Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Deciphering the genetics mechanisms that inhibit synapse formation Fortes, Ethan 2020

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Notice for Google Chrome users:
If you are having trouble viewing or searching the PDF with Google Chrome, please download it here instead.

Item Metadata

Download

Media
24-ubc_2020_may_fortes_ethan.pdf [ 2.72MB ]
Metadata
JSON: 24-1.0390272.json
JSON-LD: 24-1.0390272-ld.json
RDF/XML (Pretty): 24-1.0390272-rdf.xml
RDF/JSON: 24-1.0390272-rdf.json
Turtle: 24-1.0390272-turtle.txt
N-Triples: 24-1.0390272-rdf-ntriples.txt
Original Record: 24-1.0390272-source.json
Full Text
24-1.0390272-fulltext.txt
Citation
24-1.0390272.ris

Full Text

DECIPHERING THE GENETIC MECHANISMS THAT INHIBIT SYNAPSE FORMATION by  Ethan Fortes  B.Sc., The University of British Columbia, 2016  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Zoology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2020  © Ethan Fortes, 2020   ii  The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, a thesis entitled:  DECIPHERING THE GENETIC MECHANISMS THAT INHIBIT SYNAPSE FORMATION  submitted by ETHAN FORTES in partial fulfillment of the requirements for the degree of MASTER OF SCIENCE in ZOOLOGY  Examining Committee: KOTA MIZUMOTO, ZOOLOGY Supervisor  DON MOERMAN, ZOOLOGY     Supervisory Committee Member BEN MATTHEWS, ZOOLOGY  Additional Examiner   Additional Supervisory Committee Members:  DOUG ALLAN, CELLULAR & PHYSIOLOGICAL SCIENCES Supervisory Committee Member  iii  Abstract In animals, precise behavioural outputs require the development and maintenance of precise synaptic contacts between neurons and their cellular targets. Understanding how cells in the nervous system exhibit fine communication at the synapse level is crucial to elucidating principles underlying fine control in biological systems. The development and maintenance of precise synapses requires inhibitory mechanisms to prevent excessive neuronal inputs. Impaired inhibition of synapse formation is implicated in serious neurological conditions including autism spectrum disorder and intellectual disability. One mechanism for the inhibition of synapse formation involves Traf2- and NCK- Interacting Kinase (TNIK). Mammalian TNIK is required for proper axon guidance and cell migration, and more recently has been found by our group to negatively regulate the formation of synapses in C. elegans orthologue mig-15. However, the mechanism for mig-15-mediated loss of synapses is unknown. Here we show that mig-15/TNIK functions to inhibit synapse formation via wdr-24, a component of the GATOR2 complex upstream of TORC1, a master regulatory hub controlling growth and development. These results implicate TORC1 as a possible negative regulator of synapse formation and may lead to a deeper understanding of how a highly conserved growth and development signalling network controls the formation of neural circuits.    iv  Lay Summary The nervous system is comprised of many cells, called neurons, which form connections (called synapses) with other cells. Synapses must form with the correct targets, and many neurological disorders such as autism and intellectual disability are linked to uninhibited synapse formation. In this thesis, I studied the microscopic nematode C. elegans to explore this process. The worm contains only 302 neurons and is transparent, hence it is easy to observe their neurons. I describe a gene called mig-15, which inhibits the formation of synapses. I engineered worms with excess mig-15, reducing synapses and disrupting locomotion. Then, using random mutations, I screened worms for normal locomotion, to find a gene which mig-15 requires to function. I identified a gene I call wdr-24 as ‘downstream’ of mig-15. Like mig-15 worms, wdr-24 mutants have more synapses. wdr-24 is part of the TOR pathway which is not known to regulate synapse formation in worms.  v  Preface All work was performed in Dr. Kota Mizumoto’s lab at UBC. Dr. Kota Mizumoto and I conducted all experiments. Dr. Mizumoto cloned mig-15 and generated pan-neuronal expression clones used to generate mizIs33, mizIs34, and mizIs34. Niousha Gazor, an undergraduate research assistant, and Clare Venn Skillman, an undergraduate volunteer, assisted with generating strains. Dr. Stephane Flibotte conducted the analysis of whole genome sequencing. I performed all other experiments and data analysis.  Data in Figure 1 was featured in a peer reviewed journal, on which I am a co-author: Chen, Xi, Akihiro C E Shibata, Ardalan Hendi, Mizuki Kurashina, Ethan Fortes, Nicholas L Weilinger, Brian A Macvicar, and Hideji Murakoshi. 2018. “Rap2 and TNIK Control Plexin-Dependent Tiled Synaptic Innervation in C . Elegans.” ELife 7:e38801: 1–25.   vi  Table of Contents  Abstract ......................................................................................................................................... iii Lay Summary ............................................................................................................................... iv Preface .............................................................................................................................................v Table of Contents ......................................................................................................................... vi List of Tables ..................................................................................................................................x List of Figures ............................................................................................................................... xi List of Abbreviations .................................................................................................................. xii Acknowledgements .................................................................................................................... xiii Chapter 1: Introduction ................................................................................................................1 1.1 Caenorhabditis elegans .................................................................................................. 1 1.1.1 Nervous system of C. elegans ..................................................................................... 2 1.1.2 Genetics of C. elegans ................................................................................................ 2 1.2 Traf-2 and NCK-Interacting Kinase/mig-15 in the nervous system ............................... 3 1.2.1 Structure of TNIK ....................................................................................................... 3 1.2.2 Clinical relevance of TNIK ......................................................................................... 4 1.2.2.1 Neurological disorders associated with TNIK .................................................... 4 1.2.2.2 TNIK and cancer ................................................................................................. 4 1.2.3 TNIK and Wnt signaling ............................................................................................. 4 1.2.4 Axon guidance ............................................................................................................ 5 1.2.5 The role of TNIK in synapse formation and specificity ............................................. 6 1.3 Patterned synapse formation ........................................................................................... 7  vii  1.3.1 Synaptic tiling in C. elegans ....................................................................................... 7 1.3.1.1 The role of plx-1, rap-2, and mig-15 in synaptic tiling ....................................... 9 1.4 The TOR pathway ......................................................................................................... 10 1.4.1 TOR in C. elegans..................................................................................................... 13 1.4.2 The role of TOR in the nervous system .................................................................... 13 1.4.2.1 TOR in the developing brain............................................................................. 13 1.4.2.2 TOR at the synapse ........................................................................................... 15 1.4.2.3 Autophagy at the synapse ................................................................................. 16 1.5 Thesis Objectives .......................................................................................................... 18 1.5.1 Identification of a genetic mechanism for the negative regulation of synapse number by TNIK/mig-15.................................................................................................................... 18 1.5.2 Characterization of the GATOR complex in relation to mig-15-mediated synapse formation ............................................................................................................................... 18 Chapter 2: Materials & Methods ...............................................................................................19 2.1 Worm Culture and Strains ............................................................................................ 19 2.2 Nomenclature of C. elegans .......................................................................................... 20 2.3 DNA Extraction and Polymerase Chain Reaction ........................................................ 20 2.4 Confocal microscopy .................................................................................................... 23 2.5 Plasmid construction ..................................................................................................... 23 2.6 Multiple Sequence Alignment ...................................................................................... 23 2.7 Strain integration ........................................................................................................... 24 2.8 EMS mutagenesis.......................................................................................................... 24 2.9 Chromosomal mapping using SNPs ............................................................................. 24  viii  2.10 CRISPR/Cas9 ................................................................................................................ 25 Chapter 3: Screening for genetic mechanism of MIG-15-mediated inhibition of synapse formation ......................................................................................................................................28 3.1 mig-15 is a negative regulator of synapse formation .................................................... 28 3.1.1 Pan-neuronal overexpression of mig-15 dramatically reduces locomotor coordination .......................................................................................................................... 29 3.1.2 Pan-neuronal overexpression dramatically reduces synaptic fluorescence .............. 30 3.2 Forward screen to identify suppressors for mig-15-mediated inhibition of synapses .. 31 3.2.1 Suppression of mizIs33 Unc phenotype rescues loss of synaptic fluorescence ........ 32 3.2.2 SNP mapping reveals the location of sup3 and mizIs33 to be chromosome III ....... 33 3.2.3 Whole Genome Sequencing identifies Y32H12A.8 as a candidate gene .................. 33 3.2.4 Multiple sequence alignment of Y32H12A.8 and its orthologs ................................. 36 3.2.5 Y32H12A.8 or wdr-24 rescues loss of synapses in mizIs33 ...................................... 38 3.2.6 Conserved domain of wdr-24 rescues increased synapse number in wdr-24;mizIs33 38 3.2.7 wdr-24 mutants have increased synapse number in DA9 ......................................... 38 Chapter 4: Separating MIG-15-mediated inhibition of synapse formation from synaptic tiling ...............................................................................................................................................40 4.1 MIG-15-GATOR axis controls synapse formation independently of synaptic tiling ... 40 Chapter 5: Discussion ..................................................................................................................41 5.1 wdr-24 suppression of mig-15(OE) phenotypes ........................................................... 41 5.2 mig-15 functions through GATOR2 to inhibit synapse formation ............................... 42 5.3 Neuron-specific knockdown of Raptor/DAF-15 .......................................................... 42  ix  5.4 Amino acid sensing role of GATOR2 .......................................................................... 43 5.5 Non-conserved domain of wdr-24/Y32H12A.8 ........................................................... 44 5.6 Inhibition of GATOR1 by GATOR2 may be necessary for mig-15(OE) induced loss of synapses and coordinated locomotion. ..................................................................................... 45 References .....................................................................................................................................48   x  List of Tables  Table 1. List of strains used in this work ...................................................................................... 19 Table 2. Primers used for genotyping and cloning ....................................................................... 22 Table 3. Candidate mig-15(OE) suppressor genes from whole genome sequencing. .................. 34   xi  List of Figures  Figure 2. mig-15 is a negative regulator of DA9 synapse formation. ........................................... 29 Figure 3. Pan-neuronal overexpression of mig-15 decreases pan-neuronal synaptic fluorescence........................................................................................................................................................ 31 Figure 4. Forward screen of the Unc phenotype of mig-15(OE) reveals a suppressor mutation. . 32 Figure 5. wdr-24 rescues synaptic defects of mig-15 overexpression .......................................... 35 Figure 6. Multiple alignment reveals a SF missense mutation outside the conserved domain of wdr-24. .......................................................................................................................................... 37 Figure 7. wdr-24 enhances synaptic tiling defect of plx-1 ............................................................ 47  xii  List of Abbreviations  L1,L2,L3,L4 Larval stage 1,2,3,4 C. elegans Caenorhabditis elegans DA Dorsal A class DNC Dorsal Nerve Cord GDP Guanosine diphosphate GEF GTP Exchange Factor GFP Green Fluorescent Protein GTP Guanosine triphosphate JNK c-Jun N-terminal kinase KD Knockdown MAPK4 Mitogen-Activated Protein Kinase 4 mTOR mechanistic Target of Rapamycin OE Overexpression PCR Polymerase Chain Reaction Ste20 Sterile 20 protein TNIK Traf2- and Nck-interacting kianse Unc Uncoordinated VNC Ventral Nerve Cord     xiii  Acknowledgements I extend my deep gratitude to Dr. Kota Mizumoto for his guidance and support. His unrelenting curiosity and drive makes him an inspiring mentor. I am also very thankful for the members of my supervisory committee, Drs. Don Moerman and Doug Allan, who encouraged me and offered helpful suggestions. Furthermore, I am grateful to Drs. Doug Altschuler, Mike Gordon, Linda Matsuuchi, and Dolph Schluter for their sage advice. I am thankful for the support and comradery of my lab mates, both past and present. My fellow graduate students, Kelly Chen, Menghao Lu, Ardalan Hendi, and Mizuki Kurashina, have all generously shared their time and energy to improve my daily productivity and sanity. Likewise, I’m lucky to have worked (albiet briefly) with our talented new post-doctoral fellow, Riley St. Clair. I would also like to extend my heartfelt appreciation for undergraduate students and research assistants in our lab. Jane Wang, Niousha Gazor, Clare Venn Skillman, Arpun Johal, Minnie Kim, Ali Murtaza, and Jeffrey Lin have all, in their own ways, made my experience one I will always cherish.  A special thanks is owed to members of the Moerman Lab. Mark Edgley, whose good humour and invaluable advice I could not have done without, Dr. Stephane Flibotte, who generously offered his bioinformatic expertise to assist in my genetic screen, Vinci Au and Erica Li-Leger, both of whom have been a delightful presence in the lab and patiently taught me microinjection and CRISPR.  I am eternally grateful for the friendships I have made in my time here: Abdalla, Amelia, Mriga, Meghan, Glory, Tashana, Priya, Payel, and Kevin. Deserving a special mention is Peter, who greeted me every evening with a warm smile and kind words. My most heartfelt thanks are reserved for my loving family and partner. My parents, Colin and Anne-Maree, have always been supportive in every way possible. Finally, my partner Zephi has kept me grounded through this entire journey, and for this my words cannot express my thanks.   1  Chapter 1: Introduction In this chapter I will describe Caenorhabditis elegans as a model organism, as well as Traf-2 and NCK-Interacting Kinase (TNIK), its role in the nervous system and (specifically synapse formation). I will also introduce the mechanistic Target of Rapamycin (mTOR) pathway and its roles in the developing nervous system. Lastly, I will outline my objectives for this thesis.  1.1 Caenorhabditis elegans  Since the pioneering work of Sydney Brenner in the 1960s and 70s, the roundworm Caenorhabditis elegans has become a powerful model organism in fields of biology. Culturing and propagating worms in a lab environment is relatively simple and efficient. Strains are grown on agar plates streaked with E. coli, and at room temperature a worm will hatch and reach adulthood in three days (Brenner 1974). Worms are transparent and, taken together with their low cost, high fecundity, short life span, and freezable nature, are truly an optimal model system for research in the life sciences. The invertebrate nature of C. elegans obviates the ethical constraints of higher order organisms and thus allows for large-scale screening.  Observation and experimentation on single cells are a challenge in most systems, including cell culture system, in part because the extracellular cues (including that of neighbouring cells) present in vivo are absent ex vivo. One powerful advantage of C. elegans is its invariant cell lineage; all 959 somatic cells have a known cell fate, which can be traced from the single-cell stage (Sulston et al. 1983). Thus, individual cells with unique properties can be observed, and subtle phenotypes may be elucidated. One remarkable example of complexity within a single cell is the thermosensory neuron AFD. Worms cultured at a certain temperature prefer that temperature in a choice assay. However, the thermosensory threshold of the AFD   2  neuron corresponds with the holding temperature (15 minutes leading up to the assay) and not the cultured temperature (Hawk et al. 2018). In fact, preference for cultured temperature is determined by pkc-1-mediated connectivity between AFD and AIY. This work shows that AFD functions both in sensory adaptation and presynaptic plasticity, and illustrates the remarkable capacity, complexity, and function of individual cells (which often function in even more complex networks). 1.1.1 Nervous system of C. elegans  With only 302 neurons, the nervous system of the worm is simple but generates complex behaviours. The relatively small number and invariant morphology and function of neurons in the worm allows researchers to study the nervous system with great resolution, both in terms of microscopy and function. The motor circuit of the worm consists of only 113 neurons in 8 functionally distinct classes (Pérez-Escudero and De Polavieja 2007).  1.1.2 Genetics of C. elegans  C. elegans is one of the most genetically tractable biological systems used today. Decades of collaboration and cooperation from a dedicated community of biologists have resulted in a powerful set of resources, methods, and tools. The Caenorhabditis Genetics Center (CGC) collects strains from labs across the globe and makes them available to all other labs at low cost. In 1998, the C. elegans Sequencing Consortium published the first fully sequenced genome of any organism. Recently, the C. elegans reference genome was re-sequenced and published (Yoshimura et al. 2019).    3  1.2 Traf-2 and NCK-Interacting Kinase/mig-15 in the nervous system 1.2.1 Structure of TNIK In 1999, researchers from the pharmaceutical company Rigel, Inc. identified a novel germinal center kinase (GCK) family kinase which interacts with both Traf2 (TNF Receptor Associated Factor 2), and NCK (Non-Catalytic region of tyrosine Kinase adaptor protein) in human cells, and thus named it Traf2- and NCK-Interacting Kinase (Fu et al. 1999). The authors found that TNIK is a Ste20p kinase with shared homology with the kinase domain of NIK, and that overexpression of TNIK in kidney cell culture resulted in activation of the Jun-C Kinase (JNK) pathway. Interestingly, the role of TNIK as a kinase was unclear, because kinase-dead mutant isoforms of TNIK also activated the JNK pathway. Kinase-dead isoforms of kinases can retain certain functions, as in the case of PAT-4. The worm homologue of integrin-linked kinase (ILK), pat-4, functions as an adaptor protein in muscle assembly, and transgenic expression of kinase-dead pat-4 rescued the null mutant (Mackinnon et al. 2002). While this does not rule out important kinase-related functions of TNIK, it suggests that TNIK may function in multiple pathways, and that its kinase activity may be dispensable for some of its functions. Homologs of TNIK are present in most organisms, including mice, flies, and worms, and humans (Dan, Watanabe, and Kusumi 2001).  TNIK contains a kinase domain near the N-terminal and a citron homology (CNH) domain at the C-terminal (Taira et al. 2004). The functional roles of these domains will be explored in the following sections.    4  1.2.2 Clinical relevance of TNIK 1.2.2.1 Neurological disorders associated with TNIK It is worthwhile to note that TNIK has been implicated in several clinical conditions. Patients with schizophrenia have higher levels of TNIK mRNA in the dorsolateral prefrontal cortex and genetic studies have found that TNIK binds to DISC1, a gene associated with risk of schizophrenia and is involved in brain development (Potkin et al. 2010; Glatt et al. 2005). Furthermore, nonsense mutations in TNIK have been identified as a possible contributor to intellectual disability, although the molecular underpinnings of TNIK activity have not been fully explored (Anazi et al. 2016).  1.2.2.2 TNIK and cancer Furthermore, TNIK has been implicated in cancer growth. Specifically, TNIK’s enzyme activity appears to be essential for the maintenance of colorectal cancer growth through its interaction with the TCF4 and β-catenin complex, which has been implicated in colorectal carcinogenesis (Mahmoudi et al. 2009; Shitashige et al. 2010). TNIK inhibitors have even been proposed as a potential pharmacological treatment for colorectal cancer (Mahmoudi et al. 2009). Further research into the molecular targets and mechanisms of TNIK function will be crucial for our understanding of the disorders with which it is associated. 1.2.3 TNIK and Wnt signaling Wnt signaling is involved in several important developmental processes, including body-axis formation, stem cell maintenance, and cytoskeleton regulation (Reya and Clevers 2005). One mechanism underlying the activation of Wnt target genes is the transcriptional co-activator TCF4/β-catenin (W. Liu et al. 2000). In the intestinal crypt of mice, TNIK interacts with TCF4 to regulate Wnt signaling (Mahmoudi et al. 2009). β-catenin and TCF4 both interact with TNIK,   5  and TNIK phosphorylates TCF4 resulting in transcriptional activation of Wnt target genes (Mahmoudi et al. 2009; Shitashige et al. 2010).  1.2.4 Axon guidance In the nervous system, neurons and glia generate functional circuits by transmitting electrical and chemical signals, forming the cellular basis of the nervous system. During development, neurons project cellular processes called axons. Axon guidance describes the phenomenon of a developing axon extends to its target as a result of multiple cues, both intrinsic and extrinsic (Serafini et al. 1994; Chisholm et al. 2016; Mohamed et al. 2012; Limerick et al. 2018). Molecular guidance cues such as netrins, ephrins, semaphorins, cell adhesion molecules (CAMs), all function in concert to coordinate the construction of connectivity of the nervous system.  Due to its simple and stereotyped nervous system and accessible genetics, research in C. elegans has been effective at discovering the identity and role of genes involved in axon guidance. Using C. elegans as a model organism has revealed that TNIK functions in the nervous system to regulate the actin cytoskeleton and axon guidance. In the absence of β1 integrin, loss of the C. elegans TNIK homolog mig-15 results in more severe axon guidance defects due to impaired interaction between the actin cytoskeleton and the extracellular matrix (ECM) (Poinat et al. 2002). mig-15 acts in parallel with the actin regulator unc-34 to regulate axon pathfinding in C. elegans (Shakir, Gill, and Lundquist 2006). mig-15 is also required for Q neuroblast cell migration, functioning independent of the Wnt signal controlling posterior cell migration (Chapman, Li, and Lundquist 2008). Subsequent research found mig-15 mutants have quantifiable defects in every motor neuron axonal commissure examined (Teuliere et al. 2011). In mammalian neuron cell culture the JNK pathway requires TNIK (among other Ste20 family   6  kinases) for neurodegeneration, and a knock-down of these kinases protects neurons from neurodegeneration (Larhammar et al. 2017).  1.2.5 The role of TNIK in synapse formation and specificity Neurons form connective junctions between their axons and their cellular targets, and these junctions are known as synapses. While these connections are abundant in the nervous system (forming n the order of trillions in higher order mammals), neural circuits require precise synapse formation to give rise to complex brain function. The currently understood mechanisms of the formation and specialization of synapses in C. elegans are reviewed by Hendi et al. (2019). In addition to regulating axon guidance, TNIK also functions at the synapse. In hippocampal neuron cell culture, TNIK is enriched at the post-synaptic density (PSD), and RNAi of TNIK results in reduced synapse number and AMPA receptor (one of two glutamate receptors, crucial for excitatory synapses) function (Hussain et al. 2011). TNIK interacts with Disrupted in Schizophrenia 1 (DISC1), a genetic risk factor for schizophrenia, at the PSD to regulate post-synaptic protein levels and synaptic strength (Q. Wang et al. 2011). Specifically, DISC1 binds TNIK which inhibits its kinase activity; a knockdown of DISC1 resulted in increased TNIK activity and abundance and increased GluR1 and GluR2/3 expression (Q. Wang et al. 2011). In C. elegans, the Pam/Highwire/RPM-1 (PMR) gene rpm-1 requires mig-15 to impair synapse formation in the mechanosensory neuron PLM (Crawley et al. 2017). Recently, our lab found that MIG-15 negatively regulates synapse formation in all C. elegans neurons. Loss of mig-15 increased synapse number in the cholinergic motor neuron DA9 and overexpression of mig-15 reduces synapse number and overall synaptic fluorescence in the DNC (Chen et al. 2018). In addition to regulating synapse number, mig-15 also functions in the Plexin-  7  Rap2 pathway controlling tiled presynaptic innervation of the body wall muscle. While this will be explored in greater detail in chapters 1.3 and 4, it is important to mention that the genes upstream of mig-15 in the synaptic tiling pathway (plx-1 and rap-2) do not affect synapse number, suggesting mig-15 functions in multiple pathways to regulate patterning and formation of synapses (Chen et al. 2018).   1.3 Patterned synapse formation In both vertebrates and invertebrates, complex functions of the nervous system rely on patterning of synaptic connections in neural circuits. In the vertebrate retina, for example, cells form strict positional boundaries and synaptic partners necessary for projecting complex visual stimuli to higher order brain regions (Sanes and Zipursky 2010). In this system, synaptic specificity allows cells to connect with partners of specific subtypes. In the visual system of the fruit fly Drosophila melanogaster, a similar map of finely organized neuronal and synaptic projections is evident in the retina, lamina, and six medullar layers (Sanes and Zipursky 2010). In the developing fly, synapse formation is characterized by patterned stimulus-independent firing in specific cell types (Akin et al. 2019).  1.3.1 Synaptic tiling in C. elegans Fine motor control in living organisms requires precise synaptic connections between motor neurons and their muscular targets. Using the stereotyped development and relatively simple neuronal structure of the nematode C elegans, it is possible to closely observe patterned localization of synapses in neurons. Synapse patterning in C. elegans describes a phenomenon where, for some groups of motor neurons, each neuron of a given class forms synapses within specific segments of their axon that tile with each other to create a non-overlapping set of   8  synaptic domains (White et al. 1976). For example, nine cholinergic motor neurons of the DA-class form tiled synaptic domains along the dorsal nerve cord (DNC).  The most posterior of the DA-class motor neurons, DA9, forms neuromuscular junctions with the body wall muscle in a restricted segment of its axon (its synaptic domain) anterior to the axonal commissure (a ventral-to-dorsal projection of the DA9 axon at its most posterior point) and terminates near the posterior end of the DA8 synaptic domain (Figure 1A). The DA8 neuron soma, axon, and dendrite have very similar morphologies to that of DA9, with the notable exception of its synaptic domain, which forms anterior to the synaptic domain of DA9. The DA8 and DA9 synaptic domains exhibit minimal overlap. This phenotype is referred to as synaptic tiling and requires inter-axonal interaction between adjacent motor neurons. Mutant worms with misguided DA8 or DA9 axons do not exhibit synaptic tiling, demonstrating that axon-axon interaction is necessary for tiled synapse formation (Mizumoto and Shen 2013). This organized and segmented synaptic innervation of the body wall muscle hints at a possible mechanism for the sinusoidal locomotion characteristic of C. elegans, however, a spatiotemporal neural circuit of DA neuron signalling has not been shown. Semaphorin (SMP-1) and its receptor, plexin (PLX-1), along with the signaling pathways this interaction activates, regulate synaptic tiling between DA8 and DA9 motor neurons by inhibiting synapse formation at putative asynaptic axonal domains (Mizumoto and Shen 2013).  Patterning of neural circuits is also important is higher order animals. For instance, dendritic patterning in vertebrates is controlled by a number of intracellular signals and extrinsic cues, including semaphorins (Ledda and Paratcha 2017). Many of these genes are disrupted in neurodegenerative and psychiatric disorders, including Alzheimer’s disease, Parkinson’s disease, Huntington’s disease, intellectual disability, and autism spectrum disorder (ASD) (Ledda and   9  Paratcha 2017). Understanding the signaling mechanisms underlying synaptic tiling will offer a greater understanding of the genetic basis of patterned synaptic organization in other models and hopefully contribute to our understanding of the molecular pathways underlying brain disorders such as autism, schizophrenia, and intellectual disability. 1.3.1.1 The role of plx-1, rap-2, and mig-15 in synaptic tiling The transmembrane protein Semaphorin receptor Plexin (plx-1) is required for tiled presynaptic innervation in DA8 and DA9 motor neurons (Mizumoto and Shen 2013). Sema-plexin signalling is important for inhibitory cues at the axon growth cone during neuron development (Winberg et al. 1998; Takahashi et al. 1999). Loss of plx-1 resulted in anterior expansion of the DA9 synaptic domain and posterior expansion of the DA8 domain. PLX-1 exhibits GTPase-activating protein (GAP) activity specific to Rap GTPases (RapGAP) (Y. Wang et al. 2012).  Further research implicated the RapGAP domain of PLX-1 and its downstream partner RAP-2, a small GTPase involved in neurite outgrowth and spine formation in mammalian cells (Kawabe et al. 2010; Hussain et al. 2011), in the synaptic tiling pathway. Mutations in rap-2 (null, constitutively GDP-, and GTP-bound) resulted in synaptic tiling defects similar to that of plx-1, suggesting cycling of rap-2 GTPase activity is necessary for synaptic tiling (Chen et al. 2018). TNIK is a known downstream effector of Rap2 (Taira et al. 2004), and its C. elegans ortholog, mig-15, functions downstream of RAP-2 in the synaptic tiling pathway. Indeed, worms lacking mig-15 kinase activity (allele rh148) exhibited a severe synaptic tiling defect (Chen et al. 2018). Interestingly, mig-15 animals exhibit a more severe synaptic tiling defect than either rap-2 or plx-1 mutants. Double mutants of plx-1;mig-15 and rap-2;mig-15 show the same exaggerated synaptic tiling defect. One possible reason for this is that mig-15   10  mutants have an increase in synapse number, driving a larger expansion of the synaptic domains (Chen et al. 2018). This idea will be explored in greater detail in chapter 4.4. 1.4 The TOR pathway Discovered in 1964 from a sample of soil on the island of Rapa Nui (or Easter Island), Rapamycin (or sirolimus) is a macrolide compound produced by the bacterium Streptomyces hygroscopicus and was initially studied and developed for its antifungal and immunosuppressive properties (Chung et al. 1992). However, independent biochemical analyses of the molecular targets of Rapamycin revealed the mammalian (or mechanistic) Target of Rapamycin (mTOR, generalized as TOR) (Sabers et al. 1995; Brown et al. 1994). TOR is a serine/threonine protein kinase which has since been found to function at the centre of a large and highly conserved signalling network controlling cell growth and metabolism (Saxton and Sabatini 2017). TOR is in the PI3K-related kinase (PIKK) family and forms two distinct protein complexes, TOR complex 1 (TORC1) and TOR complex 2 (TORC2).   Diagram illustrating signaling network of TOR pathways.     11  TORC2 is primarily involved in cell proliferation and survival and is characterized by the binding of its adaptor protein Rictor (rapamycin insensitive companion of TOR). PKCα, a protein kinase which regulates the actin cytoskeleton, was the first identified substrate of TORC2 (Jacinto et al. 2004; Sarbassov 2005). TORC2 also regulates cytoskeletal remodeling and cell migration through the phosphorylation of other PKC family members, namely PKCδ, PKCγ, and PKCε (Gan et al. 2012; X. Li and Gao 2014; Thomanetz et al. 2013). Furthermore, TORC2 activates the key insulin/PI3K effector Akt. Once phosphorylated, Akt promotes cell survival, proliferation, and growth through FoxO1/3a, TSC2, and GSK3β (Guertin et al. 2009; Jacinto et al. 2006). TORC1 comprises three elements: TOR, Raptor (regulatory protein associated with TOR), and mLST8 (Tokunaga et al. 2004; D. H. Kim et al. 2003). TORC1 regulates processes critical for cell growth and proliferation: the synthesis of proteins, lipids, and nucleotides and the suppression of catabolic pathways (e.g. autophagy) (Saxton and Sabatini 2017). TORC1 phosphorylates S6K1 and elF4E Binding Protein (4EBP) to promote protein synthesis. Subsequently, S6K1 promotes the initiation of mRNA translation by phosphorylating a number of substrates, including elF4E (Saxton and Sabatini 2017). TORC1 also promotes synthesis of de novo lipids via the sterol responsive element binding protein (SREBP), which responds to low sterol levels (Saxton and Sabatini 2017). Additionally, TORC1 is a key regulator of protein turnover, more specifically autophagy. Autophagy is the catabolic process by which components of the cell are degraded and recycled. An early step in the autophagic process is the activation of the UNC-51-like-kinase ULK1 by AMPK, which drives autophagosome formation by forming a complex of ULK1, ATG13, FIP2000, and ATG101. TORC1 negatively regulates autophagy by   12  phosphorylating ULK1, preventing its activation by AMPK, when nutrient availability is normal (J. Kim et al. 2011; Dunlop and Tee 2014).  While TORC1 controls several important downstream cellular processes, the upstream mechanisms that regulate TORC1 activation provide critical insights into the role TOR plays in environmental sensing. In the past few years, ground-breaking work has revealed a well-conserved amino acid sensing branch of the TOR pathway. TORC1 activation occurs at the lysosomal surface: Four RagGTPases (RagA and RagB, and their respective homologues RagC and RagD), the Ragulator protein, and vacuolar adenosine triphosphatase (V-ATPase) form the Ragulator complex. This complex functions via GEF activity toward RagA and RagB GTPases, and receives inhibitory input from the GATOR  (GTPase-activating protein (GAP) activity toward Rags) complex (Bar-peled et al. 2013). The GATOR complex is comprised of two subcomplexes, GATOR1 (containing DEPDC5, Nprl2, and Nprl3), which is inhibited by GATOR2 (Mios, WDR24, WDR59, Seh1L, Sec13) (Bar-peled et al. 2013). The GATOR complex receives input from three known nutrient sensors, which ultimately serve to regulate TORC1 activation when the cell is replete with nutrients: Sestrin1/2, a leucine sensor (Chantranupong et al. 2014) and Castor1/2, an arginine sensor (Saxton et al. 2016) both inhibit GATOR2 in the absence of their respective amino acids, while SAMTOR (S-adenosylmethionine sensor upstream of TORC1) signals methionine starvation (sensed through the binding of SAM) by interacting with GATOR1 to inhibit TORC1 activity (Gu et al. 2017). To this end, TORC1 integrates environmental cues surrounding nutrient availability to regulate growth and development. Recent work from the Sabatini research group used cryo-electron microscopy to resolve the structure of the supercomplex of Raptor with Rag-Ragulator to 3.2 Å (Rogala et al. 2019).   13  The authors found that three α-helices of Raptor binds with the switch-I face of GTP-loaded RagA. Furthermore, they report a ‘claw’-like structure of Raptor between the two GTPase domains of RagA and RagC, which serves as a nucleotide detector to ensure that RagC is GDP-bound (a necessary condition for TORC1 activation). Rogala et al (2019) conclude that the Rag-Ragulator complex binds on the top and sides of mTORC1 and functions as a clamp, pushing mTORC1 down onto the lysosome.  1.4.1 TOR in C. elegans The worm system has proved powerful in uncovering the physiological functions of the TOR pathway (Blackwell et al. 2019). Many of the proteins in the TOR pathway are functionally conserved in C. elegans. Three core proteins of the TOR pathway are LET-363/TOR, DAF-15/Raptor, and RICT-1/Rictor. In the aging field, C. elegans has implicated TOR with lifespan; worms with decreased TOR activity have longer lifespans (Robida-Stubbs et al. 2012). DAF-15 is the worm homologue of mammalian Raptor and, interestingly, daf-15(m81) worms have arrested development and form specialized L3 dauers (Hara et al. 2002; Albert and Riddle 1988). Given that starved worms can enter a dauer state, where growth and reproduction ceases, this has raised the question of the role of TORC1-mediated nutrient sensing in dauer formation. Despite the significant work in aging, development, and metabolism, there is relatively little research on the role of TOR in the worm nervous system. 1.4.2 The role of TOR in the nervous system 1.4.2.1 TOR in the developing brain While much has been discovered about the cellular functions controlled by TOR, the full range of its biological roles has yet to be understood. One emerging area of research is the role of TOR in nervous system development.   14  In the developing brain, neural progenitor cells undergo proliferation, giving rise to neuronal and glial cells, eventually forming the adult nervous system. TOR is critical for neural progenitor cell proliferation; loss of mTOR causes hypertrophy (or overexertion) of these cells, while overactivation of mTOR results in depletion of stem cell niches and causes significant neurodevelopmental defects (described below). Controlled overactivation of mTORC1 during embryonic stages induces cortical atrophy due to apoptosis of progenitors, while overactivation during adulthood (in post-mitotic neurons) caused cortical hypertrophy, inducing seizures (Kassai et al. 2014).  Loss of tuberous sclerosis proteins 1 and 2 (TSC1/2) or phosphatase and tensin homolog (PTEN) both increases mTORC1 signalling and results in defects in neural development and morphology. In mice, a deletion in Pten caused macrocephaly and neuronal hypertrophy as well as ectopic axon and dendrite branches and increased synapses (Kwon et al. 2006). Loss of Tsc1 caused an increase in neural cell body size and larger spine heads corresponding with increased AMPAR-mediated synaptic currents (Tavazoie et al. 2005). Tsc1 mutant mice also have enlarged or dysplastic cortical and hippocampal neurons which grow ectopically throughout the cortex (Meikle et al. 2007). More recently, work to elucidate the relative contributions of mTORC1 and mTORC2 in neurodevelopment employ Raptor or Rictor specific knockouts, respectively. Loss of mTORC1 induced by a deletion in the Raptor gene in mice caused microcephaly due to reduced cell number and size (Cloetta et al. 2013). Rictor knockout mice, on the other hand, also have smaller brain size, smaller soma size, and shorter dendritic arbours (Thomanetz et al. 2013). That both loss of mTORC1 and mTORC2 result in similar brain defects in mice may indicate a balance between TORC1 and TORC2 activation is necessary for correct proliferation of developing neurons.   15   1.4.2.2 TOR at the synapse As discussed in previous sections, the nervous system functions through neural circuits comprised of millions of synaptic connections. Understanding the mechanisms by which synapses are formed, maintained, eliminated, and prohibited is a major goal of basic neurobiology research and provides powerful insights into synaptic pathologies in brain disorders. Indeed, TOR signaling has been implicated in aspects of synapse formation. As mentioned above, Pten and Tsc1 loss resulted in increased synapses and larger spine heads respectively (Meikle et al. 2007; Tavazoie et al. 2005). One possible mechanism for the maintenance of synaptic connectivity is logcal protein translation, including TORC1-regulated mRNA translation (Jung et al. 2014).  Activity-dependent changes to neuronal circuitry may also engage TOR signaling. The N-methyl-D-aspartate (NMDA) antagonist ketamine has recently been used as a rapid therapeutic agent for patients with treatment-resistant major depressive disorder (MDD). Early work found that ketamine rapidly activates mTOR signaling, resulting in increased synaptic protein abundance, synapse number and function, and new spines in the rat prefrontal cortex. This corresponded with rapid behavioural improvements in depressed rats, and was abolished when treated with rapamycin, an mTORC1 inhibitor (N. Li et al. 2010). Subsequent research found that ketamine-induced antidepressant effects are the result of AMPA receptor-mediated activation of mTOR and brain derived neurotrophic factor (BDNF) (Zhou et al. 2014). Recently, a synthetic leucine analogue which selectively binds sestrin, NV-5138, was shown to function as a selective mTORC1 activator. mTORC1 activation with NV-5138 results in ketamine-like   16  antidepressant effects in rats, rescues synapse loss associated with depression, and increases dendritic spine number in control animals (Kato, Hahm, and Duman 2019). The role of mTOR at the functional level of the synapse is the focus of more recent work. In mice, loss of mTOR in ventral tegmental area (VTA)-specific neurons decreases dopamine release and reuptake in the shell of the nucleus accumbens. This VTA-specific deletion of mTOR did not affect neuron morphology, but increased firing from and inhibition by GABAergic neurons in the VTA, resulting in decreased cocaine preference and cocaine-induced potentiation of excitatory over inhibitory dopaminergic neural activity (X. Liu et al. 2018). Synaptic plasticity driven by presynaptic activity, or Hebbian plasticity, is an important principle that drives the refinement and specificity of neural circuits. Mice reared in enriched environments have retinoic-acid receptor (RARα)-dependent increases in Hebbian synaptic plasticity, and this results in specific learning deficits. In RARα deletion mice, environmental enrichment causes an increase in mTOR signaling, which increased AMPAR levels, and rapamycin treatment rescues AMPAR levels and Hebbian plasticity defects (Hsu et al. 2019). 1.4.2.3 Autophagy at the synapse Protein degradation is a critical catabolic process and macroautophagy (hereafter autophagy) is a major component of this process. Autophagy is necessary for intracellular homeostasis as it degrades cellular material that may be harmful (e.g. proteins prone to aggregation) (Menzies et al. 2017). Autophagy-related (ATG) proteins form a double-membraned autophagosome which engulfs its targets, traffics to the lysosome, and undergoes autophagosome-lysosome fusion, resulting in degradation of the autophagic cargo (Menzies et al. 2017). One downstream critical role of TORC1 is its negative regulation of autophagy (Saxton and Sabatini 2017). TORC1 inhibits ULK1, an early gene in the autophagy cascade, directly via   17  phosphorylation and indirectly via phosphorylation of autophagy/Beclin-1 regulator 1 (AMBRA1) (Dunlop and Tee 2014; Menzies et al. 2017). In this way, TORC1 functions as a negative regulator of autophagy. A growing body of evidence implicates autophagy in the formation and function of neurons and their synapses (D. N. Shen et al. 2015; Birdsall and Waites 2019; Lieberman et al. 2018). In autism spectrum disorder (ASD) individuals and an ASD mouse model (Tsc+/-), elevated mTORC1 activity correlated with reduced autophagy and increased spine density, suggesting that synaptic pruning defects in ASD may be the result of TORC1-mediated autophagy (Tang et al. 2014). In a Fragile X syndrome mouse model (Fmr1-KO), defects in dendritic spine structure and synaptic plasticity are rescued by increased autophagy via Raptor shRNA (Yan et al. 2018). Both studies suggest that autophagy functions as a positive regulator of synaptic elimination or pruning, or put another way, a negative regulator of spine formation. However, other evidence points to autophagy as necessary for proper synapse formation. Shen and Ganetzky found that autophagy promotes synapse formation in the invertebrate Drosophila melanogaster (2009). Further, they found that Rapamycin-induced increase in autophagy significantly increased the number of boutons at the neuromuscular junctions (NMJs) in wildtype but not atg18 flies, and that these changes to the NMJ were independent of translation defects (as rapamycin inhibits TORC1 and thus decreases protein translation) (W. Shen and Ganetzky 2009). In C. elegans, autophagy regulates vesicle clustering at the AIY interneurons by forming autophagosomes at presynaptic sites during development (Stavoe et al. 2016). Unlike results found in mammalian models, these results suggest autophagy is required for synapse formation. One possible explanation for this discrepancy is alternate pathways: in mouse studies, autophagy is manipulated in disease model backgrounds, where loss of TSC1 or Fmr1 may serve as confounds and the role of autophagy in wildtype animals cannot   18  be elucidated. Another possible confounding variable is timing: knockdowns of autophagy in worms and flies affect the development of the nervous system, whereas post-embryonic treatment of rapamycin in mice may not elucidate the developmental role of autophagy in synapse formation. Taken together, these results implicate autophagy as possibly playing multiple roles at the synapse and should be further studied.  1.5 Thesis Objectives 1.5.1 Identification of a genetic mechanism for the negative regulation of synapse number by TNIK/mig-15. We aimed to gain a greater understanding of the mechanisms by which the kinase TNIK/mig-15 functions to inhibit the formation of synapses. Specifically, we wanted to identify genes which are necessary for mig-15 to reduce synapse formation in C. elegans. To this end, we did a forward genetic screen to identify potential genes of interest.  1.5.2 Characterization of the GATOR complex in relation to mig-15-mediated synapse formation We aimed to elucidate the role of the amino acid sensing branch of the TOR pathway, which turned up on our genetic screen. The TOR pathway is highly conserved, ubiquitously expressed, and implicated in several disease, and as such its role in nervous system development is of great interest.   19  Chapter 2: Materials & Methods 2.1 Worm Culture and Strains C. elegans strains were cultivated ~22℃ on NGM plates seeded with OP50 Escherichia coli. N2 Bristol worms were used as the reference wildtype strain. Strains were grown and maintained as described in Brenner (1974).  Table 1. List of strains used in this work strain gene (allele) or genotype VC2010 N2  NJ490 mig-15 (rh148) X ST36 plx-1 (nc36) IV ZB2774 Y32H12A.8 (or wdr-24) (tm419) III ZB1748 Y32H12A.8 (or wdr-24) (tm422) III UJ699 mizIs33 Prab-3::mig-15, Podr-1::GFP UJ735 mizIs34 Prab-3::mig-15, Podr-1::GFP UJ736 mizIs35 Prab-3::mig-15, Podr-1::GFP TV18675 wyis685 Pmig-13::tdTomato::rab-3, Pmig-13::2xnovoGFP::fife UJ98 mizis1 Pitr-1::mCherry::rab-3, Pitr-1::zf1-GFPnovo2::CAAX, Pvha-6::zif-1 with Pord-1::RFP UJ124 mizis3 Punc-4::zf1-GFPnovo2::rab-3, Pmig-13::zif-1, Pmig-13::mCherry::rab-3, Podr-1::RFP TV14517 wyIs446 Punc-4::2xGFP::rab-3, Pmig-13::2xmCherry::rab-3, Podr-1::RFD  VC40837 npp-20 (gk840106) IV VC2517 npp-18 (ok3278) III  grk-2  VC20647 R08D7.5 (gk368845) III TM5932 nprl-2 tm5932 UJ814 mizIs33; miz27  UJ815 mizIs33; miz28  UJ816 mizIs33; miz29  UJ817 mizIs33; miz30  UJ820 mizIs33; miz31  UJ821 mizIs33; miz32  UJ822 mizIs33; miz33  UJ823 mizIs33; miz34  UJ950 wdr-24(tm422) III; mizIs34   20  UJ951 wdr-24(tm422) III; mizIs34 UJ1052 wdr-24(tm422) III; mizIs34 UJ1146 unc-51(e1189) V; wyIs685 UJ1188 daf-15(miz64Ins)   UJ1192 daf-15(ok1412)/tmC5[tmIs1220]; wyIs685 UJ1216 mizIs33; wyIs685  UJ1217 mizIs33; wyIs685  UJ1228 Y32H12A.8(tm419) mizIs33 III; wyIs685    2.2 Nomenclature of C. elegans  Genetic nomenclature in C. elegans is standardized. All genes are named with either three or four letters followed by a hyphenated number. Genes and mutants are written as lower-case and italicized, with the option denoting the chromosome number in Roman numerals. For example, ‘daf-16 I’ refers to the gene which encodes for the (capitalized) protein DAF-16, located on chromosome I. Worms described as daf-16 carry a loss of function mutation in the daf-16 gene. The alleles of mutants may be parenthetically denoted next to the gene. For example, the daf-16 mutant allele mu86 would be written as daf-16(mu86). While gene names sometimes refer to the phenotype of their mutants, phenotypes are described with a capital initial letter followed by lower-case letters (e.g. unc-119 would be described as uncoordinated or Unc).  Transgenic strains are described using the two or three letter code of the origin lab (e.g. miz = Mizumoto Lab, wy = Kang Shen lab), followed by a designation Is (if integrated) or Ex (if expressed as an array). For example, ‘mizIs1’ describes the first integrated transgene generated in the Mizumoto Lab.  2.3 DNA Extraction and Polymerase Chain Reaction Genotypes of all mutant strains were confirmed via electrophoresis of amplified DNA fragments from polymerase chain reaction (PCR), except where a known phenotype was used for selection. DNA extraction was conducted by transferring at least 20 worms into PCR tubes with   21  20 µL of lysis buffer (50 mM KCl, 10 mM Tris pH 8.3, 2.5 mM MgCl2, 0.45% NP-40, 0.45% Tween-20, 20mg/mL proteinase K). These worms were then lysed at 65℃ for 60 minutes, followed by 95℃ for 15 minutes for Proteinase K inactivation. The lysed DNA was subsequently used as template DNA for PCR. PCR: DreamTaq polymerase and buffer were used for PCR. For each sample, 1µL of template DNA was added to a 9 µL aliquot of master mix (H2O: 5.95 µL, DreamTaq buffer: 1 µL, dNTP: 1 µL, forward and reverse primers: 0.5 µL at 0.1 mM, 50% glycerol: 0.5 µL, DreamTaq polymerase: 0.05 µL). PCR protocol was conducted according to instructions on Thermo Fisher website. Annealing temperatures were optimized for each primersets, typically between 55℃ and 63℃.     22  Table 2. Primers used for genotyping and cloning genotype forward reverse (outside deletion) reverse (inside deletion) OR enzyme grk-2(gk268) TAGTGCCAGCGTTTCTCCTT CATAACTCGATTTGGACGGG  R08D7.5 GCCAAATTTGCTCAAGCTTTCAC TTCCACAGTTCGTCAACGTCCATGG EcoRI Y32H12A.8 (S1794F/sup3) CTTTACGAGAACGATCCTGAACTGC CACATTGAGAGCTCAACAGCTG PstI Y32H12A.8(tm419) GGAGCTCTTCGTACTCTTCGTC GAAGGCAATCCGCCAAATTCC CAGAATCTGAAGAGGGCTCTG Y32H12A.8(tm422) GGAGTCTCTCGTCTTGCAATCGG AGAAAAAGGCCATTGCGGCAACC GATTGTTGCTCTATCACGCCGAG npp-18(ok3278) GCATTGCGAAGGACTGTGC GTTCTTCGGATCCATTGGGA TTAAAGGCGCAGGCACTTTC raga-1(ok386) TTTGCCAACTACATCGCCAG AGCAGCCACCTCCATCTG CGTCGCTCTCTCGAATAGAA let-363(ok3018) CGATCAGAAACGAGCCGGTA AGGACAAGCCATTCAACACC GTAATGCATCAATTCCGGCG daf-15(ok1412) GGCAGTAGATCCATCGTGTC CCTGACGAGATGTATTGGTT GTTTGATTTCAGGATCGCCG npp-20(gk840106) CATCGCCAGCAACTGAAAGG ATGGCCGCTGAAACTTGTC HpaII nprl-2(tm5955) GGACCTTTTGCAACAGGTGA TCCAACGGGAAACATGAGAAC TCTGCCAACAGAATTCCGAGA epg-5(tm3425) GCGAAGAGCAACTCTACCCGGTGT CACCGGAACCTGAAGATCAGTAAGTGGTAAA GGAGACAACAATGATGGTACAGGTAGAG Y32H12A.8/wdr-24 conserved cDNA for pSM TTCAGCGGCGCGCCATGGAAGAACCGGAT TACCATGGTACCTATTCTGTCCACATGTA  daf-15::AID 5' homology arm AACGACGGCCAGTGAATTCCCGCGGGATTCCCGAATTCTTGCAAGTGCTG GGGCTCCGGCTCCGGCTCCGGCTCCCAGTGATTGCGAGGGAGGGGAACTG  daf-15::AID 3' homology arm GAAGTTATAGTTGCAGGACCACTGCATCTCCTCTGTAAATTATTTGAATT ATGATTACGCCAAGCTTGCGGCCGCAATCATTTGAAGAATCATCTTATAG  daf15 aid integrant WT fwd 1kb TCCTTGGGAATACGGAGCTCATTCTCATATG   daf15 aid integrant WT rev 1kb AATTGAATCGCCAGCTCCACCTGCTCCACA     23   2.4 Confocal microscopy Images used for quantification of synapse number and synaptic fluorescence were acquired using confocal microscopy. The Carl Zeiss LMS800 confocal microscope (Carl Zeiss, Germany) was used to acquire images, with oil immersion lenses of either 63X or 40X magnification (Carl Zeiss, Germany). Immobilization of animals was performed on 2% agarose pads using a mixture of 0.225 M 2,3-butanedione monoxime (BDM) and 25 mM levamisole (Sigma-Aldrich, USA). Laser wavelengths of 488 nm and 568 nm was used to excite GFP and mCherry signals respectively. The software used to capture and process images was Zen Blue Edition (v2.1) running on the Windows 7 operating system, with an Intel Core™ i5-4670 CPU @ 3.40 GHz, with 32 GB of memory (RAM).  2.5 Plasmid construction All expression clones were derived from the pSM vector. Constructs were used via standard microinjection techniques (Mello et al. 1991).  Plasmid used: pFM3 (Prab-3::wdr-24conserved::pSM)  2.6 Multiple Sequence Alignment Orthologs of Y32H12A.8 were identified from a BLAST search (NIH), and an isoform of a conserved ortholog was selected from Caenorhabditis elegans, Homo sapiens, Mus musculus, Danio rerio, and Drosophila melanogaster and the FASTA sequences of their protein transcripts were compiled in a text file. Next, I conducted an alignment of these protein sequences using Constraint-based Multiple Alignment Tool (COBALT) and visualized in the COBALT tool on the NIH website (ncbi.nlm.nih.gov/tools/cobalt/cobalt.cgi). Then, I conducted a multiple   24  alignment of the conserved domain of the gene (the first 838 a.a.) with the protein sequences of the orthologs using T-Coffee. Lastly, I used ESPript to render the alignment.   2.7 Strain integration To achieve stable expression of transgenes and to avoid mosaic expression of extrachromosomal arrays, mizIs33, mizIs34, and mizIs35 were integrated using chemical mutagenesis. Briefly, worms were washed and resuspended in Psoralen. After incubation, worms were seeded and screened for fully transgenic populations. Integrants were backcrossed with N2 five times to remove background mutations. 2.8 EMS mutagenesis ~8 plates of plates with L4 stage mizIs33(Prab-3::mig-15) animals were transferred to a 15 mL conical tube to a final volume of 2 mL. 20 µL of ethyl methanesulfonate (EMS) is added and worms are rotated at a final volume of 4 mL for 4 hours. Then, 10 mL of M9 is added, spun down, and aspirated. This is repeated 5 times. Mutagenized mizIs33 animals were seeded and L4 animals were isolated onto plates and their progenies propagated. The Unc phenotype of mizIs33 worms renders the worms unable to engage in ordinary sinusoidal locomotion. I screened the mutagenized progeny for worms exhibiting wildtype or improved locomotion. Candidates were chosen if they were able to engage in more than two locomotor ‘cycles’ in response to tap, and if their progenies exhibit improved locomotion they were considered ‘suppressed’. 2.9 Chromosomal mapping using SNPs Briefly, I crossed sup3 mizIs33 animals (which have overexpressed mig-15 but wildtype locomotion) with a Hawaiian strain CB4856 and isolated several F2s of sup3 mizIs3. When compared to the N2 (Bristol) strain, CB4856 has many SNPs scattered throughout the genome.   25  Several of these SNP sites are differentially digestible by the restriction enzyme DraI. Thus, when mutant animals are crossed with the Hawaiian strain, one can expect ~50% of the SNPs to be Hawaiian in origin and ~50% to be from N2. However, in genomic regions close to the selected-for mutation or transgene, there will be low rates of Hawaiian SNPs due to lower rates of recombination. I crossed mizIs33;sup3 animals with Hawaiian males, and singled ~15 cross-progeny F1s, from which I singled ~60 F2s expressing mizIs33 and exhibiting rescued locomotion (thus selecting for mizIs33;sup3 animals). Once propagated, I genotyped these F2s for DraI-recognizable Hawaiian SNPs on the left, centre, and right arms of each chromosome. For all genomic regions except the region of the transgene and suppressor mutation, ~50% of PCR products will digest like N2 and ~50% will digest like Hawaiian strains. Genomic regions near the genes/transgenes selected for in the cross will have few Hawaiian products and mostly N2 products. 2.10 CRISPR/Cas9 To tag the endogenous copy of daf-15 with the AID degron at the C-terminus, I used CRISPR-Cas9 gene editing technology. Experimental procedures were conducted as described in Au et al. (2019). The repair template plasmid contains an upstream and downstream homology arm (500 bp in length) cloned from C. elegans gDNA. The upstream arm was cloned with a SNP at the PAM site to prevent secondary genomic cuts after homology directed repair. Between the arms are AID::BFP and, flanked by loxP sites, a selection cassette (Pmyo-2::GFP and Prps-27::neoR). This repair template plasmid   26  was injected with Cas9 protein, guide RNA (cagaggagatttacagtgat) selected from the guide RNA selection tool (http://genome.sfu.ca/crispr), and co-injection markers Pmyo-2::RFP and Pmyo-3::RFP. Drug selection was conducted with G418 (geneticin), and worms expressing neoR are resistant to its toxicity. Surviving worms at the  generation (either integrants or array-carriers) were screened for dim, uniformly expressed pharyngeal GFP and for absence of RFP (which would indicate an extrachromosomal array), and these candidate integrants were singled and propagated. ~50 worms were injected resulting in two integrants. To confirm the perfect insertion of AID::BFP and the selection cassette, I used Sanger sequencing the 5’ and 3’ junctions of the insertion and aligned the results to my predicted sequence. I acquired one perfect integrant from this process.     27  Upstream homology arm: gattcccgaattcttgcaagtgctggtggtggagaattagcaatctacgacgttgctcaaaataagctaatggctcgtatcttaccacccggagagcccaattcaatgaggagagataaaaattcaagcacatttggattgttcagctcacaaaggaaactatcacaagttcaaatgagtggaagaatgtcatcgatgtcttcagcaacagttggagaagatcgacaaccaaaagttatttcattgacaatgcatcaaatgaggttagttgattaatatctaacattatgaaaaataatttcatttaattctttcttttttttgaagctcttttttaaagacttgataaaatactaacaatcaacttcaacatttctttctagactgctcactgtcgctggatacgatgacaatacagtttgtgtatatggatccccgaaacccggacttacaggccactacgaaaacacccaaggcgccagttcccctccctcGcaatcactg note: The capitalized and underlined Guanine at the protospacer adjacent motif (PAM) was modified to prevent sgRNA-DNA binding and stabilize the Cas9 nuclease activity. Downstream homology arm: atctcctctgtaaattatttgaatttgtagtttctaaaaccgttacagaatactttaggatgtttctccattgttctacatttagattcttttagacttgaaaaaaaaactacatgattctttcttttcgttttcctacttcatctacagtgccttattaatttcttttttgaaaaacccctaaattctattaccctgtaaatactagcaacctagtatttgtattttgtgttgttcattaatagtatttaacccactttttttgaaaacttcttctaactacctgctcagctatctatttgccaccattttcatgtcttgttttgccaatgttgtttctttatatcctatattttatttttcaccctttgtaggaatagataaagttacaaactttaaaaaataatgtaaacacaatgttaacaaattttattaaaataagaatgattaatgtccaaagaaacttctataagatgattcttcaaatgatt  28  Chapter 3: Screening for genetic mechanism of MIG-15-mediated inhibition of synapse formation  3.1 mig-15 is a negative regulator of synapse formation Previous research from our group found that mig-15 is necessary for patterned synaptic innervation of the DA8/DA9 motor neurons to the body wall muscle (Chen et al. 2018). In addition, mig-15 animals have more synapses in DA9 motor neurons. To test the possibility that mig-15 is a negative regulator of synapse formation, we overexpressed mig-15 under the DA9-specific promotor Pmig-13. These animals showed a significant reduction in synapse number in DA9, suggesting that mig-15 negatively regulates synapse formation (Chen et al. 2018). One limitation of this finding is that the marker strain used to quantify synapse number only tagged the synaptic vesicle protein RAB-3 with GFP. RAB-3 puncta in DA9 can vary significantly in size, thus precluding precise quantification. In order to quantify these results using a more precise reporter, I reproduced these findings using the DA9-specific active zone marker wyIs685, which tags RAB-3 with tdTomato and CLA-1 with GFP (Figure 1B-H). CLA-1/Clarinet1 is a novel active zone protein necessary for synaptic vesicle clustering and release. Tagging these two proteins in the same marker allows for colocalized RAB-3 and CLA-1 puncta to be considered a bona fide synapse (Xuan et al. 2017).    29   Figure 1. mig-15 is a negative regulator of DA9 synapse formation. (A) Schematic diagram of DA9 motor neuron. (B-D) Representative images of presynaptic terminals in DA9 in wildtype, mig-15 loss of function, and mig-15 overexpression worms in wyIs685. Green = CLA-1 active zone protein, magenta = RAB-3 synaptic vesicles. Co-localized CLA-1 and RAB-3 puncta are denoted with a yellow arrow in (B). (E-F) Stacks of DA9 synaptic domains, cropped and straightened in ImageJ (NIH) of wildtype, mig-15, and mig-15(OE) in wyIs685. (H) Quantification of synapses in (E-G) using a one-way ANOVA followed by multiple comparisons. The observed data was normally distributed. Asterisks denote comparison to N2, except when accompanied with a bar, which denotes the two genotypes being compared: ***: p = 0.0001, ****: p < 0.0001. Scale bars = 10μm.  3.1.1 Pan-neuronal overexpression of mig-15 dramatically reduces locomotor coordination I then asked if mig-15 inhibits synapse formation across the entire nervous system. To test this, I conducted chromosomal integration of a construct of pan-neuronal overexpression of mig-15 (Prab-3::mig-15) to create three stable transgenic lines: mizIs33, mizIs34, and mizIs35.   30  These worms demonstrate a severe Unc phenotype when cultured on seeded NGM plates (Figure 4E).  3.1.2 Pan-neuronal overexpression dramatically reduces synaptic fluorescence To test whether the Unc phenotype of mizIs33 is the result of a reduction in the number of synapses, I crossed these animals into the pan-neuronal synaptic vesicle marker jsIs682(Prab-3::GFP::RAB-3). Synaptic fluorescence was calculated by averaging the mean pixel brightness, subtracted for background fluorescence, in the dorsal nerve cord (DNC) of both wildtype (N2) and mig-15(OE) (mizIs34) worms. Pan-neuronal overexpression of mig-15 reduced synaptic fluorescence in the DNC by 55% (Figure 2B), suggesting that mig-15 may function as a negative regulator of synapse formation.      31   Figure 2. Pan-neuronal overexpression of mig-15 decreases pan-neuronal synaptic fluorescence. (A) Quantification of DNC synaptic fluorescence in (B-D). N2: 14719±1919 a.u., mizIs33: 6572.91±1784 a.u., and sup3 mizIs33: 16755±2207 a.u. Error bars represent SD. Statistical comparisons were done using one-way ANOVA followed by multiple comparisons between N2 and mizIs33 and between mizIs33 and sup3 mizIs33. (B-D) Images of dorsal cord of pan-neuronal synapse-labelled worms (B: wildtype, C: mig-15 overexpression, D: mig-15(OE) + suppressor mutation). Images were straightened and processed in ImageJ (NIH). Scale bars = 10μm.  3.2 Forward screen to identify suppressors for mig-15-mediated inhibition of synapses To identify other genes that function with mig-15, I took advantage of the Unc phenotype induced by pan-neuronal overexpression of mig-15. mizIs33 worms were mutagenized (see   32  chapter 2.6 for methods) and progeny were screened for rescued locomotion. Animals with rescued locomotion may carry mutations in genes required for mig-15-mediated loss of synapses. I screened ~20,000 F2 animals and identified eight potential suppressor lines. All suppressor mutations identified were recessive. I then conducted complementation tests and found that the majority of suppressors were in the same complementation group (Figure 3, below).    Figure 3. Forward screen of the Unc phenotype of mig-15(OE) reveals a suppressor mutation. Above: Illustrative diagram of suppressor screen of mig-15 pan-neuronal overexpression in mizIs33. Lightning bolt represents EMS-induced mutagenesis. Under each stereoscope image is a representative image of synaptic fluorescence of the DNC. Below: suppressor complementation chart. 3.2.1 Suppression of mizIs33 Unc phenotype rescues loss of synaptic fluorescence  To confirm that the suppressor phenotype is the result of rescued synapse formation, I crossed the strongest suppressor line, sup3 mizIs33, into the pan-neuronal synaptic marker jsIs682. These animals displayed synaptic fluorescence indistinguishable from wildtype,   33  suggesting that the mutation in sup3 is reversing the effects of mig-15 overexpression on synapse number (Figure 2A-D). 3.2.2 SNP mapping reveals the location of sup3 and mizIs33 to be chromosome III  Next, I roughly mapped the location of the suppressor mutation using chromosomal mapping described in Davis et al. (2005). The candidate suppressor sup3 mizIs33 was chosen due to its relatively coordinated locomotion. Using this method, I found that both the sup3 mutation and mizIs33 are on chromosome III (data not shown). 3.2.3 Whole Genome Sequencing identifies Y32H12A.8 as a candidate gene sup3 mizIs33 was sent for whole genome sequencing (The Centre for Applied Genomics, Ontario). The results are outlined in Table 2. From this list, three candidate genes were tested for suppression of the Unc phenotype of mizIs33: grk-2, R08D7.5, and Y32H12A.8. Of these candidates, I used two deletion alleles of Y32H12A.8, tm419 and tm422 (Figure 5) and found that Y32H12A.8 was able to suppress the Unc phenotype of pan-neuronal mig-15 overexpression. Since Y32H12A.8 and mizIs33 are genetically linked on chromosome III, and integrated transgenes have lower rates of recombination, I used an alternate integrant of the Prab-3::mig-15 transgene, mizIs34. Y32H12A.8(tm419);mizIs34 and Y32H12A.8(tm422);mizIs34 animals both exhibit rescued locomotion compared with mizIs34 (Figure 4D,E). In the process of generating these strains I observed that heterozygous Y32H12A.8 mutant worms with homozygous mizIs34 do not have rescued locomotion, indicating that Y32H12A.8 functions as a recessive gene. To further confirm the identify of Y32H12A.8 as the suppressor identified in the genetic screen, I performed a complementation test between Y32H12A.8(tm419);mizIs34 and sup3 mizIs33 (i.e. one copy of tm419 and one copy of sup3 is sufficient to rescue the Unc phenotype in mig-15(OE)   34  worms, expressing one copy of mizIs33 and one copy of mizIs34) and found that Y32H12A.8 and sup3 fail to complement. This suggests they are both mutants in the same gene. Table 3. Candidate mig-15(OE) suppressor genes from whole genome sequencing. (The Centre for Applied Genomics, Ontario) Calls were filtered to only include missense or nonsense mutations in coding exons of genes. N2 strain was used as a reference. LG position wt mut type CDS gene name amino acid change III 73786 G A SNV F54C4.3 attf-3 H->Y III 679111 G A SNV W02B3.2 grk-2 A->T III 5028954 A G SNV R144.2 pcf-11 N->S III 5396683 G A SNV Y32H12A.8 Y32H12A.8 S->F III 5565332 G A SNV F09F7.2 mlc-3 E->K III 5723996 G A SNV B0336.1 wrm-1 V->I III 6197507 G A SNV C23G10.7 C23G10.7 L->F III 7113204 G A SNV B0280.8 nhr-10 H->Yx III 7878444 A T SNV C02C2.5 sup-18 V->D III 8974511 G A SNV R08D7.5 R08D7.5 G->E III 9812125 G A SNV ZK632.5 ZK632.5 G->E III 12182974 G A SNV Y75B8A.10 Y75B8A.10 P->S     35    Figure 4. wdr-24 rescues synaptic defects of mig-15 overexpression (A) Diagram of amino acid-sensing branch upstream of TORC1. (B-D) Representative images of presynaptic terminals in DA9 in wdr-24, and wdr-24;mizIs33 worms in wyIs685. Green  = CLA-1 active zone protein, magenta = RAB-3 synaptic vesicles. (E,F) Stereoscope images of mig-15 overexpression (mizIs34), and wdr-24;mizIs34.  (G-I) Stacks of DA9 synaptic domains, cropped and straightened in ImageJ (NIH) of wildtype, mig-15, and mig-15(OE) in wyIs685. (J) Quantification of DA9 synapse number for groups in (B-D). N2: 19.5±3.8, mig-15(rh148): 24.7±4.9, wdr-24(tm419): 23.3±3.2, mizIs33: 11.6±2.2, wdr-24:mizIs33: 16.64±4.1,  wdr-24;mizIs33;Ex[Prab-3::wdr-24]: 10±2.2. Scale bars = 10μm    36  3.2.4 Multiple sequence alignment of Y32H12A.8 and its orthologs I next asked whether Y32H12A.8 is orthologous to genes in related organisms. I conducted a Basic Local Alignment Search Tool (NCBI BLAST) search (NIH) and found that Y32H12A.8 is conserved with orthologs of the mammalian WDR24 in other landmark organisms. To confirm that this orthology is robust, I used the DRSC Integrative Ortholog Prediction Tool to compare mammalian WDR24 and C. elegans Y32H12A.8 are found that they are reciprocally orthologous to each other (DIOPT). I conducted a multiple sequence alignment with amino acid sequences from M. musculus, D. melanogaster, D. rerio, and H. sapiens (Clustal Omega). T-Coffee was used to generate a multiple alignment sequence file and the alignment was rendered in ESPript. Surprisingly, while the other species have protein lengths ranging from 776 to 790 amino acids, C. elegans Y32H12A.8 has a predicted structure of 3849 amino acids (Figure 5). The non-conserved domain is conserved only with three closely related Caenorhabditis species C. briggsae, C. remanei, and C. sp34 (NCBI BLAST).   37        (A) Multiple alignment of conserved amino acid sequence of C. elegans Y32H12A.8 with WDR24 orthologs from D. melanogaster, D.rerio, H. sapiens, and M. musculus. (B) Full alignment of the Y32H12A.8 amino acid sequence. The S1794F mutation induced from the forward screen in chapter 3.2 is labelled in black and denoted with an arrow.  Figure 5. Multiple alignment reveals a SF missense mutation outside the conserved domain of wdr-24.   38   3.2.5 Y32H12A.8 or wdr-24 rescues loss of synapses in mizIs33 I tentatively named Y32H12A.8 wdr-24, after its mammalian ortholog WDR24. To confirm that the rescued locomotion in wdr-24;mizIs34 animals is due to increased synapse number, I compared the number of synapses in the DA9 motor neuron in mizIs33;wyIs685 and wdr-24,mizIs33;wyIs685 worms. Loss of wdr-24 significantly increased synapse number in mizIs33 animals, suggesting that mig-15 requires wdr-24 (at least in part) to inhibit synapse formation (Figure 4C,H,J).  3.2.6 Conserved domain of wdr-24 rescues increased synapse number in wdr-24;mizIs33 To understand whether the conserved region of wdr-24 described in section 3.2.4 is required for the suppression of the mig-15(OE) synaptic phenotype, I cloned the wildtype cDNA of the conserved domain of wdr-24 under the pan-neuronal promoter of rab-3. I expressed this clone (pFM3) in wdr-24(tm419);mizIs33;wyIs685 animals via microinjection at a concentration of 1 ng/µL. Indeed, pan-neuronal expression of wdr-24 conserved cDNA recapitulated the loss of synapses seen in mizIs33 animals (Figure 4D,I,J). This suggests that wdr-24 functions with mig-15 to inhibit synapse formation via its conserved domain, and therefore implying that this pathway may be functionally conserved in other organisms. 3.2.7 wdr-24 mutants have increased synapse number in DA9 Loss of mig-15 results in increased synapse number in DA9, and overexpression of mig-15 in neurons requires wdr-24 for its characteristic Unc phenotype and coinciding loss of synapses (Figure 4B,G,J, Chen et al. 2018). This suggests that mig-15 functions through wdr-24 to decrease synapse number. Therefore, I expected wdr-24 mutants to show a similar synaptic   39  phenotype to mig-15 animals (i.e. increased synapse number). Indeed, wdr-24 animals have a significant increase in synapses in DA9 compared to N2 and similar to that of mig-15 (Figure 4J).    40  Chapter 4: Separating MIG-15-mediated inhibition of synapse formation from synaptic tiling  4.1 MIG-15-GATOR axis controls synapse formation independently of synaptic tiling One puzzling aspect of mig-15 function is that it controls both synaptic tiling and synapse formation. In Chen et al. we observed that mig-15 mutants have a more severe synaptic tiling defect than rap-2 and plx-1, and that mig-15;rap-2 double mutants have the same tiling defect as mig-15 single mutants (Chen et al. 2019, Figure 6D,E,G). The simplest explanation for this is that mig-15 has a larger synaptic tiling defect due to its increased number of synapses. In the absence of plx-1, rap-2, or mig-15, the border between the DA8 and DA9 synaptic domains is abolished, resulting in posterior and anterior of DA8 and DA9 synaptic domains respectively. If the length of the synaptic domain depends (in part) on the number of synapses in that domain, then without a mechanism to maintain the synaptic tiling border, the length of overlap between the domains may depend on how many synapses exist in each domain. Interestingly, wdr-24 mutants do not have a synaptic tiling (Figure 5C,G) defect but do have increased synapse number in DA9 (Figures 4H). If it is the case that mig-15 has an enhanced synaptic tiling defect due do its increased synapse number, then increasing synapse number (with wdr-24) in a tiling defective background mutant (plx-1) should enhance the synaptic tiling defect. Indeed, I found that wdr-24;plx-1 worms have a larger synaptic tiling phenotype similar to that of mig-15 single mutants, suggesting that in the absence of synaptic tiling, an increase in synapse number can enhance the synaptic tiling defect (Figure 6).   41  Chapter 5: Discussion I report a novel role for the previously uncharacterized gene Y32H12A.8. Named wdr-24, this gene encodes a protein orthologous to mammalian WDR24, a key signaling component upstream of TORC1. I found that wdr-24 is necessary for mig-15 overexpression-induced Unc phenotype and decrease in synapse number, and that wdr-24 mutants have an increase in synapse number akin to mig-15 animals. These findings provide novel insights into the function of WDR24, a relatively novel protein not yet characterized in worms and whose function is not yet fully understood.  5.1 wdr-24 suppression of mig-15(OE) phenotypes I was able to show that wdr-24;mizIs33 worms have significantly more synapses than mizIs33 worms. However, the rescue is incomplete as wdr-24;mizIs33 animals have fewer synapses than wdr-24 single mutants. Furthermore, wdr-24;plx-1 fails to enhance plx-1 synaptic tiling defect to the same significance as mig-15 defects. Together these discrepancies suggest that mig-15 inhibits synapse formation through a number of pathways, of which WDR24 is one component. To test this hypothesis, we could conduct additional forward screens to find other suppressors of mig-15 overexpression. However, it is possible that our screen reached saturation, and that mutations which suppress mig-15 in the TOR pathway are largely lethal, precluding their identification in a forward screen. Another possible explanation is that a complete loss of GATOR2 is necessary for suppression of mig-15(OE) induced loss of synapses. The 893 bp in-frame deletion in wdr-24(tm419) (Figure 5) may knock down GATOR2 function, but not abolish it altogether. In this case, the locomotor phenotype may rescue while the synaptic phenotype only partially rescues.   42  To test this hypothesis, I would generate double or triple mutants of wdr-24, npp-18, and npp-20, to abolish GATOR2 activity in a mig-15(OE) background and assess the synaptic phenotype.  5.2 mig-15 functions through GATOR2 to inhibit synapse formation I demonstrated that mig-15 requires wdr-24 to inhibit synapse formation. Mammalian WDR24 is known to form a GATOR2 complex with four other proteins: SEH1L, SEC13, MIOS, and WDR59. In C. elegans, orthologs exist for SEC13 (npp-18) and SEH1L (npp-20).  wdr-24 may function in the GATOR2 complex downstream of mig-15 to inhibit synapse formation. To test the possibility that mig-15 requires GATOR2 to inhibit synapses, I crossed npp-18 and npp-20 mutants into mizIs34, a mig-15 overexpression line with an Unc phenotype. Both npp-18;mizIs34 and npp-20;mizIs34 worms had rescued locomotion, suggesting that mig-15 overexpression requires GATOR2 to induce an Unc phenotype. However, since this result was not quantified and the strains were lost, these conclusions remain to be validated.  5.3 Neuron-specific knockdown of Raptor/DAF-15 I have shown a possible role for GATOR1 and GATOR2 in mig-15-mediated inhibition of synapse formation. The GATOR1 and GATOR2 complexes function to regulate TORC1 activation, and thus the role of TOR in synapse formation is of interest. While TOR is implicated in several critical cellular processes including autophagy, protein synthesis, and lipid synthesis (Lipton and Sahin 2014), it is not known whether mig-15/TNIK functions via the TOR pathway to inhibit synapse formation. GATOR2 activates TORC1 via GATOR1, which is defined by the binding of daf-15/Raptor to let-363/TOR. Therefore, mig-15 may inhibit synapse formation through daf-15. In C. elegans, daf-15 mutants are homozygous lethal (arresting at L3 dauer-like stage) (Albert and Riddle 1988). To address the role of daf-15 in the nervous system, I used   43  CRISPR/Cas9 gene editing technology to endogenously tag DAF-15 with the AID degron (Zhang et al. 2015). Briefly, a protein of interest is tagged with a protein motif important for targeted degradation, or degron, of 48 amino acids in length and TIR1, the substrate recognition component of the SCF E3 ubiquitin ligase complex, is expressed either as an extrachromosomal array (relatively efficient to acquire but with mosaic expression) or a chromosomally integrated transgene (offering stable expression levels but labour intensive to acquire). In the presence of the phytohormone auxin, TIR1 recognizes auxin and targets them for degradation by the proteasome. This system is ideal because it exploits the endogenous degradation machinery of the worm and allows for tissue-specific protein degradation (by expressing TIR1 under the promoter of interest) with temporal specificity (by treating worms to auxin at any timepoint).  5.4 Amino acid sensing role of GATOR2 One key aspect of WDR24 that I have not addressed is its role in nutrient sensing. Perhaps the most important function of the GATOR2 complex is its input by amino acid sensors such as Sestrin, CASTOR, and SAMTOR (Chantranupong et al. 2014; Cai et al. 2016; Lee, Cho, and Karin 2016; Saxton et al. 2016; Saxton and Sabatini 2017). Given the critical role TORC1 plays in nutrient sensing, wdr-24 likely receives input from nutrient sensing genes C. elegans homologs. Indeed, I have observed that when starved, mizIs33, mizIs34, and mizIs35 animals have near wildtype locomotion, hinting at the inhibition of wdr-24 by amino acid sensors in the absence of food. To test whether this is a result of changes in wdr-24 function and not a signaling cascade resulting in the rewiring of the motor circuit in response to starvation, I would propose imaging mig-15(OE) worms grown off food at a variety of time points. If synapse number increases with time off food, that suggests a role of nutrient sensing in synaptic maintenance.   44  Given the importance of synapse formation in normal brain development as well as issues of food insecurity and malnutrition in developing countries, this is an area worthy of further investigation. 5.5 Non-conserved domain of wdr-24/Y32H12A.8 Our forward screen of mig-15(OE)/mizIs33 worms revealed that a missense mutation in the gene Y32H12A.8 can suppress the locomotor and synaptic phenotype of mizIs33. An alignment of the amino acid sequence of this protein revealed that the C. elegans Y32H12A.8 has a conserved domain at the N-terminus, as well as a much larger non-conserved domain making up the rest of the gene. Interestingly, the suppressor mutation (S1794F) is in the non-conserved region of the gene. It is puzzling that the missense mutation which induces the rescue of mizIs33 locomotion exists outside the conserved domain of wdr-24. One possible explanation is that Y32H12A.8(S1794F) confers a conformational change to the protein structure and prevents the function of WDR24. This could mean that the Y32H12A.8(S1794F) allele is unable to form a functional GATOR2 complex with NPP-18 and NPP-20, resulting in a strong loss of GATOR2 activity. It is also possible that WDR24, which is required for inhibition of GATOR1, requires an C-terminal region for its interaction with GATOR1. A conformational change of the non-conserved protein sequence induced by a missense mutation could explain why six of the eight suppressors identified were in the same complementation group; any missense mutation expressed far enough on the 5’ end of the 9033 bp non-conserved protein-coding region of Y32H12A.8 which confers a conformational change may have a significant effect on protein function. To test this hypothesis, I would propose testing the effect of a variety of missense mutations in the non-conserved domain (available from the Million Mutation Project collection) on the Unc phenotype of mig-15(OE). If these mutations rescue the Unc phenotype, it would   45  suggest that the non-conserved domain of the Y32H12A.8 protein is required for Y32H12A.8 function in the MIG-15 synapse inhibition pathway. 5.6 Inhibition of GATOR1 by GATOR2 may be necessary for mig-15(OE) induced loss of synapses and coordinated locomotion. GATOR2 functions as a positive regulator of TORC1 complex via inhibiting the GATOR1 complex which negatively regulates TORC1. The GATOR1 complex is comprised of three subunits: Nprl2, Nprl3, and DEPDC5 (Bar-peled et al. 2013). Ordinarily, GATOR2 inhibits activity of GATOR1 via WDR24 (Cai et al. 2016). GATOR1 then inhibits TORC1 activity via GAP activity, stimulating GTP hydrolysis by the Rag GTPases. Nprl2 is necessary for this process as its Arg78 residue functions as the GAP of GATOR1 (K. Shen et al. 2019). To test whether wdr-24 functions with mig-15 via its inhibition of GATOR1, I would examine the locomotor and synaptic phenotypes of nprl-2;wdr-24;mizis33 worms. If loss wdr-24 rescues the Unc phenotype of mizIs33 via its inhibition of  GATOR1, then ‘reinhibition’ of GATOR1 in a wdr-24;mizIs33 background should reverse the rescue and restore the Unc phenotype. In other words, I reason that nprl-2;wdr-24;mizIs34 worms should have an Unc phenotype similar to that of mizIs34 and more severe than wdr-24;mizIs33. In essence, nprl-2;wdr-24 animals should phenocopy nprl-2 animals in a mig-15(OE) background. If so, this would suggest that mig-15 inhibits synapse formation via GATOR2 inhibition of GATOR1. To confirm that this is not the result of the pre-existing Unc phenotype in nprl-2 mutants, I propose comparing DA9 synapse number in nprl-2 worms to wdr-24;nprl-2. If the number of synapses in both genotypes reflect that of nprl-2 worms, this would suggest that nprl-2 (and thus GATOR1) functions downstream of GATOR2 to regulate synapse number.   46  Taken together, these findings may implicate the amino acid sensing pathway in the TOR network in a novel pathway underlying the inhibition of synapse formation, and may even be functionally conserved across species.     47    Figure 6. wdr-24 enhances synaptic tiling defect of plx-1 (A) Schematic diagram of DA8/DA9 synaptic tiling as described in Chen et al. (2018). (B-E) Representative images of synaptic tiling marker mizIs3 (RFP: DA9 synaptic domain, GFP: DA8 synaptic domain). (G) Quantification of synaptic tiling defect in N2, wdr-24, plx-1, plx-1;wdr-24, and mig-15 worms.      48  References  Akin, Orkun, Bryce T Bajar, Mehmet F Keles, Mark A Frye, S Lawrence Zipursky, Orkun Akin, Bryce T Bajar, Mehmet F Keles, Mark A Frye, and S Lawrence Zipursky. 2019. “Cell-Type-Specific Patterned Stimulus-Independent Neuronal Activity in the Drosophila Visual System during Synapse Formation Report Cell-Type-Specific Patterned Stimulus-Independent Neuronal Activity in the Drosophila Visual System during Synapse Formatio.” Neuron, 1–11. https://doi.org/10.1016/j.neuron.2019.01.008. Albert, Patrice S., and Donald L. Riddle. 1988. “Mutants of Caenorhabditis Elegans That Form Dauer-like Larvae.” Developmental Biology 126 (2): 270–93. https://doi.org/10.1016/0012-1606(88)90138-8. Anazi, Shams, Hanan E. Shamseldin, Dhekra AlNaqeb, Mohamed Abouelhoda, Dorota Monies, Mustafa A. Salih, Khalid Al-Rubeaan, and Fowzan S. Alkuraya. 2016. “A Null Mutation in TNIK Defines a Novel Locus for Intellectual Disability.” Human Genetics 135 (7): 773–78. https://doi.org/10.1007/s00439-016-1671-9. Au, Vinci, Erica Li-leger, Greta Raymant, Stephane Flibotte, George Chen, Kiana Martin, Lisa Fernando, et al. 2019. “CRISPR/Cas9 Methodology for the Generation of Knockout Deletions in Caenorhabditis Elegans.” Genes, Genomes, Genetics 9 (January): 135–44. https://doi.org/10.1534/g3.118.200778. Bar-peled, Liron, Lynne Chantranupong, Andrew D Cherniack, and Walter W Chen. 2013. “A Tumor Suppressor Complex with GAP Activity for the Rag GTPases That Signal Amino Acid Sufficiency to MTORC1.” Science 340 (6136): 1100–1106. https://doi.org/10.1126/science.1232044.A. Birdsall, Veronica, and Clarissa L. Waites. 2019. “Autophagy at the Synapse.” Neuroscience Letters 697 (May): 24–28. https://doi.org/10.1016/j.neulet.2018.05.033. Blackwell, T. Keith, Aileen K. Sewell, Ziyun Wu, and Min Han. 2019. “TOR Signaling in Caenorhabditis Elegans Development, Metabolism, and Aging.” Genetics 213 (2): 329–60. https://doi.org/10.1534/genetics.119.302504. Brenner, S. 1974. “The Genetics of Caenorhabditis Elegans.” Genetics 77 (1): 71–94. https://doi.org/10.1002/cbic.200300625.   49  Brown, Eric J., Mark W. Albers, Tae Bum Shin, Kazuo Ichikawa, Curtis T. Keith, William S. Lane, and Stuart L. Schreiber. 1994. “A Mammalian Protein Targeted by G1-Arresting Rapamycin–Receptor Complex.” Nature 369 (6483): 756–58. https://doi.org/10.1038/369756a0. Cai, Weili, Youheng Wei, Michal Jarnik, John Reich, and Mary A. Lilly. 2016. “The GATOR2 Component Wdr24 Regulates TORC1 Activity and Lysosome Function.” PLoS Genetics 12 (5): 1–28. https://doi.org/10.1371/journal.pgen.1006036. Chantranupong, Lynne, Rachel L. Wolfson, Jose M. Orozco, Robert A. Saxton, Sonia M. Scaria, Liron Bar-Peled, Eric Spooner, Marta Isasa, Steven P. Gygi, and David M. Sabatini. 2014. “The Sestrins Interact with Gator2 to Negatively Regulate the Amino-Acid-Sensing Pathway Upstream of MTORC1.” Cell Reports 9 (1): 1–8. https://doi.org/10.1016/j.celrep.2014.09.014. Chapman, Jamie O., Hua Li, and Erik A. Lundquist. 2008. “The MIG-15 NIK Kinase Acts Cell-Autonomously in Neuroblast Polarization and Migration in C. Elegans.” Developmental Biology 324 (2): 245–57. https://doi.org/10.1016/j.ydbio.2008.09.014. Chen, Xi, Akihiro C E Shibata, Ardalan Hendi, Mizuki Kurashina, Ethan Fortes, Nicholas L Weilinger, Brian A Macvicar, and Hideji Murakoshi. 2018. “Rap2 and TNIK Control Plexin-Dependent Tiled Synaptic Innervation in C . Elegans.” ELife 7:e38801: 1–25. Chisholm, Andrew D., Harald Hutter, Yishi Jin, and William G. Wadsworth. 2016. The Genetics of Axon Guidance and Axon Regeneration in Caenorhabditis Elegans. Genetics. Vol. 204. https://doi.org/10.1534/genetics.115.186262. Chung, Jongkyeong, Calvin J Kuo, Gerald R. Crabtree, and John Blenis. 1992. “Rapamycin-FKBP Specifically Blocks Growth-Dependent Activation of and Signaling by the 70 Kd S6 Protein Kinases.” Cell 69 (7): 1227–36. https://doi.org/10.1016/0092-8674(92)90643-Q. Cloetta, D., Venus Thomanetz, Constanze Baranek, Regula M Lustenberger, Shuo Lin, Filippo Oliveri, Suzana Atanasoski, and M. A. Ruegg. 2013. “Inactivation of MTORC1 in the Developing Brain Causes Microcephaly and Affects Gliogenesis.” Journal of Neuroscience 33 (18): 7799–7810. https://doi.org/10.1523/jneurosci.3294-12.2013. Crawley, Oliver, Andrew C. Giles, Muriel Desbois, Sudhanva Kashyap, Rayna Birnbaum, and Brock Grill. 2017. “A MIG-15/JNK-1 MAP Kinase Cascade Opposes RPM-1 Signaling in   50  Synapse Formation and Learning.” PLoS Genetics 13 (12): 1–27. https://doi.org/10.1371/journal.pgen.1007095. Dan, Ippeita, Norinobu M. Watanabe, and Akihiro Kusumi. 2001. “The Ste20 Group Kinases as Regulators of MAP Kinase Cascades.” Trends in Cell Biology. https://doi.org/10.1016/S0962-8924(01)01980-8. Davis, M Wayne, Marc Hammarlund, Tracey Harrach, Patrick Hullett, Shawn Olsen, and Erik M Jorgensen. 2005. “Rapid Single Nucleotide Polymorphism Mapping in C. Elegans.” BMC Genomics 6: 118. https://doi.org/10.1186/1471-2164-6-118. Dunlop, E. A., and A. R. Tee. 2014. “MTOR and Autophagy: A Dynamic Relationship Governed by Nutrients and Energy.” Seminars in Cell and Developmental Biology 36: 121–29. https://doi.org/10.1016/j.semcdb.2014.08.006. Fu, C Alan, Mary Shen, Betty C B Huang, Joe Lasaga, Donald G Payan, Ying Luo, C Alan Fu, et al. 1999. “TNIK, a Novel Member of the Germinal Center Kinase Family That Activates the c-Jun N-Terminal Kinase Pathway and Regulates the Cytoskeleton.” Cell Biology and Metabolism 274 (43): 30729–37. Gan, Xiaoqing, Jiyong Wang, Chen Wang, Eeva Sommer, Tohru Kozasa, Srinivasa Srinivasula, Dario Alessi, Stefan Offermanns, Melvin I Simon, and Dianqing Wu. 2012. “PRR5L Degradation Promotes MTORC2-Mediated PKC-δ Phosphorylation and Cell Migration Downstream of Gα 12.” Nature Cell Biology 14 (7): 686–96. https://doi.org/10.1038/ncb2507. Glatt, S. J., I. P. Everall, W. S. Kremen, J. Corbeil, R.  a ik, N. Khanlou, M. Han, C.-C. Liew, and M. T. Tsuang. 2005. “Comparative Gene Expression Analysis of Blood and Brain Provides Concurrent Validation of SELENBP1 Up-Regulation in Schizophrenia.” Proceedings of the National Academy of Sciences 102 (43): 15533–38. https://doi.org/10.1073/pnas.0507666102. Gu, Xin, Jose M. Orozco, Robert A. Saxton, Kendall J. Condon, Grace Y. Liu, Patrycja A. Krawczyk, Sonia M. Scaria, J. Wade Harper, Steven P. Gygi, and David M. Sabatini. 2017. “SAMTOR Is an S -Adenosylmethionine Sensor for the MTORC1 Pathway.” Science 358 (6364): 813–18. https://doi.org/10.1126/science.aao3265. Guertin, David A., Deanna M. Stevens, Maki Saitoh, Stephanie Kinkel, Katherine Crosby, Joon   51  Ho Sheen, David J. Mullholland, Mark A. Magnuson, Hong Wu, and David M. Sabatini. 2009. “MTOR Complex 2 Is Required for the Development of Prostate Cancer Induced by Pten Loss in Mice.” Cancer Cell 15 (2): 148–59. https://doi.org/10.1016/j.ccr.2008.12.017. Hara, Kenta, Yoshiko Maruki, Xiaomeng Long, Ken-ichi Yoshino, Noriko Oshiro, Sujuti Hidayat, Chiharu Tokunaga, Joseph Avruch, and Kazuyoshi Yonezawa. 2002. “Raptor, a Binding Partner of Target of Rapamycin (TOR), Mediates TOR Action.” Cell 110 (2): 177–89. https://doi.org/10.1016/S0092-8674(02)00833-4. Hawk, Josh D, Ana C Calvo, Ping Liu, Agustin Almoril-Porras, Ahmad Aljobeh, María Luisa Torruella-Suárez, Ivy Ren, et al. 2018. “Integration of Plasticity Mechanisms within a Single Sensory Neuron of C. Elegans Actuates a Memory.” Neuron 97 (2): 356-367.e4. https://doi.org/10.1016/j.neuron.2017.12.027. Hendi, Ardalan, Mizuki Kurashina, and Kota Mizumoto. 2019. “Intrinsic and Extrinsic Mechanisms of Synapse Formation and Specificity in C. Elegans.” Cellular and Molecular Life Sciences 76 (14): 2719–38. https://doi.org/10.1007/s00018-019-03109-1. Hsu, Yu-Tien, Jie Li, Dick Wu, Thomas C. Südhof, and Lu Chen. 2019. “Synaptic Retinoic Acid Receptor Signaling Mediates MTOR-Dependent Metaplasticity That Controls Hippocampal Learning.” Proceedings of the National Academy of Sciences 116 (14): 7113–22. https://doi.org/10.1073/pnas.1820690116. Hussain, Natasha K, Honor Hsin, Richard L Huganir, and Morgan Sheng. 2011. “MINK and TNIK Differentially Act on Rap2-Mediated Signal Transduction to Regulate Neuronal Structure and AMPA Receptor Function.” Journal of Neuroscience 30 (44): 14786–94. https://doi.org/10.1523/JNEUROSCI.4124-10.2010.MINK. Jacinto, Estela, Valeria Facchinetti, Dou Liu, Nelyn Soto, Shiniu Wei, Sung Yun Jung, Qiaojia Huang, Jun Qin, and Bing Su. 2006. “SIN1/MIP1 Maintains Rictor-MTOR Complex Integrity and Regulates Akt Phosphorylation and Substrate Specificity.” Cell 127 (1): 125–37. https://doi.org/10.1016/j.cell.2006.08.033. Jacinto, Estela, Robbie Loewith, Anja Schmidt, Shuo Lin, Markus A. Rüegg, Alan Hall, and Michael N. Hall. 2004. “Mammalian TOR Complex 2 Controls the Actin Cytoskeleton and Is Rapamycin Insensitive.” Nature Cell Biology 6 (11): 1122–28. https://doi.org/10.1038/ncb1183.   52  Jung, Hosung, Christos G. Gkogkas, Nahum Sonenberg, and Christine E. Holt. 2014. “Remote Control of Gene Function by Local Translation.” Cell 157 (1): 26–40. https://doi.org/10.1016/j.cell.2014.03.005. Kassai, Hidetoshi, Yuki Sugaya, Shoko Noda, Kazuki Nakao, Tatsuya Maeda, Masanobu Kano, and Atsu Aiba. 2014. “Selective Activation of MTORC1 Signaling Recapitulates Microcephaly, Tuberous Sclerosis, and Neurodegenerative Diseases.” Cell Reports 7 (5): 1626–39. https://doi.org/10.1016/j.celrep.2014.04.048. Kato, Taro, Seung Hahm, and Ronald S Duman. 2019. “Sestrin Modulator NV-5138 Produces Rapid Antidepressant Effects via Direct MTORC1 Activation Graphical Abstract Find the Latest Version :” 129 (6): 2542–54. Kawabe, Hiroshi, Antje Neeb, Kalina Dimova, Samuel M. Young, Michiko Takeda, Shutaro Katsurabayashi, Miso Mitkovski, et al. 2010. “Regulation of Rap2A by the Ubiquitin Ligase Nedd4-1 Controls Neurite Development.” Neuron 65 (3): 358–72. https://doi.org/10.1016/j.neuron.2010.01.007. Kim, Do Hyung, Dos D. Sarbassov, Siraj M. Ali, Robert R. Latek, Kalyani V.P. Guntur, Hediye Erdjument-Bromage, Paul Tempst, and David M. Sabatini. 2003. “GβL, a Positive Regulator of the Rapamycin-Sensitive Pathway Required for the Nutrient-Sensitive Interaction between Raptor and MTOR.” Molecular Cell 11 (4): 895–904. https://doi.org/10.1016/S1097-2765(03)00114-X. Kim, Joungmok, Mondira Kundu, Benoit Viollet, and Kun-Liang Guan. 2011. “AMPK and MTOR Regulate Autophagy through Direct Phosphorylation of Ulk1.” Nature Cell Biology 13 (2): 132–41. https://doi.org/10.1038/ncb2152. Kwon, Chang Hyuk, Bryan W Luikart, Craig M Powell, Jing Zhou, Sharon A Matheny, Wei Zhang, Yanjiao Li, Suzanne J Baker, and Luis F Parada. 2006. “Pten Regulates Neuronal Arborization and Social Interaction in Mice.” Neuron 50 (3): 377–88. https://doi.org/10.1016/j.neuron.2006.03.023. Larhammar, Martin, Sarah Huntwork-Rodriguez, York Rudhard, Arundhati Sengupta-Ghosh, and Joseph W. Lewcock. 2017. “The Ste20 Family Kinases MAP4K4, MINK1 and TNIK, Converge to Regulate Stress Induced JNK Signaling in Neurons.” The Journal of Neuroscience 37 (46): 0905–17. https://doi.org/10.1523/JNEUROSCI.0905-17.2017.   53  Ledda, Fernanda, and Gustavo Paratcha. 2017. “Mechanisms Regulating Dendritic Arbor Patterning.” Cellular and Molecular Life Sciences 74 (24): 4511–37. https://doi.org/10.1007/s00018-017-2588-8. Lee, Jun Hee, Uhn Soo Cho, and Michael Karin. 2016. “Sestrin Regulation of TORC1: Is Sestrin a Leucine Sensor?” Science Signaling 9 (431): 5–10. https://doi.org/10.1126/scisignal.aaf2885. Li, Nanxin, Boyoung Lee, Rong-jian Liu, Mounira Banasr, Jason M Dwyer, Masaaki Iwata, Xiao-yuan Li, et al. 2010. “MTOR-Dependent Synapse Formation Underlies the Rapid Antidepressant Effects of NMDA Antagonists Published by : American Association for the Advancement of Science Stable URL : Http://Www.Jstor.Org/Stable/40799879 REFERENCES Linked References Are Available.” Science 5994 (August 2010): 959–64. Li, Xin, and Tianyan Gao. 2014. “MTORC2 Phosphorylates Protein Kinase Cζ to Regulate Its Stability and Activity.” EMBO Reports 15 (2): 191–98. https://doi.org/10.1002/embr.201338119. Lieberman, Ori J., Avery F. McGuirt, Guomei Tang, and David Sulzer. 2018. “Roles for Neuronal and Microglial Autophagy in Synaptic Pruning during Development.” Neurobiology of Disease, no. December 2017: 0–1. https://doi.org/10.1016/j.nbd.2018.04.017. Limerick, Gerard, Xia Tang, Won Suk Lee, Ahmed Mohamed, Aseel Al-Aamiri, and William G. Wadsworth. 2018. “A Statistically-Oriented Asymmetric Localization (SOAL) Model for Neuronal Outgrowth Patterning by Caenorhabditis Elegans UNC-5 (UNC5) and UNC-40 (DCC) Netrin Receptors.” Genetics 208 (1): 245–72. https://doi.org/10.1534/genetics.117.300460. Lipton, Jonathan O., and Mustafa Sahin. 2014. “The Neurology of MTOR.” Neuron 84 (2): 275–91. https://doi.org/10.1016/j.neuron.2014.09.034. Liu, Wanguo, Xiangyang Dong, Ming Mai, Ratnam S Seelan, Ken Taniguchi, Kausilia K Krishnadath, Kevin C Halling, et al. 2000. “Mutations in AXIN2 Cause Colorectal Cancer with Defective Mismatch Repair by Activating β-Catenin/TCF Signalling.” Nature Genetics 26 (2): 146–47. https://doi.org/10.1038/79859. Liu, Xiaojie, Yan Li, Laikang Yu, Casey R. Vickstrom, and Qing Song Liu. 2018. “VTA MTOR   54  Signaling Regulates Dopamine Dynamics, Cocaine-Induced Synaptic Alterations, and Reward.” Neuropsychopharmacology 43 (5): 1066–77. https://doi.org/10.1038/npp.2017.247. Mackinnon, A. Craig, Hiroshi Qadota, Kenneth R. Norman, Donald G. Moerman, and Benjamin D. Williams. 2002. “C. Elegans PAT-4/ILK Functions as an Adaptor Protein within Integrin Adhesion Complexes.” Current Biology 12 (10): 787–97. https://doi.org/10.1016/S0960-9822(02)00810-2. Mahmoudi, Tokameh, Vivian S.W. Li, Ser Sue Ng, Nadia Taouatas, Robert G.J. Vries, Shabaz Mohammed, Albert J. Heck, and Hans Clevers. 2009. “The Kinase TNIK Is an Essential Activator of Wnt Target Genes.” EMBO Journal 28 (21): 3329–40. https://doi.org/10.1038/emboj.2009.285. Meikle, Lynsey, Hiroaki Onda, Kristen Pollizzi, David J Kwiatkowski, Delia M Talos, Alexander Rotenberg, Mustafa Sahin, and Frances E Jensen. 2007. “A Mouse Model of Tuberous Sclerosis: Neuronal Loss of Tsc1 Causes Dysplastic and Ectopic Neurons, Reduced Myelination, Seizure Activity, and Limited Survival.” Journal of Neuroscience 27 (21): 5546–58. https://doi.org/10.1523/JNEUROSCI.5540-06.2007. Mello, Craig C, James M Kramer, Dan Stinchcomb, and Victor Ambros. 1991. “Efficient Gene Transfer in C.Elegans: Extrachromosomal Maintenance and Integration of Transforming Sequences.” EMBO Journal 10 (1): 3959–70. Menzies, Fiona M, Angeleen Fleming, Andrea Caricasole, Carla F Bento, Stephen P Andrews, Avraham Ashkenazi, Jens Füllgrabe, et al. 2017. “Autophagy and Neurodegeneration: Pathogenic Mechanisms and Therapeutic Opportunities.” Neuron. https://doi.org/10.1016/j.neuron.2017.01.022. Mizumoto, Kota, and Kang Shen. 2013. “Interaxonal Interaction Defines Tiled Presynaptic Innervation in C. Elegans.” Neuron 77 (4): 655–66. https://doi.org/10.1016/j.neuron.2012.12.031. Mohamed, Ahmed M., Jeffrey R. Boudreau, Fabian P S Yu, Jun Liu, and Ian D. Chin-Sang. 2012. “The Caenorhabditis Elegans Eph Receptor Activates NCK and N-WASP, and Inhibits Ena/VASP to Regulate Growth Cone Dynamics during Axon Guidance.” PLoS Genetics 8 (2). https://doi.org/10.1371/journal.pgen.1002513.   55  Pérez-Escudero, Alfonso, and Gonzalo G. De Polavieja. 2007. “Optimally Wired Subnetwork Determines Neuroanatomy of Caenorhabditis Elegans.” Proceedings of the National Academy of Sciences of the United States of America 104 (43): 17180–85. https://doi.org/10.1073/pnas.0703183104. Poinat, Patrice, Adèle De Arcangelis, Saris Sookhareea, Xiaoping Zhu, Edward M. Hedgecock, Michel Labouesse, and Elisabeth Georges-Labouesse. 2002. “A Conserved Interaction between Β1 Integrin/PAT-3 and Nck-Interacting Kinase/MIG-15 That Mediates Commissural Axon Navigation in C. Elegans.” Current Biology 12 (8): 622–31. https://doi.org/10.1016/S0960-9822(02)00764-9. Potkin, Steven G., Fabio Macciardi, Guia Guffanti, James H. Fallon, Qi Wang, Jessica A. Turner, Anita Lakatos, et al. 2010. “Identifying Gene Regulatory Networks in Schizophrenia.” NeuroImage 53 (3): 839–47. https://doi.org/10.1016/j.neuroimage.2010.06.036. Reya, T., and H Clevers. 2005. “Wnt Signalling in Stem Cells and Cancer. 434, 843–850 (2005).” Nature 434: 843–50. Robida-Stubbs, Stacey, Kira Glover-Cutter, Dudley W. Lamming, Masaki Mizunuma, Sri Devi Narasimhan, Elke Neumann-Haefelin, David M. Sabatini, and T. Keith Blackwell. 2012. “TOR Signaling and Rapamycin Influence Longevity by Regulating SKN-1/Nrf and DAF-16/FoxO.” Cell Metabolism 15 (5): 713–24. https://doi.org/10.1016/j.cmet.2012.04.007. Rogala, Kacper B., Xin Gu, Jibril F. Kedir, Monther Abu-Remaileh, Laura F. Bianchi, Alexia M. S. Bottino, Rikke Dueholm, et al. 2019. “Structural Basis for the Docking of MTORC1 on the Lysosomal Surface.” Science, October, eaay0166. https://doi.org/10.1126/science.aay0166. Sabers, C. J., M. M. Martin, G. J. Brunn, J. M. Williams, F. J. Dumont, G. Wiederrecht, and R. T. Abraham. 1995. “Isolation of a Protein Target of the FKBP12-Rapamycin Complex in Mammalian Cells.” Journal of Biological Chemistry 270 (2): 815–22. https://doi.org/10.1074/jbc.270.2.815. Sanes, Joshua R., and S. Lawrence Zipursky. 2010. “Design Principles of Insect and Vertebrate Visual Systems.” Neuron 66 (1): 15–36. https://doi.org/10.1016/j.neuron.2010.01.018. Sarbassov, D. D. 2005. “Phosphorylation and Regulation of Akt/PKB by the Rictor-MTOR   56  Complex.” Science 307 (5712): 1098–1101. https://doi.org/10.1126/science.1106148. Saxton, Robert A., Lynne Chantranupong, Kevin E. Knockenhauer, Thomas U. Schwartz, and David M. Sabatini. 2016. “Mechanism of Arginine Sensing by CASTOR1 Upstream of MTORC1.” Nature 536 (7615): 229–33. https://doi.org/10.1038/nature19079. Saxton, Robert A, and David M Sabatini. 2017. “MTOR Signaling in Growth, Metabolism, and Disease.” Cell. Elsevier Inc. https://doi.org/10.1016/j.cell.2017.02.004. Serafini, T, T E Kennedy, M J Galko, C Mirzayan, T M Jessell, and M Tessierlavigne. 1994. “The Netrins Define a Family of Axon Outgrowth-Promoting Proteins Homologous to C-Elegans Unc-6.” Cell 78 (3): 409–24. Shakir, M. Afaq, Jason S. Gill, and Erik A. Lundquist. 2006. “Interactions of UNC-34 Enabled with Rac GTPases and the NIK Kinase MIG-15 in Caenorhabditis Elegans Axon Pathfinding and Neuronal Migration.” Genetics 172 (2): 893–913. https://doi.org/10.1534/genetics.105.046359. Shen, Dan Na, Li Hui Zhang, Er Qing Wei, and Yi Yang. 2015. “Autophagy in Synaptic Development, Function, and Pathology.” Neuroscience Bulletin. https://doi.org/10.1007/s12264-015-1536-6. Shen, Kuang, Max L Valenstein, Xin Gu, and David M Sabatini. 2019. “Arg-78 of Nprl2 Catalyzes GATOR1-Stimulated GTP Hydrolysis by the Rag GTPases.” Journal of Biological Chemistry 294 (8): 2970–75. https://doi.org/10.1074/jbc.AC119.007382. Shen, Wei, and Barry Ganetzky. 2009. “Autophagy Promotes Synapse Development in Drosophila.” Journal of Cell Biology 187 (1): 71–79. https://doi.org/10.1083/jcb.200907109. Shitashige, Miki, Reiko Satow, Takafumi Jigami, Kazunori Aoki, Kazufumi Honda, Tatsuhiro Shibata, Masaya Ono, Setsuo Hirohashi, and Tesshi Yamada. 2010. “Traf2- and Nck-Interacting Kinase Is Essential for Wnt Signaling and Colorectal Cancer Growth.” Cancer Research 70 (12): 5024–33. https://doi.org/10.1158/0008-5472.CAN-10-0306. Stavoe, Andrea K.H., Sarah E Hill, David H Hall, and Daniel A. Colón-Ramos. 2016. “KIF1A/UNC-104 Transports ATG-9 to Regulate Neurodevelopment and Autophagy at Synapses.” Developmental Cell 38 (2): 171–85. https://doi.org/10.1016/j.devcel.2016.06.012.   57  Sulston, J.E., E. Schierenberg, J.G. White, and J.N. Thomson. 1983. “The Embryonic Cell Lineage of the Nematode Caenorhabditis Elegans.” Developmental Biology 100 (1): 64–119. https://doi.org/10.1016/0012-1606(83)90201-4. Taira, Kiyohito, Masato Umikawa, Kirniko Takei, Bat Erdene Myagmar, Manabu Shinzato, Noriko Machida, Hiroshi Uezato, Shigeo Nonaka, and Ken Ichi Kariya. 2004. “The Traf2- and Nck-Interacting Kinase as a Putative Effector of Rap2 to Regulate Actin Cytoskeleton.” Journal of Biological Chemistry 279 (47): 49488–96. https://doi.org/10.1074/jbc.M406370200. Takahashi, Takuya, Alyson Fournier, Fumio Nakamura, Li-Hsien Wang, Yasunori Murakami, Robert G. Kalb, Hajime Fujisawa, and Stephen M. Strittmatter. 1999. “Plexin-Neuropilin-1 Complexes Form Functional Semaphorin-3A Receptors.” Cell 99 (1): 59–69. https://doi.org/10.1016/S0092-8674(00)80062-8. Tang, Guomei, Kathryn Gudsnuk, Sheng Han Kuo, Marisa L. Cotrina, Gorazd Rosoklija, Alexander Sosunov, Mark S. Sonders, et al. 2014. “Loss of MTOR-Dependent Macroautophagy Causes Autistic-like Synaptic Pruning Deficits.” Neuron 83 (5): 1131–43. https://doi.org/10.1016/j.neuron.2014.07.040. Tavazoie, Sohail F, Veronica A Alvarez, Dennis A Ridenour, David J Kwiatkowski, and Bernardo L Sabatini. 2005. “Regulation of Neuronal Morphology and Function by the Tumor Suppressors Tsc1 and Tsc2.” Nature Neuroscience 8 (12): 1727–34. https://doi.org/10.1038/nn1566. Teuliere, J., C. Gally, G. Garriga, M. Labouesse, and E. Georges-Labouesse. 2011. “MIG-15 and ERM-1 Promote Growth Cone Directional Migration in Parallel to UNC-116 and WVE-1.” Development 138 (20): 4475–85. https://doi.org/10.1242/dev.061952. Thomanetz, Venus, Nico Angliker, Dimitri Cloëtta, Regula M Lustenberger, Manuel Schweighauser, Filippo Oliveri, Noboru Suzuki, and Markus A Rüegg. 2013. “Ablation of the MTORC2 Component Rictor in Brain or Purkinje Cells Affects Size and Neuron Morphology.” Journal of Cell Biology 201 (2): 293–308. https://doi.org/10.1083/jcb.201205030. Tokunaga, Chiharu, Ken-ichi Yoshino, Kazuyoshi Yonezawa, Sujuti Hidayat, Yoshiko Maruki, Noriko Oshiro, Xiaomeng Long, Kenta Hara, and Joseph Avruch. 2004. “Raptor, a Binding   58  Partner of Target of Rapamycin (TOR), Mediates TOR Action.” Cell 110 (2): 177–89. https://doi.org/10.1016/s0092-8674(02)00833-4. Wang, Q., E. I. Charych, V. L. Pulito, J. B. Lee, N. M. Graziane, R. A. Crozier, R. Revilla-Sanchez, et al. 2011. “The Psychiatric Disease Risk Factors DISC1 and TNIK Interact to Regulate Synapse Composition and Function.” Molecular Psychiatry 16 (10): 1006–23. https://doi.org/10.1038/mp.2010.87. Wang, Yuxiao, Huawei He, Nishi Srivastava, Sheikh Vikarunnessa, Yong Bin Chen, Jin Jiang, Christopher W. Cowan, and Xuewu Zhang. 2012. “Plexins Are GTPase-Activating Proteins for Rap and Are Activated by Induced Dimerization.” Science Signaling 5 (207): 1–25. https://doi.org/10.1126/scisignal.2002636. White, J. G., E. Southgate, J. N. Thomson, and S. Brenner. 1976. “The Structure of the Ventral Nerve Cord of Caenorhabditis Elegans.” Philosophical Transactions of the Royal Society B: Biological Sciences 275 (938): 327–48. https://doi.org/10.1098/rstb.1976.0086. Winberg, Margaret L, Jasprina N Noordermeer, Luca Tamagnone, Paolo M Comoglio, Melanie K Spriggs, Marc Tessier-Lavigne, and Corey S Goodman. 1998. “Plexin A Is a Neuronal Semaphorin Receptor That Controls Axon Guidance.” Cell 95 (7): 903–16. https://doi.org/10.1016/S0092-8674(00)81715-8. Xuan, Zhao, Laura Manning, Jessica Nelson, Janet E Richmond, Daniel A Colón-Ramos, Kang Shen, and Peri T Kurshan. 2017. “Clarinet (CLA-1), a Novel Active Zone Protein Required for Synaptic Vesicle Clustering and Release.” ELife 6 (November). https://doi.org/10.7554/eLife.29276. Yan, Jingqi, Morgan W Porch, Brenda Court-Vazquez, Michael V L Bennett, and R Suzanne Zukin. 2018. “Activation of Autophagy Rescues Synaptic and Cognitive Deficits in Fragile X Mice.” Proceedings of the National Academy of Sciences 115 (41): E9707–16. https://doi.org/10.1073/pnas.1808247115. Yoshimura, Jun, Kazuki Ichikawa, Massa J. Shoura, Karen L. Artiles, Idan Gabdank, Lamia Wahba, Cheryl L. Smith, et al. 2019. “Recompleting the Caenorhabditis Elegans Genome.” Genome Research 29 (6): 1009–22. https://doi.org/10.1101/gr.244830.118. Zhang, Liangyu, Jordan D Ward, Ze Cheng, and Abby F Dernburg. 2015. “The Auxin-Inducible Degradation (AID) System Enables Versatile Conditional Protein Depletion in C. Elegans.”   59  Development 142: 4374–84. https://doi.org/10.1242/dev.129635. Zhou, W., N. Wang, C. Yang, X. M. Li, Z. Q. Zhou, and J. J. Yang. 2014. “Ketamine-Induced Antidepressant Effects Are Associated with AMPA Receptors-Mediated Upregulation of MTOR and BDNF in Rat Hippocampus and Prefrontal Cortex.” European Psychiatry 29 (7): 419–23. https://doi.org/10.1016/j.eurpsy.2013.10.005.  

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            data-media="{[{embed.selectedMedia}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
https://iiif.library.ubc.ca/presentation/dsp.24.1-0390272/manifest

Comment

Related Items