneurons is characterized by a highly
complex but stereotypic axonal trajectory. Initial commissural axon outgrowth occurs towards
the ventral midline of the spinal cord, resulting in axons crossing a particular structure called
the floor plate. Once on the contralateral side commissural axons perform a sharp orthogonal
turn projecting rostrally towards the brain finally making connections with higher brain
regions (Kaprielian et al., 2001). However, sensory neurons not only form functional
connections with spinal interneurons but in special cases also with motoneurons located in the
ventral horn of the spinal cord (Davis et al., 1989). As for sensory neurons and spinal
interneurons, motor neurons can acquire a variety of different identities that are reflected in
the choice of specific axon pathways and synaptic targets (Schneider and Granato, 2003).
3.1 Introduction
59
While initially axons from neurons of different pools project along a common pathway, their
arrival in the peripheral mesoderm is accompanied by multiple sorting events into different
motor axon trajectories. The trajectory of all the above mentioned neuronal subtypes seems to
be determined by a variety of different guidance cues and cell adhesion molecules, including
slits (Wang et al., 1999), different members of the IgCam super family (Perrin et al., 2001)
and semaphorins (Masuda et al., 2003).
Sema3A was isolated more than a decade ago based on its collapse inducing activity for
axons of DRG neurons (Kolodkin et al., 1993; Luo et al., 1993). Subsequently, other
experiments demonstrated that Sema3A selectively repels NGF responsive axons that
normally terminate in the dorsal half of the spinal cord (Messersmith et al., 1995).
Additionally, a zebrafish homologue of Sema3A has been shown to inhibit growing motor
axons (Roos et al., 1999). However, the analyses of Sema3A function in vivo using gene
disruption in mice gave rather surprising results in respect to nervous system development.
Mutant mice exhibit no or only modest defects in sensory, commissural or motor axon
projections, suggesting that either the functional significance for Sema3A in axon guidance
has been over estimated or that due to functional redundancy, other class III semaphorins can
compensate for the loss of Sema3A (Taniguchi et al., 1997). So far, only few or no additional
semaphorins have been shown to be involved in guiding sensory, commissural or motor axons
(Kikuchi et al., 1999). Zebrafish Sema4E has been demonstrated to act as a repulsive
boundary, guiding brachiomotor axons to their targets, and Sema6A acts as a repellent on
sensory NT-3 and NGF sensitive DRG neurons, whereas Sema4D seems to stimulate axonal
outgrowth of sensory neurons in a autocrine way, (Masuda et al., 2004; Xiao et al., 2003;
Zhou et al., 1997). While the number of semaphorins with demonstrated functional roles in
sensory, motor and interneuron guidance is limited, several other family members are also
expressed in and around the spinal cord about which no functional data is available (Bagnard
et al., 1998; Giger et al., 2000; Takahashi et al., 1998). Chicken Sema3D as well as mouse
Sema3C are highly expressed in developing motor neurons whereas high levels of Sema3B
message are found in sensory neurons, (Bagnard et al., 1998; Takahashi et al., 1998). Several
of these transcripts are also expressed in adjacent mesodermal tissue, suggesting that they
might also influence outgrowth of sensory and motor axons.
Over the last couple of years, several Semaphorin-binding proteins have been identified.
While neuropilins serve mainly as co-receptors transmitting guidance responses of Class III
semaphorins in a PlexinA/neuropilin/semaphorin complex, different members of the plexin
family have been shown to convey semaphorin signaling in a neuropilin independent manner
60
Part III: Paper 2
(Tamagnone et al., 1999). As for semaphorins, neuropilins and plexins are widely expressed
throughout the entire nervous system, suggesting functional roles in a variety of guidance
events. Despite various studies describing a wide range of interactions between semaphorins,
neuropilins, and plexins, our understanding of how these interactions are implicated in axon
targeting is still fragmentary. Moreover, for some semaphorins and plexins, binding partners
have not yet been described, suggesting that a certain number of interactions still await its
discovery. Finally, it seems clear that several plexins have to form interactions with multiple
semaphorins, as already described for different class A plexins and PlexinB1, and that some
semaphorins might have additional binding partners besides the described ones.
As a first step towards a more complete understanding of semaphorin functions, we
performed a detailed spatiotemporal expression analysis in developing spinal cord and
surrounding tissue. Surprisingly, the chicken genome only encodes seventeen different
semaphorins, whereas mammalian genomes code for twenty different semaphorin genes.
Interestingly the number of class III semaphorin genes is identical in avian and mammals,
however conservation within the subclass IV is very low, and the avian genome lacks a
Sema6C homologue. The absence of several semaphorins in chicken is reflected by an altered
expression pattern of chicken semaphorins in the different subclasses, suggesting that these
semaphorins have slightly altered functions and binding properties in higher vertebrates.
3.2 Material and Methods
3.2.1 Assembly of chicken semaphorin cDNAs
cDNA sequences for chicken semaphorins were assembled using the combined information
from the chicken EST (http://www.chick.umist.ac.uk) and the chicken genomic database
(http://www.ensembl.org/Multi/blastview?species=Gallus_gallus).
17
different
genomic
regions coding for semaphorins were identified using the tblastx alignment algorithms on
available vertebrate semaphorins or existing chicken semaphorins. The corresponding
genomic fragments were downloaded and analyzed using the genescan gene prediction
program (http://genes.mit.edu/GENSCAN.html). The obtained putative cDNA and protein
sequences were compared to the corresponding mammalian homologues and genescan
prediction errors were corrected by manual inspection of the intron/exon boundaries in false
predicted regions. Gaps in the assembled sequences due to inaccurate or incomplete genome
sequencing were wherever possible filled by corresponding EST sequences. A total of 162
chicken ESTs encoding different semaphorins were identified. Among these 55 covered only
3.2 Material and Methods
61
parts of the 3’UTR sequence whereas the rest contained part of the coding sequence.
Sequence alignment of genomic and EST sequences was done using the SeqMan software
(Lasergene, DNASTAR, Madison WI). 3’UTR sequences were added to the coding sequence
based on overlapping EST sequences that were supplemented with genomic sequences. UTR
sequences were terminated at the first polyadenylation AATAAA/ATTAAA sequence that
followed verified chicken UTR EST sequences. Using this combined approach 12 complete
and 5 partial cDNA sequences for semaphorins could be assembled.
3.2.2 Phylogenetic tree analysis
The domain structure of representative members of the chicken and mouse semaphorin super
family were obtained using the smart program (http://smart.embl-heidelberg.de). Individual
domains were extracted from the sequence using domain boundaries as predicted. Conserved
domains from the different semaphorin subfamilies were aligned using the CLUSTAL W
alignment algorithm (Higgens and Sharp, 1989; Thompson et al. 1994) provides by the
MagAlign software (Lasergene, DNASTAR, Madison WI). Obvious mistakes in domain
boundary prediction were manually adjusted. For better representation, alignment files were
exported into TREEVIEW software, enabling the graphical representation of the unrooted tree
(Page, 1996).
3.2.3 Cloning of semaphorin cDNA fragments
cDNA fragments of semaphorin genes with no hits in the EST database were cloned using
RT-PCR. A putative cDNA assembled based on genomic information was used as a template
to design sense and antisense primers. Total RNA was prepared using spinal cord and DRG
tissue isolated from stage 30 chicken embryos. Random and oligodT primed first strand
cDNAs were generated using the SuperscriptII reverse transcription kit (Invirtogen, Carlsbad
CA) according to the manufacturers’ instruction. A 669 bp long cDNA fragment for Sema3G
was amplified by PCR using the following sense 5’CTGTCAAGCGCCAAAAGC and
antisense 5’GGCACTGCTCCTCCACC primers. In analogy to this, a 656 bp long fragment
for Sema6B was amplified using the following two primers 5’ ATCCAGCGCATCCTCAAG
(sense) and 5’ CCCATGTCGTTCTTGCAC (antisense). Obtained PCR fragments were
cloned into the Topo TA cloning vector (Invirtogen, Carlsbad CA) and subsequently
sequenced to verify the identity of the insert.
62
Part III: Paper 2
3.2.4 In situ hybridization
3.2.4.1 cRNA probe labeling
Chicken DNA plasmids derived from the EST clones ChEST399E25, ChEST771A21,
ChEST578C18,
ChEST375M12,
ChEST7787C4,
ChEST659J12,
ChEST986M15,
ChEST585F20, ChEST225N10 and ChEST1004D4 corresponding to Sema3A, Sema3B,
Sema3C, Sema3D, Sema3E, Sema3F, Sema5A, Sema5B, Sema6D and Sema7A, found by
database searches, were linearized using the restriction endonucleases NotI or EcoRI.
Linearized plasmids were DIG labeled by incubating 2 µg of each DNA with 2 µl digoxigenin
(DIG) labeling mix (Roche), 2 µl of T3 or T7 RNA polymerase (Roche), 2 µl of 10 X
transcription buffer (Roche), and H2O added to a final volume of 20 µl for each reaction, at
37°C for 2 hours. After incubation, 2 units of Rnase free DNaseI (Roche, 10U/µl) was added
to the mix, and incubated at 37°C for 30 min, after which 2 µl of 0.2 M EDTA, pH 8.0, was
added to stop the nuclease treatment. The cRNA probe was ethanol-precipitated and dissolved
in 50 µl of Rnase-free H2O.
3.2.4.2 RNA in Situ Hybridization
Chick embryos at the designated stages were dissected in PBS and fixed in 4% PFA-PBS for
1 hour. After 30 minutes washing in PBS, tissues were embedded in OCT and quickly frozen
in isopentane on dry ice. Sections of 25 µm thick were cut, collected on Super frost Plus
(Fisher Scientific) microscope slides, dried at room temperature and stored at -20°C until use.
Alternatively, embryonic tissue from different stages were collected after dissection in PBS
and immediately embedded in OCT prior to quick freezing in isopentane on dry ice.
Tissue sections were post-fixed half an hour in 4% PFA-DEPC PBS before a single 5 minutes
wash in PBS followed by a 5 minutes wash in DEPC water were carried out. Sections were
subsequently acetylated for 10 minutes, washed for 5 minutes once in PBS and once in 2X
SSC-DEPC and subsequently incubated with the prehybridization buffer containing 40%
formamid , 5X SSC- DEPC, 5X denhardts’ solution, 0.5 mg/ml yeast tRNA, 0.5 mg/ml
salmon sperm DNA at 54°C for 3 hours.
cRNA probes, diluted in prehybridization buffer at final concentration of 3 ng/µl, were added
to the slides and incubated over night at 54°C. The next morning, slides were washed as
following: 5 minutes in 5X SSC, 5 minutes in 2X SSC, 5 minutes in 0.2X SSC, 20 minutes in
0.2X SSC containing 40% formamid at 54°C followed by one wash for 5 minutes in 2X SSC
at room temperature. All the following steps were carried out at ambient temperature. Slides
were afterwards washed twice for 10 minutes in detection buffer (0.1 M Tris-base, 15 mM
3.3 Results
63
NaCl, pH 7.5) before incubation in blocking buffer (3% milk in detection buffer) to block non
specific binding. The anti-DIG phosphatase-conjugated antibody diluted in blocking buffer at
1:2000, was added to slides and left for 1 hour at room temperature prior to washing twice in
detection buffer and one wash in alkaline phosphatase buffer (0.1 M Tris-Base, pH 9.5, 0.1 M
NaCl and 50 mM MgCl2) for 5 minutes each wash. The bound probe was detected by adding
NBT/BCIP substrate (Roche). For each ml of alkaline phosphatase buffer, 4.5 µl NBT and
3.5µl BCIP were added and the mixture was added to tissue sections and developed in the
dark over night at 4°C. Images were recorded on a Zeiss Axioskop.
3.3 Results
3.3.1 Identification of Sema3G, a novel member of the class III semaphorins
Due to its easy accessibility and its high regeneration capability, the chicken embryo has been
the model organism of choice to study a variety of developmental phenomena, ranging from
neuronal differentiation to axon guidance and synaptogenesis. However, the lack of
appropriate tools to knockout or knockdown gene function has strongly limited the use of
chicken embryos in recent years. Lately, there has been a revival in the use of the chicken
systems, mainly due to three different reasons. While the establishment of an extraordinarily
large EST database and the near completion of the chicken genome sequencing have provided
the molecular bases for obtaining a great variety of chicken gene sequences, the combination
of in ovo electroporation with functional RNAi knockdown have overcome the limitations for
chicken research (Bourikas and Stoeckli, 2003; Krull, 2004; Pekarik et al., 2003). Despite
these advances, the use of chicken embryos as a model system for the study of molecular
mechanisms of axon guidance is still limited compared to other experimental systems such as
mouse and rat. This is represented also by the fact that even if the first vertebrate semaphorin
to be identified has been chicken Sema3A (Luo et al., 1993), information about the existence
and the expression of different chicken semaphorins remains fragmentary. Until now, only
five different avian semaphorins have been characterized (Luo et al., 1995), preventing a
careful analysis of chicken semaphorin expression and function. To narrow this gap we have
now performed extensive databank searches using the combined information from the EST
and the genomic database to predict the number of chicken semaphorin genes. Searches for
different class III semaphorins resulted in the identification of all previously described
mammalian semaphorins (Fig. 1). Both databases provided a very complete set of information
as seen by the fact that neither the genomic nor the EST database failed to identify the
64
Part III: Paper 2
different class III semaphorins (Table 1). Indeed, the genomic database reveals the presence
of an additional class III semaphorin member that we called Sema3G. Subsequent databank
searches in mouse, rat, and human confirmed the existence of this novel semaphorin member.
Phylogenetic tree analysis of the conserved semaphorin domain revealed that the novel
Sema3G is most closely related to Sema3E, however the conservation of the sema domain of
chicken Sema3G and mouse Sema3G is the lowest between all class III semaphorins (Fig. 1).
Sema3A
Sema3B
Sema3C
Sema3D
Sema3E
Sema3F
Sema3G
ESTs
8
2
10
14
1
6
0
Chicken Ch
1
Un
1
1
1
Un
12
5a2
9f2
5a2
5a2
5a2
9f1
14a3
Mouse Ch
Sema4A
Sema4B
Sema4C
Sema4D
Sema4F
Sema4G
Sema4X
ESTs
n.d
0
n.d
12
n.d
5
7
Chicken Ch
n.d
28
n.d
Un
n.d
6
10
Mouse Ch
3f1
7d1
1b
13a5
6c3
19c3
n.d
Sema5A
Sema5B
Sema6A
Sema6B
Sema6C
Sema6D
Sema7A
ESTs
22
15
14
0
n.d
29
10
Chicken Ch
2
7
z
28
n.d
10
10
15b2
16b1
18c
17c
3f2.1
2e5
9b
Mouse Ch
Table 1: Chromosomal localization and EST representation of different semaphorin gene products. The number
of identified chicken ESTs for each gene is indicated and chromosomal location of the chicken and its
corresponding mouse gene are given. Not detected genes are abbreviated by n.d, whereas identified
chromosomal sequences on not yet localized chromosomes are indicated by the letters Un.
3.3.2 The chicken genome has a reduced number of semaphorin genes
In contrast to the class III semaphorins, the number of class IV semaphorins in chicken was
significantly reduced (Fig. 1; Table 1). While in rat, mouse and human we could identify six
different class IV semaphorins, only four such family members could be identified in chicken.
Moreover, these semaphorins were neither as highly conserved as the class III semaphorins,
3.3 Results
65
nor could we align the identified members to a mammalian counterpart. We were unable to
obtain a corresponding sequence for chicken Sema4A, Sema4C, and Sema4F; however, we
identified an additional class IV semaphorin that shows the highest homology to mouse
Sema4D and Sema4A, without being a clear homologue.
Fig. 1: Phylogenetic tree of the members of the semaphorin super family. Alignment is done on the bases of the
semaphorin domain of each family member using the CLUSTAL W program. The scale bar represents the
substitution rate of 10 amino acids per 100 amino acid residues. Please note that only mouse (m) and chicken (c)
semaphorin sequences are depicted, but that alignment of rat or human sequences gave similar results (data not
shown).
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Part III: Paper 2
In contrast to semaphorin members found in the subclass IV, no changes in gene number have
been observed for members of the subclasses V and VII. Nevertheless, as already observed
for class IV semaphorins the number of class VI semaphorin genes in chicken is lower
compared to mammals. While Sema6A, 6B, and 6D are present and highly conserved in the
chicken genome, Sema6C gene is absent.
3.3.3 Class
III
semaphorins
are
highly
expressed
in
developing
motoneurons
Seven secreted Class III semaphorins have been identified in the chicken genome. At stage
18, the earliest stage we have analyzed, no expression of sema3A, 3D, 3E, and 3F is
detectable in the embryonic chicken spinal cord (Fig. 2). Sema3B and 3C are however
expressed in motoneurons at this early developmental stage (Fig. 2, 3B and 3F white arrows).
By stage 22, motoneurons also express sema3A (Fig. 3, 3A white arrow) and 3D (Fig. 3, 3D
black arrow) and at stage 30 Sema3E transcripts are up regulated in these cells (Fig. 4, 3E
white arrow). However, Sema3F transcripts are never expressed in motoneurons at all
analyzed stages (Fig. 2, 3, 4, 5, and 6). In addition to motoneurons, Sema3A is expressed also
in cells of the ventral ventricular zone (Fig. 3, 3A white arrowhead) and transiently in
interneurons of the intermediate area of the spinal cord (Fig. 3, 3A star) and (Fig. 4, 3A). In
addition, by stage 30 ventral interneurons also express Sema3D (Fig. 5, 3D white arrowhead)
and Sema3F transcripts (Fig. 4, 3F white arrowhead) are expressed in a small population of
cells adjacent to the roof plate at stage 26. Interestingly, at this stage another semaphorin
Class III member, Sema3D is substantially expressed in sensory neurons of the dorsal root
ganglia (Fig. 4, 3D white arrowhead) and this expression remains high until stage 35 (Fig. 6,
3D white arrowhead).
3.3.4 Floor plate cells expresses high levels of semaphorin V transcripts
Until now, little is known about functions, and expression patterns of chicken Class V
semaphorins. We have looked at the distribution of Sema5A and Sema5B during development
of the spinal cord. At stage 18, shortly before commissural neurons located in the dorsolateral
spinal cord start to extend their axons toward the ventral midline, both semaphorin Class V
members are expressed in the floor plate (Fig. 2, 5A and 5B black arrows). Strong expression
of both Sema5A and Sema5B in the floor plate persists through stage 24 (data not shown).
While the level of Sema5A transcripts starts to decrease after stage 24, Sema5B RNA is still
expressed very strongly in the floor plate at stages 26 (Fig. 4, 5B) and are still detectable at
stage 30 (Fig. 5, 5B). In contrast to Sema5A, whose expression is restricted to the floor plate,
3.3 Results
67
Sema5B is also transiently expressed in neuroblasts of the dorsal spinal cord (Fig. 2, 5B white
star) and commissural interneurons. By stage 26, when commissural axons have crossed the
floor plate and are just about to exit and to turn into the longitudinal axis Sema5B mRNA is
strongly down regulated in commissural neurons (Fig. 4, 5B).
Motoneurons become post mitotic between stages 18 and 26 (Krull and Koblar, 2000). While
Sema5A expression can be detected as early as stage 20 in motor neurons (data not shown)
and remains later in specific subpopulations (Fig. 4, 5A), Sema5B mRNA is never detected in
this neuronal population.
Fig. 2: Expression patterns of chicken semaphorins in spinal cord and peripheral nervous system at stage 18.
Cross sections of lumbosacral chicken spinal cord were incubated with Dig labeled RNA antisense probes.
Semaphorin probes used are indicated. While Sema3B (3B, white arrow) and Sema3C (3C, white arrow)
transcripts are detected in the motor neurons, the floor plate expresses Sema5A (5A, black arrow), Sema5B (5B,
black arrow) and Sema7A (7A, black arrow). Additionally, Sema5B is expressed in dorsal interneurons (white
star). Scale bar correspond to 200 µm.
68
Part III: Paper 2
Fig. 3: Expression patterns of chicken semaphorin members in spinal cord and peripheral nervous system at
stage 22. Cross sections of lumbosacral chicken spinal cord were incubated with Dig labeled RNA antisense
probes. Semaphorin probes used are indicated. By stage 22 Sema3A expression appears in motoneurons (3A,
white arrow) and in cells of the ventral ventricular zones (3A, white arrowhead) as well as in interneurons of the
intermediate ventricular zone (3A, star). While Sema3D (3D, black arrow) and Sema3C (3C, white arrow)
expression persists in motoneurons, Sema5A mRNA is now up regulated in motoneurons (5A, white arrow),
Sema5B is strongly expressed in ventricular zone (5B, white arrow) and Sema7A is very weakly expressed in
post mitotic motoneurons (7A, white arrow). Scale bar correspond to 200 µm.
Moreover, sensory neurons in the dorsal root ganglia appear to express Sema5A transcript
during the time when they form collaterals to target their specific layers in the gray matter
(Fig. 5, 5A black arrow). While this expression is subpopulation specific, Sema5B transcript
was never detected in any sensory neurons (Fig. 2, 3, 4, 5, 6).
3.3.5 Class VI semaphorins are highly expressed in boundary cap cells
In contrast to mammals, which contain four different class VI semaphorin genes, the chicken
genome encodes only three of them (Fig. 1). While Sema6A has a striking expression pattern,
3.3 Results
69
being detectable only in boundary cap cells (Fig. 4, 6A black arrows and Fig. 5, 6A), Sema6B
is diffusely expressed in motoneurons starting at stage 22 (data not shown). Additionally
Sema6D transcripts are also detected in motoneurons as early as stage 22 (data not shown).
Fig. 4: Expression patterns of chicken semaphorins in spinal cord and peripheral nervous system at stage 26.
Cross sections of lumbosacral chicken spinal cord were incubated with Dig labeled RNA antisense probes.
Semaphorin probes used are indicated. Expression levels of Sema3A (3A white arrow) and Sema3D (3D, white
arrow) transcripts remain unaltered between stage 22 and stage 26 in motoneurons. In contrast Sema3C (3C,
white arrow) expression has been strongly down regulated in motoneurons and Sema3D RNA levels are up
regulated in DRGs (3D, white arrowhead). Moreover, Sema3F transcripts start to be expressed in a population of
cells adjacent to the roof plate, most likely representing associational interneurons (3F, white arrowheads).
Sema5A and Sema5B expression persist in the floor plate (5A and 5B arrowheads), in addition Sema5A is
expressed in motoneurons (5A, white arrowhead) and Sema5B remains highly expressed in the ventricular zone
(5B white arrowhead). In addition strong expression of Sema6A and Sema6D is now seen in boundary cap cells
(6A, black arrowheads). High levels of Sema6D are also detected throughout the gray matter of the spinal cord
(6D). Finally, Sema7A expression is strongly up regulated in motoneurons and in the dorsal lateral spinal cord
(7A, white arrowhead). Scale bar correspond to 200 µm.
70
Part III: Paper 2
Sema6D expression broadens as transcripts can be detected during later stages of
development (Fig. 4, 6D) in the dorsal spinal cord, the dorsal root ganglia, and interestingly
also in the boundary cap cells (Fig. 4, 6D, black arrow) and (Fig. 5, 6D). Subsequently,
Sema6D expression extends throughout the gray matter, with stronger signals in lamina І of
the dorsal horn (Fig. 6, 6D white arrowhead).
Fig. 5: Expression patterns of chicken semaphorin members in spinal cord and peripheral nervous system at
stage 30. Cross sections of lumbosacral chicken spinal cord were incubated with Dig labeled RNA antisense
probes. Semaphorin probes used are indicated. Little changes in expression patterns of different semaphorin
transcripts are detected between stage 26 and stage 30. The most obvious change can be seen for Sema3D (3D,
white arrowhead) and Sema5A (5A, white arrow) where transcripts are also detected now in ventrally located
interneurons. Note that Sema3E transcripts are transiently located in a subpopulation of motoneurons (3E, white
arrow) and Sema5A is expressed in a subpopulation of sensory neurons (5A, black arrow). Scale bar correspond
to 200 µm.
3.3 Results
71
Fig. 6: Expression patterns of chicken semaphorins in spinal cord and peripheral nervous system at stage 35.
Cross sections of lumbosacral chicken spinal cord were incubated with Dig labeled RNA antisense probes.
Semaphorin probes used are indicated. By this stage of spinal cord development most semaphorins transcript are
strongly down regulated, with the exception of Sema3D whose expression is maintained in motoneurons as well
as sensory neurons (3D, white arrowhead) and Sema6D, which is now detected in lamina I of the dorsal horn
(3D, white arrowhead). Interestingly, a subpopulation of motoneurons still express Sema3C and low levels of
Sema3B are still detectable in the gray matter and in DRGs. Scale bar correspond to 200 µm.
3.3.6 Semaphorin7A is expressed in endothelial cells and motoneurons
Only one member, Sema7A is found in semaphorin Class VII. Sema7A expression in the
nervous system is highly regulated and starts at stage 18 (Fig. 2, 7A black arrow) with a very
weak signal in the floor plate that is maintained through stage 26 (Fig. 4, 7A), but is no longer
visible at stage 30. Post mitotic motoneurons in the ventral spinal cord express Sema7A
weakly at stage 22 (Fig. 3, 7A white arrow), later the expression levels are up regulated very
72
Part III: Paper 2
strongly (Fig. 4, 7A) and persists until stage 30 (Fig. 5, 7A). Sensory neurons in DRGs
express Sema7A only during the time when they form collaterals (data not shown).
Interestingly, a transient expression of Sema7A is also seen in the dorsolateral spinal cord at
stage 26 (Fig. 4, 7A white arrowhead) corresponding to the location of associational neurons.
3.4 Discussion
3.4.1 The chicken genome has fewer semaphorin genes that the mammalian
genome
While the mammalian genome encodes for at least 20 different semaphorin proteins, our
databank searches only identified 17 chicken counterparts. One of the major questions in this
respect is whether we were simply unable to identify corresponding chicken fragments or
whether the chicken genome indeed contains a reduced number of semaphorin genes. The fact
that we did not obtain EST sequences for several chicken semaphorins while identifying a
genomic sequence for it, demonstrates that the EST database coverage is not yet complete,
making it possible that the failure of identifying chicken counterparts for Sema4A, Sema4C,
Sema4F and Sema6C could be due to the incompleteness of the available databases. While
this is certainly a possibility, it seems rather unlikely, since we were able to identify a
genomic sequence for each of the chicken semaphorins that showed a successful EST hit.
Several other circumstances suggest that we indeed identified all chicken semaphorins.
Despite the fact that the haploid chicken genome contains 38 autosomes plus the Z and W sex
chromosomes, compared to only 19 autosomes and the X and Y sex chromosomes in mouse,
the chicken genome is about 50% (1.2 x 109 bp) reduced in size when compared to the mouse
genome (2.56 x 109 bp). This reduction in size is mainly based on the fact that 30 of the 38
chicken autosomes, called microchromosomes, are extremely small, ranging only between 5
and 20 Mb (Venkatesh et al., 2000; Vinogradov, 1999). While a reduction in size is not per se
an indication for a reduced number of genes, as demonstrated for many teleost, two
observations we made suggest that in chicken the reduced size goes hand in hand with a
partial loss of genes. First, the overall gene length between corresponding chicken and mouse
genes is not significantly reduced (Vinogradov, 1999), second when we analyzed the
conservation of chicken genes belonging to either the cadherin or the IgCam super family, we
observed a similar reduction in genes (data not shown). Attempts to clone chicken
homologues that could not be identified by database screening using degenerate primers in
RT-PCR reactions never did yield to a corresponding gene product. Finally, yet importantly,
3.4 Discussion
73
two of the four mouse genes absent in the chicken genome are located in the same
chromosomal area, suggesting that a homologous part for this chromosome is not present in
the chicken genome. Nevertheless only the completion of the chicken genome sequencing
project as well as the extension of the number of sequenced chicken EST can finally answer
the question whether the chicken genome indeed encodes fewer class IV semaphorins than the
mammalian genome does.
3.4.2 Expression patterns of semaphorins are dynamically regulated during
spinal cord development
3.4.2.1 Motoneurons express high transcript levels of different class III
semaphorins
Sema3A was isolated more than a decade ago based on its collapse inducing activity for
sensory axons o (Kolodkin et al., 1993; Luo et al., 1993). Initial experiments demonstrated
that Sema3A selectively repels NGF responsive axons that normally terminate in the dorsal
half of the spinal cord (Messersmith et al., 1995). While many semaphorin family members
have been isolated, most studies carried out to learn more about functional roles of
semaphorins in vivo and in vitro still concentrate on secreted members belonging to the
subclass III (for reviews see , (Fiore and Puschel, 2003). Class III semaphorins mainly signal
through the formation of large signaling complexes involving neuropilins and plexinAs
(Tamagnone et al., 1999). However there is also evidence that class III semaphorins can act in
a neuropilin independent way (Gu et al., 2004).
In chicken spinal cord and adjacent sensory systems, we demonstrated the expression of
several different class III semaphorins. While almost all family members are expressed in
developing motoneurons, Sema3s are never found in the floor plate, although commissural
axons extending toward the floor plate express plexinAs. This is, however, consistent with the
absence of neuropilins in commissural axons that would be expected as co-receptors to bind
secreted semaphorins (Tamagnone et al., 1999). Interestingly, the subpopulation specific
expression of Class III semaphorins in motoneurons supports a potential implication of these
molecules in guiding motor axons. An interesting hypothesis would be that SemaIIIs are
constantly secreted by different motor nerves and that the presence of different semaphorins
will be used to drive defasciculation events of selective nerves at particular choice points,
when attractive or adhesive forces within the fascicle are weakened. Such a mechanism would
be in analogy to results seen in the Drosophila embryo, where the transmembrane Semaphorin
Іa is required for the selective defasciculation of specific motoneurons (Yu et al., 1998). Even
74
Part III: Paper 2
if class III semaphorins are supposed to be secreted they have been shown to bind strongly to
neuronal surfaces via their highly charged C-terminal end, making it possible that these
molecules are indeed highly concentrated on axon fascicles driving selective defasciculation
events at various choice points (Bagnard et al., 2000).
3.4.2.2 Expression of class V semaphorins in the floor plate, commissural
interneurons and sensory neurons suggest conserved as well as novel functions
for these molecules
We have shown that class V semaphorins are highly expressed at the ventral midline,
suggesting several potential functions for these molecules. While they can simply serve as
repulsive or attractive guidance cues, providing a signal for contralaterally and ipsilaterally
projecting neurons they may also be important as floor plate intrinsic molecules taking part in
determining floor plate morphology. Such a function would mainly be achieved when class V
semaphorins would act as cell adhesion molecules rather than as axon guidance cues.
However, such a function has never been shown so far, and no semaphorin/semaphorin
interactions have been documented. Nevertheless, the facts that binding between semaphorins
and their plexin receptors occur most likely through functional interactions between their
sema domains (Raper, 2000) and that plexins are capable of forming homophilic complexes,
make a speculation of homophilic semaphorin interactions quite plausible. However, only
functional assays using either RNA interference or knockout techniques will provide evidence
about the functional roles class V semaphorins play in floor plate cells.
Interestingly, during initial stages of commissural axon outgrowth Sema5B mRNA is also
transiently expressed in this interneuron subpopulation. However, the exact timing of Sema5B
transcript regulation in commissural neurons has not been carefully determined. It seems
feasible that Sema5B is expressed in commissural interneurons while they extend axons
toward the floor plate, but that this expression is down regulated upon contact with floor plate
cells. Analysis of Sema5B expression in stage 24 embryos (data not shown) indeed points in
this direction as significantly lower expression levels are observed in these embryos when
compared to stage 22 animals. Given the fact that growing axons seem to express Sema5B
during periods of axonal growth could suggest a novel function for this molecule potentially
serving as a receptor mediating reverse signaling. Such a function would also explain the
presence of Sema5A in a subpopulation of sensory neurons. While reverse signaling has been
already shown for many Ephrin ligands (Murai and Pasquale, 2003), this concept is rather
new for semaphorins. Nevertheless, recent reports have confirmed that semaphorins
3.4 Discussion
75
belonging to another subfamily of transmembrane semaphorins have indeed the capacity to
serve as receptors (Toyofuku et al., 2004b). However, so far no interactions for Sema5B have
been documented. Interestingly a recent report suggests Sema5A might indeed be
bifunctional, serving as attractive as well as repulsive axon guidance cue (Kantor et al., 2004).
This dual function strongly depends on interactions of heparan and chondroitin sulfate
proteoglycans with the conserved thrombospondin repeats of Sema5A (Kantor et al., 2004),
but it remains to be determined whether Sema5B is capable of similar interactions. However,
while it seems clear that either proteoglycan loaded Sema5A can serve as an attractant or
repellant in Trans, it would be interesting to see whether these complexes also function in a
Cis interaction, conferring either positive or negative guidance for commissural neurons.
3.4.2.3 Class VI semaphorins as potential gatekeeper between neurons located in
the central and peripheral nervous system
Very striking expression patterns are especially found for two class VI semaphorin members,
namely Sema6A and Sema6D. While Sema6D is expressed in a subpopulation of
motoneurons as well as in sensory neurons, exceptionally high mRNA levels for Sema6D and
Sema6A are detected in a special subpopulation of migrating neural crest cells called
boundary cap cells. Boundary cap cells settle at four different locations along the spinal cord
in the area of the dorsal root entry zones and the motor neuron exit points (Golding and
Cohen, 1997). The fact that Sema6A and Sema6D transcript up regulation occurs only after
neural crest cells have initiated their migration suggests that Sema6A and 6D are not required
during early events of boundary cap cell development. However, transcript levels for both
molecules are strongly increased by the time these cells aggregate in the area of the ventral
and dorsal roots. This expression pattern suggests several possible functions for these class VI
semaphorin family members. Thus, it seems reasonable to speculate about a possible role for
Sema6A and Sema6D in initial events of boundary cap cells aggregation, by means of
conferring either attractive or adhesive interactions. However, another appealing possibility
remains to be considered about the potential role class VI semaphorin in boundary cap cells as
“gate keepers” keeping neuronal cell bodies confined within specific locations by repulsive
interactions. The latter hypothesis is strengthened by the fact that a potential Sema6D
receptor, PlexinA1, is highly expressed in developing motor and sensory neurons (Toyofuku
et al., 2004b), (see also chapter 3). Conversely, class VI semaphorins might also act as
attractants for motor and sensory neurons, specifically guiding these axons towards the entry
zones or exit points of the spinal cord. That semaphorins can indeed act as attractant has been
76
Part III: Paper 2
shown in several recent reports (Dent et al., 2004; Wolman et al., 2004). However, only
functional assays will provide evidence about the role of class VI semaphorins in boundary
cap cells.
Part IV:
Paper 3
Expression patterns of plexins and neuropilins suggest cooperative and
separate functions in spinal cord development
4 Dummyheading
77
Expression patterns of plexins and neuropilins suggest cooperative and
separate functions in spinal cord development
1
Olivier Mauti, 2Joelle Gemayel 1Rejina Sadhu, 2Matthias Gesemann, and 1Esther T. Stoeckli
1
Brain Research Institute, University of Zurich, and department of biology, ETH Zurich, and
2
Institute of Zoology, University of Zurich,
Winterthurerstrasse 190, 8057 Zurich, Switzerland
correspondence to:
[email protected]
phone:
+41 44 635 4840
fax:
+41 44 635 6879
[email protected]
phone:
+41 44 635 3283
fax:
+41 44 635 3303
key words:
78
chicken embryo, dorsal root ganglia, motoneurons, commissural neurons
Abstract
Semaphorins and their receptors, neuropilins and plexins, play important roles in providing
positional and guidance information for growing axons. While class III semaphorins are
mainly using a plexin/neuropilin complex as signaling platform, other described interactions
suggest that most semaphorins act as guidance cues in a neuropilin independent way.
However, only few interactions between plexins and semaphorins have been described so far,
suggesting that additional cross talk exists between these molecular groups. In order to
analyze potential functions and interactions of plexins and neuropilins, we now performed a
combined EST and genomic database search to identify chicken plexins and analyze their
expression patterns in the developing chicken spinal cord. As already observed for
semaphorin genes, the chicken genome contains a reduced number of plexins when compared
to mouse, human and rat. Only seven plexins were found in chicken, whereas nine plexins
were identified in mammals. While both plexins in the single protein subfamilies C and D are
conserved between species, the A subfamily and B subfamily each lack one family member as
neither PlexinA3 nor PlexinB3 could be identified in the chicken genome. Expression
analysis of chicken plexins and neuropilins during spinal cord development revealed a
strikingly different expression for plexins and neuropilins. While PlexinAs are widely
expressed throughout the entire spinal cord, PlexinBs seemed to be most abundant in glia and
PlexinC1 seems to be expressed only during late stages of neuronal development.
Interestingly PlexinD1, which has previously been reported to be exclusively expressed in
endothelial cells of the vascular system, seems also be expressed by developing motoneurons.
In contrast to this, neuropilin expression domains are far more restricted, suggesting in
addition to cooperative also separate function for both molecular groups.
4.1 Introduction
Semaphorins constitute a large family of secreted and membrane bound molecules that can
function as repellents or attractants regulating fasciculation, axonal growth and branching as
well as terminal arborisation (Fujisawa, 2004). Two different classes of receptors, called
neuropilins and plexins, mediate these described functions. While plexins constitute a large
family of molecules that can be grouped into four major subfamilies, PlexinA, B, C and D,
neuropilins form only a small family with two known members (Tamagnone et al., 1999),
(Puschel, 2002). Neuropilins have been shown to be involved in mediating various sema III
79
80
Part IV: Paper 3
signals by forming a complex with different plexins. However, class IV and VII semaphorins
have been shown to act in a neuropilin independent manner (Tamagnone et al., 1999).
Neuropilins are transmembrane proteins acting as co-receptors conferring ligand specificity in
a larger signaling complex. Due to the lack of any obvious signaling component in the
neuropilin cytoplasmic part, activity is mediated by complex formation with either plexins,
L1, or off track tyrosine kinase (Castellani, 2002; Castellani and Rougon, 2002). While
neuropilin-1 (NP-1) is expressed in peripheral sensory neurons, autonomic neurons of the
sympathetic ganglia and a subpopulation of ventral motoneurons, neuropilin-2 is expressed
throughout the entire ventral half of the spinal cord including motoneurons and several
subpopulations of interneurons (He and Tessier-Lavigne, 1997; Giger et al., 2000). In sensory
neurons, neuropilin-2 (NP-2) seems only to be transiently expressed during initial stages of
sensory axon outgrowth, as no expression has been documented in mice older than E13 (Chen
et al., 1997; He and Tessier-Lavigne, 1997; Kolodkin et al., 1997). In accordance with the
expression of the two neuropilin RNAs, neuropilin-2 mutant mice display a strikingly reduced
size of the dorsal funiculus, whereas Neuropilin-1 deficient mice are embryonic lethal
showing abnormal defasciculation of cranial nerves and peripheral nerves as well as a looser
DRG packaging (Giger et al., 2000; Kitsukawa et al., 1997).
Among the different plexin subfamilies, members of the A group have been best characterized
in terms of expression and function. All four mammalian PlexinAs are widely but somehow
complementarily expressed in the developing spinal cord and adjacent sensory ganglia
(Murakami et al., 2001). While PlexinA3 seems strongly expressed throughout the entire
spinal cord, PlexinA2 is selectively expressed in the dorsal spinal cord. Moreover, PlexinA3
and PlexinA4 are most abundant in DRGs, whereas PlexinA1 and PlexinA2 are also
expressed in sensory neurons, but the expression is limited to a small subpopulation and
expression is absent from sympathetic ganglia (Murakami et al., 2001). Finally, PlexinA3
appears to be expressed in all peripheral ganglia, including trigeminal, vagal and otic in
addition to dorsal root and sympathetic ganglia (Cheng et al., 2001). Interestingly, PlexinA2expressing cardiac neural crest cells are patterned abnormally in Sema3C mutant mice
whereas PlexinA3 null mice display no gross defects in pathfinding of dorsal root sensory
neurons but show fasciculation defects in the ophthalmic branch of the trigeminal nerve and
aberrant development of hippocampal projections(Brown et al., 2001; Cheng et al., 2001).
In contrast to PlexinAs, PlexinBs transcripts are only expressed at low levels in different
regions of the spinal cord and peripheral nervous system. While PlexinB2 and B3 transcripts
are absent in sensory neurons, PlexinB1 mRNA can be found in both, DRG as well as
4.2 Material and methods
81
trigeminal ganglia (Worzfeld et al., 2004). In the spinal cord, PlexinB1 and B2 messages are
strongly expressed in the ependymal neuroepithelium whereas no PlexinB3 transcript is
detectable in the spinal cord before birth. Until now, also no expression in the developing
spinal cord has been reported for the unique PlexinC and PlexinD subfamily members.
Despite the fact that neuropilins and plexins are expressed in various populations of spinal
inter- and motoneurons as well as in sensory neurons of the peripheral nervous system, only
few functional interactions between these molecules have been shown to be of biological
significance (Chedotal et al., 1998; Kawasaki et al., 2002). In addition, no binding partners for
several plexins and a large number of semaphorins have been found, suggesting that a large
number of plexin/semaphorin ligand complexes are not yet identified. In order to learn more
about potential functions and interactions of neuropilins, plexins and semaphorins
participating in spinal cord and peripheral nervous system development, we have performed a
detailed spatiotemporal expression study of all semaphorins (see chapter 2), neuropilins, and
plexins present in the chicken genome. Interestingly, when compared to the mammalian
genome the chicken genome lacks two plexin family members, namely PlexinA3 and
PlexinB3 and the conservation within the PlexinB subclass is rather low. Expression of
chicken plexins and neuropilins during spinal cord development revealed that neuropilin
mRNAs are expressed only in restricted regions, only partially overlapping with the
expression of PlexinA, B, C, and D, suggesting in addition to cooperative also separate
functions for both molecular groups.
4.2 Material and methods
4.2.1 Assembly of chicken plexin cDNAs
cDNA sequences for chicken plexins were assembled using the combined information from
the chicken EST (http://www.chick.umist.ac.uk) and the chicken genomic database
(http://www.ensembl.org/Multi/blastview?species=Gallus_gallus).
7
different
genomic
regions coding for plexins were identified using the tblastx alignment algorithms on available
vertebrate plexins. The corresponding genomic fragments were downloaded and analyzed
using the genescan gene prediction program (http://genes.mit.edu/GENSCAN.html). Putative
cDNA and protein sequences were compared to the corresponding mammalian homologues
and genescan prediction errors were corrected by manual inspection of the intron/exon
boundaries in false predicted regions. Gaps in the assembled sequences due to inaccurate or
incomplete genome sequencing were wherever possible filled by corresponding EST
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Part IV: Paper 3
sequences. A total number of 85 chicken ESTs covering parts of 7 different plexins were
identified. Among these 65 contained part of the coding sequence whereas the rest covered
only parts of the 3’UTR sequence. Sequence alignment of genomic and EST sequences was
done using the SeqMan software (Lasergene, DNASTAR, Madison WI). 3’UTR sequences
were added to the coding sequence based on overlapping EST sequences that were
supplemented with genomic sequences. UTR sequences were terminated at the first
polyadenylation AATAAA/ATTAAA sequence that followed verified chicken UTR EST
sequences. Using this combined approach a total of 5 complete and 2 partial cDNA sequences
for plexins could be assembled.
4.2.2 Phylogenetic tree and domain identity analysis
The domain structure of representative members of the chicken and mouse plexin super
family was obtained using the smart program (http://smart.embl-heidelberg.de). Individual
domains were extracted from the sequence using domain boundaries as predicted. Conserved
domains from the different plexin subfamilies were aligned using the CLUSTAL W
alignment algorithm (Higgens and Sharp, 1989; Thompson et al. 1994) provides by the
MagAlign software (Lasergene, DNASTAR, Madison WI). Obvious mistakes in domain
boundary prediction were manually adjusted. For better representation, alignment files were
exported into TREEVIEW software, enabling the graphical representation of the unrooted tree
(Page, 1996). Identical and conserved amino acids within individual domains were
determined
by
pairwise
alignment
using
the
bl2seq
blast
program
(http://www.ncbi.nlm.nih.gov/-blast/bl2seq/wblast2.cgi).
4.2.3 In situ hybridization
4.2.3.1 cRNA probe labeling
The Chicken DNA plasmids derived from the EST clones (ChEST53D13, ChEST128L21,
ChEST1014M19,
ChEST890P9,
ChEST799I19,
ChEST860K1,
ChEST110K21
and
ChEST675H12) corresponding respectively to PlexinA1, PlexinA2, PlexinA4, PlexinB1,
PlexinB2, PlexinC1, Neuropilin-1 and Neuropilin-2, found by data base search, were
linearized using restriction endonucleases (NotI or EcoRI; Roche). The linearized plasmids
were DIG labeled by incubating 2 µg of each DNA with 2 µl digoxigenin (DIG) labeling mix
(Roche), 2 µl of T3 or T7 RNA polymerase (Roche), 2 µl of 10 X transcription buffer
(Roche), and H2O added to a final volume of 20 µl for each reaction, at 37°C for 2 hours.
After incubation, 2 units of Rnasefree DNaseI (Roche, 10U/µl) was added to the mix, and
4.2 Material and methods
83
incubated at 37°C for 30 min, after which 2 µl of 0.2 M EDTA, pH 8.0, was added to stop the
nuclease treatment. The cRNA probe was ethanol-precipitated and dissolved in 50 µl of
Rnase-free H2O.
4.2.3.2 RNA in Situ Hybridization
Chick embryos at the designated stages were dissected in PBS and fixed in 4% PFA-PBS for
1 hour. After 30 minutes washing in PBS, tissues were embedded in OCT and quickly frozen
in isopentane on dry ice. Sections of 25 µm thick were cut, collected on Super frost Plus
(Fisher Scientific) microscope slides, dried at room temperature and stored at -20°C until use.
Alternatively, embryonic tissue from different stages were collected after dissection in PBS
and immediately embedded in OCT prior to quick freezing in isopentane on dry ice.
Tissue sections were post-fixed half an hour in 4% PFA-DEPC PBS before a single 5 minutes
wash in PBS followed by a 5 minutes wash in DEPC water were carried out. Sections were
subsequently acetylated for 10 minutes, washed for 5 minutes once in PBS and once in 2X
SSC-DEPC and subsequently incubated with the prehybridization buffer containing 40%
formamid , 5X SSC- DEPC, 5X denhardts’ solution, 0.5 mg/ml yeast tRNA, 0.5 mg/ml
salmon sperm DNA at 54°C for 3 hours.
cRNA probes, diluted in prehybridization buffer at final concentration of 3 ng/µl were added
to the slides and incubated over night at 54°C. The next morning, slides were washed as
following: 5 minutes in 5X SSC, 5 minutes in 2X SSC, 5 minutes in 0.2X SSC, 20 minutes in
0.2X SSC containing 40% formamid at 54°C followed by one wash for 5 minutes in 2X SSC
at room temperature. All the following steps were carried out at ambient temperature. Slides
were afterwards washed twice for 10 minutes in detection buffer (0.1 M Tris-base, 15 mM
NaCl, pH 7.5) before incubation in blocking buffer (3% milk in detection buffer) to block non
specific binding. The anti-DIG phosphatase-conjugated antibody diluted in blocking buffer at
1:2000, was added to slides and left for 1 hour at room temperature prior to washing twice in
detection buffer and one wash in alkaline phosphatase buffer (0.1 M Tris-Base, pH 9.5, 0.1 M
NaCl and 50 mM MgCl2) for 5 minutes each wash. The bound probe was detected by adding
NBT/BCIP substrate (Roche). For each ml of alkaline phosphatase buffer, 4.5 µl NBT and
3.5µl BCIP were added and the mixture was added to tissue sections and developed in the
dark over night at 4°C. Images were recorded on a Zeiss Axioskop.
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4.3 Results
4.3.1 Plexin and Neuropilin genes in chicken
4.3.1.1 The avian genome lacks homologues for two mammalian plexin
counterparts
In order to identify chicken plexins and neuropilins we have performed extensive databank
searches using the combined information from the EST and the genomic database. As
previously observed for chicken semaphorin genes, database searches provide evidence that
the chicken genome encodes a reduced number of plexins compared to its mammalian
counterpart. While homologous chicken genes for the two unique family members PlexinC1
and PlexinD1 could be readily identified, neither an identical number of chicken PlexinAs nor
PlexinBs is present in the chicken genome when compared to mammalians (Table 2). While
the PlexinA subfamily contains no matching chicken sequence for PlexinA3, no counterpart
for PlexinB3 could be extracted from chicken databases (Fig. 1 and Table 1). Interestingly,
while conservation between different chicken PlexinAs was in the range of 70 to 90%,
depending on the plexin parts used for alignment, conservation between chicken PlexinBs
was only around 50% (Table 2). This value is not much higher than the values obtained when
PlexinBs were compared to plexins in other subclasses, suggesting that even when put into
the same subclass PlexinB1 and PlexinB2 might actually be members of different subclasses.
PA1
PA2
PA3
PA4
PB1
PB2
PB3
PC1
PD1
ESTs
16
23
n.d
4
4
23
n.d
6
9
Chicken Ch
12
26
n.d
1
12
Un
n.d
1
12
Mouse Ch
6d1
1h6
Xa7.1
6a3.3
9f2
15e3
Xa7.1
10c2
6e3
Table 1: Chromosomal localization and EST representation of different plexin gene products. Numbers of
identified chicken ESTs for each gene are indicated and chromosomal location of the chicken and its
corresponding mouse gene are given. Not detected genes are abbreviated by n.d, whereas identified
chromosomal sequences on not yet localized chromosomes are indicated by the letters Un.
4.3 Results
85
Sema Domain
cPlexinA1
cPlexinA2 57 / 71
cPlexinA4 56 / 70
59 / 76
cPlexinB1 28 / 47
28 / 48
27 / 47
cPlexinB2 28 / 47
30 / 49
30 / 48
36 / 55
cPlexinC1 13 / 23
12 / 22
13 / 22
12 / 20
12 / 21
cPlexinD1 24 / 38
23 / 40
23 / 40
27 / 43
24 / 40
15 / 28
cPlexinA1 cPlexinA2 cPlexinA4 cPlexinB1 cPlexinB2 cPlexinC1 cPlexinD1
PSI Domain
cPlexinA1
cPlexinA2 64 / 73
cPlexinA4 74 / 87
62 / 77
cPlexinB1 50 / 61
44 / 55
50 / 58
cPlexinB2 44 / 65
39 / 56
39 / 60
48 / 66
cPlexinC1 42 / 70
42 / 76
46 / 65
48 / 65
38 / 57
cPlexinD1 37 / 53
41 / 61
37 / 55
40 / 51
39 / 53
39 / 56
cPlexinA1 cPlexinA2 cPlexinA4 cPlexinB1 cPlexinB2 cPlexinC1 cPlexinD1
Table 2: Conservation of different regions between plexin super family members. Pairwise alignment of different
individual domains within the plexin proteins indicate that the intracellular SCOP domain is highly conserved
between the different plexin family members, whereas the two analyzed extracellular domains, sema and PSI,
show far less conservation. Interestingly the conservation between individual domains of chicken PlexinB1 and
PlexinB2 is only slightly higher than the homology between class A plexins and class B plexins, suggesting that
chicken PlexinB genes are evolutionary quite distinct. The left numbers indicate the identical amino acids,
whereas the right numbers indicate the conserved amino acids.
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SCOP Domain
cPlexinA1
cPlexinA2 88 / 95
cPlexinA4 82 / 94
86 / 95
cPlexinB1 56 / 72
56 / 73
53 / 73
cPlexinB2 56 / 74
56 / 75
56 / 76
66 / 79
cPlexinC1 49 / 69
48 / 68
49 / 70
45 / 65
48 / 66
cPlexinD1 56 / 75
56 / 74
58 / 75
55 / 74
55 / 74
57 / 77
cPlexinA1 cPlexinA2 cPlexinA4 cPlexinB1 cPlexinB2 cPlexinC1 cPlexinD1
Table 2 (continued): Conservation of different regions between plexin super family members. Pairwise
alignment of different individual domains within the plexin proteins indicate that the intracellular SCOP domain
is highly conserved between the different plexin family members, whereas the two analyzed extracellular
domains, sema and PSI, show far less conservation. Interestingly the conservation between individual domains
of chicken PlexinB1 and PlexinB2 is only slightly higher than the homology between class A plexins and class B
plexins, suggesting that chicken PlexinB genes are evolutionary quite distinct. The left numbers indicate the
identical amino acids, whereas the right numbers indicate the conserved amino acids.
4.3.1.2 Plexins and Neuropilins expression at early developmental stages of the
chicken spinal cord
During early developmental stages, (stage 18 to stage 22), axons of motoneurons start to leave
the spinal cord to form the ventral roots and neural crest cells migrate ventrolaterally
aggregating into bilaterally paired DRGs (Krull, 2001; Krull and Koblar, 2000).
At stage 18, only PlexinA1 (Fig. 2, A1 black arrow) and PlexinA2 (Fig. 2, A2 black arrow)
mRNAs are detected in motoneurons. While the expression patterns of these two plexins are
originally similar in motoneurons by stage 22, PlexinA1 and PlexinA2 transcript levels are
differently regulated. PlexinA1 transcript levels remain high in motoneurons whereas
PlexinA2 mRNA expression decreases and subsists in only a subgroup of motoneurons (Fig.
3, A1, and A2). Interestingly, PlexinA4 mRNA expression in motoneurons is also detected at
stage 22 (Fig. 3, A4 white arrow) whereas no other plexins subfamily members are expressed
during early motor axonal outgrowth (Fig. 2 and Fig. 3). Neuropilin-1 (NP-1) and Neuropilin2 (NP-2) are also expressed during early motor neuron development (Fig. 3, NP-1 and NP-2
white arrows), however only neuropilin-1 can be detected at stage 18 (Fig. 2, NP-1 black
arrow).
4.3 Results
87
Fig. 1: Phylogenetic tree of the members of the plexin super family. Alignments are done on the bases of the
plexin sema domain (A) or the first PSI domain of each family member using the CLUSTAL W program. The
scale bar represents the substitution rate of 10 amino acids per 100 amino acid residues. Please note that only
mouse (m) and chicken (c) semaphorin sequences are depicted, but that alignment of rat or human sequences
gave similar results (data not shown).
However, PlexinA1 and PlexinA2 expression is not restricted to the ventral spinal cord but
extends to the dorsal spinal cord, where PlexinA1 and PlexinA2 mRNA are also detected in
commissural interneuron precursors (Fig. 3, A1, and A2, white arrowheads). Commissural
neurons are generated at stage 19 (Stoeckli and Landmesser, 1995) and express all PlexinA
members by stage 22 (Fig. 3, A1, A2 and A3, white arrowheads), the time when the majority
of commissural axons reach the floor plate. Surprisingly, around stage 22 all PlexinAs,
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PlexinB1 (PB1) and PlexinC1 (PC1) are also expressed in the floor plate (Fig. 3, A1, A2, A4,
B1 and C1, black arrows), a structure that represents an intermediate target for commissural
interneurons. Interestingly, PlexinA1 (PA1) is only expressed in lateral but not medial floor
plate cells, and PlexinB1 is expressed in the entire ventricular zone (Fig. 3, B1 and Fig. 6,
B1), PlexinB2 is strongly expressed in cells surrounding the spinal cord and this expression is
rapidly down regulated later. In addition, PlexinA1, PlexinA4 (PA4), and both neuropilins
transcripts are detected in dorsal root ganglia at stage 22.
Fig. 2: Plexin and Neuropilin expression patterns in chicken spinal cord and peripheral nervous system at stage
18. Cross sections of chicken lumbosacral spinal cord were incubated with Dig labeled antisense probes.
Different probes used are indicated. At this early stage of development, only PA1, PA2, and NP-1 transcripts are
detected in the ventral spinal cord (A1, A2 and NP-1 black arrow). Scale bars correspond to 200 µm.
4.3.1.3 Plexins and Neuropilins expression during hindlimb innervation
After stage 23, motor axons reach the plexus region where they sort out extensively according
to their muscle targets. While the first decision is primarily a choice to grow either dorsally or
ventrally, pathways that are more refined are chosen by stages 25/26, when individual muscle
nerves begin to form (Landmesser, 2001). At that time, motoneurons can based on their
rostrocaudal position within the spinal cord axis and their transcriptional profile be separated
into different subpopulations (Jessell, 2000).
4.3 Results
89
Fig. 3: Plexin and Neuropilin expression patterns in chicken spinal cord and peripheral nervous system at stage
22. Cross sections of chicken lumbosacral spinal cord were incubated with Dig labeled antisense probes.
Different probes used are indicated. At stage 22 all PlexinAs PB1 and PC1 are expressed in the floor plate (A1,
A2, A4, B1 and C1 black arrows). While, PlexinAs are expressed in dorsal interneurons (A1, A2 and A4 white
arrowheads), PA4 and NP-2 messages are also detected in motoneurons (A4, NP-2 white arrows). PA1, PA2,
and NP-1 (NP-1 white arrow) messages persist in motoneurons at this stage and all PlexinAs and both
neuropilins are expressed in DRGs. PlexinBs transcripts are also up regulated at this time. PB1 (B1) is highly
expressed in the ventricular zone and PB2 messages (B2) are detected transiently in cells surrounding the spinal
cord. Scale bars correspond to 200 µm.
At the lumbosacral level, the region we studied, the motor column can mainly be divided into
the lateral motor column (LMC) and medial motor column (MMC), with the later motor
column being further subdivided into lateral LMCL and medial LMCM parts depending on the
motoneurons position within the LMC (Jessell, 2000; Landmesser, 1978b).
At stage 25, we can clearly detect neuropilin-2 and PlexinA1 messages in a subpopulation of
motoneurons belonging to the LMCL (Fig. 4, NP-2, and A1, black arrows), whereas
neuropilin-1, PlexinA4 and PlexinD1 (PD1) display a broader expression corresponding to all
LMCs (Fig. 4, NP-1, A4 and D1, white arrows). Additionally, while PlexinB1 transcripts are
not detected in motoneurons at this stage, PlexinB2 and PlexinC1 display weak expression in
some motoneurons that do not appear to be clearly segregated into pools (Fig. 4, B2, and C1,
black arrowheads).
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Plexin and neuropilin expression in different motoneuron subpopulations is also observed at
stage 30, although the distribution of individual plexins has changed (Fig. 5). PlexinA4
mRNA is no longer expressed homogenously in motoneurons but rather in a gradient with
increasing transcription levels from the medial to the lateral motoneuron pool (Fig. 5, A4
white arrow). The expression of PlexinD1 is lost in motoneurons and remains only in
interneurons along the ventricular zone that overlaps with NP-1 expression (Fig. 5, D1, and
NP-1, black arrows). Strikingly NP-1 and NP-2 expression shifts to distinct subsets of
neurons with almost complementary patterns (Fig. 5, NP-1, and NP-2, white arrowheads).
While the expression of PlexinB1 changes little compared to earlier stages, PlexinB2
expression is down regulated in cells surrounding the spinal cord but is weakly up regulated
in motoneurons. Such an up regulation in the lateral part of the motor column is also seen for
PlexinC1 (Fig. 5, C1).
Fig. 4: Plexin and Neuropilin expression patterns in chicken spinal cord and peripheral nervous system at stage
26. Cross sections of chicken lumbosacral spinal cord were incubated with Dig labeled antisense probes.
Different probes used are indicated. At stage 26, PA1 and NP-2 expression shift to a subpopulation of the lateral
motor column (A1, black arrow), whereas PA2 transcripts are highly detected in interneurons. PA4, NP-1, and
PD1 display a broader expression in the lateral motor column (A4, NP-1 and D1 white arrows). PB2 and PC1
messages (B2 and C1 black arrowheads) are also detectable in motoneurons although at a lower level. PlexinA
and neuropilin transcript are also detected in DRGs (A1, A2, A4, NP-1 and NP-2) and PB1 (B1) messages is still
detectable in the ventricular zone. In addition, a weak expression of PB1 is detected in a subpopulation of
sensory neurons in DRGs. Scale bars correspond to 200 µm.
4.3 Results
91
PlexinA2 and PlexinC1 expression in the floor plate persist through stage 26, when
commissural axons have crossed the midline and turned into the longitudinal axis (Fig. 4, A2
and C1). At the same time, PlexinC1 expression is also seen in the dorsal spinal cord in a
position that overlaps with the position of the dorsolateral commissural neurons. During this
stage PlexinA1, PlexinA4, neuropilin-1 and neuropilin-2 expression is unchanged in DRGs
compared to stage 22.
Fig. 5: Plexin and Neuropilin expression patterns in chicken spinal cord and peripheral nervous system at stage
30. Cross sections of chicken lumbosacral spinal cord were incubated with Dig labeled antisense probes.
Different probes used are indicated. At stage 30, PlexinA, PlexinB and PC1 expression patterns remain largely
unchanged however PA4 transcript are now detectable only in a lateral subpopulation of motoneurons (A4 white
arrowhead). Strikingly, NP-1 and NP-2 display complementary expression patterns in motoneurons (NP-1 and
NP-2 white arrowheads), whereas PD1 messages are now only detected in interneurons close the ventricular
zone (D1 black arrow). Only PA1, PA4, and NP-1 continue to be expressed in DRGs. Scale bars correspond to
200 µm.
4.3.1.4 Plexins and Neuropilins at late stages of spinal cord development
During late stages of spinal cord development, between stages 35 (Fig. 6) and 40 (data not
shown), when motoneurons have reached their muscle targets and sensory afferents terminate
in their specific target layers of the gray matter, expression patterns of plexins and neuropilins
are still changing.
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Fig. 6: Plexin and Neuropilin expression patterns in chicken spinal cord and peripheral nervous system at stage
35. Cross sections of chicken lumbosacral spinal cord were incubated with Dig labeled antisense probes.
Different probes used are indicated. During this late stage of spinal cord development only PA1 transcripts are
highly expressed in motoneurons (A1), whereas PA2, PA4 and NP-1 are expressed in the dorsal horn of the
spinal cord (A2 white arrowhead, A4 white arrow and NP-1 black arrow). PA2 is strongly detected in layer І to
layer III (A2 white arrowhead) whereas NP-1 and PA4 expression are less intense. NP-2 messages become
restricted to few cells in the ventral horns of the spinal cord (NP-2 black arrow). While PB2 and PD1 are not
detected any more at this stage, PB1 expression remain unchanged and PC1 messages are up regulated in DRGs.
Scale bars correspond to 200 µm.
The expression of neuropilins gets more and more restricted to only very distinct populations
of cells. Neuropilin-1 is expressed in the dorsal horn, whereas neuropilin-2 becomes restricted
to cells in the ventral horns of the spinal cord (Fig. 6, NP-1 and NP-2, black arrows).
Similarly, the expression patterns of PlexinA get more restricted, where PlexinA4 is detected
only in the dorsal horn at stage 35 (Fig. 6, A4 white arrow) but disappears between stages 35
and 40 (data not shown). In contrast, PlexinA1 remains more or less diffusely expressed in the
gray matter. However, a slight expression is seen also in a small area of the LMC. PlexinA2 is
expressed predominantly in layers I to III (Fig. 6, A2 white arrowhead), whereas PlexinC1, at
stage 26 widely expressed in the gray matter, retracts more and more dorsally to become
totally absent from the intermediate and ventral spinal cord at stage 40 (data not shown).
In contrast to PlexinAs, which are expressed during the time when neurons extend their
axons, PlexinC1 is expressed only during late stages of neuronal development. Neither
4.4 Discussion
93
commissural neurons nor motor neurons express PlexinC1 during the time when they
approach their first intermediate target, the floor plate, and the plexus region respectively.
After stage 30, PlexinC1 is expressed transiently throughout the gray matter (Fig. 6, C1)
before it becomes restricted to the dorsal horns at stage 40 (data not shown), where it overlaps
with PlexinA1 and PlexinA2 (data not shown). Interestingly, a massive increase in PlexinC1
expression in dorsal root ganglia at stage 35 (Fig. 6, C1) is observed, and this high expression
persists at stage 40 (data not shown). At these late stages, PlexinD1 expression in endothelial
cells in no longer detected (Fig. 6, D1 and data now shown) and neuropilin-2 expression is
down regulated in DRGs (Fig. 6, NP-2). In contrast, PlexinA1, PlexinA4 and neuropilin-1
expression remain unchanged in DRGs until stage 35 (Fig. 6, A1, A4 and NP-1).
4.4 Discussion
4.4.1 Mouse plexin genes located on the X sex chromosome are absent in
chicken
Based on their sequence homology and domain organization plexins can be grouped into 4
different subclasses namely A to D. While the C and D subfamilies contain only a single
representative, four different PlexinAs and three different PlexinBs are present in the
mammalian genome. However, extensive databank searches only identified 7 chicken
counterparts compared to the nine mammalian plexins. One of the major questions in this
respect is whether we were simply unable to identify corresponding chicken fragments or
whether the chicken genome indeed contains a reduced number of plexin genes. The fact that
we did identify all the seven chicken plexins using either the EST or the chicken genomic
database demonstrates that the coverage of these databases is, even while not complete, quite
high. Interestingly the mouse variants of the two missing plexin genes are located on the X
chromosome, implying that a homologous region of this chromosome is absent in chicken.
This hypothesis is strengthened by the fact that avian use a Z/W sex determination system
instead of the normal X/Y system found in mammals (Gilgenkrantz, 2004). Moreover, in
contrast to mammals, in chicken the female is the heterogametic sex (ZW), whereas the male
is homogametic (ZZ), further suggesting that intense chromosomal rearrangements did
happen during evolutionary development (Smith and Sinclair, 2004). These changes are also
reflected by the fact that the haploid chicken genome contains 38 autosomes compared to only
19 autosomes in mouse (Vinogradov, 1999). Remarkably, the chicken genome is still only
about half the size (1.2 x 109 bp) of the mouse genome (2.56 x 109 bp), a difference that can
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Part IV: Paper 3
be largely explained by the finding that 30 of the 38 chicken autosomes, called
microchromosomes are extremely small in size ranging only between 5 and 20 Mb
(Venkatesh et al., 2000; Vinogradov, 1999). While a reduction in size is not per se an
indication for a reduced number of genes as demonstrated for many teleost, two indications
point towards a reduced number of active chicken genes. In contrast to the pufferfish Fugu
rubripes which has been shown to have extremely small gene sizes in which intron sequence
length has been reduced to a minimum (Taylor and Semple, 2002), the overall gene length in
chicken and mouse are not significantly different . Moreover, all attempts to clone chicken
homologues that could not be identified by database screening using degenerate primers in
RT-PCR reactions failed, again implying that the missing genes are indeed not represented in
the chicken genome. Nevertheless only the completion of the chicken genome sequencing
project as well as the extension of the number of sequenced chicken ESTs can finally answer
the question whether the chicken genome indeed encodes fewer plexins than the mammalian
genome does.
4.4.2 Expression patterns of plexins and neuropilins are dynamically
regulated during spinal cord development
4.4.2.1 PlexinAs and neuropilins expression patterns in chicken suggest possible
functions for PlexinAs independently from neuropilins
A variety of studies report about the role of plexins during development (Fujisawa, 2004).
While the first plexin was described quite some time ago (Ohta et al., 1995) their expression
patterns are not known in details. In particular, little is know about temporal changes in the
expression of different plexins. We have now performed a detailed spatiotemporal expression
analysis for all different chicken plexins. Most plexins and neuropilins are highly expressed
during different stages of motoneuron development, suggesting multiple roles for these
molecules in motor nerve segregation and/or defasciculation. Interestingly, motoneurons also
express different class III semaphorins and high levels of Sema5A and Sema7A (see chapter
2), implying that in analogy to results seen in the Drosophila embryo, where the
transmembrane semaphorin Іa is required for the selective defasciculation of specific
motoneurons (Yu et al., 1998), plexin- semaphorin interactions are required for similar events
in higher vertebrates.
While the function of PlexinAs has been studied predominantly in the context of their role as
a co-receptor together with neuropilins (Tamagnone and Comoglio, 2000), it is clear that in
chicken PlexinAs must have functions that are independent of neuropilins, because they are
4.4 Discussion
95
distributed much more widely in the developing nervous system than neuropilin-1 and -2. In
the dorsal spinal cord for example, commissural neurons express all three members of the
plexin-A class but neither neuropilin-1 nor neuropilin-2. This expression pattern is divergent
from the widespread expression of neuropilin-2 in the spinal cord of the E10.5-12.5 mouse
(Brown et al., 2001; Chen et al., 2000), where neuropilin-2 is expressed very strongly in
dorsal commissural neurons and in all ventral populations of interneurons.
Interestingly, all PlexinAs are expressed also in the floor plate. PlexinA1 is expressed only in
lateral but not medial floor plate cells. Expression of plexins in the floor plate is rather
surprising, as the floor plate is the intermediate target of commissural axons, and therefore,
the site where ligands for axonal receptors are expected. A receptor function of plexins in
floor plate cells at stage 22 is less obvious, as these cells do not migrate or undergo structural
remodeling at this time. Floor plate development seems to be terminated much earlier
(Briscoe and Ericson, 2001; Briscoe et al., 2000; Jessell, 2000; Wijgerde et al., 2002).
Interestingly, the floor plate is also the earliest site of PlexinC1 expression in the spinal cord
where the expression is observed by stage 22 and last until stage 30, although much weaker.
The expression of several plexins in the floor plate might suggest a possible role for these
plexins as a ligand rather than a receptor as always presumed.
4.4.3 PlexinBs and PD1 are only transiently expressed in neurons
The expression of class-B plexins is difficult to link to any specific function. Both plexin B
members are expressed in neurons. The most prominent and longest lasting expression for a
class-B plexin is seen in the ventricular zone, where PlexinB1 is expressed from stage 22 to
stage 35. This pattern is consistent with PlexinB1 expression seen in mouse, where PB1
transcript was found in the ventricular zone of the spinal cord at E13.5 (Worzfeld et al.,
2004). The expression of chicken PlexinB2 is generally very weak with the exception of cells
surrounding the spinal cord at stage 22. These cells can be crest cell derivatives, however no
expression in sensory ganglia neurons is observed at any stage, or endothelial cells of the
perineural vascular vessels sprouting from the primitive arterial tract. In contrast to the
observations in mouse, PlexinBs are not expressed during the time when sensory afferents
target their specific layers in gray matter, as collaterals of primary sensory axons in chicken
do not form before stage 29 (Perrin et al., 2001), an age where PlexinB1 and PlexinB2 are
absent in DRG. Thus, although a contribution of both PlexinBs to the formation of the central
sensory connections is still possible it seems rather unlikely. Taken together, chicken
PlexinBs seem to have slightly altered expression patterns and functions when compared to
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Part IV: Paper 3
mouse PlexinBs a result that is hardly surprising since the chicken genome lacks one PlexinB
family member.
PlexinD1 has been reported to be expressed in endothelial cells in mice and zebrafish and
play an important role in vivo in vascular system development (Gitler et al., 2004; TorresVazquez et al., 2004). In chicken, PlexinD1 is expressed as early as stage 18 in endothelial
cells of the perineural vascular plexus and this expression persists until stage 30. In addition
PlexinD1 is expressed in developing vessels throughout the entire embryo suggesting an
important role for this protein during early stages of angiogenesis as earlier observed in mice
and zebrafish (Gitler et al., 2004; Gu et al., 2004; Torres-Vazquez et al., 2004). However,
PlexinD1 in chicken is also expressed in the central nervous system in spinal motor neurons at
the time motor axons sort in the limb plexus before entering the limb muscle mass. This is in
concordance with earlier published report demonstrating the expression of PlexinD1 in mouse
brain during embryonic development (van der Zwaag et al., 2002).
4.4.4 PlexinC1 is not expressed in early stages of neuronal development
In contrast to PlexinAs, which are expressed during the time when neurons extend their
axons, PlexinC1 is expressed only during late stages of neuronal development. Neither
commissural neurons nor motor neurons express PlexinC1 during the time when they
approach their first intermediate target the floor plate and the plexus region respectively.
Interestingly, strong expression of PlexinC1 is seen in the floor plate at stage 22, i.e. when the
majority of the axons from dorsolateral commissural neurons are in the floor plate. The
expression in the floor plate persists through stage 25, when commissural axons have crossed
the midline and turned into the longitudinal axis (Bourikas, 2005). At that time expression is
also seen in the dorsal spinal cord in a position that overlaps with the position of the
dorsolateral commissural neurons (Bourikas, 2005; Stoeckli and Landmesser, 1995). While
PlexinC1 expression in commissural interneurons is consistent with the expression of
Sema7A in the floor plate, the semaphorin known to bind to PlexinC1 (Tamagnone et al.,
1999), PC1 messages in the floor plate cells suggest homo or heterophilic plexin interactions
rather than binding with Sema7A.
Interestingly, there is a massive increase in PlexinC1 expression in dorsal root ganglia at stage
35, a time when all other plexins are down regulated and expression persists until stage 40.
Thus, PC1 starts to be expressed when axons have already completed the navigation to their
targets suggesting that PC1 might be involved in target recognition or synaptogenesis rather
than pathfinding.
Part V:
Conclusion and Outlook
97
Conclusion and Outlook
The adult nervous system is organized in remarkably complex networks capable of achieving
a broad range of activities including very complicated functions such as learning. The proper
functioning of this elaborate network results from the establishment of functional connections
between the numerous neuronal cells that are generated during development. Since the early
1990s, a constantly increasing number of studies describe axon guidance molecules and their
implication in vitro as well as in vivo in steering different types of neural axons or in affecting
the migratory paths of neural cells (Johansen and Johansen, 1997; Young et al., 2004).
However, the precise details of how neuronal wiring takes place remain to a large part
unknown.
This is due in part to the fact that the path of a growing axon appears to be very complex in
vivo and implicates several families of guidance molecules. Moreover, several members of the
same family of axon guidance may act in concert to direct the growth cone, through a series
of attractive and/or repulsive events. Additionally, the expression of guidance molecules
outside the nervous system and their implication in the growth of other tissues makes the
understanding of how axon guidance works even harder.
Plexins and their ligands, the Semaphorins, have been intensively characterized during the last
years. They have been demonstrated to play crucial roles in directing axonal outgrowth as
well as neural crest cells migration during brain and/or spinal cord development (Fiore and
Puschel, 2003; Fujisawa, 2004). However, their expression patterns extend beyond the
nervous system, suggesting their involvement in heart, bone, lung, intestine, immune system
and vascular system development (Park et al., 2004; Serini and Bussolino, 2004). Such wide
spread expression in several vital tissues made the generation of mutant mice very
complicated, since conventional knock out techniques often resulted in early embryonic
lethality related to vital function disruptions (Behar et al., 1996; Gitler et al., 2004).
Recent progresses, in alternative procedures to knock down a particular gene function, have
provided new possibilities to study plexin and semaphorin activities. We employed the
chicken embryo as a model system to knock down PlexinD1 transcripts using in ovo RNAi, a
technique based on the electroporation of long double stranded RNA injected into the central
canal of the chicken embryo (Pekarik et al., 2003). This system provides a powerful tool to
explore exclusively the role of axon guidance molecules within the spinal cord, circumventing
the problem of embryonic lethality due to severe defects in vital tissues. This seems certainly
necessary since our spatio-temporal expression analysis demonstrates that semaphorin and
98
Conclusion and Outlook
99
plexin expression in the chicken spinal cord is very complex, suggesting multiple functional
interactions within the same system (chapter 2 and chapter 3).
PlexinD1 activity in vivo has been extensively investigated during vasculogenesis.
Nevertheless, PlexinD1 is also expressed in the central nervous system, however no
functional role in nervous system development has been reported (Gesemann et al., 2001; van
der Zwaag et al., 2002). Our study demonstrates that PlexinD1 is expressed in spinal motor
neurons of chicken embryos between stages 23 and 26, the period motor axons sort within the
limb plexus before initiating the limb innervation. PD1 knock down results in severe defects
in the formation of the crural nerve, suggesting a functional role for PlexinD1 in axon sorting
at the plexus level (chapter 1). Immunostaining of chicken spinal motoneurons on cross
sections using MNR2 antibody that marks all motoneurons subtypes show no differences in
the positioning of motoneurons within the spinal cord in PlexinD1 knock down embryos.
However, MNR2 marker is a general marker for all motoneurons subtypes and does not give
any indication about the arrangement of different motoneuron pools. To exclude that PD1
knock down in chicken spinal indeed causes incorrect positioning of motoneurons in different
motor pools which could lead to abnormal motor axon outgrowth, supplementary experiments
using specific markers for each motor pool such as different cadherins or ETS transcription
factors are required. However, a function for PlexinD1 in motoneurons sorting seems unlikely
because PD1 mRNA is expressed in motoneurons only after stage 23, at this time all
motoneurons are post mitotic and have already migrated to their final position within different
pools.
Surprisingly, PlexinD1 loss of function also leads to abnormalities in the outgrowth of
sensory afferents although PlexinD1 was never detected in the dorsal sensory neurons.
Additionally, PD1 knock down embryos exhibit defasciculation defects at the ventral motor
roots and disorganization of the boundary cap cells in the region of ventral motor exit points.
Two different hypotheses could be raised to explain the observed phenotypes. In the first
hypothesis, PlexinD1 positive cells outside the spinal cord are not exclusively endothelial
cells but include a migratory subpopulation of crest cells, notably Schwann cells and/or
boundary cap cells. PlexinD1 knock down in crest cell derivatives might affect the normal
migration of these cells and subsequently lead to their incorrect positioning which can explain
the dorsal root abnormal arrangement as well as the incorrect ventral motor roots formation.
A potential role for PlexinD1 in cell migration goes along with earlier observations
demonstrating that PD1 in postnatal rats is exclusively expressed in three brain nuclei, the
100
Conclusion and Outlook
pons, the inferior olive and the dorsal column nuclei; structures that form by tangentialy
migrating cells originating from the precerebellar neuroepithelium (Altman and Bayer, 1987).
To verify this hypothesis, several additional experiments are required. In a first step and in the
absence of available antibody against PlexinD1 protein, double in situ hybridization
experiments to co-localize PlexinD1 expression outside the spinal cord with a crest cell
marker such as Sox-10 or HNK-1 are required. Additionally, these same markers can be used
also to investigate the migratory pattern of crest cells derivatives in PD1 knock down embryos
in comparison to controls. In addition, several specific boundary cap cells markers can be
used such as Krox20 or Sema6A (that is exclusively expressed in boundary cap cells in
chicken) to check the integrity of boundary cap cells formation in PlexinD1 knock down
animals.
Alternatively, the aberrant dorsal roots formation as well as the ventral motor roots defects
could result from the abnormal formation of certain vessels running in close contact and/or
interacting with growing axons. This hypothesis is based on observations reported in many
recent publications demonstrating an implication of PlexinD1 in vascular system formation
(Gitler et al., 2004; Gu et al., 2004; Torres-Vazquez et al., 2004). PlexinD1 knock out mice,
exhibit a severe disorganization of the normally highly structured and ordered intersomitic
vessels. In the case that PD1 is exclusively expressed in endothelial cells, PlexinD1 knock
down in chicken embryos might lead to the disorganization of intersomitic vessels, and
consequently these alterations may cause abnormal sensory axonal growth. This idea is
further supported by evidence that a tight cellular and/or molecular interaction between
vasculogenesis and axonal outgrowth especially between intersomitic arteries and outgrowing
sensory axons in xenopus embryos exist (Levine et al., 2003).
To investigate this hypothesis, a very important issue is to check whether in our system
PlexinD1 loss of function is achieved also in endothelial cells surrounding the neural tube. To
date we do not have any indication about the possible targeting of intersomitic vessels cells
using in ovo RNAi. To the contrary, our data seems to demonstrate a restricted targeting of
cells, exclusively within the spinal cord and dorsal root ganglia, as seen by the electroporation
of YFP plasmids. However, we cannot exclude the possibility that long double stranded RNA
targets more cells than cDNA constructs, as it is smaller and might have a different charge
compared to DNA plasmids.
In order to resolve this issue, we need to use a different approach replacing the use of long ds
RNA by a recombinant construct, containing an YFP reporter gene followed by a PlexinD1
Conclusion and Outlook
101
small interfering RNA. However, to date we did not assess the efficiency of small interfering
RNA to knock down PlexinD1 expression.
Another important matter required for verifying the plausibility of this hypothesis, is to
investigate the integrity of intersomitic vessels patterning in PD1 knock down animals.
Unfortunately, no antibodies recognizing the chicken vascular system are yet available
making the use of alternative staining methods, such as the perfusion of certain lectins that
can bind to endothelial cells and allow the staining of the chicken vascular system, necessary
(Hagedorn et al., 2004).
Furthermore, it is of high interest to investigate whether the identified PlexinD1 binding
partners (Sema3A, Sema3C, and Sema3E) are used also in chicken embryo and are implicated
in PlexinD1 signaling in spinal motoneurons. In this regard, the chicken embryo represents an
excellent model system, as the RNAi technique offers a major advantage to gene disruption in
mice, which is the possibility of knocking down several genes simultaneously. This is
important in order to avoid redundancy and to tackle the combinatorial activity of multiple
axon guidance molecules acting in chorus. In this respect, it will be interesting to investigate
axon outgrowth in single and multiple semaphorins knock down embryos. However, in order
to assess the binding between PlexinD1 and Sema3A, 3C and 3E in chicken, the cloning of
these genes is required. The cloning of chicken PlexinD1 recombinant construct offers also
the possibility to over express PlexinD1 in chicken spinal cord in vivo in order to unravel the
potential function of this protein during development.
In conclusion, our current results demonstrate clearly the expression of PD1 mRNA in
chicken spinal motoneurons during the time motor axons sort in the limb plexus, suggesting a
potential role for this protein in motor axon guidance. Indeed PlexinD1 knock down in
chicken embryos using in ovo RNAi lead to motor axon pathfinding defects in the crural
nerve trunk. Further experiments will allow us: to explain the aberrant formation of the dorsal
and ventral roots in PlexinD1 knock down animals and to unravel whether PlexinD1 plays a
novel role in the correct migration of a subpopulation of neural crest cell derivatives and/or a
tight relation between intersomitic vessels sprouting and dorsal and motor root formation.
102
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List of Publications
P1
J. Gemayel, A. Geloen, F. Mion, “Propofol-induced cytochrome P450 inhibition: an in
vitro and in vivo study in rats“, Life Science, 68, 2957–65, 2001.
P2
O. Matui, J. Gemayel, R. Sadhu, M. Gesemann, E. Stoeckli, „Expression patterns of
plexins and neuropilins suggest cooperative separate functions in spinal cord
development”. Submitted in June 2005 to Developmental Dynamics.
P3
J. Gemayel, R. Sadhu, O. Matui, E. Stoeckli, M. Gesemann, „Developmental regulation
of semaphorins in spinal cord and peripheral nervous system suggests additional
semaphorin functions and interactions in chicken embryo. In preparation (will be
submitted in July 2005 to Developmental Dynamics).
P4
J. Gemayel, R. Sadhu, R. Babey, E. Stoeckli, M. Gesemann, „Functional knock down
of Plexin D1 in chicken results in misguided motor axons and alterations in dorsal and
ventral roots organization“. In preparation.
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Curriculum Vitae
Personal data
Name:
Joelle Gemayel
Nationality:
Lebanese
Date of birth:
May 27, 1974
E-mail:
[email protected]
Education
11/00 − 02/05:
ETH Zürich, Zurich, Switzerland
Ph. D. student in Neurobiology at Brain Research Institute
Specialization: “Role of PlexinD1 in nervous system development”
09/98 − 10/00:
Université Claude Bernard, Lyon, France
DEA “Metabolism, Endocrinologie et Nutrition”
09/97 − 08/98:
Université de Brest, Brest, France
Maitrise in Physiology and Cell Biology
10/91 − 09/95:
Lebanese University Fanar, Beirut, Lebanon
Maitrise in Animal Physiology
Professional experience
09/98 − 10/00:
Université Claude Bernard, Lyon, France
Teaching assistant in practical physiology courses for sports’ athletes
01/96 − 06/97:
Tamer Group, Beirut, Lebanon
Sales representative for Tamer Group dental division
Responsible for marketing and sales of dental products to dentists
119