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Molecular and cellular mechanisms of inhibitory synapse formation in developing rat hippocampal neurons Dobie, Frederick Andrew 2012

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MOLECULAR AND CELLULAR MECHANISMS OF INHIBITORY SYNAPSE  FORMATION IN DEVELOPING RAT HIPPOCAMPAL NEURONS   by   Frederick A. Dobie  B.Sc., The University of Victoria, 2004    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 2012   © Frederick A. Dobie, 2012  ii Abstract  The proper functioning of the brain and central nervous system (CNS) requires the precise formation of synapses between neurons The two main neurotransmitter systems for fast synaptic communication in the CNS are excitatory glutamate and inhibitory g-aminobutyric acid. A growing body of evidence has begun to uncover several shared and divergent rules for the establishment of each of these two types of synapses.  At the molecular level, a number of key proteins have been shown to be involved in the initial formation and subsequent development of synaptic connection, including cell adhesion molecules (CAMs). Among the CAMs, neurexins and neuroligins are important synaptogenic proteins that act trans-synaptically to organize synapses: binding of axonal b- neurexins by neuroligins is sufficient to cause development of a presynaptic specialization at that site, while binding of dendritic neuroligin-1 or neuroligin-2 by b-neurexins is sufficient to cause development of postsynaptic excitatory or inhibitory specializations, respectively. In Chapter 2, we explore the role of a-neurexins in synapse organization. We find a-neurexins are able to specifically induce the formation of inhibitory synapses, presumably through clustering of postsynaptic neuroligin-2. Moreover, we find that the expression of various splice variants of a- and b-neurexins is regulated both during development and by activity, suggesting a physiological role for alternative splicing in the modulation of synapse assembly.  At the cellular level, it is now clear from live imaging studies that synapses and their formation are highly dynamic processes. A number of studies have established the temporal recruitment of pre- and postsynaptic components to nascent synapses and how synapse formation can influence neuron growth. However, these studies have focused on excitatory synapses. In Chapter 3, we explore the cellular mechanisms of inhibitory synapse formation and modulation. We find that entire synapses are highly mobile and can undergo dynamic structural modulation. New synapses are formed by gradual accumulation of components from diffuse cytoplasmic pools, with a significant contribution of presynaptic vesicles from previously recycling sites. These results provide new insights into the mechanisms of inhibitory synapse formation and how it is both similar and different from excitatory synapse formation.   iii Preface  A version of Chapter 2 has been previously published. Kang Y, Zhang X, Dobie F, Wu H, Craig AM (2007) Induction of GABAergic Postsynaptic Differentiation by a -Neurexins. J Biol Chem. Kang, Y. and Zhang, X., who contributed equally to the work, performed all experiments and analysis for cell-based assays (Figures 2.1-2.6, 2.9-2.11). I performed experiments and analysis for RNA isolation and RT-PCR analysis (Figures 2.7-2.8). The manuscript was originally drafted by Craig, A.M. and Kang, Y., with subsequent editing by myself.  A version of Chapter 3 has been previously published. Dobie FA, Craig AM (2011) Inhibitory synapse dynamics: coordinated presynaptic and postsynaptic mobility and the major contribution of recycled vesicles to new synapse formation. J Neurosci 31:10481-10493. I performed all experiments and analysis (with the exception of Figure 3.6B,C which was analyzed by Craig, A.M.). I drafted the original manuscript under the supervision of Craig, A.M.  I assisted in publication of the following manuscript: Linhoff MW, Lauren J, Cassidy RM, Dobie FA, Takahashi H, Nygaard HB, Airaksinen MS, Strittmatter SM, Craig AM (2009) An unbiased expression screen for synaptogenic proteins identifies the LRRTM protein family as synaptic organizers. Neuron 61:734-749. A majority of experiments were carried out by Linhoff, M.W. Transgenic mice were generated by Lauren, J. under the supervision of Airaksinen, M.S. and Strittmatter, S.M. I generated constructs for expression of truncated proteins. I assisted in characterization of newly-generated antibodies. I performed the biochemical fractionations and assisted with other biochemical experiments. I assisted Cassidy, R.M. with immunohistochemistry pilot experiments and analysis. The manuscript was originally drafted by Craig, A.M. and Linhoff, M.W., with subsequent editing by myself.  I assisted in publication of the following manuscript: Sun X, He G, Qing H, Zhou W, Dobie F, Cai F, Staufenbiel M, Huang LE, Song W (2006) Hypoxia facilitates Alzheimer's disease pathogenesis by up-regulating BACE1 gene expression. Proc Nat Acad Sci. 103:18727-32. I performed in vitro experiments for assessment of expression from various promoters.  I assisted in publication of the following manuscript: Dobie F, Craig AM (2007) A fight for neurotransmission: SCRAPPER trashes RIM. Cell. 130:775-7. This was a preview article describing the findings of a paper published in that issue of the journal, including outlining the significance of the authors’ findings in the broader context of the field so as to serve as an introduction for the layperson or uninitiated. I drafted the original manuscript, including artwork. Subsequent editing and refinement was performed by Craig, A.M. before finalization.  All experiments described in this dissertation that involve the use of animals were approved by the UBC Animal Care Committee under certificate A09-0280.  iv Table of Contents  Abstract .............................................................................................................................. ii Preface ................................................................................................................................iii Table of Contents ...............................................................................................................iv List of Figures ...................................................................................................................vii List of Abbreviations ........................................................................................................ viii Acknowledgements ...........................................................................................................xii Dedication ......................................................................................................................... xiii 1 Introduction .................................................................................................................. 1 1.1 Synapse structure and composition ......................................................................... 2 1.1.1 Presynaptic vesicles and neurotransmitter release machinery .......................... 2 1.1.2 Excitatory glutamatergic postsynapses ............................................................. 5 1.1.3 Inhibitory GABAergic postsynapses .................................................................. 9 1.2 Molecular mechanisms of synapse organization .....................................................13 1.2.1 Trans-synaptic cell adhesion molecules ...........................................................14 1.2.2 Secreted factors ...............................................................................................21 1.3 Central role of neurexins, neuroligins, and LRRTMs in   synapse induction and maturation .........................................................................25 1.3.1 Gene and protein strucutre ..............................................................................26 1.3.2 Mechanisms of synapse organization ..............................................................29 1.3.3 Physiological roles of Nrxns, Nlgns, and LRRTMs ...........................................33 1.4 Cellular mechanisms of excitatory synapse formation .............................................35 1.4.1 Initial contact and filopodial dynamics ..............................................................35 1.4.2 Transport of presynaptic proteins .....................................................................36 1.4.3 Transport of postsynaptic proteins ...................................................................37 1.4.4 Maturation of nascent synapses .......................................................................40 1.4.5 The synaptotropic model of axon and dendrite growth .....................................43 1.5 Molecular and cellular mechanisms of inhibitory synapse  formation and modification ......................................................................................44 1.5.1 Assembly and forward trafficking of GABAARs .................................................45 1.5.2 Regulation of GABAAR surface expression ......................................................47 1.5.3 Synaptic anchoring of GABAARs ......................................................................48 1.5.4 Dynamic regulation of inhibitory synapses .......................................................50 1.6 Rationale and hypothesis ........................................................................................52 2 Induction of GABAergic postsynaptic differentiation by a-neurexins .....................55 2.1 Introduction .............................................................................................................55 2.2 Materials and methods............................................................................................57 2.2.1 Primary neuronal culture and COS cell coculture .............................................57 2.2.2 Construction of expression vectors ..................................................................58 2.2.3 Immunocytochemistry and imaging ..................................................................59 2.2.4 Image analysis .................................................................................................60 2.2.5 RNA isolation and RT-PCR ..............................................................................61 2.3 Results ...................................................................................................................62 2.3.1 Specific induction of GABAergic postsynaptic differentiation  by a-neurexins .................................................................................................62  v 2.3.2 Structural basis for the differential synaptogenic activity  of a-neurexins versus b-neurexins ...................................................................68 2.3.3 Expression patterns of neurexins .....................................................................72 2.4 Discussion ..............................................................................................................77 3 Inhibitory synapse dynamics: coordinated pre- and  post-synaptic mobility and the major contribution of  recycled vesicles to new synapse formation ............................................................83 3.1 Introduction .............................................................................................................83 3.2 Materials and methods............................................................................................85 3.2.1 Hippocampal neuron culture and nucleofection ................................................85 3.2.2 DNA constructs ................................................................................................85 3.2.3 Immunocytochemistry and live antibody labeling ..............................................86 3.2.4 Live imaging .....................................................................................................87 3.2.5 Image and data analysis ..................................................................................88 3.3 Results ...................................................................................................................88 3.3.1 Clustering of Gephyrin along developing dendrites occurs  slowly over time ...............................................................................................88 3.3.2 Inhibitory postsynaptic scaffolds show dynamic mobility ..................................91 3.3.3 Inhibitory pre- and postsynaptic structures show correlated  mobility as a single unit ....................................................................................93 3.3.4 The number of inhibitory synapses is variable over  time due to apparent splitting and merging of pre-  and postsynaptic structures..............................................................................95 3.3.5 Development of inhibitory synapses occurs by slow  accumulation of pre- and postsynaptic components  and utilizes recycled synaptic vesicles .............................................................99 3.3.6 Inhibitory synapse formation can occur on both dendritic  shafts and dendritic protrusions and on newly formed  dendrite branches .......................................................................................... 104 3.4 Discussion ............................................................................................................ 105 4 Discussion ................................................................................................................. 112 4.1 Summary of findings ............................................................................................. 112 4.2 A glut of synaptogenic molecules .......................................................................... 115 4.2.1 Synapse initiation versus synapse maturation ................................................ 116 4.2.2 Synapse specificity ........................................................................................ 117 4.3 Dynamics of synaptogenesis ................................................................................ 119 4.3.1 Spatiotemporal order of inhibitory synapse formation ..................................... 119 4.3.2 Putative mechanisms of synapse mobility ...................................................... 120 4.4 Implications for health and disease ....................................................................... 121 4.4.1 Synapse organizing molecule mutations in ASD and schizophrenia ............... 122 4.4.2 GABAergic dysfunction in neurological and psychiatric disorders................... 123 4.5 Perspectives and future directions ........................................................................ 123 References ....................................................................................................................... 125   vi List of Figures   Figure 1.3    Molecular components of the inhibitory GABAergic postsynaptic  compartment .............................................................................................................................. 11  Figure 1.4    Synaptic organizing complexes ................................................................................. 20  Figure 1.5    Neurexin-Neuroligin transsynaptic interaction ........................................................ 28  Figure 1.6    Multiple cellular mechanisms for CNS synaptogenesis ........................................ 39  Figure 2.1    a-Neurexins induce clustering of the inhibitory synaptic  scaffolding protein gephyrin but not the excitatory synaptic  scaffolding protein PSD-95 ...................................................................................................... 63  Figure 2.2    b-Neurexins induce clustering of the inhibitory synaptic  scaffolding protein gephyrin and the excitatory synaptic  scaffolding protein PSD-95 ...................................................................................................... 65  Figure 2.3    Quantitation of induced clustering of gephyrin but  not PSD-95 by a-Neurexins .................................................................................................... 67  Figure 2.4    a-Neurexins induce clustering of neuroligin 2 but not  neuroligin 1/3/4 .......................................................................................................................... 69  Figure 2.5    a-Neurexins induce clustering of GABAA receptor g2 subunit .............................. 70  Figure 2.6    Postsynaptic protein clustering by neurexin-1a requires  LNS6 and is negatively modulated by upstream sequences ............................................. 71  Figure 2.7    Expression of neurexin splice variants lacking the S4 insert  increases with development in hippocampus and cortex ................................................... 73  Figure 2.8    Alternative splicing at the S4 site is regulated by NMDA receptor  activity in hippocampal cultures .............................................................................................. 73  Figure 2.9    Neurexin 2a induces clustering of the inhibitory synaptic  scaffolding protein gephyrin but not the excitatory synaptic  scaffolding protein PSD-95 ...................................................................................................... 74  Figure 2.10   Neurexin 2b induces clustering of the inhibitory synaptic  Scaffolding protein gephyrin and the excitatory synaptic scaffolding  protein PSD-95 .......................................................................................................................... 75  Figure 2.11   a-Neurexins did not induce clustering of GABAA  receptor a5 subunit ................................................................................................................... 76  Figure 3.1    Long-term imaging of inhibitory synapse dynamics ............................................... 90  Figure 3.2    Mobility of YFP-Gephyrin puncta .............................................................................. 92  vii  Figure 3.3    Coordinated movement of inhibitory pre- and  postsynaptic components ........................................................................................................ 96  Figure 3.4    Flux of YFP-Gephyrin puncta in developing neurons ............................................ 98  Figure 3.5    Complex dynamic behaviour of inhibitory synaptic components ...................... 100  Figure 3.6    Formation of new inhibitory synapses ................................................................... 102  Figure 3.7    Inhibitory synapses form on dendritic shafts, dendritic protrusions  and newly formed dendritic branches ................................................................................. 106     viii List of Abbreviations  AChE acetylcholinesterase AMPAR 2-amino-3-(5-methyl-3-oxo-1,2-oxazol-4-yl)propanoic acid AMPAR AMPA receptor ANOVA analysis of variance AP2 adaptor protein 2 APP amyloid precursor protein APV 2-amino-5-phosphopentanoic acid ASD autism spectrum disorders ATP adenosine triphosphate BDNF brain-derived neurotrophic factor BSA bovine serum albumin CA1 cornu ammonis 1 CAM cell adhesion molecule CaMKII Ca2+/Calmodulin dependent kinase II CAZ cytomatrix of the active zone Cbln1 cerebellin 1 precursor protein CCD charge-coupled device cDNA complementary DNA CFP cyan fluorescent protein CMV cytomegalovirus CNQX 6-cyano-7-nitroquinoxaline-2,3-dione CNS central nervous system DABCO 1,4-diazabicyclo[2,2,2]octane DG  dentate gyrus DGC dystrophin-associated glycoprotein complex DIV days in vitro  ix DNA deoxyribonucleic acid E18 embryonic day 18 EGF epidermal growth factor EPSC excitatory postsynaptic current ER endoplasmic reticulum FGF fibroblast growth factor FGFR FGF receptor GABA gamma aminobutyric acid GABAAR GABA type A receptor GABARAP GABAR associated protein GAD glutamic acid decarboxylase GAPDH glyceraldehyde 3-phosphate dehydrogenase GDNF glial cell line-derived neurotrophic factor GEF guanine nucleotide exchange factor GFP green fluorescent protein GFRa1 GDNF receptor a1 GK guanylate kinase GKAP GK domain associated protein GlyR glycine receptor GRIP glutamate receptor interacting protein GSK3b glycogen synthase kinase 3b GTP guanosine triphosphate HEK human embryonic kidney HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid Hsc70 heat shock cognate protein 70 Ig immunoglobulin IgSF Ig superfamily IPSC inhibitory postsynaptic current  x IPSP inhibitory postsynaptic potential JNK c-Jun N-terminal kinase KCC2 Potassium chloride co-transporter 2 LAR leukocyte common antigen-related protein LNS laminin/neurexin/sex hormone LRR leucine-rich repeat LRRTM LRR transmembrane protein LTD long-term depression LTP long-term potentiation MAGUK membrane-associated guanylate kinase MAP2 microtubule associated protein 2 mEPSC miniature EPSC MHC major histocompatibility complex mIPSC miniature IPSC mIPSP miniature IPSP MSD mean-squared displacement nAChR nicotinic acetylcholine receptor Narp neuronal activity-regulated pentraxin NCAM neural cell adhesion molecule NGL Netrin-G ligand Nlgn neuroligin NMDA N-methyl-d-aspartate NMDAR NMDA receptor Nrxn neurexin NSF N-ethylmaleimide Sensitive Factor P11 postnatal day 11 PBS phosphate-buffered saline PCR polymerase chain reaction  xi PDZ PSD-95/Discs-large/Zona occludens PKA protein kinase A PKC protein kinase C PSD postsynaptic density PTP protein tyrosine phosphatase PTV piccolo-bassoon transport vesicle RT-PCR reverse transcriptase PCR SALM synaptic adhesion-like molecule SAP synapse associated protein SH3 Src homology 3 SIRP signal regulatory protein SNAP Soluble NSF Attachment protein SNARE SNAP Receptor SS splice site S-SCAM synaptic scaffold molecule STV synaptic vesicle protein transport vesicle SynCAM synaptic cell adhesion molecule SynGAP synaptic Ras GTPase-activating protein TARP transmembrane AMPAR regulatory protein TrkC neurotrophin receptor tyrosine kinase C Tsp thrombospondin VAMP Vesicle associated membrane protein VASP vasodilator-stimulated phosphoprotein VGAT vesicular GABA transporter VGluT vesicular glutamate transporter YFP yellow fluorescent protein    xii Acknowledgements  I am grateful for the scientific enthusiasm, sage guidance, and patience of my supervisor, Dr. Ann Marie Craig, especially during the trying times when completing this degree seemed like it was impossible.  I am thankful to my supervisory committee members, Drs. Shernaz Bamji, Kurt Haas, and Tim O’Connor for their valuable input and discussion throughout my research.  A large number of people roaming the halls of the Brain Research Centre have provided not only helpful scientific discussion but also levity and wit that made it a true pleasure to come in to the lab. Included among these are members of the Craig Lab: Michael Linhoff, Yunhee Kang, Amanda Rooyakkers, Xiling Zhou, Kevin She, Kate Pettem, Tabrez Siddiqui, Daisaku Yokomaku, and Hideto Takahashi. Other denizens of the BRC that have been a joy to work with include Ainsley Coquinco, Simon Chen, Sharmin Hossain, Blair Duncan, Derek Dunfield, and Sesath Hewapathirane. In spite of, or perhaps because of, our shared plight as trainees, we have had some of the most memorable lunch time conversations of my life.  My appreciation for my dear, sweet Alexandra Tan is immense. Her support and understanding while I have gone on this journey being “curious about the world” have undoubtedly kept me sane over these many years. I can’t wait to share more adventures with her as we become grown-ups.  My deepest thanks to my parents does not seem like enough. Over the seemingly endless years of my schooling, they have been my rocks, my benefactors, my landlords, my inspiration, and my dearest friends. There are so many things in my life that I could only have accomplished with their support and wisdom. They are two of the most kind, sincere, funny, delightful people I know. I would be so lucky if the genetics I learned in school is true.  Finally, my research would not have been possible without the financial support of NSERC, CIHR, and MSFHR. These funding agencies allow Canadian scientists to flourish, and it pains me to know the well is drying up.   xiii Dedication         For my parents, Ted and Susan Dobie.  If I fell ten times, they would help me stand eleven.   1 1  Introduction One of the most stunning features of the mammalian central nervous system (CNS) is its exquisitely complex organization. This organization is derived from highly specific connections between neurons from and within well-defined regions of the brain. This specialized connectivity, known as neural circuitry, forms the basis for proper brain functioning. As might be expected for such an intricate system, the mechanisms by which circuits arise are themselves complex and tightly regulated at many levels. First, during development, specific types of neurons must be generated and migrate to appropriate brain regions (Valiente and Marín, 2010). These neurons’ axons must respond to guidance cues so as to innervate defined brain nuclei or laminae, and often these axons project to only a specific type of neuron within the target region (Tessier-Lavigne, 2002). Finally, innervating axons must form synapses at appropriate subcellular localizations on the target neuron, such as the dendritic arbour, cell body, or axon initial segment (Shen and Scheiffele, 2010). The chemical synapse is the fundamental unit of communication between neurons. The most salient features of a synapse are a presynaptic specialization tightly apposed to a postsynaptic specialization across the ~ 20 nm synaptic cleft (Waites et al., 2005). At its most basic level, the presynaptic specialization contains the machinery for exocytic release and recycling of neurotransmitter-containing vesicles in response to action potential propagation down the axon. The postsynaptic specialization contains ligand-gated ion channels that open in response to neurotransmitter binding, allowing flux of ions to generate a change in membrane potential, along with other scaffolding and signaling complexes that modulate biochemical and biophysical properties of the synapse. Although a growing body of evidence has increased our understanding of the mechanisms by which synapses are  2 initially formed following axo-dendritic contact, mature, and are subsequently maintained or eliminated (Kondo and Okabe, 2011), much is still poorly understood.  1.1 Synapse structure and composition 1.1.1 Presynaptic vesicles and neurotransmitter release machinery The principle means of fast communication between neurons at the synapse is the conversion of a chemical signal to an electrical signal. These chemical signals, neurotransmitters, of which glutamate and GABA are the two main classes for fast synaptic transmission in the CNS, are packaged into specialized membrane-enclosed synaptic vesicles (Rosenmund et al., 2003; Sudhof, 2004).  Synaptic vesicle structure and protein complement  Synaptic vesicles are small (20-80 nm) membrane-enclosed organelles with unique lipid and protein composition (Sudhof, 2004; Takamori et al., 2006). Many synaptic vesicle proteins, either integral to the vesicle membrane or anchored to it by way of lipid modifications, are unique to this organelle and function specifically in the fusion and recycling of vesicles at presynaptic sites. Recent quantitative analysis has shown the presence of over 400 different proteins associated with synaptic vesicles (Takamori et al., 2006) (Fig. 1.1). Many of these proteins can be classified based on their predominant functions. SNARE proteins involved in vesicle fusion with the presynaptic membrane include the synaptobrevins/VAMPs (Rossi et al., 2004), syntaxins (Teng et al., 2001), and SNAPs (Matteoli et al., 2009). Synaptotagmins act as calcium sensors, changing their topology in response to influx of Ca2+ in the nerve terminal and, by interacting with the SNARE complex, promoting vesicle fusion (Pang and Sudhof, 2010). Proteins involved in the re-uptake of synaptic vesicles by endocytosis include AP-1 to AP-3, dynamins (McClure and Robinson,  3 1996), sorting nexin-5 (Carlton and Cullen, 2005), and synaptojanin 1 (Song and Zinsmaier, 2003). Synapsins are thought to modulate synaptic vesicle docking, fusion and recycling (Cesca et al., 2010), while synaptophysins may regulate vesicle fusion by interacting with synaptobrevins (Valtorta et al., 2004). A large assortment of Rab GTPases is required for triggering vesicle docking with the plasma membrane (Ng and Tang, 2008). Additionally, many other proteins, though not considered unique to synaptic vesicles, are nonetheless intimately associated with them. Signaling proteins, like CaMKII (Wang, 2008), casein kinase 1 (Gross et al., 1995), and protein kinase C (Vaughan et al., 1998), are involved in the tight regulation of vesicle fusion and recycling, while cytoskeletal proteins, like actin, tubulin, Arp 2/3 (Ziv and Garner, 2004), kinesin 5A and 5B, dynein, and myosin Va and VIIb (Hirokawa et al., 2010), contribute to vesicle trafficking to and within axonal presynaptic boutons. Finally, a number of channel proteins span the vesicle membrane, involved in filling the vesicle with the appropriate neurotransmitter, driven up their concentration gradients by a V-type H+-ATPase (Gasnier, 2000). In glutamatergic neurons, one of several variants of the protein VGluT pumps glutamate into synaptic vesicles (Takamori, 2006), while in GABAergic neurons, VGAT is responsible for loading vesicles with GABA (McIntire et al., 1997). Additionally, GABAergic vesicles are enriched with GAD65, the enzyme responsible for synthesizing GABA from glutamate (Tobin and Pinal, 1998).  Architecture of the active zone  At the synapse, release of vesicles is localized to a specialized structure called the active zone (Rosenmund et al., 2003). Here, synaptic vesicle fusion and recycling are coordinated by changes in Ca2+ concentration elicited by influx of Ca2+ through voltage- activated calcium channels that respond to action potentials transmitted along the axon. The active zone not only concentrates these calcium channels sublocally, but also acts to tether synaptic vesicles in their vicinity, and to facilitate vesicle docking, fusion, and recycling (Ziv  4 and Garner, 2004). The active zone is rich in cytoskeletal proteins and curves outward into varicosities known as boutons. In the mammalian CNS, these boutons frequently form en passant synapses with many target cells (Ziv and Garner, 2004). A number of other proteins involved in active zone formation have been discovered. Piccolo (Garner et al., 2000) and bassoon (Gundelfinger et al., 1998) are large multi-domain proteins whose multiple protein- protein interaction motifs facilitate emergence of a vast web of connected molecules. Included in this web are ELKS, RIMs, Mints, CASK, Ribeye, Liprin-a, and munc13 (Ohtsuka et al., 2002; Zhen and Jin, 2004; Jin and Garner, 2008). These proteins are implicated in priming synaptic vesicles for docking by their interactions with proteins on the synaptic vesicles themselves. Since the > 200 synaptic vesicles at a given axonal bouton (Sudhof, 2004) may exist in one of several pools based on their releasability (Stevens and Murthy, 1999), it is clear that a high degree of coordination is required to maintain the fidelity of neurotransmission  Figure 1.1. Molecular components of the presynaptic compartment. A. Schematic of some of the proteins that are present in the presynaptic cytomatrix of the active zone (CAZ). B. Molecular model of an average synaptic vesicle (SV) studded with numerous proteins. A adapted with permission from Zhen and Jin, 2004. B adapted with permission from Takamori et al., 2006.      5 1.1.2 Excitatory glutamatergic postsynapses As noted above, glutamate is the key neurotransmitter for fast excitatory transmission in the mammalian CNS. Upon glutamate release from the presynaptic side, a complex network of ion channels, signaling molecules, and scaffolding proteins allow for the transduction of physiological impulses.  Gross anatomy of glutamate synapses A unique aspect of many glutamate postsynapses is their localization to a specialized dendritic protrusion known as a spine (Peters and Kaiserma.Ir, 1970). Spines project from the dendrite shaft roughly 0.5 to 2 mm and may be classified by their general shape: stubby, thin, mushroom-shaped, or cup-shaped (Hering and Sheng, 2001). In many cases, either the maturity or robustness of an excitatory synapse correlates with spine shape (Engert and Bonhoeffer, 1999; Malenka and Bear, 2004; Zito et al., 2009), and indeed spines have been shown to be structurally plastic in intact brains in live animals in response to various stimuli (Zito et al., 2004; Holtmaat and Svoboda, 2009). This structural plasticity is effected by dynamic changes to the underlying actin cytoskeleton, in which spines are rich, along with lipid addition due to insertion of various transmembrane proteins (Yuste and Bonhoeffer, 2004). It is suspected that spines play an important role in compartmentalizing the vast array of molecules involved in excitatory synaptic transmission and that, furthermore, spine geometry may affect the transduction of electrical impulses to the dendrite (Kasai et al., 2005).  Glutamate-gated ionotropic receptors Two main classes of ionotropic receptors exist at glutamate synapses: 2-amino-3-(5-methyl- 3-oxo-1,2-oxazol-4-yl)propanoic acid (AMPA) receptors (AMPARs) and N-methyl-D- aspartate (NMDA) receptors (NMDARs) (Traynelis et al., 2010). A majority of depolarizing  6 current at excitatory synapses is mediated by AMPARs (Sheng and Lee, 2001; Bredt and Nicoll, 2003). Functional AMPARs are assembled as tetramers from up to four subunits, GluR1-4. Each subunit contains a large extracellular N-terminal domain, three transmembrane domains, one “re-entrant” domain that only partially spans the membrane and forms the channel pore, and a C-terminus that is involved in protein-protein interactions and regulation of trafficking by phosphorylation. AMPARs mainly conduct Na+ into the postsynaptic cell to depolarize it; however, if a tetrameric AMPAR lacks the GluR2 subunit, it also becomes permeable to Ca2+ (Seeburg et al., 1998; Malinow and Malenka, 2002). The dynamic insertion or removal of AMPARs in response to different trains of stimuli, and, in turn, the degree of postsynaptic depolarization, are hallmarks of long-term potentiation (LTP) and long-term depression (LTD), respectively, that many speculate are cellular correlates of learning and memory (Malinow and Malenka, 2002; Malenka and Bear, 2004). NMDARs have similar structure to AMPARs, in that they are tetrameric with each subunit have a large N-terminus, three transmembrane domains, a pore-forming re-entrant loop, and C-terminus involved in protein-protein interactions and regulation (Dingledine et al., 1999; Traynelis et al., 2010). There are seven NMDA receptor subunits: NR1, NR2A-D, and NR3A-B (Stephenson et al., 2008). The obligate NR1 subunit, which binds the NMDAR co- agonist glycine, and the NR2 subunit, which binds glutamate, comprise most functional heterotetrameric NMDARs. The precise subunit composition of endogenous NMDARs and the role of NR3 subunits is still unclear (Stephenson, 2001). A critical characteristic of NMDARs are their permeability to Ca2+. Calcium acts as a messenger in the postsynaptic cell to trigger a number of biochemical cascades, including cytoskeletal rearrangement and AMPAR trafficking, that can lead to induction of LTP (Malenka and Bear, 2004). Moreover, NMDAR activation may be important in synaptic development, as recruitment of NMDARs to nascent synapses often precedes their maturation (Washbourne et al., 2002).   7  Figure 1.2. Molecular components of the excitatory glutamatergic postsynaptic compartment. Organization of proteins and protein-protein interactions in the postsynaptic density (PSD). Schematic diagram of the network of proteins in the PSD, with edge of PSD depicted at right. Only major families and certain classes of PSD proteins are shown [in approximate stoichiometric ratio and scaled to molecular size, if known (see text)]. Contacts between proteins indicate an established interaction between them. Domain structure is shown only for PSD-95 (PDZ domain, SH3 domain, GuK domain). Other scaffold proteins are colored yellow; signaling enzymes, green; actin binding proteins, pink. CaMKII (calcium/calmodulin-dependent protein kinase II) is depicted as dodecamer. Unnamed proteins signify the many other PSD proteins that are not illustrated in this diagram. Abbreviations: AKAP150, A-kinase anchoring protein 150 kDa; CAM, cell adhesion molecule; Fyn, a Src family tyrosine kinase; GKAP, guanylate kinase-associated protein; H, Homer; IRSp53, insulin receptor substrate 53 kDa; KCh, K+ channel; mGluR, metabotropic glutamate receptor; nNOS, neuronal nitric oxide synthase; RTK, receptor tyrosine kinases (e.g., ErbB4, TrkB); SPAR, spine-associated RapGAP. Adapted with permission from Sheng and Hoogenraad, 2007.  Scaffolding and signaling molecules of the Postsynaptic Density Another hallmark of glutamate synapses is a vast array of proteins so dense it can be seen by transmission electron microscopy, having a thickness of 30-60 nm (compared to the synaptic cleft of ~ 20 nm) (Harris et al., 1992). Recently, biochemical methods have led to  8 estimates that 200-1000 different proteins are present in this Postsynaptic Density (PSD) (Peng et al., 2004; Cheng et al., 2006; Sheng and Hoogenraad, 2007), including AMPA and NMDARs (Fig. 1.2).  A major class of PSD proteins are scaffolding molecules, making up roughly 6% of the total proteins in the PSD (Sheng and Hoogenraad, 2007). Included among the large host of scaffolding proteins are PSD-95, PSD-93, SAP102, SAP97, GRIP1, PICK1, Shank1, Homer, CASK, and MINT1 (Kim and Sheng, 2004). Composed of numerous protein-protein interaction domains, these scaffolds bind numerous other targets at the PSD to coordinate synaptic transmission and intracellular signaling. One of the most well-studied and abundant protein interaction domains is the PDZ (PSD-95/Discs-large/Zona occludens) domain, with at least 60 proteins containing PDZ domains described in invertebrates (Sheng and Sala, 2001). PDZ domains interact with a specific peptide motif (often E-T/S-D/E-V) at the very C-terminus of a protein, and often self-associate to form large macromolecular complexes (Hsueh et al., 1997). Perhaps the best-characterized PDZ domain-containing proteins are the PSD-95 family, known as membrane-associated guanylate kinases (MAGUKs). These proteins have three PDZ domains, a Src homology 3 (SH3) protein interaction motif, and a guanylate kinase-like domain (GK) (Sheng and Sala, 2001). MAGUKs interact directly with a wide range of other PSD proteins, including NR2 subunits (Kornau et al., 1995) and other surface neurotransmitter receptors (Marshall et al., 1998; Becamel et al., 2004), voltage-gated ion channels (Kim et al., 1995), cell adhesion molecules (Irie et al., 1997; Biederer et al., 2002), motor proteins (Garner et al., 2002; Mok et al., 2002), cell signaling proteins (Chen et al., 1998; Huganir et al., 1998; Tezuka et al., 1999; Eipper et al., 2001), regulators of the cytoskeleton (Sheng et al., 1999a; Firestein et al., 2004), anchoring proteins (Colledge et al., 2000), and other scaffold molecules (Sheng et al., 1999b). Furthermore, PSD-95 affects synaptic transmission by binding stargazin, a regulator of AMPAR surface expression (Schnell et al., 2002b). A number of key cell signaling functions are carried out by other proteins present in the PSD and dendritic spines.  9 CaMKII, a Ca2+-activated kinase thought to be essential for synaptic plasticity (Lisman et al., 2002), is anchored in the PSD by Densin-180 (Walikonis et al., 2001). The synaptic Ras GTPase-activating protein (SynGAP), which binds to PSD-95 and is inhibited by CamKII (Chen et al., 1998), is another important regulator of synaptic plasticity (Komiyama et al., 2002). The A-kinase anchoring protein 79/150 serves as a scaffold for the signaling molecules protein kinase A, protein kinase C, and the protein phosphatase calcineurin (Tavalin et al., 2002), important regulators of AMPAR trafficking. Spine shape, dynamics, and cargo trafficking are also regulated by PSD proteins through their integration of cytoskeletal signaling, particularly through the actin regulatory proteins a-actinin, cortactin, and IRSp53 (Sala et al., 2001). It thus becomes clear that, by bringing in close proximity a number of critical proteins involved in synaptic transmission and cell signaling, the PSD is uniquely suited for signal amplification.  1.1.3 Inhibitory GABAergic postsynapses As compared to glutamatergic synapses, far less is known of the molecular composition of GABAergic synapses, the major sites of fast inhibitory synaptic transmission. Whereas most glutamate synapses are localized to dendritic spines, a majority of GABA synapses are present on the dendritic shaft and have a poorly understood structure. Despite this, GABA postsynaptic sites are well-apposed to GABAergic axonal inputs for high fidelity inhibitory transmission.  GABA-gated ionotropic receptors The main fast inhibitory effect of GABA release onto postsynaptic neurons is through the GABAA receptor (GABAAR). GABAARs share homology with a superfamily of receptors including nicotinic acetylcholine receptors and 5-hydroxytryptamine type 3 receptors (Grenningloh et al., 1987; Maricq et al., 1991). GABAARs assemble into heteropentamers  10 from up to 19 different subunits: a1-6, b1-3, g1-3, d, e, p, r1-3, and q (Michels and Moss, 2007). Each subunit is composed of a large extracellular N-terminus, four transmembrane domains, a cytoplasmic loop between transmembrane domains 3 and 4, and a short extracellular C-terminus (Moss and Smart, 2001). Although the precise stoichiometry of receptor subunits is unclear, studies suggest that most mature GABAARs exist with the subunit ratio 2a:2b:1g (Chang et al., 1996; Tretter et al., 1997). The interface between a and b subunits contains the binding site for GABA, while the second transmembrane domain of each subunit lines the channel pore (Michels and Moss, 2007). The expression pattern and subunit assembly stoichiometry play critical roles in the localization and function of GABAARs. A majority (60%) of GABAARs are composed of a1b2g2 subunits, are located apposed to GABAergic presynaptic terminals on the cell soma and dendrites in numerous brain regions, and are responsible for phasic inhibition; that is, the fast inhibitory effects of GABA released at synaptic sites (Möhler, 2006). Receptors composed of a2b3g2 (15-20% of total) are also widely distributed, with synaptic localization including at the axon initial segment, while receptors composed of α3bng2 subunits (10-15%) share similar subcellular localization as a2b3g2 receptors but in different brain regions (Möhler, 2006). Another fraction of GABAARs (15%) containing various combinations of a,b,g, and d subunits are localized extrasynaptically, effecting  tonic inhibition, the response to ambient GABA in the neuropil (Möhler, 2006). GABAARs are permeable to Cl-. At resting membrane potential in mature neurons, the influx of Cl- causes hyperpolarization of the postsynaptic membrane. Interestingly, in immature neurons that express lower levels of the K+/Cl- co-transporter KCC2, the driving force for chloride is reversed from that of mature neurons, leading to activation of GABAARs becoming depolarizing (Rivera et al., 1999). This phenomenon appears important for the maturation of GABAergic synapses and neural circuits (Cherubini et al., 2011).  11   Figure 1.3. Molecular components of the inhibitory GABAergic postsynaptic compartment. Synaptic GABAA receptors are stabilized by a submembranous lattice of gephyrin by direct interaction. Cytoskeleton associated proteins are Dlc1/2 and Mena/VASP. Collybistin, a guanine nucleotide exchange factor is membrane associated and interacts with gephyrin. The dystrophin- glycoprotein complex (DGC) stabilizes the synapse and neuroligins bridge the synaptic cleft by interaction with presynaptic neurexins. Adapted with permission from Tretter and Moss, 2008.  Accessory GABAergic postsynaptic proteins Unlike glutamate synapses, GABA synapses lack a distinct PSD. Moreover, far fewer proteins have been identified that localize specifically to inhibitory synapses, likely due to a lack of biochemical methods for purification of GABAAR-associated complexes. Several  12 proteins localized to GABAergic synapses, though, have been identified (Fig. 1.3). The protein which most similarly resembles the macromolecular complexes of glutamate synapse PSDs is gephyrin (Kneussel and Loebrich, 2007). Along with being involved in molybdenum cofactor biosynthesis, gephyrin forms a hexagonal lattice by trimerization of its G domain and dimerization of its E domain (Sola et al., 2004; Saiyed et al., 2007). Gephyrin may be alternatively spliced, with alterations in splice isoforms affecting oligomerization and, putatively, gephyrin function (Paarmann et al., 2006). Gephyrin also interacts with many proteins at GABA postsynapses, including GABAARs themselves through direct interactions with the a1, a2 or a3 subunits (Tretter et al., 2008; Mukherjee et al., 2011; Tretter et al., 2011). Gephyrin oligomerization is regulated through its interactions with proteins including heat shock cognate protein 70 (Hsc70) (Triller et al., 2011), protein phosphatase 1 (Bausen et al., 2010), and protein kinases (Langosch et al., 1992). Gephyrin also interacts with glutamate receptor interacting protein 1 (GRIP1), although the physiological role of this interaction is not well understood (Yu et al., 2008). The clustering of gephyrin at synapses is further regulated by collybistin, a RhoGEF that interacts with both gephyrin and Cdc42 (Kins and Kirsch, 2000; Saiepour et al., 2010; Tyagarajan et al., 2011). Interestingly, gephyrin has also been shown to interact directly with neuroligin 2, a synaptogenic molecule known to selectively promote formation of inhibitory synapses; the mechanism of this neuroligin 2- dependent synaptogenesis was found to be depending on collybistin activation (Poulopoulos et al., 2009). While gephyrin is at least partially dispensable for GABA synapse formation (Yu et al., 2007; O'Sullivan et al., 2009), collybistin is not (Papadopoulos et al., 2007; Papadopoulos et al., 2008). Another protein complex observed selectively at a subset of GABAergic synapses is the dystrophin-associated glycoprotein complex (DGC) (Waite et al., 2009). A member of this complex, dystroglycan, binds both the extracellular matrix and the presynaptic protein neurexin (Sugita et al., 2001). Although they may not be critical for formation of GABAergic synapses in cultured neurons (Levi et al., 2002), the ablation of  13 dystroglycan or dystrophin in vivo alters baseline synaptic transmission and synaptic plasticity (Knuesel et al., 1999; Moore et al., 2002). Similar to the PSD of excitatory synapses, postsynaptic specializations of inhibitory synapses are intimately linked to the underlying cytoskeleton. Indeed, gephyrin directly interacts with both Mena/VASP, a protein that recruits G-actin for actin filament formation (Giesemann et al., 2003), and profilin 1 and 2, a protein that regulates actin polymerization in response to phosphoinositol diphosphate (Neuhoff et al., 2005). Gephyrin is also tightly associated with polymerized tubulin at synaptic sites (Ramming et al., 2000). Although there is a growing understanding of how these known GABAergic postsynaptic proteins contribute to inhibitory synapse formation and physiological function, much is still to be learned.  1.2 Molecular mechanisms of synapse organization As noted previously, synapse formation is a complex process that, among other factors, requires proper spatiotemporal, stable neuron-neuron contact, followed by recruitment of presynaptic vesicles and release machinery in an axon apposed to the matching neurotransmitter receptors and accessory proteins in the postsynaptic target cell, and, in most cases, ultimately requires functional release of and response to neurotransmitters for that connection to be maintained. If one of these criteria is not met, a synapse will not form or persist. While a complete picture of synaptogenesis remains elusive, a number of key molecules that play one or more roles in this process have recently been characterized. These molecules include a growing catalogue of both cell-surface attached cell adhesion molecules (CAMs) and diffusible factors, each of which influences synapse organization in multiple ways (Fig. 1.4).    14 1.2.1 Trans-synaptic cell adhesion molecules Cadherin family proteins Cadherins are adhesion molecules that form trans-cellular, homophilic interactions in a Ca2+- dependent manner at the junctions of many cell types, including neurons and glia (Geiger and Ayalon, 1992; Fannon and Colman, 1996; Koch et al., 1997). Intracellularly, neuronal N- cadherins are coupled to the cell-signaling proteins a-, b-, and p120-catenin, which are key regulators of cytoskeletal dynamics (Jou et al., 1995). Related to the “classical” cadherins, protocadherins bring the total number of known proteins in this family to over 100 (Wu and Maniatis, 1999). N-cadherins are localized to synapses in both the pre- and postsynaptic compartments (Yamagata and Sanes, 1995; Fannon and Colman, 1996), although most likely at the borders of synapses (Uchida et al., 1996). Interestingly, N-cadherin accumulates at sites of initial axo-dendritic contact prior to synaptogenesis (Benson and Tanaka, 1998). Expression of a broadly acting dominant negative cadherin in neuron cultures disrupts synapse formation (Togashi et al., 2002), but disruption of specific cadherins in vivo has primarily affected axon targeting, synapse specificity, and synapse plasticity rather than synapse assembly (Inoue and Sanes, 1997; Lee et al., 2001). Moreover, N-cadherin is not able to directly induce hemi-synapse formation in a co-culture system (Sara et al., 2005). Confounding the issue, deletion of the downstream cadherin signaling molecule p120- catenin does reduce synapse density in hippocampal pyramidal neurons in vivo, although it is uncertain whether this affect results directly from N-cadherin interactions (Elia et al., 2006). Protocadherins are also localized to synapses (Colman et al., 2003), where they directly interact intracellularly with the kinase fyn to possibly regulate synaptic function (Kohmura et al., 1998). Like N-cadherins, though, it seems that protocadherins do not directly affect synapse formation, but rather act in target specification or stabilization of initial contacts (Lee et al., 2003). Although the role of cadherin family proteins in initial synapse formation remains unclear, evidence suggests they play important roles in synapse stability,  15 maturation, and plasticity. Acting through Rho GTPases downstream of a-catenin, N- cadherin signaling regulates dendritic filopodia motility and spine formation (Togashi et al., 2002; Abe et al., 2004), while interaction of b-catenin with AMPARs affects their trafficking to synapses (Nuriya and Huganir, 2006). Presynaptic maturation is also affected by b-catenins, whose ablation causes a decrease in the number of synaptic vesicles in the reserve pool (Bamji et al., 2003).  EphB receptors and ephrinBs Ephs and their ligands ephrins are another family of cell adhesion molecules with widespread distribution and biological function, including well-known roles in axon guidance during early development (Kullander and Klein, 2002). While a number of classes exist, EphB and ephrinB, transmembrane receptor tyrosine kinases, are the best characterized synaptic organizing factors (Dalva et al., 2007). Postnatally, EphB2 and EphB3 have been shown to localize postsynaptically (Buchert et al., 1999), while it is suggested that ephrinBs localize to the presynaptic terminal (Yancopoulos et al., 1998). Multiple intracellular domains of both EphBs and ephrinBs facilitate downstream signaling or interaction with other signaling proteins, including the kinase domain of EphBs, PDZ-binding domains, and SH2/SH3 domains that interact with guanine nucleotide exchange factors that regulate the cytoskeleton (Kullander and Klein, 2002). Furthermore, the extracellular domain of EphBs can interact directly with the NR1 subunit of NMDARs, and clustering of EphBs leads to co- aggregation of NMDARs (Dalva et al., 2000), while the PDZ-binding domain of EphB2 is involved in AMPAR synaptic incorporation (Kayser et al., 2006). Knockout of various EphBs also leads to a decrease in excitatory synapses in vivo and impaired LTP (Grunwald et al., 2001; Henderson et al., 2001; Henkemeyer et al., 2003; Kayser et al., 2006). EphB-ephrinB interaction has also been implicated in spine development (Huganir et al., 2003). Apart from  16 acting in postsynaptic development, recent evidence also suggests that signaling by ephrinB in the presynaptic compartment may influence its maturation (Contractor et al., 2002).  SynCAMs Synaptic Cell Adhesion Molecules (SynCAMs) belong to a superfamily of proteins containing multiple extracellular immunoglobulin (Ig) domains (IgSF proteins) (Rougon and Hobert, 2003). SynCAM1 was first reported to be expressed at excitatory synapses and contain a PDZ-binding domain that can interact with CASK (Biederer et al., 2002). Initially thought to form trans-synaptic homophilic interactions, it has now been shown that SynCAMs 1-4 preferentially form heterophilic adhesion complexes (Fogel et al., 2007) by first oligomerizing in cis (Fogel et al., 2011). SynCAM1 expression on the surface of heterologous cells presented to axons is able to induce presynaptic assembly at their contact sites (Biederer et al., 2002), and SynCAM1 is rapidly recruited to nascent axo-dendritic contacts (Stagi et al., 2010). While overexpression of SynCAM1 does not increase excitatory synapse number in cultured neurons (Sara et al., 2005), it does in transgenic mice, where its knockout conversely reduces excitatory synapse number and influences LTD (Robbins et al., 2010). Postsynaptically, SynCAM1 is able to recruit both NMDA and AMPARs through the interacting proteins 4.1B and 4.1N, respectively (Hoy et al., 2009).  NCAM Another IgSF protein, neural cell adhesion molecule (NCAM), plays a prominent role in axon guidance in hippocampal development (Cremer et al., 1997) and is synaptically localized in the postnatal developing striatum (Uryu et al., 1999), suggesting a possible role in synaptogenesis. While NCAM has been shown to be important for synaptic physiology (Ronn et al., 2000), it does not appear to be directly involved in synapse formation as knockout of NCAM in cultured neurons does not affect synapse density (Schachner et al.,  17 2000). Interestingly, though, disruption of the interaction between NCAM and the extracellular matrix molecules heparan sulfate proteoglycans affects synapse density (Dityatev et al., 2004), highlighting potential novel roles for the extracellular matrix in shaping synapse development.  SALMs Synaptic adhesion-like molecules (SALMs) also contain an extracellular Ig domain, but are mainly classified by their repeated extracellular leucine-rich repeat (LRR) motifs (Kim and Ko, 2007). Also containing an intracellular PDZ-binding domain, SALM1-4 were first identified in a screen for proteins that interact with PSD-95 and related family members (Wang et al., 2006), and were further found to both localize to the synaptic density and interact with the extracellular domain of the NR1 subunit of NMDARs. SALM2 was subsequently shown to regulate excitatory synapse development: artificial aggregation of SALMs on dendrites induces the clustering of several postsynaptic proteins, and SALM1 knockdown decreases synapse density measured both microscopically and electrophysiologically (Ko et al., 2006). Puzzlingly, presentation of SALM2 on the surface of heterologous cells to the axons of cultured neurons does not result in presynaptic differentiation, an effect shown for several other synaptogenic proteins (Ko et al., 2006), nor is there a presynaptic ligand known for SALMs. Therefore, while SALMs are potentially important regulators of synapse organization, further investigation is required to decipher their mechanisms of action.  Netrin-Gs, LAR/PTPRs, and Netrin-G ligands Netrin-G ligands (NGLs), like SALMs, are LRR-containing transmembrane proteins (Siddiqui and Craig, 2010). Along with their canonical binding partners, the GPI-anchored netrin-G proteins, they are known to be important for neural connectivity during development (Yin et  18 al., 2002). NGL-1 and 2 contain a cytoplasmic PDZ-binding domain that interacts with PSD- 95, and are thus localized postsynaptically (Kim et al., 2006). While overexpression of NGL- 2 increases both pre- and postsynaptic density in a netrin-G2-dependent manner, direct aggregation of netrin-Gs alone is insufficient to induce presynaptic development, suggesting the possibility of another factor involved in NGL-1,2/netrin-G1,G2 bidirectional signaling during synapse induction (Kim et al., 2006). Another NGL isoform, NGL-3, does not bind either netrin-G1 or netrin-G2, but does have potent excitatory postsynaptic inducing properties (Woo et al., 2009). Recently, several receptors for NGL-3 have been discovered, the receptor tyrosine phosphatase leukocyte common antigen-related (LAR) (Woo et al., 2009), a protein already implicated in excitatory synapse development (Dunah et al., 2005), and the protein tyrosine phosphatases PTPd and PTPs (Kwon et al., 2010). While the first LRR motif of NGL-3 binds the first two fibronectin III domains of all three receptors, the effects of binding are different. NGL-3/LAR and NGL-3/PTPs signal bidirectionally to induce both pre- and post-synaptic development, while the NGL-3/PTPd interaction is sufficient only to induce presynaptic development (Kwon et al., 2010). The differential effects of NGL-3 and its receptors, along with the unique laminar distribution of netrin-Gs (Nishimura-Akiyoshi et al., 2007), make these proteins interesting subjects of further study on their potential roles in synapse specificity.  Other synaptogenic transmembrane proteins Several other trans-synaptic signaling complexes have recently been implicated in synaptic organization. The well-known immune system protein major histocompatibility complex I (MHCI), although not traditionally thought of as a CAM, is present both pre- and postsynaptically in the developing CNS (Needleman et al., 2010) and may act as a negative regulator of synapse formation (Glynn et al., 2011), although the mechanisms of this activity  19 are still poorly understood (Garay and McAllister, 2010). The postsynaptic receptor ErbB4 interacts with PSD-95 to regulate NMDAR surface expression (Garcia, 2000), and interaction with its receptor neuregulin also modifies synapse development (Krivosheya et al., 2008). A protein heavily implicated in the pathogenesis of Alzheimer’s Disease, amyloid precursor protein (APP), now also has a newfound role in synaptogenesis. Its deletion both pre- and postsynaptically leads to defects in neuromuscular synapse formation (Wang et al., 2009). Moreover, presentation of APP to contacting axons in a co-culture assay led directly to presynaptic development, putatively acting through its intracellular domain which binds both CASK and Mint1 (Wang et al., 2009). Finally, the neurotrophin receptor tyrosine kinase C (TrkC), whose response to neurotrophins in early development is well-established, has also recently been implicated in synapse development. Independent of neurotrophin binding, postsynaptic clustering of TrkC induces excitatory synapse development (Takahashi et al., 2011). This effect is dependent on binding to the presynaptic protein PTPs, whose clustering also induces presynaptic differentiation (Takahashi et al., 2011) and a receptor already discovered to be important in mediating the excitatory synapse development effects of NGL- 3 (Kwon et al., 2010). It appears that the list of proteins involved in synapse initiation and maturation will continue to grow, as a recent screen for synaptogenic molecules using the neuron/heterologous cell co-culture assay revealed a number of proteins putatively involved in aspects of synapse organization that remain to be characterized (Linhoff et al., 2009). The investigation of these and other new proteins will provide great insight into the increasingly complex problem that is synapse development.      20 Figure 1.4. Synaptic organizing complexes. An inventory of synaptogenic molecules, defined here as proteins that induce presynaptic (←) or postsynaptic (→) differentiation when presented to axons or dendrites, respectively. Many of the adhesion complexes have bidirectional synaptogenic activity (↔). The main receptors are also shown for the secreted synaptogenic factors. PDZ domain binding sites and common protein domains are indicated. Adapted with permission from Siddiqui and Craig, 2011.  21 1.2.2 Secreted factors Narp Neuronal activity-regulated pentraxin (Narp) was first discovered, as the name implies, as a transcript induced by activity in the hippocampus (Tsui et al., 1996). Narp is expressed and putatively secreted by neurons both pre- and postsynaptically (O'Brien et al., 1999). It binds the extracellular domain of AMPARs (O'Brien et al., 1999) and can potently induce their surface clustering (O'Brien et al., 2002). The role of Narp as a diffusible factor in synapse induction is evident from the data that its secretion from heterologous cells can induce AMPAR clustering on co-cultured spinal cord neurons, and inhibition of neuronal Narp secretion occludes its synaptogenic effects (O'Brien et al., 2002). A related protein, the pentraxins NP1, when in a complex with Narp, also displays synaptogenic activity, inducing clustering of AMPARs and, through their indirect coupling to AMPARs, NMDARs (Xu et al., 2003). Narp’s role in recruiting AMPARs to developing synapses has been shown to be crucial for their functional maturation and, in turn, establishment of appropriate connectivity in the mature visual system (Koch and Ullian, 2010). The exact sources and targets of Narp, however, remain unclear, as evidence suggests Narp acts primarily in the formation of excitatory synapses on interneurons but not on pyramidal cells (Mi et al., 2002). Moreover, the function of Narp may not be specifically synaptogenic per se, but rather in scaling of synapses as a homeostatic mechanism in response to changes in neural circuit activity (Chang et al., 2010).  Thrombospondins The growth of neurons in dissociated culture has been a common technique for over 30 years (Banker and Cowan, 1977). Pioneering studies on the mechanisms of synapse formation in culture led to the discovery that molecules secreted by glial cells, either by co- culturing neurons with glia or treating neurons with media conditioned by glia, could greatly  22 increase synapse density (Pfrieger and Barres, 1997; Nagler et al., 2001). Initial characterization of glia-conditioned media showed that cholesterol secreted by glia and bound to apolipoprotein-E was an essential factor in meeting the high metabolic demands of neuron growth and synaptic development, although not itself directly synaptogenic (Mauch et al., 2001). However, recent evidence has implicated glia-secreted thrombospondins-1 and -2 (TSPs) as synaptogenic factors (Christopherson et al., 2005). TSPs are extracellular matrix proteins with well-characterized roles in inflammation in the body’s periphery (Lopez- Dee et al., 2011). Treatment of cultured neurons with recombinant TSP is able to induce proliferation of synapses that are presynaptically normal but are postsynaptically silent, having only clustered NMDARs but not AMPARs at these sites (Christopherson et al., 2005). Interestingly, treatment of neurons with astrocyte-conditioned media is able to induce formation of both pre- and postsynaptically normal synapses, suggesting another factor secreted by glia can enhance synapse formation. Nonetheless, a receptor for TSPs on dendrites has recently been discovered, the auxiliary calcium channel subunit a2d-1 (Eroglu et al., 2009), shown to be necessary for mediating the effects of TSPs in postsynaptic induction.  It remains unclear what downstream signaling occurs that leads to recruitment of NMDARs to these synapses, although it has recently been suggested that the action of TSPs requires neuroligin-1, leading to an increase in the rate of synapse formation (Xu et al., 2010). How TSPs induce normal presynaptic maturation, either through a direct axonal receptor or retrograde signaling from dendrites, is unknown. Furthermore, TSPs, rather than being locally synaptogenic, may act by increasing a neuron’s global competency for synapse formation by engaging certain gene expression paradigms (Siddiqui and Craig, 2010).     23 FGFs and FGFRs In an elegant unbiased biochemical screen, fibroblast growth factor 22 (FGF22) was found to induce presynaptic differentiation in cultured motoneurons (Umemori et al., 2004), along with the related species FGF7 and FGF10. This recapitulated previous findings that showed FGF2 could enhance excitatory synaptogenesis in hippocampal neurons (Li et al., 2002). FGFs are thought to be secreted from target dendrites, where they subsequently bind axonal FGF receptors (FGFRs). In vivo, FGF22 is expressed by cerebellar granule cells during a critical period of synaptogenesis, while FGFR2 is expressed by incoming mossy fiber axons. Disruption of FGF7, -10, -22 and FGFR2 all inhibit presynaptic differentiation of mossy fiber-granule cell synapses (Umemori et al., 2004). Surprisingly, hippocampal CA3 pyramidal neurons express both FGF22 and FGF7, which have differing roles in presynaptic development onto these cells: FGF22 promotes organization of excitatory synapses while FGF7 promotes organization of inhibitory synapses (Terauchi et al., 2010). In line with these findings, mice lacking FGF22 are resistant to epileptic seizures while mice lacking FGF7 are prone to them (Terauchi et al., 2010). How these distinct roles of FGFs and their receptors in excitatory versus inhibitory synapse organization are carried out mechanistically is a fascinating problem that requires further investigation.  Wnt7a and Frizzled Early in development, the Wnt family of secreted proteins have well-characterized roles in neuronal morphogenesis, but also play key roles in neural circuit formation (Salinas and Zou, 2008). Roles for Wnt-7a in synapse organization and maturation have recently been uncovered. First, acting through canonical signaling pathways that involve Wnt binding to the transmembrane receptor Frizzled and downstream activation of b-catenin and glycogen synthase kinase 3b (GSK3b), Wnt-7a both increases clustering of synaptic vesicles and, in  24 turn, vesicle release properties (Cerpa et al., 2008), but also is involved in clustering of presynaptic a7-nicotinic acetylcholine receptors (nAChRs) important for modulating presynaptic release (Farias et al., 2007). On the postsynaptic side, Wnt-5a activates a non- canonical signaling pathway upon binding to Frizzled, involving activation of c-Jun N- terminal Kinase (JNK) that leads to redistribution of PSD-95 (Farias et al., 2009). However, since only the acute effects of application of Wnt-5a on postsynaptic remodeling have been addressed, it is unclear whether this non-canonical pathway is engaged in vivo during synaptogenesis or synaptic plasticity. Strikingly, Wnt has been shown to be important for positioning of synapses in C. elegans, where Wnt/Frizzled signaling actually inhibits presynaptic assembly in a subset of neurons (Klassen and Shen, 2007). Whether this function is present in mammals whose synaptic organization is not as stereotypical as the worm remains to be seen.  Other secreted factors A growing body of work has begun to highlight other key diffusible molecules important in synapse organization. Brain-derived neurotrophic factor (BDNF) has been shown to promote inhibitory synaptogenesis by acting through TrkB receptors both pre- and postsynaptically (Seil and Drake-Baumann, 2000; Luikart et al., 2005), and, perhaps as a homeostatic response, increase formation of excitatory synapses as well (Vicario-Abejon et al., 1998). The related neurotrophin NT-3 also affects synaptogenesis through TrkC receptors (Martinez et al., 1998). However, it is generally thought these molecules prime neurons to become more permissive to synaptogenesis (Waites et al., 2005). Although transmembrane proteins, signal regulatory proteins (SIRPs) a, b, and g have extracellular domains that are cleaved at the postsynaptic membrane and diffuse to the presynaptic membrane, where they induce vesicle clustering in a partially CD47-dependent manner (Umemori and Sanes,  25 2008). Recently, a ligand-induced cell adhesion molecule was discovered.  Acting as a bridging molecule, glial cell line-derived neurotrophic factor (GDNF) binds to GDNF receptor a1 (GFRa1) present on both the pre- and postsynaptic membrane to induce their transsynaptic interaction, which subsequently leads to clustering of presynaptic vesicles (Ledda et al., 2007). Another transsynaptic bridging molecule has also recently been discovered, cerebellin 1 precursor protein (Cbln1). In cerebellum, it was shown that the critical role of postsynaptic GluRd2 in synaptogenesis is mediated by direct binding of its extracellular domain to the well-characterized presynaptic synaptogenic protein neurexin, in turn inducing bidirectional synapse organization, but this occurs only in the presence of the diffusible molecule Cbln1 (Mishina et al., 2010). It was subsequently shown that Cbln family members can interact with various neurexin splice variants in a brain region-specific manner to induce synaptogenesis (Mishina et al., 2011). Moreover, Cblns may simultaneously either compete or cooperate with other postsynaptic neurexin ligands, neuroligin 1 and leucine-rich repeat transmembrane protein 2 (LRRTM2), respectively, for their synaptogenic properties (Mishina et al., 2011; Yuzaki and Matsuda, 2011). These results, along with those which will be presented below, have begun to highlight the critical central role neurexins may play in presynaptic organization.  1.3 Central roles of neurexins, neuroligins, and LRRTMs in synapse induction and maturation  As seen from the extensive catalogue shown above, an ever-increasing number of proteins are important for synapse organization. However, perhaps the best characterized synaptogenic proteins are the presynaptic neurexins (Nrxns) and postsynaptic neuroligins (Nlgns). Recently, another family of receptors for neurexins with potent synaptogenic properties, the LRRTMs, have been described, along with the Cbln1/GluRd2 interaction outlined above. It is thus becoming ever clearer how muddled the questions surrounding  26 synapse organization are. Nonetheless, groundbreaking studies on the biochemistry, cell biology, and genetics of these synaptogenic proteins has begun to reveal their importance for synaptic function in health and disease.  1.3.1 Gene and protein structure Neurexins Neurexins were initially characterized as being a receptor for the black widow spider venom component a-latrotoxin, which induces massive release of neurotransmitter from presynaptic terminals (Ushkaryov et al., 1992). Mammals have three genes for Nrxn. Each gene has both an upstream and downstream promoter that drives expression of the longer a- and shorter b-Nrxns, respectively. Considerable Nrxn protein diversity is generated by alternative splicing. Each Nrxn gene contains five potential sites of alternative splicing, termed SS#1-5, all located in the mature protein’s extracellular region. Given the three NRXN genes, each with two promoters and five alternative splice sites, there may be as many as 3908 different Nrxn polypeptides (Tabuchi and Sudhof, 2002). In addition, Nrxns may be differentially N- or O-glycosylated, conferring further diversity (Craig and Kang, 2007a). Mature a-Nrxns are ~ 150-170 kDa polypeptides composed of six extracellular Laminin/Neurexin/Sex-hormone (LNS) domains arranged into modules with three epidermal growth factor (EGF)-like domains (Fig. 1.5). They also contain a highly glycosylated extracellular domain, followed by a transmembrane domain, and an intracellular domain that includes a PDZ-binding domain. This intracellular domain is known to bind the synaptic vesicle protein synaptotagmin (Hata et al., 1993) along with the presynaptic scaffolding/signaling proteins CASK and Mint (Hata et al., 1996; Biederer and Sudhof, 2000), thereby acting as a putative nucleating factor for coupling synaptic vesicles and their release apparatus (Dean et al., 2003; Missler et al., 2003). The b-Nrxns are ~ 50-60 kDa  27 polypeptides that are essentially N-terminally truncated versions of the a-Nrxns, composed of only the sixth LNS domain and sharing the rest of their C-terminal structure with a-Nrxns (Craig and Kang, 2007a). It is becoming clear that alternative splicing of Nrxns is critical in determining their transsynaptic interaction with other proteins, including Nlgns and LRRTMs, but the localization of these splice forms and how they are generated remains poorly understood (Ullrich et al., 1995; Kang et al., 2008).  Neuroligins Neuroligin-1 was first discovered using immobilized Nrxn as bait on an affinity purification column (Ichtchenko et al., 1995). Humans have five Nlgn genes, NLGN1-4 and NLGN4Y (Jamain et al., 2003). Each gene encodes an ~ 90 kDa polypeptide with an extracellular domain homologous to acetylcholinesterase (AChE) but which lacks catalytic activity, a glycosylated domain, a transmembrane domain, and an intracellular domain that includes a PDZ-binding domain (Craig and Kang, 2007a). Along with binding Nrxns, the AChE-domain of Nlgns mediate their dimerization in the postsynaptic plasma membrane (Comoletti et al., 2003). Although this dimerization appears to be required for Nlgn function, it remains to be shown whether Nlgns exist predominantly as homo- or hetero-dimers. Additional Nlgn diversity is conferred through alternative splicing in the AChE domain mediated through two splice sites, A and B (Boucard et al., 2005). Intracellularly, Nlgns are able to bind a number of proteins, including PSD-95, S-SCAM, Shank, and PICK1 (Irie et al., 1997; Meyer et al., 2004). Recently, a 15 amino acid gephyrin-binding motif was also discovered to be conserved in Nlgns (Poulopoulos et al., 2009). Despite the fact that all Nlgns contain domains for interacting with excitatory and inhibitory synaptic scaffolds, respectively, in culture and in vivo, different Nlgns have distinct localizations to excitatory or inhibitory synapses. Nlgn1 localizes to excitatory glutamatergic synapses (Brose et al., 1999); Nlgn2  28 and Nlgn4 localize to inhibitory GABAergic and glycinergic synapses (Graf et al., 2004; Varoqueaux et al., 2004; Hoon et al., 2011); and Nlgn3 is localized to both excitatory and inhibitory synapses (Budreck and Scheiffele, 2007; Levinson et al., 2010). Initial studies have identified protein motifs that at least partially mediate synapse specificity of various Nlgns (Silverman et al., 2005; Levinson et al., 2010).    Figure 1.5. Neurexin-Neuroligin transsynaptic interaction. Structure of neurexins and neuroligins. In humans, there are three neurexin genes and five neuroligin genes. Each neurexin gene uses an upstream promoter to generate the larger α-neurexins and a downstream promoter to generate the smaller β-neurexins. Thus, β-neurexins can be thought of as N-terminally truncated α-neurexins that have a short β-specific leader (βN). In α-neurexins, the LNS (laminin, neurexin, sex-hormone- binding protein) domains are organized with EGF (epidermal growth-factor)-like domains into three homologous modules, I–III. The position of each of five sites of alternative splicing (SS1–SS5) is indicated. Neuroligins contain an extracellular acetylcholinesterase(AChE)-homologous domain that contains one or two sites of alternative splicing (SSA, plus SSB in the case of neuroligin 1). Both neurexins and neuroligins contain a highly glycosylated region (CH) and a transmembrane domain (TM; not present in some splice variants of neurexin 3), and terminate in PDZ-domain-binding sites (PDZ BD). Shown between the neurexins and neuroligins are structures of AChE, a model for the AChE-homologous domain of neuroligins, and the neurexin 1β LNS domain. The position of splice sites SS2–SS4 is shown on a single LNS domain for simplicity, although SS2 and SS3 actually occur in different LNS domains of α-neurexins. Note also that the left face of the neurexin LNS as shown here binds neuroligin but the precise structure and interacting region of neuroligin has not been reported yet. Adapted with permission from Craig and Kang, 2007.   29 LRRTMs Leucine rich repeat transmembrane proteins were originally identified using bioinformatics to search for proteins containing LRR domains similar to the Slit family of proteins (Lauren, 2003). Mammals have four LRRTM genes, LRRTM1-4. The extracellular regions of LRRTMs are composed of ten tandem LRR domains flanked on either side by disulfide linked capping motifs, followed by a transmembrane domain, and an intracellular domain that contains a putative PDZ-binding domain that can interact with MAGUKs and localizes LRRTM1 and 2 to excitatory postsynaptic sites  (Linhoff et al., 2009). Through their LRR domains, LRRTMs bind to certain Nrxn splice variants on the same interacting face as Nlgns (Siddiqui et al., 2010). LRRTM family members are differentially expressed in various brain regions and show laminar specificity in both hippocampus and cortex. Moreover, their expression is developmentally regulated, with LRRTM2-4 expression beginning in late embryonic stages in mouse, peaking around postnatal day 14, and persisting through adulthood.  1.3.2 Mechanisms of synapse organization Molecular cell biology Classic work by Scheiffele et al. was the first to suggest that, acting through Nrxns, Nlgns are able to directly induce synapse formation (Scheiffele et al., 2000). This was achieved by developing what has come to be known as the co-culture assay. In this assay, non-neuronal cells, such as HEK or COS-7 cells, are transiently transfected with DNA so that they express a protein of interest, usually on their plasma membrane. These heterologous cells are then dissociated and re-plated with developing neurons in culture. Under control conditions, a neuron coming in contact with a heterologous cell does not form a significant association with that cell. However, when Scheiffele et al. transfected HEK cells with Nlgn1 and presented these cells to pontine axons and cerebellar granule cells from dissociated mouse cerebellum, synaptic vesicles and functional neurotransmitter release sites were seen to  30 accumulate at these points of contact, known as hemisynapses since they lack the appropriate postsynaptic partner. Since both the extracellular domain of Nlgn was required for this phenomenon, and since application of soluble b-Nrxn abolished it, the authors speculated that cell adhesion mediated by Nrxn/Nlgn interaction was directly able to induce synapse formation. Subsequent studies by Graf et al. showed the converse: that presentation of Nrxns on heterologous cells to hippocampal neurons was able to induce postsynaptic specialization on contacting dendrites, including recruitment of postsynaptic scaffolds PSD-95 or gephyrin and neurotransmitter receptors NMDAR NR1 subunit or GABAR g2 subunit to excitatory or inhibitory synapses, respectively (Graf et al., 2004). The selectivity of excitatory vs. inhibitory clustering is dependent on Nrxn and Nlgn isoforms with respect to their localization and binding affinities (see below). The mechanisms of bidirectional synapse induction by Nrxns/Nlgns presumably rely largely on each protein’s intracellular interactions with other molecules. These interactions act to recruit critical pre- or postsynaptic components to synapses by trapping them at nascent or maturing contact sites, and many of these recruited molecules are known to affect intracellular signaling at the synapse. In axons, a key regulator of Nrxn-induced presynaptic differentiation is Ca2+/calmodulin-activated Ser-Thr kinase, CASK, which binds to the C-terminal PDZ-binding domain of Nrxn (Hata et al., 1996). CASK binds the proteins Velis, Mint-1, and protein 4.1 which, when in complex with Nrxn, acts as a nucleation site for actin at active zones (Butz et al., 1998; Biederer and Sudhof, 2001). Moreover, Mint-1 directly binds munc18, a component of active zones (Biederer and Sudhof, 2000), while Nrxns can recruit synaptic vesicles by their direct interaction with synaptotagmin (Hata et al., 1993). CASK is also able to bind to and influence clustering and physiology of presynaptic Ca2+ and K+ channels (Maximov et al., 1999; Atlas, 2001; Marble et al., 2005), potentially tuning synaptic vesicle release. At excitatory synapses, dendritic Nlgn-1 directly interacts  31 with both PSD-95 and S-SCAM (Irie et al., 1997; Hirao et al., 1998) through its C-terminal PDZ-binding domain. S-SCAM not only interacts with NMDARs and Nlgn-1 to regulate their clustering (Hirao et al., 1998; Iida et al., 2004), but may also be involved in cytoskeletal remodeling by interacting with guanine nucleotide exchange factors and other actin- associated proteins (Kawabe et al., 1999; Hirabayashi et al., 2004; Iida et al., 2007). Similarly, PSD-95 also directly interacts with NMDARs (Kornau et al., 1995) and may regulate the cytoskeleton through interaction with guanylate kinase domain-associated protein, GKAP, which in turn interacts with Shank and cortactin (Kuriu et al., 2006). In addition, PSD-95 interacts with K+ channels (Kim et al., 1995) and can control AMPAR trafficking through interaction with transmembrane AMPAR regulatory protein (TARP) family members like stargazin (Schnell et al., 2002a). At inhibitory synapses, dendritic Nlgn-2 directly interacts with gephyrin (Poulopoulos et al., 2009). In turn, gephyrin can bind and recruit various GABAAR subunits to target them to synaptic sites, including a1, a2, and a3 subunits (Tretter et al., 2008; Mukherjee et al., 2011; Tretter et al., 2011). Gephyrin may also control the cytoskeleton by binding collybistin, a guanine nucleotide exchange factor that acts on Cdc42 (Harvey et al., 2004), along with Profilin and Mena, actin binding proteins (Giesemann et al., 2003). Extending the co-culture approach to screen for cell adhesion proteins that could induce clustering of presynaptic vesicles, similar to Nlgns, Linhoff et al. found LRRTM1/2 had such an ability (Linhoff et al., 2009). Moreover, direct aggregation of LRRTMs on the dendrite surface was able to induce postsynaptic specialization, implying a bidirectional signaling complex involving LRRTMs for pre- and postsynaptic organization. Three independent groups ultimately discovered a mechanism by which LRRTMs induce presynaptic development is through their interaction with Nrxns (de Wit et al., 2009; Ko et al., 2009). Although far less is known of the intracellular signaling by LRRTMs, their C-  32 termini, like Nlgn-1, bind PSD-95, and could thus serve as nucleation sites for recruitment of postsynaptic components (de Wit et al., 2009; Linhoff et al., 2009).  A splice code for trans-synaptic interaction and signaling An emerging theme among Nrxn-Nlgn-LRRTM interactions in synaptic organization is the influence alternative splicing has on their interactions. As mentioned above, both Nrxns and Nlgns have a number of splice forms, including at sites that appear to be critical for their binding. Nrxn SS#4 is located in the sixth LNS domain of all a- and b-Nrxns; Nlgn SS#A is located in the AChE domain of all Nlgns, and SS#B is located in the same domain of Nlgn1 (Craig and Kang, 2007a). A number of recent studies have explored how alternative splicing affects both protein-protein interactions and synaptogenic activity of these proteins. Using binding and co-culture assays, it was found that b-Nrxns lacking the 30 amino acid insertion at SS#4 (-S4) interact with all Nlgns and LRRTM1/2 with high affinity (Boucard et al., 2005; Chih et al., 2006; Graf et al., 2006; Ko et al., 2009; Siddiqui et al., 2010). In contrast, LRRTMs do not significantly bind Nrxn1b(+S4), while all Nlgns do. Nlgn1 containing an insert at SS#B (+B) has marginally reduced affinity for Nrxn1b(+S4) but drastically reduced ability to cluster inhibitory presynaptic proteins (Levinson et al., 2005; Chih et al., 2006). The a-Nrxns interact with Nlgns(-B), while binding to Nlgn1(+B), the most strongly expressed isoform in adult brain, is abolished (Chih et al., 2006). As for b-Nrxn, binding of LRRTMs to a-Nrxn is abolished by inclusion of the splice site insertion at SS#4(Siddiqui et al., 2010). These molecular cell biological means of defining the differential binding patterns of Nrxn/Nlgn have recently been corroborated by crystallographic studies showing the structural requirements and constraints of these trans-synaptic interactions regulated by alternative splicing (Arac et al., 2007; Fabrichny et al., 2007; Chen et al., 2008; Comoletti et al., 2008; Koehnke et al., 2010; Leone et al., 2010; Tanaka et al., 2011).  33 1.3.3 Physiological roles of Nrxns, Nlgns, and LRRTMs In culture, overexpression of Nlgns induces formation of increased numbers of excitatory and inhibitory synapses that also have larger presynaptic terminals (Prange et al., 2004; Chih et al., 2005). Conversely, reduction of Nlgn-1,2,3 levels by RNA interference decreases both the amplitude and frequency of miniature postsynaptic currents, with a more profound effect on inhibitory synapses (Chih et al., 2005). This effect on inhibitory synapses is largely influenced by the levels of PSD-95, as its overexpression reduces inhibitory synapses but increases the formation of excitatory synapses (Prange et al., 2004), ostensibly by altering the distribution of Nlgn-2 from inhibitory to excitatory synapses (Levinson et al., 2005). It thus appears that the balance of excitation/inhibition is modulated by a complex interplay between Nrxn/Nlgn/PSD-95. Knockdown of LRRTM2 in vitro appears to affect only excitatory synapses, decreasing only the frequency of mEPSCs (de Wit et al., 2009). Nlgn-1 and LRRTMs are now thought to have redundant, overlapping roles in excitatory synapse development, as their concomitant reduction decreases excitatory synapse number further, but this effect is dependent on CaM kinases (Ko et al., 2011). It is thus clear that Nrxns/Nlgns/LRRTMs play essential roles in vitro for synapse development; however, their precise roles in vivo remain less obvious.  This is exemplified by the finding that triple knockout of Nlgns-1,2,3 in vivo does not change the number or morphology of synapses in neurons cultured from embryonic cortex or hippocampus (Varoqueaux et al., 2006). These mice, however, die shortly after birth due to respiratory failure, possibly a result of altered excitatory/inhibitory synapse numbers and synaptic transmission in brainstem. These data suggest that Nlgns are not critical for the initial stages of synapse formation. Knockout of LRRTM1 in vivo leads to increased size of excitatory synaptic vesicle clusters only in stratum radiatum and stratum oriens of CA1 of the hippocampus, but not other brain regions examined (Linhoff et al., 2009). Further LRRTM knockout mice, deleting more than just LRRTM1, may reveal stronger phenotypes.  34 Knockout of a-Nrxns has revealed severe reductions in N- and P/Q-type Ca2+ channel function that impair presynaptic neurotransmitter release, leading to perinatal lethality (Missler et al., 2003; Zhang et al., 2005). A modest decrease in the number of symmetric, inhibitory synapses was observed (Missler et al., 2003), although there was no apparent disruption of excitatory synapse number or morphology (Dudanova et al., 2007). b-Nrxn knockout mice have not been analyzed, leaving open questions as to how Nrxns function in synapse development in vivo. Recent data suggest that LRRTMs may play more prominent roles in synapse initiation while Nlgns may be more important in synapse maturation and maintenance (Soler-Llavina et al., 2011); however, a clear answer is not yet apparent, and will require extensive analysis of combinations of multiple Nlgn and LRRTM knockout mice.  The exact roles of these cell adhesion molecules in synapse organization may be further obscured by the contribution of the many other known synaptic organizing complexes, as described above. Thus, Nrxns/Nlgns/LRRTMs may function beyond initiation of synapse formation following cell-cell contact, playing a role in synapse stabilization, maturation, maintenance, or all of the above, or in specifying or validating which initial contacts persist. Indeed, while there may be interplay between Nlgn and LRRTM (Ko et al., 2011; Soler-Llavina et al., 2011), it appears also that N-cadherin, while not synaptogenic itself per se, is required for Nlgn-mediated synapse organization (Stan et al., 2010; Aiga et al., 2011). Further suggesting that Nlgn alone is insufficient for development of a full-fledged synapse, Nrxn-Nlgn interaction in vitro appears to recruit only functional NMDARs to nascent so-called “silent” synapses; only when glutamate is applied to the cells or CaMKII activated are AMPARs recruited and synapses unsilenced (Nam and Chen, 2005). Moreover, retrograde signaling by Nlgns through presynaptic Nrxns can lead to increases in vesicle release probability and changes in short-term synaptic plasticity (Futai et al., 2007). Recent studies have further implicated the Nrxn-Nlgn interaction in modulating behaviour. In  35 Aplysia, depletion of Nrxn or Nlgn serves to abolish sensitization of the gill-withdrawal reflex, a form of fear conditioning (Choi et al., 2011). Likewise, suppression of Nlgn-1 in the amygdala of adult rats does not impair basal synaptic transmission, but reduces NMDAR- mediated currents, impairs high-frequency stimulation-induced LTP, and diminishes retrieval of fear-conditioned memory (Kim et al., 2008). These findings all suggest roles of Nrxn/Nlgn beyond development as mediators of activity-dependent synaptic plasticity. It is becoming ever more apparent that in vitro assays have only begun to hint at the possible roles and mechanisms of action of Nrxn/Nlgn/LRRTM and that our understanding of their roles will greatly benefit from thorough physiological and behavioral examination of newly-generated transgenic mice.  1.4 Cellular mechanisms of excitatory synapse formation  1.4.1 Initial contact and filopodial dynamics Prior to synapse formation, axons and dendrites contact one another. A majority of these initial contacts are transitory, with the cells not forming a stable adhesion (Ziv and Smith, 1996; Okabe et al., 2001a; Niell et al., 2004). However, the contacts that are appropriately reinforced ultimately form functionally active synapses. Several mechanisms of initial axo- dendritic contacts that differentiate into synapses have been proposed. First, protrusions from incoming axonal growth cones can be stabilized at sites where they contact dendrites (Washbourne et al., 2002; Meyer and Smith, 2006). Alternatively, dendritic growth cones can seek out and be stabilized at nearby axons (Sabo et al., 2006). However, a majority of synapses in the CNS are formed en passant; that is, not by the contribution of growth cone- like structures. In these instances, filopodia arising from both axons and dendrites are thought to be the major sites of initial axo-dendritic contact (Saito et al., 1992; Dailey and Smith, 1996; Ziv and Smith, 1996; Fiala et al., 1998; Ahmari et al., 2000; Jontes et al., 2000; Washbourne et al., 2002; Portera-Cailliau et al., 2003; Niell et al., 2004; Gerrow et al.,  36 2006). Alternatively and to a lesser extent, contacts can form without the participation of either axonal or dendritic filopodia, instead forming directly on shafts (Friedman et al., 2000; Washbourne et al., 2002; Gerrow et al., 2006). It is thought that the multitude of transient contacts formed are only stabilized and/or mature into synapses when the appropriate complement of pre- and postsynaptic CAMs match up, although what determines a valid “match” remains esoteric and is of growing interest in the field of synapse specificity (Shen and Scheiffele, 2010). Ultimately, the core components of a synapse – presynaptic vesicles and release machinery apposed to postsynaptic neurotransmitter receptors – are themselves recruited to and consolidated at these stabilized contact sites (Fig. 1.6).  1.4.2 Transport of presynaptic proteins It is now widely accepted that vesicular delivery of presynaptic components is the primary means by which these proteins are transported through axons and arrive at synapses (Ziv and Garner, 2004). These presynaptic proteins localize to one of two kinds of so-called “transport packets”: piccolo transport vesicles (PTVs) and synaptic vesicle protein transport vesicles (STVs). PTVs are dense-core vesicles containing the active zone proteins piccolo and bassoon, along with other vesicle release machinery proteins like syntaxin, RIM, SNAP25, and N-type voltage-gated calcium channels (Zhai et al., 2001; Ohtsuka et al., 2002; Lee et al., 2003; Shapira et al., 2003). PTVs are transported in axons bidirectionally at rates up to 0.35 mm/s, and can split and merge into smaller or larger structures, respectively (Friedman et al., 2000; Bresler et al., 2004). STVs comprise various physical structures, ranging from tubulovesicular organelles to aggregates of small vesicles resembling synaptic vesicles (Kraszewski et al., 1995; Ahmari et al., 2000). They are composed of synaptic vesicle associated proteins that are distinct from those found in PTVs (Zhai et al., 2001). STVs are transported at rates of 0.1-1 mm/s (Kraszewski et al., 1995; Dai and Peng, 1996;  37 Nakata et al., 1998; Ahmari et al., 2000; Kaether et al., 2000; Sabo et al., 2006) and, like PTVs, can split and merge (Bresler et al., 2004). Both types of vesicles are transported by molecular motors: the anterograde microtubule-based kinesin family motors; the retrograde microtubule-based dyneins; and certain bidirectional actin-based myosin motors (Hirokawa et al., 2010). Recent studies have shown that synaptic vesicles can be shared among presynaptic boutons (Darcy et al., 2006; Staras et al., 2010). The physiological role of this vesicle “superpool” is unknown, and it remains to be seen whether these locally shared synaptic components can also contribute to development of presynaptic specializations. Additionally, stable synaptic vesicle recycling sites unapposed to postsynaptic specializations have been observed (Matteoli et al., 1992; Zakharenko et al., 1999). These orphan release sites are competent for neurotransmitter release, and have recently been shown to preferentially stabilize incoming dendritic growth cones (Sabo et al., 2006). It thus appears that specific sites along axons may contain “prototerminals” that are more amenable to synapse formation than other parts of the axon (Ziv and Garner, 2004).  1.4.3 Transport of postsynaptic proteins Disparity exists about the nature of delivery of postsynaptic proteins to nascent synaptic sites. Several studies have suggested that postsynaptic scaffold proteins like PSD-95 and Homer are trafficked to synapses early in their development through their gradual accumulation from either diffuse cytoplasmic pools or local delivery of only a few copies of the protein (Friedman et al., 2000; Bresler et al., 2001; Okabe et al., 2001b; Bresler et al., 2004). Alternatively, these scaffolds may be trafficked in discrete packets which may represent supramolecular complexes that co-transport PSD-95, GKAP, Shank, Nlgn-1, and NMDARs (Prange and Murphy, 2001; Wenthold et al., 2003; Gerrow et al., 2006). These trafficking complexes could thus represent organelles available for rapid synapse development following cell-cell contact, similar to presynaptic transport packets. In support  38 of this model and similar to orphan vesicle release sites, excitatory postsynaptic specializations lacking presynaptic input have been observed in young cultured neurons, although with low frequency (Rao et al., 1998). NMDARs have also been observed to both slowly accumulate (Bresler et al., 2001) or to be rapidly inserted at nascent synapses by vesicular delivery of transport packets observed to move > 6 mm/min in dendrites (Washbourne et al., 2002). Likewise, AMPARs may either be inserted haphazardly in the dendritic plasma membrane but subsequently trapped at synaptic sites (Borgdorff and Choquet, 2002) or delivered by local exocytosis at synapses, possibly dependent on subunit composition (Passafaro et al., 2001). As with presynaptic transport packets, molecular motors have been implicated in the trafficking of neurotransmitter receptor vesicles. These motor proteins interact with the receptors through various scaffold and chaperone proteins. For instance, NR2B-containing NMDARs are transported along microtubules by the kinesin motor KIF17, mediated through interactions with Mint1 and other proteins (Setou et al., 2000). AMPARs, on the other hand, are transported by kinesin motors through an interaction with GRIP1 (Setou et al., 2002). It remains unclear, however, whether these motor-dependent processes are critical for initial stages of synapse formation as opposed to being important for synapse maturation and plasticity. Another central postsynaptic signaling molecule, CaMKII, appears to be recruited to synapses from diffuse pools and may act as a sensor for neurotransmitter receptor activation at developing synaptic sites (Shen and Meyer, 1999; Rose et al., 2009). Although the two models of recruitment of postsynaptic proteins – accumulation from cytoplasmic or dendritic membrane pools versus local delivery by directed exocytosis – are not necessarily mutually exclusive, the contribution of either mode of trafficking to various proteins may differ, and, in turn, modify the kinetics of synapse assembly, thus requiring further investigation. Interestingly, recent studies have shown a number of postsynaptic proteins to be locally synthesized from their dendrite-targeted mRNA transcripts (Steward and Schuman, 2001; Hanus and Ehlers, 2008). The relative  39                         Figure 1.6. Multiple cellular mechanisms for CNS synaptogenesis. There appear to be multiple mechanisms for the recruitment and stabilization of pre- and postsynaptic proteins to new sites of axo-dendritic contact. (a) Glutamatergic synapses between axon and dendrite shafts of hippocampal neurons can form in about an hour after the initial accumulation of presynaptic vesicles. Presynaptic proteins, including synaptic vesicle precursors (STVs) and piccolo-transport vesicles (PTVs), are mobile in axons before synapses are formed (upper axon/dendrite pair). These precursors are the first proteins recruited to nascent synapses (second axon/dendrite pair). After 30 min, PSD-95 accumulates at these sites (third pair) followed by glutamate receptors (fourth pair). (b) In young cortical neurons, glutamatergic synapses can form even faster, on a timescale of several minutes. In these cells, STVs and NMDARs are both found in transport packets that are highly mobile in the axons and dendrites, respectively, before synapse formation (upper axon/dendrite pair). Both STVs and NMDAR transport packets cycle with the membrane during their transport. Contact between an axonal growth cone filopodium and a dendrite (right), or between axon and dendrite shafts (left), leads to the rapid and simultaneous recruitment of STVs and NMDARs at nascent synapses within 7 min of contact (second pair). PSD-95 is recruited to these sites with a variable time course, and AMPARs are recruited an hour following initial recruitment of NMDARs (third pair). (c) Glutamatergic synapses can also form at prespecified sites along dendritic shafts of hippocampal neurons, defined by stable preformed scaffold complexes associated with neuroligin. In this scenario, complexes of scaffolding proteins (including PSD-95, Shank, and GKAP) are mobile within dendrites before synapses are formed (upper axon/dendrite pair). When these complexes associate with neuroligin, they often become stabilized in the dendritic membrane (second pair). A significant proportion of these complexes then recruit STVs to form synapses within 2 h of their stabilization (third pair). (d) There are also predefined sites along the axon shaft of cortical neurons where en passant synapses selectively form. These predefined sites are stable locations along the axon where STVs cycle with the plasma membrane (first axon/dendrite pair) and presumably release diffusible molecules before synapses are formed (second pair). Filopodia from dendritic growth cones (right) and presumably also dendritic shafts (left) are selectively attracted to, and stabilized at, these sites (third pair). Following stabilization of this contact at this predefined site, the presynaptic terminal is formed and additional pre- and postsynaptic proteins are recruited to form a nascent synapse (fourth pair). Adapted with permission from McAllister, 2007.   40 contribution of newly-synthesized proteins in synapse development, though, remains to be studied.  1.4.4 Maturation of nascent synapses A number of live imaging studies have sought to determine the temporal order of recruitment of synaptic proteins to nascent sites. Several of these studies have suggested that presynaptic components are recruited to sites of axodendritic contact early. For example, Friedman et al. showed that sites where FM4-64, a dye marker of recycling synaptic vesicles, was stably present also recruited the active zone protein bassoon (Friedman et al., 2000). These active presynaptic terminals are thought to be recruited from PTVs and STVs being trafficked through the axon within 1-2 hours following contact initiation. Moreover, it appears that only a small number of both types of transport packets are recruited to these developing presynaptic sites, as their accumulation occurs rapidly in only a single step (Ahmari et al., 2000; Friedman et al., 2000; Shapira et al., 2003). This rapid recruitment of both PTVs and STVs to presynaptic sites obscures their precise order of accumulation, but new data suggest that these transport packets are trafficked together prior to synapse formation, potentially allowing for the assembly of functional presynaptic sites at a moment’s notice following cell adhesion (Bury and Sabo, 2011). However, transport packets may not be the only source of presynaptic components in new synapse formation, as both active zone proteins and synaptic vesicles from orphan release sites have been suggested to bud off from these locations and accumulate at presynaptic terminals (Krueger et al., 2003). In addition, these orphan release sites may themselves represent hot spots for synapse formation, stabilizing incoming contacting dendrites and requiring little modification to become fully mature presynaptic sites (Sabo et al., 2006).  Whereas presynaptic components are recruited soon after axo-dendritic contact, postsynaptic components appear to arrive later. Initial studies suggested that PSD-95 and  41 other scaffolding molecules are recruited 1-2 hours after axo-dendritic contact, between 20- 60 minutes after stable accumulation of recycling synaptic vesicles at those sites (Friedman et al., 2000; Okabe et al., 2001a). Both NMDARs and AMPARs are recruited either in concert with PSD-95 or within 10-20 minutes after its accumulation at nascent synapses (Friedman et al., 2000). In contract to presynaptic elements, which are recruited to new axonal boutons quantally, postsynaptic elements appear to accumulate gradually over tens of minutes, suggesting their recruitment either from a diffuse pool or from delivery by vesicles containing very few copies of each protein (Bresler et al., 2004). However, several recent studies have contradicted both the time course and mode of recruitment of postsynaptic components. For example, Washbourne et al. observed axonal growth cones to recruit NMDARs in contacting dendrites within less than 10 minutes of cell-cell contact, sometimes preceding vesicle accumulation at the apposed presynaptic site (Washbourne et al., 2002). These NMDARs are putatively delivered rapidly to synaptic sites through dendritic transport packets, whereas similar organelles were not observed for AMPARs, which accumulate slowly over roughly 1 hour following NMDAR recruitment (Washbourne et al., 2002). Additionally, a preformed complex of several postsynaptic elements, including PSD- 95, GKAP, Shank, and Nlgn-1, has been observed to be mobile in dendrites (Gerrow et al., 2006). In up to 30% of instances where these clustered elements were stationary but unapposed to presynaptic vesicle clusters, vesicles were subsequently recruited to these sites within 2 hours, suggesting postsynaptic development precedes presynaptic development in these instances. It thus appears there may be several modes of early synapse development, possibly regulated by one or many synapse organizing factors.  Following initial recruitment of the core components that make a functional synapse, synapses undergo a protracted period of maturation on both the pre- and postsynaptic sides. In developing cortex, the number of presynaptic vesicles per terminal increases up to threefold during the first postnatal month (Vaughn, 1989) with concomitant increase in  42 quantal size (Liu and Tsien, 1995; Mohrmann et al., 2003) and decrease in release probability (Bolshakov and Siegelbaum, 1995). Excitatory postsynaptic specializations undergo equally dramatic changes. While it appears that a majority of synapses initially form on highly motile and often transient dendritic filopodia, mature excitatory synapses are primarily located on more stable and shorter dendritic spines that develop directly from these filopodia (Ziv and Smith, 1996; Okabe et al., 2001a). Moreover, the morphology of spines can be further modified during normal development or in response to various stimuli in adulthood (Hering and Sheng, 2001; Bourne and Harris, 2008; Zito et al., 2009), and this is correlated with increases in postsynaptic density size (Harris et al., 1992). Glutamate receptor physiology of certain synapses also undergoes maturation. A significant proportion of synapses contain functional NMDARs without surface AMPARs and, because they do not elicit spikes at resting membrane potential, are termed “silent synapses” (Durand et al., 1996; Isaac et al., 1997; Nusser et al., 1998; Takumi et al., 1999). Activation of NMDARs can subsequently lead to synaptic insertion of AMPARs and unsilencing of these synapses. Additionally, there is a developmental switch in the proportion of NR2A- versus NR2B- containing synaptic NMDARs, with the former appearing later in development to partially replace NR2B-containing NMDARs and leading to decreased charge transfer (Tovar and Westbrook, 1999). Taken together, a growing body of evidence has begun to highlight the complex dynamic nature of synapse initiation and maturation.  1.4.5 The synaptotropic model of axon and dendrite growth In many different model systems and developing brain regions, synaptogenesis, dendritic growth and arborization, and axonal growth and appropriate innervation are concurrent processes. Numerous studies have suggested that each of these processes is activity- dependent (Rajan and Cline, 1998; Gomez and Spitzer, 1999; Wong and Wong, 2001; Knott et al., 2002; Lohmann et al., 2002; Niell et al., 2004; Hua et al., 2005; Haas et al., 2006;  43 Kerschensteiner et al., 2009), although certain activity-independent processes may persist (Rao and Craig, 1997; Verhage et al., 2000). The former observations, however, have led many to postulate that synapse formation is required for proper dendrite and axon development. This so-called synaptotropic hypothesis suggests that growing neuronal processes actively sample their environment for those subdomains that will best reinforce their growth, and that the likeliest places for their stabilization and reinforcement to occur is at a site where the neuron forms a synapse (Cline and Haas, 2008). Synapses are ideal sites of stabilization because, among other criteria, they are composed of adhesive cell contacts and are able to convey neurochemical information through receptor activation. Several lines of evidence have emerged to support the synaptotropic model. First, in vivo imaging of growing dendritic arbors has shown that filopodia, which typically are highly mobile and transient, are preferentially stabilized at sites where PSD-95 clusters, and that these filopodia are subsequently stabilized (Niell et al., 2004). PSD-95 accumulation also appears to stabilize filopodia for conversion into spines (Marrs et al., 2001; Prange and Murphy, 2001). A similar phenomenon is observed for growing axonal arbors, where stable accumulation of synaptophysin both appears to prevent retraction of axonal filopodia and to signal sites of new axon branch formation (Meyer and Smith, 2006; Ruthazer et al., 2006). The inference from these results is that synapse formation leads to stabilization of outgrowing processes that can subsequently continue to grow, whereas processes that are not stabilized retract and are eliminated. Further direct evidence for synapse formation being integral to dendrite arbor growth comes from the work of Chen et al. who showed that disruption of cell adhesive Nrxn-Nlgn interactions leads to both a reduction of synapse number and a destabilization of dendritic filopodia, ultimately leading to reduced dendrite arbor complexity (Chen et al., 2010). It thus appears that, at least locally, synapse formation is important for proper development of neuronal processes. However, it is likely that  44 additional activity-dependent processes that, perhaps, engage cell-wide transcriptional programs, are also required for sculpting the entire dendritic and axonal arbor.  1.5 Molecular and cellular mechanisms of inhibitory synapse formation and modification  Unlike the numerous findings that are arising for the development of excitatory synapses, far less is known about the equivalent cellular mechanisms of inhibitory synapse formation. For instance, only one report describes the initial contact between GABAergic axons and pyramidal neurons in the hippocampus, compared to the dozens of studies performed in various model systems and different brain regions to address how glutamatergic axons interact with dendrites. This preliminary report suggests that, in contrast to glutamatergic synapses, neither axonal nor dendritic filopodia participate in the early stages of cell-cell association (Wierenga et al., 2008). Additionally, the temporal order of recruitment of GABAergic synaptic components remains ill-defined, with researchers left to speculate on these mechanisms based on inferences from studies of glutamatergic synapses. However, a growing body of research has begun to define key aspects of GABAergic synapse modification, particularly with respect to GABAAR trafficking. Given the many parallels between initial synapse formation/maturation and plasticity, these findings may prove valuable informative tools for GABAergic synapse formation.  1.5.1 Assembly and forward trafficking of GABAARs The net surface complement of GABAARs can be viewed as the sum of numerous processes, including gene transcription in the nucleus and translation in the ER, oligomerization of receptor subunits and their exit from the ER and Golgi apparatus, exocytosis and insertion of receptors into the plasma membrane, and endocytosis of  45 membrane receptors and their post-endocytic recycling or degradation. Various GABAAR mRNAs have been shown to be dynamically regulated in both normal physiological states and in disease, and are thought to be directly coupled to the total surface expression of receptors (for review see (Luscher et al., 2011a). Following translation, GABAAR subunits assemble into heteropentamers in the ER. A detailed understanding of the mechanisms of subunit assembly in neurons has been complicated by the fact that many neurons often express a large number of different subunits. However, it appears that a and b subunits are required for proper exit from the ER, as genetic deletion of either subunit abolishes surface expression of GABAARs (Homanics et al., 1997; Kralic et al., 2002); in contrast, g2 knockout mice show largely normal receptor surface expression (Gunther et al., 1995). Assembled receptors are subsequently signaled to exit the ER through an interaction with PLIC-1 (Bedford et al., 2001). Importantly, it appears that PLIC-1, a ubiquitin-like protein, interferes with constitutive ubiquitin-dependent degradation of receptors (Saliba et al., 2007). Consequently, blockade of ubiquitination or overexpression of PLIC-1 enhances surface GABAAR levels (Bedford et al., 2001; Saliba et al., 2007). Concurrent with transfer from the ER to the Golgi, g2 subunits are palmitoylated by the enzymes GODZ or SERZ-b (Keller et al., 2004; Fang et al., 2006), a step required for correct assembly and forward trafficking of GABAARs. Both PLIC-1 and GODZ can be localized subsynaptically, suggesting that GABAARs are able to be assembled in membranous outposts in the dendrite, not just within the cell soma (Bedford et al., 2001; Keller et al., 2004). Several other mechanisms have been uncovered that regulate exocytosis of Golgi-derived GABAAR-containing vesicles to the plasma membrane. The guanine nucleotide exchange factor (GEF) BIG2 interacts with a motif in b subunits and activates the G-proteins ARF1 and 3, facilitating vesicle budding from the Golgi and GABAAR surface expression (Charych et al., 2004; Shin et al., 2004). The ubiquitin-like protein GABARAP interacts with g subunits, microtubules, and the SNARE  46 protein chaperone NSF (Wang et al., 1999; Nymann-Andersen et al., 2002). Although it does not appear to localize to synapses (Kneussel et al., 2000; Kittler et al., 2001), its overexpression leads to increased surface trafficking of GABAARs (Leil et al., 2004), presumably mediated by its interactions with the cytoskeleton and other exocytic proteins. Recent data from GABARAP knockout mice, though, showed no impairment of GABAergic synapse formation, calling into question its precise role in GABAAR trafficking (O'Sullivan et al., 2005). GABARAP also binds to PRIP1/2, an adaptor protein that can interact with kinases and phosphatases (Terunuma et al., 2004). Interestingly, in addition to binding to GABARAP, NSF and PRIP1/2 can also interact directly with b subunits (Kittler et al., 2004a; Terunuma et al., 2004; Goto et al., 2005). In turn, knockout of PRIP1/2 leads to decreased surface expression of GABAARs (Mizokami et al., 2007). Recently, the microtubule-based kinesin family motor protein KIF5 has been suggested to mediate transport of GABAARs to the plasma membrane (Twelvetrees et al., 2010). This trafficking is mediated by the adaptor huntingtin-associated protein 1 (HAP1), which binds both GABAARs and KIF5. Disruption of this motor-receptor complex leads to decreased surface levels of GABAARs. Moreover, using live cell imaging, the frequency and velocity of mobile GABAAR-containing intracellular vesicle-like structures was decreased. The contribution of vesicular transport of GABAARs to synapses, though, remains unclear, as a number of reports suggest the predominant method of recruitment of GABAARs to synapses is from extrasynaptic plasma membrane pools (see below).  In concert, these molecular steps lead to correct biogenesis and insertion of GABAARs on the dendrite surface. However, a number of other mechanisms are at play to govern the overall levels of synaptic GABAAR receptors and the efficacy of inhibitory phasic inhibition, including their regulated endocytosis and recycling and anchoring at synapses.    47 1.5.2 Regulation of GABAAR surface expression Under basal conditions in young cultured neurons, a large proportion, roughly 25%, of GABAARs are endocytosed every 30 minutes, destined for recycling back to the plasma membrane (70%) or degradation by lysosomes (30%) (Kittler et al., 2000; Kittler et al., 2004b). This constitutive endocytosis and recycling may underlie fine tuning of synaptic structure and function. Receptor internalization is mediated through interactions with AP2 in both the b and g subunits, which leads to clathrin- and dynamin-dependent endocytosis (Kittler et al., 2000; Herring et al., 2003). Endocytic and recycling processes are not simply constitutive, though, but may be regulated by several mechanisms. Numerous kinases can phosphorylate the AP2-binding site of b subunits, including protein kinase A (PKA) (McDonald et al., 1998), PKC (McDonald et al., 1998; Brandon et al., 2000), CaMKII (McDonald and Moss, 1994), and Akt (Wang et al., 2003), and thus phosphorylation of these sites leads to increased GABAAR surface levels. Conversely, dephosphorylation of b subunits can be performed by the protein phosphatases PP1a and PP2A (Terunuma et al., 2004; Kanematsu et al., 2007), which leads to decreased surface receptor expression. Similar to b subunits, phosphorylation sites exist in the AP2-binding site of g subunits (Kittler et al., 2008; Smith et al., 2008). Phosphorylation of these sites by the kinase Fyn (Jurd et al., 2010), leads to increased surface receptor expression. The question of how these phospho-dependent trafficking mechanisms are regulated upstream – namely what types of stimulation are required and which receptors are activated to elicit downstream events – remains unanswered. Additionally, whether these events can be triggered at a single synapses rather than cell-wide is unknown. Recently, activity-dependent mechanisms have also been shown to regulate exocytosis of GABAARs. Activation of NMDARs by chemical stimulation was found to increase GABAAR surface expression in a Ca2+/CaMKII-dependent manner, possibly through increasing receptor interaction with GABARAP and GRIP to drive  48 forward trafficking (Marsden et al., 2007).  Moreover, there is evidence that patterned stimulation that leads to activation of NMDARs can induce at least Akt-dependent GABAAR phosphorylation and its insertion at synaptic sites (Wang et al., 2003). Future studies into the modulation of exo- and endocytosis of GABAARs are of critical importance, as even small deficits in GABAAR surface expression can have severe behavioral effects (Crestani et al., 1999).  1.5.3 Synaptic anchoring of GABAARs Up to 75% of the total inhibitory charge transfer generated within dendrites is mediated by tonic inhibition through activation of extrasynaptic GABAARs responding to diffuse GABA in the extracellular space, largely due to the receptors’ high affinities for GABA and long channel open times (Mody and Pearce, 2004). What controls the localization of these receptors is poorly understood. On the other hand, there is a growing understanding of what controls the synaptic localization of GABAARs, which drive phasic inhibition and appear critically important for, among other things, modulation of excitatory synaptic strength and generation of intrinsic network oscillations that underlie certain behaviours (Farrant and Nusser, 2005). Depending on the type of synapse and GABAAR subunit composition, anywhere from 10-100 receptors may be localized opposite presynaptic terminals, representing a 50 to 130-fold enrichment of receptors at synapses compared to extrasynaptic membranes (Nusser et al., 1997; Tretter and Moss, 2008).  As described earlier (Section 1.1.3), synaptic GABAARs are primarily composed of a1-3, b2-3, and g2 (or 3) subunits, with differential brain regional expression and subcellular distribution based on subunit stoichiometry. This synaptic targeting is at least partially based on interactions with intracellular scaffold molecules. Each of a1-3 subunits have recently been shown to directly interact with gephyrin (Tretter et al., 2008; Mukherjee et al., 2011;  49 Tretter et al., 2011). Additionally, gephyrin directly binds to inhibitory glycine receptors (GlyR) (Langosch et al., 1992). Using live imaging of quantum dot labeled GlyRs in spinal cord, researchers found a significant increase in the mobility and extrasynaptic localization of GlyRs upon depletion of gephyrin (Meier et al., 2001). Moreover, the rate of synaptic accumulation of GlyRs is hastened upon interaction with gephyrin (Hanus et al., 2004), and gephyrin oligomerization is necessary for this process (Calamai et al., 2009), suggesting that diffuse membrane receptors are actively trapped at synapses by interaction with the postsynaptic scaffold. Alternatively, one report has suggested that GlyRs and gephyrin are co-transported to synapses as a single unit, often by budding off from another synapse (Maas et al., 2006). Such vesicular delivery is reminiscent of excitatory postsynaptic transport packets (Washbourne et al., 2002; Gerrow et al., 2006), but remains to be further validated. Recently, gephyrin-dependent regulation of GABAAR cell surface dynamics has been found, attributable to intracellular interaction with gephyrin (Jacob et al., 2005; Mukherjee et al., 2011). Interestingly, initial reports suggested the complete absence of postsynaptic GABAAR clustering in gephyrin knockout mice (Kneussel et al., 1999). However, further analysis showed much more modest effects on GABAergic synapses, where GABAAR clustering was dependent on those subunits analyzed (Kneussel et al., 2001; Levi et al., 2004). Indeed, acute knockdown of gephyrin in cultured neurons suggests that overall surface levels of GABAARs are unchanged, but their synaptic incorporation is, further suggesting synaptic accumulation of receptors by way of trapping of mobile receptors at scaffolds underlying synapses (Jacob et al., 2005; Yu et al., 2007). Further evidence for this model has come from electrophysiology experiments where GABA currents quickly recover following reversible inhibition, but recovery is abolished following irreversible GABAAR inhibition, suggesting a minimal contribution of newly exocytosed receptors to synaptic currents (Thomas et al., 2005).   50 1.5.4 Dynamic regulation of inhibitory synapses  There appear to be several further layers of complexity in GABAAR clustering at synapses. First, gephyrin itself is subject to regulation on several levels. Live imaging studies have revealed that it is mobile at synaptic sites, and this can be regulated by activity and the cytoskeleton (Hanus et al., 2006), although the implications for this are not fully understood. Conversely, gephyrin is able to regulate cytoskeletal dynamics through interactions with microtubules (Kirsch and Betz, 1995) and the actin cytoskeletal regulatory proteins profilin and Mena/VASP (Mammoto et al., 1998; Giesemann et al., 2003). Gephyrin clustering is also affected by phosphorylation mediated by glycogen synthase kinase 3b (GSK3b) (Tyagarajan et al., 2011). Disrupting this phosphorylation does not affect gephyrin cluster size, but does increase the number of clusters, and, in turn, GABAergic mIPSC frequency. Interestingly, mIPSC amplitude is also increased, suggesting incorporation of a greater number of GABAARs at synapses despite no change in gephyrin clustering. In addition, the intracellular gephyrin binding partner collybistin appears important for gephyrin and GABAAR clustering (Kins et al., 2000; Harvey et al., 2004). A GEF that activates Cdc42, collybistin exists in several splice forms. Splice forms that lack an N- terminal SH3 domain induce gephyrin and GlyR clustering at synapses in cultured neurons, and its genetic deletion greatly impairs both formation and maintenance of GABAergic synapses in CA1 of hippocampus (Papadopoulos et al., 2008). Interestingly, a majority of collybistin isoforms contain the SH3 domain, suggesting constitutive negative regulation of gephyrin clustering (Harvey et al., 2004). In a recent study, Poulopoulos et al. discovered that Nlgn-2 was a critical factor in mediating Collybistin-dependent gephyrin clustering and GABAergic synapse formation (Poulopoulos et al., 2009). Clustering of postsynaptic Nlgn-2 by presynaptic Nrxns leads to an initial nucleation of gephyrin through a conserved intracellular gephyrin interaction domain in Nlgn-2. Consequently, genetic deletion of Nlgn-2  51 leads to a decrease in the density of inhibitory synapses. This initial Nlgn-2/gephyrin clustering leads to activation of collybistin, which stabilizes gephyrin aggregates that subsequently anchor GABAARs at these nascent sites. Several unanswered questions remain from this model, though. First, Cdc42, the proposed downstream effector of collybistin activation, is not required for gephyrin clustering at synapses (Reddy-Alla et al., 2010). Moreover, the knockout of Nlgn-2 only leads to effects on inhibitory synapse formation at perisomatic sites in CA1 stratum radiatum (Poulopoulos et al., 2009), yet Nlgn-2 ablation leads to widespread deficits in inhibitory synaptic transmission in other structures like dentate gyrus, cortex, and retina (Chubykin et al., 2007; Hoon et al., 2009; Jedlicka et al., 2011). In somatosensory cortex, Nlgn-2 deletion only affects inhibitory transmission onto pyramidal cells from fast-spiking interneurons but not somatostatin-positive interneurons (Gibson et al., 2009). Finally, the GABAAR a2 subunit also appears to directly bind collybistin, preferentially co-localizes with collybistin in retina, and can “activate” the otherwise limiting SH3 domain of collybistin to induce gephyrin clustering, suggesting collybistin-dependent inhibitory synapse formation may be restricted to those sites with a2- containing GABAARs (Saiepour et al., 2010). Along with the trans-synaptic binding code for Nlgns/Nrxns, these GABAAR subunit dependent effects on their clustering indicate there is significant specificity in the formation of inhibitory synapses which remains to be elucidated. Finally, posttranslation modification of GABAARs themselves can regulate their mobility and affect inhibitory synaptic transmission. Whereas a number of proteins have been shown to phosphorylate GABAARs and enhance mIPSPs through receptor plasma membrane insertion, the phosphatase calcineurin depresses GABAergic mIPSPs, but not through receptor endocytosis (Lu et al., 2000). Rather, NMDAR activation causes dephosphorylation of a residue on the g2 subunit by calcineurin leads to an increase in GABAAR mobility, and, in particular, a dispersal of receptors from synaptic sites (Bannai et  52 al., 2009; Muir et al., 2010). This dispersal leads to a deficit in inhibitory transmission which, in turn, leads to an increase in excitation and facilitates LTP at excitatory synapses. To date, activation of GABAARs themselves has not been shown to directly modify their exo- /endocytosis, lateral mobility, or to modify the structure or function of other GABAergic postsynaptic components; rather, activation of adjacent excitatory synapses leads to changes in inhibitory synapses. It will be of particular interest in the future to investigate whether individual GABAergic synapses can be regulated in place by inhibitory neurotransmission.  1.6 Rationale and hypothesis Proper brain functioning requires appropriate connectivity between neurons and high fidelity synaptic transmission. The two main systems for rapid information transduction in the brain are excitatory glutamatergic and inhibitory GABAergic neurotransmission. It is becoming increasingly clear that dysfunction of either neurotransmitter system, either through defects in connectivity or synaptic function, are the root of many neurological and psychiatric diseases. A growing body of evidence is attempting to define precisely how synapses are formed, and a number of key molecular and cellular mechanisms have been identified that are important for the initiation, maintenance, and modification of synapses. Cell adhesion molecules have recently been implicated in synaptogenesis, and among these proteins are axonal neurexins and dendritic neuroligins. Their interaction at sites of cell-cell contact leads to the differentiation of functional presynaptic vesicle release machinery with apposed postsynaptic receptors and scaffolding molecules, defining a nascent synapse (Scheiffele et al., 2000; Graf et al., 2004; Nam and Chen, 2005). A number of genes and splice isoforms of both Nlgns and Nrxns exist, with the latter also being expressed as the shorter b-Nrxn or longer a-Nrxn form (Tabuchi and Sudhof, 2002).  53 Additionally, Nlgns are alternatively distributed, with Nlgn-1 localized to excitatory glutamatergic synapses and Nlgn-2 localized to inhibitory GABAergic synapses (Song et al., 1999; Graf et al., 2004). Initially, a-Nrxns were not known to interact with Nlgns (Ichtchenko et al., 1995), and alternative splicing of both b-Nrxns and Nlgns was shown to modulate their interaction and synaptogenic activity (Comoletti et al., 2003; Boucard et al., 2005; Chih et al., 2006; Graf et al., 2006). In Chapter 2 we sought to determine whether a-Nrxns were able to induce glutamatergic and GABAergic synapse formation through Nlgns, the contribution of alternative splicing to these interactions, and the developmental and activity- dependent regulation of a- and b-Nrxn expression so as to better understand the trans- synaptic code that defines proper apposition of pre- and postsynaptic components. It has also become increasingly clear that synaptogenesis is neither a unitary nor a protracted process, but instead is highly dynamic. It involves the contact of axons and dendrites, stabilization of these contacts when they are appropriate, recruitment of pre- and postsynaptic components necessary for synaptic transmission, and the functional and structural maturation of synapses. Several live imaging studies have sought to determine the cellular mechanisms of excitatory synapse formation. They have shown the importance of filopodia in initial synapse formation, that presynaptic components are recruited to nascent synapses by transport packets, this process generally precedes postsynaptic development, and postsynaptic components may be recruited to synapses by gradual accumulation from diffuse pools or from transport packets (for review see (Ziv and Garner, 2004; Waites et al., 2005; McAllister, 2007)). Live imaging studies have also revealed the short-term dynamic nature of pre-existing inhibitory synapses (for review see (Luscher et al., 2011a)). No studies exist, however, that describe the cellular mechanisms of inhibitory synapse development. In Chapter 3 we sought to characterize the dynamic nature of inhibitory synaptogenesis by using long-term live cell fluorescence microscopy to visualize  54 the spatiotemporal development of pre- and postsynaptic GABAergic synapse components so as to better understand the fundamental similarities and differences between glutamatergic and GABAergic synapse formation.                   55 2  Induction of GABAergic postsynaptic differentiation by a- neurexins1  2.1 Introduction  Synaptic connections in the brain require precise alignment of neurotransmitter receptors on dendrites opposite transmitter release sites on axons. Neurexin and neuroligin cell adhesion molecules are thought to function in development and maintenance of the two main synapse types, excitatory glutamatergic and inhibitory GABAergic synapses (Dean and Dresbach, 2006; Craig and Kang, 2007b). Neuroligins, normally present on dendrites, alone are sufficient to induce presynaptic differentiation when presented to axons of cultured neurons (Scheiffele et al., 2000; Fu et al., 2003; Chubykin et al., 2005). Conversely, neurexins, normally present on axons, alone are sufficient to induce postsynaptic differentiation when presented to dendrites of cultured neurons (Graf et al., 2004; Nam and Chen, 2005). These coculture studies suggest that neurexins and neuroligins play some role in synapse development in vivo, perhaps in protein recruitment and stabilization of synaptic complexes. There are 4-5 neuroligins in mammals. Neuroligin 1 localizes primarily at glutamatergic synapses (Song et al., 1999), and neuroligin 2 localizes primarily at GABAergic synapses (Graf et al., 2004; Varoqueaux et al., 2004). Six main neurexin isoforms are derived from three genes (1-3) and two promoters each (α and β) (Tabuchi and Sudhof, 2002). The shorter β-neurexins bind neuroligins via their single LNS domain (Ichtchenko et al., 1995). Alternative splicing at multiple sites also contributes to neurexin and neuroligin diversity. In particular, the absence of the splice site 4 (S4) insert in β-  1 A version of this chapter has been previously published. Kang Y, Zhang X, Dobie F, Wu H, Craig AM (2007) Induction of GABAergic Postsynaptic Differentiation by a -Neurexins. J Biol Chem.   56 neurexins and presence of the B insert in neuroligin 1 selectively promotes function at glutamatergic synapses (Boucard et al., 2005; Chih et al., 2006; Comoletti et al., 2006; Graf et al., 2006). The importance of these protein families for human cognition is suggested by the linkage of rare mutations in neuroligins 3 and 4 to autism and mental retardation (Jamain et al., 2003; Laumonnier et al., 2004) and the association of variations in neurexin 1 with autism (Feng et al., 2006; Szatmari et al., 2007). Mice lacking the major neuroligins 1/2/3 exhibit defects in inhibitory and excitatory synaptic transmission and die within 24 h of birth (Varoqueaux et al., 2006). Selective loss of neuroligin 1 or 2 results in selective defects in glutamate or GABA synapses, respectively (Chubykin et al., 2007). Overexpression or knockdown of neuroligins in cultured neurons also affects synapse development, altering the function and ratio of excitatory and inhibitory synapses (Prange et al., 2004; Chih et al., 2005; Levinson and El-Husseini, 2005; Futai et al., 2007). A role for neuroligin in development of neuronal cholinergic synapses has also recently been found (Conroy et al., 2007). α-Neurexins terminate in the same LNS domain, transmembrane region, and intracellular region with PDZ domain binding site as β-neurexins but contain additionally five LNS domains and three epidermal growth factor (EGF)-like domains organized into three modules. Analysis of triple knock-out mice for α-neurexin 1/2/3, leaving expression of the β- neurexins intact, revealed a surprising function in coupling presynaptic calcium channels to synaptic vesicle exocytosis (Missler et al., 2003). α-Neurexins function in transmitter release linked to N- and P/Q-type calcium channels at central nervous system synapses, in calcium- triggered exocytosis of secretory granules in endocrine cells, and to a lesser degree contribute to efficient neuromuscular transmission (Zhang et al., 2005; Dudanova et al., 2006; Sons et al., 2006). Intracellularly, α- and β-neurexins bind CASK, Mint, syntenin, and synaptotagmin (Hata et al., 1993; Hata et al., 1996; Biederer and Sudhof, 2000; Grootjans et  57 al., 2000). Extracellularly, α-neurexins presumably have unique binding partners to explain the calcium channel coupling phenotype that is not shared with β-neurexins. Known extracellular binding partners of α-neurexins include dystroglycan (Sugita et al., 2001) and the secreted peptides neurexophilins (Missler et al., 1998), although the significance of these interactions is not well understood. Originally, it was reported that α-neurexins did not bind neuroligins (Ichtchenko et al., 1995), but while the current work was in progress it became clear that α-neurexins also bind some neuroligins (Boucard et al., 2005). Neurexin- 1α with or without the S4 insert binds to neuroligins 2 and 3 and the minor variant of neuroligin 1 lacking a B insert but not to the major variant of neuroligin 1 containing a B insert (Fig. 3 of Boucard et al., 2005). Neurexin 1α lacking the S4 insert was also reported to induce clustering of gephyrin and neuroligin 2 but not PSD-95 when presented on COS cells to dendrites of cultured hippocampal neurons (Chih et al., 2006). We set out here to characterize the activity of α-neurexins in COS cell neuron coculture assays; that is, to test the ability of α-neurexins to induce glutamatergic and/or GABAergic postsynaptic differentiation in comparison with β-neurexins. We further explore the structural basis for the difference in synapse-promoting activity of α-compared with β- neurexins and the regulated expression patterns of neurexins with an emphasis on regulation at the S4 splice site.  2.2 Materials and methods 2.2.1 Primary neuronal culture and COS cell coculture Dissociated primary hippocampal neuronal cultures were prepared from embryonic day 18 (E18) rats as previously described (Kaech and Banker, 2006). Hippocampi were dissociated by trypsinization and trituration. Dissociated neurons were plated onto poly-L- lysine-coated glass coverslips in 60-mm culture dishes at a density of 300,000 cells/dish and  58 cocultured over a monolayer of glia. After 2 days, cytosine arabinoside (5 μM) was added to neuron cultures to prevent the overgrowth of glia. Cultures were maintained in serum-free minimal essential medium with N2 supplements, 0.1% ovalbumin, and 1 mM pyruvate, with replacing ⅓ of the media per dish once per week. Neurons were treated with 100 μM 2- amino-5-phosphonopentanoic acid (APV, Research Biochemicals) beginning on day 7. COS-7 and HEK293 cells were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum and transfected with Lipofectamine 2000 (Invitrogen). The flat shape of COS cells was preferred for coculture compared with the rounder shape of HEK cells preferred for confocal analysis of surface association. Transfected COS cells were trypsinized 24 h after transfection, washed, and plated at 200,000 cells/dish onto neurons pre-grown for 8-12 days in vitro (DIV). After 24 h of coculture, cells were fixed for 15 min in warm phosphate-buffered saline with 4% paraformaldehyde and 4% sucrose and permeabilized with 0.25% Triton X-100. For experiments involving immunocytochemistry for neuroligins in the cocultures, cells were fixed in -20 °C methanol for 10 min.  2.2.2 Construction of expression vectors Neurexin-1β-cyan fluorescent protein (CFP) (+S4) was described previously . To generate neurexin-1α-CFP, the N-terminal portion of rat neurexin 1α was cloned by RT-PCR and joined with the C-terminal portion of the mouse cDNA (BC047146; Open Biosystems) at the internal BstEII site and inserted in-frame into the pECFP-N1 vector (Clontech). Neurexin 2α (AK129239) and neurexin 3α (BC060719) were first corrected for apparent errors and then cloned into pECFP-N1. For neurexin 2α, the Stratagene site-directed mutagenesis kit was used to restore CAG in place of a premature TAG at residue 1301. For neurexin 3α, the splice site 1 insert consisting of an apparent duplication of exons 2 and 3 was removed by an overlap PCR method. To generate neurexin 2β′ and 3β′, the LNS domain of neurexin 1β  59 (residues 84-261) was replaced with the equivalent residues of LNS6 of neurexin 2α or 3α, altering the junctional amino acids GT to EF and amino acids EV to ST to facilitate cloning. Neurexin 1αBC contained a deletion of residues 31-473, and neurexin 1αC contained a deletion of residues 31-899 (numbering according to (39), each with the addition of an extra LV at the junction. All neurexin variants used in this paper have the insert at the splice site 4 position.  2.2.3 Immunocytochemistry and imaging Fixed neuron/COS cell cocultures were blocked with 10% bovine serum albumen (30 min; 37 °C) and incubated with appropriate primary antibodies in phosphate-buffered saline with 3% bovine serum albumen (overnight; room temperature) and then with secondary antibodies (45 min; 37 °C). Coverslips were mounted in elvanol (Tris-HCl, glycerol, and polyvinyl alcohol with 2% 1,4-diazabicyclo(2,2,2)octane). For Figs. 2.1 and 2.2, cocultures were stained with anti-gephyrin (mAb7a, IgG1, 1:500; Synaptic Systems), anti-PSD-95 (6G6-1C9, IgG2a, 1:500; Affinity Bioreagents), and anti-synapsin (rabbit, 1:1000; Chemicon) followed by secondary antibodies conjugated to Alexa 488, Alexa 568, and Alexa 647 (Molecular Probes), respectively. For Fig. 2.4, cocultures were labeled for neuroligin-1/3/4 (4F9, IgG2a, 1:1000; Synaptic Systems) or neuroligin-2 (Graf et al., 2006, rabbit, 1:400) in the Alexa 568 channel and anti-synapsin (46.1, IgG1, 1:100; Synaptic Systems) in the Alexa 647 channel. For Fig. 2.5, cocultures were labeled for GABAA receptor γ2 (rabbit; 1:200; Alomone) in the Alexa 568 channel and glutamic acid decarboxylase (GAD) 65 (GAD6, IgG2A, 1:100; Developmental Studies Hybridoma Bank) in the Alexa 488 channel. For Fig. 2.6, cocultures were labeled for neuroligin-2, gephyrin, or PSD-95 in the Alexa 568 channel and anti-synapsin in the Alexa 488 channel. For  Fig. 2.11, cocultures were incubated live with anti-GABAR α2 (kind gift of J. M. Fritschy (Brunig et al., 2002)) or γ2 antibodies for 30 min in the neuronal medium plus 50 mM HEPES, pH 7.4, at room temperature, washed  60 extensively, fixed, permeabilized, and then incubated with anti-synapsin followed by secondary antibodies. Fluorescence and phase contrast images were captured with a Photometrics Sensys-cooled CCD camera mounted on a Zeiss Axioplan microscope with a 63× 1.4 numerical aperture oil objective using MetaMorph imaging software (Molecular Devices) and customized filter sets. Controls lacking specific antibodies confirmed no detectable bleed-through between channels CFP, Alexa 488 (imaged through a yellow fluorescent protein filter set), Alexa 568, and Alexa 647. Images were acquired as grey scale from individual channels, and pseudo-color overlays prepared using Adobe Photoshop software. For quantification, sets of cells were cocultured and stained simultaneously. Images of transfected COS cells showing extensive contact with dendrites were taken in all fluorescence channels using the same exposure time for all constructs. For Fig. 2.12, optical sections were captured on an Olympus Fluoview FV500 confocal on a BX61W microscope with a 60× 1.4 numerical aperture oil objective, 442-nm laser, and customized filter set.  2.2.4 Image analysis For quantification for Figs. 2.3 and 2.4, images were randomized so that the experimenter was blind to the transfection group. The area for measuring was defined by the perimeter of the transfected COS cell. Images of the presynaptic and postsynaptic proteins were thresholded. For each postsynaptic protein cluster, a region was drawn around each cluster, and the area and total gray value was measured. Thresholded synapsin was measured through postsynaptic protein regions to determine which clusters were synaptic. Postsynaptic protein clusters that were apposed to synapsin were excluded from the final quantification. Analysis was performed using MetaMorph and Microsoft Excel. All data are reported as mean ± S.E. For quantification for Fig. 2.6, cocultures were assessed blind to the transfection group. On each coverslip all neurexin-CFP-transfected COS cells having significant contact  61 with neuronal dendrites based on phase contrast were assessed (up to 40 cells/coverslip). The area associated with each chosen COS cell was then visually scored as either positive or negative, positive indicating the presence of any clusters of postsynaptic protein (PSD-95, gephyrin, or neuroligin 2) that lacked associated staining for synapsin.  2.2.5 RNA isolation and RT-PCR For RNA extraction from brain, either two hippocampi or roughly 50 mg of cortex was dissected from rats at embryonic day 18 (E18), postnatal day 11 (P11), or adult and rapidly homogenized in 1 ml of Trizol reagent (Invitrogen), and RNA was prepared according to the manufacturer's protocol. For RNA extraction from hippocampal cultures, roughly 1 × 106 cells that were either untreated or chronically treated with 100 μM APV were harvested by briefly washing in cold phosphate-buffered saline and either scraping directly in Trizol reagent or by using the RNeasy mini kit (Qiagen) according to the manufacturer's protocol. First-strand cDNA synthesis was performed on 3 μg of total RNA for brain tissue samples or 300 ng of total RNA for hippocampal culture samples in a total volume of 50 μL using 500 units of SuperScript III reverse transcriptase (Invitrogen) and oligo(dT) primer. Extension of cDNA was performed at 50 °C for 60 min. The cDNA was then treated with 4 units of ribonuclease H at 37 °C for 30 min. PCR amplification was carried out on 0.5 μL of each above cDNA template in a total volume of 20 μL using 1 unit of Taq DNA Polymerase (Invitrogen). Thermal cycling parameters were denaturation at 95 °C for 30 s, annealing at 57 °C for 30 s, extension at 72 °C for 40 s. PCR was performed for 25-40 cycles, and the entire reaction was loaded on 2% agarose gel for electrophoresis. After staining and visualization with ethidium bromide, semiquantitation of band intensities was performed using ImageJ software. Two independent RNA preparations were made from independent animals or hippocampal cultures, and RT-PCR was performed in triplicate. All data are reported as the mean ± S.E.  62 Primers for each neurexin isoform were designed to include one reverse priming site downstream of splice site 4 that would be common to both the α and β isoforms and a unique forward priming site upstream of splice site 4 that would be exclusive to either the α or β isoform. The primers used in this study were (5′ to 3′): Nrxn1αF, ATGGGATGGCTTTAGCTGTG; Nrxn1βF, GGTCACCAGCATCCTTGC; Nrxn1R, CCCGCCAATTATTATGGTTGC; Nrxn2αF, ACTTCCTATGGAGGCCCTGT; Nrxn2βF, CCACCACGTCCACCACTT; Nrxn2R, TGGCTGTTGAAGATGGTCAG; Nrxn3αF, ACGATGCTCTCCACAGGAGT; Nrxn3βF, TCTACACCTGGCCAGCAAAT; Nrxn3R, AGAGAGTTGGCCTTGGAAGA; GAPDH forward, CCCTTCATTGACCTCAAC TACATG; GAPDH reverse, TGGTGAAGACGCCAGTAGACTC. For neurexins 1 and 2, a closely spaced doublet PCR product was obtained for the +S4 PCR product even using cloned +S4 cDNA template; thus, both bands were included in quantitation.  2.3 Results 2.3.1 Specific induction of GABAergic postsynaptic differentiation by α-neurexins  We determined here the synaptogenic activity of neurexin 1α, 2α, and 3α in COS cell neuron coculture (Figs. 2.1 and Fig. 2.9). The sixth LNS domain of α-neurexins, in common with β- neurexins, mediates binding to specific neuroligins (Boucard et al., 2005). Thus, for comparison, we assayed in the same experiments the sixth LNS domain of neurexin 1α, 2α, and 3α each in the context of the neurexin 1β-flanking sequences, corresponding to neurexin 1β itself, and constructs termed neurexin 2β′ and neurexin 3β′ (similar to neurexins 2β and 3β but with the regions flanking the LNS domain derived from 1β; Figs. 2.2 and Fig. 2.10). We expressed the neurexin variants tagged intracellularly with CFP in COS cells,    63 Figure 2.1. α-Neurexins induce clustering of the inhibitory synaptic scaffolding protein gephyrin but not the excitatory synaptic scaffolding protein PSD-95. A. COS cells expressing the neurexin variants tagged intracellularly with CFP were overlaid on hippocampal neurons pre-grown for 8-10 days in culture. After 1 day of coculture, cells were fixed and immunolabeled for PSD- 95, gephyrin, and synapsin; overlays are shown of PSD- 95 (green) plus synapsin (blue) or gephyrin (red) plus synapsin (blue). Phase contrast images (top row) show the positions of transfected COS cell nuclei (C), neuronal processes, and occasional neuron cell bodies (N). Endogenous interneuronal synapses have apposed clusters of postsynaptic PSD-95 or gephyrin and presynaptic synapsin; thus, clusters appear turquoise or purple, respectively, in the color overlays (arrows). In contrast, clusters of gephyrin lacking associated synapsin appear red in the color overlay and were induced by all α-neurexins (arrowheads). By alignment with the phase contrast images, induced clusters of gephyrin can be seen corresponding to sites of neuronal process contact with the neurexin-expressing COS cells and were frequently but not always localized near the edges of transfected COS cells, presumably where the COS cells come into closest contact with the neighboring neurons. Clusters of PSD-95 lacking associated synapsin were not induced by any α-neurexins. B. In sister coculture experiments, control COS cells expressing mCFP did not induce clustering of either PSD-95 or gephyrin. All neurexin constructs used in this paper contain the splice site 4 insert. See also Fig. 2.9 for single channel larger field of view and phase contrast images corresponding to the cropped field shown here for neurexin 2α-CFP. Scale bar, 10 μm.   cocultured these with hippocampal neurons, and assayed for ability to induce postsynaptic differentiation (as in Graf et al., 2004 and Graf et al., 2006). The ability of each neurexin to induce clustering of the inhibitory postsynaptic scaffolding protein gephyrin or the excitatory postsynaptic scaffolding protein PSD-95 in contacting dendrites was determined. Co- labeling for the presynaptic marker synapsin was used to identify the few endogenous  64 interneuronal synaptic clusters that may underlie the transfected COS cells since these bona fide synapses label for PSD-95 or gephyrin plus synapsin. Additional clusters of PSD- 95 or gephyrin lacking apposed synapsin immunofluorescence were specifically associated with COS cells expressing neurexins and are referred to as “induced clusters” throughout this paper. Such non-synaptic clusters of PSD-95 or gephyrin were not observed associated with control COS cells expressing membrane-associated CFP (mCFP; Fig. 2.1B). Induced clusters of PSD-95 or gephyrin were frequently but not always localized near the edges of neurexin-expressing COS cells, presumably where the COS cells come into closest contact with the neighboring neurons. Most often, the inducing neurexin was highly expressed over the entire COS cell and not visibly clustered, although rarely neurexin clusters on the COS cell were observed associated with dendritic protein clusters (e.g. Fig. 3.3 of Graf et al. 2004). All three α-neurexins induced prominent clustering of gephyrin but no detectable clustering of PSD-95 in contacting dendrites (Figs. 2.1 and 2.9). In contrast, like neurexin 1β (Graf et al., 2006), neurexin 2β′ and 3β′ induced prominent clustering of both gephyrin and PSD-95 (Figs. 2.2 and Fig. 2.10). Although the induced clusters of gephyrin and PSD-95 appear to overlap at low resolution, at higher resolution they usually resolve into separate clusters side by side (Fig. 2.10 and Graf et al., 2004). Quantitation of clusters of PSD-95 or gephyrin lacking synapsin was performed on random sets of COS cells expressing each neurexin variant in comparison with the base-line clusters associated with COS cells expressing only mCFP. This quantitation confirmed a significant induction of PSD-95 clusters only by the three β-neurexin constructs but none of the α-neurexins and induction of gephyrin clusters by all neurexin variants tested (Fig. 2.3, p < 0.05 or p < 0.005 as indicated, n = 32-35 cells each). For induction of gephyrin clustering, the α-neurexins exhibited weaker activity than the β-neurexin constructs but were still 2-7-fold above the background of mCFP. In this same set of experiments showing differences in the ability to induce  65 postsynaptic protein clustering, expression levels per COS cells did not differ among constructs (average relative CFP intensity values for each neurexin were 92 for 1α-CFP, 103 for 1β-CFP, 85 for 2α-CFP, 97 for 2β′-CFP, 113 for 3α-CFP, and 110 for 3β′-CFP; ANOVA p > 0.1). Figure 2.2. β- Neurexins induce clustering of the inhibitory synaptic scaffolding protein gephyrin and the excitatory synaptic scaffolding protein PSD-95. COS cells expressing the neurexin variants tagged intracellularly with CFP were overlaid on hippocampal neurons pre-grown for 8-10 days in culture. After 1 day of coculture, cells were fixed and immunolabeled for PSD-95, gephyrin, and synapsin; full field overlays are shown of PSD-95 (green) plus synapsin (blue) or gephyrin (red) plus synapsin (blue). Endogenous inter- neuronal synapses have apposed clusters of postsynaptic PSD-95 or gephyrin and presynaptic synapsin; thus, clusters appear turquoise or purple, respectively, in the color overlays (arrows). In contrast, clusters of PSD-95 or gephyrin lacking associated synapsin appear green or red, respectively, in the color overlays and were induced by all β-neurexins (arrowheads). By alignment with the phase contrast images, induced clusters of PSD-95 and gephyrin can be seen corresponding to sites of neuronal process contact with the neurexin-expressing COS cells and were frequently but not always localized near the edges of transfected COS cells, presumably where the COS cells come into closest contact with the neighboring neurons. Although the induced clusters of gephyrin and PSD-95 appear to overlap at low resolution, at higher resolution they usually resolve into separate clusters side by side (enlarged regions at the bottom of triple overlay of PSD-95 (green), gephyrin (red), and synapsin (blue)). All neurexin constructs used in this paper contain the splice site 4 insert. See also Fig. 2.10 for single channel larger field of view and phase contrast images corresponding to the cropped fields shown here for neurexin 2β-CFP. Scale bar, 10 μm.   66 The same type of coculture assay can be used to test the recruitment of neuroligin 1/3/4 (using an antibody that recognizes a common epitope) compared with neuroligin 2 to contact sites of dendrites with neurexin-expressing COS cells. Neuroligin clusters induced by neurexin contact were again differentiated from endogenous interneuronal synaptic clusters by the absence of associated synapsin immunoreactivity. We again observed a differential activity between α compared with β neurexins. Although the area of induced clusters of the neuroligins tends to be larger than that of gephyrin or PSD-95, covering a larger contact zone rather than being split into clusters more typical of a synaptic size ((Craig et al., 2006) and compare Figs. 2.1, 2.2, and 2.4), we could detect no significant clustering of neuroligin 1/3/4 by the α-neurexins (Fig. 2.4). All three α-neurexins induced prominent clustering of neuroligin 2 but not neuroligin 1/3/4 in contacting dendrites. In contrast, like neurexin 1β (Graf et al., 2006), neurexin 2β′ and 3β′ induced prominent clustering of both neuroligin 2 and neuroligin 1/3/4. Although this neuroligin 1/3/4 antibody recognizes all three neuroligins(Bolliger et al., 2001), neuroligins 1 and 3 are present at similar levels, but neuroligin 4 is not detectable in neonatal rodent tissue (Varoqueaux et al., 2006). Furthermore, neuroligin 1 is mainly expressed in the +B splice form (Chih et al., 2006), and neuroligin 1 +B binds neurexin 1β with 2-6-fold greater affinity than neuroligin 3 (Comoletti et al., 2006). Thus, the major variant clustered by β-neurexins but not α- neurexins and recognized by the neuroligin 1/3/4 antibody in this study is likely neuroligin 1 +B. Finally, we tested for the ability of each neurexin construct to induce clustering of the two other major components of GABAergic synapses, GABAA receptors and dystroglycan. The major synaptic GABAA receptor subunit γ2 (Essrich et al., 1998), like gephyrin, was clustered by all α-neurexins and β-neurexins tested. One example of induced clustering of GABAA receptor γ2 is shown for neurexin 2α in Fig. 2.5. Qualitatively, the relative clustering ability of the different constructs for GABA receptor γ2 appeared to parallel that for gephyrin  67 (data not shown). Unlike GABAA receptor γ2, which is mainly synaptic, GABAA receptor α5 subunit is mainly extrasynaptic, contributing to tonic GABAergic signaling (Brunig et al., 2002; Caraiscos et al., 2004). In a coculture experiment of neurons with COS cells expressing neurexin 2α-CFP, immunostaining of sister coverslips revealed obvious induced nonsynaptic clustering of GABAA receptor γ2 but no detectable clustering of GABAA receptor α5 (Fig. 2.11). As previously described, endogenous GABAA receptor α5 was detected more diffusely along dendrites compared with γ2, which was clustered at synapses. Some inhomogeneity in α5 immunoreactivity was observed along many dendrites, but at sites unrelated to COS cell contacts, and strong clusters were never observed. Thus, the effects of neurexins are selective for clustering GABAA receptor γ2 but not α5 subunit. Because Fig. 2.11 was performed with live cell primary antibody incubation, this also shows that the α- neurexin-induced clusters of GABAA receptor γ2 are on the dendrite surface. Dystroglycan is present at only a subset of mature GABA synapses (Knuesel et al., 1999; Levi et al., 2002) but was also reported to bind directly to α-neurexins (Sugita et al., 2001). However, in the coculture assay we did not observe clear induced clustering of dystroglycan by any neurexin, α or β, on neurons at 2-4 weeks in culture.  Figure 2.3. Quantitation of induced clustering of gephyrin but not PSD-95 by α-neurexins. Quantitation from random transfected COS cells was performed to assess the total integrated intensity of associated PSD-95 or gephyrin clusters that did not overlap with synapsin; values were normalized to the integrated intensity associated with COS cells expressing membrane-targeted CFP in sister cultures (gray lines at 1). This quantitation confirmed significant clustering of gephyrin by all neurexin variants tested but of PSD-95 only for the β-neurexin constructs and not the α-neurexins (p < 0.005 (**) and p < 0.05 (*) compared with mCFP by t test; n = 32-35 cells per construct combined from 2 independent cultures).  68 2.3.2 Structural basis for the differential synaptogenic activity of α-neurexins versus β-neurexins  The C-terminal region of neurexin-1α is identical to that of neurexin-1β from LNS6 on, and yet we show here that the synapse-promoting activity of neurexin 1α is weaker and more specific for GABAergic postsynaptic proteins than that of neurexin 1β in the coculture assay. Previous work has shown that a construct containing only the region common to neurexin 1α and 1β (deleting the β-specific sequence from neurexin 1β) had postsynaptic clustering activity equal to that of neurexin 1β (Graf et al., 2004). Thus, the difference in synaptogenic activity between neurexin 1α and 1β is due to an inhibitory effect of α-specific sequences rather than an enhancing effect of β-specific sequences. To explore the basis of this differential activity, we made sequential deletions from the mature N terminus of neurexin 1α, leaving the signal sequence for proper surface expression (Fig. 2.6). All deletion and mutant neurexin-CFP constructs assayed in this paper appeared to reach the cell surface as efficiently as neurexin 1α and neurexin 1β, as assayed by confocal optical sections through transfected HEK293 cells (Fig. 2.12). Furthermore, expression levels did not vary significantly among neurexin-CFP constructs in the COS cell expression studies (average relative CFP intensity values for each neurexin were 111 for 1α-CFP, 95 for 1αBC-CFP, 102 for 1αC-CFP, 103 for 1αD1176A-CFP, and 89 for 1β-CFP; n = 15-21, ANOVA p > 0.1). Deleting module A containing LNS1, LNS2, and the intervening EGF-like domain had no effect; the ability of this construct termed neurexin 1αBC to cluster postsynaptic proteins was indistinguishable from that of full-length neurexin 1α (Fig. 2.6, A and B). In contrast, further deleting module B containing LNS3, LNS4, and the intervening EGF-like domain resulted in increased synapse promoting activity. The ability of this construct, termed neurexin 1αC, to cluster GABAergic postsynaptic proteins gephyrin and neuroligin 2 was intermediate between that of neurexin-1α and neurexin-1β. However,  69 Figure 2.4. α-Neurexins induce clustering of neuroligin 2 but not neuroligin 1/3/4. A. COS cells expressing the neurexin variants tagged intracellularly with CFP (blue) were overlaid on hippocampal neurons pre-grown for 10-12 days in culture. After 1 day of coculture cells were fixed and immunolabeled for neuroligin 1/3/4 (with an antibody that recognizes all three) or neuroligin 2 (red) and synapsin (green). Whereas the β-neurexin constructs induced clustering of neuroligin 1/3/4 and neuroligin 2 in contacting dendrites (red or pink not associated with green), the α-neurexins induced clustering of neuroligin 2 but not neuroligin 1/3/4. Scale bar, 10 μm. B, cocultures were scanned, and the COS cells with the strongest apparent associated clusters were imaged and quantitated. The total integrated intensity of associated neuroligin that did not overlap with synapsin was normalized to a value of 100 for neurexin 1β. Significant differences among constructs were found (p < 0.001 ANOVA; n = 4-11 cells per construct).   70 Figure 2.5. α-Neurexins induce clustering of GABAA receptorγ2 subunit. A COS cell expressing neurexin-2α- CFP (blue) is shown here in contact with two major hippocampal neuron dendrites. Clustering of the GABAA receptor γ2 subunit (red) in the absence of glutamic acid decarboxylase (GAD; green) is prominent where the dendrites contact the neurexin-2α-CFP- expressing COS cell (pink). In contrast, the endogenous synaptic clusters of GABARγ2 at axon-dendrite contacts are associated with glutamic acid decarboxylase-positive GABAergic input (yellow). Similar induced clustering of GABARγ2 in COS cell-neuron coculture was observed for the other α and β neurexin constructs (not shown). Scale bar, 10 μm.  although deletion of modules A and B together significantly increased clustering of gephyrin and neuroligin 2 compared with full-length neurexin 1α, it did not increase clustering of PSD- 95 (Fig. 2.6). Thus, LNS5 and/or the third EGF-like domain within module C apparently inhibit the ability of neurexin 1 to cluster PSD-95, contributing synapse-type specificity. Boucard et al. (Boucard et al., 2005) found that LNS6 was the only LNS domain of neurexin 1α able to bind neuroligin when tested individually or in modules. We showed previously that a single point mutation to a predicted calcium binding residue D137A of this LNS domain in the context of neurexin 1β abolishes binding to neuroligins and abolishes synaptogenic activity (Graf et al., 2006). We, thus, tested whether this single point mutation in LNS6 in the context of full-length neurexin (i.e. 1α D1176A) affects synaptogenic activity. Indeed, neurexin 1α D1176A completely lacked the ability to cluster postsynaptic proteins (Fig. 2.6B). Thus, LNS6 is essential for mediating postsynaptic protein clustering in neurexin 1α, and the upstream sequences in modules B and C, regardless of additional LNS domains, inhibit rather than enhance the synaptogenic activity of neurexin 1α.  71  Figure 2.6. Postsynaptic protein clustering by neurexin-1αrequires LNS6 and is negatively modulated by upstream sequences. A, neurexin 1β, 1α, and derivatives of 1α mutated in a predicted calcium binding residue in LNS6 (1αD1176A), lacking module A (1αBC), or lacking modules A and B (1αC) were tagged intracellularly with CFP (blue) and tested for postsynaptic protein clustering activity in the COS cell-neuron coculture assay. Activity was considered positive if the contacting dendrites exhibited any clusters of PSD-95, gephyrin, or neuroligin 2 (red) lacking synapsin (green). Neurexin 1αD1176A had no detectable clustering activity. For PSD-95, all neurexin- 1α derivatives also had no detectable clustering activity; PSD-95 clustering was only observed in response to neurexin 1β (compare these images with Figs. 2.1, 2.2, 2.4). For gephyrin and neuroligin 2, the clustering activity of neurexin 1αBC was indistinguishable from that of 1α. In contrast, the clustering activity of neurexin 1αC was higher than that of 1α and intermediate between that of 1α and 1β. Scale bar, 10 μm. B, quantitation of the percentage of expressing COS cells exhibiting non- synaptic clusters in contacting dendrites confirmed these differences in relative synaptogenic activity (p < 0.001 by ANOVA; *, p < 0.001 by t test compared with neurexin 1α; n = 4 cultures with 10-78 cells scored each). PDZBD, PDZ binding domain; TM, transmembrane domain. CH, glycosylated region.   72 2.3.3 Expression patterns of neurexins Neurexin expression is brain-specific (Ushkaryov et al., 1992). At the cellular level the six major neurexin forms 1-3 α and β show fairly broad overlapping expression patterns in brain such that most neurons express multiple neurexins (Ullrich et al., 1995). How the expression patterns change with development has not been reported for the mammalian central nervous system. We addressed this question here with emphasis on the S4 insert given its importance in regulating the synaptogenic activity of β-neurexins (Boucard et al., 2005; Chih et al., 2006; Graf et al., 2006). By RT-PCR, we could readily detect all six neurexin forms 1-3 α and β from E18 to adult in rat hippocampus and cortex (Fig. 2.7A). There was a consistent increase in the percentage of all neurexins lacking the S4 insert compared with containing the S4 insert through development in both hippocampus and cortex. The difference in the ratio of splice variants was greatest for neurexin 3 α and β, for which the percent lacking the S4 insert changed from <5% at E18 to 35-50% at P11 (Fig. 2.7B). Neurons from E18 rat grown in culture for 7-22 DIV showed a similar trend with the percent of neurexins lacking the S4 insert increasing with development (Fig. 2.8). To assess potential activity regulation, we grew hippocampal neurons in culture under control conditions or chronically in the presence of the NMDA receptor antagonist APV. Semiquantitative RT-PCR in comparison to GAPDH revealed no significant difference in the total level of each neurexin mRNA (-S4 and +S4 forms combined) due to the presence or absence of APV (data not shown). However, blockade significantly increased the percentage of neurexin 2β lacking the S4 insert (from ∼20 to ∼40%) while reducing the percentage of neurexins 3α and 3β lacking the S4 insert (Fig. 2.8B). Thus, NMDA receptor activity differentially regulates splicing of neurexins at the S4 splice site.    73 Figure 2.7. Expression of neurexin splice variants lacking the S4 insert increases with development in hippocampus and cortex. The relative levels of neurexin variants containing (+) or lacking (-) inserts at splice site 4 was determined by RT-PCR from rat hippocampus and cortex at embryonic day 18 (E18), postnatal day 11 (P11), and adult (Ad). A, representative agarose gel electrophoresis of RT-PCR products. B, quantitative analysis revealed a significant increase in the percentage of each variant lacking the S4 insert between E18 and P11. Statistical analyses represent differences between P11 and other groups as determined by ANOVA with Tukey's post hoc test; *, p < 0.05; †, p < 0.005; n = 6.  Figure 2.8. Alternative splicing at the S4 site is regulated by NMDA receptor activity in hippocampal cultures. Neurons from E18 rat hippocampus were cultured in the chronic presence (+) or absence (-) of the NMDA receptor antagonist APV. Cultures were harvested at the indicated ages, and RT-PCR was performed to assess the relative levels of neurexin variants containing or lacking inserts at splice site 4. A, representative agarose gel electrophoresis of RT-PCR products. G, GAPDH loading control. DIV, days in vitro. B, quantitative analysis revealed a significant increase in the percentage of neurexin 2β and decrease in the percentage of neurexin 3α and 3β lacking the S4 insert with chronic NMDA receptor blockade. Statistical analyses represent differences between untreated cells and age-matched APV-treated cells as determined by ANOVA with Tukey's post hoc test; *, p < 0.05; †, p < 0.005; n = 6. Semiquantitative RT-PCR in comparison with GAPDH did not reveal any significant differences in total expression level of neurexin 1α, 1β, 2α, 2β, 3α, or 3β (+ and - S4 forms combined) between control cells or cells chronically treated with APV (data not shown).   74 Figure 2.9. Neurexin 2α induces clustering of the inhibitory synaptic scaffolding protein gephyrin but not the excitatory synaptic scaffolding protein PSD-95. Larger field of view single channel and additional overlay images are shown to more clearly illustrate the results of Fig. 2.1 (with synapsin image duplicated to aid visual alignment of clusters in columns). COS cells (C in phase contrast image, upper left) expressing neurexin 2α tagged intracellularly with CFP were overlaid on hippocampal neurons (N in phase contrast image, upper left) pre- grown for 8-10 days in culture. After one day of coculture, cells were fixed and immunolabeled for PSD-95, gephyrin, and synapsin. Endogenous inter- neuronal synapses have apposed clusters of postsynaptic PSD- 95 or gephyrin and presynaptic synapsin (arrows), thus clusters appear turquoise or purple, respectively, in the color overlays. In contrast, clusters of gephyrin lacking associated synapsin (arrowheads) appear red in the color overlay and were observed specifically at contact sites with COS cells expressing neurexin 2α. Clusters of PSD-95 lacking associated synapsin were not induced by neurexin 2α. Scale bar, 10 μm.        75 Figure 2.10. Neurexin 2β induces clustering of the inhibitory synaptic scaffolding protein gephyrin and the excitatory synaptic scaffolding protein PSD-95. Larger field of view single channel and additional overlay images are shown to more clearly illustrate the results of Fig. 2.1 (with synapsin image duplicated to aid visual alignment of clusters in columns). COS cells (C in phase contrast image, upper left) expressing neurexin 2α tagged intracellularly with CFP were overlaid on hippocampal neurons (N in phase contrast image, upper left) pre-grown for 8-10 days in culture. After one day of coculture, cells were fixed and immunolabeled for PSD-95, gephyrin, and synapsin. Endogenous inter- neuronal synapses have apposed clusters of postsynaptic PSD-95 or gephyrin and presynaptic synapsin (arrows), thus clusters appear turquoise or purple, respectively, in the color overlays. In contrast, clusters of PSD-95 or gephyrin lacking associated synapsin (arrowheads) appear green or red, respectively, in the color overlays and were observed specifically at contact sites with COS cells expressing neurexin 2β. While the induced clusters of PSD-95 and gephyrin appear to overlap at low resolution, at higher resolution they typically resolve into separate clusters side by side (see enlarged regions in Fig. 2.2). Scale bar, 10 μm.       76 Figure 2.11. α-Neurexins did not induce clustering of GABAA receptor α5 subunit. COS cells expressing neurexin 2α tagged intracellularly with CFP were overlaid on hippocampal neurons pre- grown for 16-18 days in culture. After one day of coculture, cells were incubated live with antibodies against either the α5 or the γ2 GABAA receptor subunit. After washing, cocultures were fixed, permeabilized, and incubated with antisynapsin and then secondary antibodies. Neurexin 2α induced surface clusters of GABAA receptor γ2 lacking apposed synapsin (arrowheads, red in color overlay), which tended to be of greater intensity than the endogenous GABAA receptor γ2 clusters apposed to synapsin (arrows, yellow in color overlay). In contrast, neurexin 2α did not induce detectable clustering of GABAA receptor α5. As previously described, endogenous GABAA receptor α5 was distributed more diffusely along dendrites compared with γ2. Some inhomogeneity in α5 immunoreactivity was observed along many dendrites, but the inhomogeneity was not more pronounced at contacts with COS cells expressing neurexin 2α. Scale bar, 10 μm.     77 2.4 Discussion We show here that α-neurexins promote GABAergic but not glutamatergic postsynaptic specialization. Specifically, α-neurexins expressed on COS cells cluster gephyrin and GABAA receptor but have no detectable effect on the distribution of PSD-95 in contacting dendrites of cultured hippocampal neurons (Figs. 2.1, 2.2, 2.3 and 2.5). This specificity presumably arises at least in part from the specific clustering of neuroligin 2 but not neuroligins 1/3/4 on the contacting dendrites by α-neurexins (Fig. 2.4). The postsynaptic clustering activity of α-neurexins is dependent on LNS6, being abolished by a single point mutation in the predicted calcium binding residue D1176A (Fig. 2.6). The central region of α- neurexin containing LNS 3-5 and two EGF-like domains inhibits the synaptogenic activity of LNS6 in the context of full-length α-neurexin (Fig. 2.5). Finally, we report that splicing of all neurexins at the S4 site is developmentally regulated, with levels of neurexins lacking the S4 insert low just before birth and increasing during the first two postnatal weeks in rat cortex and hippocampus (Fig. 2.7). Major differences between neurexins 1, 2, and 3 were not observed in these assays except for activity regulation of splicing at the S4 site, with differential regulation in 2β compared with 3α,β (Fig. 2.8). A major conclusion from this study is that α-neurexins are completely specific for promoting GABA but not glutamate postsynaptic differentiation in coculture assays. β- Neurexins containing the S4 insert were also previously shown to be more active for promoting GABA than glutamate postsynaptic differentiation in coculture assays (Chih et al., 2006; Graf et al., 2006). Chih et al. (Chih et al., 2006) did not observe induction of PSD-95 clusters by either neurexin 1β containing the S4 insert or neurexin 1α lacking the S4 insert and suggested that α-neurexins have a similar GABAergic specificity for synaptogenic activity as β-neurexins containing the S4 insert. However, we show here and in Graf et al. (Graf et al., 2006) that neurexin 1β containing the S4 insert does cluster PSD-95  78 significantly above the background level assessed by mCFP expression in COS cells. In the same experiment (Figs. 2.1, 2.2 and 2.3) where we observe significant clustering of PSD-95 by all β-type neurexin constructs (all +S4 variants), we find no significant clustering of PSD- 95 by α-neurexins. It is possible that we have observed these more subtle distinctions due to differences in experimental conditions from Chih et al. (Chih et al., 2006) including differences in construct expression level or neuron culture style. In experiments where we tried to bias toward seeing any effect of α-neurexins on distribution of glutamatergic postsynaptic proteins, assaying the more strongly clustered neuroligins rather than the scaffolding proteins, we could still find no effect of α-neurexins on distribution of neuroligin 1/3/4 (Fig. 2.4, noting that the major variant recognized by the common antibody in this experiment is thought to be neuroligin 1 +B as detailed under “Results”). In the same experiment, the α-neurexins could induce robust clustering of neuroligin 2 (Fig. 2.4). Previous evidence shows that neuroligin 2 is normally clustered only at GABA synapses (Varoqueaux et al., 2004; Graf et al., 2006) and neuroligin 1 primarily at glutamate synapses (Song et al., 1999). Subcellular localization of endogenous neuroligins 3 and 4 has not been reported, but the distribution of yellow fluorescent protein-tagged forms of neuroligins 3 and 4 is similar to that of neuroligin 1 and not 2 (Graf et al., 2004). We conclude here that α- neurexins induce clustering of the GABAergic proteins gephyrin, GABAA receptor and neuroligin 2, but not of the glutamatergic protein PSD-95 or neuroligin 1/3/4. These differences in ability to cluster specific postsynaptic components agree well with the binding selectivity of neurexins to neuroligin splice variants as reported recently (Boucard et al., 2005). The majority of neuroligin 1 in the brain is present in the form with the B insert (Chih et al., 2006). β-Neurexins with the S4 insert have lower binding affinity for neuroligin 1 with the B insert than without the B insert (Boucard et al., 2005; Chih et al., 2006; Graf et al., 2006). However, this difference is not an absolute effect, since in the most sensitive binding assay incubating soluble neurexin forms to neuroligins expressed on the  79 surface of HEK293 or COS cells, β-neurexins with the S4 insert still bound specifically to cells expressing neuroligin 1 with the B insert compared with untransfected control cells or compared with binding of an LNS deletion variant of neurexin (Boucard et al., 2005; Graf et al., 2006). In contrast, in the same experiment binding of α-neurexins with or without the S4 insert to neuroligin 1 with the B insert was not detected (Boucard et al., 2005). In short, all β neurexins bind neuroligin 1 +B but α neurexins do not. Thus, we conclude that the binding to neuroligin 1 with the B insert and ability to cluster glutamatergic postsynaptic proteins in coculture occurs in parallel. Furthermore, the addition of the S4 insert to β-neurexins reduces but does not abolish these activities, whereas the presence of the additional sequences in the longer α-neurexins abolishes these activities. Regulation of neurexin splicing has not yet been rigorously studied, although differential regulation can occur among mammalian brain regions (Ullrich et al., 1995) and during development in other systems (Patzke and Ernsberger, 2000; Zeng et al., 2006). Specifically with respect to the S4 site, differences in ratios of +S4 compared with -S4 variants of selected α- or combined α-plus β-neurexins have been reported among adult rat brain regions (Ichtchenko et al., 1995) and during development and neurotrophin exposure in chick sympathetic neurons (Patzke and Ernsberger, 2000). Here we show that the percentage of each α and β neurexin lacking the S4 insert increases between E18 and P11 in rat cortex and hippocampus. This developmental increase in the more glutamate-selective -S4 β variants correlates with the developmental increase in glutamatergic synaptogenesis. The earlier expression of the more GABA-selective +S4 β variants correlates with the earlier onset of GABAergic synaptogenesis (Tyzio et al., 1999; Ben-Ari et al., 2004). Neurexin 3 α and β, which showed the greatest developmental regulation, showed a similar increase in - S4 variants with development of E18 hippocampal neurons in dissociated culture from DIV 7 to 14 (Fig. 2.8). Interestingly, in the culture system where we could manipulate activity, we found that NMDA receptor blockade differentially regulates splicing, decreasing -S4 variants  80 of 3α,β and increasing -S4 variants of 2β. The significance of this differential activity regulation is not yet clear but may be important in combination with regulation of splicing at site 5, which is predicted to generate secreted forms of neurexin 3 but not 1 or 2 (Ushkaryov and Sudhof, 1993). No obvious developmental changes were observed for overall expression levels of α-neurexins, perhaps reflecting more their calcium channel coupling function at many synapse types rather than their additional GABA-specific roles. Analysis here of the single point mutation D1176A in the predicted calcium binding residue of LNS6 indicates that LNS6 is necessary for GABAergic synaptogenic activity of neurexin-1α; no other LNS domains are active (Fig. 2.6). Again, this coculture result is in good agreement with the binding data of Boucard et al. (Boucard et al., 2005). These authors found that LNS6 and a short upstream region is the minimal sequence necessary and sufficient from α-neurexin for binding neuroligin lacking the B insert. The combination of this published binding data and our coculture studies strongly suggest that α-neurexins cluster gephyrin and GABAA receptor through clustering neuroligin 2 on dendrites via binding of neuroligin 2 to LNS6. Although the α-neurexin-specific region upstream of LNS6 presumably interacts with other proteins and functions in coupling presynaptic calcium channels to release (Zhang et al., 2005), our deletion analyses show that this region of α- neurexin inhibits the synaptogenic activity of the C-terminal portion containing LNS6 (Fig. 2.6). The inhibitory activity did not reside in module A but in modules B and C. These upstream regions may either participate in an intramolecular interaction inhibiting the activity of LNS6 or may bind an additional inhibitory factor. More generally, in the coculture assays all neurexins induce clustering of GABAergic proteins to the same or a greater extent than that of glutamatergic proteins (e.g. comparing the increase in integrated intensity relative to the background on mCFP cells (Fig. 2.3) or comparing total area of induced clusters since non-normalized intensity would not be directly comparable (Chih et al., 2006; Graf et al., 2006)). This equal or greater activity for promoting  81 GABA postsynaptic development applies to all neurexins tested to date, 1-3, α and β, and containing or lacking the S4 insert. Although all splice forms of α-neurexins have not been tested, given that LNS6 of the α-neurexins bears the synaptogenic activity, it is highly unlikely that any forms not yet tested would have stronger activity for inducing glutamatergic than GABAergic postsynaptic protein clustering. These data are consistent with the idea that the neurexin-neuroligin system may be more important for the development and maintenance of GABA synapses than glutamate synapses. Support for this idea comes from the finding that inhibitory synaptic transmission is more severely impaired than excitatory synaptic transmission in P0 brainstem of the neuroligin 1/2/3 triple knockout mice (Varoqueaux et al., 2006). Spontaneous glutamatergic transmission frequency was reduced 4-fold and evoked failure rate unaffected, whereas spontaneous GABAergic/glycinergic transmission frequency was reduced 10-fold and evoked a failure rate markedly increased. At the same time, survival of the individual neuroligin knockouts but neonatal death of the triple knockout (Varoqueaux et al., 2006) indicates some redundancy in function rather than an absolute requirement for neuroligin 2 function at GABA synapses. In addition to the calcium channel coupling phenotype, α-neurexin 1/2/3 triple knock-out mice had a 2-fold reduction in the density of type II symmetric GABAergic synapses in neonatal brainstem with no change in density of type I asymmetric glutamatergic synapses (Missler et al., 2003). A 30% reduction in type II symmetric synapses but not type I asymmetric synapses was also found in adult neocortex of neurexin 1/2α and 2/3α double knock-out mice (Dudanova et al., 2007). It was not clear whether this loss of inhibitory synapses reflected a direct function of α-neurexin in GABA synapse development or an indirect effect of reduced synaptic activity due to the presynaptic calcium channel coupling defect. Based on the coculture results presented here, we suggest that the loss of inhibitory synapses in these mice may reflect a direct function of α-neurexins in linking GABAergic presynaptic and postsynaptic components. The apparent requirement for α-neurexin LNS6 to bind neuroligins for  82 coculture activity (D1176A in Fig. 2.6 combined with (Boucard et al., 2005; Graf et al., 2006)) suggests that the function of α-neurexins in GABA synapse formation and/or stabilization occurs via binding neuroligins. Although we could not observe clustering of dystroglycan by neurexins in the coculture assay, and GABA synapses can form in the absence of dystroglycan (Levi et al., 2002; Moore et al., 2002), the association of α- neurexins with dystroglycan (Sugita et al., 2001) may also contribute to stabilization of a postsynaptic complex at some GABA synapses (Knuesel et al., 1999). Members of the SynCAM, ephrin, Eph receptor, and netrin G ligand families are reported to trigger presynaptic and/or postsynaptic differentiation specifically for glutamate and not GABA synapses (Dalva et al., 2000; Sara et al., 2005; Kayser et al., 2006; Kim et al., 2006). The stronger functional association of the neurexin-neuroligin complex with GABA compared with glutamate synapses in coculture and in the knock-out mice may reflect a greater redundancy of synaptic organizing molecules at glutamate synapses. In Drosophila, which lack β neurexins and have less redundancy in general, null mutations in the single α neurexin gene lead to a reduction in numbers of central and neuromuscular synapses, altered ultrastructure, and defects in synaptic transmission and associative learning (Li et al., 2007; Zeng et al., 2007).       83 3  Inhibitory synapse dynamics: coordinated pre- and post-synaptic mobility and the major contribution of recycled vesicles to new synapse formation2   3.1 Introduction  Synapse development is a tightly regulated process, involving initiation, maturation, and maintenance. Key steps include accumulation of cell adhesion molecules at sites of axo- dendritic contact, presynaptic recruitment of vesicle cycling machinery, and postsynaptic recruitment of scaffolding complexes and ionotropic receptors (Shen and Scheiffele, 2010; Siddiqui and Craig, 2010). Regulated plasma membrane insertion, endocytosis and recycling, as well as local trapping and anchoring control trafficking of synaptic receptors (Malinow and Malenka, 2002; Triller and Choquet, 2005). Emerging evidence suggests that the entire synaptic structure, including scaffolding proteins and vesicles is, highly dynamic during both development and periods of plasticity (Goda and Davis, 2003; McAllister, 2007).  A growing understanding of the dynamics of excitatory synapses has been garnered through long-term live-cell microscopy of fluorescently-tagged synaptic proteins. At excitatory synapses in cultured neurons, clusters of PSD-95-GFP and other postsynaptic scaffold proteins show dynamic behaviours over periods of minutes to several days. New PSD-95 clusters formed mainly in dendrite protrusions by gradual accumulation from cytosolic pools, with a slight delay following the appearance of apposed synaptic vesicle clusters (Bresler et al., 2001; Okabe et al., 2001a). However, mobile clusters of postsynaptic  2 A version of this chapter has been previously published. Dobie FA, Craig AM (2011) Inhibitory synapse dynamics: coordinated presynaptic and postsynaptic mobility and the major contribution of recycled vesicles to new synapse formation. J Neurosci 31:10481-10493.  84 scaffolds and stationary postsynaptic clusters that can recruit presynaptic vesicles have also been observed (Gerrow et al., 2006). Stable postsynaptic scaffold clusters were associated with stabilization of dendritic protrusions (Prange and Murphy, 2001) and developed along newly grown dendrite branches over several days (Ebihara et al., 2003). While developing dendrite arbors showed net increases in postsynaptic scaffold cluster density with time, a fraction of clusters were also eliminated (Okabe et al., 1999). The size of individual PSD-95 clusters tended to normalize over time (Minerbi et al., 2009). Postsynaptic scaffold clusters could be mobile, translocating up to a few microns along dendrites, or between shafts and dendritic protrusions, and could exhibit complex behavior including merging and splitting (Bresler et al., 2001; Marrs et al., 2001; Prange and Murphy, 2001; Gerrow et al., 2006). The dynamics of excitatory synapse formation have also been shown in vivo. In young neurons of Zebrafish optic tectum, PSD-95 accumulation frequently preceded stabilization of dendritic filopodia, which then grew into new dendritic branches (Niell et al., 2004). Mechanisms of axonal arbor stabilization occurring at sites of synaptic vesicle accumulation have also been observed (Meyer and Smith, 2006; Ruthazer et al., 2006). In developing mouse barrel cortex, some PSD-95-GFP clusters imaged in vivo appeared or disappeared while other clusters were stable and maintained relative differences in intensity over days (Gray et al., 2006).  Despite this growing body of evidence on excitatory synapse dynamics, far less is known of inhibitory synapses. Because the balance of excitation and inhibition may be a key factor in the etiology of neurological and cognitive disorders (Rogawski and Löscher, 2004; Sudhof, 2008), it is critical to understand how inhibitory synapses are formed, maintained, and modified. We thus assess here the dynamic behaviour of inhibitory pre- and postsynaptic components using long-term live-cell fluorescence microscopy, exploring modification of existing synapses and development of new ones.    85 3.2 Materials and Methods  3.2.1 Hippocampal neuron culture and nucleofection. Dissociated hippocampal neuron culture from E18 rats of both sexes was performed essentially as described (Kaech and Banker, 2006), with the following modifications. Freshly dissociated neurons were pelleted at 800 x g for 5 min and subjected to Amaxa nucleofection according to the manufacturer’s protocol (Lonza, Basel, Switzerland) using 500,000 cells and 3 mg of DNA. Transfected neurons were mixed 1:1 with untransfected neurons and plated at a density of 150,000 cells/35 mm dish in Plating Medium (phenol red-free Minimum Essential Medium, 1 X Glutamax-I, 1 mM sodium pyruvate, 0.6% glucose, 10% horse serum). Dishes had been previously prepared by drilling a 22 mm round hole in the center of a 35 mm BD Falcon tissue culture dish and adhering a pre-washed 25 mm round glass coverslip to the bottom of the dish using a warm 3:1 paraffin:Vaseline mixture. Dishes were extensively washed, coated with poly-D-lysine, and pre-incubated with Plating Medium. Three h after neurons were plated onto these dishes, they were briefly washed with phenol red-free Neurobasal Medium, and then cultured in Maintenance Medium (phenol red-free Neurobasal Medium, 1 X Glutamax-I, 2% Neurocult SM1 supplement (StemCell Technologies, Vancouver, British Columbia)). This medium had been pre-conditioned for at least 3 days by glial culture growing on a 24 mm square Thermanox coverslip (Ted Pella Inc, Redding, California), which was also transferred to the 35 mm dish into which neurons were plated. After 2 days in vitro (DIV), glial proliferation was stunted by addition of 4 mM cytosine-arabinoside. Neurons were maintained by replacement of one third of culture media with fresh Maintenance Medium every 3 days. All reagents were from Invitrogen (Carlsbad, California) or Sigma (St. Louis, Missouri).  3.2.2 DNA constructs. cDNA for rat Gephyrin isoform 2,6 was obtained from Dr. Antoine Triller (Meier et al., 2000). mCitrine (a monomeric, chloride-insensitive variant of YFP, but  86 called YFP hereafter for simplicity) was fused to the N-terminus of Gephyrin separated by a 12 amino acid spacer (GGGSGGGSGGGS) using site-directed mutagenesis. This construct was subcloned into the NheI/PacI sites of a modified pLentiLox3.7 vector in which the CMV promoter had been replaced with the human synapsinI promoter (Huang et al., 2009), allowing for low-level expression in neurons. For construction of mCherry-Gephyrin, the coding sequence of mCherry (Shaner et al., 2004) was amplified by PCR and inserted into the NheI/BsrGI sites of the above construct to replace mCitrine. cDNA for rat GABAARg2 subunit, short form, was obtained from Dr. Peter Seeburg (Pritchett et al., 1989). Site- directed mutagenesis and PCR were used to add His and FLAG epitopes (ASGHHHHHHGMDYKDDDDKGLG) then EYFP between amino acids 4 and 5 of the mature g2 peptide coding sequence. This entire construct was subcloned into the AgeI/PacI sites of a modified pLentiLox3.7 vector in which the CMV promoter had been replaced with the chick b-actin CAG promoter (Takahashi et al., 2011).  3.2.3 Immunocytochemistry and live antibody labeling. Neurons were fixed for 15 min with warm 4% formaldehyde and 4% sucrose in PBS (pH 7.4) followed by permeabilization with PBST (PBS + 0.25% Triton X-100). Fixed and permeabilized cultures were blocked in 10% BSA in PBS for 30 min at 37oC and primary antibodies applied in 3% BSA in PBS for 2 h at room temperature. Coverslips were washed with PBS and incubated in secondary antibodies in 3% BSA in PBS for 1 h at 37oC.  The coverslips were then washed and mounted in elvanol (Tris-HCl, glycerol, and polyvinyl alcohol, with 2% DABCO (1,4- diazabicyclo[2,2,2]octane)). The following primary antibodies were used: mouse anti- Gephyrin (IgG1; 1:500; mAb7a; Synaptic Systems), rabbit anti-VGAT, cytoplasmic domain (1:500; 131 003; Synaptic Systems),  guinea pig anti-VGAT, cytoplasmic domain (1:500; 131 004; Synaptic Systems), rabbit anti-GABAARg2 (1:500; 224 003; Synaptic Systems), and anti-MAP2 for labeling dendrites (chicken polyclonal IgY; 1:10,000; Abcam; ab5392). Labeled secondary antibodies used were raised in goat against the appropriate species and  87 monoclonal isotype, highly cross-adsorbed, and conjugated to Alexa-568 and Alexa-647 dyes (1:500; Molecular Probe). To visualize dendrites, we used aminomethylcoumarin acetate conjugated anti-chicken IgY (donkey IgG; 1:200; Jackson ImmunoResearch; 703- 155-155).  For live-staining of actively recycling inhibitory presynaptic terminals, neuronal Maintenance Medium and glial feeder coverslip were first removed to a separate dish. Neurons were washed briefly in fresh Maintenance Medium, then briefly in normal Extracellular Solution (140 mM sodium chloride, 4 mM potassium chloride, 20 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid )), 10 mM D-glucose, 2 mM calcium chloride, 1.3 mM magnesium chloride, pH 7.4). Rabbit anti-VGAT directed towards the luminal domain of VGAT and labeled with Oyster-550 (131 103C3; Synaptic Systems) was suspended at a dilution of 2.5 mg/mL in high potassium Extracellular Solution (89 mM sodium chloride, 55 mM potassium chloride, 20 mM HEPES, 10 mM D-glucose, 2 mM calcium chloride, 1.3 mM magnesium chloride, 20 mM AP5  ((2R)-amino-5-phosphonovaleric acid), 5 mM CNQX (6-cyano-7-nitroquinoxaline-2,3-dione)). Neurons were labeled in this solution for 1 min at 37oC then washed extensively with fresh Extracellular Solution and Maintenance Medium before the conditioned Maintenance Medium and glial coverslip were replaced. Neurons were allowed to recover for 30 min before imaging.  3.2.4 Live imaging. Glass-bottomed neuron dishes were imaged on a modified Nikon TE- 2000 inverted epifluorescence microscope (Nikon Instruments, Melville, New York). The microscope was equipped with the Perfect Focus System to maintain the plane of focus over various regions of the coverslip for extended periods. Further, the microscope was fitted with a motorized Prior stage (Cambridge, UK) for repeated imaging of multiple coverslip positions and OKOLabs stage-top environmental control chamber (Quarto, Italy), allowing the imaging environment to be maintained at 37oC, 5% CO2/95% air, 100% humidity. All imaging was performed with a 63 X oil-immersion objective at lowest light levels  88 and exposure times to allow detection of fluorescence signal without inducing photobleaching or photodamage. Microscope hardware was controlled by and images obtained using MetaMorph (Molecular Devices, Sunnyvale, California).  3.2.5 Image and data analysis. Fixed cell immunocytochemistry images were analyzed using custom journals in MetaMorph. Live imaging movies were first aligned in ImageJ (National Institutes of Health). Fluorescent puncta intensity, size, and density were measured by first applying a “Mexican hat” filter (LoG3D, Daniel Sage, http://bigwww.epfl.ch/sage/soft/LoG3D/) to aid detection of spots; detected spots were then used as a mask for quantitation. Motion tracking of fluorescent puncta (Fig. 3.2, 3.3, 3.5) was either performed by hand or by using mass-center tracking based MTrackJ plugin for ImageJ (Erik Meijering, http://www.imagescience.org/meijering/ software/mtrackj/). The mean-squared displacement and diffusion coefficients were calculated according to the method of Hanus, et al. (Hanus et al., 2006). Formation of new puncta and merging events (Figs. 3.5, 3.6, 3.7) were analyzed by eye and measured by hand. Rapidly mobile YFP- Gephyrin puncta in transfected axons were excluded from analysis. All data were analyzed in Excel and GraphPad Prism (GraphPad Software, La Jolla, California). All graphs with error bars represent the mean ± standard error.  3.3 Results  3.3.1 Clustering of Gephyrin along developing dendrites occurs slowly over time We characterized the development of inhibitory synapses by expressing the major inhibitory postsynaptic scaffolding protein Gephyrin (Fritschy et al., 2008) tagged with YFP in cultured hippocampal neurons and assessing the changes in YFP-Gephyrin clustering over time. In this system, YFP-Gephyrin clustering becomes prominent on most cells around DIV 10. Beginning at this time point, we imaged fluorescent protein expressing neurons  89 consecutively every hour for 3 days (Fig. 3.1A). The integrated fluorescence intensity of individual Gephyrin clusters increased modestly over time at a mean rate of roughly 0.4%/hour (Fig. 3.1B), suggesting gradual addition of YFP-Gephyrin molecules to their sites of clustering. We observed a slow but steady increase in YFP-Gephyrin cluster density of 0.00232 ± 0.00016 puncta/mm dendrite/hour (Fig. 3.1C). Assuming a dendritic arbour of roughly 600 mm in length, this equates to roughly 30-35 new YFP-Gephyrin clusters forming per cell per day.  To assess whether these YFP-Gephyrin clusters represent bona fide synaptic sites, their co-localization with an inhibitory presynaptic marker was assessed. First, we labeled living DIV12-15 neurons with an antibody to the luminal domain of vesicular GABA transporter VGAT (Martens et al., 2008), allowing specific uptake by actively recycling inhibitory terminals (Fig. 3.1D). Immunocytochemistry of untransfected cultures and staining for endogenous Gephyrin and VGAT (cytoplasmic domain) showed 54 ± 2% colocalization of Gephyrin with VGAT, while colocalization of  YFP-Gephyrin with endogenous VGAT showed 66 ± 2% colocalization (Fig. 3.1E). Interestingly, live cultures stained with the antibody towards the luminal domain of VGAT showed 74 ± 2% colocalization of YFP- Gephyrin with VGAT fluorescent puncta, suggesting this assay may be better suited to visualizing inhibitory synaptic sites than immunocytochemistry methods. To assess the effect of this antibody labeling on inhibitory synapse formation, we stimulated cells with the hyperkalemic loading solution with or without Oyster-550-VGAT antibody, and allowed cells to recover for 18 hours before fixation and immunocytochemistry for endogenous inhibitory synapse markers. Neither treatment showed significant differences in postsynaptic Gephyrin or presynaptic VGAT densities compared to control (Fig. 3.1F; ANOVA p > 0.05). No significant difference was found in the puncta density of endogenous Gephyrin between untransfected control cells and YFP-Gephyrin expressing cells (Fig. 3.1G; t-test p > 0.05), showing that exogenous expression of Gephyrin in this system from the time of plating does not alter overall inhibitory synaptogenesis. This may be due to the fact that expression of  90 Figure 3.1. Long-term imaging of inhibitory synapse dynamics. A. Hippocampal neurons expressing YFP-Gephyrin imaged at 1-h intervals for 3 days. Images at left show the first and last time points imaged. The boxed dendrite region is magnified and straightened at right with individual frames representing intervals of 7-10 h. B-C. The integrated fluorescence intensity of YFP-Gephyrin puncta over time shows a modest increase, while puncta density shows a slow but steady increase. n = 21 dendrites from 7 cells. D. A YFP-Gephyrin expressing neuron co-stained live with Oyster-550- labelled antibody directed towards the luminal domain of VGAT. The boxed dendrite region is magnified at right. E. Quantitation of colocalization of endogenous Gephyrin or YFP-Gephyrin with VGAT. Left, untransfected DIV 12-15 cells fixed and immunostained for endogenous Gephyrin and VGAT. n = 71 dendrite regions from 20 cells. Middle, DIV 12-15 cells expressing YFP-Gephyrin fixed and stained for VGAT. n = 90 dendrite regions from 20 cells. Right, DIV 12-15 cells expressing YFP- Gephyrin and stained live with the Oyster-550-anti-VGAT. n = 57 dendrite regions from 7 cells. F. Effect of antibody labeling protocol on inhibitory synaptic markers. Neurons were either untreated (Control), treated with 55 mM KCl solution for 2 min (Stim), or treated with 55 mM KCl solution containing 2.5 mg/ml Oyster-550-anti-VGAT antibody for 2 min (Stim + Ab), then returned to their home dishes and fixed for immunocytochemistry 18 h later. Neither treatment significantly affected endogenous inhibitory postsynaptic (left) or presynaptic (right) density per dendrite area. n = 30 cells. G. Quantitation of Gephyrin-immunoreative puncta density per dendrite area from untransfected control or YFP-Gephyrin expressing cells. n = 20 cells. H. Normalized average fluorescence intensity of clusters immunostained for Gephyrin from untransfected control or YFP-Gephyrin expressing cells, showing relative overexpression of Gephyrin of ~ 4-14 %. n = 20 cells. All values represent mean ± SEM. Scale bars in A, D, 10 mm (left) and 5 mm (right).  YFP-Gephyrin from the human synapsinI promoter results in Gephyrin overexpression of only roughly 4-14% above endogenous levels (Fig. 3.1H). Together, these data show that this system is suitable for studying the developmental dynamics of inhibitory synapses.  91 3.3.2 Inhibitory postsynaptic scaffolds show dynamic mobility YFP-Gephyrin was imaged at frame intervals of 1 second (Fig. 3.2A), 1 minute (Fig. 3.2B), and 1 hour (Fig. 3.2C) for at least 18 frames. Mass center tracking was used to quantitate the mobility of YFP-Gephyrin clusters. For DIV12 neurons, the average instantaneous velocity, the gross displacement of a cluster between consecutive frames, was 0.077 ± 0.003 mm/s, 0.193 ± 0.007 mm/min, and 0.261 ± 0.015 mm/h (Fig. 3.2D). The diffusion coefficient of clusters imaged every 1 second, calculated from the plot of mean-squared displacement (not shown), was found to be 1.26 ± 0.006 x 10-4 mm2/s, similar to the value found previously in spinal cord neurons (Hanus et al., 2006). Roughly 5% of clusters imaged moved > 1 mm/min or > 2 mm/h (Fig. 3.2E). Mobile YFP-Gephyrin puncta were typically constrained within a relatively small radius, as seen from their average displacement from their original position (0.133 ± 0.012 mm/s, 0.307 ± 0.021 mm/min, and 0.763 ± 0.093 mm/h for 12 DIV; Fig. 3.2F). Roughly 10% of clusters translocated > 5 mm from their origin at some time during an 18-h imaging period (Fig. 3.2G). Total path length, the sum of all incremental displacements during the entire imaging period, averaged 1.38 ± 0.06 mm in 18 s, 3.47 ± 0.12 mm in 18 min, and 4.71 ± 0.27 mm in 18 h for DIV12 cells (Fig. 3.2H). A small subset of puncta was observed to move > 10 mm during an 18 hour imaging period. Although the three imaging epochs (every second, minute, or hour) show differences in the apparent mobility of YFP-Gephyrin puncta and occasional rapid mobility, it is clear from the high confinement index (the ratio of total path length to displacement from origin) that Gephyrin clusters are generally constrained to a well-defined sub-region of dendrite (Fig. 3.2I). However, clusters imaged every 1 hour showed a significantly lower confinement index than those imaged every 1 second or 1 minute (Fig. 3.2I), suggesting a greater involvement of non-stochastic processes in Gephyrin movement during longer imaging intervals.     92  Figure 3.2. Mobility of YFP-Gephyrin puncta. A, B, C. Montages of straightened dendrites obtained by imaging YFP-Gephyrin clusters from DIV12 neurons for 19 frames at intervals of 1 sec, 1 min, and 1 h, respectively. Bottom frame represents the maximum intensity projection of all time points imaged. D-I. Quantitative analysis by mass center tracking of individual puncta. n = 60-80 puncta from 3-4 cells. DIV12 /h and DIV15 /h not significantly different, t-test (D, F, H, I). D. Mean instantaneous velocity, averaged over all frames. E. Cumulative probability distribution of the maximum instantaneous velocity within the entire imaging period. F. Mean displacement of puncta from their original positions, averaged over all frames. G. Cumulative probability distribution of the maximum displacement of puncta from their origins within the entire imaging period. H. Scatter dot-plot of the total path length of individual puncta over the entire imaging period. Lines show the mean of all tracked puncta. I. Confinement index, defined as the total path length divided by the average displacement from origin of a puncta. Puncta imaged at 1 h intervals showed significantly less confinement than puncta imaged at 1 sec or 1 min intervals (*** ANOVA with Bonferroni post hoc test, p < 0.001). Scale bar in A,B,C, 5 mm.     93 3.3.3 Inhibitory pre- and postysynaptic structures show correlated mobility as a single unit Since inhibitory synapses in living neurons could be reliably labeled for both their presynaptic (VGAT) and postsynaptic (Gephyrin) components (Fig. 3.1D-E), we monitored their mobility in tandem using dual channel fluorescence microscopy. Apposed YFP- Gephyrin and 550-VGAT puncta showed unified mobility (Fig. 3.3A-J). Mass center tracking of both pre- and postsynaptic puncta showed these two components to be in lock-step with one another, as can be seen from their correlated instantaneous velocities, displacements from origin, and total path length, and maintenance of low distance between centres of mass (Fig. 3.3B-E, G-J). Over the entire imaging period, while both YFP-Gephyrin and 550-VGAT puncta were displaced, on average, > 5 mm, the distance between their centres of mass remained constant at mean 0.43 ± 0.08 mm (Fig. 3.3K). The total displacement of a given YFP-Gephyrin puncta and its apposed 550-VGAT puncta were highly correlated, even for puncta traveling > 10 mm (Fig. 3.3L). Moreover, regardless of how much a YFP-Gephyrin puncta was displaced during a single imaging frame, the distance between its center of mass and that for 550-VGAT remained relatively constant (Fig. 3.3M). A small proportion of cluster pairs (150/3626, 4.1%) displayed transient interpuncta distances > 2 mm that persisted for only a single frame, likely due to large fluctuations in the VGAT center of mass rather than dissociation and re-association of appositions (see below). Although we hypothesized that smaller YFP-Gephyrin and 550-VGAT puncta might display higher mobility, there was little correlation (r2 = 0.032, p = 0.055; and not shown). To ensure we could reliably track apposed pre- and postsynaptic clusters and not structures that were disassembled and re-formed in between images, we followed low frequency imaging (every 1 hour) by high frequency imaging (every 7 minutes and every 10 seconds) (Fig. 3.3N). Apposed YFP-Gephyrin and 550-VGAT clusters were visualized in every imaging frame, and the interpuncta distance remained constant over the different imaging epochs (Fig. 3.3O), suggesting these tracked cluster pairs represent genuine  94 appositions. We also assessed mean-squared displacement as a way to objectively compare mobility of Gephyrin and VGAT clusters. Of the clusters that were most mobile during the initial low frequency imaging period (Fig. 3.3O, right), calculated diffusion coefficients, D, for YFP-Gephyrin and 550-VGAT were not significantly different over any imaging epoch (D1h(Gephyrin) = 0.50 ± 0.10 vs. D1h(VGAT) = 0.45 ± 0.10 mm2/h; D7min(Gephyrin) = 0.12 ± 0.06 vs. D7min(VGAT) = 0.14 ± 0.08 mm2/min; D10s(Gephyrin) = 0.022 ± 0.027 vs. D10s(VGAT) = 0.052 ± 0.030 mm2/s; t-test, p > 0.05 for all comparisons). Of the clusters that were least mobile during the initial low frequency imaging period (Fig. 3.3O, left), calculated diffusion coefficients of YFP-Gephyrin and 550-VGAT during the initial low frequency imaging period were significantly different (D1h(Gephyrin) = 0.028 ± 0.003 vs. D1h(VGAT) = 0.097 ± 0.028 mm2/h, p < 0.05), while other diffusion coefficients were not (D7min(Gephyrin) = 0.053 ± 0.010 vs. D7min(VGAT) = 0.084 ± 0.024 mm2/min,  p > 0.05; D10s(Gephyrin) = 0.011 ± 0.024 vs. D10s(VGAT) = 0.033 ± 0.011 mm2/s,  p > 0.05). This apparently higher diffusion coefficient for VGAT than for Gephyrin for the least mobile cluster fraction may represent greater shifting of the centre of mass of the VGAT cluster due to transport of vesicle packets in and out of the VGAT cluster  (Ahmari et al., 2000; Staras et al., 2010). In contrast, if YFP-Gephyrin clusters are maintained from cytoplasmic pools (see Fig. 3.6 and related discussion), exchange of individual molecules would have little effect on centre of mass of the Gephyrin cluster. Importantly, for the most mobile fraction of cluster pairs, and for the average (Fig. 3.3O, middle), calculated diffusion coefficients for Gephyrin and VGAT did not differ, supporting a dominance of coordinated movement. To further investigate whether these mobile structures represent synapses, we co- expressed mCherry-Gephyrin together with the inhibitory postsynaptic GABAA receptor g2 subunit tagged with YFP (Fig. 3.3P). The g2 subunit is present in the majority of synaptic GABAA receptors and contributes to synaptic localization in hippocampal neurons (Essrich et al., 1998; Luscher and Keller, 2004). Mobile YFP-GABAARg2 clusters were observed in tight association with mCherry-Gephyrin clusters, moving essentially in unison, and  95 displaying patterns of mobility similar to co-clusters of YFP-Gephyrin/550-VGAT. We found 74 ± 3% of mCherry-Gephyrin clusters co-localized with YFP-GABAARg2 clusters (n = 8 cells), similar to the value of 78 ± 2% co-localization of YFP-Gephyrin with endogenous GABAARg2 (Fig. 3.6J; n = 18 cells). Together, these data show that entire mature inhibitory synapses, composed of both pre- and postsynaptic components, can be highly mobile as a single entity.  3.3.4 The number of inhibitory synapses is variable over time due to apparent splitting and merging of pre- and postsynaptic structures Long-term imaging of YFP-Gephyrin and 550-VGAT puncta revealed fluctuations in their cluster density along dendrites over time (Fig. 3.4A). Spot detection using a “Mexican hat” filter revealed both the appearance and disappearance of puncta between successive 30- min frames, leading to observed changes in puncta density (Fig. 3.4B, C). Such methods may overestimate transient changes, due to individual clusters crossing paths at distances below resolution or transiently falling below threshold for detection, but present an objective way to compare multiple dendrites and cells, and have been commonly used to quantitate excitatory synapses (Ma et al., 1999). Despite apparent appearance or disappearance of multiple puncta per frame within individual dendrites, the average YFP-Gephyrin puncta density for all cells remained relatively stable for an entire neuron, increasing slightly over the 18 h imaging period (Fig. 3.4D). The measured rates of addition and elimination of YFP- Gephyrin clusters averaged hourly from multiple dendrites per cell ranged from 0.0088 ± 0.0020 to 0.0261 ± 0.0014 events/mm/h. These short-term rates of addition and elimination were considerably greater than the net rate of change in puncta density determined by comparing the 0 and 18 hour times with an average net rate of increase in YFP-Gephyrin cluster density from all dendrites of all cells being 0.00227 ± 0.00082 puncta/mm/h. Individual dendrite regions from different cells showed a continuum of flux behaviours (Fig. 3.4E-F). For example, all three dendrites of one cell showed little change in YFP-Gephyrin  96 Figure 3.3. Coordinated movement of inhibitory pre- and postsynaptic components. A. Example image montage from a DIV12 YFP-Gephyrin expressing neuron co-labeled with Oyster-550- VGAT luminal antibody imaged every 0.5 h for 18 h. The lower right frame shows the entire tracked path of both YFP-Gephyrin (green) and 550-VGAT (red) for the entire imaging period. B-E. Measures from mass centre tracking for both YFP-Gephyrin and the corresponding apposed 550-VGAT puncta over time for A. Interpuncta distance (E) is distance between the centers of mass of YFP-Gephyrin and 550-VGAT puncta. F. Example image montage as in A showing a YFP-Gephyrin/550-VGAT cluster pair with greater total path length. G-J. Measures from mass centre tracking of the cluster pair in F. For reference, the YFP-Gephyrin cluster in A is shown on each graph as a grey dotted line. K. Averaged puncta tracking results from 40 YFP-Gephyrin (green)/550-VGAT (red) cluster pairs from DIV12 neurons. Left y-axis shows the total path length of both particles over time. Right y-axis (cyan) shows the Gephyrin-VGAT interpuncta distance. L-M. Correlation analysis shows positive association of 550-VGAT puncta total displacement with apposed YFP-Gephyrin puncta total displacement (p < 0.0001), and no association of individual frame-by-frame YFP-Gephyrin/550-VGAT interpuncta distance with YFP-Gephyrin instantaneous velocity (p > 0.5). n = 114 clusters, >3600 frames. N. Example image montage of a DIV 12 YFP-Gephyrin expressing neuron labeled with 550-anti-VGAT imaged every 1 h for 4 h, then every 7 min for 56 min, then every 10 sec for 2 min. Time notation, h:mm:ss. O. Quantitation of mean-squared displacement (MSD, left y-axis) and interpuncta distance (right y-axis) over the entire imaging period. Vertical dashed lines represent, from left to right, the per 1 h, per 7 min, and per 10 sec imaging epochs. The left graph shows quantitation from the 20% of cluster pairs showing the least mobility during the first 4 h of imaging, the middle graph all cluster pairs, and the right graph the 20% of cluster pairs showing the most mobility. n = 126 cluster pairs from 6 cells. P. Example image montage from a DIV 12 neuron co-expressing mCherry-Gephyrin (red) and YFP-GABAARg2 subunit (green) imaged every 1 h for 18 h. Scale bars in A, F, N, 1 mm; P, 2 mm.   97 puncta density, with values of 0.0000 to 0.00045 puncta/mm/h, or a change of only 0% to +5% from initial puncta density. In another example, different dendrites of a single cell showed both loss of up to 0.0064 puncta/mm/h and gain of up to 0.0030 puncta/mm/h, or a change of -16% to +20%. Finally, another cell showed different dendrites with a large range of changes in YFP-Gephyrin puncta density, from 0.0000 to 0.0181 puncta/mm dendrite/h, or 0% to +90% from initial puncta density. Hypekalemic loading of the Oyster-550-VGAT antibody did not affect synapse flux, as cells expressing YFP-Gephyrin but unlabeled for VGAT showed similar rates of addition and elimination with high variability (Fig. 3.4F). These data suggest that the level of flux is highly variable not only between different neurons but also between dendrites within an individual neuron.  Appearance and disappearance of fluorescent puncta was often observed associated with splitting of a single puncta into multiple puncta, or merging of several puncta into one, respectively. The interpretation of such observations is limited by the resolution of light microscopy. While we cannot rule out the possibility that apparently merged clusters are co-existing and co-migrating one above the other in the z plane, the apparent unified mobility of such clusters following merging could persist for many hours, even days in the long time-lapse series, suggesting a true merge. Apparent merging of clusters was observed for both YFP-Gephyrin and 550-VGAT in tandem (Fig. 3.5A, B). We less frequently observed splitting of a single puncta into multiple puncta. Apparent splitting and merging of YFP-Gephyrin often occurred without change in presynaptic structure apposed to the dynamic postsynaptic structures. In such cases, we often saw a single large 550-VGAT cluster apposed to several dynamic YFP-Gephyrin puncta (Fig. 3.5C), suggestive of multi- synapse boutons. By immunofluorescence of control neurons, 3.4 ± 0.4% of VGAT clusters were apposed to greater than one Gephyrin cluster. Cells expressing YFP-Gephyrin and subjected to 550-VGAT antibody labeling showed 3.6 ± 0.6 % of VGAT clusters with multiple Gephyrin clusters, suggesting that the live imaging protocol did not affect the formation of multi-synapse boutons (t-test, p > 0.05).  98 Figure 3.4. Flux of YFP- Gephyrin puncta in developing neurons. A. Image montage of a straightened dendrite region from a DIV12 YFP-Gephyrin expressing cell imaged every 0.5 h for 18 h. Lower panel shows colocalization of YFP-Gephyrin with 550- anti-VGAT at time 3.5 h. B. Thresholded and binarized frames from the montage in A obtained by the “Mexican hat” filter of YFP-Gephyrin signal used to assist in quantitation of individual clusters. Each frame is depicted in a different color, with all time points overlaid at bottom. C. Quantitation of the number of YFP- Gephyrin puncta (green) and 550-VGAT (red) observed over time from the dendrite region in A above. D. Quantitation of average YFP-Gephyrin puncta density over time for 26 dendrite regions from 7 cells at 12-15 DIV. E. Quantitation of the net change in YFP-Gephyrin cluster density at t18h relative to t0h. Each column of colored points represents an individual neuron, while each point represents a separate dendrite. F. Quantitation of the change in YFP-Gephyrin cluster density per cell averaged over all time points, where addition (solid bars) and elimination (hatched bars) are defined as an increase or decrease, respectively, in puncta density from tn to tn+1. At left (Stim + Ab), cells were co-labeled with Oyster-550-anti-VGAT antibody, with each colour representing the corresponding cells in E. At right (Control), cells were not stimulated or exposed to antibody. n = 3-6 dendrites per cell. Scale bar in B, 5 mm.  Whereas merging occurred frequently, elimination of YFP-Gephyrin and apposed 550-VGAT by gradual diminution was rare. Such gradual elimination of YFP-Gephyrin clusters was more apparent in the 3-day time-lapse series. Careful examination of these long time series revealed a number of clusters that formed by slow accumulation (see below), were stable for many hours, but were eventually eliminated. The mean lifetime of such clusters was 23.2 ± 2.6 h (n = 18 clusters from 4 cells). Since these 3-day experiments were performed without VGAT co-label, we could not determine whether such eliminated clusters were synaptic.   99 3.3.5 Development of inhibitory synapses occurs by slow accumulation of pre- and postsynaptic components and utilizes recycled synaptic vesicles A critical factor in the net increase in YFP-Gephyrin clustering along dendrites is the formation of new synapses. Although the net increase in the number of synapses was small, we were able to observe a number of de novo synapse formation events, defined as the appearance of new stable YFP-Gephyrin clusters and associated VGAT at sites previously lacking YFP-Gephyrin or VGAT. These events could be classified by the slow accumulation of YFP-Gephyrin and 550-VGAT at the same site (Fig. 3.6A). Both YFP-Gephyrin and 550- VGAT fluorescence intensities slowly increased over >8 hours (Fig. 3.6B), with 550-VGAT beginning to accumulate at nascent synaptic sites roughly 2 hours before the appearance of YFP-Gephyrin (Fig. 3.6C). These data suggest that inhibitory presynaptic vesicle accumulation precedes postsynaptic scaffold formation. Furthermore, both processes occurred by gradual addition of key synaptic components to these sites rather than by abrupt recruitment of large preformed complexes. Ongoing flux of small 550-VGAT puncta was observed, representing mobile packets that may contribute to both pre-existing and newly formed presynaptic elements. On the contrary, discrete dendritic transport packets were not observed for YFP-Gephyrin with its accumulation at nascent synapses appearing to occur from diffuse dendritic pools. In addition to dendritic YFP-Gephyrin, for reasons we do not understand, YFP-Gephyrin occasionally formed puncta that were rapidly transported in transfected axons; such axonal clusters were excluded from the analysis in this study.  Our data are further interesting in that 550-VGAT fluorescence signal is only detected in synaptic vesicles that had been labeled during the vesicle recycling protocol prior to the imaging session. The observation that VGAT intensity at newly formed synapses could reach that of VGAT clusters at neighbouring pre-existing synapses (e.g., Fig. 3.6A) implies that a significant proportion of vesicles from active release sites could be transferred to generate newly formed presynaptic sites. However, new YFP-Gephyrin clusters could also form in the absence of associated 550-VGAT (Fig. 3.6D). The time course of YFP-  100 Figure 3.5. Complex dynamic behaviour of inhibitory synaptic components. A. Image montage of a dendrite region from a DIV 12 YFP-Gephyrin expressing neuron co-labelled with 550-anti-VGAT and imaged every 0.5 h. Two distinct YFP-Gephyrin and 550-VGAT clusters (yellow and cyan arrows) are visible at the beginning of the imaging period, which merge into one cluster at 3 h (green arrow). B. Integrated fluorescence intensity of YFP-Gephyrin puncta corresponding to matching coloured arrows in A over time. C. Image montage as in A, showing several YFP-Gephyrin puncta (yellow, magenta, cyan arrows) rearranging around one large central 550-VGAT cluster over time. Scale bars in A,C, 2.5 mm.  Gephyrin accumulation at sites without 550-VGAT labeling (Fig. 3.6E) was similar to sites where 550-VGAT also accumulated (Fig. 3.6B). Of all new YFP-Gephyrin clusters observed to form, 65% were accompanied by concomitant accumulation of 550-VGAT at the same site (28/43 clusters), while the remaining 35% formed in the absence of associated 550- VGAT clustering. However, post hoc immunocytochemistry (Fig. 3.6I) revealed that 78.5% of newly formed YFP-Gephyrin clusters (62/79 clusters) colocalized with VGAT. Since 65% of new YFP-Gephyrin clusters also accumulated 550-VGAT, it appears that roughly only  101 13.5% of new presynaptic clusters were undetectable in our live imaging assay, highlighting the important contribution of sharing of presynaptic vesicles to nascent synaptic sites. The 21.5% of new clusters of YFP-Gephyrin not apposed to VGAT immunofluorescence may represent future synapses very slow to accumulate VGAT, non-synaptic clusters, or inhibitory postsynaptic specializations misapposed to excitatory boutons (Rao et al., 2000; Christie and De Blas, 2003).  By post hoc immunocytochemistry (Fig. 3.6I), we also detected GABAARg2 at all of the newly formed YFP-Gephyrin clusters colocalized with VGAT (62/62), supporting the idea that they represent bona fide synapses. Indeed, further suggesting that many sites of YFP- Gephyrin accumulation represent genuine inhibitory synapses, imaging of neurons co- expressing mCherry-Gephryin and YFP-GABAARg2 revealed accumulation of postsynaptic receptors in concert with postsynaptic scaffolds (Fig. 3.6F-G). Both by the dual live imaging method and by the post hoc immunocytochemistry, over 87% of newly formed stable clusters of tagged Gephyrin, including those not associated with detectable VGAT, were associated with co-clustered GABAARg2.  The rate of new YFP-Gephyrin cluster formation by slow accumulation was estimated around 0.0011 ± 0.0002 new puncta/mm/h based on visual identification of new clusters from time-lapse movies of 12-15 DIV neurons. Comparing with a net rate of increase in YFP-Gephyrin cluster density of 0.00227 ± 0.00082 for 12-15 DIV (Fig. 3.4D), and of 0.00232 ± 0.00016 puncta/mm/h for 10-13 DIV (Fig. 3.1C), we suggest that this de novo slow accumulation of YFP-Gephyrin (and 550-VGAT) into clusters represents a major mode of new synapse formation. In support of this idea, new YFP-Gephyrin clusters generated on newly grown dendrite regions observed in the 10-13 DIV time-lapse images (thus without VGAT co-labeling) also appeared to form by gradual accumulation (Fig. 3.7F). Another proportion of new stable synapse may form by splitting of existing synaptic puncta, as there was considerable flux of pre-existing clusters (Fig. 3.4).   102 Figure 3.6. Formation of new inhibitory synapses. A. Dendrite region from a DIV12 YFP-Gephyrin expressing neuron co-labeled with 550-anti-VGAT imaged every 0.5 h for 18 h. Top, pseudocolour image of YFP-Gephyrin fluorescence. Center, overlay of YFP-Gephyrin (green) and 550-VGAT (red). Bottom, pseudocolour image of 550-VGAT fluorescence. Arrow denotes the site at which clustering appears, ultimately consisting of both YFP-Gephyrin and 550-VGAT. Time 0 h represents the frame at which YFP-Gephyrin begins to accumulate. B. Quantitation of fluorescence intensity of YFP- Gephyrin and 550-VGAT from 6 cluster pairs as in A. Fluorescent signal was baselined to the average fluorescence intensity at the region of interest prior to accumulation, normalized to intensity at 7-8 h, and aligned to time 0 h when YFP-Gephyrin first began to accumulate.  C. Magnification of the x-axis in B shows accumulation of 550-VGAT at synaptic sites preceding accumulation of YFP- Gephyrin. D. Another dendrite region as in A. Only YFP-Gephyrin appears at this site, with no 550- VGAT accumulation. E. Average of fluorescence intensity, baselined, (continued next page)  103 Figure 3.6 (continued) normalized, and aligned as in B, of YFP-Gephyrin and 550-VGAT for 5 clusters at sites where only YFP-Gephyrin appears to newly accumulate, as in D. F. Dendrite region from a DIV12 neuron co-expressing mCherry-Gephryin and YFP-GABAARg2, imaged every 1 h for 18 h. Top, pseudocolour image of mCherry-Gephyrin fluorescence. Center, overlay of mCherry-Gephyrin (red) and YFP-GABAARg2 (green). Bottom, pseudocolour image of YFP-GABAARg2 fluorescence. Arrow denotes the site at which clustering appears, consisting of both mCherry-Gephyrin and YFP- GABAARg2. G. Quantitation of fluorescence intensity of mCherry-Gephyrin and YFP-GABAARg2 from 12 cluster pairs as in F. Quantitation was normalized as in B and aligned to time 0 h when mCherry- Gephyrin first began to accumulate. H. Integrated intensity of individual pre-existing YFP-Gephyrin clusters was measured over an 8 h period. Shown here are the top (orange), middle (black), and bottom (cyan) 20% fractions from a total of 134 clusters from 7 cells. I. At left, example image montage of a DIV12 YFP-Gephyrin expressing neuron imaged every 1 h for 18 h, where time 0 h represents when a new YFP-Gephyrin cluster (arrow) begins to accumulate. Following live imaging, neurons were fixed and subjected to immunocytochemistry for endogenous GABAARg2 and VGAT, shown at right. The previously identified new YFP-Gephyrin cluster co-localizes with both markers. Scale bars in A, D, F, I, 2 mm.   Considering the gradual increase in YFP-Gephyrin accumulation that had not yet reached a plateau by 8 hours following new cluster formation (Fig. 3.6A-G, 3.7F), and the overall slow rate of increase in YFP-Gephyrin intensity from averaging all clusters (Fig. 3.1B), we next assessed the change in individual YFP-Gephyrin integrated intensity in relation to initial value. There was a >10-fold range in initial integrated intensity of individual clusters, so we compared specifically the top, middle, and bottom 20% of all clusters. The top and middle fractions essentially maintained their original YFP-Gephyrin intensities, showing little increase (101.6% for top fraction and 107.9% for middle fraction, compared with 100% starting value) over 8 hours (Fig. 3.6H). In contrast, the bottom fraction increased in YFP-Gephyrin integrated intensity to 286% in just 8 hours. Thus the net increase in YFP- Gephyrin puncta intensity among all clusters (Fig. 3.1B) reflects primarily continued recruitment to the smallest/dimmest clusters, without a change in the mid-range or largest/brightest clusters. Since we also show here that new synapses begin as small/dim clusters, these results suggest that, as new synapses form, YFP-Gephyrin accumulates steadily over many hours to reach a target postsynaptic density size. Further, once a target size is reached (above the bottom 20% but including the middle and top 20% in our measures), synapses generally maintain those size differences.   104 3.3.6 Inhibitory synapse formation can occur on both dendritic shafts and dendritic protrusions and on newly formed dendrite branches Imaging of YFP-Gephyrin in DIV 10-15 neurons revealed a majority of clusters stably present on the dendritic shaft, including at the base of protrusions. However, stable clusters were also observed on dendritic protrusions, sometimes with more than one YFP-Gephyrin cluster located on a single protrusion. New clusters were able to form both on the dendritic shaft and on dendritic protrusions (Fig. 3.7A, B). Accumulation of apposed 550-VGAT was associated with some new clusters of YFP-Gephyrin both on dendrite shafts (Fig. 3.6A) and on dendritic protrusions (Fig. 3.7B). We were unable to observe clear transition of YFP- Gephyrin clusters from shaft to protrusion or, conversely, protrusion to shaft. Thus, it is likely that new inhibitory synapses that form on dendrite shafts remain on shafts, and those that form on filopodia or spines remain on the filopodia or spine. We next sought to observe how newly formed dendrites develop inhibitory synapses. Significant dendrite branch growth was observed only in the long 10-13 DIV time-lapse series of YFP-Gephyrin, thus without VGAT co-label. Using phase contrast microscopy to image all neurites, the growth of dendritic arbours was tracked with respect to inhibitory synapse formation. In one example (Fig. 3.7C-E), new YFP-Gephyrin clusters appeared between 20-36 hours after the new dendrite region was established. From observations of other newly growing dendrite branches, YFP-Gephyrin clusters could form as early as 1 hour after the branch’s origin, or even as the growth cone passed. The time of appearance of new YFP-Gephyrin clusters with respect to their position on the new dendrite branch (i.e., their distance from the dendrite’s branch point) was also unpredictable. In the example shown (Fig. 3.7C-E), the last cluster to form was closest to the branch point, while the first cluster to form was halfway between the second and last formed clusters. These observations imply that synapse formation does not proceed in a simple proximal to distal direction on a newly formed dendrite branch in hippocampal culture. New clusters on new dendrites showed accumulation of YFP-Gephyrin with similar kinetics to new clusters  105 forming on pre-existing dendrites (compare Fig. 3.7F with 3.6B, E). Considering as well our observation that VGAT clustering precedes Gephyrin clustering at new synapses (Fig. 3.6), it may be that GABAergic innervation drives the spatiotemporal arrangement of inhibitory synaptogenesis.   3.4 Discussion  We report here multiple features of the long-term dynamics of Gephyrin inhibitory postsynaptic scaffold clusters and associated GABAARg2 and presynaptic VGAT in cultured hippocampal neurons. Entire synapses translocated as a unit over mean path lengths of 5 microns in less than a day, with the Gephyrin, GABAARg2, and VGAT puncta remaining tightly associated. The densities of Gephyrin and VGAT puncta along dendrites fluctuated, mainly representing merging and splitting of pre-existing clusters. Multiple dynamic Gephyrin puncta often rearranged around a large VGAT puncta. There was a slow net increase in Gephyrin cluster density due largely to development of new synaptic clusters. Synapse formation occurred by gradual accumulation of components over several hours, with VGAT accumulating prior to Gephyrin and GABAARg2. Comparing active uptake of a VGAT luminal antibody with post hoc immunofluorescence revealed significant contribution of recycled vesicles to the majority of nascent synapses. New synapses formed predominantly on dendrite shafts, but also on dendritic protrusions, without apparent interconversion, and formed on newborn dendritic branches non-sequentially with respect to dendrite lifetime. Together, these data reveal a complex long-term dynamic nature of inhibitory synapses, with implications for synaptic integration and signaling.  We consistently observed sub-micron translocations of YFP-Gephyrin clusters within seconds to minutes. Previous studies indicate that such mobility is promoted by F-actin and  106  Figure 3.7. Inhibitory synapses form on dendritic shafts, dendritic protrusions, and newly formed dendritic branches. A. Image montage of a DIV 10 YFP-Gephyrin expressing neuron imaged every 1 h. Dendrite shaft and protrusions are outlined in the first frame. A majority of stable YFP-Gephyrin clusters are present on the dendritic shaft, with a few also on dendritic protrusions. Two new YFP-Gephyrin clusters appear during the imaging period, one on the dendritic shaft (arrowhead) at 3.5 h, and another on a dendritic protrusion (arrow) at 9 h. (continued next page)  107 Figure 3.7 (continued) B. Image montage from a DIV12 YFP-Gephyrin expressing neuron co-labeled with 550-anti-VGAT imaged every 0.5 h for 18 h. Dendrite shaft and protrusions are outlined in the first frame. A new YFP-Gephyrin cluster appears on a dendritic protrusion at 5.5 h associated with appearance of apposed 550-VGAT at 4.5 h (arrow). Note also the appearance of a new YFP- Gephyrin cluster on another dendritic protrusion to the left of this highlighted cluster that appears at time 4.5 h without the appearance of apposed 550-VGAT. C. Image montage of a DIV10 YFP- Gephyrin expressing neuron imaged every 1 h for 68 h. Upper panels show the phase contrast image of neurites of both transfected and untransfected cells merged with a pseudocolour representation of YFP-Gephyrin. Lower panels show YFP-Gephyrin fluorescence. Emerging from a primary dendrite at the lower left of the frame is the leading edge of a new dendritic branch (red arrowhead) of a YFP- Gephyrin expressing cell crosses the field of view by 28 h. Several YFP-Gephyrin clusters emerge on the new dendritic branch (magenta, cyan, and yellow arrows, in order of appearance). D. Kymograph analysis of the dendrite region outlined in the last frame of C, pseudocoloured for fluorescence intensity of YFP-Gephyrin. The dotted line on the kymograph represents the leading edge of the growing dendritic branch. E. Quantitation of normalized YFP-Gephyrin fluorescence intensity of clusters corresponding to their coloured arrows in C and aligned to time 0 h defined as the time when the growing dendritic arbour crossed the point at which the YFP-Gephyrin cluster later developed. F. Average of fluorescence intensity of YFP-Gephyrin for 4 clusters appearing on newly-formed regions of dendrite from DIV10-13 neurons. Fluorescent signal was baselined to the average fluorescence intensity at the region of interest prior to accumulation, normalized to intensity at 7-8 h, and aligned to time 0 h when YFP-Gephyrin first began to accumulate.   Scale bars in A, C, 5 mm; B, 2.5 mm.  inhibited by microtubule cytoskeletal networks, and regulated by calcium and synaptic activity (Hanus et al., 2006). Here, by extending the imaging period to many hours, we found a reduction in confinement index with longer imaging intervals (Fig. 3.2), indicating a greater contribution of non-stochastic processes to long term translocations.  Importantly, these translocations of postsynaptic scaffolds were tightly coordinated with corresponding translocations of associated GABAARs and apposed synaptic vesicles, even over distances of several microns and timescales of several hours (Fig. 3.3). Similar movements of entire synapses have been observed for excitatory synaptic structures (Bresler et al., 2001). It is unlikely that this coordinated movement is a passive process, but rather that it involves cross-talk between pre- and postsynaptic components to accomplish rearrangements of both the cytoskeleton and membrane domains to maintain axo-dendritic synaptic specializations in close apposition. Cell adhesion molecules such as Neurexins-Neuroligins and Cadherins may function to maintain the close apposition (Huang et al., 2007; Arikkath and Reichardt, 2008; Siddiqui and Craig, 2010). By controlling their precise positions on the dendritic arbor, this mobility would regulate the integrative contribution of each inhibitory  108 synapse to network signaling. Dynamic positioning of synapses, either as a by-product of neuronal cytoskeletal and membrane dynamics or a tightly locally regulated process, could be one means of regulating inhibitory/excitatory balance within dendritic segments (Liu, 2004). Synapse mobility is likely to be greatest early in development and during the peak phase of synaptogenesis, when synaptic spacing is largely determined, as in our relatively immature (10-15 DIV) cultures. Whether the dense neuropil of an adult brain affects synapse mobility requires further investigation.  In addition to this coordinated mobility of YFP-Gephyrin and 550-VGAT clusters, we also observed apparent merging and splitting events. While we were able to track a number of these events individually (Fig. 3.5), their frequency is also reflected in the high flux of puncta with only a relatively small net change in puncta density (Fig. 3.4). Flux in the density of excitatory postsynaptic scaffold clusters over time along dendritic segments has also been reported, with an interesting synchronization within a given cell (Ebihara et al., 2003). Although merging of clusters has been observed (Gerrow et al., 2006), a major mode for reduction of excitatory postsynaptic scaffold clusters is by elimination or disappearance without obvious merging (Okabe et al., 1999; Niell et al., 2004). For inhibitory postsynaptic scaffold clusters, our observations suggest that merging may be a major mode for reducing the number of synaptic clusters while concomitantly increasing individual synapse size due to the merge (Fig. 3.5A-B), whereas transient, potentially non-synaptic clusters may gradually disassemble.  The frequently observed mobility as well as potential merging and splitting of multiple YFP-Gephyrin clusters surrounding a single large 550-VGAT cluster (e.g., Fig. 3.5C) may indicate flux in postsynaptic elements on a single dendrite region associated with a multiple synapse bouton. GABAergic boutons bearing multiple active zones, each with associated postsynaptic specialization, have been previously described in several brain regions (Studler et al., 2002; Biro et al., 2006). At such GABAergic synapses, spillover of transmitter released  109 from one active zone may activate receptors opposite another active zone and may contribute to high persistent inhibition over stimulus trains (Telgkamp et al., 2004; Wanaverbecq et al., 2008). Dynamic rearrangements of postsynaptic elements associated with multiple synapse boutons, as observed here, may influence the response to spillover and thus modify properties of transmission.  We observed new inhibitory synapse formation on previously established and newly grown dendritic regions, mainly on dendrite shafts but also on protrusions (Fig. 3.7). The gradual increase in YFP-Gephyrin cluster density we observed parallels previously observed developmental maturation of presynaptic release and GABAAR clustering and functionality in culture (Deng et al., 2007). Our results support a mechanism in which YFP-Gephyrin is recruited to new synaptic sites from a cytoplasmic pool over several hours. We were unable to observe small, rapidly moving transport packets of YFP-Gephyrin, which have been suggested to be precursors for synaptic clusters at glycinergic synapses (Maas et al., 2006). Although transport packets of excitatory synaptic scaffold proteins have also been reported (Gerrow et al., 2006), other groups have suggested that, as we observed for inhibitory scaffolds, excitatory scaffolds become clustered from a diffuse pool (Bresler et al., 2001; Okabe et al., 2001b). It is generally agreed that small, rapidly mobile transport packets of synaptic vesicle clusters contribute to new synapses (Ahmari et al., 2000; Bresler et al., 2004), and our 550-VGAT imaging was consistent with this idea. Presynaptic development is thought to precede postsynaptic development at excitatory synapses (Friedman et al., 2000; Okabe et al., 2001a); likewise, we observed clustering of inhibitory synaptic vesicles prior to formation of Gephyrin and GABAAR clusters. However, whereas synaptic vesicles, active zone components, postsynaptic scaffolds and neurotransmitter receptors were recruited within tens of minutes to nascent excitatory synapses (Ahmari et al., 2000; Bresler et al., 2004), accumulation of VGAT, Gephyrin and GABAARs occurred over a period of many hours (Fig. 3.6). Once YFP-Gephyrin clusters reached a certain size, as reflected by  110 integrated intensity, differences in size tended to be maintained (Fig. 3.6H). Similarly, PSD- 95-GFP clusters in vivo were observed to maintain differences in size over days (Gray et al., 2006). The gradual accumulation of postsynaptic components may be triggered in part by Gephyrin oligomerization via its N-terminal G domain and C-terminal E domain (Fritschy et al., 2008), Gephyrin interaction with GABAA receptor subunits (Essrich et al., 1998; Tretter et al., 2008; Saiepour et al., 2010), and Gephyrin interactions with Neuroligin-2 and Collybistin (Harvey et al., 2004; Poulopoulos et al., 2009). While we used only one Gephyrin splice variant in these studies, other variants with different affinities of interaction (Meier et al., 2000) may influence kinetics of Gephyrin accumulation. Perhaps one of the most striking findings here is the involvement of recycled vesicles in new synapse formation (Fig. 3.6); to appear at newly formed synapses, the Oyster-550 anti-VGAT antibody label had to bind the luminal domain of VGAT and be endocytosed during stimulation-induced loading of active boutons. Retrospective immunocytochemistry revealed that contribution of vesicles from previously active terminals to nascent synapses represents the major mode of inhibitory presynaptic development. Whether recycled vesicles contribute to new excitatory synapse formation has not been determined, although recent work has shown that recycling vesicles can be shared among multiple pre-existing neighbouring excitatory boutons (Darcy et al., 2006; Staras et al., 2010). Further quantitative assays to determine the precise contributions of different vesicle pools to nascent inhibitory and excitatory synapses may prove insightful. Whereas the majority of excitatory synapses form on dendritic protrusions (Prange and Murphy, 2001), we observed the majority of inhibitory synapses to form on dendritic shafts. Indeed, inhibitory synaptogenesis is thought to take place without the participation of axonal or dendritic protrusions (Wierenga et al., 2008). However, a proportion of inhibitory synapses on pyramidal neurons in hippocampus and cortex are located on spines (Megias et al., 2001; Knott et al., 2002). We also observed a proportion of YFP-Gephyrin clusters  111 located on dendritic protrusions, and indeed saw new YFP-Gephyrin clusters form on both dendritic shafts and protrusions (Fig. 3.7). We did not observe transitioning of YFP-Gephyrin clusters from shafts to dendritic protrusions or vice versa, and therefore suggest that inhibitory synapses may remain on the morphological structure on which they initially form. Excitatory synapse formation on dendritic protrusions can stabilize those protrusions, promoting further dendrite elongation in a synaptotropic model of dendrite growth in some (Niell et al., 2004; Chen et al., 2010) but not all (Kerschensteiner et al., 2009) neuron types. In cultured rat hippocampal neurons, excitatory synapse formation can stabilize filopodia (Prange and Murphy, 2001), but most new synapses form on pre-existing or newly-formed dendritic protrusions (Okabe et al., 2001a) that develop on dendrite branches over several days following the branch’s outgrowth (Ebihara et al., 2003). Here, we observed inhibitory synapses form anywhere from <1 hour to several days after new dendrite growth, with the temporal sequence of synapse formation not strictly coupled to dendrite lifetime (Fig. 7C-E). Further investigation will be required to better understand any relationships between inhibitory synaptogenesis and dendrite morphogenesis, particularly in vivo where contact by GABAergic axons is more precisely patterned than in culture. Taken together, our results show complex dynamics of inhibitory synapses over long time courses. These dynamics may be reflective of the perpetual remodeling of synapses that takes place in response to signals arising from molecular interactions and synaptic activity (Liu, 2004; Hartman et al., 2006; Huang et al., 2007; Siddiqui and Craig, 2010). Understanding these complex dynamics may be critical in deciphering a number of neuropsychiatric diseases that arise from GABAergic dysfunction or imbalances in inhibition/excitation.    112 4  Discussion 4.1 Summary of findings The objectives of this work were to determine: a) What are the functions of a-Nrxns in induction of glutamatergic and GABAergic synapses, and how is alternative splicing of Nrxns modulated by development and activity? b) How do inhibitory synapses develop over time in living neurons with respect to the dynamics of synapses and the spatiotemporal recruitment of core inhibitory synapse components? Previous studies have shown the utility of the co-culture assay in determining the synaptogenic activity of Nrxns and Nlgns (Scheiffele et al., 2000; Graf et al., 2004; Nam and Chen, 2005). However, since initial reports suggested that a-Nrxns did not bind to Nlgns (Ichtchenko et al., 1995), a majority of studies have focused on the interaction between b- Nrxn isoforms and various Nlgns. Extension of co-culture experiments into other in vitro systems have highlighted the important roles for different splice forms of Nrxns and Nlgns in initiation and maturation of glutamatergic and GABAergic synapses. Strikingly, a trans- synaptic splice code of not only Nrxn-Nlgn interaction has been established, but this splice code appears to directly influence the strength of the synaptogenic activity conferred by these proteins. For instance, b-Nrxns that contain an insert at S4 have lower affinity for Nlgns containing an insert at site B and those lacking an insert at site B (Boucard et al., 2005; Chih et al., 2006; Graf et al., 2006). However, these b-Nrxn(+S4) isoforms are still able to modestly cluster Ngln-1/3/4 and the excitatory postsynaptic scaffold PSD-95 (Graf et al., 2006), and fairly robustly cluster Ngln-2 and the inhibitory scaffold gephyrin (Graf et al., 2006). In contrast conflicting reports suggest that Nlgn-1(+B) overexpression may greatly  113 enhance excitatory but not inhibitory innervation onto the overexpressing neuron (Graf et al., 2006) or have only modest effects (Levinson et al., 2005), while Nlgn-1(-B) appears to enhance both excitatory and inhibitory innervation (Graf et al., 2006). Contradictory results have also been found for overexpression of Nlgn-2, which constitutively lacks an insert at site B; its overexpression can either greatly enhance both excitatory and inhibitory innervation (Graf et al., 2006) or have less pronounced effects (Levinson et al., 2005). These discrepancies may arise from variations in culture conditions or expression levels of each protein, and thus inherently highlight the complexity of the splice interaction code in regulating synaptogenesis. The results of Chapter 2 show a new role for a-Nrxns in synapse induction ability, through previously uncharacterized interactions with Nlgns, and provide further insight into the roles of different Nlgn isoforms in synapse organization. We have shown that, as with previous findings using the co-culture assay, b-Nrxns are able to induce clustering of both excitatory and inhibitory postsynaptic components. However, a-Nrxns are only able to induce clustering of the inhibitory postsynaptic component gephyrin, along with the inhibitory synapse specific Nlgn-2 but not other Nlgn isoforms. Moreover, a-Nrxn can lead to clustering of the synaptic GABAAR subunit g2, but not the extrasynaptic subunit a5. Although these clustering abilities were not as profound as for b-Nrxns, they suggest that a- Nrxns have a specific role in induction of GABAergic synapses. A study published just prior to the results described here further confirmed the binding of a-Nrxns to Nlgns (Boucard et al., 2005). They observed a diminished affinity of a-Nrxns/Nlgns compared to b-Nrxns, paralleling the results of synaptogenic activity found here. However, they further showed that inclusion of an insert at site B of Nlgns completely abrogated their  binding to a-Nrxns, dissimilar from the finding that site B inclusion only reduces binding to b-Nrxns without complete abolishment (Boucard et al., 2005). We have further shown a developmental increases in the proportion of both a- and b-Nrxns lacking insertion at S4, mirroring the  114 developmental increase in excitatory versus inhibitory synaptogenesis (Tyzio et al., 1999). Although these methods for determination of alternative splicing of Nrxns were relatively crude, they provide the first evidence that alternative splicing may be modulated in a physiologically relevant manner.  The dynamic aspects of synapse formation are just beginning to be understood, due primarily to advances in live imaging techniques. However, a majority of these studies have focused on excitatory synapses (Okabe et al., 1999; Ahmari et al., 2000; Friedman et al., 2000; Bresler et al., 2001; Marrs et al., 2001; Okabe et al., 2001a; Okabe et al., 2001b; Prange and Murphy, 2001; Ebihara et al., 2003; Niell et al., 2004; Gerrow et al., 2006; Graf et al., 2006; Gray et al., 2006; Meyer and Smith, 2006; Ruthazer et al., 2006; Minerbi et al., 2009). Several key findings have emerged from these studies. First, following initial axo- dendritic contact, presynaptic development appears in a majority of cases to precede recruitment of core postsynaptic components. While presynaptic development occurs through rapid recruitment of mobile transport packets to nascent sites of contact, postsynaptic development lags behind presynaptic development by tens of minutes, and occurs primarily through accumulation of components from diffuse cytoplasmic pools. Several reports, though, suggest that not only may postsynaptic development precede presynaptic development, but that these postsynaptic elements may be rapidly recruited from vesicular transport packets. These newly formed synapses appear competent for bona fide synaptic transmission, although a protracted maturation process may continue for several hours to days before a fully functioning synapse, capable of differentially responding to various types of stimuli, is apparent. Our results from Chapter 3 have extended the work on the dynamic aspects of excitatory synapses to inhibitory synapses. We have shown that entire synapses comprised of both pre- and postsynaptic elements were mobile as a single structure over paths of greater than 5 mm over several hours, reminiscent of excitatory synapses (Bresler et al., 2001). A majority of synapses were confined to specific  115 subdomains while still being mobile within that region, as has previously been shown for gephyrin clusters on short time scales (Hanus et al., 2006). However, upon longer imaging intervals, non-stochastic movements of gephyrin clusters were observed, leading to escape of clusters from their confined region and movement over greater path lengths. Overall, new synapse formation on developing dendritic arbours was seen to occur gradually, with a high frequency of cluster splitting and merging events showing that there was significant flux in the number of inhibitory clusters with only a modest net change in their density. When newly-forming individual synapses were observed closely, both pre- and postsynaptic elements were seen to accumulate gradually over time, with presynaptic development preceding postsynaptic development. Although the time course of component accumulation is not identical to that for excitatory synapses, the general features of pre- preceding post and gradual accumulation from diffuse pools are similar  (Friedman et al., 2000; Bresler et al., 2001; Okabe et al., 2001b). Strikingly, our results show that previously recycling synaptic vesicles comprise a major component of new vesicle accumulation at nascent synaptic sites, with nearly 80% of nascent inhibitory synapses containing presynaptic vesicles from previously recycling sites. Altogether, our results highlight the considerable dynamics of existing and nascent inhibitory synapses, and provide new context for the general cellular mechanisms of synapse formation in the existing milieu of excitatory and synapses.  4.2 A glut of synaptogenic molecules  An obvious question that arises from the study of synaptogenic proteins is: why are there so many? Results shown here (Chapter 2) and elsewhere have found that, even for Nrxn/Nlgn, a complex binding code exists, where trans-synaptic interaction affinity and synapse organizing function is dictated by different Nrxn and Nlgn gene products and splice variants. If this were not complicated enough, recent evidence has shown there are yet other  116 postsynaptic receptors for presynaptic Nrxns. LRRTM1/2 specifically promote excitatory synapse formation by binding to a- or b-Nrxns lacking an insert at S4, with no appreciable binding to splice variants containing the S4 insert (de Wit et al., 2009; Ko et al., 2009; Siddiqui et al., 2010). Additionally, the newfound tripartite interaction of Nrxn-Cbln1-GluRd2 has been shown to mediate synapse formation in several brain regions. Now we must ask not only why there are so many different classes of synapse organizing molecules, but why there are so many different postsynaptic receptors for Nrxns.  4.2.1 Synapse initiation versus synapse maturation One simple explanation for the large number of synaptogenic molecules is that there is functional redundancy between them so as to ensure that genetic defects in one or more synaptogenic molecules do not have overwhelmingly deleterious effects on nervous system function. This is an appealing explanation if we consider the phenotype of many synaptogenic proteins in the co-culture assay: their presentation to axons or dendrites induces the clustering of pre- or postsynaptic proteins, respectively. These findings imply that most synaptogenic molecules function as nucleation factors; their clustering leads to locally increased concentrations of protein-protein interaction domains that, in turn, recruit other proteins to form a network of interacting proteins that are the hallmark of a functional synapse. Consider the phenotype of the Nlgn1,2,3 triple knockout mouse: synapses are grossly normal except in one specific brain region (Varoqueaux et al., 2006). These results imply that there are compensatory mechanisms, such as other synaptogenic molecules, that act in synapse formation in these mice, despite their lack of Nlgns. In fact, the phenotype of triple knockout mice is more severe than the Nlgn-1 or Nlgn-2 knockout mice alone (Chubykin et al., 2007; Poulopoulos et al., 2009), suggesting redundancy even among the Nlgns.  117  It is unlikely that this basic explanation is complete, though. The role of certain synaptogenic molecules may be beyond initial cell-cell adhesion and recruitment of core synaptic components, and rather in regulating maturation of synapses or synaptic plasticity. Recent evidence for convergent and divergent roles of Nlgns and LRRTMs have highlighted these potential differences. Whereas knockdown of Nlgn-1 and -3 early in development did not lead to alterations in excitatory synapse numbers or synaptic transmission, knockdown of LRRTM1 and 2 specifically decreased AMPAR transmission (Soler-Llavina et al., 2011). Combined knockdown of Nlgns and LRRTMs led to a further reduction in excitatory synapse maturation, also decreasing NMDAR activity. In contrast, knockdown of LRRTMs later in development caused no appreciable synaptic defects, while Nlgn knockdown decreased NMDAR signaling. These results suggest that Nlgns and LRRTMs may cooperate early in development for synapse maturation rather than initial formation, with LRRTMs more strongly influencing synaptic functions, while Nlgns may function later in development in maintaining proper synaptic physiology. Considering that Nlgns and LRRTMs share several overlapping Nrxn ligands, how these alternate functions are resolved remains an outstanding question. Moreover, N-cadherin appears to be important for modulating Nlgn- dependent synaptogenesis in hippocampus (Aiga et al., 2011). Whether and how other synapse organizing molecules function in concert or opposition is of continued interest.  4.2.2 Synapse specificity Another possible reason for the multitude of synaptogenic molecules is in conferring synapse specificity. The regional and subcellular expression of subsets of synapse organizing factors may lead to their involvement in validation of synapses at appropriate spatial locations. For instance, Nlgn-2 is localized specifically to inhibitory synapses, but its knockout mainly affects perisomatic hippocampal synapses (Poulopoulos et al., 2009). Nlgn- 2 interacts with all a- and b-neurexin isoforms, yet Nlgn-1(+B), the predominant  118 hippocampal isoform which specifically induces excitatory synapses, interacts with only the b-Nrxns (Siddiqui and Craig, 2010). However, what governs the appropriate matching of Nlgn and Nrxn isoforms remains poorly understood. It is plausible to imagine a scenario where an innervating axon expresses only the Nrxn isoform(s) that will significantly bind the appropriate Nlgn or LRRTM isoforms on a target dendrite, the postsynaptic receptors even being targeted to specific subcellular domains by as yet undetermined factors. Indeed, LRRTM2 and other synaptogenic molecules often display laminae-specific staining patterns, suggesting their potential involvement in synapse specificity (Appendix A). Only those axon- dendrite Nrxn-Nlgn/LRRTM interactions that are reinforced will lead to synapse formation. For instance, an innervating axon that expresses Nrxn-1b(+S4) will not lead to development of an LRRTM-induced excitatory synapse, but may lead to development of a Nlgn-1(+B)- induced excitatory synapses. We have attempted to initially address this problem in Chapter 2 by assaying the gross expression patterns of various Nrxn isoforms both during development and in different brain regions. Further profiling of regional expression patterns of the different splice variants of Nrxns and Nlgns as well as LRRTMs and other synaptogenic molecules may provide insight into their roles in synapse specificity.  Further specificity may be conferred by negative regulatory factors. The recently characterized tripartite Nrxn-Cbln1-GluRd2 interaction has been implicated in synapse formation in the cerebellum (Uemura et al., 2010). New data suggest that Cbln1 may also act in inducing Nrxn-GluD1-dependent excitatory synapses specifically in entorhinal projections into the hippocampus (Ryu et al., 2011). Interestingly, Cbln1 binding to Nrxn(+S4) variants can interfere with Nlgn-dependent synaptogenesis, sparing LRRTM- dependent synapse formation, presumably since LRRTMs only bind Nrxn(-S4) variants (Matsuda and Yuzaki, 2011). It thus appears that the complicated trans-synaptic code for  119 Nrxn-Nlgn/LRRTM-dependent synaptogenesis is subject to further regulation, and it is possible that other molecules can act similarly to limit synaptogenesis.  4.3 Dynamics of synaptogenesis It is now clear that synapse formation is a kinetic process, consisting not of discreet steps but of a fluid continuum of events. Because of this fact, we may be unable to ascribe hard and fast rules to the cellular mechanisms of synaptogenesis; instead, we may only be able to infer general principles from experimental data. We must now also face the challenges of bridging our understanding of molecular and cellular mechanisms of synapse organization in an effort to more completely describe these complex processes.  4.3.1 Spatiotemporal order of inhibitory synapse formation For excitatory synapses, varying results from different laboratories using subtly different experimental systems have lead to conflicting views as to the precise order in which synapses form. Our results for inhibitory synapses, shown in Chapter 3, suggest that presynaptic differentiation precedes postsynaptic differentiation, and that both processes occur slowly over several hours. These data are in agreement with several studies of excitatory synapses showing that innervating axons cluster presynaptic proteins shortly after dendritic contact, and that postsynaptic proteins accumulate from diffuse cytoplasmic pools only after presynaptic differentiation (Bresler et al., 2001; Okabe et al., 2001a; Bresler et al., 2004). Unlike other studies of excitatory synapses, we saw no evidence of postsynaptic development preceding presynaptic development, nor of rapid recruitment of inhibitory postsynaptic protein transport packets to nascent contact sites. Our data are consistent with a model whereby presynaptic structures instruct postsynaptic development. In this model, axo-dendritic contact leads to accumulation of PTVs and STVs at contact sites, possibly by nucleation mediated by synaptogenic molecules like Nrxns. Subsequently, postsynaptic  120 proteins begin to accumulate, possibly as a result of clustering of postsynaptic synaptogenic molecules like Nlgns by presynaptic Nrxns. The temporal order of CAM recruitment to nascent contacts remains unclear and will require further investigation. Furthermore, it is unknown whether synaptic vesicle release and postsynaptic receptor activation is required for synapse development per se, although if this is the case it may explain our observed discrepancies in the timing of postsynaptic protein recruitment between excitatory and inhibitory synapses. Whereas excitatory scaffolds may be recruited in tens of minutes after presynaptic development, inhibitory scaffolds accumulate over at least two hours. At excitatory synapses, receptor activation, particularly NMDAR signaling, may lead to downstream signals directly activated by calcium influx that may hasten postsynaptic development. On the contrary, activation of GABAARs at developing inhibitory synapses does not have any known direct signaling functions. Therefore, postsynaptic development would be dependent solely on the biophysical properties of protein aggregation or, conceivably, upon activation of adjacent excitatory synapses and propagation of their signals to nearby inhibitory synapses.  4.3.2 Putative mechanisms of synapse mobility We consistently observed large-scale movements of entire inhibitory synaptic structures over minutes to hours. At least two possible mechanisms can explain this mobility. In one scenario, proteins are rapidly recruited to the leading edge of mobile synapses while proteins at the lagging edge are disassembled, resulting in a net observed change in the position of the synapse. However, imaging of mobile synapses at high frequency suggested that synapses move as a complete structure spanning both the pre- and postsynaptic compartment. Since both compartments are intimately associated with the cytoskeleton, this mobility likely requires regulation of actin and microtubule polymerization. What signaling mechanisms are at play to enable such cytoskeletal rearrangements are as yet unknown.  121 Moreover, we frequently saw short bursts of synapse mobility followed by periods of stability. This saltatory movement may reflect the transient engagement of molecular motor-based trafficking, although what, if any, motors are involved is yet to be determined.  An advantage of our in vitro live cell imaging approach is our ability to observe the relatively flat dissociated hippocampal cultures at various imaging intervals with high resolution and low phototoxicity. Many YFP-Gephyrin structures we observed were at the theoretical limit of resolution for conventional microscopy. However, it remains to be seen whether the dynamics of inhibitory synapses we observed in culture are also present in vivo. Clearly, the in vivo environment is vastly different from culture conditions, where not only is there a dense neuropil which may affect synapse mobility, but also there are specific patterns of GABAergic innervation onto, for example, hippocampal pyramidal cells that depend on the inhibitory cell type. However, because of the technical limitations of in vivo imaging techniques like multiphoton microscopy, such as lower numerical aperture objectives, relatively slow frame rates, and limited tissue penetration, applying certain questions that are addressable in culture to in vivo remains difficult. We look forward to the continued advances in microscopy that will eventually facilitate answering such questions.  4.4 Implications for health and disease An increasing consensus among neuroscientists is that many neurological and psychiatric disorders are diseases of the synapse; that is, improper development of synapses or their dysfunction may directly lead to the pathophysiology of disease states. The expanding field of neurogenetics is beginning to define critical genetic factors that give rise to disease. Emerging evidence has implicated several synaptic CAMs in diseases such as schizophrenia and autism spectrum disorders (ASDs), along with GABAAR dysfunction in depression, anxiety, and epilepsy.  122 4.4.1 Synapse organizing molecule mutations in ASD and schizophrenia  ASDs are characterized by deficits in social interaction, stereotyped behaviours, and impaired language skills (Kuzirian and Paradis, 2011). They occur in 0.2-1% of the population and in 80% of cases there is strong genetic influence. However, there is a considerable degree of variability in the genetic factors leading to ASDs. Along with several other genes, Nrxns and Nlgns have recently been implicated in the pathophysiology of ASDs (Jamain et al., 2003; Jamain et al., 2004; Feng et al., 2006; Gusella et al., 2008; Sommer et al., 2008a; Sommer et al., 2008b; Sudhof, 2008; Blundell et al., 2010; Voineskos et al., 2011; Yu et al., 2011). In one example, a percentage of ASD patients were found to have a variation in the Nlgn-3 gene that leads to a mutation of arginine 451 to cysteine (Jamain et al., 2003). Interestingly, introduction of the mutated gene into mice leads to increases in GABAergic function and behaviours reminiscent of ASDs (Tabuchi et al., 2007). Further in vitro and animal studies of Nrxn and Nlgn gene variants associated with ASDs in human patients have begun to unravel the potentially critical role these genes play in disease etiology and may assist in development of therapies for this cluster of disorders (Chih et al., 2004; Comoletti et al., 2004; Chubykin et al., 2005; Hines et al., 2008; Colicos et al., 2009; Ehrenreich et al., 2009; Powell et al., 2009; Sudhof et al., 2009; Zhang et al., 2009; Blundell et al., 2010; Etherton et al., 2011a; Etherton et al., 2011b).  Schizophrenia is characterized by delusions, hallucinations, disordered thoughts and speech, and social deficits (Lips et al., 2011). It, like ASDs, can affect up to 1% of the population and has roughly 80% heritability, with high variability in genetic risk factors. Although the symptoms of schizophrenia appear very different from ASDs, Nrxns and Nlgns now also seem to play a role in the former, with several gene mutations linked to schizophrenia (Sand et al., 2006; Bennett, 2008; Collier et al., 2009; Cichon et al., 2011; Lips et al., 2011; Sun et al., 2011; Voineskos et al., 2011). Considerable challenges still  123 exist in development of mouse models for schizophrenia, though, as it is difficult to assay the hallmark behaviours of schizophrenia in mice.  4.4.2 GABAergic dysfunction in neurological and psychiatric disorders The classic anxiolytic and anti-epileptic drugs, benzodiazepines, are well-known to bind to GABAARs (Tan et al., 2011). Subsequently, several mutations in GABAARs have been implicated in familial epilepsy, including in the g2 and several a subunits (Spreafico et al., 1993; Jones-Davis and Macdonald, 2003; Loup et al., 2006; Eugene et al., 2007; Fritschy, 2008). Interestingly, it appears that some mutations directly affect channel function, whereas others affect channel assembly, transport, and targeting, highlighting the critical role of protein trafficking in synaptic function. Growing evidence has implicated GABAergic signaling deficits in various mood disorders, including anxiety, major depressive disorder, bipolar disorder, and suicide (Luscher et al., 2011b). Additionally, changes in GABAARs, GABA synthetic enzymes, and GABA transporters have been observed in patients with schizophrenia (Fritschy and Brünig, 2003). Along with the observation that Nrxn/Nlgn-based mouse models of ASDs have specific deficits in GABAergic transmission, it now appears that inhibitory synapses and the balance of excitation to inhibition in the brain may be of critical importance in determining a healthy neural state.  4.5 Perspectives and future directions In this dissertation, I have investigated the role of a-Nrxns in GABAergic synapse formation and how these inhibitory synapse are highly dynamic. Because of the implications of inhibitory synapses in disease states, a further understanding of inhibitory synaptogenesis may prove valuable in treatment of the numerous disorders associated with GABAergic dysfunction.  124  A more thorough understanding of the molecular mechanisms of inhibitory synapse organization is required, though, as, compared to excitatory synapses, relatively few proteins specifically involved in inhibitory synaptogenesis are known. What other proteins can influence development of inhibitory synapses? How do these proteins effect GABAergic synapse-specific roles, namely with respect to their spatial expression and subcellular targeting? Can the multitude of known synaptogenic factors act in concert or opposition to regulate synapse formation? Beyond initial synapse development, can synapse organizing factors influence plasticity at mature synapses?  In addition, the cellular mechanisms of both excitatory and inhibitory synapse formation and synapse dynamics remain obscure. Is there a set of rules that govern the timing of synapse development? Can synaptic activation or engagement of biochemical signaling pathways modify how proteins are trafficked to nascent synapses? What are the roles of cytoskeletal modification and molecular motors in determining delivery of synaptic proteins to developing synapses and the mobility of mature synapses? Do the results observed for inhibitory synapse formation in vitro translate to the intact brain?  Taken together, the results of previous studies, those presented in this dissertation, and the remaining unanswered questions paint a sorely incomplete picture of how synapses are formed and modified. Moving forward, addressing some of these issues may seem like a daunting task, but will inevitably provide valuable insight into the mysteries of the mind.        125 References  Abe K, Chisaka O, Van Roy F, Takeichi M (2004) Stability of dendritic spines and synaptic contacts is controlled by alpha N-catenin. Nat Neurosci 7:357-363. Ahmari SE, Buchanan J, Smith SJ (2000) Assembly of presynaptic active zones from cytoplasmic transport packets. Nat Neurosci 3:445-451. Aiga M, Levinson JN, Bamji SX (2011) N-cadherin and neuroligins cooperate to regulate synapse formation in hippocampal cultures. J Biol Chem 286:851-858. 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