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Glycosaminoglycan biosynthesis
The assembly of GAG chains occurs in the endoplasmic reticulum and Golgi. It is initiated by the synthesis of the so-called GAG-protein linkage region tetrasaccharide, common to both HS and CS chains, and covalently linked to specific serine residues in different core proteins. This tetrasaccharide comprises a xylose (from precursor uridine diphosphate (UDP)-xylose), two galactose (Gal) units and a glucuronic acid (GlcA) (Sugahara et al., 2003). After this first step, monosaccharide units Gal, GlcA, N-acetylglucosamine (GlcNAc) and N-acetylgalactosamine (GalNAc) are transferred from sugar nucleotide precursors (UDP-Gal, UDP-GlcA, UDP-GlcNAc and UDP-GalNAc) to the linkage region leading to chain elongation. The transfer of GalNAc or GlcNAc to the linkage oligosaccharide is the first step that provides the corresponding specificity for CS or HS formation (Silbert and Sugumaran, 2002). HS and CS then differentiate into mature chains after various modification reactions, mainly epimerization and sulfation, and this chemical combination can give rise to a large number of highly diverse structures (Figure 7). Unlike nucleic acids and proteins, the biosynthesis of complex sugars is a non-template-driven process involving several enzymes and their tissue-specific isoforms. The list of these enzymes goes from glycosyltransferases on the basis of their sulfation patterns (adapted from Sugahara et al., 2003). that participate in the elongation and polymerization of GAG chains to sulfotransferases and epimerases that modify the glycan backbone.
Sulfotransferases add sulfate groups from 3’phosphoadenosine-5’phosphosulfate to the growing GAG chains. In HS, sulfation modifications occur in restricted domains of about 5-10 disaccharides that are hyper variable and interspersed within poorly sulfated regions. The modifications in HS biosynthesis are initiated with the N-deacetylation and subsequent N-sulfation of selected GlcNAc residues, where both reactions are carried out by a bifunctional enzyme called N-deacetylase-N-sulfotransferase. Four mammalian N-deacetylase-N-sulfotransferases have been identified, with different substrate specificities (Grobe et al., 2002). After N-sulfation, C5-epimerization of GlcA into iduronic acid can occur and can be followed by 2-O-sulfation (1 enzyme), 6-O-sulfation (three enzymes) and/or 3-O-sulfation (7 enzymes). 2-O-sulfated hexuronic acids are almost exclusively found in contiguous N-sulfated domains of the GAG chain. Similarly, CS chains are composed of several types of disaccharide units that differ in their sulfation pattern. To date, seven sulfotranferases responsible for the sulfation of CS/DS chains have been identified (Kusche-Gullberg and Kjellén, 2003). A non-sulfated unit serves as a common acceptor substrate for two types of sulfotransferases, one catalyzing the 4-O-sulfation and the other the 6-O-sulfation of GalNAc residues. Subsequent sulfation on the C-2 position of the GlcA or on the C-6 position of the GalNAc can occur, forming disulfated disaccharides. The typical disaccharide units found in CS chains are named A, C, D and E, according to their sulfation pattern (Figure 8).
The redundancy of the biosynthetic machinery based on different enzyme isoforms (Esko and Lindahl, 2001) makes it challenging to take a genetic approach by knocking in or out specific enzymes. However, most critical functions of GAGs in development have been revealed by analyses of mutants defective in GAG biosynthesis enzymes. For instance, exostosin glycosyltransferase 1 homozygous mutant mice, in which only short stubs containing the linkage region are attached to the protein, fail to gastrulate and generally lack organized mesoderm and extraembryonic tissues (Lin et al., 2000). Mutants lacking both N-deacetylase-N-sulfotransferase 1 and N-deacetylase-N-sulfotransferase 2 die during early embryogenesis (Forsberg and Kjellen, 2001). In C. elegans, animals lacking one of the three HS-modifying enzymes, glucuronyl C5-epimerase, 6O-sulfotransferase or 2O-sulfotransferase, exhibit distinct as well as overlapping axonal and cellular guidance defects in specific neuron classes (Bülow and Hobert, 2004). C. elegans mutants for the nematode orthologs of chondroitin sulfate synthase and chondroitin polymerizing factor show cytokinetic regression in early embryogenesis (Hwang et al., 2003; Izumikawa et al., 2004; Mizuguchi et al., 2003). In mouse, knockout of ß-1,3-glucuronyltransferase 1 encoding gene also results in embryonic lethality due to cytokinetic failure (Izumikawa et al., 2010). These studies show the crucial roles of GAG chains in metazoan embryonic development, from worms to mammals.
Glycosaminoglycan interaction with morphogens and growth factors
Disruption of genes involved in the synthesis of HSPG and CSPG core proteins also results in physiological defects (Lamanna et al., 2007). But although the core proteins alone can display some activity, mounting evidence indicates that most functions of CSPGs and HSPGs are largely exerted through their GAG moiety. GAG moieties vary considerably in the size and number of CS or HS per core protein and in the position and degree of modifications, primarily sulfation, which create huge molecular diversity and structural complexity. For instance, an octasaccharide can have over 1,000,000 different sulfation patterns (Sasisekharan and Venkataraman, 2000). This structural heterogeneity of GAG chains is thought to promote specific interactions with a variety of bioactive molecules such as growth factors and morphogens in the extracellular environment. As mentioned earlier, genetic studies on GAG biosynthetic enzymes have provided direct functional evidence for the involvement of GAGs in cell growth and developmental processes based in particular on Wnt, Hedgehog, bone morphogenetic protein (BMP), transforming growth factor and fibroblast growth factor (FGF) signaling pathways (Lin, 2004). For instance, in Drosophila embryos with mutations in sulfateless, the N-deacetylase-N-sulfotransferase homolog, the signaling pathways involving either Wingless (a member of the Wnt family), FGF or Hedgehog are completely silenced (Lin and Perrimon, 1999). Reduction of 3O-sulfotransferase activity in Drosophila results in compromised Notch signaling with neurogenic phenotypes (Kamimura et al., 2004). In mice lacking exostosin glycosyltransferase 1, Indian Hedgehog is not correctly distributed in the gastrulating embryo (Lin et al., 2000). GAGs can participate in growth factor signaling by assembling protein-protein complexes on the cell surface. The best example is the well-studied affinity of HS for FGFs and FGF receptors. FGF signals through cell surface tyrosine kinase receptors and its oligomerization leads to receptor oligomerization, phosphorylation of other signaling molecules and initiation of signaling cascades (Raman et al., 2005). HSPGs directly interact with FGFs and their receptors to form a ternary complex at the cell surface thus facilitating FGF interaction with its receptor and/or stabilizing FGF oligomerization (Lin, 2004). Similarly, HS chains bind to both the axon guidance molecule Slit2 and its receptor Robo1 to facilitate the ligand-receptor interaction (Zhang et al., 2013).
X-ray crystal structures of GAG-protein complexes have provided information on the structural features required for GAG-protein interaction. The specificity of GAG-protein interactions is governed by the ionic interactions of the sulfate and carboxylate groups in the GAG chains with the basic amino acids on the protein, as well as by the optimal fit of the GAG chain into the binding site of the protein. The topology and distribution of basic amino acids on the protein sequence influences the specificity of its recognition of GAG sequences. Two principal consensus sequences have been presented as potential binding sites for GAG interaction: XBBXBX and XBBBXXBX, where B is a basic residue (lysine or arginine, sometimes histidine) and X is a hydrophilic residue (Cardin and Weintraub, 1989). Despite their identical charges, arginine residues bind more tightly to GAGs than lysine residues. The spacing of these residues may determine protein-GAG interaction affinity and specificity: GAG binding sites can be formed by basic amino acids distant in sequence but brought close together in the folded protein (Gandhi and Mancera, 2008). The binding affinity of GAGs depends on the ability of their oligosaccharide sequence to provide optimal charge interaction with the protein. Under physiological conditions, all carboxylic acid and sulfate groups are deprotonated, giving a very high negative charge to GAG chains (Capila and Linhardt, 2002). These negatively charged groups form ion pairs with the clusters of positively charged basic amino acids on proteins. However, ionic interactions contribute only in part to the GAG-protein interaction (30% for FGF-2 and heparin (Gandhi and Mancera, 2008)) as other types of interaction are also involved: van der Waals forces, hydrogen bonds and hydrophobic interactions.
Conformation studies have shown that variations in the primary sequence (chain length and sulfation pattern) are crucial for protein binding. The minimum GAG chain length required for binding is typically 4-12 monosaccharide units (Gandhi and Mancera, 2008). Binding affinity increases with the size of the oligosaccharides (Rusnati et al., 1999). Tetra- and hexa-saccharides of HS are sufficient to bind FGF1 and FGF2 with high affinity. However, octosaccharides or longer sugar chains are required for bridging a dimeric FGF2 along with its receptor to form the ternary signaling complex (Pellegrini et al., 2000; Schlessinger et al., 2000). The relative proportions of N- and O-sulfation in HS chains impact on their interaction with proteins. FGF1 and FGF2 interactions with HS oligosaccharides require 2-O- and -N- Hyaluronan is attached to the cell surface and provides a scaffold for the perineuronal net. The lectican family of chondroitin sulfate proteoglycans (aggrecan, neurocan, phosphacan, versican, brevican, …) bind hyaluronan through their N- terminal domain (interaction stabilized by link proteins) and tenascin-R via their C-terminal domain (adapted from Galtrey and Fawcett, 2007).
sulfate groups. However, the 6-O sulfation is critical for FGF1 signaling, but not completely necessary for FGF2 signaling (Guerrini et al., 2002). CS chains with disulfated disaccharides (CS-D and/or CS-E) bind with high affinity to midkine, pleiotrophin, FGF, hepatocyte growth factor and brain-derived growth factor (BDNF) (Tamura et al., 2012). CS-A chains negatively regulate axon guidance (Wang et al., 2008), while CS-C chains appear to be relatively permissive for neurite regeneration (Lin et al., 2011). CS-E, but not CS-A, CS-C nor HS, binds to contactin-1 and leads to neurite outgrowth (Mikami et al., 2009). The oligosaccharide motif that is recognized by a protein is often found in low abundance which, given the diversity of GAG sequences, supports the idea of a specificity in GAG-protein interactions (Raman et al., 2005).
Glycosaminoglycans in the postnatal brain
Complex sugars are precisely distributed in the developing and adult brain and the spatiotemporal expression and composition of GAGs changes throughout development. GAG-protein interactions in the central nervous system have been shown to play critical roles in proliferation and differentiation of neural progenitor cells, neuronal migration, axon guidance, synaptogenesis, neural plasticity and regeneration (Maeda et al., 2011). Sulfation patterns of GAG chains change dramatically during brain development. In the adult brain, the content of CS/DS is about 9 times higher than the HS one. The proportion of 6-O-sulfated CS decreases and that of 4-O-sulfated CS progressively increases (Kitagawa et al., 1997; Mitsunaga et al., 2006). Furthermore, while during embryonic development, GAGs are diffusely distributed in the extracellular matrix, they gradually accumulate around subsets of neurons during postnatal development.
In the postnatal and adult brain, CSPGs are the major components of a form of condensed extracellular matrix called perineuronal nets (PNNs) (Galtrey and Fawcett, 2007). PNNs surround cell soma and proximal dendrites of distinct neuron subpopulations in the brain (Celio and Blümcke, 1994). PNNs constitute a highly organized complex composed of HA, link proteins, CSPGs (mainly lecticans) and tenascin-R (Figure 9) (Carulli et al., 2007; Köppe et al., 1997b). Hyaluronan synthases, located on the plasma membrane, anchor the PNN structure to the cell surface (Carulli et al., 2007). They synthesize and secrete hyaluronan in the extracellular space. There are three hyaluronan synthase isoforms that synthesize hyaluronan chains of various lengths and at different speeds (Itano et al., 1999; Spicer et al., 1997). Hyaluronan synthase isoforms have different spatiotemporal expressions and confer different structure and mechanical strength to the PNNs (Carulli et al., 2006; Galtrey et al., 2008). The lecticans (aggrecan, neurocan, versican and brevican), bind to hyaluronan through the N-terminal domain of their core protein (Iozzo, 1998). Aggrecan, which is necessary for PNN formation (Giamanco et al., 2010; Kwok et al., 2010), is present on almost all PNN-positive neurons, while other lecticans are found only in subpopulations of PNN-bearing neurons (Galtrey et al., 2008). Furthermore, glycosylation of aggrecan in PNNs varies between different areas of the nervous system (Matthews et al., 2002). The differential distribution of lecticans in PNNs probably affects their properties and functions (Kwok et al., 2011). Some CSPGs of the PNNs are produced by neurons, some by glia and some by both cell types (Carulli et al., 2006; Giamanco and Matthews, 2012). However, cortical neurons in culture are able to construct PNN-like structures without glial cells (Miyata et al., 2005). The interaction between hyaluronan and lecticans is stabilized by link proteins (Binette et al., 1994), which belongs to a family of proteins that bind to both hyaluronan and CSPGs (Kwok et al., 2011). Crtl1 and Bral2 are the two members of this family that colocalize with PNNs; they are expressed exclusively by PNN-bearing neurons (Bekku et al., 2003; Rauch, 2004). Link proteins are crucial for the condensed nature of the PNNs (Kwok et al., 2010). Indeed, mice lacking Crtl1 in the central nervous system show reduced PNNs throughout the nervous system, with no dendritic staining and a faint staining around neuronal soma (Carulli et al., 2010). Bral2-deficient mice also have attenuated PNNs (Bekku et al., 2012). To complete the formation of the PNNs, the lecticans bind tenascin-R through their C-terminus. Tenascin-R knock-out mice exhibit abnormal PNN staining (Weber et al., 1999). The sulfation pattern of PNNs differs from the sulfation pattern of GAGs of the diffuse matrix (Deepa et al., 2006). The proportion of CS/DS (71%) is higher than that of HS (29%). In PNNs, unsulfated CS and 4-O-sulfated CS disaccharides are predominant, while the disulfated CS-D and CS-E only represent 1.2% and 2.1% of the CS GAGs, respectively. Unsulfated HS disaccharide is the major disaccharide in the PNNs, followed by 2-N-sulfated HS disaccharide units. Di- and tri-sulfated HS disaccharides represent about 15% of the PNN HS GAGs (Deepa et al., 2006).
Cellular and molecular mechanisms of cortical plasticity
Binocular interactions are detected by the integrated action of local excitatory and inhibitory connections in the visual cerebral cortex. This excitatory/inhibitory balance is dynamically adjusted by the cortical layer circuits where inhibitory connections develop later than the excitatory ones (Turrigiano and Nelson, 2004). An optimal excitatory/inhibitory balance is required for plasticity and critical period onset is determined by the maturation of local inhibitory circuits (Fagiolini and Hensch, 2000; Hensch, 2005). Consequently, directly manipulating inhibitory transmission shifts the timing of the critical period for ocular dominance plasticity (Figure 11). Developing inhibitory cells are susceptible to early life sensory experience. Partial or total loss of activity in a sensory cortex – dark-rearing from birth (Morales et al., 2002) but also whisker trimming during the critical period (Jiao et al., 2006) and early hearing loss (Kotak et al., 2008; Takesian et al., 2010) – generally leads to a In the upper panel, inputs from the eyes converge in the binocular zone of the primary visual cortex. The right scheme illustrates the cortical microcircuits between excitatory pyramidal cells (green), parvalbumin inhibitory cells (blue) and non-parvalbumin inhibitory cells (grey). GABAA receptors containing the α1-subunit are enriched at somatic synapses from large basket PV-cells onto pyramidal cells. Many factors controlling plasticity are found within the extracellular matrix surrounding PV-cells.
The lower panel presents the mechanisms controlling the onset and closure of the critical period. Early factors (PSA-NCAM) prevent precocious plasticity. Critical period onset is triggered when factors (BDNF, NARP, GAD65) promote PV-cell maturation. This triggers a sequence of molecular events (CaMKII, ERK, protein synthesis), which ultimately induce structural changes. The critical period closes when molecular brakes (PNN, PirB, NgR and epigenetic changes) gradually emerge to dampen plasticity (adapted from Sugiyama et al., 2009 and Takesian and Hensch, 2013). down-regulation of inhibitory transmission. The primary inhibitory neurotransmitter in the brain, γ-aminobutyric acid (GABA), is synthesized by glutamic acid decarboxylase produced by two distinct genes, Gad65 and Gad67. Mice lacking the synaptic isoform of the GABA-synthetic enzyme, Gad65, as well as immature wild-type animals just after eye opening, exhibit weak GABA release and no loss of visual responsiveness to an eye deprived of vision (Hensch et al., 1998). Weak GABAergic signaling inhibition in the visual cortex before P20 prevents experience-dependent plasticity. These mice show no response to monocular deprivation until inhibitory transmission is restored by enhancing the postsynaptic sensitivity to GABA with benzodiazepines (Fagiolini and Hensch, 2000; Iwai et al., 2003). These agonists of GABA, such as diazepam, effectively compensate for poor presynaptic GABA release.
GABA inhibitory interneurons account for nearly 20% of cortical neurons, but not all GABA circuits are involved in critical period regulation. Critical period onset corresponds closely to the emergence of parvalbumin inhibitory interneurons (PV-cells) (del Río et al., 1994). Parvalbumin is one of the three calcium-binding proteins which, together with calretinin and calbindin, are expressed in most GABA-producing neurons in the cortex in largely non-overlapping groups. PV-cells are fast-spiking interneurons, which means that they can fire non-adapting action potentials at rates of up to several hundred of Hertz, due in part to unique potassium conductances (Kv3 class) (Hensch, 2005). The specific blockade of the potassium channel Kv3.1 leads to impaired ocular dominance plasticity, thus mimicking the Gad65 knock-out phenotype but in a cell type-specific manner (Hensch, 2005). There are two types of PV interneurons: the chandelier cells which form synapses onto the initial segment of excitatory pyramidal cell axons and the large basket cells that mostly contact excitatory pyramidal cell soma. PV-cell induced-inhibition triggers cortical plasticity through GABAA receptors containing α1 subunit (Fagiolini et al., 2004). These receptors are located on excitatory pyramidal cell soma and preferentially targeted by large basket PV-cells (Figure 12) (Ali and Thomson, 2008). PV-cell maturation thus leads to an optimal ratio of excitatory and inhibitory circuit activity. Precocious plasticity is prevented during the pre-critical period by factors such as alpha 2,8-polysialic acid (PSA) bound to the neural cell adhesion molecule (PSA-NCAM) (Figure 12). PSA expression in the mouse primary visual cortex declines shortly after eye opening and premature PSA removal promotes the maturation of GABAergic innervation and triggers a precocious critical period for ocular dominance (Di Cristo et al., 2007).
Critical period onset is triggered when factors such as BDNF and neuronal activity-regulated pentraxin (NARP) promote PV-cell maturation (Figure 12). Genetic enhancement of BDNF triggers an early critical period in the visual cortex by promoting PV inhibitory circuit maturation (Hanover et al., 1999; Huang et al., 1999). Instead, blocking of BDNF signaling prevents the development of ocular dominance in the kitten primary visual cortex (Cabelli et al., 1997). BDNF production is activity-dependent (Greenberg et al., 2009) and BDNF is kept at immature levels in the visual cortex by dark-rearing (Castren et al., 1992). NARP is also an activity-regulated protein that regulates the expression of a class of glutamate receptor in PV-cells (Chang et al., 2010) and NARP knock-out mice fail to install ocular dominance plasticity throughout life (Gu et al., 2013). PV-cell maturation is accompanied by a sequence of structural and molecular events that lead to circuit rewiring and physiological consolidation (Hensch, 2005). Tissue-type plasminogen activator is a major serine protease, which, upon release, cleaves the physical connections between pre- and postsynaptic partners and induces dendritic spine motility (Mataga et al., 2004; 2002; Oray et al., 2004). The deprivation-induced increase in tissue-type plasminogen activator, along with spine pruning, fails to occur in mice lacking Gad65 (Mataga et al., 2002; 2004). Multiple protein kinases (CaMKII, PKA, ERK (Antonini et al., 1999; Di Cristo et al., 2001; Taha and Stryker, 2002; Trachtenberg and Stryker, 2001)) and homeostatic regulators (TNF (Kaneko et al., 2008)) eventually converge on gene transcription programs mediated by CREB to ultimately strengthen open eye connections.
The critical period closes as molecular brakes that dampen plasticity gradually emerge (Figure 12). The brake factors include structural brakes, which physically prevent neurite pruning and outgrowth (PNNs, myelin factors) and functional brakes acting on neuromodulatory systems. PNNs gradually enwrap PV-cell cell bodies and proximal dendrites (Härtig et al., 1999). In the visual cortex, the progressive increase in PNNs across postnatal development coincides with PV-cell maturation and is thought to contribute to the closure of the critical period (Pizzorusso et al., 2002; Sur et al., 1988). Mice lacking the Crtl1 link protein show extended plasticity into adulthood (Carulli et al., 2010). Myelin and myelin-associated inhibitors have also been shown to control critical period: proteins found in myelin (Nogo, myelin-associated glycoprotein, myelin oligodendrocyte glycoprotein) limit axonal sprouting by their binding to Nogo receptor and paired immunoglobulin-like receptor B complex (PirB) (Atwal et al., 2008). Nogo receptor knock-out mice exhibit ocular dominance plasticity beyond the critical period. In mice lacking functional PirB, ocular dominance plasticity is more robust at all ages (Syken et al., 2006). Developmental changes in DNA methylation and histone modifications may also be involved in critical period transitions. DNA methylation represses transcription by interfering with transcription factors binding or by recruiting repressor complexes containing histone deacetylases to condense chromatin structures (Fagiolini et al., 2009). Closure of the critical period for ocular dominance is associated with a downregulation of visual experience-induced histone acetylation and phosphorylation (Putignano et al., 2007). Windows of plasticity therefore arise between the maturation of an optimal excitatory/inhibitory balance and a later set of emerging brake factors that persistently offset the synaptic pruning machinery.
Reopening plasticity in the adult
Targeting excitatory/inhibitory balance or the molecular brakes using pharmacological or genetic approaches (as the Gad65 knock-out mouse) can accelerate or delay plasticity onset and even open windows of plasticity in the adult cortex. Indeed, the emerging view is that the brain is intrinsically plastic; adult brain plasticity is dampened (reversibly) by molecular brakes that limit excessive rewiring after critical period closure (Takesian and Hensch, 2013).
Plasticity can be reopened after the critical period by reinstalling immature lower levels of inhibition. Intracortical infusion of a Gad inhibitor or of a GABAA receptor antagonist promotes plasticity in adult primary visual cortex (Berardi et al., 2003). Exposure of adult rats to complete darkness enhances visual cortex plasticity, due to reduced expression of GABAA receptors relative to AMPA receptors that alters the excitatory/inhibitory balance in the visual cortex (He et al., 2007). Transplantation of embryonic inhibitory neurons in the postnatal visual cortex also promotes ocular dominance plasticity after critical period closure (Southwell et al., 2010). Raising adult amblyopic rats (rats monocularly deprived during the critical period) in an enriched environment also restores normal visual acuity and ocular dominance (Sale et al., 2007). A similar effect is obtained when animals are treated with fluoxetine, a selective serotonin recapture inhibitor (Maya Vetencourt et al., 2008). Both treaments reduce GABAergic inhibition in the visual cortex, accompanied by an increased expression of BDNF.
Table of contents :
LIST OF ILLUSTRATIONS
INTRODUCTION
I. Homeoproteins
A. Homeoproteins as classical transcription factors
B. Homeoproteins Otx1 and Otx2 and their roles in brain and eye development
C. Homeoproteins unconventional intercellular transfer
D. Homeoproteins as signaling proteins with in vivo roles
II. Glycosaminoglycan
A. Glycosaminoglycan structures
B. Glycosaminoglycan biosynthesis
C. Glycosaminoglycan interaction with morphogens and growth factors
D. Glycosaminoglycans in the postnatal brain
III. Critical period plasticity
A. Critical periods
B. Cellular and molecular mechanisms of cortical plasticity
C. Reopening plasticity in the adult
IV. Otx2 homeoprotein and visual cortex plasticity
A. Otx2 transfer in the visual cortex during the critical period
B. Otx2 transfer in the adult consolidated visual cortex
MATERIALS AND METHODS
I. Plasmid and proteins
II. Animals
III. Cell culture
IV. Animal experiments
V. Cerebrospinal fluid sampling
VI. Cell surface biotinylation
VII. Western blot
VIII. RNA extraction and deep sequencing
IX. Immunohistochemistry
X. Glycosaminoglycan experiments
XI. Statistical analysis
RESULTS AND DISCUSSION
I. Otx2 transgenic mice to investigate Otx2 transfer roles in the visual system
A. Otx2-AA protein activity
B. Otx2 activity in eye development and maintenance
C. Otx2 GAG-binding motif has a role in critical period timing
D. Towards the identification of targets of Otx2 in the visual cortex
Discussion
II. Identifying Otx2-binding GAGs
A. Otx2-binding GAGs in the cortex
B. GAG-binding homeoproteins: Engrailed2 versus Otx2
Discussion
III. Tools for disrupting Otx2 transfer in the visual cortex
A. A synthetic CS-E hexasaccharide interferes with Ox2 cortical transfer
B. Blocking Otx2 transfer in vivo in a single chain antibody mouse
Discussion
CONCLUSION AND PERSPECTIVES
I. Otx2, a master regulator of cortical plasticity?
II. Otx2-GAG interaction, from the source to the cortex?
REFERENCES
