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Regulation of filopodia dynamics is critical for proper synapse formation Gauthier-Campbell, Catherine 2008

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Regulation of filopodia dynamics is critical for proper synapse formation  by CATHERINE GAUTHIER-CAMPBELL B.Sc. Concordia University, 2001  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in THE FACULTY OF GRADUATE STUDIES (Neuroscience)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2008 © Catherine Gauthier-Campbell, 2008  ABSTRACT Despite the importance of proper synaptogenesis in the CNS, the molecular mechanisms that regulate the formation and development of synapses remain poorly understood. Indeed, the mechanisms through which initial synaptic contacts are established and modified during synaptogenesis have not been fully determined and a precise understanding of these mechanisms may shed light on synaptic development, plasticity and many CNS developmental diseases. The development and formation of spiny synapses has been thought to occur via filopodia shortening followed by the recruitment of proper postsynaptic proteins, however the precise function of filopodia remains controversial. Thus the goal of this study was to investigate the dynamics of dendritic filopodia and determine their role in the development of synaptic contacts.  We initially define and characterize short lipidated motifs that are sufficient to induce process outgrowth. Indeed, the palmitoylated protein motifs of GAP-43 and paralemmin are sufficient to induce filopodial extensions in heterologous cells and to increase the number of filopodia and dendritic branches in neurons. We showed that the morphological changes induced by these FIMs (filopodia inducing motifs) require ongoing protein palmitoylation and are modulated by a specific GTPase, Cdc42, that regulates actin dynamics. We also show that their function is palmitoylation dependent and is dynamically regulated by reversible protein palmitoylation. Significantly, our work suggests a general role for those palmitoylated motifs in the development of structures important for synapse formation and maturation.  ii  We combined several approaches to monitor the formation and development of filopodia. We show that filopodia continuously explore the environment and probe for appropriate contacts with presynaptic partners. We find that shortly after establishing a contact with axons, filopodia induce the recruitment of presynaptic elements. Remarkably, we find that expression of acylated motifs or the constitutively active form of cdc-42 enhances filopodia number and motility, but reduces the recruitment of synaptophysin positive presynaptic elements and the probability of forming stable axodendritic contacts. We provide evidence for the rapid transformation of filopodia to spines within hours of imaging live neurons and reveal potential molecules that accelerate this process.  iii  TABLE OF CONTENTS ABSTRACT .............................................................................................................................................. ii LIST OF TABLES.................................................................................................................................... vi LIST OF FIGURES ................................................................................................................................. vii LIST OF ABBREVIATIONS................................................................................................................. viii ACKNOWLEDGEMENTS...................................................................................................................... xi DEDICATION......................................................................................................................................... xii CO-AUTHORSHIP STATEMENT........................................................................................................ xiii 1.  CHAPTER I: Overview and Summary............................................................................................ 1 1.1. Wiring of the Brain...................................................................................................................... 1 1.2. Dendritic Filopodia...................................................................................................................... 2 1.2.1. Structure and Function of Dendritic Filopodia ...................................................................... 2 1.2.2. Filopodia Development: the Actin Cytoskeleton and Actin-Binding Proteins........................ 4 1.2.3. The Rho family of GTPases .................................................................................................... 5 1.2.4. Signaling Pathways Connecting Cdc42 to the Actin Cytoskeleton ......................................... 9 1.2.5. A Role for Acylated Proteins in Filopodia Formation.......................................................... 11 1.2.6. Palmitoylated Proteins and their Role in Filopodia Formation........................................... 14 1.3. A Role for Filopodia in Synaptogenesis.................................................................................... 15 1.3.1. Axo-Dendritic Contact Formation........................................................................................ 15 1.3.2. Synaptogenesis at Filopodial Contact Sites.......................................................................... 15 1.4. Molecules that Regulate Synaptogenesis .................................................................................. 17 1.4.1. Trans-Synaptic Signaling: Neuroligins and Neurexins ........................................................ 18 1.4.2. Scaffolding molecules ........................................................................................................... 19 1.4.3. The Role of Shank1B in Synaptogenesis ............................................................................... 20 1.5. A Role for Filopodia in Dendritic Spine Formation .................................................................. 21 1.5.1. Structure and Function of Dendritic Spines ......................................................................... 21 1.5.2. Spine abnormalities and brain disorders.............................................................................. 23 1.5.3. Spinogenesis: Different Views on the Origin of Dendritic Spines ........................................ 24 1.5.4. The Filopodia Hypothesis..................................................................................................... 25 1.5.5. Reconciling the Different Models ......................................................................................... 27 1.6. Research Hypothesis and Objectives......................................................................................... 28 1.7. References ................................................................................................................................. 31  2. CHAPTER II: Regulation of dendritic branching and filopodia formation by specific acylated protein motifs.............................................................................................................................................. 46 2.1. Introduction ............................................................................................................................... 46 2.2. Materials and Methods .............................................................................................................. 48 2.2.1. cDNA Cloning and Mutagenesis........................................................................................... 48 2.2.2. COS Cell Culture and Transfection...................................................................................... 49 2.2.3. COS Cell Labeling and Immunoprecipitation ...................................................................... 50 2.2.4. Primary Neuronal Culture and Transfection........................................................................ 50 2.2.5. Immunofluorescence ............................................................................................................. 51 2.2.6. Quantitative Measurement of Filopodia Induction in COS-7 Cells...................................... 52 2.2.7. Quantitative Measurement of Dendritic Branching and Filopodia ...................................... 52 2.2.8. Statistical Analysis................................................................................................................ 53 2.3. Results ....................................................................................................................................... 53 2.3.1. Differential Filopodia Induction by the Palmitoylation Motifs in Heterologous Cells......... 53 2.3.2. Sequence Requirement for Filopodia Induction by Palmitoylation Motifs: Adjacent Cysteines and Nearby Basic Amino Acids ............................................................................ 57 2.3.3. Filopodia Inducing Motifs Promote Filopodia Outgrowth and Enhance Dendritic and Axonal Branching in Neuronal Cells.................................................................................... 59 2.3.4. Filopodia Induction is Palmitoylation Dependent and is Reversible in Heterologous Cells and Neurons.......................................................................................................................... 64  iv  2.3.5.  The ADP-Ribosylation Factor (ARF6) Regulates Processes Outgrowth Mediated by Palmitoylated Motifs............................................................................................................. 69 2.3.6. Filopodia Induction by FIMs is also Modulated by the GTPase Cdc42............................... 74 2.3.7. Cdc42-Induced Filopodia Involves a Palmitoylation Dependent Pathway .......................... 76 2.4. Discussion ................................................................................................................................. 79 2.4.1. Minimal Sequence Required for Induction of Process Outgrowth ....................................... 79 2.4.2. Possible Mechanisms for Induction of Process Outgrowth by FIMs.................................... 80 2.4.3. Regulation of Filopodia Formation and Dendritic Branching by the GTPases ARF6 and Cdc42.................................................................................................................................... 82 2.4.4. Possible Roles in Synaptogenesis ......................................................................................... 84 2.5. References ................................................................................................................................. 87 3. Chapter III: An active role for dendritic filopodia in the formation of stable axonal-dendritic contacts........................................................................................................................................................ 92 3.1. Introduction ............................................................................................................................... 92 3.2. Materials And Methods ............................................................................................................. 94 3.2.1. Constructs Used.................................................................................................................... 94 3.2.2. Primary Neuronal Culture and Transfection........................................................................ 95 3.2.3. Immunofluorescence:............................................................................................................ 98 3.2.4. Live Imaging:........................................................................................................................ 98 3.2.5. Quantitative Measurement of Axonal Contacts, Spine Density, Filopodia Dynamics, and Synapse Number ................................................................................................................... 99 3.2.6. Calculation of synapse number .......................................................................................... 100 3.3. Results ..................................................................................................................................... 101 3.3.1. Exploratory Role for Dendritic Filopodia in Contact Initiation......................................... 101 3.3.2. Regulation of Filopodia Induction in Developing Neurons................................................ 103 3.3.3. Modulation of Filopodia Motility and Contact Stability by FIMs, Cdc42 and NLG1 ........ 107 3.3.4. Recruitment of Synaptophysin at Contact Sites is Modulated by FIMs, Cdc42 and NLG1 111 3.3.5. Involvement of Dendritic Filopodia in Spine Formation.................................................... 114 3.4. Discussion ............................................................................................................................... 121 3.4.1. Possible Roles for Dendritic Filopodia in Synaptogenic Contact Formation. ................... 121 3.4.2. Recruitment of Presynaptic Proteins at Sites of Contact by Dendritic Filopodia. ............. 122 3.4.3. Filopodia Motility is Essential in Synaptogenic Contact Formation.................................. 122 3.4.4. Signals Required for Filopodia Outgrowth and Contact Formation.................................. 124 3.4.5. A Possible Role for Dendritic Filopodia in Spine Formation............................................. 125 3.5. References ............................................................................................................................... 128 4.  CHAPTER IV: General Discussion............................................................................................... 133 4.1. Dendritic Filopodia.................................................................................................................. 134 4.1.1. Regulation and Function of Dendritic Filopodia ............................................................... 134 4.2. Development of Central Nervous System Synapses................................................................ 136 4.2.1. Filopodia Initiate Synaptogenic Contacts .......................................................................... 137 4.2.2. Recruitment of Protein Complexes at Synaptic Contacts Sites........................................... 138 4.2.3. Role for Filopodia in Spine Formation............................................................................... 140 4.2.4. Alternative Roles for Dendritic Filopodia .......................................................................... 143 4.3. Future work ............................................................................................................................. 145 4.3.1. Does Activity Regulate Filopodia Outgrowth?................................................................... 145 4.3.2. Released Molecules Required for Guiding Filopodia......................................................... 146 4.3.3. In Vivo Work....................................................................................................................... 148 4.4. References ............................................................................................................................... 150  v  LIST OF TABLES  TABLE 1-1. COMPARISON OF THE FOUR MAJOR FORMS OF LIPID MODIFICATIONS: PALMITOYLATION, PRENYLATION, MYRISTOYLATION AND GLYPIATION. ...................................................................................12 TABLE 2-1. SUMMARY OF CHIMERIC CONSTRUCTS USED IN THIS STUDY. ..............................................................56  vi  LIST OF FIGURES  FIGURE 1-1. DENDRITIC FILOPODIA ........................................................................................................................3 FIGURE 1-2. SIGNALING PATHWAYS DOWNSTREAM OF CDCD42. ............................................................................8 FIGURE 1-3. CDC42 AND ACTIN POLYMERIZATION. ..............................................................................................10 FIGURE 1-4. A CLUSTER OF FILOPODIA EMERGING FROM A DENDRITE SHAFT. ......................................................17 FIGURE 1-5. MORPHOLOGY OF DENDRITIC SPINES AND FILOPODIA .......................................................................22 FIGURE 1-6. MODELS OF SPINOGENESIS ................................................................................................................25 FIGURE 2-1. FILOPODIA INDUCTION IN COS-7 CELLS. ...........................................................................................55 FIGURE 2-2. FILOPODIA INDUCTION AND BRANCHING IN NEURONAL CELLS. .........................................................60 FIGURE 2-3. FILOPODIA FORMATION AND DENDRITIC BRANCHING IN NEURONS OVEREXPRESSING PARALEMMIN IS PALMITOYLATION DEPENDENT...........................................................................................62 FIGURE 2-4. AXONAL BRANCHING INDUCED BY FIMS. .........................................................................................63 FIGURE 2-5. PALMITATE TURNOVER ON FIMS.......................................................................................................65 FIGURE 2-6. ON-GOING PALMITOYLATION OF FIMS REGULATES FILOPODIA FORMATION IN COS-7 CELLS...........66 FIGURE 2-7. FILOPODIA INDUCTION IS PALMITOYLATION-DEPENDENT AND PARTLY REVERSIBLE IN NEURONAL CELLS...........................................................................................................................................................68 FIGURE 2-8. ARF6 REGULATES FIM-INDUCED FILOPODIA EXTENSION IN COS-7 CELLS. ......................................71 FIGURE 2-9. ARF6 REGULATION OF DENDRITIC BRANCHING AND FILOPODIA EXTENSION IN NEURONAL CELLS. ...73 FIGURE 2-10. REGULATION OF FILOPODIA EXTENSION AND DENDRITIC BRANCHING BY CDC42............................75 FIGURE 2-11. PALMITOYLATION MODULATES CDC42-INDUCED FILOPODIA EXTENSION AND DENDRITIC BRANCHING. ................................................................................................................................................78 FIGURE 3-1. TRANSFECTION METHOD USING TWO FLUORESCENTLY-LABELED CONSTRUCTS. ...............................97 FIGURE 3-2. ROLE FOR DENDRITIC FILOPODIA IN EXPLORATION AND CONTACT INITIATION. ...............................102 FIGURE 3-3. SUMMARY OF CONSTRUCTS USED....................................................................................................104 FIGURE 3-4. DETECTION OF DENDRITIC FILOPODIA AT SITES POSITIVE FOR PRESYNAPTIC MARKERS. ..................105 FIGURE 3-5. FILOPODIA AND SPINE MORPHOLOGY. .............................................................................................107 FIGURE 3-6. FILOPODIA MOTILITY MODULATES CONTACT FORMATION. .............................................................108 FIGURE 3-7. FILOPODIA STABILITY AND CONTACT FORMATION...........................................................................110 FIGURE 3-8. AXONAL GROWTH CONE INITIATES CONTACT FORMATION..............................................................111 FIGURE 3-9. ACCUMULATION OF PRESYNAPTIC PROTEINS AT SITES OF FILOPODIA CONTACTS............................113 FIGURE 3-10. CHANGES IN SPINE DENSITY AND SYNAPSE NUMBER IN NEURONS AT DIV 14. ..............................115 FIGURE 3-11. VISUALIZATION OF FILOPODIA TRANSFORMATION INTO SPINY SYNAPSES. ....................................119  vii  LIST OF ABBREVIATIONS 2-amino-5-phosphono-valeric acid (D, L-APV) 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) ABPs = actin-binding proteins ADP = adenosine diphosphate AMPA = α-amino-5-hydroxy-3- methyl-4-isoxazole propionic acid AMPAR = AMPA receptor ANOVA = analysis of variance ARF6 = ADP-ribosylation factor 6 Arp2/3 = actin-related proteins 2 and 3 ATP = adenosine triphosphate Ca++ = calcium Cdc42 = cell division cycle 42 cDNA = complementary deoxyribonucleic acid CoA = coenzyme A COS-7 = african green monkey kidney cell line CNS = central nervous system DNA = deoxyribonucleic acid DIV = days in vitro EGFP = enhanced green fluorescent protein EPSC = excitatory postsynaptic current ERM = Ezrin/Radixin/Moesin  viii  FIMs = filopodia inducing motifs G-protein = guanine nucleotide binding protein GABA = γ-aminobutryic acid GAD-65 = glutamic acid decarboxylase 65 GAP-43 = growth associated protein 43 GFP = green fluorescent protein GPCR = G-protein coupled receptors GTPase = guanosine trinucleotide phosphatase H = hours LIMK = LIM kinase LTD = long-term depression LTP = long-term potentiation Min = minutes mM = millimolar µM = micromolar µm = micrometer µl = microliter MLC = myosin light chain N-WASP = neural Wiskott Aldrich syndrome protein NLG-1 = neuroligin 1 NMDA = N-methyl-D-aspartate NMJ = neuromuscular junction NOS = nitric oxide synthase  ix  NSF = N-ethylmaleimide-sensitive factor Pak1 = p21 activated kinase 1 PAT = palmitoyl acyl-tranferase PCR = polymerase chain reaction PDZ = PSD-95, ZO1/2, Disklarge pH = potential of hydrogen PIP2 = phosphatidylinositol 4,5-bisphosphate PKA = protein kinase A PKG = protein kinase G PSD-95 = postsynaptic density 95 Rac1 = Ras-related C3 botullinum toxin substrate 1 RhoA = Ras homologous member A RNA = ribonucleic acid ROCK = Rho associated kinase RT-PCR = reverse transcriptase PCR SD = standard deviation SEM = standard error of the mean SH3 = src homology 3 SNAP25 = soluble NSF attachment protein 25 Syn = synaptophysin TTX = tetrodotoxin  x  ACKNOWLEDGEMENTS  Many thanks go to Drs. Timothy Murphy and Alaa El-Husseini for their excellent supervision and support during my years in the lab. Furthermore, I would like to thank Drs. Tim O’Connor and Wolfram Tetzlaff of my supervisory committee for helpful suggestions and insightful discussions. Of many helpful colleagues and lab-mates, I want to give special thanks to Andy Shih, Jaqueline Shehadeh and Pamela Arstikaitis with whom I shared a significant portion of my graduate student years and who helped me with advice and lasting friendship. Furthermore, I want to thank Esther Yu and Alex Trinh for their assistance in neuronal cell culture preparation. Lastly, I deeply want to thank Danielle for being so supportive and my friends and family for their continuing encouragements.  During my dissertation I was supported by grants from the National Science and Engineering Council of Canada (NSERC).  xi  DEDICATION  First and foremost, I would like to dedicate this thesis to my beloved mother, Colette Gauthier, who passed away last year, after a courageous battle against cancer. She supported me every step of the way and has always been an inspiration.  I would also like to make a special dedication to my supervisor Alaa El-Husseini, who passed away in December 2007. Alaa was a great mentor, researcher and friend, and I will miss him greatly. I have learnt so much from Alaa and will never forget his dedication to science and passion for life.  xii  CO-AUTHORSHIP STATEMENT  I would like to thank Pamela Arstikaitis for her help in collecting images and analyzing data that was used in the preparation of this thesis. In particular, Pamela contributed expertise on some of her work with Neuroligin and Shank1b. Dr. David Bredt contributed valuable insight and comments to the preparation of the manuscript for Chapter 2.  xiii  1.  CHAPTER I: Overview and Summary  1.1. Wiring of the Brain The human brain is composed of over a hundred billion neurons that communicate with each other via connections called synapses. Synapses are specialized intercellular junctions whose specificity and plasticity provide the structural basis for the formation and maintenance of complex neural networks in the brain. In humans, the majority of synapses form during early prenatal and postnatal development.  The  processes underlying synapse formation within the central nervous system (CNS) have fascinated neuroscientists for many decades.  Historically, attempts aimed at  understanding how the CNS is ‘wired’ were based primarily on the analysis of fixed and stained tissue using light microscopy (Ramon y Cajal, 1888). Recently, the introduction of contemporary molecular and optical techniques has led to diverse and intricate studies devoted to this subject.  However, the molecular mechanisms through which initial  synaptic contacts are established and modified during synaptogenesis have not yet been fully determined.  Significant progress has been made in the characterization of the structural, functional and developmental assembly of CNS synapses, particularly that of glutamatergic synapses. Proteomic and genetic approaches by a number of researchers have led to the identification of over a thousand different proteins found at synapses (Collins et al., 2006; Husi and Grant, 2001; Scannevin and Huganir, 2000). Further, the development of sophisticated fluorescence techniques has enabled the visualization in real-time of assembly of individual molecules at the synapse. These complimentary  1  approaches are beginning to yield clues as to how synapses are formed in the CNS. Synapse assembly is considered to be a multi-step process that is initiated at sites of contact between a presynaptic and postsynaptic neuron. This initial contact is thought to be followed by recruitment of proteins at the site of contact and subsequent pre- and postsynaptic differentiation. In particular, several classes of molecules appear critical in the development and maintenance of synaptic contacts between cells. Among these are important classes of cell adhesion molecules, as well as scaffolding molecules and neurotransmitter receptors which will be discussed in greater detail below. Although we are beginning to uncover the molecules involved in synapse assembly, very little is known about the mechanisms involved in the formation of a contact between a pre- and postsynaptic neuron. Axonal growth cones are thought to extend and initiate contacts with the dendritic arbors of target neurons. Synaptogenic contacts can also be initiated by dendritic growth cones, as well as motile structures called dendritic filopodia. Indeed, recent studies revealed that numerous synapses formed during development are located on dendritic filopodia (Fiala et al., 1998; Saito et al., 1992). These observations have pointed to filopodia as possible candidates for initiating synaptogenic contacts, however, this function is still unclear and controversial (Yuste and Bonhoeffer, 2004).  1.2. Dendritic Filopodia 1.2.1. Structure and Function of Dendritic Filopodia Dendritic filopodia are cell surface extensions typically 5-20 µm in length, which are filled with tight parallel bundles of actin filaments and rapidly alter their position and  2  shape (Dailey and Smith, 1996). In developing axons, filopodia occur at the tips of growth cones and aid the axon in finding its appropriate target. In dendrites, filopodia emerge from the dendritic shaft and are thought to be precursors to spines, the site of most excitatory connections in the brain (Dailey and Smith, 1996; Saito et al., 1997; Small, 1988).  However, in contrast to spines, little is known about the molecular  organization of dendritic filopodia and the molecules that regulate their motility and shape. Technical limitations associated with the small size and short half-life of filopodia in neuronal cells (often less than a minute), has shifted much of the attention to heterologous systems. Hence, much of the understanding of filopodia motility has been inferred from experiments involving molecules present in filopodia from fibroblasts or epithelial cells (Nobes and Hall, 1995a). Further, since the lifetime of dendritic filopodia is very short and their expression in the brain is mostly limited to development, the majority of research on neuronal protrusions has focused on dendritic spines.  Figure 1-1. Dendritic Filopodia Hippocampal neurons were transfected at DIV 7 with GFP and fixed at DIV 8. Representative images show the typical morphology of dendritic filopodia (arrowheads). Scale bar 5µm.  3  The precise role and function of dendritic filopodia is still controversial. They have been seen to act as precursors to spines (Jontes and Smith, 2000b). Given their ubiquitous distribution in development and the evidence that they exist transiently in non spiny neurons, dendritic filopodia have also been assigned an exploratory role independent of spinogenesis (Dvergsten et al., 1986; Linke et al., 1994; Lund et al., 1977; Mason, 1983; Ulfhake and Cullheim, 1988; Wong et al., 1992). Other studies have proposed a potential role of filopodia in guiding the growth of dendrites (Portera-Cailliau et al., 2003; Portera Cailliau and Yuste, 2001; Vaughn, 1989; Yuste and Bonhoeffer, 2001). Thus the goal of this study is to begin to test these hypotheses by investigating the dynamics of dendritic filopodia and determining their role in the development of synaptic contacts.  1.2.2. Filopodia Development: the Actin Cytoskeleton and Actin-Binding Proteins Actin filaments constitute the main cytoskeletal element of filopodia and dendritic spines. In fact, the β- and γ-isoforms of actin, which are expressed in neurons, are selectively targeted to spines and filopodia, where they are found in higher concentrations than in the dendritic shaft (Cohen 1985, Kaech 1997, Matus 1982, Wyszynski 1997). Within spines and filopodia, actin is present as a soluble pool of monomeric G-actin and as polymerized F-actin filaments that confer the characteristic morphology of these structures (Halpain, 2000; Rao and Craig, 2000; Small, 1988). The actin filaments in filopodia and in the spine neck are arranged in longitudinal bundles, whereas those within the spine head are organized into a meshwork (Chang and De Camilli, 2001; Landis and Reese, 1983; Matus et al., 1982). It is widely believed that the regulated  4  polymerization/depolymerization of actin is responsible for filopodia and spine motility, growth and structural integrity (Fischer et al., 1998; Forscher and Smith, 1988). Hence, molecules that regulate the actin cytoskeleton, such as members of the family of Rho GTPases, likely play a role in regulating filopodia morphology and motility (Nobes and Hall, 1995b).  1.2.3. The Rho family of GTPases A variety of proteins regulate the assembly, stability and disassembly of actin filaments. Among them is a family of actin-binding proteins (ABPs), comprising proteins such as profilin that catalyze the exchange of ADP to ATP; the Arp2/3 complex that nucleate F-actin; capping proteins like ADF/cofilin and gelsolin; crosslinking proteins such as alpha-actinin and proteins of the Ezrin/Radixin/Moesin (ERM) family that serve to attach actin to the plasma membrane (Winder and Ayscough, 2005). One particular family of such proteins thought to regulate actin dynamics has been extensively characterized: the Rho GTPases.  The Rho family, which includes RhoA (Ras  homologous member A), Rac1 (Ras-related C3 botullinum toxin substrate 1), Cdc42 (cell division cycle 42), and many others, are key regulators of the actin cytoskeleton (EtienneManneville and Hall, 2002; Ridley, 1999; Ridley, 2001).  The Rho GTPases are  molecular switches that cycle between an active form (GTP-bound) and an inactive form (GDP-bound). In the active form, they are capable of binding to downstream effectors and activating signaling cascades. Some of these proteins have also been shown to regulate actin dynamics in dendritic spines and filopodia. Indeed, Meng et al. demonstrate that Lim-kinase (LIMK), through its interaction with ADF/cofilin, is  5  important in the regulation of the actin cytoskeleton and spine morphology (Meng et al., 2002). Consistent with these findings, the role of Rho GTPases in controlling actin organization and cell morphology is well established (Hall and Nobes, 2000; Luo, 2000; Nakayama et al., 2000; Nobes and Hall, 1995b; Olenik et al., 1997; Ridley and Hall, 1992). In heterologous cells, activation of wild-type RhoA promotes stress fiber formation, whereas activation of Rac1 and Cdc42 induce lamellipodia and filopodia formation, respectively (Nobes and Hall, 1995b).  Although the expression of Rho  GTPases is widespread throughout the CNS, their direct effects on neuronal morphology seem to be less clear and less characterized, partly because the small GTPases have varied effects in different cell types and systems. For example, experiments in Xenopus retinal ganglion cells show that expression of dominant negative Rac1 or Cdc42 causes a decrease in dendritic arbor complexity (Ruchhoeft et al., 1999), consistent with findings that overexpression of constitutively active forms of Rac1 and Cdc42 promote neurite extension in cortical neurons (Luo, 2000; Threadgill et al., 1997).  Conversely,  mushroom body neurons from Drosophila lacking Cdc42 show no apparent defects in dendritic length and branching (Scott et al., 2003). Further, in vitro and in vivo studies sometimes provide opposing results; RhoA activation has been shown to cause growth cone collapse and neurite retraction in hippocampal neurons in culture, Xenopus retinal ganglion cells and Drosophila mushroom body neurons (Lehmann et al., 1999; Luo et al., 1996b; Nakayama et al., 2000; Ruchhoeft et al., 1999; Sebok et al., 1999), whereas removal of RhoA in Drosophila mushroom body neurons led to increased dendritic extension and expression of a dominant negative RhoA in Xenopus tectal cells resulted in an increase in dendritic length (Lee et al., 2000; Li et al., 2000).  Nonetheless, a  6  consensus is emerging in the field that Rho GTPases are key regulators of dendritic branching and morphogenesis. In summary, the above studies are generally consistent with a role for Rac1 and Cdc42 in promoting the formation of cellular protrusions in neurons, whereas RhoA acts to control dendritic length.  Recent evidence indicates that the different Rho GTPases may also play a role in regulating the formation and remodeling of dendritic filopodia and spines (Tashiro et al., 2000). The first evidence of the effects of the Rho family of GTPases on dendritic spines came from Luo et al. who found that constitutively active Rac1 (G12V) increases spine density in Purkinje cells (Luo et al., 1996a). Consistent with these findings, expression of Rac1-G12V in hippocampal or cortical neurons led to an overproduction of spines with abnormal morphology, whereas expression of the dominant negative Rac1-T17N mutant caused a reduction in spine density (Nakayama et al., 2000; Tashiro et al., 2000). Similarly, loss of Cdc42 function in neurons of the Drosophila visual system causes a 50% reduction in the density of spine-like structures (Scott et al., 2003) and expression of dominant-negative Cdc42 in dissociated hippocampal neurons prevents spine development (Irie and Yamaguchi, 2002). These results are somewhat controversial because Irie et al. also find that expression of dominant-negative Cdc42 promotes filopodia extensions, whereas other groups have shown that expression of the constitutively active Cdc42 causes an increase in filopodia extensions (GauthierCampbell et al., 2004; Mackay et al., 1995) or have no effect on spine density (Tashiro et al., 2000). Although the precise role of Cdc42 is presently not well-defined, there is growing evidence for its involvement in filopodia extension and spine morphogenesis  7  (Calabrese et al., 2006).  Figure 1-2 illustrates a portion of signaling pathways  downstream of Cdc42 that regulate actin dynamics in dendritic spines.  Figure 1-2. Signaling pathways downstream of Cdcd42. Signaling pathways downstream of the Rho GTPase Cdc42 that regulates actin dynamics in dendritic spines. Figure adapted from (Ethell and Pasquale, 2005).  8  1.2.4. Signaling Pathways Connecting Cdc42 to the Actin Cytoskeleton In contrast to their role in regulating spine morphology and dynamics, very little is known about the role of Rho GTPases in regulating dendritic filopodia. Under normal conditions the formation of filopodia in many cell types is triggered by the binding of the active GTP bound form of Cdc42 along with phosphatidyl inositol 4,5 bisphosphate (PIP2) to N-WASP, a member of the Wiskott-Aldrich syndrome (WASP) family of proteins that is highly expressed in neurons (Irie and Yamaguchi, 2002) (see Figure 1-3). This interaction leads to the activation of N-WASP, which promotes the recruitment of G-actin, profilin and the formation of a complex with Arp2/3 (actin-related proteins 2 and 3), a protein that regulates the nucleation of actin filaments (Rohatgi et al., 1999; Suetsugu et al., 2001; Yarar et al., 1999). All these findings suggest that activation of Cdc42 at specific sites at the plasma membrane may locally increase the concentration of actin complexes to promote actin-mediated elongation and membrane-protrusion formation.  9  Figure 1-3. Cdc42 and actin polymerization. Illustration of how actin is thought to be regulated through the Cdc42 pathway. Cdc42 promotes actin polymerization by activating the Arp2/3 nucleation complex through its interaction with N-WASP.  10  1.2.5. A Role for Acylated Proteins in Filopodia Formation The covalent attachment of lipid moieties is an essential modification found on many proteins. As described below in Table 1-1, four major forms of lipid modification have been identified thus far: addition of glycosyl-phosphatidylinositol (GPI) anchors, Nmyristoylation, C-terminal farnesylation (or prenylation) and S-acylation (see Figure 1-3) (Resh, 1999; Wilcox et al., 1987; Zhang and Casey, 1996). S-acylation is one of the most frequent and most versatile post-translational modifications found on proteins.  It  contributes to protein sorting, trafficking, membrane association and regulating synaptic strength (Huang and El-Husseini, 2005).  In most cases, S-acylation involves the  attachment of the palmitic acid, a 16-carbon saturated fatty acid, to cysteine residues via thioester linkages. Although the attachment of stearic, oleic or arachidonic acids are also possible, palmitoylation is more commonly used to describe the S-acylation of proteins (Bijlmakers and Marsh, 2003). Interestingly, whereas myristoylation and isoprenylation are usually stable modifications, palmitoylation is reversible and has been shown to regulate specific signaling pathways (El-Husseini and Bredt, 2002; Resh, 1999; Smotrys and Linder, 2004).  11  Table 1-1. Comparison of the four major forms of lipid modifications: palmitoylation, prenylation, myristoylation and glypiation. Palmitoylation1  Prenylation2  Myristoylation3  Glypiation4  Group  Saturated fatty acid (almost always 16carbon palmitic acid)  Isoprenoid group (15carbon farnesyl or 20carbon geranylgeranyl)  Saturated fatty acid (always 14-carbon myristic acid)  Phosphatidyl inositol group (GPI anchor)  Bond  Thioester  Thioester  Amide  Carbohydrate linker  Location of modification  Cysteine residues, anywhere in the protein  C-terminal cysteine residue (CAAX motif)  N-terminal glycine  C-terminal amino acid after signal cleavage  Reversibility  Yes, thioesterases  No, stable bond  No, stable bond  No, stable bond  Enzymology  Palmitoyl-acyl transferases (PAT)  Farnesyltransferase or geranylgeranyltransferase  N-myristoyltransferase (NMT)  Complex synthesis  Type of proteins modified  Intracellular or transmembrane proteins  Intracellular proteins  Intracellular proteins  Cell surface proteins  Effect of modification  Trafficking of proteins, and localization to membrane compartments  Association with membranes  Association with membranes  Association of proteins to extracellular cell membrane  Examples of proteins modified  GAP-431, PSD-951, GAD-651, G-proteins1, AMPARs1, GPCRs1  Ras and Rab proteins2, Gproteins2, PSD-952  G-proteins3, PKA3, PKG3, NOS3, fyn3, src3, lyn3, lsk3  Acetylcholinesterase4, placental alkaline phosphatase4, Thy-14  1  For an excellent review of palmitoylation, see (El-Husseini and Bredt, 2002) For an excellent review of prenylation, see (Zhang and Casey, 1996) 3 For an excellent review of myristoylation, see (Farazi et al., 2001) 4 For an excellent review on GPI-anchored proteins, see (Ikezawa, 2002; Karagogeos, 2003) 2  The mechanisms involved in palmitoylation are poorly understood: indeed, unlike prenylation which only occurs at the carboxy-terminal cysteine-aliphatic-aliphatic-X (CAAX) consensus sequence (Hancock et al., 1989), there is no exact consensus sequence for palmitoylation. Further, very little is known about the enzymes involved in the process of palmitoylation, mostly due to difficulties in isolating the activity of these membrane-associated enzymes. However, very recent work has identified 23 palmitoyl  12  acyl-tranferases (PATs) expressed in mammalian cells that are responsible for the palmitoylation of various neuronal proteins (Fukata et al., 2004; Huang et al., 2004). Future work will be required to characterize these enzymes further, to identify their subcellular localization and how they are regulated.  Protein palmitoylation has emerged as an important factor in regulating protein trafficking and function. Indeed, classic studies have shown that palmitoylation controls the proper targeting of many enzymes to specialized lipid microdomains such as lipid rafts, thereby directing their integration into specific signaling pathways (Arni et al., 1998; Perez and Bredt, 1998). More recent work shows that palmitoylation occurs on various classes of proteins, including neurotransmitter receptors, synaptic scaffolding proteins and secreted molecules (Bijlmakers and Marsh, 2003; Huang and El-Husseini, 2005; Smotrys and Linder, 2004). At presynaptic nerve terminals, palmitate modifies a variety of synaptic vesicle proteins such as SNAP25, synaptotagmin I and VII, that regulate neurotransmitter release (Kang et al., 2004; Washbourne et al., 2001). On the postsynaptic side, the palmitoylation of ion channels such as L-type calcium channels and neurotransmitter receptors such as AMPARs regulates signal transduction (Chien et al., 1996; Dunphy and Linder, 1998; Hayashi et al., 2005). Furthermore, several synaptic scaffolding proteins such as postsynaptic density 95 (PSD-95), which organize receptors and their associated signaling elements, are palmitoylated, and this regulates their membrane association and receptor-clustering function (DeSouza et al., 2002; ElHusseini et al., 2002; Topinka and Bredt, 1998). Below, I discuss recent evidence that  13  protein palmitoylation may regulate trafficking and signaling pathways that are important for neuronal outgrowth and synapse formation.  1.2.6. Palmitoylated Proteins and their Role in Filopodia Formation Many acylated proteins mediate interactions between the plasma membrane and the neuronal cytoskeleton, thereby playing a role in regulating process outgrowth and dynamics. Such proteins include growth associated protein 43 (GAP-43), which has been shown to regulate growth cone dynamics (Strittmatter et al., 1994a), and paralemmin, which is implicated in the regulation of membrane dynamics and dendritic outgrowth (ElHusseini and Bredt, 2002; Resh, 1999; Strittmatter et al., 1994a). Studies on the neuronal proteins GAP-43 and paralemmin show that palmitoylation is important for localizing these proteins to filopodia (Kutzleb et al., 1998; Zuber et al., 1989a; Zuber et al., 1989b). Interestingly, recent work has explored the importance of specific protein motifs that can undergo lipid modification (El-Husseini et al., 2001). In particular, we have identified specific palmitoylation motifs (see Table 2-1) that can dramatically induce changes in cytoskeleton and regulate filopodia outgrowth and dendritic branching in heterologous cells and primary cultures of hippocampal neurons (see Chapter II) (Gauthier-Campbell et al., 2004; Kutzleb et al., 1998; Strittmatter et al., 1994a; Strittmatter et al., 1994b). Thus, palmitoylation may serve as a signal for delivery of proteins involved in the regulation of cell morphology and membrane dynamics to specific active sites of the plasma membrane.  I propose that these palmitoylation motifs may play a role in  filopodia formation and the regulation of filopodia dynamics.  14  1.3. A Role for Filopodia in Synaptogenesis 1.3.1. Axo-Dendritic Contact Formation Formation of the proper synaptic contacts during development is a critical step in the wiring of the brain (Gerrow and El-Husseini, 2006; Waites et al., 2005; Yamagata et al., 2003). To define the sequence of events that underlie synapse formation, it is critical to understand the molecular mechanisms by which initial contacts are formed between pre- and postsynaptic neurons, and how appropriate axonal and dendritic protein components are recruited to initial sites of contact.  Dendritic filopodia represent  potential key structural elements that help probe the environment and establish these neuronal contacts (Fiala et al., 1998; Jontes and Smith, 2000b).  However, the  mechanisms through which initial synaptic contacts are established and modified during synaptogenesis have not been fully determined, and the molecules that regulate this process are largely unknown. I will now focus on molecules that may play a role in regulating the initial contact formation between a dendrite and axon, and the subsequent maturation into a synapse.  1.3.2. Synaptogenesis at Filopodial Contact Sites Despite the importance of proper synaptogenesis in the CNS, the molecular mechanisms that regulate the formation and development of synapses remain relatively poorly understood.  Recent studies have revealed some of the events that regulate  synapse formation (Ahmari et al., 2000; Friedman et al., 2000; Sanes and Lichtman, 2001; Washbourne et al., 2002; Ziv and Garner, 2004). In particular, there is evidence  15  for rapid recruitment of components of the synaptic vesicle release machinery to contact sites (Friedman et al., 2000; Gerrow et al., 2006). Other studies also documented the existence of transport packets and protein complexes of postsynaptic proteins to nascent neuronal contacts (Gerrow et al., 2006; Marrs et al., 2001; Prange and Murphy, 2001; Sans et al., 2003; Washbourne et al., 2002). These studies suggest that recruitment of synaptic elements to nascent sites of contact plays a role in contact stabilization and synapse maturation.  Previous studies, primarily involving live imaging of neurons, have suggested a role for filopodia dynamics in synaptogenesis and the remodeling of connectivity (Jontes and Smith, 2000a). One model for synapse formation predicts that active dendritic filopodia contact axons to induce presynaptic boutons, followed by a period of filopodial maturation into postsynaptic spines (Harris et al., 1992; Maletic-Savatic et al., 1999; Rao and Craig, 2000; Ziv and Smith, 1996). Indeed, electron microscopy studies (EM) of developing brain show synapses at the tips and base of some dendritic filopodia (see Figure 1-4) (Fiala et al., 1998). The authors find that approximately 20% of synapses occur on dendritic filopodia in animals aged P1 to P6.  16  Figure 1-4. A cluster of filopodia emerging from a dendrite shaft. The filopodia (copper) receive synaptic contacts near the tips (1,6) and at the base (3,4,5). An additional shaft synapse (2) can be seen on the surface of the dendrite. Scale bars in insets, 0.5µm; 3D scale bar, 1µm (Fiala et al., 1998). Figure reprinted with permission from the Journal of Neuroscience, © 2008 by the Society for Neuroscience.  1.4. Molecules that Regulate Synaptogenesis While synaptogenesis in the verterbrate neuromuscular junction has been extensively studied, in comparison very little is known about the molecular mechanisms mediating synaptogenesis in the central nervous system (CNS) (Campagna et al., 1995; Dai and Peng, 1995; Porter et al., 1995; Sanes and Lichtman, 1999). For example, evidence exists that agrin is sufficient to induce presynaptic differentiation in motorneurons, however it is unclear whether agrin plays a role in synapse formation in the CNS (Campagna et al., 1995). Indeed, while agrin is found at central synapses, mice  17  lacking agrin do not show any defects in CNS synaptogenesis (Serpinskaya et al., 1999). Cell surface proteins that are involved in cell-cell interactions between pre- and postsynaptic neurons are likely to play an important role in triggering the initial formation of synapses and dendritic spines in the CNS (Kossel et al., 1997). Some molecules that have recently been implicated in the initial stages of synapse formation include members of the cadherin family of proteins (Fannon and Colman, 1996; Kohmura et al., 1998; Uchida et al., 1996), neurexins and neuroligins (Prange et al., 2004; Scheiffele et al., 2000; Ushkaryov et al., 1992), EphB receptor tyrosine kinases and their ephrin-B ligands (Buchert et al., 1999; Grunwald et al., 2001; Henderson et al., 2001; Torres et al., 1998), scaffolding molecules such as Shank and PSD-95 (El-Husseini et al., 2000b; Sala et al., 2001), and number of other potential candidate molecules (Ethell and Pasquale, 2005). Here we will briefly discuss some of these cell surface proteins that are key regulators of synapse formation.  1.4.1. Trans-Synaptic Signaling: Neuroligins and Neurexins The idea that synapses are generated following contact and in response to an exchange of signals initiated between pre- and postsynaptic adhesion elements is not novel.  Hence, a large amount of research is currently being directed towards cell  adhesion molecules as initiators of synapse formation. In fact, one class of cell adhesion molecules has been found to be highly synaptogenic, mediating recognition between dendrites and the axon, and the specification of excitatory versus inhibitory synapse formation: the neuroligins (Levinson et al., 2005; Prange et al., 2004; Scheiffele et al., 2000).  Neuroligins 1, 2, 3 and 4 are postsynaptic transmembrane proteins enriched at  18  excitatory postsynaptic sites. The neuroligins associate with the third PDZ domain of PSD-95 through their C-terminal PDZ-binding site (Ichtchenko et al., 1995) and this binding is thought to regulate neuroligin 1 and 2 clustering at excitatory and inhibitory synapses respectively (Irie et al., 1997; Prange et al., 2004). Further, neuroligins bind with high affinity to presynaptic β-neurexins, and both molecules are thought to participate in synaptic contact formation through trans-synaptic heterophilic protein interactions (Ichtchenko et al., 1995; Nguyen and Sudhof, 1997; Song et al., 1999). Remarkably, neuroligin 1 and neuroligin 2 can induce the formation of functional synapses in axons contacting non-neuronal cells, and these effects are abolished by disruption of the neuroligin-neurexin protein interaction (Scheiffele et al., 2000). Thus, is appears that neuroligins and β-neurexins may function as early initiators of synaptic contact formation between pre- and postsynaptic structures and may play a role in establishing synapse specification.  However, the signaling pathways activated  downstream of the neuroligins are largely unknown. Possible cooperative effects of neuroligins/neurexins with other trans-synaptic molecules or with scaffolding proteins may play a role in the development of synapses.  1.4.2. Scaffolding molecules Scaffolding proteins have been shown to play a role in the stabilization and maturation of spines (Prange and Murphy, 2001; Sala et al., 2001; Ziv, 2001). Indeed, dispersal of postsynaptic density (PSD) proteins by disruption of lipid rafts causes the loss of dendritic spines (Hering and Sheng, 2003). Further, perturbing the function of scaffolding proteins also affects spine morphology, as well as the clustering and  19  stabilization of receptors and other components of the PSD (Garner et al., 2000; Kim and Sheng, 2004). Several key scaffolding molecules are present at the post-synaptic density, including the PSD-95/SAP102 family of proteins, as well as the Shank and Homer families.  1.4.3. The Role of Shank1B in Synaptogenesis The Shank family of proteins (also known as ProSAP, Synamon, Spank, SSTRIP and cortBP) share common domain organization consisting of ankyrin repeats, a Src Homology 3 (SH3) domain, a PSD-95/Discs large/zona occludens-1 (PDZ) domain, a proline-rich region and a SAM domain at the C terminus (Boeckers et al., 1999; Ehlers, 1999; Lim et al., 1999; Naisbitt et al., 1999; Sheng and Kim, 2000; Tu et al., 1999; Zitzer et al., 1999). Complex alternative splicing of three Shank genes generates multiple isoforms of the Shank protein (Boeckers et al., 1999; Lim et al., 1999). Shank family of proteins have been shown to interact with multiple proteins within dendritic spines, including the guanylate kinase-associated protein (GKAP), the actin-binding protein cortactin, and the scaffolding protein Homer. Through its interactions with Homer and GKAP, Shank proteins can link the NMDA and metabotropic glutamate receptor complexes at the synapse (Boeckers et al., 1999; Naisbitt et al., 1999; Tu et al., 1999). Since it lies at the cytoplasmic face of the PSD and interacts with cortactin as well as glutamate receptors, Shank is thought to play a role in regulating the postsynaptic cytoskeleton in response to synaptic activity (Sala et al., 2001). Indeed, Shank1B was shown to promote spine maturation and spine head enlargement in hippocampal neurons (Sala et al., 2001) and is sufficient to induce spine formation in aspiny neurons  20  (Roussignol et al., 2005). The molecular mechanisms underlying these morphological changes are thought to be associated with the ability of Shank1B to recruit other proteins at synaptic sites, such as F-actin, Homer, PSD-95, GKAP, the IP-3 receptor and the NR1 subunit of the NMDA receptor (Sala et al., 2001). Interestingly, Shank1B was also shown to reduce the density of filopodia-like structures in these neurons. These findings imply that Shank protein is a limiting factor for the maturation of the spine head and may contribute to the rapid maturation of filopodia into spines.  1.5. A Role for Filopodia in Dendritic Spine Formation 1.5.1. Structure and Function of Dendritic Spines The majority of excitatory synapses in the brain occur at the tip of small protrusions called dendritic spines (Harris, 1999).  These specialized structures are  critical for proper synaptic transmission and are thought to function as biochemical compartments to isolate and amplify incoming signals, (Guthrie et al., 1991; Muller and Connor, 1991; Yuste and Denk, 1995). They have been shown to play an important role in neurotransmitter signaling and many downstream events necessary for synaptic plasticity, learning and memory (Hering and Sheng, 2001). Spines are characterized by their mushroom-like morphology, their small size (about 1-2 µm) and for housing the post-synaptic machinery necessary for neurotransmitter signaling, compartmentalizing a multitude of membrane proteins, cytoskeletal elements, scaffolding proteins and secondmessenger related molecules.  Spines protrude from the dendrites of most neurons  including glutamatergic pyramidal neurons of the neocortex and hippocampus, GABAergic cerebellar Purkinje neurons and medium-spiny neurons of the striatum 21  (Calabrese et al., 2006). Although spines come in a variety of shapes and sizes, the classical shape of most mature spines consists of a bulbous head connected to the dendrite shaft by a narrow neck (see Figure 1-5). Despite their small size (head volume ~0.001-1 µm3), dendritic spines contain all the essential molecules and organelles involved in post-synaptic signaling and plasticity (Nimchinsky et al., 2002).  Figure 1-5. Morphology of dendritic spines and filopodia Illustration of the typical morphology of dendritic filopodia and various types of spines, adapted from (Hering and Sheng, 2001). Thin and stubby spines are commonly referred to as ‘immature’ spines, whereas mushroom and cup shaped spines are thought to be ‘mature’ protrusions.  Although their existence has been known for over a century (Ramon y Cajal, 1888), their function and molecular organization have only recently started to be elucidated. Technical advances in microscopy and molecular approaches have given us insight into their ultrastructural organization and complexity.  These studies have  22  revealed the incredible diversity of proteins present within a spine, particularly at the postsynaptic density (PSD). The PSD is a compact electron-dense cytomatrix of about 50nm that lies just beneath the postsynaptic membrane.  Composition analysis has  revealed hundreds of proteins concentrated at the PSD including receptors, ion channels, cell adhesion molecules, scaffolding proteins, second-messenger signaling molecules, cytoskeletal elements and many more (Hering and Sheng, 2001; Sheng, 2001; Shepherd, 1996; Walikonis et al., 2000; Zhang and Benson, 2000). Further studies have enabled the visualization of spines in real time in vivo. These experiments have revealed the highly mutable nature of dendritic spines, their rapid motility, dynamic turnover and morphological plasticity. Recent experiments have shown the correlation of altered spine structure with modifications of synaptic strength (Bonhoeffer and Yuste, 2002; Ethell and Pasquale, 2005; Kasai et al., 2002; Matus, 2005; Segal, 2005; Yuste and Bonhoeffer, 2004).  1.5.2. Spine abnormalities and brain disorders In view of the abundance of dendritic spines (our brains contain >1013 spines) and their critical role played in synaptic transmission, it is not surprising that a number of human diseases are associated with alterations with spine morphology or density. It is expected that a variety of different gene mutations lead to abnormalities in dendritic spine formation, which has been associated with synaptic dysfunction and pronounced deficits in cognitive function (Blanpied and Ehlers, 2004).  Indeed, dendritic spines are  irregularly shaped and have aberrant densities in various neurodevelopmental disorders characterized by mental retardation, such as X-linked mental retardation, Fragile X  23  syndrome, William’s syndrome, Rett syndrome, Down’s syndrome, Angleman syndrome and autism (Barnes and Milgram, 2002; Becker et al., 1991; Becker et al., 1986; Comery et al., 1997; Ferrer and Gullotta, 1990; Irwin et al., 2000; Irwin et al., 2001; Kaufmann and Moser, 2000; Rudelli et al., 1985; Stoltenburg-Didinger and Spohr, 1983; Weitzdoerfer et al., 2001; Wisniewski, 1990; Wisniewski et al., 1984); (Becker et al., 1986; Marin-Padilla, 1972; Rudelli et al., 1985; Suetsugu and Mehraein, 1980). In the adult brain, dendritic spines are lost or distorted after seizures, strokes, brain tumors, stroke and chronic alcoholism (Corbett et al., 2006; Rensing et al., 2005; Sorra and Harris, 2000; Zhang et al., 2005). Recent studies have shown decreases in spine density in neocortical neurons of patients with schizophrenia (Garey et al., 1998; Glantz and Lewis, 2000). Interestingly, several genes that are mutated in families with mental retardation encode members of the Rho family signaling pathways, such as Cdc42 exchange factors αPIX, Pak3 and faciogenital dysplasia gene1 (FDG1) (Allen et al., 1998; Govek et al., 2004; Kutsche et al., 2000; Meng et al., 2005; Meng et al., 2003; Pasteris et al., 1994) and the synaptic adhesion molecules neuroligin 1, 3 and 4 (Chih et al., 2004; Jamain et al., 2003; Laumonnier et al., 2004).  1.5.3. Spinogenesis: Different Views on the Origin of Dendritic Spines Filopodia have be proposed to be precursors of dendritic spines (see Figure 1-6) (Dailey and Smith, 1996; Small, 1988). However, this hypothesis is still controversial as there are other different views on the origin of dendritic spines (see Figure 1-6). One model is that spines can form even without a synaptic contact (Sotelo, 1978). Another model is that dendritic spines arise from synapses that initially form on the dendritic shaft  24  (Harris, 1999; Miller and Peters, 1981). The third model is that spines originate from the dendritic filopodia that are predominant in younger neurons (Vaughn, 1989; Ziv and Smith, 1996). These models will be discussed in further detail below.  Figure 1-6. Models of spinogenesis Three models of spinogenesis: (a) the Sotelo model, (b) the Miller/Peters model and (c) the filopodial model. Adapted from (Yuste and Bonhoeffer, 2004).  1.5.4. The Filopodia Hypothesis Whether filopodia are precursors to spines still remains to be determined. Some evidence points to this hypothesis (third model, Figure 1-6): filopodia and spines both protrude from the dendrite and filopodia expression during development precedes that of spines. Interestingly, dendritic filopodia can be sites for new synapse formation and some data suggest that these filopodia may become stabilized to form spines (Dailey and Smith, 1996; Fiala et al., 1998; Yuste and Bonhoeffer, 2004; Ziv and Smith, 1996). Ziv  25  and Smith (1996) first succeeded in visualizing in real-time the formation of contacts between dendritic filopodia and nearby axons in cultures of dissociated hippocampal neurons.  Based on these observations, a model was proposed in which filopodia  encounter axons, initiate synaptic contacts and undergo maturation to spines.  This  transformation infers a decrease in motility, substantial shortening, enlargement of the ‘head’ portion of the filopodium, and recruitment of the proper postsynaptic proteins and receptors. Recent evidence supports the hypothesis that contact formation triggers the maturation of filopodia into spines (Dailey and Smith, 1996; Maletic-Savatic et al., 1999; Marrs et al., 2001; Okabe et al., 2001; Trachtenberg et al., 2002). According to these studies, the function of the highly motile filopodia is to probe the environment surrounding the dendrite for an appropriate contact site on a neighboring axon. How a filopodium is guided in this process is still largely unknown, but recent findings suggest a role for glutamate in promoting filopodia extension and thus perhaps guiding it to sites of presynaptic release (Portera-Cailliau et al., 2003).  In the model above, presynaptic contacts would be required to stabilize filopodia and initiate the recruitment of proper cytoskeletal elements, scaffolding molecules, receptors and thereby trigger spine maturation.  However, several problems remain  unexplained: for example, inhibitory neurons show extensive filopodia dynamics during development, even though these neurons develop with a distinctive lack of spines (Difiglia et al., 1980; Linke et al., 1994; Lund et al., 1977; Ulfhake and Cullheim, 1988). Further, given the number of filopodia that extend and retract within a short period, they have often been assigned an exploratory role where they establish initial axo-dendritic  26  contacts, but then collapse and spines emerge de novo (Dunaevsky et al., 1999; Fiala et al., 1998; Ziv and Smith, 1996). Indeed, spines have been proposed to arise from synapses located on the dendritic shaft, consistent with the second model of spinogenesis, the Miller/Peters model (Miller and Peters, 1981).  This view originates from  observations that the majority of synapses in young pyramidal neurons are located on the dendritic shaft rather than on filopodia (Harris et al., 1992). Interestingly, live imaging studies of pyramidal neurons have documented the emergence of mature dendritic spines from shaft synapses (Dailey and Smith, 1996; Marrs et al., 2001).  In the two models described above the formation of synaptic contact induces the development of dendritic spines.  However, it is also possible that spines arise  intrinsically, without significant influence from axonal terminals, as in the case of Purkinje cells of the cerebellum (Rakic and Sidman, 1973).  Indeed, spine-like  protrusions have been shown to form on Purkinje neurons without previous axonal contact from parallel fibers (Sotelo, 1990). Furthermore, mice with mutations causing the absence of parallel fibers still develop morphologically normal dendritic spines (Sotelo et al., 1975).  From these findings, Sotelo proposed that spines can form  independently of presynaptic contacts, suggesting that synaptic proteins are not required for the genesis of all types of dendritic spines (Sotelo, 1978).  1.5.5.  Reconciling the Different Models Clearly, controversy exists as to whether or not dendritic filopodia give rise to  spines. The different models outlined above are likely to imply different molecular  27  mechanisms of spine and synapse development. It is possible that dendritic spines may emerge through different mechanisms in different types of neurons, perhaps depending on their distinctive molecular compositions. In a model where presynaptic contact is necessary to initiate spine formation, trans-synaptic molecules such as the neuroligins and neurexins (Section 1.4.2) will play a vital role at the site of axon-dendrite contact. Alternatively, in cases where the presence of a presynaptic is not necessary for spine formation, different mechanisms that do not depend on cell contact would come into play. These might include unknown intrinsic signals that could cause the clustering of molecules at the site of a spine (O'Brien et al., 1998).  What is missing from the literature is live imaging experiments that document how initial synaptic contacts between pre- and postsynaptic neurons are formed during development. In particular, long-term imaging (for several days) of developing dendrites could directly provide insight into the origin of synapses, dendritic spines and what molecules are involved in a temporal resolution (Yuste and Bonhoeffer, 2004). Further, manipulations that specifically block or enhance the development of filopodia could be very helpful in determining the role of particular classes of proteins in spinogenesis and synaptogenesis.  1.6.  Research Hypothesis and Objectives  My principal aim is to clarify the role of filopodia in developing neurons and assess whether filopodia can actively participate in the formation of axo-dendritic  28  contacts, which may in turn lead to the development of synapses. Here we will focus on the molecules involved in this process and factors that regulate filopodia dynamics and the potential maturation of filopodia into spines. The following aims will test this hypothesis:  Aim 1: Examine the regulation of filopodia outgrowth and dynamics Determine the role of specific proteins, namely the palmitoylation motifs of GAP43 and paralemmin in the regulation of filopodia outgrowth and dendritic branching in heterologous cells and primary cultures of hippocampal neurons. This work will assess the role of palmitoylation as a signal for delivery of proteins involved in the regulation of cell morphology and membrane dynamics to specific active sites of the plasma membrane. I propose that these palmitoylation motifs may play a role in filopodia formation and the regulation of filopodia dynamics. The mechanisms by which these proteins are regulated will also be addressed.  Aim 2: Assess the role of filopodia in axo-dendritic contact formation, synaptogenesis and spine formation To date, long-term imaging of neurons, i.e. more than a few hours, has been relatively difficult. Interestingly, not many studies have been able to describe the initial filopodia contact with a presynaptic axon. Here, a system will be developed for imaging neurons over a period of several hours and subsequently over several days, using a double transfection protocol to label both pre- and postsynaptic cells. This system will  29  be employed to monitor in vitro contact formation between filopodia and axonal terminals using long-term live imaging. Several issues will be addressed, namely: a) if filopodia are postulated to being precursors of synapses, then we will be able to monitor the filopodia forming contacts where synapses develop and perhaps mature into dendritic spines over a period of hours or days, or b) filopodia may have a more exploratory role, or c) some filopodia may actually serve as growing dendritic branches. This live imaging will not only provide evidence for the filopodia hypothesis, but also for the effects of FIMs on filopodia dynamics and motility. This experimental work will help elucidate the fate of dendritic filopodia and their relationship with spines and synapses.  30  1.7.  References  Ahmari, S.E., J. Buchanan, and S.J. Smith. 2000. Assembly of presynaptic active zones from cytoplasmic transport packets. Nat Neurosci. 3:445-51. Allen, K.M., J.G. Gleeson, S. Bagrodia, M.W. Partington, J.C. MacMillan, R.A. Cerione, J.C. Mulley, and C.A. Walsh. 1998. 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CHAPTER II: Regulation of dendritic branching and filopodia formation by specific acylated protein motifs1 2.1.  Introduction Neurons possess an elaborate plasma membrane architecture that includes axons,  dendrites and synaptic sites (Da Silva and Dotti, 2002). Modulation of the plasma membrane shape and composition regulate processes outgrowth, axonal development, dendritic branching and the construction of synapses (Jontes and Smith, 2000b). These changes are regulated by interactions between the constituents of the plasma membrane, the exo/endocytic vesicles, and the cytoskeleton (Wood and Martin, 2002). In nonneuronal cells, differential modulation of membrane flow can result in the formation of processes such as microspikes, lamellopodia and filopodia (Wood and Martin, 2002). However, factors that regulate the formation and maintenance of specific classes of membranous processes in neuronal cells remain poorly understood.  Dendritic filopodia, cell surface extensions filled with tight parallel bundles of actin filaments, are thought to be precursors for developing synapses (Dailey and Smith, 1996; Small, 1988). Filopodia are typically 5-20 µm in length and rapidly alter their position and shape. In dendrites, filopodia act as precursors to spines, short bulbous protrusions of the dendrite that form excitatory postsynaptic contacts on many neurons (Jontes and Smith, 2000b). Thus, changes in membrane structure that result in the  1  A version of this chapter has been published. Gauthier-Campbell, C., D.S. Bredt, T.H. Murphy, and ElHusseini Ael. 2004. Regulation of dendritic branching and filopodia formation in hippocampal neurons by specific acylated protein motifs. Mol Biol Cell. 15:2205-17.  46  formation of filopodia are likely important in axonal guidance and spine formation. In contrast, dendritic branching is thought to be mediated via an actin and/or microtubuledependent mechanism (Jan and Jan, 2003). Recent studies showed that in early neuronal development dendrites originate from lamellipodia and short neurites (Jan and Jan, 2003). New branches are then formed through sprouting from an existing branch. Extensive remodeling of the newly formed branches results in the selective stabilization of a limited number of dendritic branches.  Several acylated proteins associated with the cytoskeleton, such as GAP-43 and paralemmin, are implicated in the regulation of membrane dynamics and process outgrowth (El-Husseini and Bredt, 2002; Resh, 1999; Strittmatter et al., 1994a). Studies on the neuronal proteins GAP-43 and paralemmin show that palmitoylation is important for the proper targeting of these proteins to filopodia (Kutzleb et al., 1998; Zuber et al., 1989a; Zuber et al., 1989b). In contrast, dual palmitoylation of PSD-95 is necessary for appropriate postsynaptic localization to dendritic spines (Craven et al., 1999; El-Husseini and Bredt, 2002; El-Husseini et al., 2000a). Overexpression studies showed that the palmitoylated motif of GAP-43 not only targets heterologous proteins to filopodia in neuron-like cells but can also induce filopodia formation (Strittmatter et al., 1994a; Strittmatter et al., 1994b). Moreover, the dually prenylated and palmitoylated motif of paralemmin is essential for its morphogenic activity and concentration at filopodia and microspikes (Kutzleb et al., 1998). Thus, palmitoylation may serve as a signal for delivery of proteins involved in the regulation of cell morphology and membrane dynamics to specific active sites of the plasma membrane.  47  To test the possibility that specific palmitoylation motifs may be an intrinsic determinant of filopodia formation, we studied the effect of overexpression of various palmitoylation motifs on process outgrowth in COS-7 cells and in primary hippocampal neurons. Here, we find that select dually acylated motifs such as the ones present in GAP-43 and paralemmin can induce filopodia in both heterologous cells and neurons. Surprisingly, these palmitoylated motifs also enhance dendritic and axonal branching. In contrast, the palmitoylation motifs of PSD-95 and PSD-93 are incapable of triggering process outgrowth. We also show that a combination of two adjacent cysteine residues and basic amino acids within 2 residues of the palmitoylated cysteines are required for filopodia formation. Furthermore, filopodia induction can be reversed by either blocking on-going protein palmitoylation or by activating specific GTPases that regulate bulk membrane cycling and actin dynamics.  2.2. 2.2.1.  Materials and Methods cDNA Cloning and Mutagenesis Generation of GW1 PSD-95, PSD-95 (C3,5S), PSD-95 (1-26) fused to GFP and  mutations within the palmitoylation motifs of GAP-43, paralemmin and PSD-95 were described previously (Craven et al., 1999; El-Husseini et al., 2000a). The addition of the GAP-43, NSP, and β2A N-terminal palmitoylation motif to PSD-95 were constructed with oligomers encoding the appropriate wild type or mutated motifs and restriction sites, that were annealed and subcloned into GW1 PSD-95-GFP at a Hind III site upstream of the starter methionine and a silent Kpn I site at amino acid 13 of PSD-95. The C-terminal prenyl-palmitoylation motif of paralemmin was added to the C-terminus of PSD-  48  95(C3,5S)-GFP or GFP alone with primers encoding the appropriate wild type or mutated motif and restrictions sites. Paralemmin was obtained by RT-PCR from mouse brain RNA and subcloned into pEGFP (Clontech) at Bgl II and Hind III sites. Wild type and mutant forms of ARF6 were kindly obtained from Dr. Julie G. Donaldson (National Institutes of Health, Maryland). Cdc42 constructs were purchased from the Guthrie Institute, PA.  2.2.2.  COS Cell Culture and Transfection COS-7 cells (COS) were maintained in Dulbecco’s Modified Eagle’s Medium  (DMEM) supplemented with 10% fetal bovine serum, 2mM glutamine and 10mM sodium pyruvate until they reach about 80% confluency. Briefly, cells were removed from a 10 cm culture dish by 0.25% trypsin digestion as previously described (ElHusseini et al., 2002) and plated on 12mm glass coverslips at a density of 50,000 cells per well.  COS cells were transfected 18 to 24hrs after plating using Lipofectamine  (Invitrogen). Briefly, 2 µg of DNA and 2 µl of Lipofectamine 2000 (Invitrogen) were mixed to 200 µl of Optimem (Invitrogen) and let to stand for 5 minutes at room temperature. 100 µl of the Lipofectamine/Optimem mix was added to the cells with 500 µl of DMEM per well of a 24-well culture plate. Cells were incubated for 6 hours at 37oC and the Lipofectamine/Optimem reagent was replaced with fresh DMEM media. Transfection efficiency varied between experiments from 0.1 to 1%.  49  2.2.3.  COS Cell Labeling and Immunoprecipitation For labeling with [3H]palmitate, COS cells  were labeled for 3 h in media  containing either 25 1mCi/ml [3H]palmitic acid as previously described (El-Husseini et al., 2002). For pulse-chase experiments using [3H]palmitate, cells were incubated for variable times in media containing 100 µM palmitate. Labeled cells were washed with ice-cold PBS and resuspended in 0.1 ml lysis buffer containing TEE (50 mM Tris-HCl pH 7.4, 1 mM EDTA, 1 mM EGTA), 150 mM NaCl and 1% SDS. After extracting for 5 min at 4oC, Triton X-100 was added to 1% to neutralize the SDS in a final volume of 0.5 ml and insoluble material was removed by centrifugation at 10,000xg for 10 min. For immunoprecipitation, the samples were then incubated with GFP antibodies (anti-guinea pig, 1:1000 dilution) for 1 hr at 4oC. After addition of 20 µl of Protein A-Sepharose beads (Pharmacia), samples were incubated for 1 h at 4oC. Immunoprecipitates were washed three times with buffer containing TEE, 150 mM NaCl and 1% Triton X-100, boiled in SDS-PAGE sample buffer with 1 mM DTT for 2 min. and analyzed by SDSPAGE. For fluorography, protein samples were separated by SDS-PAGE, and dried under vacuum. Gels were exposed to Kodak X-Omat MR with intensifying screens at 80oC for 5 to 20 days.  2.2.4.  Primary Neuronal Culture and Transfection Neuronal cultures were prepared from hippocampi of E18/E19 rats as described in  Craven et al.(Craven et al., 1999). In brief, hippocampi were dissociated by enzyme digestion with papain followed by brief mechanical trituration. Cells were plated on poly-D-lysine (Sigma) treated glass coverslips (12 mm in diameter) at a density of  50  100,000 to 125,000 cells per well and maintained in Neurobasal media (Gibco) supplemented with B27, penicillin, streptomycin, and L-glutamine as previously described (Brewer et al., 1993). Hippocampal cultures were transfected at 7 days in vitro (DIV) by lipid-mediated gene transfer. At this age, particularly between 8 and 12 DIV, neuronal cells in culture have extended dendrites and are decorated with numerous filopodia. Briefly, two micrograms of DNA and 8 µl of Enhancer (Qiagen) were mixed in 150 µl of EC Buffer (Qiagen) and let to stand for 5 minutes at room temperature. 15 µl of Effectene (Qiagen) were added and incubated at room temperature for 10 minutes, followed by 1 ml of Neurobasal media. 135 µl of the Effectene mix was added to the cells with 200 µl of Neurobasal media per well. Cells were incubated for 3 hours at 37oC and the Effectene reagent was replaced with fresh Neurobasal media. Transfected cells were fixed and immunostained at 10 days in vitro, as described below.  2.2.5.  Immunofluorescence Coverslips were removed from culture wells and fixed in 4% paraformaldehyde at  0 oC for 10-15 min. The cells were washed with phosphate-buffered saline containing 0.1% Triton-X-100 (PBST). COS-7 cells were labeled for F-actin using a rhodamine-or Alexa 488-conjugated phalloidin label (Molecular Probes). Antibodies to GFP (Qbiogene), Cy3 (Jackson Immuno) and Alexa 488 (Molecular Probes) were used. Coverslips were incubated for 1h at room temperature (RT) with primary antibodies, washed in PBST and incubated for 1h at RT with secondary antibodies. Coverslips were then mounted on slides (Frost Plus; Fisher) with Fluoromount-G (Southern  51  Biotechnology Associates), and images were taken under fluorescence microscopy with a 63x objective affixed to a Zeiss Axiovert M200 inverted microscope.  2.2.6.  Quantitative Measurement of Filopodia Induction in COS-7 Cells Images were taken using a 63x objective affixed to a Zeiss Axiovert M200  inverted microscope and AxioVision software. Cells were scored according to the following criteria: COS-7 cells with a minimum of 5 filopodia measuring at least 10µm, or cells with 20 filopodia measuring at least 5µm in length were scored as being “with filopodia”. All other cells were scored as being “without filopodia”. All analysis was conducted on at least 100 cells obtained from at least three individual experiments. Changes were compared using one-way analysis of variance (ANOVA) and filopodia induction is expressed as the average number of COS-7 cells scored as being “with filopodia”. Error bars on graphs are SEM.  2.2.7.  Quantitative Measurement of Dendritic Branching and Filopodia Images were taken using a 63x objective affixed to a Zeiss inverted microscope  and AxioVision software. For dendritic branching, all dendrites within a field of view were counted, including primary and secondary dendrites. In the data presented here, the total number of dendrites was pooled and counted as branches, regardless of primary or secondary classification.  All filopodia within the same field of view were counted manually and the dendritic length was measured using Northern Eclipse (Empix Imaging Inc.). Both  52  dendritic branches and filopodia were counted by an observer blinded to the transfection type. Images were obtained from at least 60 cells from experiments repeated at least three times. Changes were compared using one-way analysis of variance (ANOVA) and filopodia density is expressed as the average number of filopodia per 100 µm. Error bars on graphs are SEM.  2.2.8.  Statistical Analysis All experiments were repeated at least three times. Results are presented as mean  + SEM. Statistical analysis of raw data was performed using Excel. Experimental groups were compared by one-way ANOVA, two-way ANOVA followed by Bonferroni’s post test, Kruskall-Wallis test, or t test as appropriate. A statistical probability of p < 0.05 was considered significant.  2.3.  Results  2.3.1. Differential Filopodia Induction by the Palmitoylation Motifs in Heterologous Cells Previous studies have shown that the dually palmitoylated N-terminus of GAP-43 induces filopodia in heterologous cells (Strittmatter et al., 1994b).  We first asked  whether other dually acylated motifs induce the same effect. For this analysis, we used GFP fusion proteins that contain dually acylated motifs from various neuronal proteins, including the N-terminal palmitoylation motifs of GAP-43, PSD-95 and PSD-93 and the C-terminal motif of paralemmin. Filopodia induction was assessed according to criteria described in Methods. As shown in Figure 2-1, we find that the palmitoylation motif of  53  PSD-95 fused to GFP (PSD1-26) has no effect on the morphology of transfected cells. Filopodia extensions are found in only 11% of PSD1-26-expressing COS-7 cells, comparable to GFP alone (9%). In contrast, 39% of COS-7 cells transfected with the GAP-43 palmitoylation motif (GAP 1-14) show filopodia.  Similarly, 35% of cells  transfected with the dually palmitoylated and prenylated motif of paralemmin fused to GFP (Para CT) displays filopodia. Moreover, mutations of the palmitoylated cysteines in GAP-43 and paralemmin blocked the formation of filopodia mediated by these motifs (Table 2-1). The differential induction of filopodia by specific dually acylated motifs (i.e. GAP-43 and paralemmin, but not PSD-95 and PSD-93) suggests that palmitoylation is necessary but not sufficient to induce filopodia in heterologous cells. These results are surprising because filopodia induction was thought to be a unique property of the dually acylated motif present at the N-terminus of GAP-43 (Strittmatter et al., 1994b). This finding suggests that a larger number of acylated motifs may exhibit this property and that there may be a consensus sequence for filopodia induction.  54  Figure 2-1. Filopodia induction in COS-7 cells. (A) COS-7 cells were transfected with various constructs fused to GFP (green) as described in table 2-1 and immunolabeled for F-actin with a rhodamine-conjugated phalloidin antibody (red). When transfected with palmitoylation motifs of GAP-43 (GAP1-14) or of paralemmin (Para CT), COS-7 cells show extensive filopodial outgrowth, compared to cells transfected with the palmitoyaltion motif of PSD-95-GFP (PSD126). Filopodia induction was quantified by counting the number of cells showing extensive filopodia outgrowth (at least 5 filopodia of ≥10 µm per cell, or at least 20 filopodia of ≥5 µm) and expressed as a percent of cells “with filopodia”. (B) A graph showing that filopodial induction by GAP1-14, Para FL, Para CT and GAP1-14/PSD-95 are statistically different from GFP (*= p<0.05; ** = p<0.01; *** = p<0.01). N > 500. Scale bar, 10µm.  55  Constructs  Palmitoylation Motif  Filopodia Induction  GAP-43 FL Paralemmin FL PSD-95 FL PSD-93 FL  ML MRRTKQVEKN-DMKKHR K CSIM MD L IVTTKKYRYMFFA Y ALRTNVKK-  +++ ++++ ---  GAP-43 NT (GAP1-14) PSD-95 NT (PSD1-26) PSD-93 NT (PSD1-64) β2a (1-11) NSP (416-427) Paralemmin CT (Para CT)  ML MRRTKQVEKNMD L IVTTKKYRYMFFA Y ALRTNVKKMQCCGLVHRRRY MLTCCCLWAFKT-KKYR-DMKKHR K CSIM  ++++ --++ ++ ++++  GAP-43 NT (GAP1-14 C3,4S) Paralemmin FL (Para FL C334S) Paralemmin FL (Para FL C336S) Paralemmin FL (Para FL C334,336S)  MLSSMRRTKQVEKN-DMKKHRSK CSIM -DMKKHR KSCSIM -DMKKHRSKSCSIM  -++ ---  GAP-43 To PSD-95 like: PSD-93 NT/PSD-95 GAP-43 NT/PSD-95 GAP-43 NT (InsL4)/PSD-95 GAP-43 NT (R6,7I)/PSD-95 GAP-43 NT (C3L)/PSD-95 GAP-43 NT (InsL4, R6,7I)/PSD-95  MFFA Y ALRTNVKKML MRRTKQVEKNML L MRRTKQVEKNML MIITKQVEKNMLL MRRTKQVEKNML L MIITKQVEKN-  PSD-95 to GAP-43 like: PSD-95 D2L, Del L4 PSD-95 T8R PSD-95 CCT8R  ML IVTKKYRYMD L IVRTKKYRYML IVRTKKYRY-  -+++++ +++ + +++ -++ + ++++  Table 2-1. Summary of chimeric constructs used in this study. Summary of the chimeric constructs used in this study. All constructs are fused to GFP. GAP-43 FL: full length GAP-43. Paralemmin FL: full length paralemmin. PSD-95 FL: full length PSD-95. PSD-93 FL: full length PSD-93. GAP1-14: residues 1-14 of of GAP-43. PSD1-26: residues 1-26 of PSD-95. PSD1-64: residues 1-64 of PSD-93. β2a (1-11): residues 1-11 of the β2a subunit of L-type calcium channel. NSP (416-427): residues 416 - 427 of the viral protein NSP. Para CT: 13 residues from the C-terminus of paralemmin. GAP1-14 C3,4S: palmitoylation deficient motif of GAP-43. Para FL C334S: Cys 334 Ser mutant full length paralemmin. Para FL C336S: Cys 336 Ser mutant full length paralemmin. Para FL C334,336S: Cys 334,336 Ser mutant full length paralemmin. The following constructs were fused to PSD95 FL: PSD-93 NT/PSD-95: palmitoylation motif of PSD-93; GAP-43 NT/PSD-95: palmitoylation motif of GAP-43 (GAP1-14); GAP-43 NT (InsL4)/PSD-95: GAP1-14 with a Leu inserted between cys3 and cys4; GAP-43 NT (R6,7I)/PSD-95: GAP1-14 with mutations of Arg 6 and Arg 7; GAP-43 NT (C3L)/PSD-95: Cys3Leu mutant GAP1-14; GAP-43 NT (InsL4, R6,7I)/PSD-95: GAP1-14 with an insertion of Leu at position 4 and mutations of Arg 6 and Arg 7 to Ileu. PSD-95 D2L, Del L4: full length PSD-95 with Asp 2 mutated to Leu and deletion of Leu 4. PSD-95 T8R: full length PSD-95 with a mutation of Thr 8 to Arg. PSD-95 CCT8R: full length PSD-95 with a mutation of Asp 2 to Leu, Thr 8 to Arg and deletion of Leu 4. Constructs were scored from “--” to “+++++” according to their relative ability to induce filopodia outgrowth, where “--” means a lack of filopodia inducing properties and “+++++” shows maximal filopodia inducing properties. Palmitoylated cysteines are in bold and basic residues are underlined.  56  2.3.2. Sequence Requirement for Filopodia Induction by Palmitoylation Motifs: Adjacent Cysteines and Nearby Basic Amino Acids To define the consensus sequence for filopodia induction, we examined the amino acid sequences surrounding specific acylated motifs. Previously, we showed that the palmitoylation motifs of GAP-43 and PSD-95 preferentially, but not exclusively, target chimeric proteins to axons and dendrites, respectively (El-Husseini et al., 2001). Systematic mutagenesis of these motifs showed that the spacing of the palmitoylated cysteines and the presence of adjacent basic residues determine the differential targeting by these motifs (El-Husseini et al., 2001).  To test whether these features are also  essential for filopodia induction, we assessed filopodial induction by a panel of chimeric and mutant constructs (Table 2-1. Summary of chimeric constructs used in this study.). We first found that a chimeric protein containing the palmitoylation motif of GAP-43 fused to PSD-95 (GAP-43 / PSD-95) robustly induces filopodia (Table 2-1). This finding suggests that the palmitoylation motif of GAP1-14 can induce filopodia outgrowth to a greater extent when fused to PSD-95. The mechanism by which this enhanced filopodia induction occurs is not known, but may involve changes in protein conformation or increased targeting to areas of active membrane rearrangement. However, addition of a single amino acid between the contiguous cysteines of the GAP-43 palmitoylation motif fused to PSD-95 (GAP-43 NT (InsL4)/PSD-95), which maintains palmitoylation, reduces its ability to induce filopodia (Table 2-1). This suggests that the proximity between the two cysteines is essential but not sufficient to induce filopodia extension. Consistent with this, palmitoylation motifs such as those present in the viral nonstructural protein NSP (Laakkonen et al., 1996) and the β2a subunit of L-type calcium channel (Chien et al., 1996), which contain 2 adjacent cysteines can modestly increase filopodia density (see  57  Table 2-1). To address whether the two basic amino acids one residue away from the palmitoylated cysteines are important for filopodia induction, we replaced those basic amino acids with the hydrophobic residue isoleucine (Ile) (see construct GAP-43 NT (R6,7I)/PSD-95). These changes reduced the ability of the GAP-43 motif to induce filopodia (Table 2-1). Hence, substituting basic residues that are normally positively charged at physiological pH with residues that are not charged likely restricts the association of the palmitoylated motifs with the plasma membrane and effectors present at the plasma membrane. These results indicate that a combination of two adjacent acylated cysteines and two nearby basic residues are essential for the maximum morphogenic effects induced by the GAP-43 motif.  Next, we tested whether minimal sequence changes to PSD-95 that generate a combination of two adjacent acylated cysteines and a nearby basic residue were sufficient to induce filopodia. For this analysis, we used a PSD-95 mutant with no spacing between the cysteines (del L4) and a T8R mutation (CCT8R). Strikingly, this mutant form is also palmitoylated (Figure 2-5) and induced filopodia as observed with the GAP-43 and paralemmin acylated motifs (Table 2-1). Conversely, mutating the basic residues of the GAP-43 palmitoylation motif to hydrophobic ones and insertion of an amino acid between the palmitoylated cycteines (GAP43 NT (InsL4;R6,7I)/PSD-95) blocked its ability to induce filopodia. Moreover, mutating the palmitoylated cysteine adjacent to the prenylated one in paralemmin abolished its effects on filopodia induction (Table 2-1 and data not shown). Thus, the presence of two adjacent cysteines that are either modified by palmitate or by a combination of palmitoyl and prenyl groups can serve the same function on filopodia induction. These results demonstrate that the spacing between the two-  58  acylated cysteines and nearby basic residues are two features critical for creating a filopodia inducing motif (FIM).  2.3.3.  Filopodia Inducing Motifs Promote Filopodia Outgrowth and Enhance  Dendritic and Axonal Branching in Neuronal Cells To characterize further the role of dually acylated motifs in the regulation of filopodia extension, we transfected a Filopodia Inducing Motif (FIM) into primary cultured hippocampal neurons at 7 days in vitro (DIV 7), a stage at which neurons possess a relatively small number of filopodia and few synapses. Over the next few days, from DIV 8 to 12, neuronal dendrites will gradually become covered with numerous filopodia  Similar to what was observed in COS cells we find that FIM transfection  significantly increases the number of dendritic filopodia. Hippocampal neurons showed a 79% increase in filopodia density when transfected with GAP1-14 versus a palmitoylation deficient mutant motif, GAP1-14 (C3,4S) (Figure 2-2). Similarly, GAP114 transfected neurons show a 74% increase in filopodia density when compared to red fluorescent protein (RFP) transfected cells (Figure 2-2).  For this analysis we co-  transfected cells with RFP, which shows a homogenous distribution in neuronal processes and permits visualization of filopodia in neuronal cells.  59  Figure 2-2. Filopodia induction and branching in neuronal cells. (A) Cultured hippocampal neurons transfected at day in vitro 7 (DIV 7) with either the palmitoylation motif of GAP-43 (GAP 1-14), or paralemmin (Para CT) show extensive branching and high filopodial density at DIV 10. In contrast, neurons transfected with RFP alone or RFP together with either the isolated palmitoylation motif of PSD-95 fused to GFP (PSD 1-26) or the palmitoylation-deficient motif of GAP-43 fused to GFP (GAP C3,4S) show less branching and lower filopodial density. Higher magnifications of dendritic branches and filopodia are shown in insets. (B) Graph shows the average number of filopodia per 100µm. (C) Graph shows the extent of branching in tranfected neurons with various constructs (*** = p<0.001). N > 50 cells (n > 500 filopodia) for each transfection. Scale bars, 10µm.  60  Surprisingly, we find that FIMs not only increase the number of dendritic filopodia but also promote dendritic branching. Hippocampal neurons overexpressing the palmitoylation motifs of GAP-43 or paralemmin show a 71% and 76% increase in the number of dendritic branches, respectively (Figure 2-2).  In contrast, neurons  overexpressing GFP, PSD-95 or the palmitoylation deficient motif of GAP1-14 (C3,4S) do not show altered dendritic branching. Similarly, mutating the palmitoylated cysteine at position 336 present at the C-terminus of full length paralemmin abolishes the filopodia formation and dendritic branching induced by paralemmin (Figure 2-3). These changes in the density of filopodia and branches were not restricted only to dendrites. Axons from neurons overexpressing a FIM also showed enhanced filopodia density and axonal branching, as compared to GFP-transfected neurons (Figure 2-4).  61  Figure 2-3. Filopodia formation and dendritic branching in neurons overexpressing paralemmin is palmitoylation dependent. Cultured hippocampal neurons expressing full-length paralemmin (Para FL; top panel) show extensive branching and high filopodial density after 10 days in vitro. In contrast, neurons transfected with a palmitoylation-deficient paralemmin (Para C336S; lower panel) show less branching and a lower filopodial density. Scale bar, 10µm.  62  Figure 2-4. Axonal branching induced by FIMs. Images of axons from neurons co-transfected with the palmitoylation motif of GAP-43 (GAP 1-14) and RFP show extensive axonal branching and filopodia-like protrusions (arrowheads). In contrast, neurons cotransfected with GFP and RFP show significantly less branching and a lower filopodial density. Scale bar, 10µm.  63  2.3.4. Filopodia Induction is Palmitoylation Dependent and is Reversible in Heterologous Cells and Neurons To test whether the palmitoylation of FIMs is dynamic, we conducted a pulsechase experiment as previously described. This allowed us to determine the rate of palmitoylation and de-palmitoylation on various proteins. Breifly, COS cells transfected with various GFP fusion proteins were metabolically labeled using [3H] palmitate. We find that the half-life of palmitate on PSD-95 and FIMs is approximately 2-4 h (Figure 2-5). To further analyze the role of palmitoylation in filopodia extension, we acutely blocked palmitoylation with 2-bromopalmitate. 2-bromopalmitate is a non-metabolizable palmitate analog that blocks palmitate incorporation into proteins (Coleman et al., 1992; Mikic et al., 2006; Webb et al., 2000). The validity of this compound as an inhibitor of protein palmitoylation has been verified by several approaches and for at least two dozen palmitoylated proteins, including Src family kinases, Rho family proteins, G-proteins, PSD95 and transmembrane receptors such as the Nicotinic α7 Receptor and CCR5 (Resh, 2006). Treatment with 20 µM 2-bromopalmitate for 6-8 h immediately after transfection inhibits filopodia induced by FIMs in COS cells (data not shown).  Because  palmitoylation and filopodia formation are reversible processes (Milligan et al., 1995; Mumby, 1997; Ross, 1995), next we asked whether protein palmitoylation is required for the maintenance of filopodia. To test this, we treated transfected COS-7 cells that had already developed filopodial extensions (24 h post-transfection) with 2-bromopalmitate or palmitate as a control. We find that treatment with 2-bromopalmitate caused a significant decrease in the number of cells with filopodia, from 48% to 18% after 8 hours of treatment. In contrast, treatment with palmitate did not significantly change the total number of cells with filopodia (see Figure 2-6).  64  Figure 2-5. Palmitate turnover on FIMs. (A) COS-7 cells transfected with GFP fusion proteins containing the palmitoylation motif of either PSD-95 (PSD 1-26) or FIMs obtained from GAP-43 (GAP-14), paralemmin (Para CT) or a mutant form of PSD-95 (PSD-95:CCT8R). After labeling with [3H]palmitate for three hours, cells were chased with 100 µM cold palmitate for 0, 2 ,4 and 6 hours (h). GFP fusion proteins were purified by immunoprecipitation using antiGFP antibodies. [3H]palmitate incorporation was visualized by autoradiography, and GFP fusion proteins were visualized by western blotting.  65  Figure 2-6. On-going palmitoylation of FIMs regulates filopodia formation in COS-7 cells. (A) COS-7 cells transfected with the GAP-43 palmitoylation motif and treated with 50 µM 2bromopalmitate (24 h post-transfection), which blocks protein palmitoylation, show a significant decrease in filopodia outgrowth. (B) Graph shows the percentage of cells expressing filopodia for the different treatments; untreated and palmitate-treated cells are statistically different from 2-bromopalmitate-treated cells (** = p<0.01). N = 500 cells for each treatment. Scale bar, 10µm.  66  Next, we examined whether FIM-induced filopodia are also reversible in neuronal cells.  When compared to untreated or palmitate-treated cells, hippocampal neurons  treated with 20 µM 2-bromopalmitate 12 hours post-transfection with FIMs showed a 26% decrease in filopodia number (Figure 2-7 A and B). In contrast, a slight reduction in the number of branches was observed upon 2-bromopalmitate treatment (Figure 2-7 C). These data indicate that palmitoylation is also necessary for maintaining the newly formed neuronal filopodia induced by FIMs.  To determine whether palmitoylation is involved in the formation and maintenance of intrinsic dendritic filopodia that are normally formed during early neuronal development, we examined whether treatment with 2-bromopalmitate modulates the total number of filopodia number in GFP expressing neurons. Indeed, our results show a small but significant decrease in the total filopodia density as compared to palmitate-treated or untreated cells (Figure 2-7 D). In contrast, no change in dendritic branching was observed upon acute treatment with 2-bromopalmitate (Figure 2-7 E). These findings indicate that filopodia induction and maintenance in neuronal cells partially involves ongoing protein palmitoylation.  67  Figure 2-7. Filopodia induction is palmitoylation-dependent and partly reversible in neuronal cells. (A) Cells were transfected at DIV 8 the palmitoylation motif of GAP-43 (GAP 1-14) and treated with vehicle, palmitate (Palm) or 2-bromopalmitate (2-Br-Palm) were done 12 hours post-transfection. (B) Graph shows the average number of filopodia per 100 µm for the different treatments (*** = p<0.001). (C) Graph shows the average number of branches for the different treatments (* = p<0.05). (D) The total number of dendritic filopodia is reduced in GFP transfected neurons upon treatment with 2-Br-Palm treatment but not with Palm or Vehicle. (E) No significant change in dendritic branching was observed in GFP transfected neurons upon treatment with 2-Br-Palm. N > 40 cells (n > 600 filopodia) for each treatment. Scale bar, 10µm.  68  2.3.5. The ADP-Ribosylation Factor (ARF6) Regulates Processes Outgrowth Mediated by Palmitoylated Motifs Previous studies showed that plasma membrane recycling and reinsertion at defined plasma membrane sites can play a role in cortical actin rearrangements (Song et al., 1998). A candidate protein for the regulation of this process is ARF6, a non-Rho family GTPase that regulates an endosomal-plasma membrane cycling compartment and influences cortical actin remodeling. In neurons, recent studies showed that dendritic branching can be regulated by an ARF6-dependent pathway (Hernandez-Deviez et al., 2002). Here we tested whether the changes in membrane dynamics induced by the palmitoylated motifs can be regulated by ARF6.  This was first assessed in COS cells by coexpression of a FIM and an HA-tagged ARF6 construct. When compared to cells expressing a FIM alone, we observed a 55% reduction in the number of COS-7 cells with filopodia when co-transfected with the FIM of GAP-43 and wild type ARF6 (Figure 2-8). Co-transfection with a constitutively active form of ARF6 (ARF6-Q67L) resulted in a 67% decrease in the number of cells with filopodia. On the other hand, COS-7 cells co-transfected with a dominant negative form of ARF6 (ARF6-T27N) and FIM show no significant change in the number of cells containing filopodia. These results indicate that activation of ARF6 may act as a negative regulator of filopodia induction mediated by the FIM in heterologous cells. To test whether disruption of the conventional endocytic pathway may also block filopodia induced by FIMs, we assessed the effects of overexpression of a constitutively active form of Rab5 (Q79L).  Rab5 regulates trafficking of endocytosed membranes in a  pathway independent of ARF6. We find that expression of Rab5 (Q79L) had no effect on  69  the filopodia induced by FIM (see Figure 2-8). Thus, cortical actin rearrangements associated with bulk membrane cycling rather than membrane endocytosis, appear to be important for this process.  70  Figure 2-8. ARF6 regulates FIM-induced filopodia extension in COS-7 cells. (A) COS-7 cells co-transfected with the palmitoylation motif of GAP-43 and either wild-type ARF6 (ARF6wt), constitutively active form of ARF6 (ARF6-Q67L), dominant negative ARF6 (ARF6-T72N), or constitutively active Rab5a (Rab5a-Q79L). (B) Filopodia induction is quantified as the percentage of cells co-transfected showing filopodial outgrowth (* = p<0.05; ** = p<0.01). N = 200 cells for each transfection. Scale bar, 10µm.  71  To test whether a similar mechanism may regulate filopodia formation and dendritic branching in neuronal cells, neurons were transfected with FIMs in the presence of HA-tagged wild type ARF6 or a constitutively active form of ARF6 (ARF6-Q67L). We find that co-expression of wild type or ARF6-Q67L with the FIM of GAP43 or paralemmin resulted in a significant decrease in the average number of filopodia and dendritic branches (Figure 2-9). Neurons co-transfected with the GAP-43-FIM motif and with ARF6-Q67L showed a 54% decrease in filopodia number and a 49% decrease in the number of dendritic branches.  Similarly, neuronal cells co-transfected with the  paralemmin-FIM and ARF6-Q67L showed a 55% decrease in filopodial extenstions and 61% decrease in the number of dendritic branches. These results indicate that the small GTPase ARF6 contributes to the regulation of dendritic branching and filopodia extension induced by FIMs in neuronal cells.  72  Figure 2-9. ARF6 regulation of dendritic branching and filopodia extension in neuronal cells. (A) Hippocampal neurons were co-transfected with the palmitoylation motif of GAP-43 (GAP 1-14) or the palmitoylation motif of paralemmin (Para CT) and either with wild-type ARF6 (ARF6wt), or with a constitutively active form of ARF6 (ARF6-Q67L). (B) Graph shows the relative filopodia density; data are expressed as a percentage of GAP1-14 or Para CT transfected cells. (C) Graph shows the extent of branching in transfected neurons. (*** = p<0.001). N > 25 cells (n > 500 filopodia) for each transfection. Scale bars, 10µm.  73  2.3.6. Filopodia Induction by FIMs is also Modulated by the GTPase Cdc42 Filopodia extension in heterologous cells is regulated by a variety of proteins associated with the actin cytoskeleton, including members of the Rho family of GTPases (Hall, 1998b). In particular, Cdc42 has been shown to actively participate in filopodia extension.  To assess whether Cdc42 is involved in the palmitoylation-dependent  induction of filopodia outgrowth mediated by FIMs, we co-expressed the GFP-tagged FIMs with either HA-tagged wild type or mutant forms of Cdc42 in COS-7 cells. Overexpression of a constitutively active form of Cdc42 (Cdc42 G12V) resulted in approximately 5 fold increase in the number of cells with filopodia (Figure 2-10). Cotransfection of Cdc42 G12V with the FIM of GAP-43 or paralemmin further increased the number of cells with filopodia, as compared to GFP transfected cells (Figure 2-10 A and B). In contrast, the number of cells with filopodia was significantly decreased upon co-transfection of FIMs with a dominant negative form of Cdc42 (Cdc42 T17N) when compared to those expressing FIM alone. These data demonstrate that filopodia induced by FIMs can be modulated by Cdc42.  Hence blocking the Cdc42 pathway by  overexpression of a dominant negative form of Cdc42 (Cdc42 T17N) prevented filopodia outgrowth mediated by FIMs, suggesting that Cdc42 may act downstream of FIMs.  74  Figure 2-10. Regulation of filopodia extension and dendritic branching by Cdc42. (A) COS-7 cells were transfected with GFP, the palmitoylation motif of GAP-43 (GAP1-14) or the palmitoylation motif of paralemmin (ParaCT) and with either a constitutively active Cdc42 (Cdc42 G12V) or a dominant negative Cdc42 (Cdc42 T17N). Co-expression of GFP, GAP1-14 or ParaCT with Cdc42 G12V increased the number of cells with filopodia, whereas co-expression with Cdc42 T17N blocked filopodia induced by GAP1-14 and ParaCT. (B) Percentage of COS cells expressing filopodia for the different transfections (* = p<0.05). (C) Hippocampal neurons were co-transfected with GAP1-14 or GFP and either with Cdc42 G12V or Cdc42 T17N. (D) Relative filopodia density expressed as a percentage of GFP, GAP1-14 and ParaCT transfected neurons. (E) Graph shows the extent of branching in transfected neurons. (* = p<0.05). N >30 cells (n > 400 filopodia) for each transfection. Scale bars, 10µm.  75  Next, we examined whether a similar mechanism regulates filopodia extension in neuronal cells. When compared to GFP transfected cells, neurons expressing a constitutively active Cdc42 show a 74% increase in filopodia density (Figure 2-10 C and D). However, FIM expressing neurons did not show a further increase in filopodia density when co-transfected with Cdc42 G12V. These results suggest that Cdc42 may be downstream of FIMs and maximally upregulated by overexpression of the constitutively active form of Cdc42 (Cdc42 G12V).  Alternatively, a mechanism of homeostatic  regulation may limit the increase in filopodia density. Remarkably, when co-transfected with a dominant negative Cdc42, FIM expressing neurons show a 39% decrease in filopodia density and a 34% decrease in the number of dendritic branches (Figure 2-10 CE). Similarly, GFP expressing cells show a decrease in dendritic branching when cotransfected with Cdc42 T17N (data not shown). Taken together, these data indicate that Cdc42 participates in the extension and maintenance of FIM-induced neuronal process outgrowth.  2.3.7. Cdc42-Induced Filopodia Involves a Palmitoylation Dependent Pathway To determine whether filopodia induction mediated by Cdc42 requires protein palmitoylation, we treated COS cells transfected with Cdc42 G12V, 12 h posttransfection, with 20 µM 2-bromopalmitate or palmitate. We find that 8 h treatment with 2-bromopalmitate, but not with palmitate or vehicle, eliminated filopodia induced by Cdc42 G12V (Figure 2-11 A and B). These results show that filopodia formation by Cdc42 in heterologus cells involves a palmitoylation dependent pathway. To assess whether Cdc42 mediated filopodia outgrowth also relies on palmitoylation in  76  hippocampal neurons, we treated cells with 20µM 2-bromopalmitate 12 h posttransfection.  Significantly, neurons transfected with Cdc42 G12V showed a 23%  decrease in the filopodia density when treated with 2-bromopalmitate compared to untreated or cells treated with palmitate (Figure 2-11C and D). These results demonstrate that filopodia extension in neuronal cells is in part dependent on palmitoylation. Furthermore, on-going palmitoylation may participate in the regulation of neuronal processes induced by Cdc42.  77  Figure 2-11. Palmitoylation modulates Cdc42-induced filopodia extension and dendritic branching. (A) COS-7 cells were transfected with either a constitutively active or inactive Cdc42 (Cdc42 G12V or Cdc42 T17N, respectively). 12 h post-transfection, cells were treated with either 20 µM palmitate (Palm) or 2-bromopalmitate (2-Br Palm) for 8 h. Results show a significant decrease in the number of cells with filopodia outgrowth as compared to vehicle or palmitate treated cells. (B) Percentage of cells expressing filopodia for the different treatments; untreated and palmitate-treated cells are statistically different from 2bromopalmitate-treated cells (** = p<0.01). N = 500 COS cells. (C) Images of hippocampal neurons illustrate changes in the density of dendritic filopodia in cells upon transfection with Cdc42 G12V and treatment with 20 µM 2-bromopalmitate or vehicle 12 h post-transfection. (D) Data show a significant decrease in filopodia density after treatment with 2-Br Palm. (* = p<0.05). N >30 neuronal cells (n > 400 filopodia) for each transfection. Scale bars, 10µm.  78  2.4. Discussion Recent investigations have revealed several important roles for palmitoylation in protein sorting and targeting to specific cellular compartments. In neuronal cells, palmitoylation serves as a signal for differential sorting of proteins to axons or dendrites and for protein association with specialized lipid microdomains and synaptic membranes (El-Husseini et al., 2001). Here, we define a palmitoylated protein motif sufficient to induce filopodia formation in heterologous cells and to increase the number of filopodia and dendritic branches in neurons. We show that the morphological changes induced by this FIM require on-going protein palmitoylation and can be reversed by specific GTPases that regulate bulk membrane cycling and actin dynamics. Whether the increase in filopodia induced by FIMs actually translates into enhanced spine morphogenesis still remains to be confirmed. However, it would not be surprising to discover a critical role for palmitoylation in spine development and synapse formation.  2.4.1. Minimal Sequence Required for Induction of Process Outgrowth The sequence requirement for FIMs strictly relies on the presence of two adjacent acylated cysteines and nearby basic residues. Palmitoylation motifs such as those present in PSD-95 and PSD-93 lack this consensus sequence and fail to induce filopodia. Importantly, the palmitoylation motif of PSD-95 can be converted to a GAP-43 like motif and vice versa by minimal amino acid changes (Table 2-1).  These motifs remain  palmitoylated with mutations of nearby residues, but deletion of one of the two cysteine residues decreases the level of palmitoylation. Moreover, similar alterations in the FIM present in paralemmin disrupt its ability to induce filopodia. Because of the degeneracy 79  in the sequence requirements, we predict that FIMs are present in several other neuronal proteins involved in the regulation of cytoskeleton dynamics and processes outgrowth.  2.4.2. Possible Mechanisms for Induction of Process Outgrowth by FIMs The morphogenic effects of GAP-43 and paralemmin are highly dependent on protein palmitoylation.  Our analysis shows that palmitate turnover on GAP-43,  paralemmin and PSD-95 are relatively similar and with a half-life of 2-4 hours. Consistent with this, treatment with 2-bromopalmitate, a drug that blocks protein repalmitoylation, significantly disrupts filopodia induction by FIM motifs only 6-8 h after treatment with this drug. Shorter exposures had no effects on the number of FIM-induced filopodia (not shown). Moreover, mutating the palmitoylated cysteines of GAP-43 or paralemmin block their ability to induce filopodia (Aarts et al., 1999; Kutzleb et al., 1998). Thus, the presence of FIMs in proteins such as GAP-43 and paralemmin that regulate process outgrowth strongly suggest functional relevance.  Previous studies  indicate that the first 10 amino acids of GAP-43 are sufficient to induce filopodia and to stimulate Go (Arni et al., 1998; Strittmatter et al., 1990; Sudo et al., 1992). Surprisingly, synthetic di-palmitoylated peptides which include the N-terminal sequences of GAP-43 abolished Go activity (Strittmatter et al., 1994a).  The authors concluded that  palmitoylation controls a cycle of GAP-43 between an acylated, membrane associated reservoir of inactive GAP-43 and a depalmitoylated active form of the protein. However, our data show that depleting the pool of palmitoylated protein by 2-bromopalmitate treatment, abolishes the effects of FIMs indicating that modification by palmitate mediates the morphogenic effects we observe.  80  How do specific acylated motifs reorganize the cytoskeleton and generate process outgrowth? Previous evidence has shown that the GAP-43 and the paralemmin acylation motifs are efficiently sorted to lipid rafts whereas motifs like PSD-95 lacking the FIM consensus sequence are not (El-Husseini et al., 2001). Lipid rafts are specialized plasma membrane microdomains enriched in sphingolipids that aggregate with cholesterol to form packed raft-like domains within the fluid membrane bilayer. Others recently reported that protein acylation confers localization to cholesterol and sphingolipidenriched membranes (McCabe and Berthiaume, 2001). In addition, the presence of nearby basic residues has been proposed to stabilize interactions with the negatively charged phospholipids present at the plasma membrane (Resh, 1999).  Thus,  incorporation into lipid rafts and strong association with the plasma membrane through interactions with positively charged residues may contribute to the regulation of process outgrowth. A possible mechanism for the observed changes in the cell morphology mediated by FIMs is that the incorporation of these lipidated motifs into lipid rafts may create local changes in membrane tension and the extension of filopodia-like structures. Previous studies showed that alteration in the concentration of specific lipids alter membrane dynamics and fluidity.  For example, addition of sphingomyelin or  phosphatidyl ethanolamine analogs, lipids that expand the plasma membrane, increase the rate of cell spreading and lamellipodia extension and cause a decrease in membrane tension (Bershadsky and Futerman, 1994; Furuya et al., 1995; Harel and Futerman, 1993; Schwarz et al., 1995). It is possible that a similar mechanism is involved whereby the increased rate of addition of palmitate to specific plasma membrane microdomains stimulates process outgrowth by a physical alteration of membrane tension.  81  Alternatively, a change in membrane tension and expansion may stimulate the activation of elements critical for recruitment and anchoring of specific proteins associated with filopodia extension at the plasma membrane.  Filopodia outgrowth is also dependent upon proper assembly of signaling proteins that facilitate interaction between receptors and specific downstream signaling components important for the regulation of cytoskeletal dynamics (Meyer and Feldman, 2002).  Alteration of protein complex assembly may have major consequences in  regulating cell morphology. Under normal conditions the formation of filopodia in many cell types is triggered by the binding of the active GTP bound form of Cdc42 along with phosphatidylinositol 4,5 bisphosphate (PIP2) to the Wiskott-Aldrich syndrome protein (WASP) family of proteins and the recruitment of profilin and Arp2/3 (actin related proteins 2 and 3), proteins that regulate nucleation of actin filaments (Miki et al., 1998; Rohatgi et al., 1999; Yarar et al., 1999). The dynamic protrusion and retraction of filopodia is dependent on the regulation of actin binding and capping proteins (Rao et al., 2000). Although one cannot rule out that a direct interaction between FIMs and elements important for regulating cytoskeletal dynamics may trigger changes in cell morphology, the high degeneracy of FIM s makes this possibility less likely.  2.4.3. Regulation of Filopodia Formation and Dendritic Branching by the GTPases ARF6 and Cdc42 ARF6, a non-Rho family GTPase regulates an endosomal plasma membrane recycling pathway and influences cortical actin remodeling independent of Rho, Rac and  82  Cdc42 activity (Radhakrishna and Donaldson, 1997; Song et al., 1998). In neuronal cells, dendritic branching is regulated by an ARF6-dependent pathway (Hernandez-Deviez et al., 2002; Song et al., 1998). These findings are consistent with our results that AFR6 plays a critical role in regulating FIM-stimulated process outgrowth and dendritic branching (Figure 2-9). We find that overexpression of wild type or constitutively active ARF6 (Q67L) reverses the effects of FIMs both in COS-7 and neuronal cells. In contrast, expression of a constitutively active Rab5 (Q79L), a GTPase that regulates early endosome trafficking in a pathway independent from ARF6, did not affect filopodia induced by FIMs. These results indicate that cytoskeletal rearrangements and membrane dynamics associated with bulk membrane cycling rather than classical membrane endocytosis may contribute to the effects induced by FIMs. Our findings also suggest that ARF6 may act as a negative regulator of filopodia induction mediated by FIMs. The ability of ARF6 to reverse the effects of FIMs in various cell types suggests that a common signaling pathway is involved in this process.  Cdc42 regulates actin dynamics and the formation of filopodia in neuronal and non-neuronal cells (Hall, 1998b; Nobes and Hall, 1995a). In this study, we define a novel mechanism by which filopdia induction by Cdc42 also involves protein palmitoylation. Cdc42 has been shown to be prenylated at cys188, but it is unknown whether this protein is also palmitoylated (Zhang and Casey, 1996). Whether or not Cdc42 is palmitoylated, our new findings indicate that palmitoylation plays a role in the regulation of filopodia formation and maintenance induced by Cdc42.  83  Dendritic morphogenesis depends on many factors; for example, several proteins known to regulate microtubule transport or stability have been shown to affect dendritic morphology. MAP2, the microtubule-associated protein is a major component of crossbridges between microtubules and is known to stabilize them (Harada et al., 2002). Furthermore, the Rho-family of GTPases play important roles in controlling dendritic morphology (Cline, 2001; Redmond and Ghosh, 2001). Thus, coordinated changes in the assembly of microtubules and actin filaments are important for dendritic morphogenesis. Here we demonstrate that FIM-induced dendritic branching also involves a Cdc42dependent pathway. Although blocking on-going protein palmitoylation by 2-bromo palmitate dramatically reduced the effects of FIMs and Cdc42 on process outgrowth in COS cells, these effects were less dramatic in neuronal cells. The smaller change in the number dendritic filopodia is possibly due to differences in the turnover rate of palmitate on various acylated proteins involved in the regulation of process outgrowth and remodeling in neurons.  2.4.4. Possible Roles in Synaptogenesis Filopodia extensions are tightly associated with neuronal development and the initiation of synaptic contacts. Moreover, the enhanced dendritic and axonal branching mediated by FIMs increase the total surface area available for synapse formation. FIMs are present in GAP-43 and paralemmin, two proteins implicated in process outgrowth and synapse formation. GAP-43 is involved in signals that regulate cytoskeletal organization in the nerve ending (Benowitz and Routtenberg, 1997) and GAP-43-deficient mice exhibit restricted axonal pathfinding defects (Sretavan and Kruger, 1998; Strittmatter et  84  al., 1995).  Moreover, its expression is dramatically enhanced during neuronal  development and after nerve injury (Karns et al., 1987; Skene and Willard, 1981). On the other hand, paralemmin is enriched both in axons and dendrites as well as at pre- and postsynaptic sites (Kutzleb et al., 1998).  Interestingly, the expression of a specific  isoform of paralemmin is significantly enhanced during a period that correlates with synapse formation (Kutzleb et al., 1998). Also, our data show that overexpression of full length or the C-terminal palmitoylation motif of paralemmin enhanced the number of filopodia and dendritic spines. We propose that in the context of the full protein, FIMs may cooperate with other regulatory domains present in these proteins to coordinate membrane flow and cytoskeleton rearrangements important for synapse formation.  The re-shaping of synaptic contacts is an active process that continues throughout the life of a neuron. Indeed, high levels of expression of proteins containing the FIMs such as GAP-43 and paralemmin persist in the adult nervous system, in regions that have been associated with high plasticity (Kutzleb et al., 1998; Routtenberg, 1987). Recent data show a dramatic decrease in GAP-43 palmitoylation during early neuronal development suggesting that protein acylation may serve as a dynamic regulator of axonal extension and synapse formation (Patterson and Skene, 1999).  Moreover,  blocking protein palmitoylation by cerulenin disrupts the morphogenic effects induced by GAP-43 (DeJesus and Bizzozero, 2002). Consistent with these findings, our results show that inhibition of protein palmitoylation by 2-bromopalmitate, blocks filopodia formation induced by FIMs. Thus, regulated protein palmitoylation may also act as a signal for modulating membrane dynamics and protein interactions associated with structural plasticity of synaptic contacts. The reversible nature of protein palmitoylation makes it a  85  good candidate for regulating the dynamic changes within the cell. Our results show that the increase in the number of filopodia induced by FIMs in young neurons correlates with a significant increase in the number of spine-like structures.  Thus, on-going  palmitoylation may be necessary for maintaining existing filopodia and for their transformation to spines.  Synaptogenesis is a rapid process and appears to rely on the recruitment of synaptic protein complexes to developing dendritic filopodia soon after they are contacted by an axon (Anderson et al., 1995; Craig et al., 1993; Fiala et al., 1998; Prange and Murphy, 2001; Zhang and Benson, 2000). One model for synapse formation predicts that active dendritic filopodia contact axons to induce presynaptic boutons, followed by a period of filopodial maturation into postsynaptic spines (Harris et al., 1992; MaleticSavatic et al., 1999; Rao and Craig, 2000; Ziv and Smith, 1996). Whereas filopodia induction is partly palmitoylation-dependent, it remains to be clarified whether spine formation is also dependent on palmitoylation.  86  2.5. References Aarts, L.H., P. Verkade, J.J. van Dalen, A.J. van Rozen, W.H. Gispen, L.H. Schrama, and P. Schotman. 1999. B-50/GAP-43 potentiates cytoskeletal reorganization in raft domains. Mol Cell Neurosci. 14:85-97. Anderson, S.A., J.D. Classey, F. Conde, J.S. 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Price, E. Hunter, and D. Zacharias. 2006. A live cell, image-based approach to understanding the enzymology and pharmacology of 2bromopalmitate and palmitoylation. Methods Enzymol. 414:150-87. Milligan, G., M. Parenti, and A.I. Magee. 1995. The dynamic role of palmitoylation in signal transduction. Trends Biochem Sci. 20:181-7. Mumby, S.M. 1997. Reversible palmitoylation of signaling proteins. Curr Opin Cell Biol. 9:148-54. Nobes, C.D., and A. Hall. 1995. Rho, rac and cdc42 GTPases: regulators of actin structures, cell adhesion and motility. Biochem Soc Trans. 23:456-9. Patterson, S.I., and J.H. Skene. 1999. A shift in protein S-palmitoylation, with persistence of growth-associated substrates, marks a critical period for synaptic plasticity in developing brain. J Neurobiol. 39:423-37. Prange, O., and T.H. Murphy. 2001. Modular transport of postsynaptic density-95 clusters and association with stable spine precursors during early development of cortical neurons. J Neurosci. 21:9325-33. Radhakrishna, H., and J.G. Donaldson. 1997. ADP-ribosylation factor 6 regulates a novel plasma membrane recycling pathway. J Cell Biol. 139:49-61.  89  Rao, A., E.M. Cha, and A.M. Craig. 2000. Mismatched appositions of presynaptic and postsynaptic components in isolated hippocampal neurons. J Neurosci. 20:8344-53. Rao, A., and A.M. Craig. 2000. Signaling between the actin cytoskeleton and the postsynaptic density of dendritic spines. Hippocampus. 10:527-41. Redmond, L., and A. Ghosh. 2001. The role of Notch and Rho GTPase signaling in the control of dendritic development. Curr Opin Neurobiol. 11:111-7. Resh, M.D. 1999. Fatty acylation of proteins: new insights into membrane targeting of myristoylated and palmitoylated proteins. Biochimica et Biophysica Acta (BBA) Molecular Cell Research. 1451:1-16. Resh, M.D. 2006. Use of analogs and inhibitors to study the functional significance of protein palmitoylation. Methods. 40:191-7. Rohatgi, R., L. Ma, H. Miki, M. Lopez, T. Kirchhausen, T. 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Zhang, W., and D.L. Benson. 2000. Development and molecular organization of dendritic spines and their synapses. Hippocampus. 10:512-26. Ziv, N.E., and S.J. Smith. 1996. Evidence for a role of dendritic filopodia in synaptogenesis and spine formation. Neuron. 17:91-102. Zuber, M.X., D.W. Goodman, L.R. Karns, and M.C. Fishman. 1989a. The neuronal growth-associated protein GAP-43 induces filopodia in non-neuronal cells. Science. 244:1193-5. Zuber, M.X., S.M. Strittmatter, and M.C. Fishman. 1989b. A membrane-targeting signal in the amino terminus of the neuronal protein GAP-43. Nature. 341:345-8.  91  3.  Chapter III: An active role for dendritic filopodia in the formation of stable  axonal-dendritic contacts2 3.1. Introduction Formation of synaptic connections during development and their modification by experience are important steps in the wiring of the brain (Gerrow and El-Husseini, 2006; Waites et al., 2005; Yamagata et al., 2003). To define the sequence of events that underlie synapse formation, it is critical to understand the molecular mechanisms by which initial contacts are formed between pre- and postsynaptic neurons, and how appropriate axonal and dendritic protein components are recruited to initial sites of contact. Recent advances in imaging techniques have revealed some of the events that regulate excitatory synapse formation (Ahmari et al., 2000; Friedman et al., 2000; Sanes and Lichtman, 2001; Washbourne et al., 2002; Ziv and Garner, 2004). These studies revealed that several components of the synaptic vesicle release machinery travel in transport packets and are rapidly recruited to contact sites. However, the factors that signal the recruitment of transport vesicles and regulate this process is largely unknown (Ahmari et al., 2000). Other studies also documented the existence of transport packets and protein complexes of postsynaptic proteins to nascent neuronal contacts (Gerrow et al., 2006; Marrs et al., 2001; Prange and Murphy, 2001; Sans et al., 2003; Washbourne et al., 2002). These studies suggest that recruitment of synaptic elements to nascent sites of contact plays a role in contact stabilization and synapse maturation. 2  A version of this chapter will be submitted for publication. Gauthier-Campbell, C., P. Arstikaitis, A. ElHusseini, T. H. Murphy. An active role for dendritic filopodia in the formation of stable axo-dendritic contacts.  92  Dendritic filopodia represent potential key structural elements that help probe the environment and establish neuronal contacts (Fiala et al., 1998; Jontes and Smith, 2000b). Filopodia are membranous extensions filled with tight parallel bundles of actin filaments that are typically 5-35 µm in length and rapidly alter their position and shape over minutes (Harris, 1999; Matus, 2005; Portera-Cailliau et al., 2003; Ziv and Smith, 1996). Electron microscopy studies (EM) indicate that synapses can be formed at the tip and base of dendritic filopodia (Fiala et al., 1998). Filopodia are also thought to serve as precursors of small protrusions called dendritic spines (Fiala et al., 1998; Harris, 1999; Prange and Murphy, 2001; Zhang and Benson, 2000). These specialized structures are critical for proper excitatory synaptic transmission and are thought to function as biochemical compartments to isolate and amplify incoming signals (Guthrie et al., 1991; Muller and Connor, 1991). Spines are characterized by their mushroom-like morphology with bulbous tips (0.5-2 µm) connected to a dendrite by a thin neck (~0.04-2 µm in length) (Calabrese et al., 2006).  Despite the identification of several proteins that  regulate spine maturation, it remains unclear whether dendritic spines emerge from filopodia at early stages of synapse formation.  Dendritic filopodia have also been  assigned an exploratory role independent of spine formation, especially given that filopodia can exist transiently in aspiny neurons (Linke et al., 1994; Ulfhake and Cullheim, 1988). Other studies hint to a role for filopodia in guiding the growth of dendrites (Portera-Cailliau et al., 2003; Portera Cailliau and Yuste, 2001; Vaughn, 1989). Indeed, recent work by Sdrulla and Linden shows that filopodia may participate in dendrite development in the cerebellum. In this study, bridge filopodia are proposed to  93  play a role in the self-repulsion of growing dendritic branches to occupy the appropriate spatial orientation (Sdrulla and Linden, 2006).  Here we report that a significant proportion of dendritic filopodia are directly apposed to presynaptic sites that are positive for both excitatory and inhibitory presynaptic markers and that the number of synapses apposed to the tips of filopodia is augmented by expression of neuroligin-1 (NLG1). Using time-lapse microscopy, we also reveal that filopodia in contact with presynaptic axons may actively participate in the recruitment of axonal transport packets to these contact sites. Consistent with this idea, synaptogenic proteins such as NLG1 reduce dendritic filopodia motility and promote rapid stabilization and recruitment of presynaptic elements at the axonal contact site. These results reveal that filopodia play an active role in the formation of diverse types of synaptic contacts. Moreover, expression of proteins that control filopodia dynamics and their adhesive properties modulate the number of synapses formed at the tips of dendritic filopodia.  3.2. Materials And Methods 3.2.1. Constructs Used Generation of GFP fusion proteins of the palmitoylated motif of GAP-43, the constitutively active form of Cdc42, neuroligin-1 and DsRed-tagged synaptophysin were described previously (Gauthier-Campbell et al., 2004; Gerrow et al., 2006). Briefly, the palmitoylation motifs of GAP-43 and paralemmin were fused to GFP within the 94  custom GW1 vector (Craven et al., 1999; El-Husseini et al., 2000a). Cdc42 constructs were purchased from the Guthrie Institute (Pennsylvania, USA). Wild-type neuroligin-1 was a kind gift from Dr. Peter Scheiffele (Columbia University, USA), GFP-tagged Shank1B was a kind gift from Dr. Sala (University of Milan, Italy).  3.2.2. Primary Neuronal Culture and Transfection Neuronal cultures were prepared from hippocampi of E18/E19 rats as described earlier (Gerrow et al., 2006). In brief, hippocampi were dissociated by enzyme digestion with papain followed by brief mechanical trituration. Cells were plated on poly-D-lysine (Sigma) treated glass coverslips (25 mm in diameter) and maintained in Neurobasal media (Invitrogen) supplemented with B27, penicillin, streptomycin, and L-glutamine as described in Brewer et al. (Brewer, Torricelli et al. 1993). Hippocampal cultures were transfected using three different methods to determine whether the results were specific to a particular protocol. The three methods were: nucleofection (Amaxa), lipid-mediated gene transfer (Invitrogen), or calcium phosphate transfection (BD Biosciences, CA). Similar results were obtained with each protocol. Representative data is shown using the nucleofection transfection method, which provides the greatest transfection efficiency, as well as moderate levels of protein overexpression. Briefly, the electroporation protocol was carried out on DIV 0, immediately before plating: 2 to 6 million dissociated hippocampal neurons were re-suspended in 100 µl of room temperature electroporation solution (120 mM KCl, 10 mM KH2PO4, 0.15 mM CaCl2, 5mM MgCl2, 25 mM HEPES, 2 mM EGTA, 2 mM ATP, 5 mM GSSG, pH to 7.4) with 2 µg of high quality endotoxinfree DNA and transferred into AMAXA electroporation cuvettes (AMAXA Inc.,  95  Gaithersburg, MD).  Neurons were then transfected by electroporation using the  AMAXA electroporator, as described by AMAXA Inc. (Gaithersburg, MD).  The  electroporation program used was O-13. Cells were immediately re-suspended in DMEM with 10% Calf Serum, plated at a final density of 0.5 million live cells/mL on 25 mm PDL-coated coverslips and allowed to recover for 1 hour before replacement with Neurobasal Media. The final cell density was approximately 1 million neurons per 25mm coverslip (6-well plates).  In the case of transfection experiments with two  constructs, each fluorescently-labeled construct was individually transfected by electroporation. At the time of plating, the cells transfected with SynDsRed were then mixed with transfected cells expressing the various GFP-tagged constructs and plated as described above (for example: 0.5 million SynDsRed transfected cells co-plated with 0.5 million GFP transfected cells).  The cells are co-cultured together until imaging or  staining, from 7 to 14 DIV. See Figure 3-1 below for a description of this technique.  In the case of lipid-mediated gene transfer, neurons were transfected at 7 days in vitro (DIV) and imaged at 8-9 DIV. Briefly, 2 µg of DNA and 2 µl of Lipofectamine 2000 (Invitrogen) were mixed to 200 µl of OptiMEM (Invitrogen) and let to stand for 5 minutes at room temperature. 100 µl of the Lipofectamine mix was added to the cells with 1000 µl of Neurobasal media per well of a 6-well culture plate. Cells were incubated for 3 hours at 37oC and the Lipofectamine reagent was replaced with fresh Neurobasal media. Calcium phosphate transfections were done at 7 days in vitro (DIV): briefly, 2 µg of DNA and 6.2 µl of calcium phosphate buffer (4M, BD Biosciences) were mixed with 92 µl of HBSS (Hanks Balanced Salt Solution, pH 7.0) and let stand for 5 minutes at  96  room temperature. This DNA solution was added dropwise to 100 µl of distilled water and the mix was added to the cells with 1000 µl of Neurobasal media per well. Cells were incubated for 10 minutes at 37oC and the calcium phosphate reagent was replaced with fresh Neurobasal media.  Figure 3-1. Transfection method using two fluorescently-labeled constructs. Dissociated hippocampal neurons are separated and independently transfected by electroporation with either SynDsRed or with a GFP-tagged construct. The two batches of electroporated cells are then pooled and plated together to make a mixed neuronal culture, with neurons expressing SynDsRed together with neurons expressing a GFP-tagged construct.  97  3.2.3. Immunofluorescence: Coverslips were removed and fixed in 4% paraformaldehyde for 10 min. The cells were washed with phosphate-buffered saline containing 0.1% Triton-X-100 (PBST). Antibodies to GFP (Qbiogene), synaptophysin (PharMingen), GluR1 (Upstate), Cy3 (Jackson Immuno) and Alexa 488 (Molecular Probes) were used. Coverslips were incubated for 1h at room temperature (RT) with primary antibodies, washed in PBST and incubated for 1h at RT with secondary antibodies. Coverslips were then mounted on slides (Fisher) with Fluoromount-G (Southern Biotechnology Associates).  3.2.4. Live Imaging: Imaging was performed at 8-12 DIV in an environmentally controlled chamber (37° C and 5% CO2) with an automatically heated stage and objective heater. Images were taken using a 63x 1.4 NA oil immersion objective affixed to a Zeiss Axiovert M200 microscope with a monochrome 14-bit Zeiss Axiocam HR charged-coupled device (CCD) camera.  Filter sets from Chroma include: GFP (HQ470/40x, Q494LP,  HQ525/50m) and DsRed (HQ535/50x, Q564LP, HQ610/75m). No visible bleed-through or cross-excitation was detectable with even 5000ms exposures. Exposures were set to 25% of saturation and pixels were binned by a factor of 2 to minimize photo-damage to live cells. Images were taken at 14 bit as per camera specifications, exported at 16 bits and analyzed using Northern Eclipse (Empix Imaging, Mississauga, Canada). For the analysis of filopodia motility and synapse formation, images were taken every 1 minute for 60 minutes. Neurons and filopodia were kept in focus manually during the course of acquisition. In the case of spine formation in long term imaging, pictures of the same  98  neuron were taken at DIV 10, 11 and 12.  Time-lapse movies were subsequently  corrected for alignment and x- or y-drifting by using the ImageJ (NIH) plugin Register ROI (Michael Abramoff, University of Iowa, USA).  3.2.5. Quantitative Measurement of Axonal Contacts, Spine Density, Filopodia Dynamics, and Synapse Number Images were taken using a 63x objective affixed to a Zeiss Axiovert M200 microscope and AxioVision software. To correct for out-of-focus protrusions within the field of view, focal plane (z-) stacks were acquired and maximum intensity projections were used for the analysis of images. All analyses were done blinded to the transfection type. DsRed transfected neurons express high levels of the fluorescent protein within all processes and the cell body. Axons from DsRed transfected cells were distinguished from dendrites based on their unique morphological features, including lack of spines, constant caliber, smaller diameter compared to dendrites and obtuse-angle branching. Similarly, axons from SynDsRed transfected cells were identified based on their morphological features, as well as enhanced fluorescence within axons compared to the cell body or dendrites.  Dendritic filopodia and spines: all protrusions between 1-10 µm in length were counted and expressed per length of dendrite. Protrusions were scored based on their morphology: protrusions with a bulbous head wider than the neck by at least 20% were scored as spines and all other protrusions were scored as filopodia (see Figure 3-5). Spines with a head of at least 1.5 µm in width were scored as mushroom spines.  99  Filopodia density was expressed as the average number of filopodia per 10 µm of dendrite. All statistical analysis was done using the XLSTAT add-in for Microsoft Excel (Addinsoft, NY); comparisons between 2 populations were done using the MannWhitney test, two-tailed, and multiple group comparisons were done using the one-way analysis of variance (ANOVA, with Student-Newman-Keuls post-hoc correction). For analysis of filopodia motility, time-lapse imaging was performed as described above. Movies of individual neurons were analyzed as follows: the absolute change in length of at least 20 filopodia per neuron were measured every 5 min of a 1 h movie. Filopodia were classified as “stable” if they did not move more than 0.5 µm over the entire image acquisition period, regardless of whether they were in contact with a synaptophysinlabeled axon.  3.2.6. Calculation of synapse number Images were taken using a Zeiss Axiovert M200 microscope. To reduce the contribution of dim or out-of-focus synaptic puncta within a field of view, (z-) stacks were acquired over several 0.30-0.35 µm z-sections and maximum intensity projections were used for analysis of images. Exposures were set to 75% of saturation of the brightest fluorescent clusters to ensure the greatest dynamic range without over- or under-sampling. Images were exported at 16 bits and analyzed in Northern Eclipse (Empix Imaging) by using custom-designed macros. Firstly, the outline of each neuron, as determined by the expression of GFP constructs, was carefully drawn and saved as a cell outline. Individual grayscale images of the synaptic staining in the red channel (either synaptophysin or GluR1) were processed at a constant threshold level to create a  100  binary “mask” image. This binary mask image contains only pixels that are assigned either an absolute black value (binary number 0) or an absolute white value (binary number 1). This binary mask image was then multiplied by the original image. The resulting image contained a discrete number of clusters with pixel values of the original image. Only clusters which are contained within the original cell outline and that have an average pixel intensity at least 1.5 times greater than background pixel intensity were used for analysis. The average background pixel intensity was measured by averaging 5 separate dendritic areas that were manually selected and did not contain any clusters. This analysis allowed for the automatic and unbiased measurement of synaptic puncta size, brightness and density. The density of GluR1 and synaptophysin puncta is expressed per area of dendrite (µm2) and normalized to GFP-expressing neurons.  3.3. Results 3.3.1. Exploratory Role for Dendritic Filopodia in Contact Initiation Dendritic filopodia have been implicated in neuronal contact formation and spine development (Dailey and Smith, 1996; Portera Cailliau and Yuste, 2001). Here, we provide new evidence that filopodia are important not only for probing the environment, but also in the formation of initial contacts necessary for establishing synapses and spines. Using a novel double transfection system, we show in real time the formation of contacts between presynaptic DsRed tagged axons and postsynaptic dendrites expressing GFP (or any one of the GFP-tagged constructs described previously).  Time lapse  imaging of GFP transfected cells revealed that dendritic filopodia continually interact with axons and can establish new stable contacts with a presynaptic partner (Figure 101  3-22A). We find that 27.9% ± 3.9% contacts are transient, whereas 21.4% ± 4.7% of the contacts formed remain stable for periods of more than 1 h (Figure 3-22B). Further, a small portion of filopodia can initiate new contacts that remain stable for the remainder of the imaging period (3.3% ± 0.9%; Figure 3-2B). These results reveal that filopodia are important not only for probing the environment, but also for establishing the initial contacts between neuronal cells.  Figure 3-2. Role for dendritic filopodia in exploration and contact initiation. (A) Time-lapse recording showing GFP expressing filopodia continuously exploring the environment and developing transient and stable contacts with DsRed-labeled axons (arrows). Panels show three filopodia contacting an axon, two of which form stable contacts within the imaging period (arrowheads) and one unstable contact (arrow). (B) Quantification of the different events observed during the imaging period ± SEM. N = 10 cells with n = 507 filopodia. Scale bar, 5µm.  102  3.3.2. Regulation of Filopodia Induction in Developing Neurons Filopodia extension is thought to be regulated by a variety of proteins associated with the actin cytoskeleton, including members of the Rho family of GTPases (Hall and Nobes, 2000). In particular, Cdc42 has been shown to actively participate in filopodia extension, dendrite morphogenesis and spine development (Gauthier-Campbell et al., 2004; Irie and Yamaguchi, 2002; Scott et al., 2003; Tashiro et al., 2000). Recently, us and others have discovered that the palmitoylated motifs of GAP-43 and paralemmin are sufficient to induce filopodia in neurons (Gauthier-Campbell et al., 2004; Huang and ElHusseini, 2005; Strittmatter et al., 1994a). The filopodia-inducing properties of these FIMs (filopodia-inducing motifs) are dependent on palmitoylation and the presence of basic residues near the palmitoylation sites (Gauthier-Campbell et al., 2004).  Here we utilized various fluorescently-tagged proteins (described in Figure 3-3. ) that modulate filopodia formation to assess their contribution to synapse development. We find that hippocampal neurons at days in vitro 8-9 (DIV 8-9) expressing the palmitoylated motif of GAP-43 fused to GFP (GAP1-14) or the constitutively active form of Cdc42 (Cdc42-G12V) show an increase not only in filopodia number at DIV 8 (1.70 ± 0.16 and 1.68 ± 0.17 filopodia/10µm respectively, compared to 1.00 ± 0.11 filopodia/10µm for GFP, Figure 3-4), but also in the average length of their filopodia, as compared to GFP-expressing cells (8.6 ± 0.75 µm and 7.0 ± 0.56 µm, compared to 5.3 ± 0.29 µm for GFP).  103  Figure 3-3. Summary of constructs used. Full length neuroligin-1 was either tagged with HA (hemagglutinin; NLG1-HA) or with GFP (NLG1). All other constructs are fused to GFP. GAP1-14: residues 1-14 of GAP-43. Cdc42-G12V: full length constitutively active Cdc42 with a G12V mutation. Shank1B: full length wild type Shank1B.  104  Figure 3-4. Detection of dendritic filopodia at sites positive for presynaptic markers. (A) Representative images showing differential induction of filopodia in hippocampal neurons transfected with GFP alone or either with GFP tagged forms of the palmitoylated motif of GAP-43 (GAP1-14), the constitutively active form of Cdc42 (Cdc42-G12V) or neuroligin-1 (NLG1) at DIV 0 and analyzed at DIV 8 and DIV 14. (B) Quantification of the number of filopodia induced by these proteins. (C) Representative images of neurons transfected at DIV 5 with GFP or HA-tagged NLG1 (NLG1-HA), fixed and stained at DIV 8 for the excitatory presynaptic marker VGLUT (vesicular glutamate transporter 1) and the inhibitory presynaptic VGAT (vesicular GABA transporter). (D) Graph shows the percentage of filopodia in contact with endogenous synaptophysin clusters (Endo-Syn). (E) Graph shows the percentage of filopodia in contact with presynaptic clusters positive for VGLUT and VGAT. N >500 protrusions from at least 13 cells for each group were counted. *=p<0.05, **=p<0.01, *** = p<0.001, one way ANOVA. Scale bars, 10µm.  105  Since the adhesive properties of neuronal processes may influence filopodia formation and dynamics, we assessed whether expression of the adhesion molecule neuroligin-1 (NLG1), a potent inducer of synapses, influences filopodia development (Chih et al., 2005; Gerrow et al., 2006; Levinson et al., 2005; Levinson and El-Husseini, 2005; Prange et al., 2004; Scheiffele et al., 2000). Our analysis shows that expression of NLG1 significantly increases filopodia number in DIV 8 neurons (1.36 ± 0.13 filopodia/10µm compared to 1.00 ± 0.11 filopodia/10µm for GFP, Figure 3-4 B; see Figure 3-5 for a representative example of filopodia versus spine morphology). Further, NLG1 expression causes an increase in the fraction of filopodia in contacts with synaptophysin-positive clusters (26.5% ± 1.30% compared to 11.7% ± 0.9% for GFP, Figure 3-4 D), indicating that they represent emerging synapses, or ‘proto-synapses’. To further characterize synapses apposed to filopodia, we immunolabeled GFP and NLG1 transfected cells with either the excitatory presynaptic marker VGLUT (vesicular glutamate transporter-1) or the inhibitory presynaptic marker VGAT (vesicular GABA transporter).  Remarkably, we find that a fraction of VGLUT and VGAT positive  synapses are formed directly apposed to the tips of filopodia.  Moreover, NLG1  expression enhances the proportion of filopodia in contact with VGLUT and VGAT presynaptic markers when compared to GFP expressing cells (29.3% ± 2.8% and 19.6% ± 2.6%, respectively compared to 7.7% ± 2.9% and 5.2% ± 2.2% for GFP; Figure 3-4 C and E). Taken together, these data reveal that filopodia participate in synapse formation and that proteins that regulate cell-cell adhesion may accelerate the transformation of filopodia to functional synapses. These findings are consistent with a proposed role of  106  dendritic filopodia in excitatory synapse formation (Fiala et al., 1998). Our results also reveal that filopodia play an active role in the development of inhibitory contacts.  Figure 3-5. Filopodia and spine morphology. Hippocampal neurons were transfected at DIV 7 with GFP, fixed and stained at DIV 8 for endogenous synaptophysin (Endo-Syn). Representative images summarize the various protrusions observed. These include filopodia, stubby spines and mushroom spines (*mushroom spines at DIV8 look immature compared to mushroom spines at DIV 14 or older, see Figure 3-109).  3.3.3. Modulation of Filopodia Motility and Contact Stability by FIMs, Cdc42 and NLG1 Filopodia are dynamic structures that usually protrude and elongate up to 10 µm in less than a minute and can have lifetimes of less than 10 min (Portera-Cailliau et al., 2003; Ziv and Smith, 1996). Thus, filopodia are thought to serve in exploring the neuron’s environment and probing for appropriate contacts. To characterize the rate of  107  filopodia turnover, time-lapse microscopy was performed on DIV 8-9 hippocampal neurons expressing the various fluorescently-tagged proteins. This was done using an environmentally controlled chamber (37°C and 5% CO2), with images taken every 1-5 min for periods of up to 3 h. Filopodial protrusive motility was calculated by measuring the change in filopodia length over time. The motility of filopodia in neurons expressing Cdc42-G12V or GAP1-14 was greater than cells expressing GFP (0.41 ± 0.03 µm/min and 0.35 ± 0.04 µm/min respectively, compared to 0.24 ± 0.02 µm/min for GFP; Figure 3-6 A and B). In contrast, cells expressing NLG1 show reduced filopodia motility (0.19 ± 0.01 µm/min compared to 0.24 ±0.02 µm/min for GFP; Figure 3-6 A and B).  Figure 3-6. Filopodia motility modulates contact formation. (A) Time-lapse recording showing rapid changes in filopodia dynamics. Filopodia are frequently observed rapidly extend, retract and branch. Representative images of dendrites of neurons (DIV 8) showing relative filopodia motility over a 15 to 30 min time period. Filopodia of neurons expressing the palmitoylated motif of GAP-43 (GAP1-14) or the constitutively active Cdc42 (Cdc42-G12V) exhibit increased motility when compared to filopodia from GFP expressing cells. In contrast, filopodia in GFP-tagged neuroligin-1 expressing cells (NLG1) are more stable. Arrows and arrowheads point to motile filopodia, and asterisks (*) indicate filopodia that are stable over 30 minutes. (B) Graph shows the average filopodia motility expressed as the change in filopodia length over time ± SEM. The motility of at least 60 filopodia for each group was measured over 1 h, from at least 3 independent experiments; *=p=0.011, ***=p<0.001, one way ANOVA. Scale bars, 10µm.  108  To further explore the relation between filopodia motility and contact formation, we analyzed the behavior of filopodia in GFP expressing cells in contact with DsRedlabeled axons. Contacts between filopodia and axons established and subsequently lost within that period (1h) were classified as ‘lost contacts’. Time lapse imaging of GFP transfected cells revealed that dendritic filopodia continually interact with axons, potentially, to establish a contact with a presynaptic partner (Figure 3-7 A and Figure 3-9 A). 21.4% ± 4.7% of the contacts formed remain stable for periods of more than 1 h (Figure 3-7 C), however 27.9% ± 3.9% contacts are transient (Figure 3-67 A and B). Neurons expressing GAP1-14 or Cdc42-G12V show more transient filopodia-axon contacts over an hour period, as compared to GFP expressing cells (40.3% ± 6.5% and 62.3% ± 4.4%, respectively, versus 27.9% ± 3.9% for GFP). Moreover, neurons expressing GAP1-14 or Cdc42-G12V show a greater percentage of contacts lost with DsRed-labeled axons (14.7% ± 4.3% and 25.1% ± 5.2%, respectively) when compared to GFP-expressing cells (5.0% ± 1.9%; Figure 3-7 B). In contrast, neurons expressing NLG1 show a significant decrease in transient contacts (18.3% ± 3.5%). The percentage of filopodia that remain stable over a period of 1 h is much greater in cells overexpressing NLG1 (31.3% ± 4.4%) compared to GFP (21.4% ± 4.7%; Figure 3-7 A). NLG1 expressing cells also show a smaller number of lost contacts than GFP transfected cells (1.8% ± 0.9%; Figure 3-7 B). These findings demonstrate that modification of the adhesive properties of filopodia (by NLG1 overexpression for example), influences their motility and the probability of establishing contacts with presynaptic axons. This is in agreement with the finding that NLG1-expressing cells have a greater percentage of filopodia that can form synaptic contacts or ‘proto-synapses’ (Figure 3-4 C to E).  109  Figure 3-7. Filopodia stability and contact formation. (A) Hippocampal neurons were transfected at DIV 0 and imaged at DIV 8 or 9. Examples of dendritic filopodia from GFP, GAP1-14, Cdc42-G12V and NLG1 transfected neurons establishing contacts with DsRed–labeled axons (DsRed). (B) Quantitative analysis reveals that more contacts are lost in neurons expressing GAP1-14 and Cdc42-G12V, as compared to GFP cells. *=p=0.039 for GAP1-14 and *=p=0.029 for NLG1, **=p<0.01, one way ANOVA. (C) Filopodia contacts with axons expressing DsRed (DsRed) that remain stable for the entire imaging period (1 h in this experiment) were classified as “stable”. Filopodia from neurons expressing GAP1-14 or Cdc42-G12V are transient and less frequently form stable contacts with axons. In contrast, filopodia from NLG1 expressing cells more frequently form stable contacts. Graphs are expressed as a percentage of all filopodia contacts with DsRed-labeled axons ± SEM. *=p=0.023 for GAP1-14, *=p=0.016 for Cdc42-G12V and *=p=0.036 for NLG1, one way ANOVA. N >400 filopodia from at least 8-15 cells for each group. Scale bars, 10µm.  It is worth mentioning that the analysis was performed on contacts between filopodia and axons en passant. In rare occasions we also observed association of axonal growth cones with dendritic filopodia, however because only few events were observed,  110  the significance of this association could not be assessed (Figure 3-8). These data are consistent with a model in which both motile filopodia and process stabilization (following axonal contact) are necessary to induce structures that are capable of maturing into synapses.  Figure 3-8. Axonal growth cone initiates contact formation. A DsRed- labeled axonal growth cone initiates a contact with a small dendritic filopodia. Also note that at the site of contact with the axonal growth cone and dendrite a new dendritic filopodium emerges. Images were acquired every 1 min for a period of 1 h.  3.3.4. Recruitment of Synaptophysin at Contact Sites is Modulated by FIMs, Cdc42 and NLG1 Previous studies have shown that clusters of postsynaptic proteins enhance the recruitment of synaptophysin positive transport packets to contract sites (Gerrow et al., 2006). Here, we examined whether dendritic filopodia associated with synaptophysinDsRed labeled axons, help recruit presynaptic elements to contact sites. Neurons transfected with synaptophysin-DsRed show high levels of protein over-expression, which leads to its diffuse pattern of distribution in the axon, as compared to endogenous synaptophysin expression which is more clustered. Our analysis reveals that 28.0% ±  111  3.6% of stable filopodia from GFP-expressing cells were found associated with synaptophysin-DsRed positive clusters whereas 61.4% ± 7.9% of filopodia in NLG1 expressing cells were associated with synaptophysin-DsRed clusters within the imaging period (Figure 3-9 F). These data are higher than the immunostaining analysis which shows that about 11.7% ± 0.9% of total filopodia are associated with synaptophysin positive puncta (Figure 3-4 D). This suggests that a portion of filopodia associated with presynaptic clusters may subsequently be eliminated. Remarkably, 11.9% ± 1.6% of GFP-positive filopodia stably associated with axons, but lacking presynaptic protein clusters apposed to them, were found to recruit synaptophysin on the presynaptic site of contact within 1h (Figure 3-9 G). In contrast, the fraction of filopodia that recruit synaptophysin-DsRed at sites of contact were significantly altered in GAP1-14, Cdc42G12V and NLG1 expressing cells (6.5% ± 1.7%, 3.3% ± 1.7%, and 20.4% ± 4.5% respectively; Figure 3-9 A to E and G). These findings provide further evidence that enhanced contact stability by proteins such as NLG1 potentiate the recruitment of presynaptic elements to sites of contact between dendritic filopodia and axons.  112  Figure 3-9. Accumulation of presynaptic proteins at sites of filopodia contacts. Hippocampal neurons were transfected at DIV 0 and imaged at DIV 8-9. (A) Examples of dendritic filopodia from GFP and NLG1 transfected neurons contacting synaptophysin-DsRed-labeled axons (Syn). Arrowheads show dendritic filopodia in contact with presynaptic clusters of SynDsRed and arrows indicate filopodia contact with lack of presynaptic accumulation. (B) Quantitative analysis of the percentage of filopodia in contact with synaptophysin-DsRed positive clusters. *=p=0.030 for GAP1-14 and *=p=0.040 for NLG1, **=p<0.01. (C) Example showing the recruitment of a cluster positive for synaptophysin (arrowhead) at the site of contact. (D) Quantification of change in fluorescence intensity of synaptophysin in the boxed area in (C) shows recruitment of distinct clusters of synaptophysin at the site of contact. (E) Quantification of change in fluorescence intensity of synaptophysin within the boxed area in (C) shows progressive accumulation of synaptophysin at the site of contact. (F) Percentage of filopodia contacting synaptophysin-DsRed labeled axons that show enhanced accumulation of synaptophysin at sites of contact over the 1 hour imaging period. The graphs are expressed as a percentage of all filopodia contacts with synaptophysin-labeled axons ± SEM. *=p=0.04, **=p<0.01, ***=p<0.001, one-way ANOVA. N >120 filopodia from 5-8 cells for each group. Scale bars, 5 and 10µm.  113  3.3.5. Involvement of Dendritic Filopodia in Spine Formation If filopodia serve as precursors of spines, then an increase in filopodia number may lead to an increase in spine density in more mature neurons. To address this issue we examined the effects of various filopodia-inducing molecules on the development of spines in DIV 14 hippocampal neurons. At this age, neuronal cells show a significant increase in spine formation and an associated decrease in filopodia density. Despite the significant increase in filopodia density at 10 DIV, expression of GAP1-14 did not alter spine size or number at 14 DIV (Figure 3-10 A and B). In contrast, neurons expressing Cdc42-G12V showed a significant decrease in the density of mushroom-shaped spines suggesting that expression of constitutively active form of Cdc42 delays the maturation of filopodia to spines (0.090 ± 0.015 mushroom spines/10µm versus 0.22 ± 0.02 for GFP, p<0.001). Expression of NLG1 resulted in an increase in the density of mature spines (1.57 ± 0.14 spines/10µm compared to 0.93 ± 0.04 for GFP, stubby and mushroom spines combined; Figure 3-10 A and B). Taken together, these results reveal that enhanced filopodia density is not the rate-limiting step for controlling the number of spines. Rather, factors that modulate actin dynamics and the adhesive properties of filopodia may be critical for filopodia transformation to spines.  114  Figure 3-10. Changes in spine density and synapse number in neurons at DIV 14. Hippocampal neurons were transfected at DIV 7 with GFP, or GFP tagged forms of GAP114, Cdc42-G12V, NLG1 or Shank1B, fixed and stained at DIV 14 for GluR1 or synaptophysin. (A) Examples of dendrites from neurons transfected with the various constructs described above and then stained for GluR1. Overlay images are shown in the right panels. (B) Graph B shows the relative distribution of stubby and mushroom spines upon expression of the various proteins. (C) Quantification of changes in synaptic staining intensity of GluR1 and synaptophysin upon expression of these proteins (at least 10 neurons were analyzed for each protein; histograms show mean ± SEM, normalized to the GluR1 staining density of GFP transfected neurons. *=p<0.05, **=p<0.01, *** = p<0.001, one way ANOVA). Scale bar, 10µm.  115  116  To more directly explore whether filopodia serve as precursors to spines, neurons were imaged over 3 days (DIV 10, 11 and 12; 24 h time points) followed by retrospective labeling for GluR1 to identify mature spines (Figure 3-11 A). During this period, a large number of filopodia formed and disappeared per day: 46.3% ± 7.8% of existing filopodia on DIV 10 had disappeared by DIV 11 and 33% ± 6.5% of filopodia imaged on DIV 11 were new filopodia that were not present at DIV 10. Similar results were obtained between DIV 11 and 12. It is likely that these percentages are an underestimate since only a single time point was used to preserve the health of the neurons. In fact, a large proportion of dynamic filopodia will not be captured using this method. However, it does give us some evidence for the existences of ‘hot-spots’ where filopodia can be seen and those same ‘hot-spots’ later become the sites of dendritic spines.  In addition to filopodia, we also looked at spine density on these cells. During these 3 days of imaging, spine density was increased by 10.2% ± 3.1% per day. Imaging analysis of GFP transfected cells (n=6) revealed that 18 new spines formed during the imaging period. 5 of those spines appeared at sites where filopodia were present 24 h earlier, suggesting that ~30% (29.2% ± 2.9%) of new spines appear at sites that contained filopodia at least 24 h earlier (Figure 3-11 B). On the other hand, the fate of filopodia is more uncertain: out of 306 filopodia analyzed (67 of those remain visible for 3 days; Figure 3-11 B, only 1.6% ± 0.3% of filopodia visible at a given time point transformed into a spine within 24 h. These results suggest that a small fraction of existing filopodia may transform into spines. Alternatively, it is possible that ‘hot-spots’ exist on the dendrite, from which filopodia are likely to emerge and these ‘hot-spots’ promote the  117  growth of a dendritic spine. It is also possible that ‘hot-spots’ exist on neighbouring axons, whereby the release of guidance molecules or neurotransmitters promotes the growth of protrusions at these ‘hot-spots’. Recent evidence supports the presence of ‘hot spots’ along dendrites and/or axons that signal the stabilization and docking of transport packets (Bamji, 2005; Gerrow et al., 2006; Sabo et al., 2006).  Once again, it is important to note that these results are only correlative and based on analysis of time points 24 h apart; given that the imaging interval is very long, it is almost certain that many protrusion outgrowth and retraction events are not captured, and hence we can only speculate on the origin and/or fate of these protrusions. Further, one cannot preclude the possibility that the majority of dendritic spines emerge from transient filopodia that were not visible during the imaging period or directly emerge from the dendritic shaft. In contrast, it is possible that filopodia present at the site of a newly formed spine 24 hours prior may retract and disappear, and that the spine actually emerges de novo. To address these issues, we conducted a series of experiments whereby we were able to image in real-time the transformation of a filopodia into a dendritic spine (Figure 3-11 C).  118  Figure 3-11. Visualization of filopodia transformation into spiny synapses. (A) Hippocampal neurons were transfected with GFP or GAP1-14 at DIV 0. Images of the same neuron were taken every 24 h, at 10, 11 and 12 DIV post-transfection. Cells were then fixed and retrospectively immunolabeled for GluR1. (B) Graph showing the percentage of spines that emerge at sites where filopodia were present 24 h earlier. (C) Graph shows the percentage of spines that emerge de novo, at sites where no filopodia were present 24 h earlier (n >300 filopodia for each group, from at least 4 cells). (D) Quantitative analysis of changes in spine density over a period of 24 h (n >180 protrusions from at least 8 cells). (E) Hippocampal neurons were transfected with red fluorescent protein (RFP) and GFP or Shank1B at DIV 7 and imaged at DIV 8. An example of a filopodium from a Shank1B expressing neuron that rapidly transforms into a spine-like structure, which remains stable for the duration of the imaging period. (F) Graph shows the percentage of spines that directly transform from filopodia during the 3 h imaging period (N >240 filopodia from at least 8 cells; ** = p<0.01 *** = p<0.001, one way ANOVA). Scale bars, 10µ, 1, and 5µm.  119  To further explore the possibility that spines emerge from rapid transformation of existing filopodia, we employed Shank1B, a potent inducer of spine maturation (Sala et al., 2001). Long-term expression of Shank1B in immature neurons (DIV 0 to DIV 8) causes an increase in the density of spines compared to neurons expressing GFP alone (0.39 ± 0.03 and 0.12 ± 0.02 compared to 0.21 ± 0.02 and 0.063 ± 0.01, for stubby spines and mushroom spines per 10µm, respectively). A rapid increase in spine density was also observed within 2 days of expression of Shank1B in young neurons (DIV 7 to DIV 9; Figure 3-11 D). The increase in spine density was correlated with a 21.8% decrease in filopodia number, suggesting that Shank1B promotes rapid transformation of existing filopodia into spines. To explore this further, we performed live imaging of Shank1Bexpressing cells. We find that 1.9% ± 0.5% of filopodia in GFP and 4.1% ± 0.7% of filopodia in Shank1B expressing cells transform into spines within 3 h (Figure 3-11 E and F). In a few occasions, filopodia transformation into spines was also observed to occur within 30 min (Figure 3-11 E). The enhanced rate of filopodia transformation into spines in neurons expressing Shank1B suggests that during neuronal development, recruitment of scaffolding molecules involved in spine maturation may be required for the rapid transformation of newly emerging filopodia into spines.  Despite the increase in spine density in neurons expressing NLG1 or Shank1B there was no significant increase in the staining intensity of the AMPA receptor subunit GluR1 (Figure 3-10 C). In contrast, expression of the constitutively active regulator of actin dynamics Cdc42-G12V that promotes extensive filopodia elaboration and motility reduced the density of synaptic GluR1 and synaptophysin staining. Hence, disrupting the  120  normal regulation of Cdc42 prevents the proper formation of synaptic contacts and spine development.  3.4. Discussion 3.4.1. Possible Roles for Dendritic Filopodia in Synaptogenic Contact Formation. Dendritic filopodia have been implicated in neuronal contact formation and spine development (Dailey and Smith, 1996; Portera Cailliau and Yuste, 2001).  By  manipulation of expression of several molecules that modulate filopodia induction, we provide new evidence that filopodia are important not only for probing the environment, but also in the establishment of synapses and spines. Another critical finding that emerged from this analysis is that expression of a constitutively active form of Cdc42 increases filopodia motility, but reduces the probability of forming stable contacts with axons and the subsequent recruitment of presynaptic elements. In contrast, expression of the adhesion molecule neuroligin-1 dramatically reduces filopodia motility and enhances the number of stable filopodial contacts recruiting presynaptic elements. The enhanced stability of filopodia in neuroligin-1 expressing cells is most likely due to their enhanced adhesive properties that allow their rapid association with contacting axons. We further show that filopodia undergo rapid turnover and continuous remodeling and that only a small fraction of emerging filopodia transform to spines. This process happens over a period of several days, however, the transformation of filopodia to spines can occur within hours upon expression of Shank1B. These results reveal that filopodia actively participate in synapse formation and spine development. Moreover, filopodia  121  transformation to synaptic contacts is controlled by molecules that regulate filopodia formation, membrane and actin dynamics and their adhesive properties.  3.4.2. Recruitment of Presynaptic Proteins at Sites of Contact by Dendritic Filopodia. Previous studies showed that synaptic contacts can exist at the tips of some dendritic filopodia (Fiala et al., 1998; Saito et al., 1997). In agreement with these findings, we show that dendritic filopodia can recruit synaptophysin-positive transport packets to contact sites, suggesting an active role for filopodia in the early events of synaptogenesis. Another important observation is that a significant proportion of dendritic filopodia are found apposed to axonal terminals positive for excitatory (VGLUT positive) or inhibitory (VGAT positive) presynaptic markers. The ability of filopodia to establish inhibitory contacts was surprising since the majority of inhibitory synapses exist on the dendritic shaft. This suggests that the development of inhibitory synapses also involves contact formation between dendritic filopodia with axons of inhibitory neurons followed by filopodia retraction to form shaft synapses.  3.4.3. Filopodia Motility is Essential in Synaptogenic Contact Formation We further show filopodia motility is critical for establishing stable contacts with presynaptic cells. This process is not only controlled by actin-remodeling proteins, such as Cdc42, but also by palmitoylated protein motifs that induce filopodia formation. Indeed, blocking palmitoylation or mutation of the basic amino acids adjacent to the palmitoylated sites prevents the morphogenic effects of these motifs (Gauthier-Campbell  122  et al., 2004). What remains unclear, is how protein acylation influences filopodia induction and motility. One possibility is that insertion of saturated fatty acids into the phospholipid bilayer may trigger changes in membrane tension, thereby influencing filopodia outgrowth. It is also possible that changes in membrane dynamics influence the activity of GTPases that regulate filopodia formation. However, it is important to point out that despite the increase in number of newly formed filopodia in neurons expressing acylated protein motifs, the increase in filopodia number does not lead to enhanced density of spiny synapses. Most likely, the enhanced filopodial motility and reduced contact stability may have contributed to the lack of changes in the total number of synapses or spines formed. In contrast with these results, expression of the constitutively active form of Cdc42 not only enhances filopodia motility and contact instability but also reduces the number of spines formed. These results reveal that filopodia density per se is not the rate limiting step in determining the number of synapses formed, but rather other factors including regulation of actin dynamics are critical for transformation of emerging filopodia to stable synaptic contacts. Another intriguing finding is that expression of neuroligin-1 restricts filopodia motility and enhances stable contact formation. This is likely due to the adhesive properties of this protein (Irie et al., 1997; Nguyen and Sudhof, 1997; Scheiffele et al., 2000; Song et al., 1999). Thus, changes in membrane structure and its adhesive properties modulate the formation of synapses at the tip of dendritic filopodia.  123  3.4.4. Signals Required for Filopodia Outgrowth and Contact Formation It remains unclear whether specific secreted molecules induce dendritic filopodia sprouting, extension and contact formation with the appropriate presynaptic cell. In line with this possibility, motile filopodia were observed in few occasions apparently chasing transport packets or vesicles in DsRed-expressing axons. It has been proposed that filopodia extend in response to glutamate being released from nearby axons (Cooper and Smith, 1992). Focal application of glutamate leads to an increase in filopodia length (Portera-Cailliau et al., 2003). Moreover, filopodia can emerge from spines in response to brief focal pulse of exogenously applied glutamate (Richards et al., 2005). Other experiments on cultured slices showed that filopodia emerge from dendrites after synaptic activation (Maletic-Savatic et al., 1999). Thus, local bursts of glutamate release may guide filopodia to their targets.  However, others have shown that filopodia  extensions persist even after blocking glutamate receptors and synaptic activity (PorteraCailliau et al., 2003; Wong et al., 2000; Ziv and Smith, 1996). Thus, whether filopodia are required to sense glutamate gradients or other molecules for navigation to nascent presynaptic sites is still unknown. Several studies have demonstrated the release of inhibitory neurotransmitters from developing presynaptic axonal terminals. Indeed, recent work by Li et al. demonstrated the co-release of glutamate and acetylcholine from single neurons in the developing frog tadpole spinal cord (Li et al., 2004). Earlier work provided evidence for the co-release of the inhibitory neurotransmitters GABA and glycine from individual neurons at spinal synapses (Jonas et al., 1998). Other studies have shown that the release of peptides, such as dynorphin and neurotensin that are important for proper synaptic development (Conner-Kerr et al., 1993; Legault et al., 2002). The co-release of  124  glutamate and dopamine gives rise to major CNS dopaminergic connections Sulzer, Joyce, Lin 1998). Hence, there are a number of peptides and neurotransmitters released during development and synaptogenesis.  However, it is still unclear whether these  molecules serve as guidance cues for filopodia.  3.4.5. A Possible Role for Dendritic Filopodia in Spine Formation Because of the temporal order in which they appear during development, filopodia have been proposed to serve as precursors of spines (Ziv and Smith, 1996). This hypothesis however is still controversial because many adult neurons are devoid of spines, even despite bearing numerous filopodia at the time of early synaptogenesis (Dvergsten et al., 1986; Linke et al., 1994; Lund et al., 1977; Mason, 1983; Ulfhake and Cullheim, 1988; Wong et al., 1992). Other studies postulated that synapses can directly form on dendritic shafts, followed by gradual emergence of spines at these sites (Ethell and Pasquale, 2005; Fiala et al., 1998). However, this view does not preclude the possibility that a certain portion of emerging filopodia rapidly transform into spines and form long-lasting synapses. Our data reveal that the majority of filopodia do not give rise to spines (only 3.1% ± 0.3% of filopodia present at DIV 10 are replaced by a spine at DIV 12). Therefore, it is possible that the role of most filopodia is not to uniquely create spines, but rather to regulate the process of selective synaptogenesis.  Hence the  exploratory role of filopodia may help insure the formation of synaptic contacts with the appropriate presynaptic partners, but many dendritic spines may emerge de novo from dendrites, bypassing the need for filopodia. The decision whether filopodia transform  125  into stable contacts and/or spines requires the recruitment of molecules that influence membrane and actin dynamics as well as their adhesive properties.  The findings presented in this study strongly support the hypothesis that dendritic filopodia play an active role in the initiation of synaptic contacts with nearby axonal shafts. Although it is possible that not all spiny synapses are derived from filopodia, we show clear evidence that a certain portion of them are. Further, we show that filopodia dynamics play an important role in the development of appropriate synaptic contacts. Indeed, the protrusive motility of filopodia seems to be a critical aspect of proper contact initiation between a dendrite and an axon. The proper dynamics of filopodia that lead to their stabilization, as well as the critical proteins involved in filopodia motility may serve as factors important for the establishment of connections between neighboring neurons and the maturation of a spine. Recent evidence from Nishiyama et al. supports the view that the motility of axonal branches plays a role in the formation of synaptic contacts. The authors find that highly dynamic transverse branches of cerebellar climbing fibers fail to form conventional synapses, whereas the almost static ascending branches develop functional synapses with Purkinje cells (Nishiyama et al., 2007). Interestingly, our study points to dendritic filopodia as the key players in the establishment of synaptogenic contacts.  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Evidence for a role of dendritic filopodia in synaptogenesis and spine formation. Neuron. 17:91-102.  132  4. CHAPTER IV: General Discussion  On average, each of the one hundred billion neurons in the human brain receives and makes over ten thousand synaptic contacts. Synapses are specialized intercellular junctions whose specificity and plasticity provide structural and functional basis for the formation and maintenance of the complex neural network in the brain. The number, location and type of synapses are well controlled and synaptic circuits are formed in a highly reproducible way.  Although our understanding of the development of synaptic  connections has increased dramatically over the past few years, many key issues have yet to be addressed. For instance, we are only beginning to appreciate the potential role of filopodia in synaptogenesis and spine formation, and we are far from describing all the molecules involved in regulating this process. Whether filopodia are precursors to spines or play a role in synapse formation also remains elusive. In developing neurons, filopodia are gradually replaced by spines as synapse formation progresses (Matus, 2005). These observations have pointed to filopodia as possible candidates for initiating synaptogenic contacts, however, this function is still unclear and controversial (Yuste and Bonhoeffer, 2004). Indeed, it appears that some, but not all, spine synapses result when synapses initially form on filopodia and then transform into spines. However, other synapses have been shown to form on dendritic shafts void of filopodia and promote the growth of a spine from this site of contact (Fiala et al., 1998). The research projects presented in this thesis have focused on the function and regulation of dendritic filopodia and their role in the development of synaptic contacts to contribute to our understanding of developmental processes associated with neuronal maturation.  133  4.1. Dendritic Filopodia 4.1.1. Regulation and Function of Dendritic Filopodia While little is known about the molecular constituents of dendritic filopodia, axonal growth cone filopodia and filopodia from fibroblasts have been relatively wellstudied.  In axonal growth cone filopodia, the core bundle consists of ~20 actin  microfilaments which have their barbed end attached to an electron-dense submembrane compartment at the tip of the filopodium (Lewis and Bridgman, 1992). The growth cone filopodium is regulated by the small GTPase Cdc42, which has been shown to induce de novo filopodium formation through the activation of N-WASP, a member of the WASP family of proteins (Wiskot-Aldrich Syndrome Protein) (Miki et al., 1998). Additional proteins identified at the tip of the growth cone filopodium include β1 integrin; members of the ezrin-radixin-moesin (ERM) family of proteins which link the actin filaments to the membrane; Mena (mammalian enabled), an ena/VASP family member, and phosphotyrosine (Goldberg et al., 2000; Wu and Goldberg, 1993; Wu et al., 1996). All of these components at the tips of axonal growth cone filopodia may affect the rate of actin polymerization and therefore regulate filopodia extension and dynamics. Interestingly, these actin-regulating proteins found in axons have also been identified in dendrites, for instance, Mena has been found at the tip of both dendritic and axonal growth cones, and is thought to regulate actin polymerization at this site by its scaffolding activity for Arp2/3 and other core actin-binding proteins (Goldberg et al., 2000). This suggests that similar mechanisms are involved in regulating the actin cytoskeleton in dendritic filopodia.  134  Recent evidence points to the role of actin-regulating proteins in the extension of dendritic filopodia (Hall, 1998a; Nobes and Hall, 1995a). Consistent with these findings, we show that the small Rho GTPase Cdc42 plays a role in regulating filopodia dynamics and the subsequent formation of synapses. Regulation of dendritic filopodia outgrowth and motility is not exclusively influenced by actin-regulating proteins: proteins that alter the membrane composition also affect filopodia formation and dynamics. Indeed, we demonstrate that specific palmitoylated protein motifs, characterized by two adjacent cysteines and nearby basic residues, are sufficient to induce filopodial extensions in heterologous cells and to increase the number of filopodia and the branching of dendrites and axons in neurons. Such motifs are present at the N-terminus of GAP-43 and the Cterminus of paralemmin, two neuronal proteins implicated in non-cytoskeletal influence of filopodial outgrowth (El-Husseini et al., 2002; Gauthier-Campbell et al., 2004; Resh, 1999; Strittmatter et al., 1994a). This raises the possibility that insertion of a rigid saturated fatty acid such as palmitate into a more fluid phospholipids bilayer could directly trigger these morphological effects. In support of this model, a change in cell morphology has recently been shown to result from changes in membrane tension caused by perturbation of membrane flow dynamics triggered by specific lipids (Raucher and Sheetz, 2000). Filopodia induction is blocked by mutations of the palmitoylated sites or by treatment with 2-bromopalmitate, an agent that inhibits protein palmitoylation (Webb et al., 2000). However, a potential caveat of this experiment is that the effects of 2bromopalmitate treatment are non-specific since it is a potent inhibitor of all palmitoylacyl transferases (PATs) and numerous neuronal proteins are palmitoylated (Webb et al.,  135  2000). Disrupting the palmitoylation of all palmitoylated proteins likely affects their trafficking, subcellular localization and function. Inhibiting the palmitoylation of only a subset of proteins by blocking specific PATs would more directly address the role of protein palmitoylation in filopodia formation. However, individual PATs were only identified recently and there are currently no known inhibitors that have differential specificity for these enzymes (Fukata et al., 2004; Huang et al., 2004).  4.2. Development of Central Nervous System Synapses Much of our understanding of synapse formation is derived from studies on the neuromuscular junction (NMJ). At the NMJ, contacts are formed between presynaptic motorneurons and postsynaptic muscle cells (Goda and Davis, 2003; Sanes and Lichtman, 1999). Due to its size and accessibility, the development and function of this synapse are well characterized.  In contrast to the NMJ, the mechanisms of  synaptogenesis in the CNS are poorly understood, partly because of the heterogeneity of synapses and the differences in timing of their development (Rao et al., 1998). The emerging view is that events of synaptogenesis at the NMJ are presumed to be similar for formation of CNS synapses, but most likely with different molecular players. Despite the incomplete understanding, CNS synapse formation is thought to involve several characteristic steps: contact initiation, recruitment of pre- and postsynaptic proteins, stabilization and maturation of the synaptic contact. These steps will now be discussed in more detail.  136  4.2.1. Filopodia Initiate Synaptogenic Contacts Filopodia have long been thought to play a role in dendritic exploration (Dailey and Smith, 1996; Portera Cailliau and Yuste, 2001). Together with complementary findings from other laboratories, our studies provide evidence that filopodia are not only important for probing the environment, but are also key players in the development of synaptic contacts. Indeed, several electron microscopy (EM) studies have revealed synapses on the tips and shafts of dendritic filopodia, suggesting they are capable of participating in synaptogenesis (Fiala et al., 1998; Saito et al., 1997). Using live cultured hippocampal neurons and fluorescent labeling methods, we demonstrated the ability of filopodia to initiate a contact with nearby axons and this contact formation is affected by filopodia motility. Indeed, expression of molecules like GAP1-14 or cdc42-G12V that enhance filopodia motility, cause a decrease in the number of stable filopodia-axon contacts over an hour period, as compared to GFP expressing cells.  Interestingly,  neurons expressing NLG1 show a significant increase in the percentage of stable contacts suggesting that molecules that contribute to enhancement of filopodia motility may decrease the probability of forming stable contacts. These results support the hypothesis that dendritic filopodia play a role in the initiation of synaptic contacts and that this process can be accelerated by the recruitment of necessary pre- and postsynaptic proteins, including adhesion molecules. As a cautionary note, it is important to realize that these studies are done using (essentially) two-dimensional (2D) imaging, hence filopodia that are extending perpendicularly above and below a given dendrite would most likely not be captured by our imaging system. Additionally, given the resolution limits of our microscope (maximal measured resolution is ~0.5µm in the lateral plane and ~1µm in the  137  z-plane), it is possible that very small filopodia may be overlooked, especially if they contain low levels of fluorescently-labeled proteins. Therefore, the density of filopodia is likely under-represented in these studies.  4.2.2. Recruitment of Protein Complexes at Synaptic Contacts Sites Recent advances in imaging techniques have revealed some of the molecular events underlying formation of excitatory synapses in the CNS (Ahmari and Smith, 2002; Bresler et al., 2004; Friedman et al., 2000; Li and Sheng, 2003; Sanes and Lichtman, 2001; Shapira et al., 2003; Waites et al., 2005; Washbourne et al., 2002; Ziv and Garner, 2004). These elegant studies demonstrated that the major components of the synaptic vesicle release machinery travel in transport packets and are rapidly recruited to contact sites. Other studies revealed the existence of transport packets and protein complexes that regulate delivery of postsynaptic protein complexes to nascent neuronal contacts (Gerrow et al., 2006; Prange and Murphy, 2001; Sans et al., 2003; Waites et al., 2005; Washbourne et al., 2002; Washbourne et al., 2004).  This translocation of protein  complexes has been thought to be triggered by initial contact between dendritic filopodia and presynaptic axons, resulting in the stabilization of filopodia and maturation into spines. These data support a model in which both motile filopodia and contact initiation are necessary to induce structures that are capable of maturing into synapses. Consistent with this hypothesis, we find that a significant proportion of filopodia initiate a contact with nearby axons and recruit key presynaptic molecules (Chapter III). However, factors and molecules that trigger the accumulation of proteins to the site of contact remain  138  largely unknown.  Potential candidates that mediate the recruitment of pre- and  postsynaptic proteins during the initial contact between neurons are cell adhesion molecules (CAMs) (Bamji, 2005). Recent studies have shown that several CAMs can modulate synapse number, maturation and function.  Specifically, members of the  cadherin family have been shown to accumulate at sites of synaptogenic contact (Jontes et al., 2004; Togashi et al., 2002). Blocking N-cadherin results in a loss of dendritic spines (Togashi et al., 2002). Protocadherins have also been implicated in synapse formation: deletion of protocadherin-γ results in a loss of synapses (Weiner et al., 2005). Further, overexpression of neuroligins has been demonstrated to increase the size and number of excitatory and inhibitory presynaptic terminals (Levinson et al., 2005; Prange et al., 2004), as well as potentiate the clustering of postsynaptic proteins (Dean et al., 2003; Graf et al., 2004; Nam and Chen, 2005). Consistent with these findings, we show that adhesion molecules like NLG1 can enhance the recruitment of presynaptic elements to the site of contact. These findings strongly support the hypothesis that dendritic filopodia play a role in the development of synaptic contacts and that this process triggers the recruitment of protein complexes essential for the development of excitatory and inhibitory synapse formation.  Another critical aspect of synaptogenesis is specificity, and the mechanisms that determine whether a new contact develops into an excitatory or inhibitory synapse remain unknown. The limited information available on the constituents of inhibitory synapses has restricted much of the research to excitatory synapse development (Walikonis et al., 2000).  While few proteins have been identified at inhibitory synapses, such as γ-  139  aminobutyric acid (GABA) receptors and the scaffolding protein gephyrin, little is known about the mechanisms of inhibitory synapse maturation. Surprisingly, in our studies we find a significant proportion of dendritic filopodia are apposed to presynaptic clusters positive for the inhibitory protein VGAT, suggesting that filopodia may also play a role in the development of inhibitory synapses. The ability of filopodia to establish inhibitory contacts is novel and supports the hypothesis that filopodia are important for contact formation between postsynaptic dendrites and inhibitory axons.  Whether filopodia  retract and shaft synapses are formed at the site of filopodia retraction is unknown and more work is needed to clarify their role in the formation of inhibitory synapses and the molecules directly involved in this process.  4.2.3. Role for Filopodia in Spine Formation Whether filopodia are precursors to spines still remains unclear. Several points of evidence seem to point to this hypothesis: filopodia and spines both protrude from the dendrite, and filopodia expression during development precedes that of spines. Interestingly, dendritic filopodia can be sites for new synapse formation and some data suggest that these filopodia may become stabilized to form spines (Fiala et al., 1998; Ziv and Smith, 1996). Consistent with this idea, using time-lapse imaging we showed that contact between a dynamic dendritic filopodium and axon seemingly caused the stabilization of the filopodium. Although this observation was rare, it reinforces the hypothesis that dendritic filopodia are capable of initiating synaptogenic contacts with presynaptic axons and stabilize to promote the recruitment of key molecules required for synapse formation.  Further, we were able to capture for the first time filopodia 140  transforming into spines (Figure 3-11 E). Indeed, we find a small percentage of filopodia can transform into spines within 3 h, and expression of Shank1B in immature neurons promotes the rapid transformation of existing filopodia into spines. The enhanced rate of filopodia transformation into spines in neurons expressing Shank1B suggests that during neuronal development, recruitment of scaffolding molecules involved in spine maturation may be required for the rapid transformation of newly emerging filopodia into spines. This view is consistent with previous findings from Prange et al., who show that the recruitment of scaffolding proteins such as PSD-95, promotes the stabilization of dendritic filopodia (Prange and Murphy, 2001). The recruitment of PSD-95 and possibly other postsynaptic proteins within filopodia suggests the likely maturation into a spine.  Despite the evidence for the filopodia hypothesis for the origin of dendritic spines, several problems remain unexplained: for example, some neurons show extensive filopodia dynamics during development, even though many of these neurons develop with a distinctive lack of spines (Difiglia et al., 1980; Linke et al., 1994; Lund et al., 1977; Ulfhake and Cullheim, 1988). Indeed, most inhibitory neurons in the CNS which show extensive filopodia density and motility during development, but will mature without subsequent spines, suggesting that the role of filopodia is not uniquely ‘spinogenic’. Consistent with these findings, our data also suggest that filopodia density is not a factor in determining the subsequent development of spiny synapses. Indeed, we find that being able to induce filopodia outgrowth and thereby increasing the density of dendritic filopodia does not necessarily lead to enhanced synapse or spine density in older neurons. Proteins like cdc42-G12V or the palmitoylation motif of GAP43 promote  141  filopodia induction, but we find that long-term expression of these proteins does not lead to enhanced synapse formation. This suggests that the number of filopodia on a given neuron is not the limiting factor in determining the subsequent spine density and proteins that have ‘filopodia-inducing’ capacity do not necessarily have ‘spinogenic’ capacity. Other mechanisms must be involved including, but perhaps not limited to filopodia motility. Taken together, these data suggest that other mechanisms such as homeostatic plasticity may ultimately be more important in regulating the density of spines on a postsynaptic cell and hence the synaptic input a neuron receives (Turrigiano and Nelson, 2000).  Understanding synaptogenesis and spinogenesis is highly significant because a growing number of human disorders have been associated with abnormalities in dendritic spine morphology or density, such as epilepsy, stroke, trauma, schizophrenia, dementia, etc. (Fiala et al., 2002; Glantz and Lewis, 2000; Nimchinsky et al., 2002; Swann et al., 2000). However, the connection between molecular mechanisms and clinical phenotype has yet to be clearly established. Being able to uncover the functional abnormalities in synaptic function will likely shed light on a variety of disorders. Interestingly, some studies have proposed a role for neuroligins in the genetic etiology of autism (Jamain et al., 2003). It has been proposed that alterations in neuronal circuitry and/or neuronal signaling are responsible for the behavioral and cognitive aberrations in autism patients. Mutations in NLG3 and NLG4 leading to intracellular retention of neuroligin proteins have been found in some autistic patients (Chih et al., 2004). These mutations affect celladhesion molecules localized at the synapse, suggesting that defects in synaptogenesis  142  may predispose to autism. However, these findings are still controversial: more recent studies indicate that mutations in the genes encoding neuroligin are not common among all autistic patients (Gauthier et al., 2005; Talebizadeh et al., 2004; Vincent et al., 2004). Hence, the idea that autism is not uniquely due to a mutation of a single gene would not be surprising considering the heterogeneity of autistic disorders.  4.2.4. Alternative Roles for Dendritic Filopodia Despite the evidence for a role for filopodia in synapse and spine formation, several problems remain unanswered. Firstly, given the abundance of filopodia in early stages of development, and the relatively smaller number of spines on a mature neuron, it is likely that filopodia play a role in other processes. Further, evidence from our work and others shows that only a small proportion of filopodia will actually make connections with a neighboring axon and form a synapse (Fiala et al., 1998). Finally, the maturation of a single neuron may involve the formation of hundreds of dendritic branches, each hosting hundreds of filopodia with potentially different roles (Evers et al., 2006). Consequently, multiple functions have been assigned to dendritic filopodia including steering the dendritic growth cone (Portera-Cailliau et al., 2003), guiding the axon growth cone towards the dendrite, serving as precursors to new dendrite elaboration (Niell et al., 2004) and synaptogenesis and spinogenesis (Fiala et al., 1998; Yuste and Bonhoeffer, 2004). Some of these processes are interconnected: for example, new dendrite formation can occur in a ‘synaptotrophic’ fashion, whereby the formation of new synapses at the tips of dendritic filopodia leads to their stabilization and transformation into new dendrites (Konur and Yuste, 2004; Niell et al., 2004). Work within the optic tectum of  143  zebrafish larvae indicates that almost all synapses form on newly extended dendritic filopodia, a fraction of which are maintained and in turn cause the maturation of filopodia into dendritic branches (Niell et al., 2004). These findings support a role for filopodia in the growth and branching of dendritic arbors.  Hence, the evidence points to multiple roles for dendritic filopodia. To date, very few studies have explored the existence of morphologically and functionally distinct types of dendritic filopodia (Evers et al., 2006; Portera-Cailliau et al., 2003). In a recent study by Evers et al., the authors demonstrate the presence of two different types of dendritic filopodia on MN5 motorneurons of the hawkmoth, each undergoing developmentally-regulated morphogenesis (Evers et al., 2006). One type of filopodia located on dendritic shafts seems to participate in the formation of shaft synapses, by either ‘pulling’ presynaptic axon towards the dendrite or by gradual filopodia shortening and growth of the postsynaptic dendrite. The other type of filopodia, located on growing dendritic tips, also accumulates presynaptic terminals, likely reflecting the synaptotrophic growth of the dendrite.  Recent work using two-photon microscopy has explored  alternative roles for dendritic filopodia (Sdrulla and Linden, 2006). This study by Sdrulla et al. demonstrates the presence of a new ‘type’ of filopodia often found between adjacent dendrites in Purkinje cells of the cerebellum: “filopodia bridges”. Although morphologically similar to filopodia from hippocampal neurons, they are likely to be functionally unique. Indeed, these filopodia have been found to make contacts between two neighboring dendrites, suggesting an instructive role for filopodia in guiding and stabilizing the growth of dendritic arbors during early postnatal development (Sdrulla and  144  Linden, 2006). These data provide evidence for the presence of at least three distinct types of filopodia during early postembryonic dendritic growth and synaptogenesis.  4.3.  Future work  4.3.1. Does Activity Regulate Filopodia Outgrowth? Dendritic filopodia in immature neurons are highly dynamic, and this motility is commonly thought to be actin-based. Motility and turnover of these early protrusions decreases throughout development, mirroring an increase in their average lifetime and density. Recently, the motility and length of dendritic filopodia have been shown to be modulated by neuronal activity. Yuste et al. provide evidence that dendritic filopodia density and length is increased by global blockade of synaptic transmission using TTX or calcium-free solutions (Portera-Cailliau et al., 2003). This treatment has also been shown to upregulate glutamatergic synapse formation (Turrigiano et al., 1998).  Moreover,  blocking ionotropic glutamate receptors results in a decrease in the density and turnover of shaft filopodia, whereas focal glutamate application leads to an increase in the length of shaft filopodia (Portera-Cailliau et al., 2003). Interestingly, filopodial motility in mossy fibers of the hippocampus is differentially regulated by kainate receptors: synaptic stimulation of kainate receptors enhances motility in younger slices, but it inhibits it in mature slices (Tashiro et al., 2003). Further, local calcium-induced signals have been shown to play a role in controlling filopodia growth and motility. Indeed, transient increases in calcium concentration induced by synaptic activity caused the stabilization of existing filopodia and prevent the growth of new filopodia (Lohmann et al., 2005). Taken together, these results support a model by which dendritic filopodia can be 145  regulated by neuronal activity.  Whether the initial establishment of axo-dendritic  contacts is dependent on neuronal activity still remains to be determined. Interestingly, our data show that manipulating the motility of filopodia and dendritic spines results in dramatic changes in synaptogenesis (see Chapter III).  4.3.2. Released Molecules Required for Guiding Filopodia It is unclear whether dendritic filopodia play a uniquely exploratory role by randomly extending and retracting in the neuropil, or are guided by released molecules. In comparison to research on the axonal growth cone, very little is known about the molecules that guide dendritic filopodia. Indeed, since the early 1980s, a wide range of research has focused on guidance cues for axons throughout development (Mueller, 1999; Tessier-Lavigne and Goodman, 1996). Biochemical and genetic screens have identified a multitude of proteins acting as attractive and repulsive cues working in a contactdependent fashion or at a distance via secreted molecules (for a review on the mechanisms of axonal guidance, see (Chilton, 2006). These findings have led to a myriad of subtly different signals that are employed during development to ensure the precise wiring of neurons in the nervous system. Further, recent work has examined the role of axonal growth cone filopodia in sensing changes in the molecular milieu during axon pathfinding (Gomez et al., 2001). These results suggest that activation of receptors at the tips of axonal filopodia by extracellular ligands may generate signals that are transduced into changes in motility.  Whether similar molecular cues exist for the  guidance of dendritic filopodia remains unknown. It has been proposed that dendritic filopodia may extend in response to glutamate being released from nearby axons,  146  possibly secreted by extending growth cones (Cooper and Smith, 1992). Recent studies have shown that focal glutamate application leads to an increase in filopodia length (Portera-Cailliau et al., 2003), and that micro-filopodia can extend from the surface of spines in response to brief focal pulse of exogenously applied glutamate (Richards et al., 2005). Conversely, Chang et al. have shown that axonal filopodia movement is highly inhibited by the neurotransmitter glutamate (Chang and De Camilli, 2001). Further, experiments in cultured slices have suggested that filopodia can emerge from dendrites after synaptic activation (Maletic-Savatic, Malinow et al. 1999). However, some studies have shown that filopodia extensions persist even in the presence of glutamate receptor blockers (CNQX and AP-5) (Ziv and Smith 1996) or in the presence of global synaptic activity blockers (TTX) (Wong, Faulkner-Jones et al. 2000) or zero-calcium (PorteraCailliau, Pan et al. 2003). Further, some studies have failed to show the presence of any type of glutamate receptor at the tips of dendritic filopodia (Harris and Stevens, 1989). Hence, it remains unclear whether glutamate serves as the guidance cue for filopodia extension and synaptic contact formation, however, glutamate-receptor channel activation does not appear to be critical in this process. These findings may be further reconciled when one considers that activity during development can be intermittently punctuated by local bursts of glutamate release that can be sensed to guide filopodia to their targets. Thus, dendritic filopodia may be actively exploring the environment, searching for possible synaptic partners by sensing gradients of presynaptically released glutamate. If and how filopodia can sense glutamate gradients is still unknown. Some studies have failed to show the presence of any type of glutamate receptor at the tips of dendritic filopodia (Harris and Stevens, 1989). Hence, more work is required to identify other  147  guidance cues by which filopodia find their way to make contacts with the appropriate presynaptic partners.  4.3.3. In Vivo Work An increasing number of molecules and signaling pathways seem to affect the development, maintenance and plasticity of synapses and dendritic spines. However, most of the studies involving synaptogenesis and spinogenesis have been done in neuronal cultures. Although cultured cells provide a clean and powerful system for dissecting out mechanisms underlying protein assembly and contact formation, they may be unable to provide definitive answers about how synapses are formed in vivo. Because the natural surroundings of the cells have been removed, processes that depend on interaction with neighboring cells and the natural extracellular matrix may be altered in vitro. It will be important in the future to confirm and compare the significance of these findings in vivo. With the advent of new imaging techniques and powerful microscopes, several groups have recently been able to visualize dendritic spines in live animals (Grutzendler et al., 2002; Holtmaat et al., 2005; Niell et al., 2004; Portera-Cailliau et al., 2005; Zuo et al., 2005). Using transgenic mice overexpressing yellow fluorescent protein and transcranial two-photon microscopy, Zuo et al. show the gradual loss of dendritic filopodia and their replacement with spines as animals mature (Zuo et al., 2005). As the molecular control of spine morphogenesis becomes clearer, the challenge will be to use this knowledge to specifically manipulate the shape and density of dendritic spines in vivo and to address a number of outstanding questions. For example, what is the  148  functional significance of dendritic spines in learning, memory and normal cognitive function?  149  4.4.  References  Ahmari, S.E., and S.J. Smith. 2002. Knowing a nascent synapse when you see it. Neuron. 34:333-6. Bamji, S.X. 2005. Cadherins: actin with the cytoskeleton to form synapses. Neuron. 47:175-8. Bresler, T., M. Shapira, T. Boeckers, T. Dresbach, M. Futter, C.C. Garner, K. Rosenblum, E.D. Gundelfinger, and N.E. Ziv. 2004. 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