Role of the Cell Adhesion Molecule L1 during
Early Neural Development in Zebrafish
by
Wanyi Xiang
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Graduate Department of Biochemistry
University of Toronto
© Copyright by Wanyi Xiang, 2008
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Role of Cell Adhesion molecule L1
during Early Neural Development in Zebrafish
Wanyi Xiang
Doctor of Philosophy
Graduate Department of Biochemistry
University of Toronto
2008
Abstract
The neural cell adhesion molecule L1 is a member of the immunoglobulin superfamily and it mediates many adhesive interactions during brain development.
Mutations in the L1 gene are associated with a spectrum of X-linked neurological disorders known as CRASH or L1 syndrome. The objective of this thesis was to use the zebrafish model to investigate the molecular mechanisms of L1 functions and the pathological effects of its mutations. Zebrafish has two L1 homologs, L1.1 and L1.2.
Inhibition of L1.1 expression by antisense morpholino oligonucleotides resulted in phenotypes that showed resemblances to L1 patients. However, knockdown of L1.2 expression did not result in notable neural defects. Furthermore, analysis of the expression pattern of L1.1 has led to the discovery of a novel soluble L1.1 isoform,
ii L1.1s. L1.1s is an alternatively spliced form of L1.1, consisting of the first four Ig-like domains and thus a soluble secreted protein.
L1.1 morphants exhibited disorganized brain structures with many having an enlarged fourth/hindbrain ventricle. Further characterization revealed aberrations in ventricular polarity, cell patterning and proliferation and helped differentiate the functions of L1.1 and L1.1s. While L1.1 plays a pivotal role in axonal outgrowth and guidance, L1.1s is crucial to brain ventricle formation. Significantly, L1.1s mRNA rescued many anomalies in the morphant brain, but not the trunk phenotypes. Receptor analysis confirmed that L1.1 undergoes heterophilic interactions with neuropilin-1a
(Nrp1a). Peptide inhibition studies demonstrated further the involvement of L1.1s in neuroepithelial cell migration during ventricle formation. In the spinal cord, spinal primary motoneurons expressed exclusively the full-length L1.1, and abnormalities in axonal projections of morphants could be rescued only by L1.1 mRNA. Further studies showed that a novel interaction between the Ig3 domain of L1.1 and Unplugged, the zebrafish muscle specific kinase (MuSK), is crucial to motor axonal growth. Together, these results demonstrate that the different parts of L1.1 contribute to the diverse functions of L1.1 in neural development.
iii Acknowledgements
I would like to express my gratitude to my supervisor, Dr. Chi-Hung Siu, for
cultivating an environment that fosters independency, self-discipline and creativity in my
scientific career. I have gained from the opportunity to explore, inquire, and discover.
Science tells the truth as authority, in lieu of the authority as truth.
I am sincerely grateful to two other members of my supervisory committee, Dr.
Chi-chung Hui, and Dr. David Clarke. Their insightful advices and expert inputs have led to many of the discoveries during the course of this project. I am also very appreciative of their generosities and having their laboratory instruments accessible.
Many friends have come and gone in the Siu lab, and I am gratified by their kindness and help. In particular, I thank Dr. Paul Yip, Shrivani Sriskanthadevan, Eric
Huang, and Linda Sun. I enjoy sharing scientific opinions, daily life, and emotional stories with them.
I also thank the Canadian Institutes of Health Research and University of Toronto for their financial support.
My final dedication is to my family. I am forever grateful for my parents’ endless love, and their heritage recipes for nutritious soup that keep me healthy even during the toughest days. I am always inspired by my loving husband with his passion for science and enthusiasm for the future. From our debates on almost everything, I have gained many interesting ideas and am certainly marveled by his broad knowledge and interests. I am sure that we will tie on the next debate topic- as our title tells.
iv TABLE OF CONTENTS
Abstract ii Acknowledgements iv Table of Contents v List of Figures vii List of Tables ix List of Abbreviations x
Chapter I: Introduction 1. Cell Adhesion in Development and Diseases 2 2. Immunoglobulin Superfamily in Neural Development 17 3. Cell Adhesion Molecule L1 and CRASH Syndrome 29 4. L1-Associated Interactions 39 5. L1-Mutant Mouse Models 47 6. The Zebrafish Model 50 7. Embryonic Neural Development in Zebrafish 59 8. A Comparative Analysis of Embryonic Brain Development in the 71 Vertebrates 9. Identification of Zebrafish L1 Homologs 80 10. Hypothesis and Rationale of the Thesis 82
Chapter II: Knockdown of L1.1 Expression in Zebrafish Results in Hydrocephalus-Like Phenotype and Reveals a Role in Ventricular Polarization during Brain Development 1. Summary 85 2. Introduction 86 3. Materials and Methods 89 4. Results 98 5. Discussion 122
v Chapter III: Interactions between a Novel Soluble Form of L1.1 and Neuropilin 1a Regulate Brain Ventricle Formation in Zebrafish 1. Summary 128 2. Introduction 129 3. Materials and Methods 131 4. Results 141 5. Discussion 166
Chapter IV: Role of L1 during Axonal Path Finding of Primary Motoneuron in Zebrafish 1. Summary 171 2. Introduction 172 3. Materials and Methods 175 4. Results 181 5. Discussion 193
Chapter V: Conclusions and Future Perspectives 1. Differential functions of zebrafish L1 homologs in embryonic 199 development 2. Role of L1s in brain ventricle formation and the pathological 199 mechanism of L1-associated hydrocephalus 3. L1s-neuropilin 1 interaction in the brain ventricle development 203 4. Neuropilin 1-mediated signaling 206 5. Role of L1 at the neuromuscular junction 208 6. Signaling in L1-mediated postsynaptic differentiation 211 7. Role of L1s in pre-assembling postsynapse 213
References 216
Appendix 247
vi LIST OF FIGURES
Chapter I:
1.1 Major families of cell adhesion molecules 4 1.2 The structure of members of the Ig superfamily implicated in growth 18 and guidance in the nervous system 1.3 Phylogenetic analysis of the L1 gene family 20 1.4 Conserved features of the cytoplasmic tail of L1 family members 23 1.5 Schematic diagram of domain involvement to specific L1 extracellular 40 interactions. 1.6 Early development of the zebrafish embryo 60 1.7 Patterning of the zebrafish embryonic brain 72 1.8 Different stages of radial neuronal migration in the developing brain 75
Chapter II:
2.1 Alignment of zebrafish L1.1 and L1.2 with human L1 100 2.2 Neuritogenic activity of recombinant L1.1 and L1.2 102 2.3 Expression of L1.1 in the embryonic brain 103 2.4 Knockdown of L1.1 resulted in abnormal embryonic development 106 2.5 Morpholino knockdown of zebrafish L1.2 uncorrelated with distinct 108 developmental defects 2.6 Anomalies in brain ventricle formation associated the enlargement of 111 the 4th ventricle in L1.1 morphants 2.7 Abnormal cell dense zones in the L1.1 deficient brain 113 2.8 Expression of brain marker genes pax2.1 and pax6 in wild-type and 115 morphant brains 2.9 Abnormal ventricular development in the absence of L1.1 117 2.10 BrdU mapping of cell patterning 119 2.11 Effects of L1.1 deficiency on brain cell proliferation 121
vii Chapter III:
3.1 Detection of a novel alternatively spliced isoform of L1.1 142 3.2 Cloning and identification of the short alternatively spliced isoform of 143 L1.1 3.3 Expression of the L1.1s protein during embryonic development 146 3.4 Neuritogenic activity of L1.1s 14 3.5 Differential expression of L1.1s and L1.1 in the embryonic brain 151 3.6 L1.1s mRNA rescued ventricle formation but not axonal growth in L1.1 154 morphant 3.7 Involvement of L1.1 sequence YAANEL in L1-Nrp1a interaction 157 3.8 Restoration of ventricular polarization by local injections of 160 recombinant L1.1s 3.9 Inhibition of L1.1-Nrp1a interaction by synthetic peptide disrupted 161 ventricle formation in the midbrain 3.10 Involvement of L1.1-Nrp1a interaction in neuronal migration at the 164 diencephalic ventricle
Chapter IV:
4.1 Differential expression of L1.1 isoforms in spinal neurons and somites 182 4.2 Aberrant motor axonal growth in L1.1 morphants 184 4.3 Co-immunoprecipitation of L1.1 with Unplugged FL 187 4.4 Colocalization of L1.1 and Unplugged FL along the axonal path 189 4.5 Direct binding between L1.1 and Unplugged 190 4.6 Effects of pathological L1 mutations on the development of motor 192 axons
Chapter V:
5.1 Involvement of L1.1s in the development of brain ventricle 201 5.2 Schematics depicting NPC migration in vivo mediated by L1.1s-Nrp1a 205 interactions 5.3 Schematic illustration of the interaction between neuronal L1.1 and 209 Unplugged during synaptogenesis along the CaP axonal path
viii LIST OF TABLES
Chapter I:
1.1 Functional interactions of CAMs with signaling molecules 11 1.2 Phenotypes of animal models deficient in CAMs 14 1.3 Conserved characteristics in the cytoplasmic tail of L1 gene superfamily 24 1.4 Missense mutations in L1 gene associated with CRASH syndrome 35 1.5 Genotype and phenotype correlation of L1 mutations with severity of 37 CRASH syndrome 1.6 Phenotype and cell biology of selected L1 mutations 38 1.7 The zebrafish toolbox 53
Chapter II:
4.1 Abnormal axonal projection from CaP in L1.1 morphants of 24 hpf 186
ix LIST OF ABBREVIATIONS
AChR Acetylcholine receptor BMP Bone morphogenetic protein BrdU 5-Bromo-2’-deoxyuridine BSA Bovine serum albumin CAM Cell adhesion molecule CaP Caudal primary motoneuron CHL1 Close homolog of L1 CNS Central nervous system CRASH Corpus callosum hypoplasia, mental retardation, adducted thumbs, spastic paraplegia, and hydrocephalus CSF Cerebrospinal fluid CST Corticospinal tract DAB 3,3'-Diaminobenzidine DMSO Dimethyl sulfoxide ECM Extracellular matrix EGF Epidermal growth factor EGFP Enhanced green fluorescence protein ENU Ethylnitrosourea ERK Extracellular signal-regulated kinase ERM Ezrin-radixin-moesin FGF Fibroblast growth factor FGFR Fibroblast growth factor receptors FNIII Fibronectin type III repeat Hh Hedgehog hpf Hour post fertilization HRP Horse radish peroxidase Ig Immunoglobulin IgSF Immunoglobulin superfamily L1.1EC L1.1 extracellular domain
x MAG Myelin-associated glycoprotein MAGUK Membrane-associated guanylate kinase-family scaffolding protein MHB Mid-hindbrain boundary MiP Middle primary motoneuron MLF Medial longitudinal fasciculus MuSK Muscle specific kinase NMJ Neuromuscular junction NPC Neuronal progenitor cell Nrp1 Neuropilin 1 NTP/BCIP Nitro blue tetrazolium chloride/ 5-Bromo-4-chloro-3-indolyl phosphate pax Paired box PBS Phosphate buffered saline PCR Polymerase chain reaction PFA Paraformaldehyde pH3 Phosphohistone H3 RACE Rapid amplification of complementary DNA ends RoP Rostral primary motoneuron RT-PCR Reverse-transcription-polymerase chain reaction SDS Sodium dodecylsulfate SDS-PAGE SDS-polyacrylamide gel electrophoresis Sema3A Semaphorin 3A TILLING Targeting-induced local lesions in genomes TUNEL Terminal deoxynucleotidyl transferase dUTP nick end labeling UTR Untranslated region
xi
Chapter I:
Introduction
1 1. CELL ADHESION IN DEVELOPMENT AND DISEASES
Over 10 million years after the emergence of the first multicellular organisms,
evolution has given rise to remarkably diverse and sophisticated life forms that surround us. The evolution of multicellularity permitted the assemblies of specialized cells and tissues. A crucial step during tissue morphogenesis must have been the ability of cells to contact tightly and interact specifically with other cells (Chothia and Jones, 1997).
A variety of cell adhesion mechanisms are responsible for assembling cells together and, along with their connections to the internal cytoskeleton, determine the overall architecture of the tissue (Gumbiner, 1996). To establish functional adhesion units, animal cells express a variety of cell adhesion molecules (CAMs) or adhesion receptors that enable the same type of cells to adhere tightly and specifically, or otherwise
discriminate and segregate themselves into distinct tissues (Gumbiner, 1996; Chothia and
Jones, 1997). Animal cells also secrete a complex network of proteins and carbohydrates
to assemble the extracellular matrix (ECM), which creates a special environment in the
space between cells. At the extracellular surface, the cell adhesion molecules recognize
and interact either with other cell adhesion receptors on neighboring cells or with proteins
of the ECM. At the intracellular surface of the plasma membrane, cell adhesion receptors
associate with peripheral membrane proteins, which serve to link the adhesion systems to
the cytoskeleton, to regulate the functions of the adhesion molecules, and to transduce
signals initiated at the cell surface by the adhesion receptors (Gumbiner, 1996).
In these three-dimensional tissue assemblies, the cell-cell and cell-matrix
interactions mediated by CAMs participate in cell-cell communication, structural support,
locomotion, and gene expression, in addition to adhesiveness. Defects in these complex
2 interactions will result in developmental malformations and diseases (Lodish et al., 2000;
Harwood and Coates, 2004).
1.1. Cell adhesion molecules
The advance of molecular cloning has led to the discovery of a large diversity of
CAMs. Vertebrate CAMs are grouped into four major superfamilies: the cadherins,
immunoglobulin (Ig)-like superfamily (IgSF) of CAMs, integrins, and selectins (Figure
1.1). Cadherins are the major CAMs responsible for Ca2+-dependent homophilic adhesion
constituting cell-cell junctions in vertebrate tissues. The Ig CAM superfamily is
comprised of the most diverse family members of recognition molecules, which contain
one or more characteristic immunoglobulin-like domain. The integrin receptors are
crucial for cell binding to the extracellular matrix, as well as cell-cell adhesion between
adjacent cells. They are widely involved in many developmental processes. The selectins
are carbohydrate-binding proteins and play a prominent role in the circulation of
lymphocytes and neoplastic cells.
1.1.1. Cadherins
The cadherin superfamily is a group of cell surface receptors that is comprised of
more than 100 members (Takeichi, 2007). The cadherin family of proteins mediate Ca2+- dependent adhesion and play key roles in development and disease (Cavallaro and
Christofori, 2004; Halbleib and Nelson, 2006). They are the major transmembrane components of adherens junctions and desmosomes, and cluster at sites of cell-cell contact in most solid tissues (Takeichi, 1995; Gumbiner, 2000; Yagi and Takeichi, 2000).
3 Cadherins Ig CAMs Integrins Selectins
α subunit β subunit
αA domain Lectin βA domain domain β-propellor Ca2+ Ig-like domain Hybrid EGF-like domains domain domain Ca2+ S-S EC Ca2+ β-sandwich domains S-S S-S S-S S-S EGF-like CCP domains repeats modules
Plasma Plasma membrane membrane
E-cadherin NCAM α1β1 P-selectin
Figure 1.1. Major families of cell adhesion molecules. CAMs are a diverse group of integral membrane proteins, generally containing a cytoplasmic domain. There are four major families of CAMs which are based on their structural characteristics. One prototype molecule from each family is illustrated. The classical cadherins consists of five extracellular cadherin repeats (EC domains). They mediate Ca2+-dependent adhesion and interact via homophilic interaction. Members of Ig superfamily contain one or more domains which show homology to the immunoglobulin (folded by Cys- Cys disulphate bond and represented by loops). IgSF members undergo both homophilic and heterophilic interactions. Integrins are comprised of αβ heterodimers, are involved in cell-cell as well as cell-matrix interactions. Selectins recognize specific carbohydrate groups via its lectin-like domain. They also contain an EGF-like domain and multiple complement control protein (CCP) modules.
4 The first three identified cadherins are epithelial (E)-cadherin, neuronal (N)-
cadherin, and placental (P)-cadherin. Although they were named according to the main
tissues in which they were first found, all of them are found in many other tissues.
Members of the cadherin family are characterized by extracellular cadherin (EC) repeats of varying number. Classical cadherins mediate Ca2+-dependent homophilic protein-
protein interactions via the cadherin extracellular domains, elaborating both cis and trans
orientations in a zipper-like fashion (Wheelock and Johnson, 2003). This mode of
interaction is evidenced by a recent structural analysis using electron tomography, which
indicates that individual cadherin molecules form groups and interact through their tips in
a highly flexible manner (He et al., 2003). Another interesting feature of the interactions
constituted by classical cadherins is that an exquisite specificity determines a restricted
homophilic interaction exclusive to the same type of cadherin on another cell (Miyatani
et al., 1989; Cavallaro and Christofori, 2004).
The intracellular domains of classical cadherins interact with β-catenin, γ-catenin
and p120 to assemble the cytoplasmic cell adhesion complex that is critical for the
formation of extracellular cell-cell adhesion. β-catenin and γ-catenin bind directly to α-
catenin, which is linked to actin and affects actin polymerization. Therefore, the
extracellular cadherin-mediated interaction is transmitted and influences the actin
dynamics at the cell-cell contacts (Yamada et al., 2005; Halbleib and Nelson, 2006).
1.1.2. Ig SF members
Members of the IgSF consist of the largest and most diverse group of cell
recognition molecules. More than 100 members have been identified playing key roles in
5 the immune system, in the nervous system, and in other tissues as well (Maness and
Schachner, 2007). It is believed that the duplication and diversification of genes from a
limited number of genes has led to the generation of multifarious families of recognition
molecules (Thomsen et al., 1996). Proteins within this superfamily share a common
structural element, with one or more domains showing homology to those present in the
immunoglobulins (Edelman and Crossin, 1991). Most of these proteins contain a transmembrane domain and a cytoplasmic domain. However, they vary greatly in terms
of their overall size and domain organization, and the sequence identity between two Ig
domains may be as low as 20%. Members of the IgSF usually mediate cell-cell adhesion
in a Ca2+-independent manner, which may involve both homophilic and heterophilic
interactions. Although binding frequently occurs between IgSF proteins, several of them
are capable of interacting with CAMs from other families as well as the ECM (Maness
and Schachner, 2007). For example, endothelial junctional adhesion molecule 1 binds
leukocyte integrin αLβ2 during inflammatory recruitment of leukocyte (Ostermann et al.,
2002).
1.1.3. Integrins
The integrins are major cell receptors for cell-matrix and cell-cell interactions
(Hynes and Zhao, 2000; Hynes, 2002). Integrins are heterodimeric glycoproteins
constituted by the non-covalent binding of α and β subunits. At present, 8 β and 18 α
subunits have been identified in mammals but only 24 distinct pairs are known to
assemble into receptors (Hynes and Zhao, 2000; Stupack, 2007). Each of the 24 integrins
appears to play diverse role and have a specific, non-redundant function in most
6 biological processes. The binding of integrins to their ligands necessitates extracellular divalent cations (Ca2+ or Mg2+, depending on the integrin, Hynes, 2002).
Functions of integrins are known to be allosterically regulated by conformational
change upon binding of their specific extracellular ligands. Activation of the integrin
subsequently triggers intracellular signaling cascades, often acting in concert with G
protein-coupled or kinase receptors (Hynes, 2002). Paralleled to the “outside-in”
activation, the binding of an integrin receptor to its ligand can be enhanced by the effects
on the cytoplasmic domains, which are referred to as “inside-out” signaling (Ginsberg et
al., 1992; Hynes, 1992). Therefore, the coupling of ligand to the integrin receptor is
regulated in a bidirectional and reciprocal equilibrium (Hynes, 2002).
1.1.4. Selectins
The selectins are type I membrane glycoproteins. The three known selectins are
named according to the cell type on which they are primarily expressed as L (leukocyte)-
selectin, E (endothelial)-selectin and P (platelet)-selectin, which is also expressed in
endothelial cells (Foxall et al., 1992). L-selectin was first discovered in mediating
lymphocyte recognition of high endothelial venules in the lymph node and homing into
the lymph nodes (Gallatin et al., 1983; Arbones et al., 1994). Selectins are well-known
for their pivotal roles in inflammatory leukocyte trafficking in both acute and chronic
settings (Simon and Green, 2005; Wang et al., 2007). The selectin system is also
involved in other processes such as leukocyte diapedesis (Mayadas et al., 1993; Arbones
et al., 1994), hematogenous metastasis of carcinoma cells (Rosen, 2004), demyelination
7 of axons (Huang et al., 1994), and implantation of the early mammalian embryo
(Genbacev et al., 2003).
Selectins share common structure with a calcium-dependent lectin domain,
followed by an epidermal growth factor (EGF)-like motif, a series of consensus repeats, a
transmembrane domain and a short cytoplasmic tail (McEver, 2002). Selectins bind
preferentially to ligands that are modified with carbohydrates, such as sialic acid and
fucose. For examples, the sialyl Lewis X (sLex) oligosaccharide is a common epitope for all three selectins (Foxall et al., 1992). On the other hand, P- and L-selectin, but not E- selectin, also bind in a Ca2+-independent manner to sulfated glycans such as heparin
(Varki, 1997).
1.2. Regulation of extracellular interactions of CAMs
Cell adhesion molecules are pivotal to the development and maintenance of tissue
structure in multicellular organisms. Many of the adhesive contacts require temporal-
spatial regulation of focal adhesions. For example, melanoma cells migrating through the endothelial monolayer of the blood vessel requires rapid establishment of local adhesive
assembly at the leading edge and dissociation of adhesion at the rear. In the mean time,
the migratory cell changes its morphology by coupling intracellular cytoskeletal filament
in order to pass between the endothelial cells (Christofori, 2003). Mechanisms regulating
the adhesiveness in the ectodomains of CAMs include affinity modulation (the strength of non-covalent bonding between the interactors), as well as valency modulation (the surface density of the interactors (Carman and Springer, 2003).
8 With respect to affinity interaction, a unique segment or motif defines the homophilic or heterophilic interactions of CAMs. For example, α5 and αv integrins specifically recognize ligands that contain three-peptide sequence arginine-glycine- aspartic acid (RGD), such as L1 CAM and the matrix proteins vitronectin and fibronectin
(Ruoslahti, 1996). This ligand-dependent binding may often lead to conformational changes of the CAM that can enhance the interaction avidity (Humphries et al., 2003).
Valency modulation of CAM adhesiveness occurs independently of ligand engagement. Instead, it is achieved by regulating the density of the receptor on the cell surface and subsequently the number of adhesive bonds at the interaction interface. One of the mechanisms mediating prevalence of CAMs is via the interaction between their cytoplasmic tails and cytoskeletal adaptors (Carman and Springer, 2003; Cavallaro and
Christofori, 2004). Cytoskeletal anchorage strengthens the adhesiveness at the focal contact and affects the local cytoskeletal dynamics, which reciprocally facilitates the redistribution of cell surface receptors into clusters. For example, at the adherens junctions, cadherins form protein complexes with cytosolic protein catenin, which engages a strong cytoskeletal force targeting the adherent site at the cell surface
(Gumbiner, 1996; Yap et al., 1997). Endocytosis is another mechanism that effectively re-distributes CAMs and directs them to the competent zone of contact. Exocytic and endocytic trafficking of CAMs is essential in tempospatial regulation of their cell surface presentation in various adhesion processes (Ivaska et al., 2002; Kamiguchi, 2003; Bryant and Stow, 2004; D'Souza-Schorey, 2005). For instance, upon activation of NMDAR
(receptor for N-methyl-D aspartate) at the neural excitatory synapse, internalization of N-
9 cadherin is involved in regulating the synaptic strength in a β-catenin dependent manner
(Tai et al., 2007).
1.3. Intracellular signal transduction of cell-cell adhesion
Interactions mediated by CAMs contribute to physical adhesive force at the cell- cell contact, but also frequently induce intracellular signal transduction. Engagement of an extracellular ligand stimulates conformational changes in the cytoplasmic domain of the receptor. Such changes often result in exposure of a binding site for intracellular signal proteins, which become active and propagate the signal into the cytoplasm.
Integrins are the prototype CAMs that transduce intracellular signals in this manner. In the case that the CAM interacts with a receptor containing kinase domain, such as fibroblast growth factor receptors (FGFR), Eph receptor and muscle specific kinase
(MuSK), the extracellular interactions often induce the activation of the receptor’s kinase domain and subsequently the responsive cellular signal cascades. Intracellular signal transduction of cell-cell adhesion molecules ultimately results in the activation of gene transcription and alteration in cell behavior. Some of the CAM-induced signaling interactions and their physiological relevance are summarized in Table 1.1.
Moreover, cross-talk between individual CAM-induced signal pathways is essential for development. This can be exemplified by the interactive signals transduced respectively by cadherin- and integrin-mediated adhesions in a neurite outgrowth system.
The nonreceptor tyrosine kinase Fer can bind to both cytoplasmic tails of N-cadherin and
β1 integrin, coordinately regulating their signals in such a manner that the loss of an
10 Table 1.1. Functional interactions of CAMs with signaling molecules.
Signaling CAM Biological functions Reference molecule E-cadherin β-Catenin If β-catenin is present in a cadherin cell-adhesion complex, McCrea et al. 1991 increase of cell-cell adhesion; If β-catenin is not sequestered, activation of WNT signaling and Alman et al. 1997 tumor-cell invasion c-MET Disruption of intercellular adhesion Kamei et al. 1999 IGF1R Disruption of intercellular adhesion, increase metastasis; Lopez et al. 2002 Enhancement of intercellular adhesion and survival in cancer cells Mauro et al. 2001 EGFR Inhibition of ligand-dependent EGFR activation; Qian et al. 2004 Ligand-independent activation of EGFR Shen et al. 2004 Src Loss of epithelial differentiation; invasiveness Behrens et al. 1993 PI3K Activation of AKT; Bakin et al. 2000 Cell survival Bergin et al. 2000 N-cadherin FGFR1 Neurite outgrowth; Saffell et al. 1997 Cell migration and invasion; Suyama et al. 2002 Survival of ovarian epithelial cells Trolice et al. 1997 FGFR4 Neurite outgrowth; Cell-matrix adhesion Cavallaro et al. 2001 c-MET Epithelial-cell invasion Van Aken et al. 2003 Src Inactivation of N-cadherin-mediated adhesion Hamaguchi et al. 1993 PI3K Activation of AKT; Cell survival Rieger-Christ et al. 2004 Fer Crosstalk between N-cadherin and β1-integrin Arregui et al. 2000 PTP1B Regulation of N-cadherin-mediated adhesion Balsamo et al. 1998 NCAM FGFR1 Neurite outgrowth Saffell et al. 1997 FGFR4 Neurite outgrowth; Cell–matrix adhesion Cavallaro et al. 2001 p59 (Fyn) Activation of FAK in neurite outgrowth Beggs et al. 1997 PKC Neurite outgrowth Kolkova et al. 2000 GFRα1 Schwann-cell migration and axonal growth; Paratcha et al. 2003 Myelination of Schwann-cell Iwase et al. 2005 L1CAM FGFR1 Neurite outgrowth Saffell et al. 1997 Neuropilin Modulation of semaphorin 3A signaling, axon guidance Castellani et al. 2000; Castellani et al. 2002 Src Neurite outgrowth Atashi et al. 1992 Integrin ILK Cell proliferation Cruet-Hennequart et al. 2003 αvβ3 FAK Cell-matrix adhesion; Pfaff et al. 2001 Melanoma invasiveness Li et al. 2001 paxillin Cell-matrix adhesion; Pfaff et al. 2001 Cell migration Aznavoorian et al. 1996 Src Cell-matrix adhesion; Willey et al. 2003 Cancer cell survival and proliferation Huveneers et al. 2007 Rho Cell-matrix adhesion Butler et al. 2003 GPCR Cell-matrix adhesion Berg et al. 2007 PTK Cell-matrix adhesion Butler et al. 2005 E-selectin GPCR Leukocyte rolling on endothelial cells McMeekin et al. 2006 Rho Leukocyte rolling on endothelial cells Wojciak-Stothard et al. 1999 PTK Leukocyte rolling on endothelial cells Zarbock et al. 2007 c-MET: Mesenchymal epithelial transition factor; EGFR: EGF receptor; FAK: Focal-adhesion kinase; GFR: growth factor receptor; GPCR: G- protein coupled chemotactic receptor; IGF1R: insulin-like growth factor 1 receptor; ILK: integrin-linked kinase; PI3K: Phosphoinositide-3 kinase; PKC: protein kinase C; PTK: protein tyrosine kinase; PTP: Protein Tyrosine Phosphatase.
11 effector from the cytoplasmic domain of N-cadherin is accompanied by the gain of that effector by the β1-integrin complex (Arregui et al., 2000).
1.4. Cell adhesion molecules in embryogenesis
The impact of functional CAMs on embryogenesis becomes apparent quickly after the embryo is fertilized. For example, E-cadherin is expressed in the embryo at a very early stage, and contributes to the adherens junction formation and cell compaction at the morula stage (the embryo prior to the 32-cell stage). If an antibody against E- cadherin is added to the morula, integrity of the adherens junction is disrupted, and the embryo fails to undergo compaction and the subsequent formation of the blastocyst is aborted (Reima, 1990). As the embryo develops further, a variety of CAMs are expressed in a highly restricted manner and are critical in different processes, such as cell-sorting, polarity establishment and maintenance, and cell migration, during anterior-posterior axis formation, as well as neurulation and organogenesis at later stages (Thiery, 2003).
Garcia-Castro et al. (2000) showed that the cell adhesion molecule N-cadherin is one of the earliest proteins to be asymmetrically expressed in the chicken embryo and that its activity is required during gastrulation for proper establishment of the left-right axis.
Blocking N-cadherin function randomizes heart looping and alters the expression of Snail and Pitx2, later components of the molecular cascade that regulates left-right asymmetry
(Garcia-Castro et al., 2000).
Deficiency in CAMs essential in the key developmental processes will lead to delay or abnormality in development or even lethality at early embryonic stages. For example, N-cadherin-deficient mouse embryos die of several developmental defects. In
12 these mutant mice, the heart tube fails to develop normally and somites are small and
irregular (Radice et al., 1997). These phenotypic abnormalities are consistent with N-
cadherin playing a pivotal role in epithelialization of mesenchymal segmental plates that
precedes the formation of somite and myotome compartments (Linask et al., 1998).
At present, many knockout or mutant animals having deficient CAMs have been generated. Studies on these knockout and disease models have contributed tremendously to our knowledge of CAM functions during early embryonic development. Phenotypes of some of these mutants are summarized in Table 1.2.
1.5. Cell adhesion molecules in tissue survival and regeneration
In the adult, cells frequently change their adhesive properties in response to physiological or pathological processes. CAMs are evidently crucial to the maintenance and surveillance of mature tissue structure. In the injured spinal cord, neuronal CAMs are implicated in the responsive phase of rapid and substantial nerve fiber ingrowth into the lesion from both rostral and caudal spinal tissues. In the transectioned adult rat, L1 and
NCAM are found to be intensively expressed in the Schwann cells and the leptomenigeal cells proximal to the lesion, suggesting their involvement in axonal regeneration (Brook et al., 2000).
In the vascular system, both endothelial cells and neutrophils undergo changes in
CAM expression in response to inflammatory mediators, permitting rapid recruitment of phagocytes to the damaged tissue. The initial step in leukocyte recruitment involves endothelial cell activation and selectin expression on the endothelium. Simultaneously, leukocytes become activated, characterized by the activation of integrins. Activated
13 Table 1.2. Phenotypes of animal models deficient in CAMs.
CAM Animal model Deficiency resulted phenotypes Reference
N-cadherin Knockout mouse (Cdh2-/-) Die at E10; the myocytes subsequently Radice et al., 1997 dissociate and the heart tube fails to develop normally Cardiac-specific expression Rescued heart development; Luo et al., 2001 of Cdh2 in the Cdh2-/- Morphologically malformed neural tube; mouse increased apoptosis in neural and somitic tissues Zebrafish parachute (pac) Impaired neural tube development, Lele et al., 2002 mutant particularly in the midbrain and hindbrain; Zebrafish N-cad knockdown Perturbed migration of adaxial cells Cortes et al., 2003 (precursors of slow-twitch muscles) Drosophila null CadN Aberrant axonal trajectories in the CNS, Iwai et al., 1997 mutant including failure of position shifts, defective bundling, and errors in directional migration of growth cones Drosophila mosaic deletion Abnormal photoreceptor axon extension Prakash et al., 2005 of CadN in the retina by X- to the target ray irradiation
L-selectin Knockout mouse Defects in leukocyte behavior, including Mayadas et al., elevated numbers of circulating 1993 neutrophils, total absence of leukocyte rolling in mesenteric venules, and delayed recruitment of neutrophils to the peritoneal cavity upon inflammation
NCAM Knockout mouse Defects in presynaptic organization and Rafuse et al., 2000; function at neuromuscular junction, Polo-Parada et al., including synaptic vesicle dynamics, and 2001 transmitter release Reduced size of the olfactory bulb; Rolf et al., 2002 pathfinding errors of corticospinal axons; reduced exploratory behaviour and deficits in spatial learning Increased intermale aggression and Stork et al., 1999 neuroendocrine response
L1 L1-deficient mouse Reduced corticospinal tract; abnormal Dahme et al., 1997 disrupted in the Ig3 domain myelination; delayed motor response and weakness in the hind-limbs; Dilated brain ventricles, altered shape of Fransen et al., 1998; Sylvius aqueduct; Rolf et al., 2001 Vermis hypoplasia and impaired Fransen et al., 1998 exploration patterns; L1-deficient mice disrupted Reduced corticospinal tract Cohen et al., 1998 in the Ig6 domain Knockdown of L1.1 in adult Impaired locomotor recovery as well as Becker et al., 2004 zebrafish at the lesion after reduced regrowth and synapse formation spinal cord transection of axons.
14 integrins enable leukocyte rolling on the endothelium via their interactions with P-
selectin and E-selectin on the endothelial cell. Firm adhesion of leukocytes requires
interactions between leukocyte CD18, integrins, and endothelial cell ICAM-1. Finally,
leukocyte transmigration is mediated by endothelial cell PECAM (Kakkar and Lefer,
2004). Therefore, recruitment of leukocytes to the inflammation site involves complex
interactions mediated by multiple CAMs.
1.6. Cell adhesion molecules in diseases
CAMs constitute interaction complexes in the regulatory machinery that governs
individual physiological processes. Malfunction in any part of this complex inevitably
leads to disease or cancer. Changes in the expression or function of CAMs can therefore
contribute to pathological progress both by altering the adhesion status of the cell and by
affecting cell signaling (Cavallaro and Christofori, 2001; Christofori, 2003).
Recent experimental evidence indicates that such processes play a crucial role in
tumor progression, particularly during invasion and metastasis. In most, if not all, cancers
of epithelial origin, E-cadherin is functionally disrupted or its expression decreases
(Cavallaro and Christofori, 2004). The effect is likely to be direct because re-establishing
E-cadherin function in cadherin-deficient cell lines can reverse the invasive phenotype
(Vleminckx et al., 1991). This is further supported by an in vivo study on a mouse model
(Perl et al., 1998). Concomitantly, gain of mesenchymal cadherins (N-cadherin, R-
cadherin, and cadherin 11) in the converted epithelial cells potentiates cell migration and
invasiveness, possibly via a p120 catenin-dependent pathway (Yanagisawa and
Anastasiadis, 2006).
15 In addition to cadherins, integrins and some IgSF members (e.g. NCAM and L1)
are also shown to play an important role in the progression to tumor malignancy. For
example, L1 has been implicated in regulating metastatic dissemination in certain tumor
types, including breast cancer, prostate cancer and melanoma (Kenwrick and Doherty,
1998). In particular, due to its ability to interact with integrins in a heterotypic manner,
L1 has been proposed to favor the adhesion and transendothelial migration of melanoma cells, a crucial step in the metastatic dissemination of tumor cells (Voura et al., 2001).
16 2. IMMUNOGLOBULIN SUPERFAMILY IN NEURAL DEVELOPMENT
The IgSF represents one of the largest protein families in the human genome with
an estimated 765 genes (Brummendorf and Lemmon, 2001). They are widely distributed
in many tissue systems, particularly in the immune and nervous system. All members
consist of at least one Ig-like domain, which is 70–110 amino acids in length. Each Ig
domain usually contains two characteristic cysteine residues ~55 to 75 residues apart,
which provide stability by forming intradomain disulfide bonds (Walsh and Doherty,
1997). Although only 15–27% of the residues are identical, the Ig domain of this group of proteins shares 75–90% similarities in the structural conformation (Harpaz and Chothia,
1994). A second structural motif found in most Ig superfamily members is the fibronectin
type III (FnIII) repeat, which contains ~90 amino acids (Huber et al., 1994). Furthermore,
the presence of tyrosine kinase and phosphatase modules in the cytoplasmic tail
contributes to their diverse functions. Schematic presentation of the structures of some
IgSF members is illustrated in Figure 1.2.
Members of the IgSF usually mediate cell-cell adhesion in a Ca2+-independent
manner, which may involve both homophilic and heterophilic interactions. Although
binding frequently occurs between IgSF members, several of them are capable of
interacting with molecules from other families during the processes of axonal growth, guidance, and synapse formation (Brummendorf and Rathjen, 1996; Walsh and Doherty,
1997). The following section will highlight two IgSF members, L1 and muscle specific
kinase (MuSK), which are the focus of my thesis research.
17 Domain legend: Ig-like S-S S-S FnIII W Cysteine rich S-S GPI
S-S Tyrosine kinase S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S S-S WW S-S S-S S-S S-S S-S S-S S-S
MEMBRANE MEMBRANE
NCAM L1 Contactin 1 MAG P0 FGFR MuSK CHL1 Contactin 2 NrCAM
Figure 1.2. The structure of some members of the Ig superfamily implicated in growth and guidance in the nervous system. Shown here are examples of Ig CAMs and receptor tyrosine kinases (RTKs). All molecules are typified by having Ig domains of various types; in the brain they are of C2, V, or I sets. Members of the CAM subgroups all contain Ig domains, and with the exception of MAG, contain FnIII repeats. NCAM and L1 are transmembrane proteins, whereas contactins are GPI
anchored. P0 is one of the smallest Ig member, consisting only one Ig domain extracellularly. Two representative RTKs, FGFR and MuSK, are illustrated. FGFR contains three Ig domains. MuSK contains three Ig domains and a cysteine-rich domain. Both proteins possess an intracellular tyrosine kinase domain.
18 2.1. L1 family
L1 is one of the most studied neural cell adhesion molecules, and it is involved in a variety of adhesion processes integral to neural development, including neuronal migration (Lindner et al., 1983), axon growth (Lemmon et al., 1989), axonal
fasciculation (Honig et al., 1998) and myelination (Itoh et al., 2005), and neuron proliferation (Dihne et al., 2003). The L1 gene family have also been found to play critical roles in neuronal survival and axon regeneration after trauma (Maness and
Schachner, 2007). L1 is a prototypic member of this family. Vertebrate species appear to harbor several different L1-type genes in their genomes, whereas several arthropod genomes contain only one L1-type gene. All vertebrate L1 genes can be assigned to one of four phylogenetic L1 gene subfamilies, referred to as L1, neurofascin, CHL1 (close homolog of L1), and Nr-CAM (Hortsch, 2000). It is possible that two sequential duplications gave rise to four homologous sets of genes in amniotes from a single
ancestral L1 gene of the type found in arthropod genomes. These genome duplications
have resulted in a diversification of gene expression and protein structures (Pebusque et
al., 1998). In zebrafish, two L1 homologs are found, namely L1.1 and L1.2. Shown in
Figure 1.3 is the phylogenetic analysis on currently known L1 gene family proteins from
both vertebrates and invertebrates.
L1-related molecules contain an extracellular region of six Ig-like domains and
four to five FnIII repeats, followed by a highly conserved cytoplasmic domain of ~110
residues. The multidomain structure of L1 molecules suggests that L1 may undergo
complex interactions with diverse receptors that modulate cell adhesiveness and
responsiveness necessary for the attractive and repellent cues.
19 Neurofascins
Rat Mouse neurofascin neurofascin
Human neurofascin NrCAMs Chick neurofascin Mouse NrCAM CHL1 Mouse Rat CHL1 NrCAM Human Invertebrate CHL1 L1-like molecules Human Hemolin NrCAM
Chick Drosophlia neuroglian NrCAM C. elegans Lad-2
Rat F3/contactin 1
Rat TAG-1/contactin 2
Mouse L1 Chicken axonin-1/contactin 2
Rat L1 Human Chick L1 NgCAM Goldfish L1.1 Fugu L1 Zebrafish L1.2 Zebrafish L1.1 L1CAMs
Figure 1.3. Phylogenetic analysis of the L1 gene family. Full length protein sequences of L1 gene family members were directly downloaded from GenBank and aligned using the multiple alignment option of the Clustal X program package (www-igbmc.u-strasbg.fr/BioInfo/clustalx/). The phylogenetic tree was constructed from these aligned sequences by using the method Neighbor Joining (NJ) from the program package.
20 Electron micrographs of rotary-shadowed L1 reveal that the extracellular portion
of L1 has a compact structure with a horseshoe fold involving the first four Ig-domains,
which is flexible and can open into an elongated shape (Schurmann et al., 2001). It is
proposed that, similar to axonin-1 and hemolin, the sharp bend between the second Ig
domain (Ig2) and the third Ig domain (Ig3) creates a U-shaped module in which the Ig1
associates tightly with the Ig4 and the Ig2 associates with the Ig3 (Su et al., 1998;
Freigang et al., 2000; Hall et al., 2000). The conformational changes of L1 from a folded
to extended structure, may reflect changes in the adhesion potency of L1 (Drescher et al.,
1996; Schurmann et al., 2001). However, the mechanism that drives this conformational
change and influences L1 binding capacity remains unclear.
Q The cytoplasmic domain of L1 contains a conserved sequence (FIG /AY) that
reversibly binds ankyrin, a spectrin adaptor which couples L1 to the subcortical actin
cytoskeleton (Kizhatil et al., 2002; Whittard et al., 2006). Alternative splicing of L1 generates a neuronal isoform that contains a sequence Arg-Ser-Leu-Glu (RSLE), which enables AP2/clathrin-mediated endocytosis of L1 (Kamiguchi et al., 1998c).
Internalization of L1 molecules is regulated by the phosphorylation of the tyrosine
residue immediately preceding this RSLE sequence, which abrogates the binding of AP2
to the L1 tail. Interestingly, this sequence is also implicated in the interaction between L1
and ezrin, a member of the ezrin, radixin, and moesin (ERM) family of membrane-
cytoskeleton linking proteins (Dickson et al., 2002). Besides the YRSLE sequence,
another stretch of amino acids (K/RxxK) at the juxtamembrane region has been shown to
independently link L1 to ERM proteins (Cheng et al., 2005). The presence of these ERM-
binding motifs potentiates the association of L1 family members with actin, underscoring
21 their capability of modulating cytoskeleton dynamics. The involvement of these motifs in regulating the functions of L1 will be discussed in greater details in a subsequent section on “L1-associated Interaction”. Moreover, several serine resides are conserved in the cytoplasmic domain of L1, and their phosphorylation may be involved in regulation of
L1-mediated intracellular responses (Kenwrick et al., 2000). The characteristics of the cytoplasmic domain of L1 are shown schematically in Figure 1.4 and a comparison of these conserved features among different L1 molecules is summarized in Table 1.3.
2.2. Muscle specific kinase (MuSK)
MuSK is expressed in skeletal muscle, where it is concentrated in the postsynaptic membrane of neuromuscular junctions (NMJs). MuSK is a critical component of agrin-
induced postsynaptic assembly (Burden, 2002). The extracellular region of mammalian
MuSK contains three Ig-like domains and a cysteine-rich domain. An extracellular
kringle domain is also found in avians, fish, and amphibians. In the intracellular region,
MuSK contains a kinase domain which enables autophosphorylation (Jennings et al.,
1993; Valenzuela et al., 1995; Cheung and Smith, 2000).
During the formation of the neuromuscular synapse, MuSK is crucial for
orchestrating postsynaptic differentiation. Upon innervation, MuSK is activated by the
engagement of neural agrin and forms a primary synaptic scaffold to which the synapse-
specific cytoplasmic protein rapsyn recruits acetylcholine receptor (AChR). Numerous
clusters of AChR constitute the postsynaptic apparatus and stabilize the neuromuscular synapse structure (Burden, 2002). MuSK can also mediate synapse assembly in the absence of agrin. In mice that lack agrin expression (agrin-/-), MuSK assembles AChR
22 MEMBRANE MEMBRANE
K G ERM binding G K
1176 → Y c-src R S AP-2 binding L ERM binding E 1181 → S Casein kinase II
F Ankyrin binding I G Doublecortin binding Q 1229 → Y ERK 2
COOH
Figure 1.4. Conserved features of the cytoplasmic tail of L1 family members. Amino acid numbering is based on the human L1 protein. Characteristic binding sites for ERM proteins, AP-2 adaptor, ankyrin, and doublecortin are highlighted in pink. Phosphorylation sites at tyrosine and serine residues are indicated on the left. The YRSL motif of L1 is required for correct targeting to the growth cone. It interacts with the AP-2 adapter complex facilitating clathrin-mediated endocytosis which rapidly modulates L1 distribution in response to environmental changes. This motif is partly encoded by an alternatively-spliced exon 27 (RSLE). The rapid formation and dissociation of stable adhesion complexes of L1 with itself and other heterophilic ligands probably require specific interactions with two actin adaptors, ERM and ankyrin. Also, L1 contains binding consensus for doublecortin that links to microtubule. Anchorage of L1 to the cytoskeleton is regulated by phosphorylation of Ser and Tyr residues in the cytoplasmic domain. The kinases implicated for the corresponding residues are indicated on the right.
23
Table 1.3. Conserved characteristics in the cytoplasmic tail of L1 superfamilies.
(Adapted from Brümmendorf et al., 1998)
Species Site of binding to actin RSLE Site of phosphorylation Site of binding to cytoskeleton proximal to (encoded by exon 27) by casein kinase II ankyrin the plasma membrane
Vertebrates hu, rt, mo, ch, zf, L1 KxxK Yes S FIGQY ca 24 L1.2 zf, fg KxxK Yes S - CHL1 hu, mo RxxK No S FIGAY neurofascin mo, rt, ch RxxK Yes S FIGQY NrCAM hu, mo, ch KxxK Yes S FIGQY
Invertebrates
neuroglian dr RxxK No S FIGQY Lad-2 ce Rxxx No ? FIGQY
Species abbreviations: ca, goldfish (Carassius auratus); ce, Caenorhabditis elegans; ch, chick; dr, Drosophila melanogaster; fg, pufferfish (Fugu rubripes); hu, human; mo, mouse; rt, rat; zf, zebrafish (Danio rerio) clusters on the central region of muscle early in development by an unidentified mechanism (Lin et al., 2001). A recent study suggests that Dok-7, a MuSK-interacting
cytoplasmic protein, may participate in this agrin-independent synapse formation (Okada
et al., 2006). In zebrafish, a similar pre-patterned postsynapse on the somite along the
axonal path is observed before the axonal growth cone approaches (Panzer et al., 2006).
By analyzing the early stages of postsynaptic differentiation in muscles of mutant mice lacking agrin, MuSK, rapsyn and/or motor nerves, Lin et al. (2001) have proposed three
overlapping steps of the postsynapse formation at the neuromuscular junction. First, a
muscle-intrinsic, nerve/agrin-independent and MuSK-dependent mechanism initiates the
formation of postsynaptic specialization in an end-plate band. Second, nerve-derived
agrin acts through MuSK to promote apposition of nerve terminals to these nerve-
independent acetylcholine receptor clusters and to induce propagation of postsynaptic
assemblies. Finally, aneural AChR clusters are dispersed by the nerve through an agrin-
independent mechanism (Lin et al., 2001).
Mutations in the human MuSK gene are responsible for autosomal myasthenic
syndrome, a human neuromuscular transmission dysfunction (Chevessier et al., 2004). In
zebrafish, mutations in the MuSK homolog result in the unplugged phenotypes, which
manifest axonal guidance defects specific for motor neurons (Zhang et al., 2004). In
zebrafish, three pioneering motor neurons, caudal primary (CaP), middle primary (MiP)
and rostral primary (RoP), innervate each somatic hemisegment. The pathway selection
of CaP and RoP is affected in unplugged mutants. Although their growth cones navigate
correctly from the spinal cord to the somitic choice point, they fail to select their
25 subsequent cell-type-specific path. Pathway selection of MiP neurons appears unaffected
in these mutant embryos (Zhang and Granato, 2000).
2.3. Other neural IgSF members
Other IgSF members are known to play key roles in supporting neuronal survival, synapse formation and axonal guidance, including NCAM, contactins, P0, myelin-
associated glycoprotein (MAG), and Fibroblast growth factor receptors (FGFRs).
Discovered as the first neural cell adhesion molecule, NCAM was shown to be
capable of mediating adhesion of cells in the retina (Hoffman et al., 1982). Vertebrate
NCAM exists in several isoforms which are selectively expressed by the neuron and glia
and at different stages of development (Persohn et al., 1989; Doherty and Walsh, 1991;
Polo-Parada et al., 2004; Polo-Parada et al., 2005). In the mouse, three major NCAM
proteins, sharing identical extracellular domains (shown in Figure 1.2), are encoded by
alternatively spliced transcripts with distinct molecular weights of 180 kDa, 140 kDa and
120 kDa (Barthels et al., 1987). In mice lacking all forms of NCAM, subtle defects were
observed in the nervous system. These mice exhibit reduced mossy fiber density in the
hippocampus and aberrant fasciculation and pathfinding of these axons, thus supporting
the role of NCAM in synaptic plasticity (Cremer et al., 1997). An important feature of
NCAM that makes it a unique member of IgSF is the presence of α-2,8-linked polysialic
acid (PSA) in domain Ig5 (Kleene and Schachner, 2004), which modifies functional
properties of the NCAM protein backbone during neural migration, axon pathfinding and
synaptic plasticity (Eckhardt et al., 2000). PSA modulates NCAM functions probably by
affecting its homophilic interaction (Maness and Schachner, 2007).
26 Contactins are a subgroup of neural IgSF CAMs that are attached to the cell
membrane by a glycosylphosphatidylinositol anchor (Yoshihara et al., 1995; Falk et al.,
2002). Contactin 1 (also known as F3 in the mouse and F11 in the chick) and contactin 2
(known as TAG-1 in the rat and axonin-1 in the chick) are both implicated in neurite
outgrowth and axonal fasciculation (Furley et al., 1990; Falk et al., 2002). Due to the
absence of a cytoplasmic domain, these proteins rely on heterophilic interactions to
transduce signals required for axonal growth and guidance. In a given neurological
process, contactins are shown to bind specifically to the L1 subgroup of IgSF, including
L1/NgCAM, NrCAM, and neurofascin, and these interactions therefore define their
explicit function (Falk et al., 2002; Maness and Schachner, 2007). During neurite
fasciculation of chick DRG neurons, the heterophilic interaction of axonin-1/contactin 2
with NgCAM occurs only when both molecules are located in the same membrane (cis
binding). This cis-association of axonin-1 and NgCAM is able to assemble independent
of cell-cell contact (Buchstaller et al., 1996). When cells are apposing each other and
cell-cell contact is made, two axonin-1:NgCAM heterodimers on adjacent cell members
are induced to form hetero-tetrameric complexes which consequently trigger
oligomerization as a zipper-like array owing to the homophilic binding capacity of both
axonin-1 and NgCAM (Kunz et al., 1996; Kunz et al., 1998; Freigang et al., 2000).
Protein zero (P0) is one of the simplest IgSF proteins and contains only a single
IgSF domain in the extracellular region, followed by a single membrane-spanning helix
and an intracellular domain (Lemke and Axel, 1985). P0 undergoes homophilic adhesion which is implicated in the compaction of myelin, by holding together individual wraps of the myelin sheath (D'Urso et al., 1990). Mutations in the P0 gene are responsible for two
27 congenital peripheral neuropathies, namely Charcot-Marie-Tooth disease (CMT) type 1B and Dejerine Sottas (DS) syndrome (Warner et al., 1996).
The MAG is expressed in Schwann cell and oligodendroglial membranes of
myelin sheaths, consisting of two Ig domains that show potent binding to sialic acid-
containing oligosaccharides (Tropak and Roder, 1997; Quarles, 2007). MAG has a
demonstrated function in maintaining glia-axon interactions in both the peripheral and
central nervous system (CNS) by directly binding to a sialyloligosaccharide on a
glycoprotein or ganglioside present on the axonal membrane (Schachner and Bartsch,
2000; Quarles, 2007). Another important characteristic of MAG contributing to its
function is the presence of carbohydrate moieties, one of which is the HNK-1 epitope
(McGarry et al., 1983).
FGFRs are tyrosine kinase receptors that contain Ig-like domains (Coutts and
Gallagher, 1995). FGFRs possess high binding affinity to an array of FGFs and are able
to induce various cytoplasmic signal transduction pathways via autophosphorylation of
the intercellular tyrosine kinase domain. In the nervous system, FGFR1 is involved in early neural induction and patterning, including establishment of anterior-posterior axis, organization and regionalization of the developing brain (Mason, 2007). In addition,
FGFR1 has been implicated in neurite outgrowth, via the interactions with NCAM, N-
cadherin and L1 (Saffell et al., 1997).
28 3. CELL ADHESION MOLECULE L1 AND CRASH SYNDROME
L1 was first described as a 200 kDa cell surface glycoprotein present on
cerebellar granule cells and N2A neuroblastoma cells. In addition to this 200 kDa species,
immunopurification using an anti-L1 antibody reveal molecular weight components of
140 kDa and 80 kDa (Rathjen and Schachner, 1984). These fragments have also been detected in the human, rat and chick brains (Wolff et al., 1988; Liljelund et al., 1994;
Burgoon et al., 1995). The lower molecular weight fragments are likely the proteolytic
products of the 200 kDa full length molecule (Faissner et al., 1985).
During CNS development, L1 is expressed on the surface of long axons and
growth cones. It continues to be expressed in the adult on unmyelinated axons. In the
advancing neuronal growth cone, L1 protein is localized primarily to the leading edge
(Kamiguchi and Lemmon, 2000). L1 is also expressed on some migrating neurons, such
as the neurons originated from the forebrain subependymal zone (Barami et al., 1994;
Kamiguchi et al., 1998b). My thesis research is focused on the investigation of L1
functions during early neural development, with an attempt to elucidate the pathogenic
mechanisms of mutations that lead to phenotypic abnormalities in patients. The following
sections will review our current knowledge of L1 and CRASH/L1 syndrome that
associates with the L1 mutations.
3.1. L1CAM gene
In humans, the L1 gene has 28 exons (Bateman et al., 1996). Two mini exons, 2
and 27, can be alternatively spliced, resulting in an isoform having both exons and an
isoform lacking these exons (Reid and Hemperly, 1992; Jouet and Kenwrick, 1995). The
29 isoform that lacks exons 2 and 27 seems to be restricted to non-neural cells, as well as
premature oligodendrocytes, although its functional significance remains to be elucidated
(Takeda et al., 1996; Itoh et al., 2000). The isoform that contains both exons 2 and 27 is exclusive to the neuronal cells. Alternative splicing of exon 2 has been shown to influence L1 homophilic interaction, as the L1 lacking this exon 2-encoded sequence binds to each other poorly (Jacob et al., 2002). The alternatively spliced exon 27 encodes the Arg-Ser-Leu-Glu (RSLE) sequence in the cytoplasmic domain. RSLE constitutes part of a highly conserved stretch of motif, YRSLE, which can be phosphorylated at the tyrosine residue (Hortsch, 1996). As discussed earlier, phosphorylation of this Tyr regulates L1 association with the cytoskeleton and its internalization.
In humans, the mature L1 protein has 1256 amino acids. Its extracellular part consists of six Ig-like domains and five fibronectin type III-like domains, which is followed by a single-pass transmembrane domain and a short cytoplasmic tail (Moos et al., 1988). L1 homologues have been discovered in rat (also known as NILE), mouse, chicken (also known as NgCAM), zebrafish (L1.1), Fugu and goldfish (see the phylogenetic tree in Figure 1.3).
3.2. Clinical presentation of CRASH syndrome
Mutations in the L1 gene are responsible for an X-linked recessive neurological disorder that has been described as Hydrocephalus due to stenosis of the aqueduct of
Sylvius (HSAS, Bickers and Adams, 1949), X-linked hydrocephalus (XLH; Edwards et al., 1961), or spastic paraplegia type I (SPG-1; Kenwrick et al., 1986). Linkage analysis on X-linked hydrocephalus patients mapped the disease locus to Xq28 (Willems et al.,
30 1990). Subsequent analyses have revealed that these three syndromes result from
mutations in the L1 gene (Jouet et al., 1994). Various names have been given to this family of diseases including “L1 syndrome” (Jouet and Kenwrick, 1995) and “CRASH syndrome” (Fransen et al., 1995). CRASH syndrome has been adopted widely and it summarizes the major features associated with L1 mutations, namely corpus callosum hypoplasia, mental retardation, adducted thumbs, spastic paraplegia, and hydrocephalus.
The varying nomenclature is a reflection of extremely variable presentation both within
and between families (Serville et al., 1992). X-linked hydrocephalus has a prevalence of
approximately 1 in every 30,000 males and is the most common genetic form of
congenital hydrocephalus (Jouet et al., 1994).
3.2.1. Hydrocephalus
Hydrocephalus (water on the brain) presents as abnormal accumulation of
cerebrospinal fluid (CSF) in the third and lateral ventricles. The extent of ventricular
enlargement is highly variable with the most extreme cases presenting elevated
intracranial pressure and huge macrocephaly (Schrander-Stumpel et al., 1990; Fransen et
al., 1995). The excess of CSF and increased size of cerebral ventricles may partly result
from stenosis of a thin channel called the aqueduct of Sylvius through which CSF
circulates (Bickers and Adams, 1949). Those that develop hydrocephalus in utero or soon
after birth have a lower life expectancy and many die neonatally (Fransen et al., 1997).
However, stenosis of the aqueduct of Sylvius is not necessarily a constant feature of L1
syndrome. Several patients manifesting hydrocephalus are not diagnosed with stenosis of
the aqueduct of Sylvius (Yamasaki et al., 1995).
31 Besides aqueduct stenosis, inadequate cell migration or loss of neurons may also contribute to the increase in cavity size (Yamasaki et al., 1995). It is evident that, in some
L1 patients, a huge number of cells fail to migrate properly from the ventricular proliferation zone of the hydrocephalus brain (Kamiguchi et al., 1998a). Other frequently associated malformations include underdevelopment of the anterior vermis of the cerebellum and fused thalami, which imply abnormal neuronal proliferation, migration or survival (Yamasaki et al., 1995).
3.2.2. Agenesis of the corpus callosum
Perhaps the most striking pathological observation in L1 patients is hypoplasia or absence of two long axonal tracts, the corticospinal tract (CST) and the corpus callosum
(Chow et al., 1985; Yamasaki et al., 1995; Graf et al., 2000). The corpus callosum is the large bundle of nerve fibers that connects the two cerebral hemispheres, and its underdevelopment may contribute to the spastic paraplegia and mental retardation observed in patients with abnormal L1.
3.2.3. Spastic paraplegia
Spasticity of the lower limbs and hyperreflexia is a consistent feature in all cases of CRASH syndrome (Fransen et al., 1997). The cause of this has been correlated to the abnormal development of the CST. Under physiological conditions, the nerve fibers of the CST project from the cortex to the spinal cord, and its primary function is in the control of voluntary movement. Neuropathological examination of the medulla in
32 patients with X-linked hydrocephalus reveals the absence or hypoplasia of the pyramidal formation (Yamasaki et al., 1995).
3.2.4. Mental retardation
Mental retardation is present in nearly all individuals with L1 mutations
(Yamasaki et al., 1997). While the degree of deficits may vary, mental retardation is prominent in patients exhibiting severe hydrocephalus (Willems et al., 1987; Jouet et al.,
1994; Vits et al., 1994). More than 20% of patients with modest and no hydrocephalus also have grave mental retardation. The conditions of mental retardation cannot be improved by ventriculo-peritoneal shunting to remove CSF and relieve the ventricular pressure. Therefore, mental retardation associated with L1 mutations is likely due to abnormal development of the central neural system rather than a secondary consequence of hydrocephalus (Yamasaki et al., 1997).
3.2.5. Adducted thumbs
The phenotype adducted thumbs, which presents as flexion deformity of the thumbs, is a specific finding commonly associated with CRASH syndrome (Schrander-
Stumpel et al., 1995). The adduction appears to result from loss of innervation of the extensor pollicis longus (EPL) and lack of input from the appropriate spinal motor neuron pool (Holtzman et al., 1976). Nearly all patients examined showed adducted thumbs
(Yamasaki et al., 1997). Interestingly, presence of adducted thumbs is diagnosed in 94% of the patients having missense mutations in the cytoplasmic domain, suggesting that adducted thumbs might be caused by dysfunction of the cytoplasmic domain in either its
33 signaling function or an essential interaction with the cytoskeleton (Yamasaki et al.,
1997).
3.3. Disease diagnosis
Molecular diagnosis of L1 gene mutations provides definitive prenatal diagnosis
of CRASH syndrome at 10 weeks of gestation (Jouet and Kenwrick, 1995). Typical
analysis is performed on cDNAs derived from B cell samples by single-strand
conformation polymorphism (SSCP) for gene mutations followed by direct sequencing
(Jouet et al., 1994). An adapted approach applying fluorescent-assisted mismatch
analysis allows rapid detection of mutations and identification of intronic variations
(Saugier-Veber et al., 1998). On the other hand, clinical prenatal diagnosis of
hydrocephalus may be restricted to ultrasound, which is generally achieved after 16
weeks of gestation (Jouet and Kenwrick, 1995).
Over the past two decades, 146 mutations of the L1 gene have been identified:
36% missense mutations (Table 1.4), 13% nonsense mutations, 24% deletions, 3%
insertions and 24% splice site changes
(http://www.rug.nl/umcg/faculteit/disciplinegroepen/MedischeGenetica/HereditaryDiseas
es/L1cam/).
3.4. Genotype-phenotype correlations
L1 mutations give rise to a highly variable spectrum of disease states. Bateman et
al. (1996) proposed that (1) nonsense mutations in the extracellular part of L1 leading to
truncation or absence of L1 are more likely to cause a severe phenotype, (2) mutations in
34 Table 1.4. Missense mutations in L1 gene associated with CRASH syndrome.
Nucleotide change Exon Amino acid change Protein domain Reference
G26 → C 1 W9S SP Jouet et al., 1993; Jouet et al., 1995a C88 → A 2 H30N SP Finckh et al., 2000 C358 → G 4 L120V Ig1 Bateman et al., 1996 G361 → A 4 G121S Ig1 Jouet et al., 1995a T536 → G 6 I179S Ig2 Ruiz et al., 1995 C550 → T 6 R184W Ig2 Fransen et al., 1996 C550 → T 6 R184W Ig2 Finckh et al., 2000 C550 → T 6 R184W Ig2 Graf et al., 2000 G551 → A 6 R184Q Ig2 Jouet et al., 1994 G551 → A 6 R184Q Ig2 MacFarlane et al., 1997 A581 → G 6 Y194C Ig2 Gu et al., 1996 G604 → T 6 D202Y Ig2 Sztriha et al., 2000 C630 → G 6 H210Q Ig2 Jouet et al., 1994; Vits et al., 1994 T656 → C 6 I219T Ig2 Saugier-Veber et al., 1998 C719 → T 7 P240L Ig3 Gu et al., 1996 G791 → A 7 C264Y Ig3 Jouet et al., 1993 G803 → A 7 G268D Ig3 Fransen et al., 1997 G925 → A 8 E309K Ig3 Jouet et al., 1995a G925 → A 8 E309K Ig3 Straussberg et al., 1991; Fransen et al., 1997 C998 → G 9 P333R Ig3 Van Camp et al., 1996 T1003 → C 9 W335R Ig4 Saugier-Veber et al., 1998 G1005 → T 9 W335C Ig4 Finckh et al., 2000 G1108 → A 9 G370R Ig4 Kaepernick et al., 1994; Ruiz et al., 1995 G1108 → A 9 G370R Ig4 Finckh et al., 2000 C1156 → T 10 R386C Ig4 Saugier-Veber et al., 1998 T1172 → C 10 L391P Ig4 Fransen et al., 1996 A1223 → T 10 N408I Ig4 Finckh et al., 2000 G1243 → C 10 A415P Ig4 Sztriha et al., 2002 T1262 → A 10 V421D Ig4 Finckh et al., 2000 C1277 → A 11 A426D Ig5 Fransen et al., 1997 G1354 → A 11 G452R Ig5 Jouet et al., 1994 C1417 → T 12 R473C Ig5 Saugier-Veber et al., 1998 T1445 → C 12 L482P Ig5 Gu et al., 1997 G1490 → A 12 C497Y Ig5 Finckh et al., 2000 T1624 → C 13 S542P Ig6 Gu et al., 1997 G1792 → A 14 D598N Ig6 Vits et al., 1994 G1895 → C 15 R632P Fn1 Fransen et al., 1997; Vits et al., 1998 A1963 → G 16 K655E Fn1 Izumoto et al., 1996 C2021→G 16 S674C Fn1 Michaelis et al., 1998 G2071 → A 16 A691T Fn1 Finckh et al., 2000 C2072 → A 16 A691D Fn1 Du et al., 1998 G2092 → A 16 G698R Fn1 Du et al., 1998 C2215 → T 18 R739W Fn2 Finckh et al., 2000 T2222 → C 18 M741T Fn2 Gu et al., 1997 G2252 → C 18 R751P Fn2 Finckh et al., 2000 G2254 → A 18 V752M Fn2 Gu et al., 1997 G2302 → A 18 V768I Fn2 Gu et al., 1997 G2302 → T 18 V768F Fn2 Jouet et al., 1995b G2308 → A 18 D770n Fn2 Simonati et al., 2006 A2351 → G 18 Y784C Fn2 MacFarlane et al., 1997 T2804 → C 21 L935P Fn4 Du et al., 1998 C2822 → T 21 P941L Fn4 Jouet et al., 1995b A3209 → G 24 Y1070C Fn5 Jouet et al., 1993; Jouet et al., 1995b C3581 → T 28 S1194L Cyto Fransen et al., 1994 C3671 → T 28 S1224L Cyto Saugier-Veber et al., 1998 T3685 → C 28 Y1229H Cyto Fransen et al., 1997 G3716 → A 28 G1239E Cyto Finckh et al., 2000
(Adapted from http://www.rug.nl/umcg/faculteit/disciplinegroepen/medischegenetica/hereditarydiseases/l1cam/index.)
35 the cytoplasmic domain of L1 generally generate a milder phenotype than extracellular
mutations, and (3) extracellular missense mutations affecting amino acids situated on the
surface of a domain cause a milder phenotype than those affecting amino acids buried in
the core of the domain. Therefore, these mutations have been categorized into three
classes according to the structure of the expressed protein. Their correlations to the
severity of CRASH syndrome are shown in Table 1.5. Class 1 mutations disrupt only the
cytoplasmic domain. They are expected to disrupt L1-mediated signaling and
cytoskeleton interactions but not necessarily L1-mediated adhesion. The missense
mutations in the extracellular domains of L1 constitute Class 2 mutations. They would
represent a subtle to modest form of disruption. A point mutation might disrupt one form of L1 adhesion but preserve others. Class 3 mutations result in a stop codon that causes a truncation in the extracellular domain. This would lead to a secreted L1 molecule that does not function in either adhesion or signaling (Yamasaki et al., 1997). While accepting these generalizations about genotype-phenotype correlations, it is noteworthy that clinical findings in L1 can range from mild to severe even in the same family, indicating that other factors must influence the clinical presentation (Finckh et al., 2000).
Studies on missense mutations of L1 have suggested specific L1 functions, particularly, L1 interactions involving the affected amino acid residues. In order to investigate some of the unknown functions of L1, several missense mutations of L1 have been constructed for study and they are summarized in Table 1.6.
36
Table 1.5. Genotype and phenotype correlation of L1 mutations with severity of CRASH syndrome.
Modified after Yamasaki et al., (1997); Fransen et al., (1998b)
Class I Class II Class III
Genotype Mutations in the Missense mutations in Mutations resulting in cytoplasmic domain the extracellular domains extracellular truncation 37 Putative L1 protein function Extracellular functions Cell adhesion and Total loss of protein and adhesion conserved functions partially function conserved
Clinical phenotype Hydrocephalus absent to moderate moderate to severe mostly severe
Survival to 1st year all survive 30% die before 1 year 50% die before 1 year
Mental retardation ~25% of patients ~25% of patients 100% of patients (IQ score < 50)
Adducted thumbs ~95% ~85% ~100%
Table 1.6. Phenotype and cell biology of selected L1 mutations.
Mutation Domain Phenotypes in human Cell surface expression in comparison with wild type (WT) location Predicted Deaths<1 year Hydrocephalus Other phenotypes protein in CHO cells effect a (cases) (cases) L120V Ig1 surface 0/1 1/1 SP, AT, MR, ACT WT (De Angelis et al., 2002) R184Q Ig2 structural 10/15 15/15 AT, SP, MR, ACT Retarded in ER (De Angelis et al., 2002; Michelson et al., 2002) Y194C Ig2 surface 2/3 3/3 SP, ACC, AT WT (Michelson et al., 2002) D202Y Ig2 structure 0/1 1/1 SP, MR, AT, ACC, ACT ―
38 G268D Ig3 structure ― ― ― ― E309K Ig3 surface 0/2 1/2 SP, MR, AT Reduced (Cheng and Lemmon, 2004) N408I Ig5 structure ― 2/2 ― ― D544N Ig6 surface ― ? SP, MR, AT Retarded in ER (De Angelis et al., 2002) D598N Ig6 surface 0/3 0/3 SP, AT, MR, Reduced (De Angelis et al., 2002)
SP, spastic paraplegia; MR, mental retardation; AT, adducted thumbs; ACC, agenesis of corpus callosum; ACT, agenesis of corticospinal tract. a The effects of equivalent mutations are predicted by biophysical characterization of mutations on well-characterized model protein systems (Randles et al., 2006) 4. L1-ASSOCIATED INTERACTIONS
Pathogenic L1 mutations associated with CRASH syndrome are distributed along the whole molecule, implying that different L1 interactions are involved in various processes integral to neural development. L1 fulfills these roles via its multiple domains by interacting extracellularly with a variety of cell surface receptors and ECM molecules, conferring neuronal adhesion and responsiveness for neural guidance cues, as well as by coupling to cytoskeletal elements and signal cascades.
It has been shown that L1 can homophilically interact with another L1 in both cis and trans manner. Homophilic L1-L1 interaction has drawn a great deal of attention because it is believed to be the most common mode of action in promoting axon growth along a bundle of preexisting axons (Stallcup and Beasley, 1985; Landmesser et al., 1988;
Lemmon et al., 1989). Furthermore, L1 has been shown to heterophilically interact with several other cell surface molecules, including members of the IgSF, integrins, proteoglycans, and tyrosine kinase receptors (Maness and Schachner, 2007). The capacity for interacting with this large variety of potential partners, in both trans- and cis-binding manners, enables L1 to perform diverse functions at different times and locations during development. Several extracellular interactions involving L1 are summarized in Figure
1.5. In addition to adhesive interactions via the extracellular domains, L1 is able to mediate intracellular responses by modulating cytoskeleton dynamics, signaling and endocytic pathways via several conserved sites within the cytoplasmic tail, which are illustrated in Figure 1.4.
39 Homophilic Heterophilic
L1 Nrp-1 Integrin Neurocan TAG-1/ F11/ axonin-1/ Contactin 1
FASNKL Contactin 2 1
2
3
4
5 RGD 6
I II III IV V MEMBRANE
Figure 1.5. Schematic diagram of domain involvement to specific L1 extracellular interactions. Homophilic interaction of L1 involves Ig2 and the third FnIII domains. L1 also interacts heterophilically with several cell surface molecules including: Nrp-1 (through a sequence flanking FASNKL in Ig1 of L1), integrins (via the RGD motif in Ig6), neurocans (via Ig1), and contactins (requiring the integrity of a number of domains). The red bar represents the center of the correspondent interaction, while the grey bar represents regions that have also been implicated in the interaction.
40 4.1. Characteristics and functions of homophilic interaction
The earliest evidence of L1 homophilic interaction was provided from self-
aggregation of NgCAM-conjugated Covaspheres which are specifically disrupted by anti-
L1 Fab (Grumet and Edelman, 1988). Further experiments have demonstrated that L1-L1
interaction is able to mediate neurite outgrowth. As a substrate, L1 is potent at inducing
neurite extension from neurons obtained from chick and mouse, and its potency is
abrogated by the addition of anti-L1 antibody to the cell culture (Lemmon et al., 1989).
The mechanism of L1 homophilic interaction has been studied extensively. Initial
studies showed that adhesion of small cerebellar neurons to L1 fragments involved the domains Ig 1-2, Ig5-6, and the FnIII repeats 3-5 (Appel et al., 1993). A further study employing protein-conjugated microspheres refined the adhesion to domains Ig1-2 and the third FnIII domain (Holm et al., 1995). In the human L1 molecule, the homophilic binding site is thought to be centered at the second Ig-like domain. Covaspheres
covalently coupled with Ig2 domain showed a great extent of self-aggregation. As a
substrate, Ig2 is a potent inducer for neurite growth (Zhao and Siu, 1995). Moreover,
deletion of Ig2 rendered human L1 inactive in promoting neurite outgrowth (Zhao et al.,
1998). Similarly, L1 chimeric proteins bearing mutations in Ig2 domain do not engage in homophilic binding (Zhao and Siu, 1996; Kenwrick and Doherty, 1998).
Besides Ig2, FnIII repeat-3 is thought to be another key site for cis-homophilic binding, because this single domain has been shown to undergo spontaneous homomultimerization to form trimers and higher order complexes (Silletti et al., 2000).
The L1 antibody 557, which recognizes a sequence at the joint region of FnIII repeats 2 and 3, inhibits homophilic binding of L1. The synthetic peptide derived from this
41 sequence was shown to be as efficacious at neuritogenesis as L1, and potent to increase intracellular levels of Ca2+ and stimulate the turnover of inositol phosphates in neurons
(Appel et al., 1995).
Based on these observations, Silletti et al. (2000) proposed a model of induction of L1 homophilic interaction. First, a distal ligation event causes the conformation of L1 to change from a closed globular molecule (with a horseshoe fold structure) to an open extended molecule. This switch in conformation requires the involvement of Ig domains, which may be centered at the Ig2 domain. As the molecule opens, exposure of more sites for subsequent avid binding, including FnIII domains, potentiates a greater adhesive state of the molecule. Therefore, engagement of bonding at multiple sites on the molecule stabilizes the homophilic interaction.
4.2. Heterophilic interactions with integrins
Mammalian L1 molecules have a conserved RGD motif in their Ig6 domain for recognition by integrins. The mouse and rat forms of L1 contain another RGD sequence within their Ig6. In chick, one RGD sequence is present in the third FnIII repeat of
NgCAM (Hortsch, 2000). Regardless of its location, the presence of an RGD motif on L1 suggests L1’s capability of binding to integrins (Hynes, 2002). The α5β1 integrin is the first integrin identified to interact with murine L1 (Ruppert et al., 1995). Cells that express both L1 and integrin α5β1 were induced to aggregate upon the activation of the integrin receptor with anti-CD24 antibody (heat-stable antigen). This process of cell aggregation was inhibited by a synthetic RGD-containing peptide (Ruppert et al., 1995).
The αVβ3 integrin has also been demonstrated to interact with human and rat forms of L1
42 (Ebeling et al., 1996; Montgomery et al., 1996). Functionally, the neuritogenic property of L1-integrin binding via RGD motif was first demonstrated by Yip et al. (1998). Chick dorsal root ganglion cells were able to extend neurites on Ig6 substrate, and their projections were disrupted by addition of either a RGD-containing peptide or specific antibody against αVβ3 (Yip et al., 1998).
It is not known whether L1 modulates integrin-dependent cell adhesion by increasing integrin avidity or affinity for ligands, or whether cis binding or intracellular signaling is involved. Nevertheless, the fact that L1 is capable of interacting with integrin receptors implies that L1 is potent to modulate intracellular processes via integrin- dependent mechanisms. Several studies have demonstrated that L1-integrin binding mediates activation of signal pathways via Src, PI3 kinase, MEK and ERK during neuronal process extension (Yip et al., 1998; Schaefer et al., 1999; Schmid et al., 2000;
Ridley et al., 2003). However, the RGD motif is not present in the primary sequences of zebrafish L1 homologs, suggesting that this integrin-binding mechanism may not apply to the zebrafish.
4.3. L1-neuropilin interaction
Emerging evidence suggests that L1 participation in the signaling of a secreted guidance cue of the semaphorin family involves direct interaction of L1 with neuropilin.
L1-deficient mice show axon guidance errors during the corticospinal tract crossing the midline (Cohen et al., 1998), which is known to respond to repulsive semaphorin 3A
(Sema3A) from the spinal cord (Fournier et al., 2000). When cortical slices from the L1 mutant mouse were co-cultured with spinal cord explant or COS7 cell aggregates
43 expressing Sema3A, the axons extending from the cortical slice showed little
responsiveness to the repulsive effects of Sema3A (Castellani et al., 2000). Further
studies has revealed that Sema3A-induced repulsion requires cis-interacting L1 and
neuropilin 1 (Nrp1), whereas trans binding of L1 to cis-interacting L1-Nrp1 complexes converts Sema3A-mediated axon repulsion to Sema3A-mediated axon attraction. This conversion process involves a sequence Phe-Ala-Ser-Asn-Lys-Leu (FASNKL) in the first
Ig-like domain of L1 (Castellani et al., 2002). A mutation (L120V) in this region disrupts
the interaction between L1 and Nrp1 and conversion of the Sema3A repellent effect,
demonstrating a possible pathological mechanism of this mutation for CRASH syndrome
(Castellani et al., 2002).
Switching from repulsiveness to attraction of Nrp1/Sema3A may be mediated by the endocytic pathway. When Sema3A exerts its repulsive effect, it promotes endocytosis of Nrp1 and co-internalization of L1, and subsequently induces growth cone collapse
(Fournier et al., 2000; Castellani et al., 2004). When its repulsive response is converted to attraction by L1-mimetic peptide, both endocytosis and growth cone collapse are blocked (Castellani et al., 2004).
4.4. FIGQY motif and L1-ankyrin binding
Neural migration and neurite outgrowth rely on the dynamics of cytoskeletal elements, including actin filaments and microtubules that interact with an array of cytoskeleton-associated proteins (Wen and Zheng, 2006).
The cytoplasmic tail of L1 harbors a docking site, SFIGQY, for cytoskeletal ankyrin/spectrin, the significance of which is illustrated by two pathological mutations at
44 this site (Y1229H and S1224L) that disrupt ankyrin binding (Needham et al., 2001).
Association with ankyrin is regulated by phosphorylation of the tyrosine residue in the sequence FIGQY, which is conserved among L1 family members including L1, NrCAM and neurofascin (Davis and Bennett, 1994). Phosphorylation of this tyrosine leads to the abolition of ankyrin binding that occurs in regions of neuronal migration and axon extension (Garver et al., 1997; Jenkins et al., 2001), and this process appears to be regulated by the MAP kinase pathway (Whittard et al., 2006). In the initial formation of neurites, the membranous protrusions surrounding the soma contain filamentous actin and ankyrin B that continuously move toward the soma (retrograde flow). By the interaction with ankyrin B, therefore, L1 is coupled to the retrograde F-actin flow in these perisomatic structures to mediate neuritogenesis (Nishimura et al., 2003).
The sequence FIGQY is also demonstrated to interact with doublecortin, a microtubule-associated protein important in the migration of cortical neurons.
Doublecortin recognizes specifically the phospho-FIGQY sequence, providing additional evidence for the pivotal role of FIGQY modulating cytoskeleton dynamics (Kizhatil et al.,
2002).
4.5. RSLE and L1 endocytosis
As discussed previously, the alternative splicing of exon 27 in the L1 gene generates two forms of L1, a neuronal form containing the amino acids Arg-Ser-Leu-Glu
(RSLE) and a non-neuronal form lacking this sequence. In the neuronal form, the phosphorylation of Tyr1176 immediately N-terminal to the RLSE sequence affects the interaction of L1 with the AP-2 adaptor, thus linking L1 to the clathrin-based endocytic
45 machinery (Kamiguchi and Lemmon, 1998). Endocytosis plays an important role in growth cone migration by modulating spatial asymmetry in the adhesive forces of CAMs.
Endocytosis via clathrin-vesicles changes the density and avidity of L1 in a migrating growth cone. L1 is internalized at the central domain of the growth cone, followed by anterograde vesicular transport and recycling to the plasma membrane of the leading front (Kamiguchi and Yoshihara, 2001). Phosphorylation of the tyrosine residue by the non-receptor tyrosine kinase c-src is shown to inhibit L1 binding to AP-2 (Schaefer et al.,
2002). Furthermore, inhibition of L1 endocytosis by abrogation of the endocytic pathway decreases the rate of L1-dependent growth cone migration (Kamiguchi et al., 1998a).
A recent study demonstrates that Ser1181, which immediately follows RSLE motif can be phosphorylated by casein kinase II and that phosphorylation of Ser1181 confers binding ability to AP-2. In addition, mutations at Ser1181 lead to deficits in both
L1 internalization and L1-stimulated axon growth (Nakata and Kamiguchi, 2007). Taken together, phosphorylation of the YRSLE motif plays an important role regulating L1 trafficking in the growth cone critical for axon elongation.
46 5. L1-MUTANT MOUSE MODELS
The L1 protein in mice is very similar in sequence to human L1 protein with 88% identity (Tacke et al., 1987). So far, two transgenic mouse models have been generated that have a targeted null mutation in the L1 gene in the Ig3 and Ig6 domains, respectively
(Dahme et al., 1997; Cohen et al., 1998).
5.1. Resemblances to CRASH syndrome in L1 mutant mouse models
L1 mutant mouse models have produced a phenotype similar to that observed in man. As in the case of L1 patients, the CNS of the mice was largely intact with major malformation limited to hypoplasia of the corticospinal tract, corpus callosum and cerebellar vermis and a degree of ventricular enlargement, which is dependent on the genetic background of the mouse (Dahme et al., 1997; Cohen et al., 1998; Fransen et al.,
1998a; Demyanenko et al., 1999). The mutant mice had a reduced size of the medullary pyramids, consistent with an overall reduction in the corticospinal projection (Dahme et al., 1997). There were no corticospinal axons detected in the spinal cord caudal to the cervical level (Cohen et al., 1998). Moreover, a substantial proportion of corticospinal axons failed to cross the midline at the medullary decussation and instead passed ipsilaterally into the dorsal column (Cohen et al., 1998).
Hydrocephalus can be produced only in the mice of certain genetic background.
Nonetheless, the presence of hydrocephalus shows no correlation with the diminution of the aqueduct of Sylvius that is common in X-linked hydrocephalus patients (Rolf et al.,
2001). In the L1-deficient mice that show dilated ventricles, it is evident that numbers of hippocampal neurons (e.g. pyramidal neurons and granule neurons) are significantly
47 decreased. However, it is unlikely that the decrease in these neurons is associated with
cell death within the neuronal populations because no distinguishable difference between the mutant and wild type has been observed by the Terminal deoxynucleotidyl
Transferase dUTP Nick End Labeling (TUNEL) assay (Demyanenko et al., 1999).
5.2. Discrepancies among mouse models
Mouse models generated by truncation in the Ig3 and Ig6 domains are predicted to give rise to nonfunctional secreted proteins. Therefore, no membrane-bound L1 is expected in the mutants. Although many studies from the mutant mice have contributed greatly to our understanding of L1 function in various neurological processes, major discrepancies are known to exist. First, abnormal decussation of corticospinal projections is found to be apparent in mutants (Cohen et al., 1998). However, based on a clinical study on L1 mutation carrier females and hemizygous males (L1 patients), corticospinal projections have been demonstrated both neurophysiologically and histologically to the lumbar spinal segments, and there is no anatomical or neurophysiological evidence to support significant failure of decussation of corticospinal axons (Dobson et al., 2001). In an L1 patient that has a nonsense mutation affecting Ig 6, which is similar to the mutation used to create the knock-out mouse of Dahme and colleagues (1997), the majority of corticospinal axons were able to decussate at the medulla. It was therefore argued that L1 functions in corticospinal axon projection/elongation instead of guidance (Dobson et al.,
2001).
Secondly, hydrocephalus in CRASH patients has been categorized as obstructive hydrocephalus. One of the possible mechanisms is that impaired outgrowth and/or
48 pathfinding of axons lead to subsequent death of those neurons that are unable to
innervate, resulting in an enlarged chamber. Although defects in the corticospinal tract
are prominent, no significant cell death inside the hippocampus and septum (including
pyramidal neurons which axons form the corticospinal tract) has been detected by
TUNEL assay in the L1 mutant mice (Demyanenko et al., 1999). Therefore, cell death is
less likely to constitute a pathological mechanism for hydrocephalus.
On the other hand, hydrocephalus in L1 patients is frequently found to be
associated with prominent stenosis of cerebral aqueduct (Fransen et al., 1997; Weller and
Gartner, 2001). However, the current mouse models fail to correlate the blockage of the
aqueduct to the resultant enlargement of lateral ventricles (Rolf et al., 2001).
Furthermore, the current mouse models do not show the phenotype that neuronal cells position aberrantly within the ventricular proliferative zone of the hydrocephalus brain (Kamiguchi et al., 1998a). There is no supporting evidence from the mouse studies for abnormal neuronal proliferation or migration in the anterior vermis of the cerebellum and thalami that are phenotypic in some L1 patients (Yamasaki et al., 1995).
These discrepancies may rise from the presence of residual fragmented proteins that possibly possess partial functions of the wild-type protein. Indeed, these mutants contain residual proteins that are immunoreactive to polyclonal anti-L1 antibodies raised against the extracellular domains of L1 (Dahme et al., 1997; Cohen et al., 1998).
Moreover, genetic background may also contribute to the variable phenotypes
(Kamiguchi et al., 1998a). Therefore, it is necessary to explore L1 functions in a new in vivo system, where examination of a large population of mutants is permissible and phenotypes of various disease states of CRASH syndrome are assessable.
49 6. THE ZEBRAFISH MODEL
The zebrafish (Danio rerio) is an excellent model for study of vertebrate development and disease. During the late 1960s, phage geneticist George Streisinger from the University of Oregon began to look for a model system to study the genetic basis of vertebrate neural development. His passion for tropical fish led him to the humble zebrafish. It took him almost ten years to publish his first zebrafish paper, which provided a detailed description of the normal morphological and functional development of the zebrafish embryo. He also developed a technique for producing homozygous diploid fish, which makes it possible to detect rare recessive mutations and to produce large clones of genetically identical fish (Streisinger et al., 1981).
In the early 90s, two large genetic screens were performed in Tübingen, Germany and Boston, Massachusetts to identify genes with unique and essential functions in zebrafish development. Thereafter, the zebrafish has quickly emerged as a model organism important for the identification and characterization of genes and pathways involved in development, organ function, behavior, and disease (Eisen, 1996).
6.1. Advantages of using zebrafish as a model system
Many attributes of zebrafish make it an attractive and quickly accepted model system for studying development and diseases. First, its fertilization and development are external, permitting direct observation and manipulation of embryos under a wide variety of laboratory conditions. Secondly, zebrafish embryos and early adults are optically transparent, a characteristic that facilitates direct observation of internal organs by light microscopy. Combination of these two advantages allows the behavior of single cells or
50 groups of cells in the embryo to be followed by real-time imaging in the native
environment of the developing embryo, when the specific cells or tissues are labeled with
a fluorescence tag. Third, zebrafish is a vertebrate with simple and cost-effective
husbandry requirements. Fourth, it is relatively fecund and generates clutches of 100–200 embryos weekly. Moreover, its embryonic development is rapid. Most major organs including the gut and the vasculature are in place by 2 days post fertilization and
embryogenesis is complete 5 days after fertilization.
Furthermore, during organogenesis, zebrafish embryos are permeable to small
molecules and drugs, providing easy access for drug administration and vital dye staining
(Zon and Peterson, 2005).
6.2. Overview of current applications and techniques in the zebrafish research
In the past decade, thousands of mutations have been generated in large-scale
forward genetic screens of zebrafish (Eisen, 1996). Such large-scale genetic screens had
previously been limited to invertebrates such as flies, worms and yeast. Zebrafish made it
possible to take the forward genetic approach to investigate vertebrate-specific processes
that affect development and disease. Mutations obtained from the large scale genetic
screens affect organogenesis, physiology as well as behavior (Driever et al., 1996;
Haffter et al., 1996). These mutations have proved to be a rich source of information
about the relationships between genes and function. The zebrafish genome has been
sequenced (http://www.sanger.ac.uk/Projects/D_rerio/), also substantial annotation of the
genome is accessible through the trans-National Institutes of Health Zebrafish Genome
Initiative (http://www.nih.gov/science/models/zebrafish/). Several additional newly
51 developed tools, as summarized in Table 1.7, have greatly increased the utility of the zebrafish as an experimental organism.
6.2.1. Forward genetics-ENU mutagenesis screening
Large-scale genetic screens have been performed in zebrafish using ethylnitrosourea (ENU) as a chemical mutagen to generate mutations. More than 7000 mutations in 600 genes have been generated, affecting every aspect of development
(Driever and Fishman, 1996; Haffter et al., 1996).
There are a few peculiarities of genetic screens that are eminently suitable for extrapolation to human diseases. First, phenotype is the entrance filter for genetic screens.
Combinations of defects may reflect syndromes; i.e., patterns of multiorgan dysfunction that coexist because of shared underlying pathways. In addition, many common human diseases, even if heritable, are not due to null mutations. Unlike targeting of genes for disruption in mice, which intentionally renders them null, ENU often causes partial hypomorphic (loss-of-function) mutations, which may reveal more subtle effects.
Therefore, ENU-induced mutations are potentially more relevant to human disease than null mutations in which early or widespread dysfunction may obscure the role of a gene in later organ formation. Furthermore, because functional analysis comes first in genetic screens with no presupposition of the protein structure, ENU-induced mutations may reveal previously undefined important domains (Shin and Fishman, 2002).
52 Table 1.7. The zebrafish toolbox
Adapted from Zon and Peterson (2005)
Technology Description References
Forward genetics Chemical mutagenesis Ethylnitrosourea (ENU) as the mutagen; high mutation Solnica-Krezel et al., 1994 rates, large-scale screens Insertional mutagenesis Insertion is facilitated by retrovirus, efficient cloning of Amsterdam, 2003 mutations in the presence of a molecular tag Reverse genetics Morpholinos Antisense oligos against transcription initiation or splicing; Nasevicius et al., 2000 rapid, inexpensive gene knock-downs TILLING Directed identification of permanent mutations in genes of Wienholds et al., 2002 53 interest Expression profiling Gene chip Zebrafish genome arrays from Affymetrix are used to study http://www.affymetrix.com/products/arrays/specific/zebrafish.affx gene expression of over 14,900 Danio rerio transcripts Spotted microarrays cDNA and oligonucleotide microarrays Stickney et al., 2002; Ton et al., 2002 Other tools Genomic sequence 5.7-fold coverage of the zebrafish genome The Sanger Institute Danio rerio Sequencing Project: http://www.sanger.ac.uk/Projects/D_rerio Substantial genome annotation Trans-NIH Zebrafish Initiative, http://www.nih.gov/science/models/zebrafish Transgenesis Rapid production of stable transgenic lines by the injection Meng et al., 1999; Davidson et al., 2003; Kurita et al., 2004 of plasmid, retrovirus and transposon cDNA collections Full-length cDNA collections http://zebrafish.org/zirc/est/estAll.php Mutant collections Thousands of categorized mutant lines http://zfin.org Hundreds of lines available through public stock centers http://zebrafish.org/zirc/fish/lineAll.php Physical and genetic maps Radiation hybrid and microsatellite genetic linkage maps Geisler et al., 1999; Hukriede et al., 1999 6.2.2. Reverse genetics-morpholino knockdown approach
Morpholino knockdown screens constitute an effective means of systematically
assessing the roles of individual genes in many developmental and disease processes.
Morpholino oligonucleotides typically contain 25 morpholino subunits, each of which is
comprised of a nucleic acid base, a morpholine ring and a non-ionic phosphorodiamidate
intersubunit linkage (Summerton, 1999). Differing from RNA interference, morpholinos
efficiently exert knockdown of gene function via an RNase-independent mechanism.
When injected into the cytosol of zebrafish embryo, they can block translation initiation
by targeting the 5' untranslated region (UTR) through the first 25 bases of the coding
sequence (Nasevicius and Ekker, 2000). Once in the cytosol, morpholinos also freely
diffuse between the cytosol and the nucleus, where they can modify pre-mRNA splicing
in the nucleus (when splice junctions are targeted) or they can block microRNA (miRNA)
activity (Bruno et al., 2004; Flynt et al., 2007). Morpholino oligonucleotides have been found highly stable in cells and are widely used as an effective and specific method to knockdown gene functions in the zebrafish.
6.3. TILLING project (Targeted Induced Local Lesions IN Genomes)
TILLING is a recent approach to knockout the targeted gene activity in zebrafish.
It involves random mutagenesis with ENU, followed by targeted screening for induced
mutations at the genomic DNA level. The first step in TILLING is to mutagenize male zebrafish and mate them with untreated female fish. When the offsprings are raised to adulthood, genotyping will be performed to examine any fish that carries a mutation within the gene of interest (Sood et al., 2006). Once a carrier fish bearing a desired
54 mutation is identified, it will be out-crossed to obtain homozygous mutant fish, in which
the correlated phenotypes will be studied (Wienholds et al., 2002).
TILLING appears to be a powerful alternative approach to study the function of
genes which are difficult to examine by genetic screens. So far, more than 146 mutant
fish lines have been collected using this target-selected mutagenesis technique by the
European ZF-MODELS consortium, including a splice mutation in the L1.1 gene
(http://www.niob.knaw.nl/researchpages/cuppen/zfmodels/).
6.4. Transgenic reporters
The first stable lines of transgenic zebrafish were first generated over a decade ago (Stuart et al., 1988; Stuart et al., 1990). Recently, techniques that reliably produce transgenic animals on a routine basis have been developed by injecting plasmid or using retrovirus and transposons. When a fluorescent tag is coupled to the reporter or the desired biological marker under a tissue-restricted or inducible promoter, these techniques have generated numerous transgenic animals in which developmental stage- and tissue-specific gene regulation, cell migration and targeted misexpression can be examined. The power of transgenic reporters is magnified when combined with advanced confocal imaging technologies to provide a non-invasive system to study dynamic physiological processes in the context of the intact organism (Udvadia and Linney, 2003).
Recent studies utilizing the transgenic reporter system have contributed significantly to our understanding of the cellular mechanisms of vertebrate gastrulation (Solnica-Krezel,
2005), brain morphogenesis (Koster and Fraser, 2001; Langenberg and Brand, 2005) and organogenesis of the heart (Trinh and Stainier, 2004).
55 Beyond cell behavior, transgenic reporter systems in the zebrafish have greatly
improved our understanding of the molecular and cellular mechanisms of the protein of
interest. For example, Scholpp and colleagues (2004) utilized fluorescence-tagged Fgf8
protein to “visualize” its effective signaling range in nascent neuroectoderm, which
appears to depend on internalization of Fgf8 in the responsive cells.
Furthermore, in combination with other modifications, such as RNA caging (a
modification on RNA synthesized in vitro to control the initiation of RNA translation,
Ando et al., 2001) and Fluorescent Timer (a recently developed fluorescent protein that shifts colors of different wavelengths over time, providing maturation information of the tagged protein, Terskikh et al., 2000), transgenic reporters permit a much more sophisticated real-time live imaging system in the study of cell behavior during development.
6.5. Zebrafish as a model for human disease
As discussed earlier, forward genetic screens in zebrafish have generated tremendous information on embryonic development in the vertebrate. In addition, several disease-related screens have been performed to extend our understanding of these pathogenic processes. These screens have collected recessive mutants with lethal developmental phenotypes that usually represent the most severe form of the corresponding human syndrome. In some instances, the mutant phenotypes have been sufficiently similar to the disease pathology in humans to allow the mutation to be identified by a candidate gene approach. For example, the cellular pathology of zebrafish muscle degeneration mutants presented superficial similarities to human muscular
56 dystrophies. One of these mutants, sapje, carries a defect that impairs the zebrafish dystrophin gene, which is homologous to the human gene affected in Duchenne muscular dystrophy (Bassett et al., 2003). Human heart disease can be recapitulated in several zebrafish mutants. In both zebrafish and humans, TTN (titin) mutations cause
cardiomyopathy (Xu et al., 2002), while TBX5 (T-box 5) mutations cause congenital
heart defects (Garrity et al., 2002).
In the nervous system, mutations in the cadherin23 gene are responsible for
sputnik phenotypes showing defects in hair cell mechanosensation (Nicolson et al., 1998).
Mutations in cadherin23 also cause deafness and vestibular defects in mice and humans,
therefore, sputnik is regarded as a model for human deafness (Sollner et al., 2004). Other
zebrafish models of nervous system disorders have been described, including retinal
degeneration and anxiety (Li and Dowling, 1997; Peitsaro et al., 2003).
Examination of zebrafish mutations can inversely lead to discovery of the
candidate gene(s) responsible for the human disease. An excellent example is the
identification of ferroportin (an iron transporter) in a form of anaemia. By positional
cloning, the gene responsible for hypochromic anaemia in the zebrafish mutant
weissherbst has been identified to be ferroportin1, which functions in iron transport from
mother to embryo (Donovan et al., 2000). Subsequently, it was found that ferroportin is
defective in patients with haemochromatosis type 4 (Njajou et al., 2001).
Once a zebrafish disease model is generated, the ease of embryo manipulation
enables further investigation of the molecular and cellular basis of the disease, as well as
drug screening to reverse the disease phenotypes. Furthermore, zebrafish embryos and
juveniles are highly accessible for high-throughput functional and morphological
57 analyses in the multi-well assay plate (Langheinrich, 2003; Zon and Peterson, 2005; Kari
et al., 2007). Therefore, the zebrafish system may provide a shorter path to developing novel therapies for human disease.
58 7. EMBRYONIC NEURAL DEVELOPMENT IN ZEBRAFISH
The zebrafish presents an excellent model for early embryonic development. The
zebrafish brain possesses conserved anatomy shared by other vertebrates: forebrain
(subsequently divides into telencephalon and diencephalon), midbrain or mesencephalon,
hindbrain or rhombocephalon, specialized sensory organs (eye, olfactory system and ear),
peripheral nervous system with motor and sensory components, and enteric and
autonomic nervous systems. Also, the zebrafish exhibits “higher” behaviors and integrated neural function, including memory, conditioned responses and social behaviors such as schooling (Lieschke and Currie, 2007).
7.1. Landmarks of zebrafish neural development
Following fertilization of the egg, zebrafish development proceeds through a number of morphologically distinct stages including zygote (0-0.75 hour post fertilization, hpf), cleavage (0.75-2.25 hpf), blastula (2.25-5.25 hpf), gastrula (5.25-10.33 hpf), and segmentation (10.33-24 hpf; Kimmel et al., 1995). Several key stages during early development of the nervous system are illustrated in Figure 1.6. The fertilized zebrafish
egg is telolecithal. The cytoplasm (the blastodisc, which gives rise to the embryo proper)
sits on the large mass of yolk, defining the animal-vegetal axis. Like other teleost zygotes,
it undergoes meroblastic (incomplete) cleavage until the 16-cell stage. Blastomeres
mount on the animal pole until about 4 hpf, when the blastodisc begins to thin and spread
over the surface of the yolk cell. This process is called epiboly and continues until the
yolk cell is completely engulfed (Kimmel et al., 1995).
59 1-cell Non-yolky cytoplasm begins to stream towards the (0 hpf) animal pole, segregating the blastodisc from the clearer yolk granule.
Shield The blastoderm spreads by epiboly around the (6 hpf) yolk, resulting in two layers of cells. The outer cell layer (epiblast) gives rise to the neural system.
Bud stage First neuronal markers are detectable in the (10 hpf) polster, indicating the onset of neural system development.
18 somite stage Primary neurons are programmed and start to (18 hpf) project their axons. Ventricle formation of the brain has initiated.
25 somite stage Posterior lateral line neurons develop central and (20 hpf) peripheral axons. Contractions of myotomes become more coordinated and more frequent.
30 somite stage Brain structures become prominent. Ventricles are (24 hpf) connected and filled with cerebrospinal fluid. Heart beats.
250 μm
Figure 1.6. Early development of the zebrafish embryo. Living embryo are staged according the morphology and hours post fertilization (hpf) shown on the left. The corresponding developmental landmarks in the nervous system are presented on the right (Kimmel et al., 1995).
60 Gastrulation begins at 50% epiboly, when cells involute within the thickened
marginal region. Involution of germ cells gives rise to two layers of cells. The outer layer
is called epiblast and the inner layer is called hypoblast. The cells in the epiblast
correspond to the ectoderm and will give rise to such tissues as epidermis, the central
nervous system, neural crest, and sensory placodes. The hypoblast gives rise to both the
mesoderm and endoderm (Kimmel et al., 1995).
A specific group of cells in the shield, which resides in a small region of the
margin, are identified as an organizer that is capable of defining the dorsal-ventral axis
and inducing the primitive nervous system (Shih and Fraser, 1996). When the organizer
is transplanted to the ventral side of the embryo, it will induce ectopically a secondary
axis (Saude et al., 2000). It is believed that the organizer expresses the molecules
Chordin and Noggin which antagonize Bone morphogenetic protein (BMP) signaling and
induce the formation of neural tissue from the dorsal ectoderm. Subsequently, the
organizer produces signals, including Wnts, FGFs and retinoic acid, that further influence
anterior and posterior division of neural tissues (Appel, 2000).
At the end of gastrulation (~10 hpf), the dorsal epiblast begins to thicken at the
anterior-most region called polster and at the midline (Kimmel et al., 1995). At this stage, expression of early neuronal determinators (such as the basic helix-loop-helix transcription factors neurogenin-related protein 1, neuroD, and Islet-1) is initiated, defining the rudiment of nervous system, i.e. the neural plate (Inoue et al., 1994; Korzh et al., 1998).
Soon after the completion of gastrulation, the brain primordium develops at the polster and has distinctively thickened into the neural keel, which is a solid mass or rod
61 of cells. In the anterior trunk, neural keel formation occurs in between the 6- and 10-
somite stages. Subsequently, the lumen of the neural tube opens, beginning ventrally and moving dorsally. Formation of the neural tube in the zebrafish trunk is previously known as "secondary neurulation", as opposed to “primary neurulation” which involves columnarization of existing epithelial neural plate and then rolling and folding the epithelium (Schmitz et al., 1993). However, recent evidence argues for the involvement of a primary neurulation mechanism. First, the neural tube arises clearly from epithelium, differing from condensation of mesenchyme that occurs in secondary neurulation. The midline of the neural keel is distinct with a pseudo-stratified columnar epithelium on either side (Lowery and Sive, 2004). Furthermore, confocal time-lapse imaging shows a clear rolling of the neural plate in vivo, which is a characteristic of primary neurulation
(Geldmacher-Voss et al., 2003).
At 17-18 hpf, four prominent subdivisions of the brain, the telencephalon, diencephalon, midbrain and hindbrain, become distinguishable (Kimmel et al., 1995).
Initiation of the brain ventricle formation occurs at this stage (Lowery and Sive, 2005).
During the early morphogenesis of the brain, striving proliferation and coordinating migration of the cells in the developing ventricles are essential, sharing regulatory mechanisms highly conserved in vertebrates (Wullimann and Knipp, 2000). In the trunk, primary neurons are differentiated into three classes (sensory neurons, interneurons and motoneurons) as early as 15 hpf (Kuwada, 1986; Eisen et al., 1989; Kuwada et al., 1990).
They are repeated in the spinal cord segments along the anteroposterior axis (Kuwada et al., 1990). At 18 hpf, primary motoneurons start extending their central axons as well as peripheral projections along the somite in a wave proceeding rostrocaudally (Bernhardt et
62 al., 1990). Being innervated, contractions of myotomes become progressively stronger and more frequent at 20 hpf (Hanneman and Westerfield, 1989).
By the end of the first day, most of the brain structures have become prominent, including the telencephalon, diencephalon, mesencephalon (including tegmentum, optic tectum), epiphysis, mid-hindbrain boundary, cerebellum and rhombencephalon (Haffter et al., 1996). The ventricular system has developed as a free passage ready for circulating the CSF. Primary neurons, including sensory neurons, interneurons and motoneurons, all initiate axons and synaptically interconnect each other. Furthermore numerous neurons, regarded as secondary population because they develop later and are smaller in soma size, have arisen in the hindbrain and spinal cord. They will project their trajectories at a later stage (Westerfield, 1992; Kimmel, 1993).
7.2. Brain domain organization of early zebrafish brain
During early development, the simple sheet of neuroepithelial cells that constitutes the neural plate is transformed into the highly complex and elaborative structures of the central neural system. Brain structures become noticeable under the stereomicroscope after 16-17 hpf, while positional identity of the cells constituting these structures have been specified as early as onset of gastrulation (Kimmel et al., 1995; Woo et al., 1995). Identification of the origin of the brain structures is permissible by tracing labeled neural plate cells for their destinations in the brain at later developmental stages
(Woo et al., 1995). From the fine-tuned fate map of 6 hpf (Woo and Fraser, 1995), progenitors of forebrain neurons are clustered around the dorsal midline near the animal pole; midbrain progenitors lie vegetal, or posterior, to forebrain precursors at dorsal and
63 lateral positions; whereas cells with hindbrain fate lie closer to the germ ring, lateral to the shield. Prospective hindbrain and spinal cord cells are displaced laterally and move to dorsal positions by 10 hpf at the end of gastrulation (Woo and Fraser, 1995).
How do the cells arrive at their final destination? First, they must acquire information regarding their location along the antero-posterior and dorso-ventral axes. In some cases, they are determined to reside on the left or right sides of the brain. The positional identity of cells is known to be mediated mainly by three signaling pathways, the BMP pathway, the Hedgehog (Hh) pathway and the Nodal pathway (Schier and
Talbot, 2005).
The earliest DV pattern within the anterior neural plate may be established by a gradient of BMP activity (Barth et al., 1999). Subsequent to this, the Nodal and Hh pathways act to specify ventral fates and refine dorso-ventral pattern throughout the forebrain. For instance, Nodal signaling is required both for the forward movement of ventral CNS cells (Varga et al., 1999) and specification of optic stalks and ventral telencephalic identity (Rohr et al., 2001; Take-uchi et al., 2003). With respect to early antero-posterior patterning of the forebrain neuroepithelium, recent research has implicated the Wnt signaling pathway in this process (Wilson and Houart, 2004). It is evident that suppression of Wnt activity is required for establishing the fate of anterior forebrain, which gives rise to the telencephalon (Houart et al., 2002), whereas the posterior portion of the forebrain develops into diencephalon. For the left/right positioning of the cells in the forebrain, Nodal signaling has been implicated in regulating asymmetry development (Halpern et al., 2003). Additional to the Nodal pathway, Wnt
64 signaling was recently found indispensable in the regulation of left/right asymmetry (Carl et al., 2007).
Along the anteroposterior axis, another well-studied organizing center is the isthmic organizer or mid-hindbrain boundary (MHB) organizer. It appears that the MHB organizer is specified during the initial subdivision of the neuroectoderm. It is positioned in response to a Wnt8 gradient (Rhinn et al., 2005). At a later stage, MHB is induced by
Fgf8 signals, which are responsible for the maintenance of gene expression, MHB morphogenesis and lineage restriction. Isthmic Fgf8 expression regulates the MHB patterning and later aspects of development of the adjacent territories and maintains positional cell identities in the midbrain and hindbrain (Rhinn et al., 2006). The importance of the isthmic organizer to brain regionalization can be demonstrated in the no isthmus (noi) mutant. In noi mutant embryos, lack of the isthmic organizer leads to absence of MHB and, concomitantly, loss of midbrain and hindbrain structures (Brand et al., 1996).
7.3. Cell migration during embryonic brain formation
Upon acquisition of positional identity, newly born neurons often migrate long distances to reach their final destinations. Although the full extent of migrations within the zebrafish brain remains to be illustrated, several migratory processes have been
characterized by confocal time-lapse imaging. Many aspects of the neuronal migration in
the zebrafish embryo are shown to be conserved among vertebrates.
During the early morphogenesis of the midbrain, MHB and hindbrain, newborn
neurons generated from the neuroepithelium migrate radially, coinciding with constant
65 expansion of the ventricles (Langenberg et al., 2006). Following the lineage of the migrating cells, it is apparent that these cells acquire the positional identity at birth and their migration is highly restricted within the boundaries of Fgf8, Wnt1, Otx2 and Gbx2 expression (Langenberg and Brand, 2005).
In the dorso-anterior hindbrain, the upper rhombic lip (URL) is the germinal zone for neurons that constitute the vertebrate cerebellum. The migratory route of these cells has been defined by labeling newborn cells with fluorescence protein and tracing their migration in the 2-day embryo. Cells leaving from the germinative URL first migrate rostrally toward the MHB. Once they reach the MHB, they move ventrally along the
MHB until they finally settle in the region of the ventral brain stem (Koster and Fraser,
2001).
Neuronal migration has also been demonstrated during several specified processes in brain development, including parapineal precursors crossing the midline to form a left- sided nucleus in response to Nodal signal (Concha et al., 2003) and posteriorization of cranial motor neurons within the hindbrain under the synergic regulation of Prickle1 and trilobite proteins (Jessen et al., 2002; Carreira-Barbosa et al., 2003).
7.4. Ventricle formation of the zebrafish brain
Zebrafish brain ventricle formation involves an intrinsic process independent of
CSF circulation and subsequently the additive effects of ventricle expansion and the force generated by the circulating CSF. The intrinsic process of ventricle morphogenesis requires establishment and maintenance of proper polarity at the ventricular epithelia and regional cell proliferation (Lowery and Sive, 2005), as well as coordinated migration of
66 newborn neurons from the germinative zone (Koster and Fraser, 2001; Langenberg and
Brand, 2005). Disruption of either polarity at the neuroepithelium or proliferation inhibits
this initial step of ventricle formation. When CSF begins to circulate in the ventricle following the establishment of heart beat, the circulation pressure on the ventricle and the intrinsic morphogenesis synergistically expand the ventricle (Lowery and Sive, 2005).
Polarization of the ventricular neuroepithelia is found to be intrinsic and essential to brain ventricle formation. During embryonic brain development, polarized distribution of supernumerary centrosomes and adherens junctions is observed along the ventricular surface of the ventricular zone in the ferret (Chenn et al., 1998) and the mouse (Yang et al., 2003). In zebrafish, a membrane-associated guanylate kinase-family scaffolding protein (MAGUK), which is required in establishing neuroepithelial polarity (Wei and
Malicki, 2002), is indispensable to proper brain ventricle formation. Mutations in nagie
oko (nok), which encodes a MAGUK family protein, result in the failure of initial
ventricle opening in all the brain regions (Lowery and Sive, 2005).
Migration of newborn neuronal cells from the ventricular proliferation zone is
also fundamental to the formation of ventricles. In the case of the Nok mutant, absence of
the ventricle is associated with disorganization of neurons (Wei and Malicki, 2002;
Lowery and Sive, 2005). Zebrafish share conserved migration mechanisms with other
vertebrates, including radial migration and tangential migration. ECM and cell adhesion
molecules play an important role in migration of neuronal cells, and therefore influence
ventricle formation. For example, laminin acts as a permissive migratory substrate for
telencephalic neural progenitors migrating in the mouse neocortex (Mizuno et al., 2005),
possibly via the interaction with integrin α3β1 (Hatten, 2002). Interestingly, mutations in
67 three zebrafish laminin subunits, including bashful/laminin α1, grumpy/laminin β1 and sleepy/laminin γ1, lead to similar abnormalities that manifest an irregularly shaped brain with a severely enlarged hindbrain ventricle. These results suggest that the functions of laminin in mediating neuronal migration are conserved among vertebrates and its malfunctioning may disrupt normal ventricle formation (Schier et al., 1996; Parsons et al.,
2002; Paulus and Halloran, 2006).
At the stages post heart beating (24 hpf), the application of CSF pressure on the ventricle wall facilitates further inflation of the ventricle. Implication of CSF on ventricle formation is based on morphological characterization of the mutant snakehead, which has defects in heart-beating. In snakehead, the ventricle fails to inflate despite the fact that initial morphologies appear normal (Schier et al., 1996; Lowery and Sive, 2005).
7.5. Axogenesis in the early zebrafish CNS
The earliest neurons become postmitotic in the later stage of gastrulation, characterized by the presence of intracellular enzyme acetylcholineesterase (Ross et al.,
1992). These early differentiated neurons are termed primary neurons because of their distinctive appearance, which includes large soma size, long axons, and paucity in number (Kimmel, 1993). Primary neurons include all three classes, sensory neurons,
interneurons and motoneurons. Their development in the embryo appears in a rostral-to-
caudal wave. At 16 hpf, the first axons emerge and start to navigate to the
neuroepithelium of the zebrafish brain. The earliest axons grow from the clusters of cells
and form bundles. These bundles fasciculate and ultimately establish the connection between the brain and spinal cord and between the CNS and periphery. Wherever axons
68 grow in the early brain, the pattern is discrete, precise, and stereotyped (Hjorth and Key,
2002).
By 24 hpf, a simple scaffold of bilaterally symmetric axon tracts and commissures are already present in the brain and the spinal cord (Kimmel, 1993). A single prominent longitudinal system of collected bundles, called medial longitudinal fasciculus (MLF), appears continuous along the whole length of the ventral part of the brain. Descending axons from the telencephalon, diencephalon and the midbrain are bundled to the MLF
(Chitnis and Kuwada, 1990). In the hindbrain, MLF is pioneered by caudal hindbrain reticulospinal interneurons and projects caudally into the spinal cord (Mendelson, 1986).
Another prominent longitudinal bundle present in the hindbrain is the lateral longitudinal fascicle (LLF). The LLF arises from axons from primary sensory neurons.
Caudally growing axons from the trigeminal ganglion neurons meet rostrally growing axons from Rohon-Beard cells located in the spinal cord (Metcalfe et al., 1990).
Primary motoneurons in the spinal cord are present in a single bilateral pair of clusters in each spinal segment and innervate muscle fibers in the corresponding pair of myotomes (Eisen, 1991). Three specified primary motoneurons are present in every hemisegment of spinal cord and, according to the locations of the muscle fiber they innervate, they are CaP (caudal primary motoneuron innervating the ventral myotome),
MiP (middle primary motoneuron innervating the dorsal myotome), and RoP (rostral- most primary motoneuron innervating the region between the two myotomes). The CaP initiates outgrowth ventrally from the spinal cord and pioneers the common path for the axonal processes extending from MiP and RoP (Eisen, 1991; Melancon et al., 1997). CaP axon grows to the horizontal myoseptum, and pauses at muscle pioneer cells, a group of
69 specific adaxial muscle cells, before it continues specifically to the ventral myotome,
where it branches to form its terminal arbor (Eisen, 1991). Primary motoneurons functionally interact with muscle cells at the synaptic structure, called neuromuscular junctions (NMJs), while extending their growth cones into the periphery. Blocking postsynapse formation by cholinergic antagonists abolishes the axonal growth
(Westerfield et al., 1986). Evidence from time-lapse imaging suggests that the growth cone of CaP axons navigates on pre-patterned synapses. Clusters of acetylcholine receptors (AChRs) pre-exist along the prospective axonal path in the central region of the myotome and are incorporated into NMJs as axons advance (Flanagan-Steet et al., 2005;
Panzer et al., 2005; Panzer et al., 2006).
70 8. A COMPARATIVE ANALYSIS OF EMBRYONIC BRAIN DEVELOPMENT
IN THE VERTEBRATES
Many aspects of brain development are conserved among vertebrates, including neuroanatomy and gene expression patterns. However, there are distinct specializations, such as the six-layered cortex for the mouse, the everted telencephalon for the zebrafish, and the dorsal ventricular ridge for the chick (Wullimann and Mueller, 2004). In this section, the conserved features among vertebrates will be examined to assess common processes in brain function, development and evolution.
8.1. Comparison of brain organization
Zebrafish forebrain gives rise to telencephalon as well as diencephalon. Similar to other teleosts, the telencephalon has only a rudimentary cortex and it everts. However, in vertebrates, the telencephalon gives rise to the cerebral cortex, basal ganglia and hippocampus by the process of evagination (Salas et al., 2003). Contrary to the telencephalon which is remarkably different from other vertebrates, the zebrafish diencephalon displays a series of brain subdivisions characteristic of all vertebrates, including the epithalamic structures (epiphysis, habenula) and the hypothalamus with the attached pituitary (Wullimann and Mueller, 2004).
Vertebrates share conserved organization of the midbrain and MHB regions in terms of gene expression patterns (Figure 1.7), having the expression of orthodenticle homologue (Otx) family homeobox genes at the midbrain and paired box 2 (Pax2) at the
MHB anterior to a narrow Fgf8 region (Wurst and Bally-Cuif, 2001). Downstream of
Pax2, the homeodomain transcription factor Engrailed is required to maintain midbrain
71 Otx2 Pax2 M HB Engrailed
Midbrain H r1 i D r2 nd Fgf8 r3 br ai r4 n Krox20 r5 T r6 r7
Figure 1.7. Patterning of the zebrafish embryonic brain. Expression of putative regulatory genes subdivide the brain in distinctive patterns that are clearly related to the neuromeres. Left side view at 18 hpf. Otx2 is expressed in the diencephalon and midbrain (Mori et al., 1994). At the MHB, Pax2 expression (Krauss et al 1991b) regulates the downstream signaling of Engrailed (Hatta et al., 1991), which is rostral to the region governed by Fgf8 (Reifers et al.,1998). The expression of Krox20 is restricted to the rhombomeres 3 and 5 (Oxtoby and Jowett, 1993). T: telencephalon; D: diencephalon; MHB: mid-hindbrain boundary. r: rhombomere.
72 fate in chicken, mice and zebrafish (Araki and Nakamura, 1999; Scholpp and Brand,
2001; Wurst and Bally-Cuif, 2001). MHB serves as a prototypical local organizer of the
embryonic brain and induces the morphogenesis of the midbrain and hindbrain.
Conversely, complete or partial knockout of the engrailed and Fgf8 genes expressed only
on either side of the mid-hindbrain junction lead to the gradual disappearance of both
mesencephalic and metencephalic structures in mouse and zebrafish, indicating a positive
feedback loop in the maintenance of the MHB (Lun and Brand, 1998; Meyers et al., 1998;
Reifers et al., 1998; Picker et al., 2002).
The hindbrain is the most evolutionarily ancient part of the vertebrate brain
(Jackman et al., 2000). Its basic organization into a series of seven or eight segments
termed rhombomeres is similar across vertebrate species (Moens and Prince, 2002).
The morphological segmentation of the zebrafish hindbrain is transiently visible at the 18
hpf (Kimmel et al., 1995), when five prominent bulges along the anterior-posterior extent
of the hindbrain, namely rhombomeres (r)2-r6, are detectable in the vicinity of the
developing otic vesicle, which lies lateral to r5. The order of boundary formation is
stereotypical for embryos of a given species. In zebrafish, the r4 territory is defined first
with the appearance of the boundary between r3 and r4 (r3/4) and then the r4/5 boundary.
At the anterior end of the hindbrain, r1 is usually described as a large segment extending
from the MHB junction to the r1/2 boundary (Moens and Prince, 2002). Fate mapping
experiments in the mouse, zebrafish, and chick have shown that cells in this region
contribute to the cerebellum (Zinyk et al., 1998; Wingate and Hatten, 1999; Koster and
Fraser, 2001). The organization of r1 is complex, with distinct classes of neurons
differentiating along its anteroposterior axis (Koster and Fraser, 2001). Regarding to
73 conserved gene expression pattern, krox-20, which encodes a zinc-finger transcription factor with an important role in hindbrain patterning, is discretely expressed in the third and fifth rhombomeres in many vertebrate embryos, including the zebrafish (Oxtoby and
Jowett, 1993).
8.2. Coordinated proliferation and migration of neuronal progenitors at the
ventricle of the embryonic brain
During the development of the brain, neurons are generated from precursor cells that line the walls of the ventricular system deep within the brain, then they often migrate long distances from their birth place to reach their final destination. Disruption of either proliferation or migration of the neuronal cells will reciprocally affect the other process and subsequently lead to failure in ventricle formation. Our current knowledge of the migration route map of these new born neurons that leave the germinal ventricular zone comes largely from fate-mapping studies in rodent models (Figure 1.8). Recently, time- lapse confocal imaging in the live zebrafish embryo facilitates resolving the migration route map in vivo.
8.2.1. Neuroepithelial proliferation
Proliferation at the ventricular neuroepithelia involves both symmetric and asymmetric divisions of progenitor cells (Huttner and Kosodo, 2005). Time-lapse microscopy of dividing cells in slices of developing cerebral cortex reveals that cleavage orientation predicts the fates of daughter cells. Vertical cleavages produce behaviorally and morphologically identical daughters that resemble precursor cells; these symmetric
74 A. C. Locomotion Ventricle
MZ CP SP
B. Translocation IZ Pia
PP
VZ VZ
Ventricle Ventricle
Mitotic neuron Glia Migrating neuron
Figure 1.8. Different stages of radial neuronal migration in the developing brain. (A) Schematic diagram of a section through the developing rodent forebrain. The dorsal forebrain gives rise to the cerebral cortex. Radial migration in the boxed area is shown in B at early stage and C at later stage. (B) During early corticogenesis, translocation is the predominant mode of movement. Mitosis occurs at the surface of the neuroepithelium, with the division plate vertical or horizontal to the ventricle surface. Neuronal migration begins when the first cohort of postmitotic neurons moves out of the ventricular zone (VZ) to form the preplate (PP). (C) At later stages, glia- guided migration is more prevalent. Subsequent cohorts of neurons (pyramidal cells) migrate through the intermediate zone (IZ) to split the PP into the outer marginal zone (MZ) and inner subplate (SP). CP, cortical plate. Adapted from Nadarajah and Parnavelas (2002).
75 divisions may serve to expand or maintain the progenitor pool. In contrast, horizontally
dividing cells produce basal daughters that behave like young migratory neurons and apical daughters that remain within the proliferative zone (Chenn and McConnell, 1995).
The fate of the daughter cells is intrinsically determined, as the mitotic cell is polarized in
terms of distribution of cell fate determinant molecules, such as Numb, Notch1 (Chenn
and McConnell, 1995; Shen et al., 2002; Rasin et al., 2007). Inhibition of proliferation evidently disrupts ventricle formation in zebrafish and mouse (Lowery and Sive, 2005;
Camarero et al., 2006).
It is evident that proliferation of the neuronal progenitors necessitates coordinate nuclear migration within the ventricular zone. The position of the nucleus correlates with progression of the cell cycle (Ueno et al., 2006). In particular, the cells retract their bipolar processes and round up the ventricular surface during mitosis (Takahashi et al.,
1993). Therefore, interkinetic nuclear migration of the neuronal progenitor depends on the dynamic organization of the cytoskeleton and requires maintenance of cellular polarity that involves junctional components (Chenn et al., 1998; Tsai and Gleeson,
2005). Remarkably, distribution of centrosomes is polarized, presented as supernumerary centrosome demarcating the ventricular surface of the neuroepithelia (Chenn et al., 1998).
8.2.2. Radial migration
Neuronal migration in the cerebral cortex begins when the first cohort of postmitotic neurons leaves the germinal ventricular zone (VZ) to form the preplate (PP), which is a transient layer lining the outer surface of the cortex (Figure 1.8.B). Preplate is later split by the arrival of accumulating cortical plate neurons into the superficial
76 marginal zone and the deeper subplate (Figure 1.8.C). Two modes of radial migration
have been identified in corticogenesis in mouse, namely, somal translocation and glia-
guided locomotion (Nadarajah and Parnavelas, 2002).
Somal translocation is the predominant mode of movement during early corticogenesis, when the cortical anlage is small. The cells that undergo somal translocation typically have a long, radially oriented basal process terminating at the pial surface, and a short, transient trailing process. The migratory behavior of translocating cells is also distinctive, being characterized by continuous advancement that results in a faster rate of migration, compared with a radial-guided migrating cell (Nadarajah et al.,
2001). It has been proposed that translocation involves three dynamic steps typical in cell
movement: (1) extension of the leading process that explores the immediate environment
for attractive or repulsive cues, (2) 'nucleokinesis', or the movement of the nucleus into
the leading process, and (3) retraction of the trailing processes (Nadarajah and Parnavelas,
2002). However, cell adhesion mechanisms mediating translocation of neuronal progenitors remain largely unknown.
Glia-guided migration is more prevalent at later stages, when the cerebral wall is considerably thicker (Hatten, 2002). Cells that adopt glia-guided locomotion have a
shorter radial process that is not attached to the pial surface. As the soma moves forward,
the leading process with a growth-cone-like tip maintains its length. These cells show a
characteristically slow saltatory pattern of locomotion (Nadarajah et al., 2001). Integrity
of gap junction is essential in maintaining this neuron-glia interaction. Acute down-
regulation of gap junction proteins, such as connexin 26 and connexin 43, impairs the
migration of neurons to the cortical plate (Elias et al., 2007). Also, the interactions
77 between the ECM protein reelin and integrin α3β1 mediate neuronal adhesion to radial glial fibers and radial migration (D'Arcangelo et al., 1995; Dulabon et al., 2000).
Interestingly, in the reeler mouse (a spontaneous mutant in reelin), the development of the preplate appears normal (D'Arcangelo et al., 1995). The fact that mutations affecting glia-guided migration do not severely affect the early formation of the preplate (which appears to rely on somal translocation) agrees with the hypothesis that the molecular mechanisms underlying somal translocation and glia-guided locomotion are different.
Existence of two distinct modes of radial migration is necessary in the higher vertebrate which have a more advanced and complicated cortex (Marin and Rubenstein, 2003).
8.2.3. Tangential migration
Tangential migration is a process in which cells migrate orthogonal to the direction of radial migration. This migration behavior appears to be glia-independent, and comprises distinct types of cell movement that diverge primarily in the type of substrate used by migrating cells. In some cases, groups of neurons migrate using each other to promote their migration, as in the case of olfactory bulb interneuron precursors. In other cases, tangentially migrating neurons follow growing axons to reach their destination
(Marin and Rubenstein, 2003). Tangentially migrating elements include pioneer neurons and interneurons (both of subpallial origin) and Cajal-Retzius cells mostly of pallial origin (Nadarajah and Parnavelas, 2002). Tangential migration of interneurons in the mouse neocortex is one of the best studied processes. Interneurons that arise in the ventral forebrain migrate dorsally into different telencephalic structures (Marin and
Rubenstein, 2001).
78 Guidance of tangentially migrating interneurons involves the coordination of multiple guidance cues. The general direction of interneuron migration (ventral to dorsal) appears to be established by the simultaneous activity of chemorepulsive and chemo- attractive factors produced by the preoptic area and the cortex, respectively (Hatten,
2002). One of these guidance cues is mediated by semaphorins and neuropilin receptors on the neuron. In response to a chemorepulsive signal composed in part of sema3A and sema3F, migrating interneurons expressing neuropilins are sorted to the cortex. Loss of neuropilin function is known to decrease the number of neurons reaching the embryonic cortex (Marin et al., 2001). In the zebrafish hindbrain, branchiomotor neurons that migrate caudally are found to utilize the trangential mode of migration (Bingham et al.,
2002).
79 9. IDENTIFICATION OF ZEBRAFISH L1 HOMOLOGS
The current study employs a zebrafish model to elucidate the functions of L1
during early embryonic stages. Two zebrafish L1 homologs, L1.1 and L1.2, have been
identified. Two partial sequences showing similarity to the 3’ portion of the mammalian
L1 molecules were obtained by screening an embryonic cDNA library (Tongiorgi et al.,
1995). L1.1 and L1.2 contain most of the conserved features found in the cytoplasmic domain of other L1 proteins. For example, both L1.1 and L1.2 contain the YRSLE motif for AP-2 adapter binding, suggesting that their activities may be regulated by endocytosis.
Nonetheless, L1.1, but not L1.2, contains the consensus binding sequence for ankyrin and doublecortin, implicating its capability of modulating cytoskeleton dynamics and neuritogenic activity.
In situ hybridization using 3’-end sequences of L1.1 has revealed that L1.1
expression is first detected in premature brain regions at 16 hpf and it becomes prominent
in all classes of neurons at later stages, while L1.2 has a more restricted expression
pattern, present only in a subpopulation of neuronal cells after 22 hpf (Tongiorgi et al.,
1995).
Recent studies on L1.1 and L1.2 have focused on their involvement in axon
regeneration in adult fish. It is suggested that both proteins may participate in axonal
regeneration, but in rather different processes. After caudal spinal cord transection in the
adult fish, expression of L1.1 mRNA is greatly increased in the nuclei of the medial
longitudinal fascicle, whereas L1.2 mRNA is significantly upregulated in the glia cells in
the spinal cord. On the other hand, the hindbrain nuclei show enhanced expression of
L1.1, but not L1.2, after a proximal lesion (Becker et al., 1998). An implied role for L1 in
80 axonal regeneration is further evidenced in the mouse after spinal cord axotomy. Over- expression of exogenous L1 by adeno-associated viral infection in treated mice enhances axon regeneration and reinnervation and, importantly, improved stepping abilities and muscle coordination during ground locomotion (Chen et al., 2007).
L1.1 has also been implicated in the process of memory consolidation. In the fish trained to avoid mild electric shocks, upregulation of L1.1 is detected in the tectum soon after acquisition of training, whereas the level of L1.2 mRNA seems unchanged (Pradel et al., 2000). Consistently, injection of anti-L1.1 antibody decreases retention of the memory of the trained fish, indicating the pivotal function of L1.1 in long-term memory in zebrafish (Pradel et al., 1999; Pradel et al., 2000).
81 10. HYPOTHESIS AND RATIONALE OF THE THESIS
Clearly, the development of the nervous system requires precise positioning of the
neurons as well as proper neuronal projection and connectivity. L1 plays key roles in
these diverse processes. The importance of L1 to the early development of the nervous
system is further underscored by the implication of mutations in the L1 gene in the
congenital CRASH syndrome. Pathogenic missense L1 mutations have been found to
occur frequently at highly conserved residues among vertebrate L1 molecules, suggesting crucial contributions of these amino acids to the functions of L1. Replacement of these
amino acids in the extracellular domains may either induce conformational instability or
alter adhesive interactions with its binding partner(s), consequently leading to abnormalities in neural development.
There are aspects of pathological mechanisms of L1 mutations which have not been defined by studies on L1 mouse models. In particular, correlation between the pathogenic mutations and hydrocephalus remains to be established. Therefore, it demands
exploration of alternative animal models. Taking advantage of the zebrafish model in development, my research project aims (1) to elicit the involvement of zebrafish L1 homologs L1.1 and L1.2 during early neural development, (2) to identify specific L1- interactions that mediate neuronal processes in the context of the developing embryo, and
(3) to elucidate the pathogenic mechanisms in which the responsible missense mutations may lead to abnormalities in the developing nervous system.
The functions of zebrafish L1.1 and L1.2 during early embryonic development were examined by knocking down the specific proteins by morpholino injection into the zebrafish embryo. In chapter II, notable abnormalities in early brain development were
82 observed in L1.1 morphants, but not in L1.2 morphants. A phenotypic enlargement of the
fourth ventricle prominent in L1.1 morphants was associated with an irregular shaped
brain. Further characterizations suggested that L1 plays an important role in regulating
periventricular polarity and cell proliferation during normal brain ventricle development.
Establishment of ventricular polarity and precise neuronal positioning in the
developing brain require proper cell-cell adhesion and cell-matrix interactions. In chapter
III, L1-neuropilin interactions were examined during ventricle formation. In particular,
L1.1s, a novel isoform of L1.1 that consists of the first four immunoglobulin domains of
the full length molecule, was discovered and shown to be both necessary and sufficient to
support radial neuronal migration away from the ventricle. L1.1s mRNA was capable of rescuing the L1.1 morphant phenotypes in the brain, whereas the mutation L123V located in the binding site of L1-Nrp1 compromised the rescuing function of L1.1s. The importance of the L1-neuropilin interaction in the early morphogenesis of the brain was
further examined by in vivo treatment of the embryos using a synthetic peptide against
L1-Nrp1 interaction, which resulted in abnormal cell migration from the ventricular zone
and destruction of ventricular polarity.
The final project was designed to examine how L1 influences axon projections
into their muscle targets in the periphery. In chapter IV, the importance of L1 to primary motoneuron development was assessed. The loss of L1.1 expression led to ectopic branching of axons from CaP. A novel interaction between L1.1 and unplugged (a zebrafish homolog of MuSK) was implicated in the process of axonal growth. Pathogenic
mutations in the Ig3 domain disrupted this specific interaction and led to abnormal axonal projections, suggesting that Ig3 domain may possess a critical binding site for MuSK.
83
Chapter II:
Knockdown of L1.1 Expression in Zebrafish
Results in Hydrocephalus-like Phenotype and Reveals a
Role in Ventricular Polarization during Brain
Development
(All experimental results described in this chapter were obtained by me.)
84 SUMMARY
The neural cell adhesion molecule L1 plays an important role during embryonic brain development. Mutations in the human L1 gene are responsible for a wide spectrum of X-linked neurological disorders including hydrocephalus or dilated lateral ventricles.
However, the pathological mechanisms associated with L1 mutations remains to be elucidated. Zebrafish expresses two L1 homologues, L1.1 and L1.2. When L1.1 was knocked down by morpholino injection, a phenotype with an enlarged fourth ventricle was observed in zebrafish embryos. Morphological characterization revealed that, in association with the enlargement of the fourth ventricle, both diencephalic and mesencephalic ventricles were obstructed and exhibited irregular shapes in the morphants.
This abnormal construction of the brain could be attributed to aberrant polarization of the periventricular neuroepithelia prior to inflation of the compromised ventricles by cerebrospinal fluid. Furthermore, disorganized brain structures were evident at the mid- hindbrain boundary and the diencephalon-mesencephalon division, demarcated respectively by the expression of pax2.1 and pax6. Cell migration traced by BrdU labeling during the early stage of brain ventricle formation revealed abnormal organization in the periventricular zones and positioning of new born cells in the absence of L1.1. Concomitantly, cell proliferation in these regions was inhibited. These results provided in vivo evidence in support of a role for L1 in the regulation of periventricular polarity and cell proliferation during brain ventricle formation, which may underlie the pathological development of X-linked hydrocephalus.
85 INTRODUCTION
The neural cell adhesion molecule L1 is a member of immunoglobulin (Ig)
superfamily. L1 and L1-related proteins have been detected in a wide spectrum of animal
species and they contain six Ig-like domains and five fibronectin (Fn) type III-like repeats
in the extracellular region, a single span transmembrane domain and a highly conserved
cytoplasmic tail (Hortsch, 1996; Maness and Schachner, 2007). L1 plays an important
role in many neurobiological processes, including neural cell survival (Haney et al., 1999;
Chen et al., 2005; Chen et al., 2007) and migration (Lindner et al., 1983), neurite
outgrowth (Lemmon et al., 1989; Dong et al., 2003), axon fasciculation (Honig et al.,
1998; Ohyama et al., 2004; Wiencken-Barger et al., 2004) and myelination (Wood et al.,
1990; Itoh et al., 2005).
In human, L1 is expressed in post-mitotic neurons of the central and peripheral neural systems, on pre- or nonmyelinating Schwann cells of the peripheral system, and on glial cells. It is enriched in the growth cone and in the endocytic vesicles along axons
(Joosten and Gribnau, 1989; Minana et al., 2001; Wisco et al., 2003; Chang et al., 2006).
Mutations in the human L1 gene result in a broad spectrum of X-linked neurological disorders, manifesting corpus callosum hypoplasia, retardation, adducted thumbs, spastic
paraplegia and hydrocephalus, collectively known as CRASH or L1 syndrome (Fransen
et al., 1997; Fransen et al., 1998b; Weller and Gartner, 2001). To date, more than 140
mutations associated with CRASH syndrome have been identified in the L1 gene (Finckh
et al., 2000; Sztriha et al., 2002; Silan et al., 2005). These mutations including missense,
86 nonsense and frameshift, spread across the entire L1 coding region, implicating that all its protein domains are involved in one or more L1 functions (Weller and Gartner, 2001).
CRASH patients are commonly diagnosed with abnormal development of the major axonal tracts, including corticospinal tract and corpus callosum (Yamasaki et al.,
1997). Severe hydrocephalus is found predominantly in patients carrying mutations that affect protein conformation or key binding sites and those that cause premature protein termination in one of its extracellular domains, whereas no hydrocephalus or only a slight ventricular dilation is detected in patients carrying mutations in the cytoplasmic domain
(Fransen et al., 1998a; Kamiguchi et al., 1998a).
Mice that produce mutant L1 truncated in the extracellular Ig3 domain (Dahme et al., 1997) or the Ig6 domain (Cohen et al., 1998) display some of the human CRASH phenotypes, including failure in axonal pathfinding by corticospinal axons while crossing of the midline, vermis hypoplasia, impaired learning and hydrocephalus (Dahme et al.,
1997; Cohen et al., 1998; Fransen et al., 1998a; Demyanenko et al., 1999; Rolf et al.,
2001). The agenesis of corticospinal tracts observed in mouse and man are thought to result from the degeneration of neurons that fail to synapse with correct targets (Cohen et al., 1998; Kamiguchi et al., 1998b). The malformation of corticospinal tracts is one of the proposed mechanisms that lead to the enlargement of the lateral ventricles (Fransen et al.,
1998a; Demyanenko et al., 1999). However, the severity of hydrocephalus is correlated strongly to the genetic background of the mouse (Dahme et al., 1997; Kamiguchi et al.,
1998a). Low incidence of hydrocephalus has been reported in these mouse models (Rolf et al., 2001). The pathogenesis of the hydrocephalus phenotype remains largely unknown.
87 In this report, zebrafish was used as a model to investigate the role of L1 during neural development. Partial cDNA clones of two L1 homologues, L1.1 and L1.2, have been reported in zebrafish (Tongiorgi et al., 1995). Complementary DNA sequences containing the entire coding regions of L1.1 and L1.2 were cloned in current study. When
L1.1 expression was knocked down by antisense morpholinos, embryos exhibited a spectrum of phenotypes shared by CRASH patients, including underdevelopment of the forebrain and midbrain and enlargement of the fourth ventricle. Morphological characterizations have revealed an obstructed ventricular system. The abnormal organization of brain domains and the enlargement of the fourth ventricle in L1.1 morphants can be attributed to defective polarization of the neuroepithelium and impaired cell proliferation.
88 MATERIALS AND METHODS
Zebrafish maintenance
Wild-type zebrafish (Danio rerio) were purchased from local pet stores. Fish were
maintained on a 14 hour light/10 hour dark cycle at 28.5°C according to Westerfield
(2000). Embryos were obtained by spontaneous spawning and staged by hours post
fertilization (hpf) at 28.5°C and number of somites (Kimmel et al., 1995).
Cloning of cDNA encoding zebrafish L1
RNA was isolated from embryos 32 hours after fertilization using the Trizol reagent
(Invitrogen, Burlington, Ontario). Complementary DNA was synthesized using
SuperScript II reverse transcriptase (Invitrogen). 5’ rapid amplification of complementary
DNA ends (RACE) was performed to obtain the 5’ portions of zebrafish L1.1 and L1.2
molecules according to manufacturer’s instruction. Specific primers were designed based on the partial sequences of zebrafish L1.1 (Genbank Accession: X89204, L1.1sp1,
GAGTCTAGAATCTCCCAGTGTAT; L1.1sp2, TCTGTTGCTGTCTTCTTCACTCA) and L1.2 (Genbank Accession: X89205, L1.2sp1, GGCTGACTGGCTGTACGCTTCT;
L1.2sp2, CAGCTCTCGGTTACAAACCCACT, Tongiorgi et al., 1995). All cDNA were cloned and sequenced (ACTG, Toronto, Ontario).
Preparation of His6-tagged fusion proteins
Complementary DNA sequences encoding extracellular domains, including the 6 Ig
domains and 5 Fn-type III domains, were obtained by PCR using VentR polymerase
89 (New England BioLab, Pickering, Ontario). Amplification Primers are: 5’-
TACCATGGGCTACATTCAGATCCCACAC-3’ and 5’-
ATACTCGAGTTCGGTCGCAAAGTTCCT-3’ for L1.1, and 5’-
TACCATGGGCCTAAAACCCGGGAACACC-3’ and 5’-
CACTCGAGCTCGGTTACAAACCCACT-3’ for L1.2. The cDNA fragments were
inserted into pET-28(a) vector (Novagene, Madison, WI) at NcoI and XhoI sites.
Plasmids were then transformed into E. coli BL21 (DE3) (Novagene). For L1.2 fusion
protein, bacteria were co-transformed with the pRARE plasmid to generate rare tRNAs
required for full-length protein synthesis.
Transformants were induced by 1 mM IPTG for 3 hours at 37°C, and lysed in the
presence of 1 mg/ml lysozyme, 25 U/ml DNase, 1 mM PMSF and 1x protease inhibitor
cocktail (Sigma, Oakville, Ontario). Fusion proteins were enriched in inclusion bodies.
Inclusion bodies were collected and resuspended in a denaturing buffer containing 8 M
urea and 20 μM β-mercaptoethanol. Fusion proteins were purified by binding to Ni-NTA
agarose beads (Qiagen, Mississauga, Ontario). For refolding, excess oxidized glutathione
(32.4 mg) was first added to 2.4 ml of elution product to prevent disulfide bond formation between cysteine residues. After 30 minutes of incubation, this mixture was dialyzed against 100 ml of urea buffer (0.1 mM NaH2PO4, 0.01 M Tris, 8 M urea, 20% glycerol,
1% TritonX-100, 0.3% N-lauroylsarcosine, pH 8.0) for 30 min at room temperature. The
urea in the dialysis buffer was slowly diluted overnight with 400 ml buffer minus the urea
to a final concentration of 1.6 M urea. After refolding, fusion proteins were dialyzed by
two changes of phosphate buffered saline (PBS). Formation of disulfide bond between
proper cysteine residues was catalyzed by the addition of 1 mM of GSH (reduced
90 glutathione) and 0.2 mM GSSG (oxidized glutathione), followed by dialysis against PBS.
Integrity of fusion proteins was verified by sodium dodecylsulfate-polyacrylamide gel electrophoresis (SDS-PAGE) under reducing and nonreducing conditions.
Production of polyclonal anti-L1.1 and L1.2 antibodies
Rabbits were immunized by injection of 0.5 mg of the purified fusion protein containing the extracellular portion of either L1.1 or L1.2, followed by four boosts of 0.2 mg of protein each at intervals of 2-4 weeks. Immune IgG was purified using refolded bacterial fusion protein according to Koelle and Horvitz (1996). Briefly, refolded fusion protein was applied on a nitrocellulose membrane (Amersham, Baie d’Urfe, Quebec) at ~0.1 mg/cm3. The membrane was blocked with 5% bovine serum albumin (BSA) in TBS for 1 hour at room temperature, followed by two washes with TBS. The crude serum was diluted 1:5 in TBS and incubated with the membrane at 4°C overnight. The membrane was sequentially washed three times with TBS and two times with PBS. The bound antibodies were eluted from the membrane using 0.1 M glycine (pH 2.5). The eluted antibody solution was then neutralized with 1/10 volume of 1M Tris (pH 8.0) and stored at -80ºC. Antibody specificity was confirmed by western blott analyses.
Neuritogenesis assay
Spinal neurons derived from 18 hpf zebrafish embryos were cultured according to
Anderson (2001). Cells (5 x 104 cells) were dissociated from the trunk region and seeded on 12-mm diameter coverslips coated with His-tagged recombinant protein (80 μl of 1
91 μM protein per coverslip). Cells were cultured for 20 hours at 28°C in Leibovitz L-15
medium (Invitrogen) supplemented with 1% N-2 supplement (Invitrogen).
Spinal neurons bearing neurites were identified by immunostaining of acetylated-
tubulin. Cells were fixed with 4% paraformaldehyde (PFA) for 10 minutes. After
blocking with 2% BSA in PBS, cells were incubated for 2 hours with mouse monoclonal
antibody against acetylated tubulin (1:500 dilution in 2% BSA to a final concentration of
2.4 μg/ml) (Sigma), followed by incubation with Alexa-488-conjugated goat anti-mouse
IgG (1:500, Invitrogen). Coverslips were mounted in Fluormount (DAKO, Mississauga,
Ontario). Samples were examined by epifluorescence microscopy and images of
acetylated-tubulin-positive neuronal cells were recorded. The longest neurite was
measured for each neuronal cell and only neurites with a length greater than two cell body width were recorded. For each substrate, ~150 neurites from 3 different coverslips were measured in each experiment. Data were collected from four independent experiments.
Morpholino oligonucleotides and mRNA injection
Morpholino antisense oligonucleotides were purchased from Gene Tools (Philomath,
OR). L1.1-MO1 (5’-CACTGGAGGCATTCTGGAGCAAACA-3’) recognized a region
flanking the initial translation start codon AUG, while L1.1-MO2 (5’-
CAGTCCCGACTCCAGACACACACAC-3’) was upstream in the 5' UTRs. Two
mismatch morpholino oligonucleotides were synthesized as controls (L1.1-MO1-mis, 5'-
CAgTGcAGGCATaCTGGAGgAAtCA-3' and L1.1-MO2-mis, 5’-CAgCTcCTG
AgAGACAgAAACAgAG-3’, where mismatched nucleotides were in lower case).
Embryos were injected with 0.5 nl of antisense morpholino oligonucleotides into the yolk
92 at the 1-2 cell stage (Nasevicius and Ekker, 2000) using the Eppendorf Transjector 5246.
The highest dose of morpholino (~2 ng per embryo at the concentration of 4 mg/ml) used in these experiments did not increase mortality significantly or cause overt non-specific necrosis.
Capped RNA of L1.1 for rescue experiments was synthesized from the restriction enzyme-linearized plasmid containing full-length L1.1 using T3 Cap Scribe (Roche,
Laval, Quebec). For injection, capped RNA was diluted to 0.1 μg/μl in the solution that contained morpholino L1.1-MO2.
Western blot analysis
Zebrafish embryos were dechorionated and deyolked before homogenization in lysis buffer (63 μM Tris-HCl, 2% TritonX-100, 5% glycerol, 1 mM DTT, 3.5% SDS) according to Westerfield (2000). Affinity-purified rabbit anti-L1.1 IgG was applied at 1
μg/ml in blocking buffer (5% dry milk in Tris-buffered saline with 0.1% Tween-20).
Secondary HRP-conjugated goat anti-rabbit IgG (1:10,000; Pierce, Ottawa, Ontario) was applied, followed by ECL detection (Amersham).
Brain imaging and ventricle visualization
Embryos of 24 hpf were fixed in 4% PFA and stained with 1 μM SYTO-16 Green
(Invitrogen) for 1 hour. To obtain dorsal and lateral brain images, embryos were orientated in 0.2% low melting point agarose for confocal microscopy (Leica TCS NT).
Image stacking was performed using ImageJ (National Institutes of Health).
93 To visualize brain ventricles, embryos at 48 hpf were anesthetized with 0.1 mg/ml of Tricaine (Sigma) dissolved in embryo medium prior to injection and imaging. The hindbrain ventricle was micro-injected with ~5 nl of Texas Red-conjugated Dextran (5% in 0.2 M KCl; Invitrogen). Micrographs were taken within 5 minutes following injection.
Areas of ventricles filled with the dye were estimated using ImageJ. Student’s t-test was performed using the statistical analysis software GraphPad Prism program version 4.
Histology and Immunohistochemistry
Embryos of 48 hpf were stained with Mayer’s hematoxylin (Sigma) following fixation in
4% PFA. Embryos were dehydrated using an ethanol series, followed by xylene, and then embedded in paraffin. Embryos were sectioned at 5 μm on a rotary microtome (Leica
Jung RM 2035) and collected on Snowcoat X-TRA slides (Surgipath, Winnipeg,
Manitoba).
Whole-mount immunostaining was performed on embryos using affinity-purified rabbit antibodies against L1.1 IgG (4 μg/ml), γ-tubulin (1.6 μg/ml; Abcam, Cambridge
MA), monoclonal antibodies against zn-12 and znp-1 (5 μg/ml; Zebrafish International
Resource Center, Eugene, OR). Alexa 568-conjugated (Invitrogen) or horseradish peroxidase (HRP)-conjugated (Pierce) secondary antibodies were used for immunostaining. To obtain fluorescence images of transverse sections, embryos were
embedded in 17% gelatin plus 10% sucrose and sections (~100-μm thick) were cut using
a scalpel blade. Sections were mounted in Fluormount and images were collected by
confocal microscopy. The enhanced 3,3'-Diaminobenzidine (DAB) color substrate
94 (Pierce) was used for non-fluorescence specimens. Embryos were mounted and
photographed under an inverted microscope (Nikon ECLIPSE TE2000).
Detection of BrdU incorporation in proliferating cells
Dechorionated embryos were incubated in embryo medium containing 10 mM 5-bromo-
2'-deoxyuridine (BrdU, Sigma) and 15% DMSO in the cold for 20 minutes. To remove unincorporated BrdU, embryos were quickly rinsed a few times in fresh embryo medium
containing 15% DMSO and incubated with 10 mM dTTP in 15% DMSO for 5 minutes in
the cold. Embryos were cultured in fresh medium at 28.5°C for different time periods
until fixation. To detect incorporated BrdU, antigen retrieval was performed by
incubation with 2 N HCl for 1 hour, followed by overnight incubation at 4°C with
primary mouse anti-BrdU antibody G3G4 (Developmental Hybridoma Bank, Iowa City,
IA), diluted to 1 μg/ml in blocking solution (2% normal goat serum and 5% BSA, 1%
DMSO and 0.1% Tween-20 in PBS) according to George-Weinstein et al. (1993). Alexa
488-conjugated goat anti-mouse IgG was applied at 1:500 dilution in blocking solution
for 1 hour at room temperature. Embryos were embedded in 17% gelatin-10% sucrose
solution and thick sections were cut manually. Sections were mounted in Fluormount for
confocal microcopy.
Proliferation assay using phosphohistone-H3 labeling
To label proliferating cells, whole embryos were probed with rabbit anti-phospho-
Histone H3.3 (Ser31, 1:200 to a final concentration of 5 μg/ml; Upstate, Temecula, CA)
overnight at 4°C, followed by incubation with Alexa 488-conjugated goat anti-Rabbit
95 IgG (1:500) for 1 hour at room temperature. Embryos were counterstained with 1 μM
SYTO-16 green for 30 minutes. The head was dissected from each embryo for flat-mount
in Fluormount. Cell proliferation was quantified within the brain region spanning from
the rostral-most forebrain to the caudal-most portion of the otic vesicle as outlined by
SYTO-16 green staining as adapted from Lowery and Sive (2005). The number of
phosphohistone-H3 (pH3) antibody-labeled cells per unit area of the image was estimated
by ImageJ. The number of labeled cells were either counted manually or estimated by the
total fluorescent pixels divided by the average pixels of individual cells. Statistical
analyses were performed using a Student’s t-test with Prism4 for Windows (GraphPad).
In situ hybridization
Antisense riboprobes specific to zebrafish Pax2.1 and Pax6 were produced by RT-PCR according to Macdonald et al., 1994 and Krauss et al., 1991b, respectively. Digoxigenin
(DIG)-labeled riboprobes were synthesized by in vitro transcription using T7 polymerase
and DIG-RNA labeling mix (Roche) according to the manufacturer’s instruction.
Embryos of 24 hpf were fixed in 4% PFA overnight at 4°C. Embryos were kept in
100% methanol at -20°C until use. For in situ hybridization, embryos were rehydrated
through serial incubations (5 min each) in 75% methanol/25% PBS-Tw (phosphate
buffered saline, 0.1% Tween-20), 50% methanol/50% PBS-Tw, and 25% methanol/75%
PBS-Tw solutions. Embryos were permeabilized with 10 μg/ml proteinase K for 5
minutes at room temperature and refixed in 4% PFA for 20 minutes (Westerfield, 2000).
After prehybridization in the hybridization buffer (50% formamide, 5 x SSC, 0.1%
Tween-20, 50 μg/ml heparin, 500 μg/ml salmon semen tRNA, 92 mM citric acid),
96 embryos were hybridized with 100 ng of DIG-labeled antisense (1:250 in hybridization buffer) overnight at 65°C. Post-hybridization washes were performed with 0.2 x SSC at
65°C for 20 min, followed with two washes with 0.1 x SSC at 65°C for 20 min each. For antibody detection, embryos were incubated overnight at 4°C with alkaline phosphatase- conjugated anti-DIG antibody (1:5000 dilution) in blocking buffer (5% sheep serum and
5% BSA in PBS-Tw). Color detection was performed using the Nitro blue tetrazolium chloride/ 5-Bromo-4-chloro-3-indolyl phosphate (NTP/BCIP) substrate (Roche) in the presence of 0.5 mg/ml levamisole (Sigma). Embryos were washed extensively in PBS-
Tw and mounted in 70% glycerol for microscopy. Manual sections of embryos were cut using a scalpel blade.
97 RESULTS
Cloning of zebrafish L1 cDNAs
The 5’-region of complementary DNA fragments for L1.1 and L1.2 were amplified by 5’ RACE. The cDNA sequences containing the complete coding regions of
L1.1 (4052 bp) and L1.2 (4112 bp) were obtained and sequenced (Genbank accession numbers AY376855 and AY376856). The L1.1 and L1.2 cDNAs contain open reading frames encoding 1269 and 1271 amino acids, respectively. Similar to most L1 species, they are organized into six Ig-like domains, five fibronectin type-III repeats, a single transmembrane domain and a short cytoplasmic tail. The deduced amino acid sequences of L1.1 and L1.2 share 48% sequence identities with each other, and they displayed 40% and 41% sequence identity, respectively, with human L1. The RSLE sequence, encoded by exon 27 in human and Fugu (Coutelle et al., 1998), is present in both homologues.
The sequence encoded by exon 2 in human, YEGHHV, can be aligned with the sequences YTYNKL in L1.1 and YKIRDL in L1.2. Most pathological missense mutations in human L1 affect conserved residues. Among the fifty-four amino acids where reported missense mutations occur, 24 are identical among L1.1, L1.2 and human
L1, 5 are highly conserved, and another 5 are less conserved (Figure 2.1).
Neuritogenic activity of L1.1 and L1.2
In order to determine whether zebrafish L1.1 and L1.2 are capable of stimulating neurite outgrowth like other L1 species, His6-tagged fusion proteins containing the extracellular regions of L1.1 or L1.2 were purified in E.coli and applied as substrates for
98 V
Ig1 VS
Ig2 S W/Q Y Q T L
Ig3 D K R R/C
Ig4 R P I D D R
Ig5 C P N
Ig6 N
Fn1 T/D R
Fn2 PM I/F C
Fn3
Fn4
Fn5 L
L
TM Cytoplasmic tail L E
99 Figure 2.1. Alignment of zebrafish L1.1 and L1.2 with human L1. Deduced amino acid sequences of L1.1 (Genbank accession: AY376855) and L1.2 (Genbank accession: AY376856) were aligned with the human L1 sequence (Genbank accession: M77640) by Clustal X. The two zebrafish L1 homologues share similar domain composition with the human orthologue, containing six Ig-like domains, five fibronectin type-III (Fn) repeats, a single transmembrane domain (TM) and a cytoplasmic tail. The conserved cysteine residues in each Ig domains are highlighted in blue and the tryptophan and tyrosine residues in the Fn repeats are highlighted in green. The conserved neural-exclusive sequences generated by exons 2 and 27 are highlighted in yellow. The amino acids that are substituted in CRASH syndrome are boxed, with completely identical residues shown in red, highly conserved residues in green and less conserved residues in purple. The corresponding pathogenic amino acid substitutions are indicated on top of each box.
100 culturing embryonic primary spinal neurons isolated from embryos at the 18-somite stage
(Figure 2.2). Post-mitotic neurons and their neurites were fixed after 20 hours of culture and visualized by immunostaining with a monoclonal antibody raised against acetylated-
α tubulin (Figure 2.2.B). Both L1.1 and L1.2 promoted neurite outgrowth from the spinal neurons. Quantitative analysis showed that 50% of the neurites on the L1.1 substrate extended more than 90 μm, with a mean length of 114 μm. Similarly, 50% of the neurites on L1.2 were longer than 90 μm with a mean neurite length of 116 μm. In contrast, neurites extending from cells cultured on 1% BSA had a mean neurite length of 42 μm
(Figure 2.2.C and D).
Expression of L1.1 in the brain
To investigate the role of L1.1 during zebrafish development, the expression pattern of L1.1 was first examined as a critical prerequisite to functions in the developing neural system. Whole-mount immunohistochemistry revealed that L1.1 was expressed in all classes of neurons in the brain at 24 hpf (Figure 2.3.A), including neurons of the epiphyseal cluster, dorso-rostral cluster and ventro-rostral cluster, trigeminal ganglion neurons, posterior lateral line ganglion and anterior lateral line ganglia, as well as their trajectories. Intensive staining of L1.1 was observed at the medial longitudinal fasciculus and dorsal longitudinal fasciculus. Interestingly, L1.1 expression was also detected on the ventricular surfaces, the anterior of the isthmic fold and the cerebellum, as shown in the dorsal view of the midbrain and mid-hindbrain boundary regions (Figure 2.3.B). In transverse sections, L1.1 was localized to the luminal surface of the neuroepithelium of
101 A abL1.1EC L1.2EC Signal Ig-like domains Fn III-like repeats cyto peptide 130 100 Nascent L1 73 54 L1.1EC 40 L1.2EC 35 B a b c
BSA L1.1EC L1.2EC
C D m) 100 μ 140 zL1.1 EC 80 zL1.2 EC 1% BSA 100 60 40 60
neurites >X axis >X neurites 20 20
Percentage of cells with with cells of Percentage 0
0 100 200 300 ( neurites of length Mean 1%BSA L1.1EC L1.2EC Neurite length (μm) 400
Figure 2.2. Neuritogenic activity of recombinant L1.1 and L1.2. (A) His6-tagged L1.1EC and L1.2EC fusion proteins were produced in E.coli. (a) Schematic diagram of the L1.1EC and L1.2EC constructs, which contained the extracellular portion of mature proteins L1.1 and L1.2, respectively. (b) Coomassie blue-stained profiles of the purified and refolded L1.1EC and L1.2EC proteins. (B) Embryonic spinal neurons isolated from 18-hpf embryos were cultured on different protein substrates for 20 hours and then fixed for staining with an anti-acetylated tubulin antibody. Fluorescence images of neurons cultured on (a) BSA, (b) His6-tagged L1.1EC and (c) His6-tagged L1.2EC were recorded for neurite measurements. Scale bars: 50 μm. (C) Accumulative plots of neurites extending from neurons cultured on BSA (○), L1.1EC (■), or L1.2EC (∆). (D) Mean neurite lengths projecting from neurons cultured on different substrates. Data represent the mean ± S.E.M. of four experiments.
102 A MHB B
MLF THD M M H ec ec VIII PLL V MHB
CDE
4 V
ot eye eye MLF MLF DVDT TPOC TPOC
Figure 2.3. Expression of L1.1 in the embryonic brain. Embryos at 24 hpf were subjected to whole mount immunohistochemistry using immuno-purified rabbit anti- L1.1 antibodies, followed by Alexa 568 anti-rabbit IgG (pseudocolored in purple). SYTO-16 Green was used as counterstain to reveal the brain structure. (A, B) Dorsal views of the brain with rostral to the left. Confocal stacks of the brain were superimposed in (A). Image shown in (B) was taken from a section 17 μm ventral from the dorsal longitudinal fasciculus. White arrow heads indicate the position of mid-hindbrain boundary (MHB), while the open arrow denotes the medial longitudinal fasciculus (MLF). D denotes diencephalons; ec, neurons of the epiphyseal cluster; H, hindbrain; M, mesencephalon; PLL, posterior lateral line ganglion; T, telencephalon; V, trigeminal ganglion neurons; VIII, acoustic/anterior lateral line ganglia. Transverse sections of the embryonic brain were also taken at the midbrain (C), mid-hindbrain boundary (D) and hindbrain (E) regions. DVDT, dorsal- ventral diencephalic tract; TPOC, tract of post-optic commissure; ot, otic vesicle; and 4V, fourth ventricle. L1 expression was associated with the ventricular surface in different brain regions (arrows in B). Scale bars: 50 μm.
103 the ventricles (panels C-E of Figure 2.3). Embryos stained with the pre-immune serum did not reveal any immunoreactive staining in the brain (data not shown).
Phenotype of L1.1-knockdown embryos
To investigate the role of L1.1 in brain development, L1.1 expression was inhibited by the injection of L1.1 morpholino antisense oligonucleotides into embryos.
All most identical results were obtained with both L1.1MO1 and L1.1MO2. Western blot analysis of pooled embryos showed that injection of 2 ng morpholino per embryo resulted in 95% reduction of L1.1 expression at 24 hpf and 90% at 48 hpf (Figure 2.4.B), while lower amounts of morpholino (0.5 ng and 1 ng) reduced L1.1 expression by ~90 % at 24 hpf (data not shown). Since injection of 2 ng per embryo did not cause significant developmental delay or necrosis, this amount was used for most of the subsequent experiments.
Phenotypes of varying degree of severity were observed and they were classified as wild-type-like, class I and class II phenotypes (Figure 2.4.A). Class I morphants had a smaller head, curved body trunk, and reduced motility. Class II morphants showed a more severe phenotype, manifesting a shorter and more crooked body trunk. Most of the morphants in this class were unable to move away when they were pricked by a pin.
Injection of morpholino between 0.5 ng and 2 ng per embryo consistently yielded ~60% class I morphants and ~20% class II morphants by 24 hpf (Figure 2.4.C). Deleterious effects were not observed in uninjected embryos or controls injected with the mismatch morpholino. Further characterizations were focused on class I morphants, which were able to survive various experimental treatments yet preserve the integrities and
104 A 24 hpf 48 hpf ab Uninjected
cd Ctrl MO
fg L1.1MO Class I Class L1.1MO hi L1.1MO Class II Class L1.1MO j k
0.5 mm Rescued
BC 24 h 48 h 100 Wild type-like Ctrl L1.1 Ctrl L1.1 80 Class I kDa MO MO MO MO Rescued 60 Class II 182 40 n = 76 n = 226 n = n = 86 n = n = 107 n = 116 Embryos
% 20 82 64 0 Blot: anti-L1.1 0.5 ng 1 ng 2 ng 2 ng 4mis L1.1MO MO
D a bcd
Ctrl L1.1MO L1.1MO Rescued 50 μm
105 Figure 2.4. Knockdown of L1.1 resulted in abnormal embryonic development. (A) Embryos were given differently treatments at the 1 to 2-cell stage: without morpholino injection (uninjected), injected with mismatch control morpholino (ctrl MO), L1.1 morpholino (L1.1MO) or a combination of L1.1 morpholino and L1.1 mRNA (rescued). Micrographs of live embryos were taken at 24 hpf (a, c, f, h and j) and 48 hpf (b, d, g, i and k) of development. At 48 hpf, ~55% of morphants (85/152, based on 4 independent experiments) showed enlarged 4th ventricle in the hindbrain (arrowheads in g and i). (B) Protein blots of lysates derived from control uninjected embryos, L1.1 morphants injected with 2 ng L1.1 morpholino, and rescued embryo co-injected with in vitro synthesized L1.1 mRNA, were probed with affinity-purified rabbit anti-L1.1 antibodies. (C) Quantification of embryos in each phenotype class at 24 hpf. (D) Aberrant neuronal projections from trigeminal ganglion neurons due to the loss of L1.1 expression. Trajectories were stained with mAb zn-12 (Metcalfe et al., 1990) and the images are shown with dorsal to the top and caudal to the right.
106 morphologies of the brain.
L1.1 morphants exhibited notable defects in brain structure at 24 hpf, including
obscure mid-hindbrain boundary, flattened or smaller forebrain and midbrain (Figure
2.4.A). Underdevelopment of the forebrain and midbrain persisted through 48 hpf (Figure
2.4.A). Interestingly, ~55% of L1.1 morphants of this stage showed egregious
enlargement of the fourth ventricle (Figure 2.4.A). These defects were rescued when
embryos were co-injected with L1.1 mRNA and L1.1MO2, which recognizes the 5’UTR
of the endogenous L1.1 transcripts. Western blot analysis confirmed the expression of
L1.1 in the rescued embryo.
On the other hand, injection of L1.2 morpholino resulted in relatively mild
phenotypes in a small population of injected embryos (Figure 2.5.B). In addition, none of
the L1.2 morphants manifested enlarged 4th ventricle.
L1 has been reported to play a pivotal role in axonal growth and guidance and malfunction of L1 causes abnormal axonal development in both humans and mice (Walsh and Doherty, 1997; Maness and Schachner, 2007), In order to assess the effects of L1.1 deficiency on neuronal trajectories in morphants, the mAb zn-12 was used to label sensory neurons (Metcalfe et al., 1990). At 24 hpf, trigeminal ganglian neurons in the control hindbrain projected an extensive axonal network into the periphery. In the morphant brain, although a similar number of neurons were observed in the trigeminal ganglion, there were fewer and shorter processes projecting from these neurons and forming a much less extensive axonal arbor (Figure 2.4.D, panel b). More severe defects were found in morphants manifesting the class II phenotype (Figure 2.4.D, panel c).
These defects were rescued by the co-injection of L1.1 mRNA.
107 in control embryos and L1.2morphants. in controlembryos (C) Western intheembryo. additional defects Injectionsof abnormalities. phenotypic wher L1.2 morpholinoatthisconcentration, de embryonic exhibitednormal 70% ofembryos (L1.2MO).(B) Morethan morpholino L1.2 2ng with injected hpf and(b)embryo developmental defects. Morpholinoknockdown ofzebr 2.5. Figure A C B
% of embryos 24 hpf (A) Lightmicrographsof(a 100 a kDa 116 182 20 40 60 80 49 64 82 0 Blot: anti-L1.2 L1.1MO (2 ng) Ctrl n = 226 L1.2 MO L1.2MO (2 ng)
both L1.1andL1.2morpholinoscaused n = 192 Ctrl 108 L1.2MO L1.1MO (2 ng) + (2 ng) eas ~80%ofL1.1morphants displayed (2 ng) b afish L1.2 uncorrelated withdistinct L1.2uncorrelated afish
blot analysis of analysis L1.2expressionblot (arrow) n = 33 velopment by 48 hpf after injection of injection by48hpfafter velopment ) uninjected live embryo of24 liveembryo ) uninjected Class II Class I Wild type -like L1.2 MO L1.2
Anomalous development of brain ventricles in morphants
A striking feature of the L1.1 morphants was the enlargement of the 4th ventricle in the hindbrain. To assess the size of the hindbrain ventricle, a fluorescent dye conjugated with dextran was injected into the hindbrain of 48-hpf morphant and control embryos (Figure 2.6.A). The forebrain and midbrain cavities quickly became fluorescent due to the flow of cerebrospinal fluid (CSF). In the control brain, the fluorescent dye revealed a free passage within the ventricular system, connecting the fourth ventricle and the ventricles in the forebrain and midbrain. This ventricular system was symmetric to the midline of the brain (panel a” in Figure 2.6.B). However, the shape of the ventricular system was not always symmetric and the size varied broadly among the morphants
(panels b” and c” in Figure 2.6.B). Quantification of the fluorescent areas in the morphant brains suggested that the 4th ventricle, on average, was about 3 times larger than that of the control (Figure 2.6.C). Interestingly, the fluorescence-filled regions in the morphant forebrain and midbrain were relatively smaller than those in the control brain, suggesting that the enlargement of the hindbrain ventricle was accompanied by smaller ventricles in the forebrain and the midbrain. Cross sections of the morphant brain consistently showed that the ventricles in the forebrain and midbrain were smaller and irregular in shape, and often obstructed, while the arch-like structure of the tectum was obscure in the mid-hind brain boundary (Figure 2.6.D).
In the hindbrain region of the control embryo, the 4th ventricle had its anterior portion the widest and the deepest, lying between the upper rhombomere lip and the first rhombomere segment, and it narrows sharply along the rostrocaudal axis between the
109 A B 4V Ctrl L1.1MO L1.1MO M abc F 48 hpf Lateral C ) 0.25 a’ b’ c’ 2
(mm 0.20 Dorsal 0.15 a” b” c” 0.10
0.05 Dorsal FM4V
Fluorescent area Fluorescent 0.00 WT L1.1MO
D Forebrain Midbrain MHB Hindbrain c d e ab cde b a ot ot ot ot Ctrl
d’ e’ c’ a’ b’ c’ d’ e’ b’ ot a’ ot ot L1.1MO
E Midbrain Hindbrain
a DV b c 4V d 4V DV
eye SC eye SC Ctrl L1.1MO Ctrl L1.1MO
110 Figure 2.6. Anomalies in brain ventricle formation associated the enlargement of the 4th ventricle in L1.1 morphants. (A) A Scheme depicting the injection of fluorescent Texan Red-dextran into the fourth ventricle of embryos. The fourth ventricle (4V) is positioned posterior to the forebrain ventricle (F) and the mesencephalic ventricle in the midbrain (M). (B) Light micrographs of control embryos (ctrl) and L1.1 morphants (L1.1MO) showing the lateral (a-c) and dorsal (a’- c’) views of the brain with caudal to the right. The position of mid-hindbrain boundary (MHB) is bracketed and the enlarged 4th ventricle of morphants is indicated by an arrow. Fluorescent images were captured within 5 minutes of dye injection into control or morphant 4th ventricles of different sizes (white arrows in a”-c”). The positions of the forebrain and midbrain ventricles are indicated by solid arrowheads and open arrowheads, respectively. Scale bars: 0.2 mm. (C) Quantification of the fluorescent images. The areas occupied by Texas Red-dextran in the L1.1 morphants (n=9) were significantly larger than those in WT embryos (n=10), P < 0.005. (D) Histological analysis of the L1.1 morphant brain at 48 hpf by hematoxylin staining. Transverse sections were cut from various regions of the brain as shown: forebrain (a, a’), midbrain (b, b’), mid-hindbrain boundary (c, c’), hindbrain region anterior (d, d’) and posterior (e, e’) to the otic vesicle (ot). Scale bars: 0.1 mm. (E) Micrographs of midbrain (a, b) and hindbrain (c, d) sections shown at higher magnification, revealing the more ambiguous ventricular zones in L1.1 morphants (b, d). Ventricular zones are highlighted by brackets, which contain dense cells in apposition to the ventricles. Dash lines demarcate the subpial surface of the ventricles. The intermediate zone lies between the subpial surface and the ventricular zone. SC: spinal cord. Scale bars: 20 μm.
111 third and fifth rhombomeres (Trevarrow et al., 1990). The reduction of chamber space
was especially noticeable in transverse sections of the 4th ventricle anterior and posterior
to the otic vesicle, which lies laterally to the fifth rhombomere (Figure 2.6.D, panels d
and e). In contrast, the chamber space in cross sections of the 4th ventricle was
considerably larger at the corresponding region of the morphant brain (Figure 2.6.D,
panels d’ and e’). Furthermore, the rhombomeric lips in the hindbrain were flattened,
defining a wider and deeper chamber.
At higher magnification, sections of the control brain showed distinct layers, from the ventricle surface to the subpial surface, including the ventricular zone (a dense cell layer occupied by neural progenitors) and the intermediate zone (a region consists of predominantly migratory postmitotic cells, Mueller and Wullimann, 2002; Nadarajah and
Parnavelas, 2002). In the morphant brain, however, the ventricular zone was less distinguishable from the intermediate zone. Many cells, probably of ventricular origin, were dispersed across the ventricular and intermediate zones (Figure 2.6.E).
Disorganization of brain domains in morphants
The organization of the brain during embryonic development was examined by confocal scanning of the brain scaffold after staining with SYTO-16 green fluorescent
nucleus dye. In uninjected and control morpholino injected embryos of 24 hpf, intensive
fluorescent regions were observed in the tegmentum and ventral tectum in the midbrain,
and the cerebellum and rhombomeres in the hindbrain (Figure 2.7, A-A” and B-B”). it suggests that these regions were the cell dense zones, associating with active cell
proliferation (Wullimann and Knipp, 2000). In the morphant, the cell dense zones in the
112 ot, otic vesicle; Ce,cerebellum; vesicle; ot, otic aremarkedbysolidwhite a hindbrain regions openarrow,respectively. The arrowhead and and andD”).Theforebrain (A”, B”,C” scanning Green weresubjectedtoconfocal Mid-hindbrainboundary(MHB)isbr embryos. D) brainat24hpf.(A,B,Cand embryonic mo stained withSYTO-16Greentoreveal 2.7.Abnormalcelldensezo Figure
Rescued L1.1 MO Ctrl MO Uninjected D C B A MHB H, hindbrain. Scale bars:200 Scale H,hindbrain. nes intheL1.1deficientbrain. D’ C’ B’ A’ midbrain ventricles are indicated byopen indicated are midbrain ventricles 113 rphology and the cell densityzonesinthe andthecell rphology dorsally (A’, B’, C’ and D’) and laterally andD’)laterally dorsally (A’,B’,C’ Light micrographs ofthebrainliving Light micrographs cell dense zones in the midbrain and inthemidbrain densezones cell rrows. Tg, tegmentum; Te, optic tectum; tectum; Te,optic rrows. Tg,tegmentum; acketed. Brains stainedwithSYTO-16 Brains acketed. D” C” B” A” μ m. m. Te Tg Ce Embryos were H ot ot ot ot midbrain, mid-hindbrain boundary and the hindbrain were much less prominent as the brain structures in these regions were not clearly demarcated by the fluorescence signal.
The data suggest the presence of a smaller cell population and the underdevelopment of both midbrain and hindbrain. Co-injection with full-length L1.1 mRNA was able to rescue this phenotype (Figure 2.7.D-D”).
To assess the role of L1.1 in ventricle organization, embryos were examined at 24 hpf by in situ hybridization of pax2.1 and pax6. The expression of these transcription factors is regulated in a spatially restricted manner, which begins early in the developing vertebrate CNS and confers regional identity of individual brain domains (Krauss et al.,
1991a). In zebrafish, pax2.1 expression has been localized to optic stalk and the isthmus of the mid-hindbrain boundary (Krauss et al., 1992). Although pax2.1-positive cells were present in the L1.1 morphants, the ventricle within the mid-hindbrain boundary was less well defined (Figure 2.8.A, panel i). Also, instead of a more restricted distribution as in control embryos (panels c and f in Figure 2.8.A), a substantial amount of positive signal was detected laterally and dorsally across the entire mid-hindbrain boundary region of morphants, suggesting that L1.1 influences the patterning of isthmus organizers.
The expression of pax6 is restricted to the forebrain and the diencephalon- mesencephalon junction (Krauss et al., 1991b; Macdonald et al., 1994), and Pax6 is known to control L1 gene expression in the mouse forebrain (Meech et al., 1999). In
morphant embryos, the pax6-positive domain in the forebrain, was seriously attenuated
(panels g and h in Figure 2.8.B). Furthermore, expression of pax6 at the diencephalon-
mesencephalon junction was narrower along the rostrocaudal axis. The ventricle, which
114 A
abc uninjected de f Ctrl MO
gh i L1.1 MO
B
abc uninjected def Ctrl MO
ghi L1.1 MO
Figure 2.8. Expression of brain marker genes pax2.1 and pax6 in wild-type and morphant brains. Whole-mount in situ hybridization of pax2.1 (A) and pax6 (B) revealed brain organization at the mid-hindbrain boundary (arrows) and the diencephalon-mesencephalon division (brackets), respectively. In both (A) and (B), the lateral views are shown in panels a, d, g and the dorsal views are shown in panels b, e, h. Pax6-positive region in the telencephalon is attenuated in the L1.1 morphant (arrowheads in g and h of B). The transverse sections (c, f and i) show the expression of pax2.1 at the mid-hindbrain boundary (A) and pax6 at the diencephalon- mesencephalon division (B). Open arrows demarcate the position of ventricle. Scale bars: 100 μm.
115 was normally surrounded by pax6-positive cells in control embryos, appeared to be
underdeveloped in the morphant (Figure 2.8.B, panel i).
Abnormal cell polarization in the ventricular zone of morphants
The disorganization of brain ventricles in L1.1 morphants suggested that the cell
polarization in the ventricle zone was affected in the absence of L1.1. The neural
progenitor cells in the neuroepithelium apposing the ventricles are known to contain
supernumerary centrosomes, which can be highlighted by staining cells with tubulin staining (Chenn et al., 1998). These cells are normally confined to the ventricular surface,
until they exit from the cell cycle to differentiate (Chenn et al., 1998; Yang et al., 2003).
After labeling with anti-γ-tubulin antibody, intense fluorescence staining was observed in
the outermost layer of periventricular cells of control embryos, indicating the presence of
supernumerary centrosomes that defined the polarization of the ventricular zones, as
shown in control embryos in panels A-F, Figure 2.9. However, the ventricles of the
morphant brain were poorly defined by the supernumerary centrosomes. Many cells with
dense centrosome staining were found in the areas farther away from the ventricle, and
some were even located inside the ventricle (Figures 2.9.G-I). In embryos rescued by the
co-injection of L1.1 mRNA, centrosome polarity at the ventricle was restored (Figures
2.9.J-L).
Aberrant cell patterning revealed by BrdU mapping
In zebrafish, the directed migration of neurons radially from the primitive
periventricular neuroepithelium in midbrain and hindbrain is crucial to the construction
of the embryonic brain and is conserved in zebrafish (Koster and Fraser, 2001;
116 Midbrain MHB Hindbrain Inset ABC C’ Uninjected DEFF’ Ctrl MO
GHI I’ L1.1MO
JKL L’
Rescued 50μm 10μm
Figure 2.9. Abnormal ventricular development in the absence of L1.1. Whole mount immunostaining with anti-γ tubulin antibody to label centrosomes in wild-type embryos (A-C), control morpholino-injected embryos (Ctrl MO, D-F), L1.1 morpholino-injected embryos (L1.1MO, G-I) and embryos coinjected with L1.1 mRNA and L1.1 morpholino (Rescued, J-L). Sections taken at the midbrain (A, D, G, J), mid-hindbrain boundary (B, E, H, K) and hindbrain (C, F, I, L) are shown. (C’, F’, I’, L’) The boxed areas in the hindbrain are shown at higher magnification with immunostaining of γ tubulin (purple) and the nucleus (green). In L1.1 morphants, the γ tubulin-stained cells were less polarized, with many located at a distance from the ventricular surface (open arrows). Scale bars: 50 μm. Scale bar in insets: 10 μm.
117 Langenberg and Brand, 2005). To investigate how L1.1 knockdown might affect cell patterning in the developing brain, the cells that were born at 24 hpf were pulse-labeled with BrdU and their positions were mapped after 2 or 12 hours. Before the chase, BrdU- labeled cells were confined to the vicinity of the primitive ventricle in control embryos
(Figures 2.10.A, C, E). Two hours after birth, the BrdU-positive cells in the control embryo had already migrated away from the ventricular surface (Figures 2.10.A’, C’, and
E’). This organized pattern reflects the oriented migratory path of neuronal cells from the germinal layer of the ventricular neuroepithelium (Koster and Fraser, 2001; Adolf et al.,
2006). In addition, many BrdU-labeled cells were situated in the primitive mesencephalon by 12 hours post birth (Figures 2.10.A”, C”). However, in the morphant brain, the BrdU-labeled cells showed a more dispersed pattern in all the brain regions and they mingled with poorly polarized centrosomes (Figures 2.10.B, D, F). At 2 hours after birth, the BrdU-labeled cells showed a more random distribution with some even located inside the poorly defined ventricles (insets of Figures 2.10.B’, D’). The patterns of BrdU- positive cells remained to be disorganized even at 12 hours after birth, and they failed to localize to regions ventral to the mesencephalic ventricle (Figures 2.10.B” and D”).
Reduction in brain cell proliferation
The abnormal distribution of supernumerary centrosomes in the morphant brain suggested perturbation of the cell cycle in the neuroepithelial region (Brinkley, 2001;
Yang et al., 2003; Ueno et al., 2006). To assess the effects of L1.1 knockdown on the
proliferation of brain cells, mitotic cells were labeled with the antibody directed against phosphohistone-H3 (pH3) at 20, 24 and 28 hpf, corresponding to the period of rapid
118 arrows). Scalebar:50 ventral tothemesencephalic toregions localize 2hour inventricle located wereaberrantly cells athi are shown.Insetsshowtheboxedregions (green) and (A”-G”) postlabeling.Crosssec determined was localization with BrdU,andtheir 2.10.BrdUmappingFigure ofcellpatterning. BrdU pulse-labeling at 24 hpf Inset After 12 h After 2 h 0 h ”B ”D ”F” F’ E” D” E’ C” D’ B” C’ A” B’ A’ ABCDEF A’ 10 Ctrl Ctrl μ γ m -tubulin (purple, used to label supernumerar -tubulin (purple,usedtolabel ibanMdhnbanbudr Hindbrain boundary Mid-hindbrain Midbrain B’ μ 11OL.M L1.1MO L1.1MO L1.1MO m. Scalebarininsets:10 L1.1MO tions ofthebrainsimmunos C’ Ctrl Ctrl gher magnification. Arrows denote BrdU-labeled ArrowsdenoteBrdU-labeled gher magnification. 119 s after birth. BrdU-labeled cells wereunableto cells s afterbirth.BrdU-labeled ventricle ofmorphants12hourspostbirth (open ventricle Proliferating cells at 24 hpf were pulse-labeled at24hpfwerepulse-labeled cells Proliferating at0hour(A-F),2hours(A’-F’) and12hours μ m. m. D’ L1.1MO y centrosome that outlined the ventricle) theventricle) outlined y centrosome that tained with anti-BrdU antibody withanti-BrdUantibody tained Ctrl ot ot 50 μ ot m construction and growth of the zebrafish embryonic brain (Wullimann and Knipp, 2000).
The stained cells appeared to be more frequently localized to regions apposing the developing ventricles in the control embryo than the morphant embryo (Figure 2.11.A, panels a-c). Two loci of proliferating cells were found in the forebrain at 28 hpf (Figure
2.11.A, panel c), which might represent the neurons of the dorsal and ventral rostral clusters (Ross et al., 1992). However, they were not observed in the morphant brain
(Figure 2.11.A, panel f). Quantitative analysis showed a reduction in the proliferating cell population between 20 and 28 hpf in L1.1 morphants, whereas an increase was observed in control embryos. In comparison with control brains of the same stage, a significant reduction in the number of proliferating cells in morphant brains was observed at both 24 and 28 hpf (Figure 2.11.B).
120 A B 20 h 24 h 28 h abc 40 ** 30 * Ctrl
20
ot ot ot ot ot per unit area ot 10 n=9 n=8 n=5 n=12 n=12 n=11 d e f cells % of Proliferating 0 20h 24h 28h L1.1MO
ot ot ot ot ot ot
Figure 2.11. Effects of L1.1 deficiency on brain cell proliferation. Embryos at different stages were incubated with anti-phospho-histone (pH3), which stained the nuclei of proliferating cells (purple). (A) Flat-mounted brains showing proliferating cells from the rostral-most of the telencephalon to the caudal-most of the otic vesicle of the brain. Brain structures were visualized by SYTO-16 Green staining (green). Two loci of proliferating cells (open arrows) were present in the forehead at 28 hpf, which may be the neurons of dorsal and ventral rostral clusters, but they were absent in the morphant. (B) Quantification of proliferating cells in control embryos (gray bars) and morphants (white bars) was performed by counting pH3-positive cells using ImageJ. The percentage of proliferating cells per arbitrary unit area of the brain was significantly reduced in morphants in comparison with control embryos at stages of 24 and 28 hpf: *, P < 0.001; **, P < 0.01 (Student’s t-test). Scale bar: 200 μm.
121 DISCUSSION
We report here the cloning of cDNAs for the complete coding regions of the two
L1 homologues in zebrafish, L1.1 and L1.2. Their expression patterns are distinct during embryonic development. L1.1 transcripts are present in all known classes of neurons, while L1.2 transcripts are detected only in small subpopulations of neurons with variable expression levels (Tongiorgi et al., 1995). Our immunostaining results corroborate with these earlier in situ hybridization results. The expression pattern of L1.1 and L1.2 suggests that they play a role in the development of axonal trajectories in the central and peripheral nervous system. Indeed, both L1.1 and L1.2 are potent inducers of neurite outgrowth from primary neurons. L1.1 has been found to stimulate the regeneration of axons from spinal neurons as well as synapse formation after spinal cord lesions in adult fish (Becker et al., 2004; Becker et al., 2005).
Morpholino knockdown of L1.1 synthesis in embryos leads to developmental anomalies in the embryonic brain. In contrast, the knockdown of L1.2 expression has only marginal effects on brain development. Although substrate-coated L1.2EC fusion protein displayed neuritogenic activity, axonal defects were not detected in L1.2 morphants, suggesting that the contribution of L1.2 to neural development may be quite different from that of L1.1. Further characterizations of the L1.1 morphants reveal resemblances to some of the phenotypes of human L1 syndrome. One of the most frequent pathological manifestations in human L1 patients is hypoplasia or the absence of two long axonal tracts, the corticospinal tract and the corpus callosum (Chow et al., 1985;
Yamasaki et al., 1995; Graf et al., 2000). Agenesis of these brain structures reflects the fundamental role of L1 in the development of nerve fibers. Loss of L1.1 expression in
122 zebrafish results in less extensive axonal network and shortened projections from neurons, such as the trigeminal ganglion neurons, which highlights the importance of L1.1 in brain development.
In addition to aberrant axonal outgrowth, more than half of the morphants at 48 hpf show notable enlargement of the 4th ventricle, which is associated with a disorganized
ventricular system. Many L1 mutations, especially truncation mutations in the
extracellular domain, are known to cause the abnormal accumulation of CSF in the third
and lateral ventricles, leading to the formation of X-linked hydrocephalus (Edwards et al.,
1961; Fransen et al., 1995; Yamasaki et al., 1995). Severe hydrocephalus is often
associated with stenosis of the aqueduct of Sylvius, which is located between the third
and fourth ventricles and facilitates the circulation of CSF. Aqueductal stenosis can lead
to the accumulation of CSF and the enlargement of cerebral ventricles in L1-related
hydrocephalus patients (Fransen et al., 1995). In the L1.1 morphants, histological
analyses and in situ hybridization show that the passage within the mid-hindbrain
boundary, which is the zebrafish equivalent of the aqueduct of Sylvius, is underdeveloped.
These striking similarities between the enlarged fourth ventricle in L1.1 morphants and
the hydrocephalus phenotype in L1 patients suggest that they may share a similar
etiologic mechanism.
Brain ventricle formation is known to involve two phases (Lowery and Sive,
2005). The initial morphogenesis, which is circulation-independent, requires intrinsic
neuroepithelial polarization and local cell proliferation. The second phase is characterized
by the circulation-dependent expansion of the lumen, which occurs after the beginning of
heart beats. It has often been found that many neural progenitor cells fail to migrate
123 properly from the ventricular proliferative zone in hydrocephalus patients (Kamiguchi et
al., 1998a). In zebrafish, the loss of L1.1 expression has led to disrupted ventricular
polarity as well as abnormal positioning of neural progenitor cells in brain ventricles
before the beginning of heart beats at 24 hfp. These neural cell migration events appear to
depend on L1.1-specific functions because L1.2 cannot compensate the loss of L1.1 in
morphants.
Polarization of the ventricular neuroepithelia is intrinsic and essential to brain
ventricle formation (Chenn et al., 1998; Yang et al., 2003). Molecules that establish and
maintain neuroepithelial polarity are important to ventricle development, and abnormal
expression of these molecules inevitably lead to the failure of ventricle opening in all
brain regions. Notably, Nok mutations cause similar anomalous development and result in
the absence of ventricular systems in zebrafish (Wei and Malicki, 2002; Lowery and Sive,
2005). In the mouse, loss of the lethal giant larvae homolog (Lgl1), a crucial molecule in
the maintenance of cell polarity during early embryonic development, also results in
severe hydrocephalus (Klezovitch et al., 2004). In L1.1 morphants, disturbed
polarization of supernumerary centrosomes reflects the necessity of L1.1 for
neuroepithelial integrity and polarity in the primitive brain, as well as the subsequent
initiation of ventricle formation. The ectopic positioning of neuroepithelial cells even 12
hours after birth suggests that L1 deficiency causes defects in cell migration in the
ventricular zones, possibly involving impaired motility and the loss of proper guidance cues.
Coordination of cell proliferation with migration in the neuroepithelia is also fundamental to embryonic brain ventricle development (Nadarajah and Parnavelas, 2002).
124 Our results show that defects in periventricular polarity in the L1.1 morphant brain is
accompanied by compromised cell proliferation. L1 is known to bind indirectly to actin
via its interactions with ankyrin and members of the ezrin, radixin, and moesin family,
through which L1 may mediate remodeling of the cytoskeletal scaffold (Davis and
Bennett, 1994; Burden-Gulley et al., 1997; Dickson et al., 2002). In the migratory cell at
the periventricular neuroepithelia, L1.1 may function in the establishment of cell polarity
via its interactions with cytoskeletal components, thus influencing cell proliferation.
However, we cannot exclude the possibility that L1.1 is able to transduce a proliferation
signal directly via its interaction with other membrane receptors since L1 has been
reported to promote the proliferation of neural progenitor cells (Dihne et al., 2003).
Other frequently associated malformations in hydrocephalus include the
underdevelopment of the anterior vermis of the cerebellum and fused thalami (Yamasaki
et al., 1995). It is likely that these developmental defects have also resulted from
abnormal cell proliferation and migration. The transcription factor Pax6 is known to
control L1 gene expression in the forebrain (Meech et al., 1999). In Pax6-deficient mice,
the loss of L1 expression in the forebrain has been implicated in the dispersion of cells in
the ventral thalamus and hypothalamus (Jones et al., 2002). Analyses of the L1.1
morphants have also revealed disorganized brain domains with Pax2.1- and Pax6-
positive progenitor cells distributed in much less restricted patterns, consistent with a role
for L1 in guiding neuronal cell migration.
It is, therefore, evident that L1 plays multiple roles during brain development. Our
analyses of the abnormalities in brain structures associated with the enlargement of the 4th ventricle in L1.1 morphants have revealed important roles for L1.1 in the proliferation of
125 neural progenitor cells, polarity of the neuroepithelial cells, and ventricle formation, which may underlie the pathological development of L1-related hydrocephalus. As the morphant phenotypes in zebrafish can be reversed by co-injection of L1.1 mRNA, a pathogenic mutation for CRASH syndrome may disrupt one or more L1.1 functions and compromise its rescue performance. Therefore, future characterization of malformations in morphant embryos co-injected with different L1.1 mutant mRNA species should yield further insights into the pathological mechanisms associated with the L1 mutations.
126
Chapter III:
Interactions between a Novel Soluble Form of L1.1 and
Neuropilin 1a Regulate Brain Ventricle Formation in
Zebrafish
(All experimental results described in this chapter were obtained by me.)
127 SUMMARY
Zebrafish express a novel soluble isoform of the cell adhesion molecule L1.1 prior to the expression of the full-length protein during embryonic development. L1.1s
contains only the first four immunoglobulin-like domains of L1.1. L1.1s is localized in
the periventricular cells of the embryonic brain, distinct from the expression pattern of
the full-length L1.1. Here, we show that L1.1s is involved in the development of brain
ventricles during zebrafish embryogenesis. Embryonic brain ventricle formation depends
on the oriented migration of neuronal progenitor cells from the germinative zones and the
proper establishment of ventricular polarity. Knockdown of L1.1 expression by
morpholinos leads to anomalous cell polarization and ectopic localization of neural
progenitor cells in the ventricular zone. Co-injection of L1.1s mRNA rescued the
morphant phenotype. However, the rescue function of L1.1s was compromised by a
mutation that disrupts L1.1s-to-neuropilin 1a interaction. Abrogation of L1.1s-neuropilin
1a interaction by an inhibitory peptide leads to aberrant ventricular polarity and inhibition
of ventricle formation. BrdU mapping studies revealed abnormal ventricular cell
migration in the presence of this inhibitory peptide. Taken together, the data suggest that
L1.1s-neuropilin 1a interaction regulates cell migration in the periventricle region and is
indispensable during embryonic ventricle formation.
128 INTRODUCTION
A crucial step in embryonic brain development is neuronal cell positioning at
proper destinations, which is often achieved through an active process of migration from
the germinal place to the target location (Tsai and Gleeson, 2005). In the cerebral cortex,
individual postmitotic neurons produced in the germinative neuroepithelia or ventricular
zones undergo directed radial migration, with the earliest-generated neurons positioned in
the deepest layers and later-generated neurons occupying the superficial layers
(Nadarajah and Parnavelas, 2002). Defects in early neuronal cell migration often result in
anomalous development of ventricles and brain structures (Gressens, 2006).
The proper positioning of neuronal cells in the developing brain depends on
extracellular guidance cues and proper cell-cell interactions. Mutations in neural cell
adhesion molecules or alterations in their temporal and spatial expression pattern
frequently lead to abnormal brain structures and deficits in neurological functions (Hatten,
1999). In humans, recessive X-linked hydrocephalus, which is characterized by the
narrowing of the aqueduct of Sylvius and the enlargement of the lateral ventricles, has
been attributed to mutations in the cell adhesion molecule L1 (Fransen et al., 1997;
Weller and Gartner, 2001).
L1 is a member of immunoglobulin (Ig) superfamily of cell adhesion molecules.
It consists of six Ig-like domains and five fibronectin type III-like repeats, a single transmembrane domain and a highly conserved cytoplasmic tail (Hortsch, 1996; Maness and Schachner, 2007). L1 exerts its functions via both homophilic and heterophilic interactions involving multiple domains in the extracellular portion of the protein
(Maness and Schachner, 2007). L1 plays a role in many important neuronal processes,
129 including axonal growth and navigation (Lemmon et al., 1989; Dong et al., 2003), neural
cell proliferation and survival (Haney et al., 1999; Chen et al., 2007), as well as axon
fasciculation (Honig et al., 1998; Wiencken-Barger et al., 2004) and myelination (Itoh et
al., 2005). A large number of mutations in the human L1 gene have been associated with
a variety of neurological deficits collectively known as L1 or CRASH syndrome (an
acronym for Corpus callosum hypoplasia, Retardation, Adducted thumbs, Spastic
paraplegia and Hydrocephalus; Fransen et al., 1995). In patients with severe
hydrocephalus, the regions around the ventricular zone of the brain contains a huge number of cells that have failed to migrate properly (Kamiguchi et al., 1998a), suggesting that neuronal cell migration requires L1-dependent cell adhesion.
Zebrafish express two L1 homologues, L1.1 and L1.2 (Tongiorgi et al., 1995).
Inhibition of L1.1 expression by morpholino antisense oligonucleotides leads to abnormal brain development in embryos, which exhibit enlarged 4th ventricle, underdeveloped mid-
hind brain boundary, and aberrant polarity at the ventricular surface (see Chapter II).
In this chapter, we report the discovery of L1.1s, a novel soluble isoform of L1.1,
which is comprised of the first four extracellular Ig domains. L1.1s is expressed in the
periventricular cells of the embryonic brain, and it can rescue the malformations of brain ventricles in the L1.1 morphant. A pathogenic mutation that abolishes the interactions
between L1.1 and neuropilin-1a (Nrp1a) inhibits the rescue function of L1.1s. An
inhibitory peptide that blocks L1.1s-Nrp1a interaction also disrupts the radial migration
of periventricular cells. These results support a role for L1-Nrp1 interaction in brain
ventricle development.
130 MATERIALS AND METHODS
Fish maintenance
Wild-type zebrafish (Danio rerio) were purchased from local pet store. Fish were maintained on a 14 hour light/10 hour dark cycle at 28.5°C according to Westerfield
(2000). Embryos were obtained by spontaneous spawning and raised in embryo medium.
They were staged by hours post fertilization (hpf) at 28.5°C and number of somites
(Kimmel et al., 1995).
Reverse-transcription-polymerase chain reaction (RT-PCR)
To demonstrate alternative usage of exons 2 and 27 in L1.1 and L1.2, total RNA was extracted from embryos at different development stages between 6 and 32 hpf. Five micrograms RNA of each stage were reverse-transcribed using random hexamer primers
(Invitrogen, Burlington, Ontario). Primers flanking exons 2 and 27 of L1.1 and L1.2 were the following (Figure 3.1.A): 5’-AGAAGAGGACAGAAGCGCACTC-3’ and 5’-
CAGGCCAGATTAACATCATCCA-3’ for L1.1 exon 2 (amplicons were expected to have 167 bp and 182 bp corresponding to the respective spliced forms); 5’-
GCAAACCAAAAACTGAAGGA-3’ and 5’-CCAGACTGTCGTTGCTGCT-3’ for L1.1 exon 27 (112 bp and 124 bp); 5’-CTCTAAAACCCGGGAACACCAC-3’ and 5’-
CCGAAAAGGTGGTGACAGATTT-3’ for L1.2 exon 2 (96 bp and 111 bp); 5’-
GAAGGCCAAATAGATTCCGAAG-3’ and 5’-GACTGGCTGTACGCTTCTCCTC-3’ for L1.2 exon 27 (88 bp and 100 bp). Polymerase chain reactions were carried out for 40 thermocycles consisting of denaturation at 94°C for 20 sec, annealing at 50°C for 20 sec
131 and extension at 72°C for 30 sec, followed by a final extension at 72°C for 10 min.
Amplification products were separated in poly-acrylamide gel buffered in 0.5x TAE and visualized by staining of ethidium bromide. Amplification with β-actin specific primers
(5’-GAGCTGTCTTCCCATCCATC-3’ and 5’-CCCATCTCCTGCTCAAAGTC-3’) was
performed to verify the quality of each cDNA sample.
Cloning of zebrafish soluble isoform of L1.1 (L1.1s) was performed by rapid
amplification of 3’-complementary end (3’ RACE). Primary reaction was performed
using L1.1 specific primer at the 5’ untranslated region (5’-
GACGTCCTTCAGTCTGTGTGTGT-3’), and was followed by a nested reaction using
an inner primer (5’-AGAAGAGGACAGAAGCGCACTC-3’). The presence of L1.1s
mRNA was further verified by RT-PCR using its specific primers L1.1SP18 (5’-
ATGTGCTGGAGTCTGATGATGGAG-3’) and L1.1s1 (5’-
CACACGAAATGCAGCCATACATA-3’) in different stages of embryos (Figure 3.2.B
panel a).
Mouse and human soluble L1 isoform (mL1s and hL1s) cDNA sequences were
predicted based on the donor site 5’ of intron 10 for alternative splicing in their respective
L1 gene. The presence of mL1s was examined by RT-PCR with mRNA extracted from
embryos of embryonic day 12 to 18 (E12-E18). Specific primers being used were mL1s
up (5’-CGAAGAGGACGATGGCGAGTA-3’) in the exon 8 and mL1s down (5’-
ATCCCCTAGAATCCCCAACAAG-3’) in the intron 10 (the predicted stop codon is
underlined). Human L1s was cloned from mRNA extracted from human embryonic kidney cells HEK 293. Primers were 5’-GGAAAGATGGTCGTGGCGCTGCGGTA-3’
132 and 5’-CCGAGGACACATCAGCTGCCA-3’. All cDNA were cloned and sequenced
(ACTG, Toronto, Ontario).
Preparation of His6-tagged fusion proteins
Complementary DNA sequence encoding L1.1s was obtained by PCR using PfuUltra
high-fidelity DNA Polymerase (Stratagene, La Jolla, CA). Amplification Primers were
5’-TACCATGGGCTACATTCAGATCCCACAC-3’ and 5’-
TGCTCGAGATGCTCACCAACAACATGCACGTGTGTG-3’. The cDNA fragment
was inserted into pET-28(a) vector (Novagene, Madison, WI) at NcoI and XhoI sites.
Plasmid was then transformed into E. coli BL21 (DE3). Bacterial fusion protein
production and purification was performed as described in Chapter II.
Plasmids were constructed for expression of Ig1 and Ig2 His-tagged fusion
proteins. PCR was carried out using the Ig1 primers: 5’-
accatggcaCACGACTACACATATAACAA-3’ and 5’-actcgagGGTCAGATGCACCAG-
3’; and Ig2 primers: 5’-accatggcaACAGAGCCCGTCCCAAGCTTG-3’ and 5’- actcgagATTCTTAACCACAGAGTTTGAG-3’ (extensions of restriction enzyme sequence for cloning are shown in lowercase). Mutant Ig1L123V was generated by site-
directed mutagenesis using PfuUltra high-fidelity DNA polymerase (Stratagene). Ig1 and
Ig2 fusion proteins were both soluble in the cytosol. They were purified by Ni-NTA
chromatography (Qiagen, Mississauga, Ontario) using a native protocol according to the
manufacturer’s instruction.
133 Neuritogenesis assay
Recombinant His6-tagged L1.1s protein were applied as substrate to cultures of
embryonic primary neuronal cells (Andersen, 2001). Cells (5x 104 cells) derived from the
trunk region were seeded on to 12-mm-diameter coverslips coated with the appropriate substrate. Cells were cultured in Leibovitz L-15 medium (Invitrogen) supplemented with
1% N2 (Sigma) and allowed to develop for 20 hours at 28°C.
Spinal neurons bearing neurites were identified by immunostaining of acetylated- tubulin as described in Chapter II. For each substrate, ~150 neurites from 3 different coverslips were scored in each experiment. Data were collected from four independent
experiments.
In situ hybridization
Antisense riboprobes specific to zebrafish L1.1 and L1.1s were designed. They recognized sequences 1563-2301 of the L1.1 cDNA (Genbank accession: X89024) and
1449-1780 of L1.1s cDNA located in the intron 10 (Figure 3.5.A). Templates for riboprobes were obtained by PCR methods with the T7 promoter sequence attached to the forward primers. For L1.1 antisense probe, the primers used were 5’-
TAATACGACTCACTATAGGGTGCTGGAGTGCTGTAGGA-3’ and 5’-
GATTCTAGACTCTATTCCAGCTCTGT-3’. For L1.1s antisense probe, primers being
used were 5’-TAATACGACTCACTATAGGGGGTTTCATTTTACTTGGTTT-3’ and
5’-GTGAGCATTAATAGTGTAAA-3’. The template for sense probe was generated by
primers and 5’-TAATACGACTCACTATAGGGGTGAGCATTAATAGTGTAAA-3’
and 5’-GGTTTCATTTTACTTGGTTT-3’ (the T7 promoter sequence is underlined).
134 Digoxigenin (DIG)-labeled riboprobes were synthesized by in vitro transcription using
T7 RNA polymerase and DIG-RNA labeling mix (Roche, Laval, Quebec) according to the manufacturer’s instruction.
Embryos of 24 hpf were collected and fixed in 4% PFA overnight at 4°C.
Embryos were kept in 100% methanol at -20°C until use. For in situ hybridization, embryos were rehydrated through serial 75% methanol/25% PBS-Tw (phosphate buffered saline, 0.1% Tween-20), 50% methanol/50% PBS-Tw, and 25% methanol/75%
PBS-Tw. Embryos were permeabilized with 10 μg/ml proteinase K for 5 minutes at room temperature and fixed again in 4% PFA for 20 minutes. After incubation in hybridization buffer (50% formamide, 5 x SSC, 0.1% Tween-20, 50 μg/ml heparin, 500 μg/ml salmon semen tRNA, 0.092M citric acid, pH 6.0), embryos were hybridized with 100 ng of DIG- labeled antisense riboprobes (1:250 in hybridization buffer) overnight at 65°C. To decrease background, embryos were incubated with fresh hybridization buffer for one hour and followed by stringent post-hybridization washes, which included once with
0.2xSSC at 65°C for 20 min and twice with 0.1xSSC at 65°C. For antibody detection, embryos were incubated with alkaline phosphatase-conjugated anti-DIG antibody
(1:5000) in blocking buffer (5% sheep serum and 5% BSA in PBS-Tw) overnight at 4°C.
For fluorescence detection post-hybridized embryos were incubated overnight in
Rhodamine-conjugated anti-DIG Fab (1:100 dilution, Roche). Embryos were embedded in 17% gelatin-10% sucrose and sectioned. Sections were mounted in Fluormount fluorescence mounting medium (DAKO, Mississauga, Ontario) for confocal microscopy.
135 Embryonic cell culture
Fish embryos were allowed to develop to 6 hours post fertilization (hpf) at 28ºC and were
dechorionated manually. Cells were dissociated in phosphate buffered saline (PBS)
containing 2% bovine serum albumin (BSA) and collected by centrifugation at 300 g for
5 minutes. Cells were washed twice and then resuspended in Leibovitz L-15 medium
(Invitrogen). Cells derived from 20 embryos were seeded into each 9 mm-well of the gasket (Electron Microscopy Sciences, Hatfield, PA) and allowed to settle on a poly-L- lysine slide (Fisher, Ottawa, Ontario). Unattached material was removed after 15 min, and 40 μl of fresh L-15 medium was applied to each well. Cells were further cultured for
2 hours at 28ºC. For proteinase K treatment of membrane proteins, cells were added with
10 μg/ml of proteinase K (Fermentas, Burlington, ON) in Hank’s balanced salt solution
(HBSS) after removal of the medium and incubated for 5 minutes till cells started to float up. Proteolysis was terminated by the addition of ice-cold 2% BSA in HBSS containing 3 mM phenylmethylsulphonyl fluoride. Cells were pelleted and washed twice in HBSS preceding western blotting.
For immunocytochemistry detecting cell surface presentation of L1.1s, embryonic cells were fixed in 4% PFA for 10 min. Without permeabilization, cells were subjected to immunostaining with anti-L1.1 antibody (5 μg/ml).
Western blotting and identification of L1.1s dimer
Lysates from hand-deyolked embryos of various stages or cultured cell were prepared
and subjected to SDS-PAGE and Western blot. Affinity purified rabbit anti-L1.1
antibody was applied as 1 μg/ml in block buffer (5% dry milk in TTBS, Tris-buffer saline
136 with 0.1% Tween20). Immunodectection was processed with goat anti-rabbit IgG
(1:10,000, Pierce) and subsequently ECL (Amersham, Baie d’Urfe, Quebec) following a standard protocol. To dissociate L1.1s dimer, lysates were first boiled in loading buffer and treated with 1% TritonX-100 in NaCl (0.5 M for recombinant protein and 1 M for embryonic culture lysate) before they were applied on the gel. For anti-actin immunostaining, monoclonal anti-actin mouse IgM (JLA20, Developmental Studies
Hybridoma Bank, Iowa City, IA) was applied at 1:500.
In vitro mRNA production and microinjection
Complementary DNA containing the complete open reading frame of L1.1s was cloned into pBSIISK vector downstream of T7 promoter. Pathogenic mutations in L1.1s
(L1.1sL123V and L1.1sE314K) were generated by site-directed mutagenesis using PfuUltra high-fidelity DNA polymerase (Stratagene). Capped RNA was synthesized using mMESSAGE mMACHINE T7 kit (Ambion, Foster City, CA). For embryo injection, capped RNA was diluted to 0.1 mg/ml in a solution containing 4 mg/ml morpholino against L1.1 5’-untranslation region (5’-CAGTCCCGACTCCAGACACACACAC-3’).
Approximate 0.5 nl of injection volume was applied to each embryo at 1-2 cell stage (2 ng morpholino and 0.05 ng mRNA). The 4-mismatch morpholino control in use was 5’-
CAgCTcCTG AgAGACAgAAACAgAG-3’ (mismatched bases are in lower case). The expression of neuropilin 1a (Nrp1a) was knocked down using the morpholino as described (2 ng per embryo, Lee et al., 2002). All injections were performed using the
Eppendorf Transjector 5246.
137 Whole mount immunohistochemistry
Whole-mount immunostaining was performed on embryos using affinity-purified
rabbit antibodies against L1.1 IgG (4 μg/ml), γ-tubulin (1.6 μg/ml; Abcam, Cambridge,
MA), monoclonal antibodies against zn-12 (5 μg/ml; Zebrafish International Resource
Center, Eugene, OR). Alexa 568-conjugated (Invitrogen) or HRP-conjugated (Pierce,
Ottawa, Ontario) secondary antibodies were used for immunostaining. Embryos were
embedded in 17% gelatin plus 10% sucrose and sectioned. Sections were mounted in
Fluormount and images were collected by confocal microscopy. The enhanced DAB
color substrate (Pierce) was used for non-fluorescence specimens. Embryos were
mounted and photographed under an inverted microscope (Nikon ECLIPSE TE2000).
Cell binding assay
COS7 cells were transiently transfected with plasmids expressing enhanced green
fluorescence protein (EGFP) fused zebrafish neuropilin 1a (Nrp-1a) or EGFP only.
Transfectants were seeded as monolayer on the Teflon-coated 8-mm well slides (Fisher).
For the binding assay, His6-tagged fusion proteins (0.1 μM) were first incubated with
mouse anti-His6 monoclonal IgG (0.3 μg/ml, Novagen) in binding buffer (1x HBSS, 10
mM Hepes, 0.5 mM MnCl2, 2% BSA) for 1 hour at room temperature. Fifty μl of
recombinant proteins were added to each well of transfected cells and allowed to bind for
45 min at room temperature. At the end of incubation period, the unbound material was
removed with three washes in HBSS. Cells were fixed in 4% PFA for 15 min without
permeabilization, extracellular bound His6-tagged proteins were detected by Alexa 568 conjugated goat anti-mouse IgG (1:500, Invitrogen). For the peptide inhibition assay, 10
138 μM of each peptide, YAANEL and YAANEV (Advanced Protein Technology Centre, the
Hospital for Sick Children, Toronto, Ontario), was included in the incubation mixture
before applying to the transfectants.
Injection of recombinant proteins into the brain
Bacterial His6-tagged fusion proteins containing L1.1s and individual Ig1 and Ig2
domains of L1.1 were diluted to 2 μM in PBS containing 0.1% neutral red. Each fusion
protein (~0.5 nl) was injected into the midbrain of 24 hpf L1.1 morphants, which showed
flattened forebrain and midbrain and lacked distinct mid-hindbrain boundary. The
morphants that received the fusion protein were cultured for another 3 hours at 28ºC, and
the effect of the protein on ventricular polarity was examined by immunostaining with
anti-γ-tubulin (1.6 μg/ml).
Treatment of embryos with inhibitory peptides
Wild-type embryos were raised to 17 hpf and treated with different concentrations of
peptide (YAANEL or YAANEV as control) in embryo medium containing 4% DMSO,
which enhances the penetration of peptide into embryos, but does not cause lethality.
After 5 hours, embryos were fixed for further morphological analyses. SYTO16 green
(1μM, Invitrogen) was used to stain the nucleus and reveal the brain scaffold.
Polarization of the ventricle was revealed by anti-γ tubulin immunostaining. F-actin was
stained using Alexa 568-labeled phalloidin (1:40 dilution, Invitrogen).
139 Monitoring ventricular cell migration with BrdU
Wild-type 48 hpf embryos were pulse-labeled with BrdU (Sigma, 10 mM in 15% DMSO)
at 4ºC for 20 min, followed by several quick rinses in fresh embryo medium containing
15% DMSO and incubation with 10 mM dTTP in 15% DMSO for another 5 minutes in the cold. Next, embryos were placed in a solution containing 2 mM peptide in 4% DMSO and incubated at 28ºC for 5 hours before fixation. Labeled cells were detected using an anti-BrdU antibody (G3G4, 1 μg/ml; Developmental Studies Hybridoma Bank).
Northern blot
The L1.1 sequence 1-1556 bp was used as the template for anti-sense riboprobe, which is labeled with Digoxigenin.
Ten μg of total RNA isolated from 48 hph embryo were electrophoresed in 1% agarose/5% formaldehyde gels in MOPS buffer, and were transferred overnight onto positively charged nylon membrane (Roche Diagnostics). Following cross-linking, the membranes were hybridized at 68°C with the antisense riboprobe in hybridization buffer containing 50% deionized formamide, 5x SSC, 0.1% N-lauroylsarcosine, 0.02% SDS and
2% blocking reagent. After overnight hybridization, low stringency washes in 2x SSC,
0.1% SDS at room temperature followed by high stringency washes in 0.5x SSC, 0.1%
SDS at 68°C were employed to eliminate nonspecific binding. Riboprobe binding was detected by reaction with anti-DIG conjugated to alkaline phosphatase (anti-DIG-AP)
(Roche Diagnostics), followed by application of disodium 3-(4-methoxyspiro{1,2- dioxetane-3,2’-(5’-chloro) tricyclo [3.3.1.1]decan}-4-yl) phenyl phosphate (Roche
Diagnostics). The chemiluminescent signal was visualized on X-ray film.
140 RESULTS
Expression of a novel soluble isoform of L1.1 during embryonic development
The mammalian L1 gene is known to contain two alternatively spliced mini exons,
exon 2 and exon 27, which are expressed exclusively in neuronal cells (Reid and
Hemperly, 1992; Jouet et al., 1995b). Both exons were shown to be alternatively spliced
in the two zebrafish L1 homologues, L1.1 and L1.2, during embryonic development
(Figure 3.1). Surprisingly, two L1.1 isoforms were detected by RT-PCR at embryonic
stages of 6 and 8 hpf. They contained the alternatively spliced exon 2 but not the exon 27
sequence, indicating they were truncated prior to the exon 27 (Figure 3.1.B).
To identify these short forms, 3’-RACE was performed using cDNA prepared
from 6 hpf embryos. In both cases, a single cDNA fragment containing part of the intron
10 sequences was obtained, suggesting that both isoforms were produced by alternative
splicing at the donor site 5’ of intron 10 (Figure 3.2.A). Two short L1.1 isoforms (1791
bp and 1776 bp) were cloned, encoding peptides of 430 and 425 amino acids,
respectively. Both predicted peptides contained the first four Ig-like domains, differing
only in the presence of the five amino acids encoded by exon 2.
To examine the expression of L1.1s during embryonic development, RT-PCR was carried out using primers specific for L1.1s. L1.1s mRNA was first detectable at 6 hpf and its expression increased at later embryonic stages (Figure 3.2.B).
As revealed by Northern blot, two transcripts were detected with the riboprobe against the 5’ sequence of L1.1, which will recognize both long and short forms of L1.1.
The size of long isoform was calculated as 12 kb, which is in agreement with the
141 A
15 bp 12 bp L1CAMs E 1 E 2 E 3 E 26 E 27 E 28
Splicing forms: E1 E2 E3 Splicing forms: E26 E27 E28 E1 E3 E26 E28 B Embryonic stage (hpf) bp 6 8 10 12 14 16 18 20 24 32 zL1.1 200 E1 E2 E3 182 bp E 2 100 E1 E3 167 bp 200 E26E27E28124 bp E 27 100 E26E28 112 bp
zL1.2 200 E1 E2 E3 E 2 111 bp 100 E1 E3 96 bp E 27 100 E26E27E28 100 bp E26E28 88 bp
β-actin 600
Figure 3.1. Detection of a novel alternatively spliced isoform of L1.1. (A) Schematic drawing depicting the RT-PCR strategies to assess the expression of alternatively spliced isoforms during embryogenesis. Specific primers were designed for regions flanking exon 2 and exon 27, respectively. The amplification products were separated by polyacrylamide gel electrophoresis. (B) Temporal expression of exons 2 and 27-containing transcripts in zebrafish L1.1 and L1.2 during embryonic development between 6 to 32 hpf. At the stages of 6 and 8 hpf, the L1.1 primers detected two isoforms, containing only the 5’ region flanking exon 2 but lacking the 3’ portion that contained exon 27.
142 AB
L1.1 gene …cacgtgcatgttgttggtgagcattaatag……agaactg… (a) Exon 10 Intron 10 Exon 11 In 8 In 9 In 10 L1.1 gene Ex 8 Ex 9 Ex10 Ex11 L1.1s Exon 10 Intron 10 RT-PCR primers HVHVVGEH L1.1 Exon 10 Exon 11 L1.1s Ex8 Ex9 Ex10
HVHVVELPA Product size: 508 bp Putative Ig4 Putative Ig5
C D (b) Embryonic stage (hpf) Mouse embryonic day L1.1 kb 0 3632 kb kb E12 E16 E18 0.7 7.0 1.5 0.5 L1.1s 4.7 0.75 0.3 0.5 mL1s 2.7 0.1 L1.1s 1.8 0.25 0.8 1.5 0.5 β-actin 1.0 0.6
E L1.1s CEAVNKHGSILINTHVHVVGEH 430 hL1s CEARNRHGLLLANAYIYVVRECPPFLTS 431 mL1s CEARNQHGLLLANAYIYVVRECTDLISCLPSFSLSLLSSHPPPYPCLIAPSLSFQSTKTTQSAACWGF 470
Figure 3.2. Cloning and identification of the short alternatively spliced isoform of L1.1. (A) Schematic drawing depicting the splicing junction between exons 10 and 11 and the amino acid sequence of the C-terminal region of L1.1s. (B) Expression of L1.1s mRNA during embryonic development. (a) PCR primers were designed specifically for L1.1s. The forward primer recognized a sequence in exon 8 and the reverse primer recognized a sequence in intron 10, amplifying a product of 508 bp. (b) L1.1s expression was detected at embryonic stages 6 - 32 hpf (arrow). The PCR products of β-actin were included as an internal control (open arrow). (C) Northern blot analysis revealed the size of L1.1s transcript is 2.2 kb. (D) RT-PCR was carried out using primers exclusive to a deduced mouse soluble L1 isoform, mL1s. mL1s (arrow) was detected in between E12 and E18 of embryonic development. (E) Alignment of the C-terminal region of the deduced peptides for L1s in mouse (mL1s), human (hL1s) and zebrafish (L1.1s). Predicted intron-exon boundary 5’ of intron 10 is marked by an open arrowhead.
143 published result (Tongiorgi et al., 1995). The size of the short isoform is calculated as 2.2 kb, according to its migration distance during the electrophoresis (Figure 3.2.C).
To determine whether a soluble L1 is also expressed in higher vertebrates, specific primers were designed for the mouse L1 gene based on a splicing scheme similar to that of zebrafish L1.1s. A product of 513 bp was obtained by RT-PCR using mouse embryonic mRNA as template. Sequencing data confirmed that the cDNA contained the
3’ portion of the predicted isoform. Therefore, a short form of mouse L1 (mL1s) is expressed during mouse embryonic development (Figure 3.2.D). A similar approach led to the discovery of a similar alternatively spliced L1 isoform (hL1s) in human embryonic kidney 293 cells (Figure 3.2.E). Both mouse and human L1s isoforms contained the open reading frame encoding the first four Ig domains of L1.
Presentation of dimeric L1.1s on the cell surface
Immunoblot analysis was carried out using an affinity purified anti-L1.1 antibody to ascertain the expression of the L1.1s protein. The full-length 200 kDa L1.1 protein and its 140 kDa degradative product (Pradel et al., 1999) appeared at 14 hpf and became clearly detectable at 18 hpf. However, a 50 kDa band, corresponding closely to the predicted size of L1.1s, was detected in embryos between 6 hpf and 48 hpf, (Figure 3.3A).
The full-length 200 kDa L1.1 protein and its 140 kDa degradative product (Pradel et al.,
1999) became detectable at 18 hpf. Interestingly, a 100 kDa band was also detected as
early as 6 hpf and increased over time, suggesting the presence of a dimer form of L1.1s.
Dimerization of L1 fusion proteins containing immunoglobulin domains has been found
less sensitive to SDS and reducing reagents, when the proteins are produced from HeLa
144 A a Embryonic stage (hpf) b Embryonic stage (hpf) L1.1 6 10 12 14 18 4824 6 10 12 14 18 kDa L1.1 171 L1.1 fragment 110 L1.1s dimer
79 60 L1.1s 47 Actin (a longer exposure) 47
B ab kDa 12 kDa 12 171 tetramer 171 110 dimer 110 L1.1s dimer 79 60 79 60 47 L1.1s L1.1s NaCl/ 47 TritonX-100: – + – +
nic C o um y br ells edi m c M E L1.1 12 3 kDa 180 115 L1.1s dimer 82 64 49 L1.1s 37
Actin 49 37 Proteinase K: +–
D a L1.1s b L1.1s DiI;DAPI
145 Figure 3.3. Expression of the L1.1s protein during embryonic development. (A) Embryo lysates from different stages of zebrafish embryonic development were prepared for SDS-PAGE followed by immunoblot analysis. (a) A shorter exposure and (b) a longer exposure of the chemiluminescence signal. The full-length L1.1 protein and its N-terminal proteolytic fragment of 200 and 140 kDa, respectively, were detected by the anti-L1.1 antibody (open arrowhead). The antibody also detected a protein band of 50 kDa, corresponding to the deduced size of L1.1s (open arrow), and a 100 kDa band, which might represent the dimer form of L1.1s (arrow). Longer exposure time showed that both 200 and 140 kDa bands were present at 14 hpf. Actin was included as the loading control. (B) Effects of salt and detergent treatments on the L1.1s dimers. (a) Dissociation of recombinant L1.1s protein treated with 0.5 M NaCl and 1% TritonX-100. (b) Gel profiles of L1.1s protein derived from embryonic cell lysate (6 hpf) in the absence (−) or presence (+) of 1 M NaCl and 1% TritonX-100. (C) Cultured embryonic cells were subjected to proteinase K digestion (10 μg/ml) for 5 min at room temperature. Culture medium and cell lysates with (+) or without (−) prior proteinase K treatment were processed for immunoblot analysis. The actin blots were included as internal controls and ascertain the equivalent amounts of cellular protein were treated either in the presence or absence of proteinase K (lanes 2 and 3). (D) Cell surface presentation of L1.1s revealed by immunocytochemistry. (a) Non- permeabilized cells dissociated from the 6 hpf embryo were immunostained with anti- L1.1 antibody (green). (b) The superimposed micrograph with cytosol (red) stained by DiI and nucleus (blue) by DAPI. Scale bar: 10 μm.
146 cells (Schurmann et al., 2001). Indeed, His-tagged L1.1s protein was present in dimer and tetramer forms in solution, which were dissociated into the monomeric form in the
presence of 0.5 M NaCl and 1% TritonX-100 (panel a in Figure 3.3.B). Similarly, the
dimeric form of L1.1s from embryonic lysate was sensitive to NaCl and Triton X-100
treatment and presented predominantly as the monomeric form (panel b in Figure 3.3.B).
The deduced peptide sequence of L1.1s does not contain a predicted
transmembrane domain, suggesting that it may be secreted by cells into the extracellular
matrix. Indeed, the monomeric form of L1.1s was detected in the culture medium two
hours after 6 hpf embryonic cells were seeded on a culture dish (lane 1 in Figure 3.3.C).
Although the dimer form was absent from the culture medium, it was found associated
with the cultured cells (lane 3 in Figure 3.3.C). To determine whether the L1.1s dimer
was associated with the endo- or ecto-surface of the cell membrane, live cells were
treated briefly with proteinase K preceding lysis and preparation for SDS-PAGE. The
L1.1s dimer was removed completely by proteinase K (lane 2 in Figure 3.3.C),
suggesting that it was associated with the outside of the cell membrane. Immunostaining
of non-permeabilized cells derived from 6-hpf embryos revealed clustering of L1.1s on
the cell membrane (Figure 3.3.D).
To investigate the potential role of secreted L1.1s, His-tagged L1.1s was expressed in E. coli and coated on culture dishes to determine its effect on neurite outgrowth. Spinal neurons isolated from 18 hpf embryos were seeded on L1.1s and cultured for 20 hours. Substrate-coated L1.1s promoted neurite growth from the primary spinal neurons to an extent comparable to that of the complete extracellular portion of
147 L1.1, indicating that the neuritogenic activity of L1.1 is associated with the first four extracellular Ig-like domains (Figure 3.4).
Association of L1.1s with the brain ventricle surface
Whole mount in situ hybridization was carried out with specific riboprobes to distinguish L1.1s mRNA from L1.1 mRNA (Figure 3.5.A). Expression of L1.1 in wild- type 24-hpf embryo was found to be exclusive to neuron clusters in the brain (Figure
3.5.B), including the dorsal and ventral rostral clusters, ephyseal cluster, posterior commissure, trigeminal ganglion, anterior lateral line ganglia and posterior lateral line ganglion. A cross-section of the hindbrain showed that no L1.1 mRNA was detectable in the regions proximal to ventricles.
Contrary to the L1.1 expression pattern, L1.1s expression seemed to be distributed more widely in the brain. In situ hybridization showed that L1.1s was present in the forebrain, the midbrain, mid-hindbrain boundary, the hindbrain and the eye (Figure 3.5.C, panels a and b). Confocal images of in situ hybridization revealed that L1.1s was expressed more abundantly in cells situated in apposition to the brain ventricles at levels of midbrain, isthmus and hindbrain.
Involvement of L1.1s in brain ventricle formation
In cells that lined the ventricular surface, only L1.1s mRNA, but not L1.1 mRNA, was expressed, suggesting that L1.1s may play a role in the brain ventricle development.
Ventricular cells in the neocortex are neural progenitor cells (NPCs) (Rakic, 1988), which divide at the ventricular edge and migrate radially by either somal translocation or glia-
148 A a b c
BSA L1.1EC L1.1s
B C m)
μ 120 100 100 80
80 60 60 40 40 20 20 axis >X neurites
Percentage of cells with with cells of Percentage 0 0
Mean length of neurites ( neurites of length Mean 1000 200 300 400 500 BSA zL1.1s zL1.1EC Neurite Length (μm) (n=4)
Figure 3.4. Neuritogenic activity of L1.1s. (A) Embryonic primary neurons isolated from the 18 hpf embryo were cultured on slides coated with substrates BSA (a), recombinant L1.1EC (b), and L1.1s (c) for 20 hours. The neurons and their extended processes were identified by anti-acetyl tubulin immunostaining. Scales bars: 50 μm. (B) Mean neurite lengths for the longest neurite extending from neuronal cells were estimated. Data represent the mean ± S.E.M. of four experiments. (C) Representative cumulative plots of neurite lengths projected on the three substrates: BSA (♦), L1.1s (○), and L1.1EC (■).
149 A 1 1563 2301 4067 L1.1 WWWWWW L1.1 riboprobe 1 1449 1780 L1.1s In 10 WWW L1.1s riboprobe
B ec a V VIII PLL b PC c ot drc * ot ot vrc ec PC
C MHB F M M MHB a H b F H
e cd e * *
Midbrain Isthmus Hindbrain
150 Figure 3.5. Differential expression of L1.1s and L1.1 in the embryonic brain. (A) Two specific riboprobes were designed to distinguish between L1.1 and L1.1s. The L1.1 probe targeted the cDNA sequences between 1563-2301, and the L1.1s probe targeted the sequences 1149-1780 of the L1.1s transcript. (B) In situ hybridization of L1.1 using a riboprobe specific to the full-length transcripts. Dorsal (a) and lateral (b) views of wild-type 24 hpf embryonic brain, rostral is to the left, showed that L1.1 was present exclusively in neuronal clusters. (c) Cross section of hindbrain showed that L1.1 expression was absent from the ventricle (*). (C) In situ hybridization specifically against L1.1s in the wild-type 24 hpf embryo. (a) Dorsal and (b) lateral views of the brain with rostral to the left. L1.1s was located in the forebrain, the midbrain, mid-hindbrain boundary, the hindbrain and the eye. (c-e) mRNA of L1.1s was detected by a fluorescence anti-DIG antibody. Transverse sections of the brain at levels of midbrain (c), isthmus (d) and hindbrain (e) revealed that L1.1s was expressed in the cells in apposition to the ventricle (*). F, forebrain; H, hindbrain; M, midbrain; MHB, mid-hindbrain boundary; ot, otic vesicle; PC, posterior commissure; PLL, posterior lateral line ganglion; V, trigeminal ganglion; VIII, anterior lateral line ganglia; drc, dorsal rostral clusters; e, eye; ec, ephyseal cluster; vrc, ventral rostral clusters. Scale bars: 0.1 mm in B and C (a, b); 20 μm in C (c-e).
151 guided locomotion during cerebral cortex development (Nadarajah and Parnavelas, 2002).
To investigate the function of L1.1s, L1.1s mRNA was co-injected into embryos with morpholino antisense oligonucleotides, which knocked down the expression of both
L1.1s and L1.1 in the embryo (Figure 3.6.C).
In the absence of both L1.1s and L1.1, embryos showed abnormal development of the brain. At 24 hpf, brain structures (including tectum, tegmentum and mid-hindbrain boundary) were underdeveloped in the L1.1 morphant (Figure 3.6.A, panel b). Analysis of confocal images of embryonic brain stained with SYTO green showed that these structures were indefinable in the L1.1 morphant (Figure 3.6.A, panel b’). In contrast, in embryos injected with the control morpholino, the wild-type pattern of fluorescence was observed in tectum, tegmentum and cerebellum (Figure 3.6.A, panels a and a’).
Significantly, these morphant brain defects were rescued by co-injection of L1.1s mRNA with the morpholino (c and c’ in Figure 3.6.A). Midbrain structures and mid- hindbrain boundary were distinct. Among the embryos that received combination injection of L1.1s mRNA and L1.1 morpholino (122 embryos from four independent experiments), less than 10% of embryos manifested abnormalities in the brain at 24 hpf, indicating rescue of the morphant phenotype. However, co-injection of L1.1s mRNA failed to rescue the morphant phenotypes in the body trunk (shortened and/or curved), and >60% of embryos exhibited malformations in the trunk (Figure 3.6.B). These results suggested that L1.1s was sufficient to re-establish proper brain domain organization in the L1.1 morphant.
Therefore, we hypothesized that the first four Ig domains contain critical components involved in normal brain domain development. The contributions of L1.1s to
152 A Ctrl L1.1 MO MO+L1.1s MO+L1.1sL123V MO+L1.1sE314K abcde
Bright 200 μm
a’Te Ce b’ c’ d’ e’ F
SYTO Tg H 200 μm
B C Brain domain abnormalities Trunk abnormalities a 48 hpf b 24 hpf L1.1s mRNA ** L1.1 rescued embryos 100% * Ctrl MO kDa 80% L1.1 kDa L1.1 181 WT L123V E314K 60% 70 L1.1 fragment 40% 55 n=84 n=47 n=186 n=122 116 L1.1s dimer 20% 40 0 82 O A O A Western blot: anti-L1.1 WT M N NA NA WT M N NA NA .1 R R R .1 R R R 1 m m m 1 m m m 64 V K V K L s 3 4 L s 3 4 2 1 2 1 .1 1 3 .1 1 3 1 L E L E s s 1 s s +L .1 .1 +L .1 .1 O 1 1 O 1 1 L L L L M + + M + + 49 L1.1s .1 O .1 1 O 1 O O L M M L M M .1 .1 .1 .1 1 1 1 1 L L L L Actin 47
Western blot
D E Ctrl L1.1MO L1.1MO +L1.1s Nrp1a MO Nrp1a MO acb d * a *
200 μm Midbrain b a’ b’ c’ d’ * * * * 200 μm 50 μm Hindbrain
F a b c d
Ctrl L1.1MO L1.1MO+L1.1s L1.1MO+L1.1 50 μm
153 Figure 3.6. L1.1s mRNA rescued ventricle formation but not axonal growth in L1.1 morphant. (A) Brain development in embryos at 24 hpf, which had been co-injected with morpholino (4 mg/ml) and wild-type or mutant L1.1s mRNA (0.1 mg/ml). Bright field images of the living embryos (a-e) and confocal scans of brain stained with SYTO16 green (a’-e’), lateral views with rostral to the left. In the control embryo (a and a’), cell dense zones showing intense fluorescence were located in the forebrain (F), tegmentum (Tg), tectum (Te), cerebellum (Ce) and hindbrain (H). (b and b’) Embryos injected with L1.1 morpholino targeting the 5’-UTR sequences. Red arrows denote the underdeveloped MHB. (c and c’) Embryos co-injected with L1.1s mRNA. The cell dense zones were restored in the tegmentum (arrow in c’) and MHB appeared normal (open arrow). (d and d’) Embryos co-injected with L1.1sL123V mRNA, showing abnormal development of midbrain (open arrowhead) and MHB (red arrows). (e and e’) Embryos co-injected with L1.1sE314K, showed normal brain development. (B) Comparison of abnormalities in the brain and the trunk of injected embryos. Abnormal trunk development included a shorter and/or curved body. **, p<0.01; *, p<0.05 (Student’s t- test). (C) Lysates of 24-hpf uninjected embryos (ctrl) and L1.1 morphants (L1.1MO) (a), and embryos co-injected with in vitro synthesized mRNA and L1.1 morpholino (b) were subjected to immunoblot analysis against L1.1. The expression of both L1.1s and L1.1 was inhibited in L1.1 morphants. Wild-type and mutant L1.1s mRNA were produced in the embryos. (D) Knockdown of Nrp1a by morpholino. (a) Left lateral view of a living embryo and (b) confocal image of brain stained SYTO16 green, showing aberrant development of MHB (red arrows). (E) Transections of midbrain (a-d) and hindbrain (a’-d’) regions showing γ-tubulin immunostaining, which revealed the polarization of supernumerary centrosomes at the ventricular surface. Red arrows denote abnormal localization of centrosomes in cells at the underdeveloped midbrain ventricle in L1.1 morphants and Nrp1a morphants. Open arrowheads mark non-polarized distribution of centrosomes in the L1.1 morphant hindbrain. Asterisks mark the ventricle. (F) Lateral views (with rostral to the left) of axonal trajectories from trigeminal ganglion neurons immunostained with mAb zn-12. Scale bars: 200 μm in A and D; 50 μm in E and F.
154 midbrain development were assessed by immunostaining cross-sections with an anti-γ tubulin, which labels supernumerary centrosomes that are intrinsically polarized to the luminal surface of neuroepithelial progenitors at the developing ventricle (Chenn et al.,
1998). At 24 hpf, immunofluorescence representing centrosomes was polarized to the ventricular surface in both midbrain and hindbrain regions of the control brain (Figure
3.6.E, panels a and a’). In L1.1 morphants, however, the peri-ventricular polarity of centrosomes was disrupted (Figure 3.6.E, panels b and b’). In particular, in the midbrain level, the ventricle opening became obscure (Figure 3.6.E, panel b). When morpholino was co-injected with L1.1s mRNA, centrosomes were polarized to the ventricular surface, in a pattern similar to that of wild-type (WT) embryos (Figure 3.6.E, panel c). Despite its ability to rescue midbrain development, L1.1s mRNA did not re-establish the polarization of centrosomes in the hindbrain (Figure 3.6.D, panel c’).
Studies of pathological missense mutations of L1 should provide invaluable insights on the functional significance of specific residues and domains of the protein.
Among the mutations that occur in the first four Ig domains, L120V and E309K have been associated with the enlargement of the lateral ventricle or hydrocephalus (De
Angelis et al., 1999) and yet have little influence on the cell surface expression of the protein (De Angelis et al., 2002). Therefore, the effects of these mutations on the rescue function L1.1s were examined by co-injecting mRNA of L1.1sL123V (corresponding to
the mutation L120V in the Ig1 domain of human L1) or L1.1sE314K (corresponding to the
human mutation E309K in the Ig3 domain) with the inhibitory morpholino. In embryos
co-injected with L1.1sL123V mRNA, both tectum and tegmentum of the midbrain
remained ambiguous (Figure 3.6.A, panels d and d’), suggesting that Leu-123 plays a
155 critical role in L1.1s function. However, the mutation E314K did not hamper the ability of L1.1s to rescue the morphant phenotype (Figure 3.6.A, panels e and e’).
It has been reported that the mutation L120V in mouse L1 disrupts its interaction with Nrp1 when corticospinal axons are crossing the midline (Castellani et al., 2002).
Nrp1 is expressed in the superficial layers of optic tectum and the protein is localized to the luminal surface of the mesencephalic ventricle in vertebrates (Takagi et al., 1987;
Takagi et al., 1991; Takagi et al., 1995; Kawakami et al., 1996). In zebrafish, two Nrp1 homologues have been identified, neuropilin-1a and 1b (Bovenkamp et al., 2004; Yu et al., 2004). The expression of Neuropilin-1a (Nrp1a) is relatively abundant in the midbrain and forebrain regions, in particular the tegmentum and tectum (Yu et al., 2004).
Knocking down the expression of Nrp1a in the zebrafish shows defects in vascular development (Lee et al., 2002) and axonal guidance (Wolman et al., 2004; Feldner et al.,
2005; Kuan et al., 2007). In order to assess the role of Nrp1a in brain ventricular development, Nrp1a was knocked down in the embryo. At 24 hpf, the Nrp1a morphant exhibited indistinct midbrain structures (Figure 3.6.D). The resultant abnormalities, including ambiguous midbrain ventricle and disrupted ventricular polarity of centrosomes, shared similarities to that observed in L1.1 morphants (Figure 3.6.E, panel d).
To determine whether L1.1 binds Nrp1a directly, Nrp1a-EGFP was constructed for transfection of COS7 cells. Transfectants were used to perform in vitro cell binding assays with His6-tagged L1.1 recombinant proteins. Results from these cell binding assays showed that recombinant L1.1s binds to COS7 cells expressing Nrp1a (Figure
3.7.A). The direct interaction between L1 and Nrp1 is, therefore, conserved in zebrafish.
Domain Ig1 achieved a similar binding level to Nrp1a as L1.1s, whereas mutation L123V
156 Nrp1a-EGFP COS7 EGFP COS7 A BCDE F G
L1.1s Ig1 Ig1L123V L1.1s L1.1s BSA L1.1s +YAANEL (10-5M) +YAANEV (10-5M)
Red: mAb anti-His6 Green: GFP Blue: DAPI
H
100 80 binding 2 60 40
>700 pixel 20 % transfectantsbearing 0 Protein: L1.1s Ig1 Ig1L123V L1.1s L1.1s BSA L1.1s + + YAANEL YAANEV Transfectant: Nrp1a-EGFP EGFP
Figure 3.7. Involvement of L1.1 sequence YAANEL in L1-Nrp1a interaction. Various
fusion proteins of L1.1 were cross-linked by anti-His6 antibody and applied on COS7 cells expressing Nrp1a-EGFP (green) to assess their ability to bind to Nrp1a. Binding of fusion
protein was detected by anti-His6 mouse IgG (red). Similar levels of L1.1s (A) and Ig1 domain (B) bound to Nrp1a-expressing cells. The mutation L123V (C), as well as the peptide YAANEL (D), disrupted the binding of Ig1 to Nrp1a. (E) The control peptide YAANEV had no effect. (F, G) The two controls showed only background level of binding. (H) Histogram showing the relative levels of L1.1s binding to the COS7 cells. Data represent the mean ± s.d. Scale bar, 50 μm.
157 (corresponding to human mutation L120V) interfered with this interaction (Figure 3.7B,
C). The sequence 115-FASNKL-120 in the Ig1 domain of L1 is apparently responsible
for L1-Nrp1 interaction and the pathological mutation L120V in this sequence disrupts
this interaction (Castellani et al., 2002). When the synthetic peptide 118-YAANEL-123,
which corresponds to the FASNKL sequence of human L1, was added to the binding assay, it inhibited the binding between L1.1s and Nrp1a (Figure 3.7.D), whereas the control peptide YAANEV, which carries the pathological mutation, showed no inhibitory effect (Figure 3.7.E).
As shown earlier, L1.1s as a substrate was a potent inducer of neurite outgrowth from primary spinal neurons (Figure 3.4). The effect of L1.1s on the development of axonal processes was further examined in embryos co-injected with L1.1s mRNA and
L1.1 morpholino. In L1.1 morphants, axonal growth from trigeminal ganglion neurons was defective, as revealed by immunostaining with mAb zn-12 (Figure 3.6.F, pane b).
When L1.1s mRNA was co-injected with L1.1 morpholino, axonal projections from the trigeminal ganglia in the injected embryos remained underdeveloped. Only a few short processes were observed (Figure 3.6.F, panel c). In contrast, co-injection of L1.1 full- length mRNA rescued axonal growth in morphants, which displayed extended axonal arbors from the trigeminal ganglion neurons into the midbrain and forebrain (Figure 3.6.F, panel d). This rescue function of L1.1 is consistent with the expression pattern of L1.1
(Figure 3.5.B) and its role in axonal growth and guidance (Maness and Schachner, 2007).
Therefore, L1.1s appeared to influence only the brain domain organization and ventricle
development, but not axonal projections from neurons.
158 Requirement of Ig1 domain for L1.1s function in brain ventricular development
The data so far showed that the L1.1s protein was secreted from cells and bound to the cell membrane as a dimer, and that L1.1s mRNA was able to rescue morphant defects in early brain formation. It was, therefore, of interest to determine whether supplementing soluble protein L1.1s to L1.1 morphant brain might rescue ventricle development. His6-tagged fusion proteins, including L1.1s, individual Ig1 or Ig2 domain,
were injected separately into the midbrain of L1.1 morphant at 24 hpf and the
morphology of ventricles was examined 3 hours afterwards (Figure 3.8.A). Whereas the
L1.1 morphant midbrain showed poorly defined ventricle with cells bearing unpolarized
centrosomes (Figure 3.8.C), morphants with local application of the L1.1s protein showed restored ventricular polarity in the midbrain (Figure 3.8.D). Notably, the Ig1 domain alone (Figure 3.8.E), but not Ig2 (Figure 3.8.F), was capable of restoring proper development of the midbrain.
Involvement of L1.1-Nrp1a interaction in embryonic brain development
Previous cell binding assays showed that recombinant L1.1s bound specifically to
COS7 cells expressing Nrp1a. Domain Ig1 achieved a similar binding level to Nrp1a as
L1.1s. To determine whether L1.1s-Nrp1a interaction was functionally involved in brain ventricle development, different amounts of the inhibitory peptide YAANEL were added to the embryo medium at 17 hpf, preceding the initial opening of the brain ventricle.
Ventricle development in these embryos was analyzed after 5 hours (Figure 3.9.A). At the concentrations of 2 mM and 4 mM, YAANEL resulted in obscure midbrain structures, including tegmentum and tectum (Figure 3.9.B, panels b and f, c and g). Furthermore, γ
159 A Fusion proteins L1.1MO 24 hpf
Develop for 3 hours
Ctrl 27h L1.1MO 27h B C *
e e e e
L1.1MO 24h + protein injection DEFG * *
+ L1.1s + Ig1 + Ig2 + PBS
Figure 3.8. Restoration of ventricular polarization by local injections of recombinant L1.1s. (A) Schematic drawing illustrating the local application of recombinant protein to the morphant brain. At 24 hpf, morphants showing abnormal brain development were injected with purified recombinant L1.1s, Ig1, Ig2, or PBS into the midbrain region. The manipulated embryos were allowed to develop for another 3 hours at 28°C before fixation for phenotyping. (B-G) Transverse sections of midbrain stained by anti-γ tubulin: uninjected control (B) and L1.1 morphant at 27 hpf (C), brains injected with L1.1s (D), Ig1 domain (E), Ig2 (F), or PBS (G). Polarization of supernumerary centrosomes at the ventricular surface was restored in the brains injected with either L1.1s or Ig1. Results were obtained from 6-10 embryos of three independent experiments. Arrows point to the disrupted polarization at the ventricle while asterisks denote the midbrain ventricle; e, eye; Scale bar: 50 μm.
160 AB YAANEL YAANEV 4% DMSO 2 mM 4 mM 4 mM abcd
17 hpf (18 somites) efTe gh Tg Peptide treatment SYTO Bright ijkl -tubulin γ mn o p
~22 hpf (26 somites) M M M F-actin
Figure 3.9. Inhibition of L1.1-Nrp1a interaction by synthetic peptide disrupted ventricle formation in the midbrain. (A) Schematic drawings depicting the in vivo inhibition assay by synthetic peptides. Prior to the commencement of intrinsic ventricle morphogenesis, embryos of 17 hpf were treated with either the inhibitory peptide YAANEL or the control peptide YAANEV for 5 hours till the time when ventricle development came under the influence of the flow of the cerebrospinal fluid. (B) Brain morphology after treatment with the inhibitory peptide. (a-h) Lateral views with rostral to the left. Bright field images of live embryos (a-d) and confocal scans of the SYTO green stained brain (e-h) revealed that ambiguous tectum and tegmentum structures in embryos treated with YAANEL at either 2 mM or 4 mM (white arrows). (i-l) Transverse section of midbrains stained with anti-γ tubulin. Each treatment were repeated five times independently and each set contained 5-10 embryos. Open arrowheads denote disrupted polarity of the ventricle. (m-p) Dorsal view of flat- mounted embryonic brains stained with F-actin to reveal the ventricle opening, rostral being to the left. Open arrow indicates impeded formation of midbrain ventricle in 4 mM YAANEL-treated embryos. Te, tectum; Tg, tegmentum; M midbrain. Scale bar: 50 μm.
161 tubulin staining showed that the supernumerary centrosomes in the periventricular cells became aberrantly unpolarized (Figure 3.9.B, panels j and k). In addition to altered brain domain organization and abnormal ventricular polarity, ventricle opening in the midbrain region was compromised as revealed by F-actin staining. Thus, failure in ventricle formation was prominent in embryos treated with 4 mM YAANEL (Figure 3.9.B, panel o). In contrast, embryos treated with 4 mM of the control peptide YAANEV did not exhibit notable aberrations (panels d, h, l, p of Figure 3.9.B).
Effects of L1.1-Nrp1a interaction on neuronal cell positioning at the diencephalic ventricle
During early brain ventricle formation, NPCs proliferate at the ventricle edge and migrate away from the ventricle to a destination in a deeper layer (Nadarajah and
Parnavelas, 2002). Cell polarity affects cell migration and proliferation in the ventricular zone (Schaar and McConnell, 2005; Solecki et al., 2006; Ueno et al., 2006). Since
YAANEL interrupted ventricle formation, its effects on ventricular polarity might result in defective cell migration in the developing brain ventricle. To address this issue, in vivo
BrdU pulse-chase assays were performed. Proliferating cells at the brain ventricle of 48
hpf embryos were pulse-labeled with BrdU and the positioning of labeled cells was
examined 5 hours after peptide treatment (Figure 3.10.A). When embryos were fixed
immediately after BrdU labeling, the labeled cells were predominantly situated at the
ventricular surface (data not shown), consistent with several recent reports (Mueller and
Wullimann, 2002; , 2003). Five hours later, labeled cells originated from the
diencephalic ventricle had migrated away, situated at a distance from the ventricle
162 A BrdU pulse labeling Peptide treatment 5 h WT 48 hpf
B a b 4% DMSO 4%
c d 2mM YAANEL 2mM YAANEV (ctrl) 2mM YAANEV ef L1.1MO 48h+5h Nrp1aMO 48h+5h
C 35 * 30 25 20 m)
μ 15 ( 10 n=5 n=7 n=5 5 0 Distance from the ventricle O V L S E E N M N A D A % YA YA 4 mM mM 2 2
163 Figure 3.10. Involvement of L1.1-Nrp1a interaction in neuronal migration at the diencephalic ventricle. (A) Scheme drawing depicting the in vivo neuronal migration assay. Embryos of 48 hpf were first pulse-labeled with BrdU, followed by peptide treatment in the presence of 4% DMSO for 5 hours. (B) Transverse sections at the midbrain level immunostained with antibodies against BrdU (purple) and γ-tubulin (green). (a) Embryo treated with 4% DMSO with the diencephalic ventricle region (boxed) shown at higher magnification (b). (c and d) Diencephalic ventricle in embryos treated with 2 mM of either YAANEV or YAANEL. Diencephalic ventricle of L1.1 morphants (e) and Nrp1a morphants (f) are shown at 5 hours after pulse- labeling with BrdU. Scale bar: 50 μm. (C) Histogram showing distances of migratory cells from the ventricle. Data represent the mean ± s.d, and the numbers of embryos examined are as indicated. Student’s t-test show significant difference between YAANEV-treated embryos and YAANEL-treated embryos; *, P<0.01.
164 (Figure 3.10.B, panels a and b). However, upon treatment with the YAANEL peptide at 2 mM, most of the BrdU-labeled cells remained in the vicinity apposing the ventricle
(Figure 3.10.B, panel d). In embryos treated by the control peptide YAANEV, labeled cells were located at a distance from the ventricle comparable to that of the buffer control
(Figure 3.10.B). When the greatest distance from the leading edge of the migratory cells to the ventricle surface was measured, the distance in the YAANEL-treated embryos was greatly reduced with an average of 14.0 μm (from 5 embryos) in comparison with that in the YAANEV-treated embryos (27.6 μm, averaged from 7 embryos) (Figure 3.10.C). In
L1.1 and Nrp1a morphants, BrdU-labeled cells were also inhibited from migrating away from the ventricle (Figure 3.10.B, panels e and f).
165 DISCUSSION
Our studies have led to the discovery of a soluble isoform of L1 in vertebrates.
Zebrafish L1.1s displays a distinct expression pattern at the ventricular surface and is
involved in the regulation of the ventricle formation during brain development. In vivo functional studies have provided evidence in support of a critical role for L1.1s in maintaining polarity and promoting cell migration in the brain ventricle via its interaction with Nrp1a.
L1.1s is secreted from cells as a matrix protein. Since recombinant fragments derived from the extracellular portion of L1 are known to promote cell migration and axonal guidance (Castellani et al., 2000; Maretzky et al., 2005), L1.1s is expected to serve similar functions. Indeed, recombinant L1.1s is a potent inducer of neurite outgrowth from primary neurons. Interestingly, L1.1s associated with the cell surface adopts a dimer form. Dimerization of L1 has been shown to promote oligomerization of the protein to form higher order complexes, which apparently magnify the neuritogenic activity of L1 (Doherty et al., 1995).
An abundance of L1.1s is associated with the neuroepithelium and the protein is confined to the luminal surface of brain ventricles, differing from the expression pattern of the full-length L1.1. Knockdown of L1.1 results in abnormal NPC positioning in the developing ventricle, concomitant with the anomalous ventricular polarization and proliferation (see Chapter II). Significantly, these phenotypes can be rescued by the co- injection of L1.1s mRNA. NPCs arise from the ventricular neuroepithelium and migrate outward away from the midbrain ventricle (Nadarajah and Parnavelas, 2002; Langenberg
166 and Brand, 2005; Langenberg et al., 2006). It is conceivable that L1.1s participates in cell-ECM interactions, which may facilitate the migration and positioning of NPCs.
It has been shown that cell positioning in the developing brain necessitates normal expression and distribution of ECM components. For instance, the ECM protein reelin, which is expressed exclusively in the pioneer neurons, guides neuronal cell migration along the radial array. In the reeler mice that bear a mutation in the reelin gene, neurons fail to reach their correct destination in the developing brain, thus affecting the organization of the cerebellar and cerebral cortices (D'Arcangelo et al., 1995; Hirotsune et al., 1995). In zebrafish, mutations in laminin subunits consistently lead to prominent anomalies that manifest irregularly shaped brain and severely enlarged hindbrain ventricle, suggesting that laminin functions in mediating neuronal migration. Therefore, its malfunctioning disrupts normal ventricle formation (Schier et al., 1996; Parsons et al.,
2002; Paulus and Halloran, 2006).
L1 has also been demonstrated as an important substrate participating in various cell migration processes, such as transendothelial migration of melanoma cells (Beer et al., 1999; Voura et al., 2001) and transepithelial migration of activated leukocytes
(Duczmal et al., 1997). In certain L1 patients, a large number of ventricular cells fail to migrate properly (Kamiguchi et al., 1998a), consistent with the notion that L1 facilitates cell migration at the ventricle. Our studies show that L1.1s deficit renders aberrant cell positioning, supporting a role for L1.1s in the brain patterning during embryonic development.
Also, abrogation of L1.1s-Nrp1a interaction disrupts positioning of cells derived from the diencephalic ventricle neuroepithelia, giving rise to the possibility that this
167 interaction is responsible for cell migration at the ventricular zone in the midbrain. It has
been shown that neuropilins facilitate neuronal cell migration in the developing brain by
sensing repellent semaphorin cues. In the mouse striatum where semaphorins 3A and 3F
are expressed, migrating interneurons lacking neuropilins 1 and 2 are populated
exclusively; whereas those expressing either neuropilins are directed to the cortex (Marin
et al., 2001). Because soluble L1 is able to interfere with the repulsive Nrp1-semaphorin
3a (sema3a) interaction (Castellani et al., 2000), it is possible that the soluble L1 acts as a
matrix component and guides the migration of new born neuron cells by fine-tuning the
cell response to the Nrp1-sema3a interaction.
Results of the peptide inhibitory assay suggest that the interaction between L1.1s
and Nrp1a is indispensable in establishing ventricular polarity that is required for ventricle formation. This function may involve a cytosolic signaling cascade triggered by
Nrp1a. The last three amino acids of neuropilin-1 (S-E-A-COOH) are responsible for
interaction with a neuropilin-1-interacting protein (NIP), which contains a PSD-
95/Dlg/ZO-1 (PDZ) domain and is able to initiate intracellular signals in response to
ligand binding (De Vries et al., 1998; Cai and Reed, 1999). This SEA motif is conserved
from Xenopus to humans. Interestingly, between the two zebrafish Nrp1 homologues,
only Nrp1a contains this binding motif (Bovenkamp et al., 2004). It is conceivable that
interactions between L1.1s and Nrp1a stimulate signals that can influence the cytoskeletal
stability and positioning of the neuronal progenitor cell via regulatory motifs in the cytoplasmic domain of Nrp1a.
The short isoform of L1 is also present in both mouse and human embryonic cells.
It is possible that vertebrate L1s may share conserved functions during the formation of
168 embryonic brain ventricles. In L1 patients, hydrocephalus is frequently associated with stenosis of cerebral aqueduct (Fransen et al., 1997; Weller and Gartner, 2001). However, recent mouse models do not necessarily correlate the blockage of the aqueduct to the resultant enlargement of lateral ventricle. It is possible that functional truncated protein fragments are present in these mutant mice, which are generated by disrupting mouse L1 at either Ig3 domain (Dahme et al., 1997; Fransen et al., 1998a; Rolf et al., 2001) or Ig6 domain (Cohen et al., 1998; Demyanenko et al., 1999). The accumulation of functional fragments in these mouse models may contribute to the low occurrence of hydrocephalus
(4 out of 70 mutant offsprings, Rolf et al., 2001). Indeed, our results suggest that the Ig1 domain is sufficient in establishing ventricular polarity in the L1.1 morphants.
In conclusion, our results provide evidence for a novel soluble isoform of L1 prior to the initiation of brain ventricle formation. Therefore, it has opened up a new and exciting area to explore the mechanisms by which L1 mediates polarity and migration of
NPCs in the ventricular zone of the developing brain.
169
Chapter IV:
Role of L1 during Axonal Path Finding of Primary
Motoneuron in Zebrafish
(All experimental results described in this chapter were obtained by me.)
170 SUMMARY
The neural cell adhesion molecule L1 plays an important role in many aspects of
neural development, including axonal growth and guidance. In this study, knockdown of
L1.1 expression by antisense morpholino oligonucleotides resulted in aberrant projection
of axons from primary motoneurons, which shared similarities with the phenotypes
observed in unplugged mutants. The unplugged gene in zebrafish encodes an ortholog of
muscle specific kinase (MuSK) and plays a pivotal role in postsynapse formation at the
neuromuscular junction. Co-immunoprecipitation and fluorescence localization studies suggested that L1.1 interacts with Unplugged. Fluosphere-to-substratum binding studies show that the heterophilic interaction between L1.1 and Unplugged involves the third Ig- like domain of L1.1. Full-length L1.1 mRNA, but not L1.1s mRNA, was able to rescue abnormal motor axonal growth in L1.1 morphants, suggesting that the full-length protein is required for the development of motoneurons. The pathological mutations G273D and
E314K, which abrogated this specific binding, disturbed axonal growth in vivo. Current results unveiled a role for L1 in the development of neuromuscular junction (NMJ) that is prerequisite for proper axonal pathfinding.
171 INTRODUCTION
The cell adhesion molecule L1 is a member of the immunoglobulin (Ig) superfamily of recognition molecules. L1 is comprised of six Ig-like and five fibronectin type III domains, followed by a single transmembrane domain and a short and highly conserved cytoplasmic tail (Hortsch, 1996). L1 has been recognized as an important molecule in a variety of neuronal processes, including axonal pathfinding and fasciculation ((Maness and Schachner, 2007). Mutations in the human L1 gene result in a broad spectrum of neurological disorders: corpus callosum hypoplasia, retardation,
adducted thumbs, spastic paraplegia and hydrocephalus, collectively known as CRASH
or L1 syndrome (Fransen et al., 1997; Fransen et al., 1998b; Weller and Gartner, 2001).
The pathological mutations spread along the entire protein, implying that individual domains of L1 are involved in one or more of its complex interactions and functions
(Fransen et al., 1998b).
L1 is localized predominantly, but not exclusively, to the developing nervous system. In vertebrates, ranging from fish to mammals, L1 is expressed on all classes of
CNS neurons, Schwann cell, and sensory neurons (Rathjen and Schachner, 1984; Martini and Schachner, 1986; Persohn and Schachner, 1987; Landmesser et al., 1988; Tongiorgi et al., 1995). L1 is targeted to the leading edge of the growth cone of neurons, underscoring its crucial role in axonal growth and navigation (Kamiguchi and Yoshihara,
2001). In L1-deficient mice, corticospinal axons fail to cross the midline because of the lack of responsiveness to the semaphorin3A/neuropilin-1 (sema3A/Nrp1)-mediated repulsion (Dahme et al., 1997; Cohen et al., 1998; Castellani et al., 2000).
172 In the migratory growth cone, L1 exerts its role in supporting adhesion and
sensing the environmental cues via its extracellular interactions with various cell surface
receptors, including L1 itself, integrins, tyrosine kinase receptors, and other Ig
superfamily cell adhesion molecules (Maness and Schachner, 2007). Disruption of L1
extracellular interactions with Fab fragments against L1 causes perturbations in the
growth of retinal ganglion cell axons (Brittis and Silver, 1995). In addition to
extracellular adhesive interactions, the cytoplasmic domain is indispensable for L1
functions (Kamiguchi and Yoshihara, 2001; Kamiguchi, 2003). L1 influences
cytoskeletal dynamics involving several motifs in its cytoplasmic tail, which include the
highly conserved motif FIGQY that recruits ankyrin, through which L1 is linked to the actin filament (Gil et al., 2003; Nishimura et al., 2003; Whittard et al., 2006).
Phosphorylation of the tyrosine in the motif enhances the interaction of L1 with doublecortin, which couples L1 to the microtubules (Kizhatil et al., 2002). Another conserved sequence YRSLE and a juxtamembrane motif (KxxK) have been found to contribute to cytoskeletal remodeling as they proved the docking sites for ezrins (Dickson et al., 2002; Cheng et al., 2005). The YRSLE motif is also involved in the internalization and recycling of L1 by clathrin-coated endocytic vesicles, therefore, is implicated in regulating L1 functions at the migrating growth cone (Kamiguchi and Lemmon, 1998;
Kamiguchi et al., 1998c).
L1 is also enriched in synaptic vesicles and L1-containing presynaptic structures have been found to align with the postsynaptic apparatus at the interface between axon and muscle (Triana-Baltzer et al., 2006). However, little is known about how L1
173 functions in synapse formation, especially when the axon of motoneuron is navigating
along the muscle.
Zebrafish expresses two L1 homologs, L1.1 and L1.2. L1.1 transcript is expressed
in all known classes of neurons, whereas L1.2 is restricted to a subpopulation of neurons
(Tongiorgi et al., 1995). L1.1s, a short transcript variant that encodes the first four Ig
domains of L1.1, is also found in zebrafish (Chapter III). Knockdown of L1.1 by
morpholino oligonucleotides results in aberrant projection of axons from primary
motoneurons, similar to the phenotypes observed in unplugged mutants. In zebrafish,
spontaneous mutations in Unplugged (a zebrafish MuSK ortholog) result in errors in
navigation of primary motor axons in zebrafish (Zhang and Granato, 2000; Zhang et al.,
2004; Lefebvre et al., 2007). Further studies show that L1.1 undergoes heterophilic interaction with Unplugged during zebrafish axonogenesis. Abolition of this interaction by missense mutations in L1.1 that are associated with CRASH syndrome disturbs normal motor axon development.
174 MATERIALS AND METHODS
Zebrafish maintenance
Wild-type zebrafish (Danio rerio) were purchased from local pet store. Fish were
maintained on a 14 hour light/10 hour dark cycle at 28.5°C, according to Westerfield
(2000). Embryos were obtained by spontaneous spawning and staged by hours post fertilization (hpf) or number of somites (Kimmel et al., 1995).
RT-PCR and cloning
A zebrafish cDNA clone containing the coding region of unplugged full length (FL)
isoform (Genbank accession: AY536052) was obtained by RT-PCR (Qiagen) with
muscle total RNA isolated from adult fish. The respective forward and reverse primers
were 5’- CTGATACGAGGCTGACCAATGGA-3’ and 5’-
GCTTGCCCTCTTGAGCATACAAT-3’. The open reading frame of Unplugged FL
fused with enhanced green fluorescent protein (EGFP) at the 3’ end was then cloned
downstream of the promoter for muscle creatine kinase (MCK, Ju et al., 1999). The
plasmid containing this construct (pMCK:Unp-EGFP) was injected into embryos, and it
expressed Unplugged in muscle cells.
For transfection and expression of L1.1 in neuronal cells, the HuC promoter (Park
et al., 2000) was placed upstream of L1.1 cDNA which was tagged with the DsRed
fluorescent protein at the 3’ end (pHuC:L1.1-Red). Mutant L1.1 plasmids carrying
mutations G273D and E314K (corresponding respectively to pathological mutations
G268D and E309K in L1 patients) were generated by site-directed mutation using PCR
175 methods (Stratagene, Mississauga, Ontario) before being cloned into the DsRed reporter system to construct pHuC:L1.1G273D-Red and pHuC:L1.1E314K-Red.
Microinjection
Morpholino oligonucleotides were purchased from Gene tools (Philomath, OR).
Antisense morpholino targeting the 5’- untranslated region (5’-
CAGTCCCGACTCCAGACACACACAC-3’) was used to knockdown L1.1 expression.
The corresponding mismatch control morpholino 5’-CAgCTcCTG
AgAGACAgAAACAgAG-3’ contained four mismatched bases shown in lower case.
Morpholino oligonucleotides were diluted into 4 mg/ml with 2 mg/ml phenol red (pH8.0) and injected into embryos at the 1-2 cell stage (~2 ng per embryo). For mRNA rescue experiments, capped in vitro transcribed mRNAs of L1.1s and full-length L1.1 were diluted to 0.1 μg/μl with 0.2 M KCl. Approximately 50 pg of each mRNA was injected into each embryo in combination with 2 ng of antisense morpholino oligonucleotides. To generate transient transgenic embryos that expressed Unplugged in muscle cells or L1.1 in neurons, 20~40 pg of the corresponding plasmid was injected into embryos at the one- cell stage. Mosaic expression of these plasmids was observed in the injected embryos.
Manipulated embryos were raised in embryo medium containing 1% penicillin/streptomycin (Westerfield, 2000) until 24 hpf and then fixed with 4% paraformaldehyde (PFA).
176 In situ hybridization
Specific digoxigenin (DIG)-labeled riboprobes for L1.1 isoforms (L1.1 and L1.1s, see
Chapter III) were synthesized by in vitro transcription using T7 RNA polymerase and
DIG-RNA labeling mix (Roche) according to manufacturer’s instruction.
Wild-type embryos at 24 hpf were collected and fixed in 4% PFA overnight at 4°C. After
a couple of washes with fresh PBS-Tw (phosphate buffer saline, 0.1% Tween-20), embryos were permeabilized with 10 μg/ml proteinase K (Roche) in PBS-Tw for 5
minutes at room temperature and fixed again in 4% PFA for 20 minutes. After incubation
with hybridization buffer (50% formamide, 5 x SSC, 0.1% Tween-20, 50 μg/ml heparin,
500 μg/ml salmon semen tRNA, 0.092M citric acid, pH 6.0), embryos were hybridized
with 20 ng of DIG-labeled antisense riboporbe in hybridization buffer overnight at 65°C.
To decrease background, embryos were incubated with fresh hybridization buffer for one
hour preceding stringent post-hybridization washes, which included once with 0.2X SSC
at 65°C for 20 min and twice with 0.1X SSC at 65°C for 20 min. For antibody detection,
embryos were incubated with alkaline phosphatase-conjugated anti-DIG antibody
(1:5000, Roche) in blocking buffer (5% sheep serum and 5% BSA in PBS-Tw) overnight at 4°C. Color detection was performed using the NTP/BCIP substrate (Roche) in the
presence of 0.5 mg/ml levamisole (Sigma) until desired color had developed. After
several washes in PBS-Tw, embryos were sectioned manually with a scalpel blade and
mounted in 70% glycerol for microscopy.
177
Immunohistochemistry
Whole mount immunohistochemistry was performed on embryos using affinity-purified
polyclonal antibody anti-L1.1 (5 μg/ml, see Chapter II) and monoclonal antibodies znp-1
(5 μg/ml, Zebrafish International Resource Center, Eugene OR), F59 and SV2 (1 μg/ml,
Developmental Studies Hybridoma Bank). HRP conjugated goat anti-mouse IgG (0.5
μg/ml, Pierce,) and Alexa 568 goat anti-rabbit IgG and Alexa 488 goat anti-mouse IgG
(10 μg/ml Invitrogen) were used as secondary antibodies. The HRP conjugated secondary antibody was detected by enhanced DAB substrate (Pierce) and the embryos were mounted in Permount (Fisher). For fluorescence detection, embryos were embedded in
17% gelatin-10% sucrose and sectioned. Sections were mounted in Fluormount fluorescence mounting medium (DAKO, Mississauga, Ontario) for confocal microscopy.
Co-immunoprecipitation
Wild-type embryos of 24 hpf were lysed in native lysis buffer (50 mM Tris-HCl pH 7.4,
1 mM MgCl2, 2.5 mM EGTA, 2.5 mM EDTA, 10% glycerol, 1% NP-40) in the presence
of protease inhibitors and phosphatase inhibitors. The homogenized material was diluted
10-fold in the lysis buffer without NP-40 and incubated with protein A agarose
conjugated with anti-L1.1 IgG (1 μg/ml), anti-MuSK (30 μl, recognizing the amino acids
529~668 of rat MuSK, Sigma) or pre-immune rabbit IgG (1 μg/ml). The immune
complexes were eluted with 0.1M glycine and separated on SDS-PAGE, followed by
western blot analysis using rabbit anti-L1.1 IgG (1 μg/ml) or anti-MuSK IgG (2 μg/ml),
178 which recognizes the epitope containing amino acids 844~859 of the human MuSK
(Stratagene).
Production of bacterial fusion proteins
To construct His6-tagged fusion proteins, the following cDNA sequences were cloned
into the pET-28(a) vector with a fused His6-tag at the 3’ end: extracellular portion of
mature Unplugged, L1.1 domains Ig 1-4, Ig 1-6, Ig1, Ig2, Ig3, and Ig3 mutants (Ig3G273D
and Ig3E314K). Unplugged, Ig1-4 and Ig1-6 fusion proteins were produced in E.coli and
purified by Ni-NTA chromatography (Qiagen) under denaturing conditions before being
refolded as described previously (Chapter II). His6-tagged fusion proteins containing individual domains of L1.1 were soluble, and were purified under native conditions
according to manufacturer’s instructions.
Fluosphere-to-substratum binding assay
Fusion protein containing the extracellular portion of unplugged was conjugated to Alexa
488 fluospheres (0.5 μm in diameter, Invitrogen) according to the manufacturer’s
instruction and stored in blocking buffer (50 mM PBS, 2% BSA, 2% dextran, 0.1%
Triton X-100, 1% glucoside).
Fusion proteins of L1.1 (1 μM) were used to coat on Superfrost Plus slides (VWR)
overnight at 4°C. Unplugged-conjugated fluospheres were diluted to 0.005% slurry in
blocking buffer and incubated with the substratum for 30 min in room temperature with
gentle agitation. Unbound and loosely attached beads were removed from the substratum
by PBS-T (0.5% Triton X-100 in PBS). Bound beads were then fixed with 4% PFA and
179 mounted in DAKO. Images of bound fluospheres were captured using fluorescence microscope (Nikon ECLIPSE TE2000). Quantification of fluospheres was performed using ImageJ (National Institutes of Health). Data were collected from four independent experiments, each with four replicates.
180 RESULTS
Differential expression of L1.1 and L1.1s in the zebrafish trunk
To investigate the role of L1.1 in the development of axons from primary
motoneurons, the expression of L1.1 isoforms, L1.1 and L1.1s, in the trunk was
examined. In situ hybridization was used to investigate the role of L1.1 during
development of axons from primary motoneurons. Riboprobes specific for L1.1s or L1.1 mRNAs were used in these studies and revealed different expression patterns for the two
L1.1 isoforms. The transcripts of full-length L1.1 were exclusively expressed in the
spinal cord, indicating its neuronal origin (Figure 4.1.B). However, Ll.1s mRNA was
expressed predominantly in muscle cells, and it was absent from the spinal cord (Figure
4.1.B).
Whole mount immunohistochemistry was performed using affinity-purified anti-
L1.1 IgG to assess the distribution of L1.1 proteins. Fluorescence labeling was revealed
in neurons in the spinal cord and their axonal projections along the medial-lateral somites
(Figure 4.1.C). At 24 hpf, there are three primary motoneurons in each hemisegment of
the spinal cord, namely, rostral primary (RoP), middle primary (MiP) and caudal primary
(CaP), which extend their axons out of the spinal cord along a common pathway which is
pioneered by that of the CaP. After reaching the muscle pioneer cells, the axonal
trajectory of the CaP motoneuron courses ventrally while the MiP axon turns dorsally
(Hjorth and Key, 2002). The staining patterns in Figure 4.1.B showed that L1.1 was
enriched at neuronal synapses demarcated by the presynaptic vesicle marker SV2 (Feany
et al., 1992). Interestingly, L1.1 staining also coincided with the staining of F59 (Figure
181 A C L1.1 SV2 Merged CaP somite abc SC RoP SV2 SC SC MiP + NC NC NC Anti-L1.1 somite
D L1.1 F59 Merged abc B SC SC
L1.1 L1.1s NC NC F59
a b +
SC
NC
Anti-L1.1 def In situ NC NC
Figure 4.1. Differential expression of L1.1 isoforms in spinal neurons and somites. (A) Schematic illustration of axon trajectories from primary motor neurons at 24 hpf (Eisen, 1991a). The three axons share a common path from the spinal cord to the choice point. At the choice point, the CaP axon continues into the ventral myotome, the MiP axon sprouts a dorsal branch, and the RoP axon pauses at the choice point. (B) Whole mount in situ hybridization using riboprobes specific for L1.1s and L1.1. L1.1 was expressed exclusively in the spinal neurons (a), while L1.1s was detected primarily in the somites (b). (C, D) Whole mount immunohistochemistry showing colocalization of L1.1 with SV2 and F59, respectively. (C) Neuronal L1.1 was enriched in the pre-synaptic vesicles labeled by SV2. In the merged image (c), L1.1 is shown in red and SV2 in green. Red dash lines define the contour of the spinal cord. (D) L1.1 antigen was associated with adaxial cells (a-c) and pioneer muscle cells (d-f). Merged images (c and f) show the colocalization of L1.1 (red) with F59 (green). Arrows denote the position of pioneer muscle cells. SC, spinal cord; NC, notochord. Scale bars: 20 μm.
182 4.1.D), which reveals adaxial cells, the slow muscle precursors (Miller et al., 1985).
These results demostrate a mutually exclusive pattern of expression for L1.1 and L1.1s.
Whereas L1.1 is expressed in the spinal cord neurons, L1.1s is expressed primarily in the
adaxial cells.
Involvement of L1.1 in the development of primary motoneuron
To further characterize the role of L1.1 in motor axonal growth, expression of
Ll.1 was inhibited using antisense morpholino oligonucleotides. Expression of both L1.1
and L1.1s in the embryo was knocked down successfully using oligonucleotides against
the 5’UTR of L1.1 (see Figure 3.6.C in Chapter III). Motoneurons in L1.1 morphants
were examined by immunostaining with mAb znp-1, which specifically recognizes
primary motoneurons and their axons (Trevarrow et al., 1990). In control embryos, the
CaP axon elongated ventrally along the medial position of the somite, and branched only
after it reached the ventral portion of the somite (Figure 4.2.A and B). In morphants,
axons from CaP motoneuron displayed irregular morphologies, including truncation and ectopic branching along the axonal path (Figure 4.2, C and D). Notably, some of these axonal extensions were stalled or branched at the position of muscle pioneer cells.
Among the axonal projections from the CaP motoneuron in the L1.1 morphant, 33% of them appeared shortened and 19% showed ectopic branching. Furthermore, the normal morphology of the MiP axon that extends dorsally was not observed in the L1.1 morphant.
Because L1.1 and L1.1s were knocked down simultaneously by the morpholino
(see Figure 3.6.C in Chapter III), it raised the question whether L1.1s or L1.1 was
183 A B
Uninjected Ctrl MO C D
L1.1MO L1.1MO E F
L1.1MO + L1.1 mRNA L1.1MO + L1.1s mRNA
Figure 4.2. Aberrant motor axonal growth in L1.1 morphants. Embryos at 24 hpf were immunostained with mAb znp-1. Lateral views of the trunk are shown with caudal to the right: (A) uninjected embryo, (B) embryo injected with the 4-mismatch control morpholino, (C and D) L1.1 morphants, (E) embryo co-injected with L1.1 mRNA and L1.1 morpholino, and (F) embryo co-injected with L1.1s mRNA and L1.1 morpholino. Results were obtained from 5 independent experiments, each of which included 8-15 embryos. Arrowheads shortened CaP axonal projections, while arrows denote axons bearing ectopic branches. Scale bars: 50 μm.
184 responsible for the abnormal development of motoneurons. To address this issue, L1.1
morpholino was co-injected into embryos with either L1.1 or L1.1s mRNAs. The
rescuing effects of L1.1 and L1.1s are shown in Figure 4.2.E. In the embryos that
received a combined injection of L1.1 mRNA and L1.1 morpholino, axons of
motoneurons displayed normal morphology, suggesting that L1.1 rescued the morphant
phenotype. However, in embryos coinjected with L1.1s mRNA and L1.1 morpholino,
CaP projections were aberrantly truncated and bore ectopic branches (Figure 4.2.F). The
data showed that L1.1s failed to support proper axonal growth in the morphants. Thus,
neuronal L1.1, and not L1.1s, appears to be responsible for motor axonal growth and
navigation (Table 4.1).
Association of L1.1 with Unplugged in developing motor axon
The phenotypic abnormalities of axonal growth from the CaP in the L1.1
morphants showed resemblance to the defects observed in Unplugged mutants (Zhang
and Granato, 2000). Unplugged is the zebrafish ortholog of MuSK, which is expressed in
adaxial muscle cells and mediates synaptic differentiation at the NMJ (Zhang et al., 2004;
Lefebvre et al., 2007). Deficits of L1.1 and Unplugged gave rise to similar phenotypes,
raising the possibility that L1.1 interacts with the muscular Unplugged during motor
axonal growth. To test this hypothesis, co-immunoprecipitation was performed using anti-L1.1 antibody, followed by immuoblotting with anti-MuSK IgG. In the immunoprecipitates, a 104 kDa protein was recognized by the anti-MuSK antibody,
corresponding to the full-length form of Unplugged (Figure 4.3.A). The splice variant-1
(SV1) of Unplugged was not detected in the precipitates. Conversely,
185
Table 4.1. Abnormal axonal projection from CaP in L1.1 morphants of 24 hpf.
Shortened Branching # of fish CaP counted axons axons
Wild-type 14 224 0 1
L1.1MO 9 144 48 (33%) 27 (19%)
L1.1MO+L1.1 mRNA 8 128 0 1 (1%)
L1.1MO+L1.1s mRNA 10 160 103 (64%) 75 (47%)
Note: Axons from the fourth to the eleventh CaP motoneurons were included. In total, 8 axons on each side of the somite and 16 axons in each embryo were counted.
186 A
IP: anti-L1.1 Crude lysate Crude Control IP L1.1 IP L1.1
115 kDa Unplugged FL 82 kDa
Blot: anti-MuSK
B
IP: anti-MuSK MuSK IP Control IP Crude lysate Crude L1.1 182 kDa
115 kDa
82 kDa Blot: anti-L1.1
Figure 4.3. Co-immunoprecipitation of L1.1 with Unplugged FL. Lysate of 24 hpf embryos was immunoprecipitated using (A) anti-L1.1 IgG or (B) anti-MuSK antibodies. The protein blots were probed for the presence of MuSK and L1.1, respectively. The full-length form of Unplugged (104 kDa) was co- immunoprecipitated with L1.1, and the full-length form L1.1 (200 kDa) was brought down together with Unplugged. The 115 kDa band in the crude lysate might represent cross-reacting material.
187 immunoprecipitation of Unplugged resulted in the co-precipitation of the 200 kDa species of L1.1, corresponding to the full-length L1.1 (Figure 4.3.B). The 140 kDa fragment of L1.1, which is processed by proteolysis and composed of the N-terminal portion (Faissner et al., 1985; Pradel et al., 1999), was not detected in the precipitates, suggesting that only the membrane-bound full-length L1.1 was associated with
Unplugged.
Association of Unplugged and L1.1 was further characterized in embryos that expressed EGFP-tagged Unplugged in the muscle fiber. In the embryos carrying
MCK:Unp-EGFP, Unplugged full-length protein was distributed en passant on muscle fibers in apposition to the axonal path. Unplugged was particularly enriched in regions predominantly constituted by L1.1 (Figure 4.4).
Involvement of the Ig3 domain in L1.1-Unplugged interaction
Whether L1.1 and Unplugged interact by direct binding to each other was examined using the fluosphere-to-substratum assay. Fluospheres coupled with the extracellular domain of Unplugged were applied on various substrate-coated L1.1 recombinant proteins. Proteins containing Ig1-6 and Ig1-4 domains displayed similar levels of binding to Unplugged (e and f in Figure 4.5.B). Interestingly, among the substrates containing single domains of L1.1, Unplugged showed the highest level of binding to Ig3 (panel d in Figure 4.5.B), whereas Ig1 and Ig2 exhibited little binding capability (panels b and c in Figure 4.5.B). However, binding of Unplugged-conjugated fluospheres was reduced to the background level on the Ig3 proteins carrying mutations
188 MCK:Unp-EGFP L1.1 Unp L1.1 Unp L1.1 ABCD
20 μm 10 μm
Figure 4.4. Colocalization of L1.1 and Unplugged FL along the axonal path. (A) Cross section of transient transgenic MCK:Unp-EGFP embryos at 24 hpf. The boxed area is enlarged to show colocalization of L1.1 and Unplugged in B-D. (B) Muscle specific expression of Unplugged FL was revealed by GFP fluorescence. (C) L1.1 was detected by immunohistochemistry. (D) The superimposed image (Unplugged in green and L1.1 in purple) shows enrichment of Unplugged FL apposing the L1.1 puncta. Scale bar: 10 μm.
189 A -6 -4 73D 14K Unp Ig1 Ig1 kDa Ig1 Ig2 Ig3 G2 E3 kDa 20 100
15 73 54 10 40 35
25 B
BSA Ig1 Ig2 Ig3
Ig1-4 Ig1-6 Ig3G273D Ig3E314K
C
5000 4000 2 3000 /mm 2000 1000
Number of bound beads 0 BSA Ig 1 Ig 2 Ig 3 Ig1-4 Ig1-6 Ig3G273DIg3E314K
Figure 4.5. Direct binding between L1.1 and Unplugged. (A) Coomassie blue stained gel profiles of His6-tagged fusion proteins purified from E. coli: individual Ig1, Ig2 or Ig3 domains of L1.1, Ig3 mutants bearing the G273D and E314K mutations, Ig1-4, Ig1-6, and the complete extracellular portion of Unplugged. (B) Representative micrographs of Unplugged-fluospheres bound on substrate-coated (a) 1% BSA, (b) Ig1, (c) Ig2, (d) Ig3, (e) Ig1-4, (f) Ig1-6 domains of L1.1, and the two mutant forms of Ig3, (g) Ig3G273D and (h) Ig3E314K. (B) Quantification of fluospheres bound per unit area (mm2) on various substrates. Data represent the mean ± S.E.M. based on four different sets of experiments.
190 G273D and E314K (panels g and h in Figure 4.5.B). The data suggested that the Ig3
domain was responsible for L1.1 direct interaction with Unplugged.
Effect of CRASH mutations on L1.1 mediated axonal growth
Abolition of L1.1-Unplugged interaction by mutations in the Ig3 domain raised
the possibility that these mutations might disrupt axonal development mediated by L1.1-
Unplugged interaction. The effects of L1.1G273D and L1.1E314K on axonal growth were
analyzed in the embryos injected with either wild-type HuC:L1.1-Red plasmid or mutant
(pHuC:L1.1G273D-Red or pHuC:L1.1E314K-Red) plasmid together with L1.1 morpholino.
As shown by immunostaining using znp-1 mAb, the axonal trajectories appeared normal
in the CaP neurons that expressed HuC:L1.1-Red (shown in red fluorescence in Figure
4.6.D), confirming the earlier results that demonstrate L1.1 capability of rescuing the
morphant phenotype.
In embryos co-injected with mutant pHuC:L1.1E314K-Red and L1.1 morpholino, the red fluorescence which showed the distribution of the mutant L1.1 protein, was observed along the axon trajectory in a similar manner as the wild-type protein. However, axonal projections from CaP neurons that expressed L1.1E314K-Red, were shortened and
ectopically branched (Figure 4.6.G). Similar abnormalities were observed in the pHuC:L1.1G273D-Red injected embryos. The L1.1G273D-Red-expressing CaP motoneurons
extended branched and truncated axons. Moreover, the mutant L1.1G273D-Red protein
seemed to be retained in the cell body (panel F in Figure 4.6). These results suggest that both G273D and E314K mutations abrogated the normal function of L1.1 during CaP axonal development.
191 ACB
ControlL1.1MO L1.1MO DE
L1.1MO+HuC:L1.1-Red L1.1MO+HuC:L1.1-Red FG
L1.1MO+HuC:L1.1G273D-Red L1.1MO+HuC:L1.1E314K-Red
Figure 4.6. Effects of pathological L1 mutations on the development of motor axons. Embryos were co-injected with L1.1 morpholino and either wild-type or mutant HuC:L1.1-Red plasmids. Embryos of 24 hpf were immunostained with mAb znp-1. Lateral views are shown with caudal to the right. The znp-1 signal is shown in green, L1.1 in red, and the overlaid signal in yellow. (A) Uninjected embryo and (B and C) L1.1 morphants. (D and E) Exogenous wild-type L1.1 was co-injected with L1.1 morpholino, and restored normal axon development. Notably, axons that failed to express L1.1 appeared abnormal. (F) Mutant L1.1G273D-Red and (G) mutant L1.1E314K-Red were injected together with L1.1 morpholino. Neurons expressing either mutant proteins extended abnormal axons. Arrows indicate the ectopic axonal branches. Arrowheads denote shortened axons. Open arrowheads demarcate the axons epxressing L1.1-Red fusion proteins. Open arrows show the accumulation of mutant L1.1G273D-Red in the neuronal cell body. Scale bar: 50 μm.
192 DISCUSSION
Current study has provided new evidence in support of a role for L1 in axonal
pathfinding from the primary motoneuron, possibly via its interaction with the
postsynaptic component MuSK. This heterophilic interaction of L1 can be disrupted by mutations in the Ig3 domain, which resulted in abnormal axonal projections from the CaP motoneurons.
L1.1 is enriched in synaptic vesicles along the motor axon, consistent with the notion that L1 is involved in establishing synapses during axon navigation
(Godenschwege et al., 2006; Triana-Baltzer et al., 2006). Immunostaining of L1.1 studies showed that L1.1 is colocalized to the regions where the muscular Unplugged fusion protein is expressed. In addition, co-immunoprecipitation studies suggest that a L1.1- containing complex may interact with Unplugged. This is compatible with the observation that L1 is juxtaposed to the AChR-rich postsynaptic apparatus at NMJs in the rat and chick (Triana-Baltzer et al., 2006).
Postsynapse differentiation at the NMJ has been ascribed to the activation of
MuSK in response to the neurotransmitter Agrin, which recruits acetylcholine receptor
(AChR) to the neuromuscular interface (Fuhrer et al., 1997; Burden, 2002). Defects in
MuSK abolish postsynaptic differentiation in vivo (Lin et al., 2001; Lefebvre et al., 2007).
In vertebrates including zebrafish, MuSK activation by an unknown mechanism, which is independent of Agrin and the presence of motor axons, induces AChRs clustering to the central region of the muscle (Lin et al., 2001; Yang et al., 2001; Lefebvre et al., 2007). In zebrafish, prepatterned AChR clusters have been observed on the axonal path in advance of CaP motor axons extending from the spinal cord (Panzer et al., 2005). In vivo imaging
193 has revealed that motor axon growth cones and filopodia are selectively extended toward these small preformed postsynaptic clusters and form contacts with them, whereby synaptogenesis occurs following the rapid engagement of presynaptic vesicles and additional postsynaptic assembly (Panzer et al., 2006). The implication here is that a neuromuscular recognition adhesion is required for guiding navigation of the axonal growth cone on this prepatterned path. It is conceivable that, during the guidance of CaP axons along the MuSK-patterned postsynaptic apparatus, L1 interaction with MuSK contributes cell-cell adhesion between the migratory growth cone and the muscle membrane. In supporting this hypothesis, our results have demonstrated clearly that L1 deficiency results in abnormal axonal growth, a phenotype similar to the defects in
Unplugged mutants (Zhang and Granato, 2000; Zhang et al., 2004).
The direct interaction of L1.1-Unplugged was mapped to the Ig3 domain of L1.1.
Mutations G273D and E314K in Ig3 that disrupted L1-Unplugged interaction were shown to affect CaP axon pathfinding. Therefore, Gly273 and Glu314 in the Ig3 domain may contribute to the interaction interface for L1.1 binding to Unplugged. Both L1.1s and
L1.1 possess this putative binding site for Unplugged, however, only the full length form of L1.1 was able to rescue the abnormal axonal projection from the CaP. It is possible that the extracellular interactions of L1.1 might induce cytoplasmic interactions that were required for axonal growth from the CaP. Consistent with this notion, in lad-1 (an
L1CAM homolog in C. elegans) mutants that produce truncated molecules prior to the cytoplasmic domain, dorsal D (DD) and ventral D (VD) motoneurons exhibit abnormal growth and branching in their dorsal projections (Wang et al., 2005). Accordingly, our co-immunoprecipitation experiments demonstrated that L1.1 and Unplugged were both
194 presented as full-length molecules interacting at the membrane of NMJs. The cytoplasmic domain of L1 contains potent features that enable signal transduction and cytoskeletal remodeling. First, the sequence FIGQY constitutes a binding site for ankyrin which couples L1 to the actin cytoskeleton (Davis and Bennett, 1994; Boiko et al., 2007). Also,
FIGQY has been shown to interact with doublecortin, a microtubule-associated protein
(Kizhatil et al., 2002). Phosphorylation of the tyrosine is critical in L1 functions, because mutation of this residue is associated with CRASH syndrome (Fransen et al., 1997). The phosphorylation state of this sequence is shown to abolish ankyrin binding but increase doublecortin interaction, therefore influencing the cytoskeleton dynamics in the migrating growth cone (Garver et al., 1997; Kizhatil et al., 2002). On the other hand, tyrosine phosphorylation in the motif FIGQY of L1 is found concentrated at the neuromuscular junction, which is overlaid with AChR localization (Jenkins et al., 2001). In the
Drosophila central nervous system, mature synapse formation requires this conserved sequence of Neuroglian (a L1-like molecule in Drosophila), ascribed to normal phosphorylation of the sequence (Godenschwege et al., 2006).
The importance of the full-length form of L1.1 on axonal growth and guidance may also be ascribed to its capabilities of transmitting extracellular interaction to intracellular signal cascades that modulate cell behavior. During the decussation of corticospinal axon tracts in response to repellant semaphorin 3A, the trans interaction between L1 and neuropilin-1 is shown to activate the nitric oxide synthase and cGMP guanylyl cyclase, which consequently converts the sema3A effect into an attractive cue
(Castellani et al., 2000; Castellani et al., 2002). Furthermore, during neurogenesis and neuroprotection, substrate L1 interacts with integrin receptors and activates signal
195 transduction pathways that involve Src family kinases, PI3 kinase, as well as mitogen-
activated protein kinases (Loers et al., 2005; Maness and Schachner, 2007). In the
process of synapse formation at the NMJ, it is possible that the L1.1-Unplugged
interaction activates the kinase activity of Unplugged.
However, the L1-Unplugged interaction may be transient. In zebrafish, expression of Unplugged mRNA is found in adaxial cells and then decreases as they start radial migration (Zhang et al., 2004), whereas L1 has been found in neuronal synaptic vesicles
at the NMJ (Triana-Baltzer et al., 2006). Therefore, L1 may play various roles during
postsynaptic differentiation. A future challenge will be the identification of L1 binding
components that mediate maintenance of the synapse at the neuromuscular junction.
196
Chapter V:
Conclusions and Future Perspectives
197 Normal development of the nervous system necessitates the temporospatially precise tuning of cellular interactions, which are involved in neural induction,
proliferation and migration of neuronal cells, subsequently in their differentiation, axonal
navigation and synaptogenesis, and ultimately in programmed cell death. Neuronal recognition molecules are crucial elements in shaping the precision of neural functions, providing not only adhesiveness by interacting with molecules on adjacent cells or the
ECM, but also their impingement on intracellular signal transduction and cytoskeletal dynamics.
The main objective of my thesis project was to investigate L1 functions in early embryonic neural development in zebrafish, which is a newly emerged vertebrate model organism. In particular, major efforts have been made to elicit the attributes of L1 that are poorly understood in other vertebrates. It has been a decade since the first mouse model for L1 deficiency was generated (Dahme et al., 1997). However, the pathological
mechanism for L1-associated hydrocephalus has not been elucidated by mouse models, largely due to great variations in phenotype and the low incidence of ventricle enlargement in mutant mice (4/70 in mutants truncated in Ig3; and 1/10 in conditional knockout mutants with a tetracycline-controlled transactivator inserted into the exon 2)
(Rolf et al., 2001). In contrast, our studies show a high occurrence (~55%) of the enlargement of the fourth ventricle in L1.1-deficient zebrafish embryos. The difference in the incidence rate may be due to the efficiency of the knock-down approach, which eliminates residual amounts of L1 fragments possessing partial function.
198 Differential functions of zebrafish L1 homologs in embryonic development
Full-length cDNAs for the two zebrafish L1 homologs, L1.1 and L1.2, were cloned at the beginning of my project. Despite the high degree of sequence similarity
L1.1 and L1.2 and their shared neuritogenic activities, many functions that are conserved among vertebrate L1 seem to belong exclusively to L1.1 and not L1.2. Knockdown of
L1.1 in the embryo gave rise to a spectrum of abnormalities, several of which phenotypically resemble the CRASH syndrome; whereas knockdown of L1.2 did not lead to similar phenotypes.
Furthermore, a novel isoform of L1.1, which consists of the first four Ig domains of L1.1 was discovered and its critical functions in brain ventricle formation were studied.
Interestingly, a soluble isoform of L1 is also detected in the mouse embryo and the human embryonic cell line HEK293. These results suggest that L1s may play a pivotal role in brain functions via a mechanism that is conserved among vertebrates.
Role of L1s in brain ventricle formation and the pathological mechanism of L1- associated hydrocephalus
L1.1s transcripts were prominently expressed in the neuronal cells apposing the ventricle and the protein functions at the luminal surface of these cells. Periventricular expression of L1.1s may be a prerequisite for brain ventricle formation. The distribution pattern of L1.1s in the midbrain ventricle coincides with that of Nrp1a (Kawakami et al.,
1996; Bovenkamp et al., 2004). Nrp1 is one of the few recognition molecules that have been characterized in neuronal progenitor cells and it has been implicated in various developmental processes of neuronal cell migration as well as axon guidance (Takahashi
199 et al., 1998; Marin et al., 2001; Luo et al., 2002; Kuan et al., 2007). L1 interaction with
Nrp1 has been demonstrated in mediating decussation of the corticospinal tract
(Castellani et al., 2000; Castellani et al., 2002). My studies demonstrate that disrupting the interaction between L1.1s and Nrp1a or knocking down either protein gives rise to similar morphological abnormalities in the embryonic brain, implying that Nrp1a is the membrane receptor which facilitates the role of L1.1s in orientating neural progenitor cells (NPCs) in the cortex. Intriguingly, as an ECM component apposing the ventricle,
L1.1s exists as a dimer when it is associated with the cell membrane, suggesting that the interaction with its surface receptor induces dimerization of L1.1s. It is conceivable that dimeric L1.1s elaborates adjacent interaction complexes to form clusters and thereby propagates adhesiveness.
A schematic illustration of how L1.1s may exert its influence during the formation of brain ventricles is presented in Figure 5.1A. At the onset of ventricle morphogenesis, NPCs are polarized with respect to the cytoskeleton, as well as the differential distribution of junctional components and cell fate determinants (Chenn et al.,
1998; Reugels et al., 2006). As revealed by immunohistochemistry, L1.1 antigen is positioned to the primitive ventricular surface of the NPC. Because the periventricular cells only express L1.1s but not L1.1, it is suggested that L1.1s presents at the ventricular surface, possibly via its interaction with a cell surface receptor. Abrogation of this interaction leads to the abnormal orientation of NPCs. When the NPCs begin to migrate outward, L1.1s anchors the rear process of the NPC to the luminal surface. Leaving the ventricular zone, NPCs retract the trailing process from the ventricular surface. After the first radial migrating wave of NPCs have left the ventricular zone, a new wave of NPCs
200 ABWild type brain L1.1 morphant brain
Stage 1: Initial ventricle morphogenesis
Stage 2: Early ventricle inflation
Stage 3: Continuous ventricle expansion
CSF CSF
Legend: L1.1s dimer Cell surface NPC receptor
Figure 5.1. Involvement of L1.1s in the development of brain ventricle. (A) In the zebrafish, development of the midbrain ventricle apparently involves three steps. L1.1s exerts its roles in defining the polarity of NPC, as well as in promoting coordinated cell migration and proliferation. At the onset of ventricle morphogenesis, NPCs are polarized with L1.1s localized to the ventricular surface, which may bind to a cell surface receptor on the NPCs. When the NPCs start to migrate, their rear process remains anchored to the ventricle surface via L1.1s interaction. Following the arrival of the first migration wave of NPCs at the preplate, a new wave of postmitotic neurons begin to migrate. As CSF starts to flow after the heart beats at 24 hpf, the combinatory effects of ventricular cell migration and CSF pressure further expand the ventricle. (B) In the absence of L1.1s, NPCs become poorly polarized and disorientated in the cortex. Migration of the NPCs is impeded and proliferation of a new wave of NPCs is inhibited. Consequently, the ventricles become disorganized and the ventricular passage for CSF is obstructed. As the local pressure increases in the fourth ventricle, it gives rise to the hydrocephalus-like phenotype.
201 that have completed mitosis at the ventricle surface start to migrate radially. L1.1s may
continuously exert its influence on the polarization of NPCs. As CSF starts circulating
after the heart beats at 24 hpf, the additional effect of pressure associated with CSF
circulation further expands the ventricle. As the cortex becomes thicker, migration of
NPCs may adopt different modes that are regulated by more complicated mechanisms.
However, NPC migrating from the neuroepithelia does not seem to involve the full-length L1.1. L1.1 is found predominantly in neuronal clusters outside the vicinity of
the ventricles. This spatial expression pattern is incompatible with a role in facilitating migration in the neuroepithelium. Its proteolytic fragments which, though contain all potent features of L1.1s and become part of the ECM, may be located at a distance too far
from the ventricular zone to exert an influence on the migration of NPC.
Loss of L1 expression may also disrupt the normal pattern of coordinated cell
proliferation and migration in the periventricular zone. A potential pathogenic
mechanism that underlies the development of L1-associated hydrocephalus is presented
in Figure 5.1B. In the L1-deficient brain, NPCs fail to polarize and become disoriented.
Following mitosis, radial migration of NPCs by somal translocation is impeded because
their rear processes are unable to adhere to the matrix at the ventricular surface. Impaired
migration of these cells results in aberrant accumulation of post-mitotic cells in the
periventricular zone and hinders subsequent waves of cell proliferation on the ventricle
surface. Consequently, coordinated cell migration and proliferation in the ventricular
neuroepithelium are disturbed in the absence of L1, resulting in a disorganized
ventricular system with an obstructed passage for CSF. As CSF circulation begins, the
increased volume of CSF that is accumulated in the hindbrain elevates the pressure
202 abnormally on the walls of the hindbrain ventricle, leading to the enlargement of the 4th ventricle (Figure 5.1.B).
Cell polarity, migration, and proliferation are inter-related processes that are under stringent regulation during brain development. A defect in molecules involved in anyone of these processes will impede the other processes. For example, genes LIS1
(encoding a component of platelet-activating factor acetyl-hydrolase) and DCX (encoding doublecortin) are responsible for lissencephaly, a severe brain developmental disease characterized by the mislocalization of cortical neurons (Vallee and Tsai, 2006).
Mutations in these two molecules result in the impairment of microtubule-dependent cell motility, as evidenced by live imaging of LIS1-deficient neural progenitor cells (Tsai et
al., 2005). Due to the impediment of interkinetic nuclear migration, the position of mitotic cells in the LIS1-defected mouse progressively becomes ectopic and away from the ventricular lumen where normal mitosis occurs. Concomitantly, the percentage of
proliferating cells decreases significantly (Gambello et al., 2003).
L1s-Neuropilin 1 interaction in the brain ventricle development
Further studies will be required to assess the impacts of L1s-Nrp1 interaction on
the migration of NPCs during ventricle formation. An important question is whether the
localization of L1.1s and Nrp1a proteins in the ventricle is affected by the absence of
their interacting counterpart. The expression of L1.1 and Nrp1a can be knocked down
individually by injecting morpholino oligonucleotides into embryos. The respective
morphants will be immunostained to assess changes, if any, in the distribution of the
other protein in the periventricular zone. Since knockdown of either protein results in
203 disturbed ventricular polarity and underdeveloped midbrain and MHB (see Chapter III), it
is likely that the absence of either L1.1s or Nrp1a influences the expression pattern of
each other reciprocally.
Experiments can also be carried out to determine in which process L1.1s-Nrp1a interaction plays a major role, whether cell proliferation, migration, or polarity formation.
First, the direct effect of this interaction on ventricular polarity should be further assessed using the inhibitory peptide YAANEL to treat embryos at 20 hpf for different time periods: 0 min, 5 min, 15 min, 30 min and 2 hours, followed by immunostaining with anti-γ tubulin antibody to visualize polarized centrosomes. If this treatment disrupts the polarity of the centrosomes in the ventricle, a pattern of disturbed supernumerary
centrosomes should become evident within a 30-min time frame (Takemura et al., 1995).
To examine whether ventricular migration is directly associated with L1.1s-Nrp1a
interaction, real-time imaging of the migration of NPCs in vivo can be performed after the addition of the inhibitory peptide. Nuclei tracking of NPCs in live embryos can be facilitated by the use of Histone H2A.F/Z:GFP transgenic fish (Pauls et al., 2001), which has been exploited successfully to demonstrate radial migration of NPC during early morphogenesis of midbrain and MHB (Langenberg and Brand, 2005). Live transgenic embryos can be mounted in 1.5% low melting point agarose in an imaging chamber
(Concha and Adams, 1998) and analyzed by a two-photon confocal system. The use of the vital dye BODIPY-ceramide should facilitate the display of cell morphology (Cooper et al., 1999). As shown in Figure 5.2, the effects of the YAANEL peptide (2 mM in 4%
DMSO) on NPC migration will be monitored for up to 5 hours. Embryos will be fixed afterwards and processed for immunohistochemistry using anti-Nrp1a antibody
204 ACB a DMSO/YAANEV YAANEL a a
~22 hpf (26 somites) b b b 2 h later 2 h eeF c c M
H later 4 h
Processes of Nucleus of NPC Nrp1a at focal Diffuse expression NPC adhesion of Nrp1a
Figure 5.2. Schematics depicting NPC migration in vivo mediated by L1.1s-Nrp1a interactions. (A) The Histone H2A.F/Z:GFP transgenic embryo of 22 hpf will be mounted in 1.5% low melting point agarose and imaged for 5 hours. Optical sections of the brain (a) will be obtained on a confocal microscope. Dorsal view of the brain is shown in (b). NPC migration will be monitored at the midbrain (M) level. E, eye; F, forebrain; H, hindbrain. (B) In the brain treated with either DMSO or the control peptide YAANEV, L1.1s-Nrp1a interactions remain intact. At the beginning, the rear process of the NPC is anchored to the ventricular surface (as revealed by BODIPY- ceramide stain). The process by which NPCs undergo somal translocation and migrate out of the ventricular zone will be monitored at 2 hours (b) and 4 hours (c). Upon arrival to their destinations, NPCs are expected to retract their trailing processes. (C) In embryos treated with the inhibitory peptide YAANEL, specific interactions between L1.1s and Nrp1a are abrogated. Nrp1a is dissociated from the adhesion complex and diffuses on the NPC surface. As NPCs fail to adhere to the ventricle surface (b), their migration will be impeded. By 4 hours, processes of NPCs will be withdrawn (c) and the nuclei will stall at their original positions.
205 (Bovenkamp et al., 2004) to reveal the distribution of Nrp1a on NPCs. Extrapolating the results from Chapter III, we can anticipate that the rear process of the migratory NPC will be retracted from the ventricular surface as a result of the abrogation of L1.1s-Nrp1a binding. The nucleus may fail to undergo somal translocation due to disrupted cell polarity and cytoskeleton organization. Moreover, immunostaining with anti-phospho-
Histone H3.3 antibody should reveal the position of mitotic cells in the ventricle. Since mitotic cells in the L1.1 morphants are mostly not confined to the ventricular surface, it is likely that many cells undergoing mitosis will be positioned outside the vicinity of the ventricle.
Neuropilin1-mediated signaling
How does L1-Nrp1 interaction regulate cellular events? Using the yeast two- hybrid system, Cai and Reed (1999) have shown that Nrp1 interacts with a PDZ
(PSD95/Discs Large/ZO-1)-containing protein, named GIPC. This interaction occurs at the C-terminal motif Ser-Glu-Ala-COOH of Nrp1a, which is conserved in all vertebrate species including zebrafish. PDZ refers to the structural domain containing a Gly-Leu-
Gly-Phe sequence motif, which is commonly found in signaling molecules involved in the localization and clustering of membrane receptors and ion-channel subunits (Kim and
Sheng, 2004). The interaction with a PDZ-containing protein suggests that Nrp1 could induce signaling cascades and modulate cytoskeleton dynamics. Indeed, results from an independent study suggest that this Nrp1 interacting protein, GIPC, binds to a GTPase- activating protein for Gα subunits (De Vries et al., 1998). Therefore, it is possible that
Nrp1 is coupled to Gα or other components in the G-protein-coupled signal transduction
206 pathway. On the other hand, a recent report shows that L1-Nrp1 interaction elevates the
intracellular level of cGMP of the nNOS pathway (Castellani et al., 2004).
Which signaling pathway is associated with L1-Nrp1 interactions? The initial identification of the involvement of potential signaling pathways in L1-Nrp1-mediated polarization of NPC would rely on pharmacological studies. NPC polarization at the periventricle can be monitored after treatment of live embryos with various pharmacological reagents. To assess the involvement of NO/cGMP signaling, embryos of
20 hpf can be incubated with the NOS inhibitor 7 nitroindazole (7NI) or the cGMP inhibitor 1H-(1,2,4)oxadiazolo (4,3-a)quinoxalin-1-one (ODQ) for 5 hours. If the NOS signaling pathway is involved, immunostaining with anti-γ tubulin should reveal an abnormal pattern of supernumerary centrosomes.
Similarly, the involvement of G-protein-mediated signaling can be tested by incubating embryos with pertussis toxin (PTx), a widely used reagent in the study of heterotrimeric G-proteins in signaling pathways. If the ventricular polarity is PTx- sensitive, downstream components of the G-protein signaling cascade can be examined using other pharmacological agents. Upon activation, the free Gα subunit (bound to GTP) dissociates from the βγ subunits. Free G subunits are functionally active and directly regulate effector proteins, such as phospholipase C, phospholipase A2 (Bloch et al.,
1989; Park et al., 1993). To test which downstream components are responsible for transmitting the signal of L1.1s-Nrp1a, embryos can be incubated with the aminosteroid
U-73122 (an inhibitor of G protein-mediated phospholipase C activation) or O4-
Bromophenacyl bromide (BPB, a generally used inhibitor for phospholipase A2) and then subjected to the examination of polarity in the ventricle.
207 Role of L1 at the neuromuscular junction
It has been well documented that L1 plays important roles in promoting neurite outgrowth and axon guidance. Its potent activity has been ascribed to its interactions with
a variety of other recognition molecules. Clearly, L1 is enriched in synaptic vesicles,
whereby it is transported to the leading edge of migrating growth cone. At the interface
between axon and muscle, L1-containing presynaptic structures has been found to align
with the postsynaptic apparatus (Triana-Baltzer et al., 2006). This observation suggests a
hitherto unidentified role of L1 in the formation of the neuromuscular junction (NMJ).
Figure 5.3 depicts the hypothetical role of L1 at the NMJ. In this scheme, neuronal L1.1
is targeted to synaptic vesicles (which can be stained with mAb SV2) for transport to the
axonal growth cone. When the growth cone meets the muscle fiber, L1.1 interacts with
pre-patterned Unplugged to initiate synaptogenesis. Interactions of L1.1 with Unplugged
have been shown by the presence of both proteins in the co-immunoprecipitated complexes using antibodies against either molecule. Furthermore, fluosphere-to- substratum binding assays have revealed that the physical interaction between the two
proteins involves the Ig3 domain of L1 (Chapter IV). The adhesive interactions between
L1.1 and Unplugged may trigger the rearrangement of intracellular cytoskeletal
components and activation of signaling cascades within both cells.
The cytoplasmic domain of L1 may be indispensable for its synaptogenic activity.
Several conserved motifs in the L1 cytoplasmic domain provide docking sites for linker proteins to the cytoskeleton and thereby modulate cytoskeleton dynamics (Maness and
Schachner, 2007). The sequence FIGQY is crucial to the activity binding of L1 to the cytoskeleton. The phosphorylation/dephosphorylation-state of the tyrosine modulates the
208 AB Neuronal Neuronal growth cone growth cone
? AChR
Muscle fiber Rapsyn
Legend: L1.1 L1.1s Unplugged Unplugged Synaptic vesicles Agrin AChR full length full length containing L1.1 (inactive) (active)
Figure 5.3. Schematic illustration of the interaction between neuronal L1.1 and Unplugged during synaptogenesis along the CaP axonal path. (A) Before the approach of a migratory growth cone, Unplugged is clustered with an en passant morphology on the muscle fiber. When the axonal growth cone meets the muscle fiber, L1.1 interacts with Unplugged and initiates synaptogenesis. L1.1s is expressed in muscle cells and its role remains to be determined. (B) Because L1.1 is able to undergo homophilic interaction, L1.1 quickly clusters Unplugged molecules to the adhesion site at the interface. Soon afterwards, upon the activation of Unplugged by the neurotransmitter Agrin, the activated Unplugged clusters expand and the signal are propagated to recruit AChR to assemble the postsynapse apparatus (Burden, 2002).
209 binding affinity of this sequence to doublecortin and ankyrin, which respectively link L1 to the microtubule and actin filaments (Garver et al., 1997; Jenkins et al., 2001; Kizhatil et al., 2002). Furthermore, the sequence RSLE and another juxtamembrane sequence
(KGGK in human L1) are putative docking sites for the actin-binding proteins ERM which may also contribute to the role of L1 in actin cytoskeleton remodeling (Dickson et al., 2002; Cheng et al., 2005). Consistent with the demonstrated impact of the cytoplasmic tail of L1 on neurite outgrowth, only the full-length form of L1.1 is capable of rescuing the morphant phenotype of aberrant axonal growth from the primary motoneurons (See Chapter IV).
The L1.1-Unplugged trans-interactions quickly clusters Unplugged molecules in the vicinity of the adhesion sites. Unplugged is further activated by the neurotransmitter
Agrin which rapidly induces large clusters of activated Unplugged to the NMJ, and the signal propagation of Unplugged is able to recruit numerous AChR to the postsynaptic apparatus (Burden, 2002). From immuno-colocalization of L1.1 and SV2 demonstrated in
Chapter IV, it is evident that L1.1 constitutes the neuronal synaptic vesicles along the axonal trajectory, suggesting that L1.1 may contribute to the stability of NMJ. This possibility is supported by an immunostaining study of the NMJ which revealed the co- localization of phosphorylated-FIGQY with AChR in the postsynaptic membrane
(Jenkins et al., 2001).
While functions of the full-length form of L1 have been emphasized, the role of
L1.1s in NMJ is not necessarily dispensable. L1.1s contains the first four Ig domains and is capable of interacting with Unplugged, as evidenced by the fluosphere-to-substratum binding data (Chapter IV). In fact, it is notable that 47% of axons exhibited ectopic
210 branching in embryos co-injected with L1.1 morpholino and L1.1s mRNA, versus 19%
of that in L1.1 morphants. The elevated number of aberrant axons in L1.1s mRNA co- injected embryos suggests an effect of L1.1s additional to the loss of its ability to induce cytoplasmic signals. The phenotype resembles that of the twister mutant in zebrafish, which show an increase of neuromuscular activity (Lefebvre et al., 2004). These observations also raise the possibility that L1.1s participates in the assembly of the
postsynaptic apparatus, which will be explored in a later section.
Signaling in L1-mediated postsynaptic differentiation
The investigate further the role of L1.1 in NMJ formation, it would be important
to determine whether L1.1 is able to induce the kinase activity of Unplugged inside of the
muscle cell, which will then trigger the recruitment of AChR to the postsynapse site
(Burden, 2002). It is known that the functions of MuSK in synaptic differentiation is
induced by the neurotransmitter agrin (DeChiara et al., 1996; Gautam et al., 1996). The
kinase that actives MuSK remains unknown. However, in the pathway of agrin-mediated
activation of MuSK, staurosporine (a relatively non-selective protein kinase inhibitor)
only affects AChR clustering without blocking tyrosine phosphorylation of MuSK,
whereas herbimycin (a specific tyrosine kinase inhibitor) abolishes the phosphorylation
of both MuSK and AChR (Ferns et al., 1996; Fuhrer et al., 1997). These results suggest
that a staurosporine-insensitive tyrosine kinase is responsible for MuSK phosphorylation.
It has been shown by mutagenesis that downstream signaling of agrin/MuSK depends
upon the phosphorylation of Tyr553 in the juxtamembrane region (NPxY) of MuSK (Zhou
et al., 1999; Herbst and Burden, 2000). Phosphorylation of this tyrosine is required to
211 fully activate the MuSK kinase activity and to recruit downstream signaling components
for all aspects of MuSK signaling, including tyrosine phosphorylation and clustering of
AChRs, and presynaptic differentiation (Herbst and Burden, 2000; Herbst et al., 2002).
However, the mechanism that regulates the phosphorylation of Tyr553 remains to be
elucidated.
What is the role of L1 in postsynaptic specialization? An important question here
is whether L1 is an activator of MuSK function. The sequence NPxY in zebrafish
Unplugged is highly conserved and the putative phosphorylation site has been assigned to
Tyr625 (Zhang et al., 2004). To determine whether Unplugged is phosphorylated by L1.1-
mediated interaction, full-length wild-type Unplugged construct fused with a HA tag to
its C-terminus (Unplugged-HA) will be transfected into COS7 cells. Lysates derived
from transfectants that have been cultured overnight on L1.1EC, L1.1ECG273D, and
L1.1ECE314K (G273D and E314K mutations disrupt the L1.1-Unplugged interaction, as
demonstrated in Chapter IV) will be subjected to immunoprecipitation with anti-HA
3F10 mAb. Unplugged activation in response to wild-type and mutant L1.1 proteins will
be assayed by immunoblotting with mAb 4G10 (anti-pY). If L1.1-mediated interaction is involved in the activation of Unplugged, phosphorylation of Unplugged will take place
when L1.1EC is used as culture substrate, whereas Unplugged will remain non-
phosphorylated when cells are cultured on the G273D and E314K mutant L1 proteins.
In order to test whether L1.1-Unplugged interaction is able to stimulate AChR
clustering, the Unplugged-EGFP construct will be used for transfection into mouse
myoblast C2C12 cells, which have been successfully utilized in many studies to
demonstrate AChR clustering upon MuSK activation (Glass et al., 1996; Jones et al.,
212 1999; Willmann et al., 2006). Either wild-type or mutant L1.1EC will be added to the
culture of transiently transfectants expressing Unplugged-EGFP fusion protein, the induction of AChR clusterization will then be examined by staining with Texas Red- conjugated α-bungarotoxin (a snake venom neurotoxin which binds specifically to
AChR). The pattern of AChR localization will be compared with that of L1.1EC (stained
with anti-His6 antibody) and Unplugged (revealed by GFP).
Role of L1s in pre-assembling postsynapse
Neurotransmitter induces clustering of AChRs which are distributed diffusely in
the plane of the muscle membrane while Agrin has been thought to be the neuron-derived
activator that mediates this process (Sanes and Lichtman, 1999). Two studies using
genetic approaches have demonstrated that the initial steps in postsynaptic differentiation
require MuSK but not Agrin (Lin et al., 2001; Yang et al., 2001). Indeed, early synaptic
specialization is apparent in the Agrin mutant mouse or in the absence of motor axons.
Further in vivo imaging studies support this notion, revealing an en passant morphology
of prepatterned AChR clusters before neuro-muscle contacts are made (Panzer et al.,
2005). Therefore, Agrin appears to be necessary to maintain, rather than to induce AChR
clusters. Interestingly, transfection of a panel of MuSK mutations into human embryonic
kidney 293T cells leads to the observation that discrete clusters of MuSK are formed
independent of its tyrosine kinase activity (Bianchetta et al., 2005). These results suggest
that an extracellular stimulus or a binding protein must be responsible for the clustering
of MuSK prior to its activation.
213 In zebrafish, L1.1s is expressed in the muscle (Chapter IV). The L1.1s protein is
localized to the migratory adaxial cells, which express a relatively high level of
Unplugged (Zhang et al., 2004). Injection of L1.1s mRNA induces ectopic branching, a
phenotype that could have been resulted from elevated postsynaptic activity in a
hyperactive AChR mutant (Lefebvre et al., 2004). Therefore, it is possible that the expression of L1.1s by mRNA injection into the embryo over-activates Unplugged and consequently elevates postsynaptic differentiation. It is, therefore, possible that, under physiological conditions, muscular L1.1s interacts with Unplugged and activates its
signaling cascades to assemble en passant synapses.
To test this hypothesis, the pre-assembly of postsynaptic apparatus will be
assessed using Texas Red-conjugated α-bungarotoxin in embryos lacking an L1.1-
Unplugged interaction. The en passant morphology of AChR clusters is expect to be
absent in the L1.1 morphant as in the case of Unplugged mutants (Lefebvre et al., 2007).
If muscular L1.1s plays a role in clustering Unplugged, the resultant AChR assembly may also be absent from the prospective motor axonal path prior to axon arrival (which can be monitored by immunostaining with mAb SV2, an axonal marker for pre-synaptic vesicles). The preassembly of AChR clusters is expected to be restored when wild-type
L1.1s mRNA is coinjected with L1.1 morpholino. On the other hand, pre-assembled
AChR clusters should not be detected in the embryos co-injected with L1.1s mRNA carrying the mutations G273D and E314K because these mutations have been found to specifically abrogate the L1.1-Unplugged interaction (see Chapter IV),
As demonstrated in Chapter III, L1.1s is prone to dimerize on the cell surface. To
evaluate the potential impact of dimeric L1.1s on Unplugged clustering, the plasmid
214 Unplugged:HA will be transfected into COS7 cells. Either the Ig3-His6 fusion protein or the induced dimer form of Ig3 (by preceding incubation with anti-His6 antibody) will be added to the transfected cell culture. The cell surface distribution of Unplugged on non- permeabilized cells will be assessed by immunocytochemistry using anti-Unplugged antibody against an extracellular epitope. Whether Unplugged is clustered by Ig3 monomer or dimer will be assessed by the percentage of cells containing puncta and clusters of MuSK. Furthermore, activation of MuSK by monomeric and dimeric Ig3 will be examined. Lysates derived from the Ig3/(Ig3)2 treated transfectants will be immunoprecipitated with anti-HA antibody and the presence of tyrosine-phosphorylated
Unplugged will be probed using mAb 4G10. As activated MuSK will induce further clustering of MuSK, punta of active MuSK will become evident quickly, inducing differentiation of numerous postsynaptic assemblies (Jones et al., 1999; Moore et al.,
2001). These attempts to elucidate the function of the dimeric L1.1s may unlock the mystery behind the initiation of MuSK function during synaptic differentiation.
215
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Appendix
247 Partial sequence of zebrafish L1.1 gene (exon 10-11)
GCACGGAT GTGGACCCGA GACGGCGTGT GAGTTCTGGG AAACTGATCC TGTCCAATGT GGAGTTCAGC GATACGGCAG TGTATCAGTG CGAGGCTGTC AACAAACACG GAAGCATCCT GATCAACACA CACGTGCATG TTGTTGGTGA GCATTAATAG TGTAAAACAC ATATTATTTA CATGGTTCCT ATTCTGCACA GGCTCATAGG GAACATGTGC CCAGGGCTAC ATATTTGAGA ACTGCAAAAT ATATACCCGG GGCTATGTAT GGCTGCATTT TGTGTGTAAA GTGTACGACA ACCTTCTCTT CCTTTTTTGC TAAAGCTTTG ACGGATTGCC TCCGTATGGA CCGCTTTTCT GGCGTAACCA GTTTGCTTAG TTTTCCATGT ATGGGATTGG ACTTGGGATG TGGAGCGGAG TGGACTTGGG TTCAAAACCA ACAAAGCACA CTTCCAGAAA CCAAGTAAAA TGAAACCAAA AAATAAATAA GGCGATCCTT GGCTCAGTTG GCGTTTCTGT GTGGAGTTTG CATGTTCTCC CTGCCTTCAC GTGGGTTTCC TCCGAGTGCT CCGGTTTCCC CCACAGTCCA AACACATGCG GTACAGGTGA ATTGGGTTAG CTAAATTGTC CGTAGTGTAT GAGTGTGTGT GTGGATGTTT CCCAGAGGTG GGTTGCGGCT GGAAGGGCAT CCGCTGCGTA AAAGCTTGCT TTATAAGTTG GCGGTTCATT CCGCTGTGGT CACCCTAGAT TCATTTCCAA ACAAAATGCA ACGTCCCCCG ACATTGAGGT ACAGCGTCAA TCTGCCGTCA TGTTAACGTC CTGGCCCTGC AGGGAAAACT GACTTTCGTA AGATGCTGTT TTACAGACTG ACCTTTGCCT TCTCTTTGGA TTGTTTTGTT TGTCTGTGTA CGTGTATTAT GTCTTGTCTC TTTGTAAGGA GCATGTAACG TGTTGTTATG CAAGAACATT TTCAGATCTA AGTCTGACAA TAAAGTACTG TATTGCAGAA CTGCCAGCTC AGATCCTGAC CCCAGATGAG CGCCTCTATC AGGCCACTGC AGGACAGACT GTAATGTTGG ACTGCAGAAC GTTCGGGTCG CCGCTGCCCA AAATACACTG
Notes: Yellow highlighted sequences: exons 10 and 11; Red highlighted sequences: two continuous stop codons; Underlined sequence: 3’ L1.1s sequence obtained by RACE; Red sequences: polyadenylation signal (AATAAA is a signal with highly significant occurrence rate 90%, Colgan and Manley, 1997; Beaudoing et al., 2000).
Reference:
Beaudoing, E., Freier, S., Wyatt, J.R., Claverie, J.M., and Gautheret, D. (2000) Patterns of Variant Polyadenylation Signal Usage in Human Genes. Genome Res.10, 1001-1010. Colgan, D.F. and Manley, J.L. (1997) Mechanism and regulation of mRNA polyadenylation Genes Dev. 11, 2755-2766.
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