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Characterization of embryonic muscle migration in Caenorhabditis elegans and of the role of unc-54 and… Viveiros, Ryan 2013

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   Characterization of embryonic muscle migration in Caenorhabditis elegans and of the role of unc-54 and mem-1 in inhibiting ectopic muscle membrane extensions.    by   Ryan Viveiros B.Sc., The University of British Columbia, 2003 M.Sc, The University of British Columbia, 2008    A THESIS SUMITTED IN PARTIAL FULFILLMENT OF  THE REQUIREMENTS FOR THE DEGREE OF   DOCTOR OF PHILOSOPHY  in   The Faculty of Graduate and Postdoctoral Studies   (Cell and Developmental Biology)   THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)   October 2013   ?Ryan Viveiros, 2013        ii Abstract  C. elegans body wall muscle is formed after a series of well-orchestrated steps. This thesis describes the plasma membrane dynamics of these migrating embryonic cells and the identification of two genes unc-54/MHB and mem-1 that appear to be involved negatively regulating post-embryonic muscle membrane extensions. During the characterization of embryonic muscle morphogenesis, whereby two rows of cells split and migrate dorsally or ventrally to form the final four muscle quadrants present upon hatching, I observed an anterior migration event, whereby the anterior-most pair of cells in each of the four muscle quadrants extends long processes to the anterior tip of the developing embryo. The anterior-most muscle cells then follow these extensions into their final positions in the developing embryo. Using RNAi and mutant analysis, I have identified laminin as being involved in mediating the dorsal-ventral muscle migrations and that the ?-integrin INA-1, the ephrin VAB-2 and its receptor VAB-1 and the Robo receptor SAX-3 indirectly promote the proper extension of the ventral anterior muscle processes by organizing the embryonic neurons so as to provide a clear path for muscle membrane extension. Post embryonically, the loss of either unc-54 (a myosin heavy chain B), or mem-1 (a WD repeat domain protein) results in ectopic membrane extensions from the mature muscle cells. These extensions can be rescued via targeted depletion of actin remodeling or cell adhesion complex components. During the analysis of these mutations I identified a predisposition for generating these ectopic membrane extensions that is conferred by using ectopic expression of PAT-3/?-integrin that is bound to GFP   iii and that using the PAT-3/?-integrin transmembrane domain to localize GFP to the plasma membrane is sufficient to generate this sensitivity.   iv Preface  Chapter 1. Portions of the introductory text are modified from previously written introductory material from my masters thesis intitled ?An Investigation Into the Genes Mediating Myoblast Migration in the Nematode Caenorhabditis elegans? (2008) completed at the University of British Columbia.  Chapter 2. A version of this material has been published as Viveiros R, Hutter H, Moerman DG (2011) Membrane extensions are associated with proper anterior migration of muscle cells during Caenorhabditis elegans embryogenesis. Dev Biol. 2011 Oct 1;358(1):189-200. Harald Hutter (Simon Fraser University, Burnaby, Canada) performed the experiments that generated the supplementary videos and was responsible for initial list of basement membrane components and interactors. I performed all additional experiments. Don Moerman and I conceived the experiments and I wrote the manuscript for the published paper.   Chapter 3. This chapter contains unpublished material. Ralf Schnabel (Technische Universit?t Braunschweig, Braunschweig) was responsible for the isolation of the GE6583 strain. Robert Barstead (Oklahoma Medical Research Foundation, Oklahoma City, USA) performed the whole genome sequencing of GE6583. Jon Taylor (University of British Columbia, Vancouver, Canada) helped perform the SNP-Chip mapping. Stephane Flibotte (University of British Columbia, Vancouver, Canada) performed the analysis of both the whole   v genome sequencing and SNP-Chip data. I performed all additional experiments. Don Moerman and I conceived the experiments.   vi Table of Contents  Abstract................................................................................................................ ii	 ?Preface ................................................................................................................ iv	 ?Table of Contents............................................................................................... vi	 ?List of Tables.................................................................................................... viii	 ?List of Figures .................................................................................................... ix	 ?List of Abbreviations ......................................................................................... xi	 ?Acknowledgements ......................................................................................... xiii	 ?1 Introduction ...................................................................................................... 1	 ?1.1	 ? Caenorhabditis elegans as a model organism................................................ 1	 ?1.2	 ? Membrane extension and adhesion ................................................................. 2	 ?1.3	 ? C. elegans body wall muscle ............................................................................ 6	 ?1.4	 ? Embryonic striated muscle migration.............................................................. 7	 ?1.5	 ? C. elegans muscle arms .................................................................................. 11	 ?1.6	 ? Thesis objectives ............................................................................................. 15	 ?2 Membrane Extensions are Associated with Proper Anterior Migration of Muscle Cells During Caenorhabditis elegans Embryogenesis .................... 17	 ?2.1	 ? Synopsis ........................................................................................................... 17	 ?2.2	 ? Introduction ...................................................................................................... 18	 ?2.3	 ? Materials and methods .................................................................................... 20	 ?2.3.1	 ? C. elegans strains ....................................................................................... 20	 ?2.3.2	 ? Molecular biology ........................................................................................ 21	 ?2.3.3	 ? Microscopy .................................................................................................. 21	 ?2.4	 ? Results .............................................................................................................. 22	 ?2.4.1	 ? A detailed description of C. elegans embryonic muscle migration.............. 22	 ?2.4.2	 ? Proper embryonic muscle migration in C. elegans requires laminin ........... 27	 ?2.4.3	 ? Proper ventral muscle process extension requires the ? -integrin INA-1, the ephrin VAB-2, and the ROBO receptor SAX-3........................................................ 30	 ?2.4.4	 ? Improper ventral muscle process extension is not due to defects in ventral hypodermal morphogenesis.................................................................................... 35	 ?2.4.5	 ? SAX-3 is required in the nervous system for proper extension of ventral anterior muscle processes ...................................................................................... 37	 ?2.4.6	 ? Mislocalized neurons are associated with ventral anterior muscle defects. 39	 ?2.5	 ? Discussion ........................................................................................................ 41	 ?2.5.1	 ? C. elegans muscle as a model to study muscle cell migration in vivo......... 41	 ?2.5.2	 ? Embryonic muscle migrations in C. elegans require laminin....................... 42	 ?2.5.3	 ? Multiple signaling pathways affect ventral anterior muscle processes........ 45	 ?2.5.4	 ? Anterior muscle processes are required for anterior migration ................... 47    vii 3 UNC-54 and MEM-1 Work in Concert to Inhibit Ectopic Membrane Extensions Away From the Nerve Cords in C. elegans Body Wall Muscle. 51	 ?3.1	 ? Synopsis ........................................................................................................... 51	 ?3.2	 ? Introduction ...................................................................................................... 52	 ?3.3	 ? Materials and methods .................................................................................... 54	 ?3.3.1	 ? C. elegans strains ....................................................................................... 54	 ?3.3.2	 ? RNAi............................................................................................................ 54	 ?3.3.3	 ? Microscopy .................................................................................................. 55	 ?3.3.4	 ? Microinjection procedure ............................................................................. 55	 ?3.3.5	 ? Isolation of temperature sensitive mutants using EMS mutagenesis.......... 56	 ?3.3.6	 ? Sequencing of GE6583 and identification of candidate mutations.............. 56	 ?3.3.7	 ? Mapping the GE6583 causative allele......................................................... 57	 ?3.4	 ? Results .............................................................................................................. 58	 ?3.4.1 	 ? Identification of t3197, a new temperature sensitive allele of the C. elegans myosin, unc-54, that results in ectopic muscle membrane extensions. .................. 58	 ?3.4.2	 ? The genes, unc-54 and mem-1 are both involved in mediating the ectopic muscle membrane extension phenotype. ............................................................... 59	 ?3.4.3	 ? Loss of mem-1 has no effect on muscle arm extension.............................. 64	 ?3.4.4	 ? GFP or YFP bound to the membrane via the ?-integrin transmembrane domain increases ectopic membrane extensions. .................................................. 66	 ?3.4.5	 ? RNAi depletion of actin remodeling or cell adhesion components reduces the severity of the ectopic membrane extensions. .................................................. 68	 ?3.4.6	 ? The unc-54(t3197) mem-1(t3198) double mutants exhibit neuronal defects. . .................................................................................................................................70	 ?3.5	 ? Discussion ........................................................................................................ 72	 ?3.5.1	 ? Both unc-54 and mem-1 are required to prevent ectopic membrane extensions ............................................................................................................... 72	 ?3.5.2	 ? Depletion of components involved in actin remodeling or cell adhesion inhibit ectopic membrane extensions. ..................................................................... 76	 ?3.5.3	 ? AVM neurons display pathfinding defects in the unc-54(t3197) mem-1(t3198) double mutants. ........................................................................................ 77	 ?3.5.4	 ? Expression of a PAT-3/?-integrin transmembrane domain::GFP fusion protein predisposes muscle cells for ectopic membrane extension defects ........... 77	 ?3.5.5	 ? Summary..................................................................................................... 80	 ?4.	 ? Conclusion.................................................................................................. 81	 ?4.1	 ? A novel anterior muscle migration event is necessary for proper embryonic muscle morphogenesis in C. elegans. ................................................ 81	 ?4.2	 ? Proper neuron organization is crucial for anterior muscle migration. ....... 82	 ?4.3	 ? Embryonic muscle migration in C. elegans is a laminin dependent, but integrin independent process.................................................................................. 83	 ?4.4	 ? UNC-54/MHCB and MEM-1 in inhibit ectopic muscle membrane extensions.  . ...........................................................................................................................83	 ?4.5	 ? Membrane-bound GFP or YFP predisposes muscle cells to ectopic membrane extensions. ............................................................................................. 84	 ?4.6	 ? Future directions.............................................................................................. 84	 ?References......................................................................................................... 87	 ?Appendices...................................................................................................... 103	 ?Appendix A: Chapter 2 supplementary data ........................................................ 103	 ?Appendix B: Chapter 3 supplementary data ........................................................ 123	 ?  viii  List of Tables  Table 2.1 Percent of 1.5 ? 2 fold embryos with defects in ventral anterior muscle process extension. .............................................................................. 32 Table 3.1: Rescue of ectopic membrane extension by unc-54 or mem-1.... 65 Table 3.2: Analysis of muscle arm extensions in the unc-54(t3197) mem-1(t3198) double mutant..................................................................................... 66 Table 3.3: Ectopic membrane extension defects can be rescued by targeted depletion of actin remodeling and cell adhesion components. ................... 71 Table A.1: Results of the RNAi feeding screen and mutational analysis .. 104 Table B.1: RNAi targets that failed to rescue ectopic membrane extensions in the unc-54(t3187) mem-1(t3198) double mutant. ..................................... 124   ix List of Figures  Figure 1.1: Organization of C. elegans body wall muscle and muscle arm membrane extensions. ....................................................................................... 9 Figure 1.2: C. elegans embryonic muscle migrations. .................................. 12 Figure 2.1: Schematic of the dorsal and ventral embryonic muscle migration............................................................................................................ 23 Figure 2.2: Migrating embryonic muscle cells extend lamellipodia and filopodia. ............................................................................................................ 24 Figure 2.3: Muscle cells extend membrane processes to the anterior of the embryo and maintain extensive contact between dorsally and ventrally migrating muscle cells. .................................................................................... 26 Figure 2.4: Distribution of perlecan coincides with the terminal position of the migrating muscle cells and is distributed beneath the extending anterior processes. ........................................................................................... 28 Figure 2.5: Depletion of laminin by RNAi results in defects in dorsal and ventral muscle migration and laminin along with vab-2, ina-1, and sax-3 is required for proper extension of the ventral anterior muscle processes. .. 31 Figure 2.6: The anterior-most ventral muscle cells in laminin(RNAi) treated embryos are displaced to the posterior.......................................................... 33 Figure 2.7: Anterior muscle processes extend during of hypodermal anterior enclosure and converge with the hypodermal leading edge at the sensory depression. ......................................................................................... 38 Figure 2.8: Displaced neurons are associated with improper extension of the ventral anterior muscle processes. .......................................................... 40 Figure 3.1: Ectopic membrane extensions in the unc-54(t3197) mem-1(t3198) double mutant strain.......................................................................... 60 Figure 3.2: The t3197 temperature sensitive mutation resides in the myosin head domain UNC-54/MHCB. ........................................................................... 61 Figure 3.3: Comparison of MEM-1 protein sequence with WDR6. ............... 64 Figure 3.4: Loss of unc-54 results in disruption of the muscle adhesion complexes in C. elegans body wall muscle. .................................................. 69 Figure 3.5: PAT-3/?-integrin based membrane bound fluorophore markers sensitize muscle cells for the production of ectopic membrane extensions............................................................................................................................. 70 Figure 3.6: AVM guidance defects in the unc-54(t3197) mem-1(t3198) double mutant. ............................................................................................................... 73   x Figure A.1. Loss of laminin results in a disorganized embryo with mislocalized muscle. ...................................................................................... 103 Figure B.1: SNP mapping of the causative allele of GE6583 indicates it is located on the right arm of chromosome 1 .................................................. 123   xi List of Abbreviations  ~  Approximately ?  Degrees %  Percent 4-D  Four-dimensional BOC  Brother of CDO C  Celsius CAM  Cell adhesion molecule  CAPS  Capricious CDO  CAM-related/down-regulated by oncogenes DNA  Deoxyribonucleic acid DEPC  Diethylpyrocarbonate dsRNA Double stranded RNA ECM  Extracellular matrix  EDTA  Ethylenediaminetetraacetic acid EME  Ectopic membrane extensions  F1  First generation progeny F2  Second generation progeny FGF  Fibroblast growth factor GFP  Green fluorescent protein hr  Hour IPTG  Isopropyl-beta-D-thiogalactopyranoside kb  kilobase  KCl  Potassium Chloride L  Lennox L1  First larval stage min  Minute  M9   Minimal media salt solution 9  MgCl2  Magnisium dichloride ml  Milliliter mM  Millimolar MRF  Myogenic Regulatory Factor  NaCl  Sodium Chloride ng  Nanogram  NGM  nematode growth medium  NMJ  Neuromusclular junction  NP-40  Nonyl phenoxypolyethoxylethanol Po  Parental generation PCR  Polymerase chain reaction RNA  Ribonucleic Acid RNAi  RNA interference SF/HGF Scatter factor/hepatocyte growth factor  SNP  Single nucleotide polymorphism  Tris-HCl Tris(hydroxymethyl)aminomethane hydrogen chloride    xii ?g  Microgram ?m  Micrometer ?l  Microliter x  Times  YFP  Yellow fluorescent protein   xiii Acknowledgements  Many strains used in this work were provided by the Caenorhabditis Genetics Center, supported by the National Center for Research Resources of the National Institutes of Health (USA). I would also like to thank Michel Labouesse (University of Strasbourg, Strasbourg, France) for a number of helpful strains. This research was supported by grants from the National Science and Engineering Research Council and the Canadian Institutes of Health Research to DGM (University of British Columbia. RV(University of British Columbia) is a research trainee of the Michael Smith Foundation for Health Research. H.H. (Simon Fraser University) is a senior scholar of the Michael Smith Research Foundation and funded by grants from the National Science and Engineering Research Council and the Canadian Institutes of Health Research. R.J.B. was supported by the United States National Institutes of Health (P41-HG-003652). DGM also received support as a Fellow of the Canadian Institute for Advanced Research. I would like to acknowledge the guidance and input from my committee  members: Guy Tanentzapf, Calvin Roskelley (University of British Columbia, Vancouver, Canada) and Harald Hutter (Simon Fraser University, Burnaby, Canada). I would also thank my collaborators Harald Hutter (Simon Fraser University, Burnaby, Canada), Robert Barstead (Oklahoma Medical Research Foundation, Oklahoma City, USA) and Ralf Schnabel (Technische Universit?t Braunschweig, Braunschweig), Germany) for their input and aid with this research. I would like to acknowledge the help from everyone in the Moerman lab   xiv and lastly I would like to acknowledge my supervisor Don Moerman for his mentorship and guidance in helping me to see this research to completion.  1 1 Introduction 1.1 Caenorhabditis elegans as a model organism  The regulation of cell protrusions plays a crucial role in human health and development. Cell migration, cell invasion, neurite outgrowth, epiboly and ingression all depend on the proper membrane extension and retraction (Pollard and Borisy, 2003; Mattila and Lappalainen, 2008). Defects in these processes have been linked to a number of human diseases, most notably cancer. A key stage in cancer progression is metastasis, whereby tumor cells delaminate, invade the surrounding healthy tissue and migrate away from the primary tumor to other tissues, where they can form new tumors, severely decreasing patient survivability. As number of genes involved in cell migration and cell invasion have already been linked to metastasis (Sahai, 2007; Nurnberg et al., 2011), further study of the processes may lead to the identification of new drug treatment targets, or of predictive markers of post tumor formation disease progression. C. elegans has proven to be an excellent model organism to answer basic questions concerning how cells move and change shape.  C. elegans is a free living species of nematode, a common soil roundworm found around the globe. Its ease of cultivation, large brood sizes, short generational time and ability to propagate by mating or self-fertilization make it an excellent genetic model to study animal behavior and development (Brenner, 1973; Brenner, 1974) . It was the first multicellular organism to have its genome    2 sequenced and approximately 38% of C. elegans genes are predicted to have human homologues (Shaye and Greenwald, 2011). The adult C. elegans hermaphrodite consist of 959 somatic cells and the transparent nature of both it and its eggs allowed for the determination of its complete embryonic and post-embryonic cell lineage (Sulston and Horvitz, 1977; Sulston et al., 1983). This transparency also allows for the labeling of the animals internal tissues or cells with fluorescent markers, facilitating the observation of biological processes, such as cell migration, in vivo and in real time. The invariant nature of the C. elegans cell lineage and the ease of labeling and observing its various cell types in vivo, has facilitated the identification and characterization of numerous proteins involved in cell migration (For example see Zallen et al., 1998; Sherwood et al., 2005; Rasmussen et al., 2008; Chihara and Nance, 2012; Marcus-Gueret et al., 2012). Thanks to studies such as these and others in different model systems, we are beginning to piece together the grand picture of how cells use their membrane to migrate and change shape. 1.2 Membrane extension and adhesion Cell membrane extensions occur in a complex and tightly regulated multistep process, beginning with a cell becoming polarized in response to extracellular signals. Actin polymerization at the leading edge of the migrating cell results in the extension of flat membrane protrusions called lamellipodia and finger-like protrusions known as filopodia. These membrane extensions form adhesions with the underlying extracellular matrix to anchor and stabilize the protrusions.    3 In brief, remodeling of the cortical actin cytoskeleton in response to extracellular cues is mediated by the by the Rho family of GTPases, which includes RhoA, Rac1 and Cdc42 (Heasman and Ridley, 2008) . Rac1 and Cdc42 stimulate the polymerization of actin by activating specific nucleation promoting factors (NPFs) such as the family of Wiskott-Aldrich syndrome proteins, which includes N-WASP, WASP, WAVE and WASH (Symons et al., 1996; Miki et al., 1998b; Miki et al., 1998a; Liu et al., 2009). These NPFs in turn interact with the actin-related protein 2/3 (Arp2/3) complex, which then binds to the tip or side of existing actin filaments and serves as nucleation side for the polymerization of actin side branches (Reviewed in Pollard and Borisy, 2003). This branching is crucial for the formation of lamellipodia, but is nonessential for filopodia formation, since filopodia lack branched actin and rely on other NPFs such as formins for their actin nucleation (Reviewed in Mattila and Lappalainen, 2008; Yang and Svitkina, 2011). Interestingly, the Arp2/3 complex appears to be dispensable for certain types of cell migration. Cells lacking the Arp2/3 complex, though unable to respond to ECM cues, have been shown to be able to perform chemotaxis by using filopodia-based migration (Wu et al., 2012).  A number of other actin-binding proteins are able to regulate the speed and organization of actin assembly in membrane extension by affecting the pool of available actin monomers and free ends (Reviewed in Welch and Mullins, 2002; Pollard and Borisy, 2003; Ridley et al., 2003; Le Clainche and Carlier, 2008). These include, profilin, which prevents the self-nucleation of actin by binding to monomers and also targets monomers to barbed ends and capping    4 proteins that terminate filament elongation and thereby restrict polymerization to nascent filaments close to the plasma membrane (Blanchoin et al., 2000; Pantaloni et al., 2000). The ADF/cofilin family, which severs filaments and promotes actin dissociation from the pointed end, frees up actin monomers from old filaments so they can be incorporated into the nascent filaments forming at the leading edge (Ichetovkin et al., 2002). Other proteins still help to support the actin network, such as cortactin that acts to stabilize branches and filamin A and a-actinin help to stabilize the whole actin network by cross-linking filaments (Xu et al., 1998; Flanagan et al., 2001; Weaver et al., 2001).  As the membrane extends outwards, adhesion complexes are formed attaching the protrusions to the surrounding extracellular environment. These adhesions are thought to act as a ?molecular clutch?, such that when adhesion complexes are adhering to the actin network, the force generated by actin polymerization drives membrane extension at the leading edge and when the adhesion complex disengage, the slippage that occurs between the actin network and adhesions results in retrograde flow and decrease protrusion rate (Lin et al., 1994; Suter and Forscher, 1998; Brown et al., 2006; Hu et al., 2007). When integrin complexes in lamellipodia first come in contact with the ECM, interactions take place with talin and kindlin which serve to enhance integrin activation and leads to stabilization of its grip to the matrix (Reviewed in Wolfenson et al., 2013). Assembly of these nascent adhesion complexes occurs quickly, involving only a small number of integrin molecules and leads to actin polymerization (Yu et al., 2011). Subsequent myosin mediated tension put on    5 these nascent complexes serves to reinforce their strength and facilitates their maturation into true focal adhesions (Roca-Cusachs et al., 2009). This maturation is thought to be due to myosin generated force inducing conformational changes in key adhesion components, which exposes binding or activation sites. One such protein is talin, which serves as a linker between the integrin complex and actin, binds vinculin when mechanically stretched (del Rio et al., 2009) and is required to translate non-muscle myosin II-generated forces to the substrate (Jiang et al., 2003). Integrins themselves are also activated by tension (Friedland et al., 2009) and collectively this activation serves to stabilize the nascent adhesions. In C. elegans muscle dense bodies are analogous to mammalian adhesion complexes and share many of the same proteins. In dense bodies, the PAT-2/PAT-3/?/? integrin heterodimer binds the ECM protein UNC-52/Perlecan (Rogalski et al., 1993; Hresko et al., 1994; Gettner et al., 1995). Additional proteins are recruited via two parallel but dependent pathways, which culminate in the binding of DEB-1/vinculin to actin filaments (Moerman and Williams, 2006). One fork of this pathway involves two proteins, UNC-112/MIG-2/Kindlin and the integrin-linked kinase (ILK) homolog PAT-4. These two cytoskeletal adapter proteins are required in concert for both to properly associate with the nascent dense body structure (Rogalski et al., 2000; Mackinnon et al., 2002) and both are required for the subsequent recruitment of the third member of the complex, PAT-6/actopaxin (Lin et al., 2003). The second pathway involves the recruitment of the LIM domain protein UNC-95 and DEB-1/vinculin, both of which require the    6 other for proper localization to the dense body (Barstead et al., 1991; Broday et al., 2004). Both of these pathways are absolutely essential for proper attachment of the dense body adhesions to actin.  Though integrin adhesions are essential for the stabilization of membrane extensions and thus for cell migration, my own work, as detailed in chapter 2 has revealed that integrin based adhesions do not appear to be involved in mediating the embryonic migrations of C. elegans embryonic muscle cells (Viveiros et al., 2011). Laminin is required for proper migration of these cells and this suggests that C. elegans muscle migration occurs through adhesion dependent, but integrin independent migration, making C. elegans body wall muscle an interesting model for the study of cell migration.  1.3 C. elegans body wall muscle C. elegans body wall muscle consists of 95, mono-nucleated, striated muscle cells arranged symmetrically in four quadrants, a ventral right, ventral left and a dorsal left and right quadrant (Figure 1.1) (Sulston and Horvitz, 1977). Of these muscle cells, 81 are derived embryonically, with the remainder arising from a posterior mesoblast cell during larval development, as the mature muscle cells do not divide (Sulston and Horvitz, 1977; Sulston et al., 1983). As in vertebrates, myogenic fate of striated muscles in C. elegans is specified by a group of basic helix-loop-helix proteins known as the myogenic regulatory factors (MRFs). In mice, four factors are required MyoD, Myf5, Myogenin and MRF4 (Ott et al., 1991; Pownall and Emerson, 1992; Sassoon,    7 1993; Kassar-Duchossoy L, 2004). In C. elegans, there is only one MRF, HLH-1, the C. elegans MyoD homologue (Chen et al., 1994), but muscle fate also depends on a pair of other myogenic factors, UNC-120/SRF and HND-1/HAND (Fukushige et al., 2006). Interestingly, these genes play prominent roles in regulating smooth and cardiac muscle development in mammals (Cserjesi et al., 1995; Niu et al., 2005). This has lead to the postulation that all major muscle types, skeletal, smooth and cardiac, may have arisen from a common ancestral cell type (Baugh and Hunter, 2006; Fukushige et al., 2006). After acquiring their cell fate, C. elegans muscle cells must migrate to organize into the body wall muscle quadrants present upon hatching.   1.4 Embryonic striated muscle migration. In vertebrates, the majority of skeletal muscle is derived from progenitors present in the somites, which arise from segmentation of paraxial mesoderm present to the sides of the notochord. Environmental signals cause the specification of myogenic and dermal progenitors in the dorsal somite forming a structure known as the dermomyotome. As myogenic fate is determined, the myogenic cells delaminate and migrate from the dermomyotome to the myotome, where they fuse to form myotubes. Delaminated myogenic cell can also migrate from the dermomyotome directly to sites of limb bud formation (Kalcheim C, 1999; Buckingham, 2001; Christ and Brand-Saberi, 2002; Bryson-Richardson and Currie, 2008).     8 Establishment of the muscle progenitor pool requires the Pax3 transcription factor, and loss of Pax3 expression results in the loss of all hypaxial muscle (Bober et al., 1994; Tajbakhsh et al., 1997), while their proliferation depends on the expression of the c-myc proliferating factor the Six1-Eya1-Dach transcriptional activation complex (Li et al., 2003). Their delamination from the dermomyotome requires the tyrosine kinase receptor c-Met and its ligand, scatter factor/hepatocyte growth factor (SF/HGF) and in c-Met or SF/HGF mutant mice there is a complete lack of limb muscles (Bladt et al., 1995; Dietrich et al., 1999).  After delamination, actively migrating muscle precursors destined for the limb buds express the homeodomain transcription factor Lbx1. Lbx1 is required for the migration of limb muscle progenitors, but is dispensable for other muscle precursor populations remain unaffected (Schafer and Braun, 1999; Gross et al., 2000), suggesting the different populations of migrating muscle progenitors must navigate and respond to different environmental cues to reach their respective sites of muscle formation. Only once these muscle precursors reach their final destination do they proceed to proliferate and finally acquire their terminal cell fates though expression of the MRFs (Buckingham, 2001; Bryson-Richardson and Currie, 2008).     9  Figure 1.1: Organization of C. elegans body wall muscle and muscle arm membrane extensions. Panel A is a cartoon of a left lateral view of C. elegans body wall muscle with the dorsal and ventral muscle quadrants of the animal shown. The rectangles illustrate the locations of the cross sections depicted in panels B and C. Panel B is a cartoon of a cross section through the animal illustrating the relative position of the 2 dorsal and 2 ventral muscle quadrants. The small circles between the quadrants represent the dorsal and ventral nerve cords that the muscle cells extend membrane processes towards. Dorsal is to the top and ventral is to the bottom. Panel C is a cartoon depicting a ventral view of a subset of ventral muscle cells extending muscle arms to the ventral nerve cord. The muscle cells more lateral to nerve cord must extend their muscle arms across their more proximal neighbours, and thus have longer membrane processes.     10 A number of cell migration and guidance factors have been implicated in the migration of these muscle progenitor cells, one of the first being laminin, which promotes myoblast migration in vitro (Foster et al., 1987; Goodman et al., 1989).  Studies in mice have shown that loss of certain cadherins, CDO (CAM-related/down-regulated by oncogenes), BOC (brother of CDO) and neogenin result in minor defects in muscle precursor migration, though these genes appear to play a larger role in promoting muscle cell fate (Krauss et al., 2005). To date, the only guidance proteins that have been implicated in playing an major role in these migrations are the chemokine receptor CXCR4, its ligand SDF-1 and Gab1, which is an adaptor protein that transduces signals elicited by tyrosine kinase receptors, (Vasyutina et al., 2005; Chong et al., 2007; Rehimi et al., 2010) and their loss does not completely disrupt myoblast migrations, indicating that there are still other proteins involved in the process yet to be identified. One approach to identify new components involved in these migrations in vertebrates is to screen for genes using a simpler model organism. Compared to the complexity observed in vertebrate myoblast migration, C. elegans embryonic muscle migration involves only 81 muscle cells, which arise from four of the six so-called ?founder cell? lineages, 32 from the C lineage, 28 from the MS lineage, 20 from the D, the only muscle exclusive cell lineage, and 1 from the AB lineage (Sulston et al., 1983).  The newly formed muscle cells initially are arranged in two clusters at the midline on the left and right sides of the embryo, lying beneath the hypodermal seam cells (Sulston et al., 1983). Starting at ~290 min of embryogenesis cells in these two populations split to migrate dorsally, or    11 ventrally to lie beneath the hypodermal cells and form the final four quadrants present upon hatching (Figure 1.2) (Sulston et al., 1983; Hresko et al., 1994; Viveiros et al., 2011). The muscle cells? decision to migrate dorsally or ventrally appears to be lineage dependent (Sulston et al., 1983), but as in higher eukaryotes, the guidance cues regulating these migrations are currently unknown. Interestingly, these migrations occur while some muscle cells are still dividing and in a dynamic environment where broad stages of C. elegans embryonic muscle cell migration have been characterized, the genes mediating it are still unknown. As well as providing an excellent model for the study embryonic migrations, C. elegans muscle is also a great model for the study of post-embryonic migratory behavior, as mature muscle cells extend specialized membrane processes to the motor neurons, which are necessary for proper muscle innervation.  1.5 C. elegans muscle arms   Muscle contraction required the establishment of neuromuscular junctions (NJMs) between motor neurons and muscle. For contact between these cell types to occur, motor neuron axons must migrate to find their correct post-synaptic targets and this recognition has been shown to involve the extension of filopodia from both the pre-synaptic axon and the post-synaptic muscle cells (Ritzenthaler et al., 2000; Ritzenthaler and Chiba, 2003). While initially identified in Drosophila, these post-synaptic muscle filopodia, often referred to as myopodia, have also been found to be extended in mammalian NMJ formation    12 and appear a widespread feature of NMJ formation (Uhm et al., 2001; Misgeld et al., 2002). These myopodia have been shown to cluster near approaching axons in response to neuronally secreted agrin (Uhm et al., 2001). Agrin signaling appears to act through p120 catenin, a cytoplasmic component of the cadherins adhesion complex. Agrin exposure leads to the dissociation of p120 catenin from the cadherins complex and ectopic expression of p120 catenin in cultured muscle    Figure 1.2: C. elegans embryonic muscle migrations. Panel A is a cartoon depicting a cross-section of an ~5hr embryos with muscle cells adjacent to the hypodermal seam cells (Based on observation from Hresko et al., 1994; Viveiros et al., 2011). Arrows indicate the direction of migration of the muscle cells in the two clusters to form the final four muscle quadrants. B-E are anti-CeMyoD antibody staining of embryos undergoing muscle cell migration (D. Moerman, unpublished). Panels are of increasingly older embryos and show the dorsal and ventral migrations that generate the final four muscle quadrants.    13 cells was able to generate myopodia in the absence of agrin (Madhavan et al., 2006). Myopodal recognition of its pre-synaptic target also plays a role in NMJ formation. In Drosophila, Capricious (CAPS), a transmembrane protein with leucine-rich repeats (LRRs) that is expressed in a subset of muscle cells, clusters at the tips of myopodia and in the absence of CAPS there is a reduction in contact between the pre and post-synaptic surfaces and a delay in the establishment of NMJs (Kohsaka and Nose, 2009). As CAPS is only expressed in a subset of muscle cells (Shishido et al., 1998), it appears that the factors involved in post-synaptic recognition of neurons are expressed in a cell specific manner. In C. elegans, motor neuron axons do not completely extend to the body wall muscle. For proper innervation to occur, the muscles cells must extend specialized membrane extensions, called muscle arms, to make contact with the neurons (Stretton, 1976; White et al., 1986; Dixon and Roy, 2005). Animals hatch with one or two of these processes, which are believed to arise passively during embryogenesis through adjacent attachment between the muscle cells and neurons followed by separation of the tissues during morphogenesis (Dixon and Roy, 2005). Following hatching the larval pass through 4 larval stages before becoming adults and the number of these muscle arms roughly doubles between the first and third larval stages. In contrast to the embryonically derived muscle arms, post-embryonic muscle arm development is an active process requiring cytoskeletal remodeling (Dixon and Roy, 2005). It has been know for quite some time that the proper extension of post-embryonic muscle cells requires UNC-   14 104/kinesin-mediated anterograde axonal transport of synaptic vesicles (Hall and Hedgecock, 1991) suggesting a neuronally derived muscle arm attractant. More recently, the details of the signaling molecules involved in regulating muscle arm extension have been elucidated. MADD-4, an ADAMTS-like protein, has been identified as a neuronally expressed attractant that signals through UNC-40/DCC to induce the extension of muscle arms (Alexander et al., 2009; Seetharaman et al., 2011). Mutations in daf-2 insulin-like and the TGF-B signaling pathways have been shown to cause an increase in the number of muscle arms (Dixon et al., 2008). These mutations appear to phenocopy what occurs during wild type dauer formation. Dauer larvae, an alternative hardy larval stage that can survive in harsh environmental conditions (Cassada and Russell, 1975; Golden and Riddle, 1984), also display supernumerary muscle arms and it appears that increasing the number of muscle arms may be involved in some as of yet uncharacterized dauer behavior (Dixon et al., 2008).  C. elegans muscle has also been shown to be a good model for the characterization of ectopic membrane extensions (EMEs) and a number of genes have been identified as being involved in inhibiting EMEs. In muscle, FGF signaling has been shown to be involved in suppressing membrane extensions with the FGF receptor, ELG-15, and SEM-5/Grb2, an adapter protein that functions downstream of receptor tyrosine kinases, both functioning cell autonomously in muscle cells to negatively regulate membrane extensions (Dixon et al., 2006). Cell adhesion also plays an important role in negatively regulating these processes, as the loss of laminin or components of integrin    15 adhesion complexes also results in the generation of EMEs and hyperactivation of the FGF pathway is able to suppress these ectopic processes (Dixon et al., 2006). The easy detection of ectopic membrane extensions in muscle makes it a great model for the identification of regulators of membrane extension.  In my own work, as detailed in Chapter 3, I have identified two other genes that appear to be negatively regulating membrane extensions. The loss of either the myosin heavy chain B protein UNC-54 or the WD motif protein MEM-1 results in the formation of ectopic muscle membrane extensions, which can be rescued through the depletion of cell adhesion or actin remodeling proteins (see Chapter 3)  1.6 Thesis objectives   Muscle cell migration is an integral part of C. elegans embryogenesis. Though the cell movements involved in embryonic muscle development are well known, the genes involved in orchestrating these events are as yet undetermined. Using C. elegans as a model, I screened for genes involved in regulating muscle cell migration. In addition, as previous characterization of C. elegans muscle migration was based on nuclear position, I used a membrane bound marker to characterize the membrane dynamics of these migrating cells.   During C. elegans embryonic muscle migration I discovered an anterior migration event orchestrated by the anterior most muscle cells. These cells extend long membrane processes to the anterior-most tip of the developing embryo. These processes are necessary for the anterior migration of these cells    16 and their disruption results in a posterior displacement of these cells. This is the first observation of anterior migration of embryonic muscle cells and has increased our understanding of how the embryonic muscle quadrants are formed. Studying post-embryonic muscle, I identified two genes that when disrupted, result in the generation of ectopic membrane extensions in the muscle cells. These two genes appear to work synergistically. In Chapter 2 I describe my characterization of embryonic muscle migration and identification of genes involved in the process. In Chapter 3 I describe the identification and characterization of two genes, unc-54 and mem-1 (Y54E5B.2) that are involved in preventing ectopic muscle membrane extension. These two chapters provide insight into the embryonic and post-embryonic behaviour and regulation of C. elegans muscle membrane dynamics.      17 2 Membrane Extensions are Associated with Proper Anterior Migration of Muscle Cells During Caenorhabditis elegans Embryogenesis 2.1 Synopsis    C. elegans body wall muscle is formed after a series of well-orchestrated steps. With the onset of specification embryonic muscle cells accumulate under the hypodermal seam cells at the left and right sides of the embryo. Shortly thereafter they begin to migrate dorsally and ventrally resting beneath the dorsal and ventral hypodermis eventually forming the four muscle quadrants present upon hatching. In this study I describe the plasma membrane dynamics of these migrating cells and observe the extension of filopodia and lamellipodia during dorso-ventral migration. I also describe an anterior migration event during embryonic muscle morphogenesis, whereby the anterior-most pair of cells in each of the four muscle quadrants extends long processes to the anterior tip of the developing embryo. Anterior-most muscle cells then follow these extensions into their final positions in the developing embryo. Using RNAi and mutant analysis, I have identified laminin as being involved in mediating the dorsal-ventral muscle migrations. Finally, I show that the ?-integrin INA-1, the ephrin VAB-2 and its receptor VAB-1 and the Robo receptor SAX-3 indirectly promote the proper extension of the ventral anterior muscle processes by organizing the    18 embryonic neurons so as to provide a clear path for muscle membrane extension. 2.2 Introduction During embryogenesis, directed cell migrations play a major role in organizing developing tissues. As new cell types arise they frequently migrate away from their place of origin to form nascent tissues. This is the case during striated muscle development. While the details vary among species, striated muscle formation requires the migration of nascent myoblasts away from where they are specified to the site of muscle formation (Reviewed in Christ and Brand-Saberi, 2002; Schnorrer and Dickson, 2004). In the nematode Caenorhabditis elegans, embryonically derived body wall muscle cells undergo a series of discrete cell movements to generate the proper tissue arrangement. Muscle cells arise from four distinct founder cells, MS (28 cells), C (32 cells), AB (1cell) and D (20), and initially arrange into two quadrants on the left and right sides of the embryo, lying beneath the hypodermal seam cells (Sulston et al., 1983). Starting at ~290 min of embryogenesis these cells migrate dorsally or ventrally to lie beneath the dorsal and ventral hypodermal cells, forming the four body wall muscle quadrants present upon hatching (Figure 2.1) (Sulston et al., 1983; Hresko et al., 1994). These migrations are occurring while some of the muscle cells are still dividing and in a dynamic environment where cell-cell contacts can differ from embryo to embryo (Schnabel et al., 1997).  This begs the question as to how these cells unerringly reach their destinations in such a dynamic environment.    19 Extracellular matrix (ECM) and cell surface components are involved in cell sorting, positioning, migration and attachment. These molecules include fibronectin, laminin, basement membrane collagens, as well as the NCAM, cadherins, integrins, UNC-6/netrins and several identified growth factor signaling pathways, like Wnt and FGF (Ishii et al., 1992; Adams and Watt, 1993; Serafini et al., 1994; Locascio and Nieto, 2001; Keller, 2005; Heisenberg and Solnica-Krezel, 2008). However, with the notable exception of Wnt signaling (Reviewed in Hardin and King, 2008), the study of mutations in C. elegans homologs of many of these genes suggests they are not essential for cell migration in the early embryo. Mutations in many of these structural proteins have quite drastic effects on cell attachment, movement and migration in later development, but, for the most part, do not appear to affect earlier developmental processes (Hedgecock et al., 1987; Guo et al., 1991; Ishii et al., 1992; Leung-Hagesteijn et al., 1992; Rogalski et al., 1993; Sibley et al., 1993; Hresko et al., 1994; Williams and Waterston, 1994; Gettner et al., 1995; Huang et al., 2003). Recent global studies using RNAi methodology affirms these earlier observations (Sonnichsen et al., 2005; Schnabel et al., 2006). Apparently, we are still missing key genes involved in embryonic cell migration. Recent work by Tucker and Han (2008) showed that ina-1/?-integrin expression in the hypodermis and vab-1/vab-2/ephrin/ephrin receptor expression in the nervous system are required for the proper positioning of a subset of the anterior muscle cells, and also identified a novel metalloprotease, MNP-1, that is required for all proper embryonic muscle migrations to occur. Their study reveals    20 more about the mechanisms orchestrating these cell movements, but we still lack a complete picture of the process. Here I present a description of the plasma membrane dynamics of migrating embryonic muscle cells in C. elegans and show that nematode embryonic muscle migrations require the basement protein laminin. Over the course of these observations, I identified a previously undescribed muscle morphogenetic event, whereby the anterior-most cells in each quadrant extend membrane processes to the anterior of the embryo and then muscle cells follow these processes to the anterior end of the embryo. I also demonstrate that proper extension of the ventral, but not the dorsal, anterior processes is dependent upon laminin, ina-1/?-integrin, vab-2/vab-1/ephrin/ephrin receptor and sax-3/Robo.  These muscle projections are required for the anterior migration of the muscle quadrants during elongation, as loss of the ventral extensions results in posterior displacement of the ventral anterior muscle cells.  2.3 Materials and methods 2.3.1 C. elegans strains  Animals were maintained as described (Brenner, 1974). The N2 Bristol strain was used as reference wild type. The strains used in this study were: PD7963, ccIs7963[phlh-1::GFP]; VH95, hdIs61[phlh-1::memGFP, rol-6(su1006)]; ML1153 mcIs46[DLG-1::RFP;unc-119(+)]; RW10055, unc-119 (ed3); stIs10055 [cnd-1(3.2kb)::HIS24::mCherry + unc-119(+)]; VC2420, lam-1(ok3139)    21 IV/nT1[qIs51](IV;V); IC464, sax-3(ky123) X; quEx100[pajm-1::SAX-3 + odr-1::RFP]; IC459, sax-3(ky123) X; quEx102 [prgef-1::SAX-3 + odr-1::RFP]; VC1782, egl-15(ok2314) X; MT5383, lin-44(n1792) I; EW72, cwn-1(ok546) II; cwn-2(ok895) IV; NW1704, smp-1(ev709) I; jcIs IV; him-5(e1490) V; MT1215, egl-20(n585) IV; NW1701, mab-20(ev778) I; muIs32 II; him-5(e1490) V.    To generate VH95, transgenic animals containing the plasmid with the membrane-tagged GFP under the control of the hlh-1 promoter (see below) were generated as described (Mello and Fire, 1995). During this study it was found that the transgene appears to be stably transmitted to all progeny suggesting that the original extrachromosomal array has become integrated into the genome. 2.3.2 Molecular biology To generate a membrane-bound GFP tag expressed in body wall muscle, the signal sequence and transmembrane portion of PAT-3 (aa1-29 and aa720-770) were PCR-amplified and inserted in front of the coding region of GFP in a plasmid also containing the hlh-1 promoter (Chen et al., 1994).  Feeding RNAi was done as per Kamath et al (2003) and injection RNAi, was done at 200ng/ml and was performed as previously described (Gonczy et al., 2000; Sonnichsen et al., 2005). Injected adults were cultured for 24hrs before being used for further analysis. 2.3.3 Microscopy 4-D recordings were done as previously described (Moerman et al., 1996; Schnabel et al., 1997; Thomas and White, 1998; Burglin, 2000) with the addition    22 of fluorescent Z-stacks. For cell tracking I used Simi Biocell (Schnabel et al., 1997). Confocal microscope imaging and fluorescent recordings were performed on a Quorum WaveFX spinning disc system mounted on a Zeiss Axioplan II microscope and an inverted Zeiss Axiovert LSM 5 confocal microscope equipped with epifluorescence, Nomarski optics, and LSM 5 Pascal software. Image acquisition and analysis was done with the LSM 5 Pascal volume-rendering software and the Volocity software package. (www.improvision.com).  2.4 Results  2.4.1 A detailed description of C. elegans embryonic muscle migration.   To reveal the mechanisms of muscle cell movements I visualized the surface of migrating muscle cells by expressing a GFP tag localized to the membrane in muscle cells. This allowed me to detect cell contacts, and various extensions like filopodia or lamellipodia, which are indicative of an active migration process. As the hlh-1/ceMyoD promoter used to drive expression of the marker activates as cells acquire their myogenic fate (Chen et al., 1994), GFP is detectable before muscle cells begin to migrate.  As the cells divide, they form two rows on the left and right sides of the embryo, with MS derived cells the furthest anterior, the C derived cells the furthest posterior and the D derived cells in between the two (Figure 2.1). Left and right MS and D derived muscle are already separated early on while the left and right C derived cells maintain a high    23 degree of contact in the posterior (Figure 2.2). Though membrane connections exist between the posterior-most C derived cells (data not shown), the cell morphology of these early muscle cells appears to be devoid of visible lamellipodia or filopodia.    Figure 2.1: Schematic of the dorsal and ventral embryonic muscle migration. Panel A is a cartoon adapted from Hresko et al. (1994) of a cross-section of an embryo, showing the starting position of the muscle cells beneath the hypodermal seam cells and direction of their migrations. Panels B and C are ball models indicating the nuclear position of the left embryonic muscle cells. Panel B shows the cells before their dorsal and ventral migration and panel C shows the same cells and their descendants after they have migrated. Red spheres mark MS derived cells, blue the D derived and green the C linage derived muscle cells. In panels B and C anterior is to the left and ventral is to the bottom. Arrows indicate direction of migration.    24  Figure 2.2: Migrating embryonic muscle cells extend lamellipodia and filopodia. A fluorescent image showing the GFP tagged muscle membranes of the dorsal right (DR) and dorsal left (DL) muscle quadrants. The left and right C derived, posterior muscle cells can be seen clustered in the posterior. GFP appears to tag both the plasma membrane, as well as, some of the internal membranes of the cell. The triangles indicate leading edges of dorsally migrating cells and the arrows indicate membrane extension to the anterior of the embryo. Anterior is to the left of the panel. Scale bar is 10?m.   At ~ 290 minute stage of development, cells in the left and right muscle rows begin to migrate dorsally or ventrally (Sulston et al., 1983; Schnabel et al., 1997). Using the membrane marker I can clearly see that these cells have the morphology of migrating cells, with lamellipodia and filopodia at the leading edges (Figure 2.2). As described previously (Sulston et al., 1983; Moerman et al., 1996; Schnabel et al., 1997), these dorsal and ventral migrations initiate from the anterior to the posterior, with the MS derived cells preceding the D derived, followed by the C derived muscle cells, which now complete their left and right separation. Extensive cell contact is maintained between the dorsal and ventral    25 contralateral cells during migration. These contacts dissipate in an anterior to posterior fashion during elongation, with the contacts in the posterior C lineage cells persisting through the onset of embryo movement (Figure 2.3).  Interestingly, while the cells are migrating away from the lateral hypodermal seam cells, I have also observed instances of cells extending processes from the lagging edge back towards the lateral hypodermal seam cells, though these are eventually lost as embryogenesis progresses (Figure 2.3). At ~330 minute of development, as the cells in the left and right muscle rows migrate dorsally or ventrally, the anterior-most cells in each of the four developing body wall muscle quadrants extend membrane processes to the anterior tip of the embryo (Figs 2.2, 2.3). In all, eight cells extend these processes: MSapappp and MSapaaap from the dorsal left and MSppappp and MSppaaap from the dorsal right quadrants, and MSapapap and MSappapp from the ventral left quadrant and MSppapap and MSpppapp from the ventral right. The processes extend forward, curving along the contour of the embryo and converge near the point of the sensory depression. At their longest, these processes extend 16 ? 1.2?m, compared to the 2.2 ? 0.4?m membrane extensions generated by the dorsally and ventrally migrating cells. As elongation progresses, their cell bodies and the adjacent, more posterior muscle cells continue to advance to the anterior, decreasing the overall length of the membrane processes. Ultimately, this results in the loss of the processes completely from these maturing muscle cells.      26  Figure 2.3: Muscle cells extend membrane processes to the anterior of the embryo and maintain extensive contact between dorsally and ventrally migrating muscle cells. After contact between dorsal (D) and ventral (V) muscle cells is disrupted, cells can re-extend processes back towards the lateral hypodermal seam cells. Panels A, C, E are Nomarksi images depicting an elongating wild type embryo with images taken at 15-minute intervals. B, D, F are fluorescent images of the same embryos visualizing the membrane-bound GFP tagged muscle cells. G, H and I are enlargements of the indicated regions of panels B, D and F. Asterisks indicate anterior membrane extension. Arrows indicate points of contact between the dorsal and ventral muscle quadrants. Triangles indicate the extension of a ventral process from a dorsal cell. In all    27 panels anterior is to the left and the ventral side of the embryo is to the bottom. Scale bar is 10?m.  UNC-52/perlecan is a key component of the basement membrane that links the body wall muscle to the hypodermis. Though null alleles of unc-52 do not result in any defects in embryonic muscle migration, it is the first component required for the initiation of proper sarcomeric assembly (Rogalski et al., 1993; Hresko et al., 1994) and as such, I was interested to see how its deposition related to the migration of the muscle cells. Looking at the perlecan distribution in concert with the muscle membrane marker reveals that perlecan appears to be laid down only as the leading edges of the dorsally and ventrally migrating muscle cells reach their destination (Figure 2.4). In the case of the anterior muscle processes, perlecan appears to be laid down beneath them as they extend, because no part of the processes appear free of perlecan and none is apparent before their leading edges (Figure 2.4). These results would indicate that perlecan is more of a marker for the final position of the muscle cells, than a component of active muscle migration.  2.4.2 Proper embryonic muscle migration in C. elegans requires laminin  To identify genes required for proper embryonic body wall muscle migration, I performed a feeding RNAi screen focusing on 768 genes known or predicted to be components of, or receptors for, the extracellular matrix (ECM) (Appendix A: Supplementary Table 1). I focused on identifying knockdowns that    28  Figure 2.4: Distribution of perlecan coincides with the terminal position of the migrating muscle cells and is distributed beneath the extending anterior processes. All panels are fluorescent images of wild type embryos. A-C are images of a two-fold embryo. D-F are images of an earlier elongating embryo in the process of rotating clockwise from resting on its dorsal surface to its right side. A and D show muscle cells labeled with membrane bound GFP. Panels B and E are embryos stained with anti-UNC-52 antibodies. Panels C and F are overlays showing the coincidence of perlecan and muscle cells. In all panels anterior is to the left and ventral is down. Scale bar is 10?m.  effected the migrations that mediate the two row to four quadrant transition, since defects of the early embryo, i.e. improper gastrulation or non-muscle cell fate changes, can result in a non-specific disorganized muscle phenotype. As defects in the migrations that generate the final four muscle quadrants should result in embryonic or larval lethality, I focused on identifying those phenotypes, though I also examined non-lethal animals for displaced muscle using a muscle specific    29 GFP marker, since C. elegans displays a muscle plasticity that allows them to compensate for absent muscle cells (Moerman et al., 1996). Of all genes screened only depletion of laminin resulted in defects in the later stage embryonic muscle cell migration.  Laminin forms a heterotrimer consisting of an ?, ? and ? subunit. C. elegans has two ? subunits, one ? and one ? subunit, forming two laminin isoforms (Huang et al., 2003; Kao et al., 2006). Targeted depletion of any of the subunits using RNAi results in a number of embryonic muscle migration defects. Both the dorsal and ventral migrations are defective, indicated by lagging cells present between the dorsal and ventral quadrants, and there is an apparent loss of muscle cells from the anterior ventral quadrant (Figure 2.5, Table 2.1). While there are extensive defects in the migrations forming the dorsal and ventral muscle quadrants, the formation of the initial left and right muscle rows appears unaffected in the laminin-depleted embryos (data not shown).  I also examined lam-1(ok3139) embryos, which are mutant for a putative null allele of the laminin b subunit. The mutant embryos show pre-hypodermal enclosure arrest and the early muscle rows do not form. The general tissue organization of these embryos appears to be defective (Appendix A: Supplementary Figure 1), which makes it impossible to determine whether loss of laminin has a direct or indirect affect on the early muscle organization. As complete loss of laminin results in embryos that arrest before the later muscle migration events occur, I used lam-1 RNAi for my further analysis.     30 To determine whether the missing anterior-most ventral cells were absent or misplaced, I used a 4-D lineaging system (Schnabel et al., 1997) to follow these cells in laminin depleted embryos. Tracing the cells' migratory pathways in these embryos, I found that the cells were still present, but displaced to the posterior, having failed to properly migrate to the anterior of the embryo (Figure 2.6). Observing the laminin depleted embryos during migration reveals that the anterior-most ventral muscle cells fail to extend membrane processes properly to the anterior of the embryos, while their dorsal counterparts do extend proper processes (Figure 2.5).  Since integrin is a well-characterized receptor for laminin mediated cell migration (Reviewed in Belkin and Stepp, 2000), I tested whether this is also true for C. elegans embryonic muscle migration. I tested a null allele of the sole C. elegans ?-integrin, PAT-3. Consistent with pervious studies (Hresko et al., 1994; Gettner et al., 1995), it did not show any defects in anterior extensions or other aspects of embryonic muscle migration (data not shown). 2.4.3 Proper ventral muscle process extension requires the ? -integrin INA-1, the ephrin VAB-2, and the ROBO receptor SAX-3  The displacement of the anterior most ventral muscle cells in the laminin RNAi treated embryos is very similar to the embryonic phenotype of the ?notched head? mutants studied by Tucker and Han (2008). I therefore examined the mutants used in that study to determine whether these mutants also are defective for anterior muscle process extension in the ventral muscle cells. I first    31   Figure 2.5: Depletion of laminin by RNAi results in defects in dorsal and ventral muscle migration and laminin along with vab-2, ina-1, and sax-3 is required for proper extension of the ventral anterior muscle processes. Panels A and B are of a wild type embryo and C - F are of RNAi treated or mutant embryos. A is Nomarski images of a developing embryo. B - F are fluorescent images of membrane-bound GFP labeled muscle cells. Arrows show misplaced cells and the brackets indicate missing anterior-most ventral muscle cells. In all panels anterior is to the left and the ventral side of the embryo is to the bottom. Scale bar is 10?m.     32 Table 2.1 Percent of 1.5 ? 2 fold embryos with defects in ventral anterior muscle process extension.  Percent of embryos with muscle process extension defects a, wt 2% sax-3(ky123) 66% sax-3(ky123); pajm-1::sax-3 60% sax-3(ky123); prgef-1::sax-3 25% vab-2(e96) 60% ina-1(gm86) 58% lam-2(RNAi) 42% lam-1(RNAi) 76% epi-1(RNAi) + lam-3(RNAi) 32% egl-15(ok2314) 0% egl-20(n585) 0% lin-44(n1792) 0% cwn-1(ok546) 2% cwn-2(ok895) 2% mab-20(ev778) 2% smp-1(ev715) 2% smp-2(ev709) 0% unc-6(e78) 4% a n = 50    33   Figure 2.6: The anterior-most ventral muscle cells in laminin(RNAi) treated embryos are displaced to the posterior. Panels A, B and C are of a wild type embryos and panels D, E and F are of a lam-2 (RNAi) treated embryo. A and D are Normaski images of developing embryos. Panels B and E are fluorescent images showing GFP labeled muscle cells. Panels C and F are plots of the migrations of the anterior-most ventral left cell indicated by the arrows in panels B and E. Dots represent the position of the cell's nuclei in the embryo at consecutive time-points, with the connecting line representing the distance traveled. Time-points were taken every 35s. Muscle cells migrate ventrally, but fail to migrate to the anterior, as in wild type. In all panels anterior is to the left and the ventral side of the embryo is to the bottom. Scale bar is 10?m. examined ?-integrin, ina-1 mutants and found the ventral muscle cells failed to properly extend anterior processes (Table 2.1) resulting in posterior displaced muscle cell similar to that seen in the laminin RNAi (Figure 2.6). As pat-3, the    34 only known ?-integrin in C. elegans mutants did not show any defects in muscle cell extension, ina-1?s involvement was unexpected. vab-2/ephrin mutants, displayed a similar phenotype to ina-1 mutants (Table 2.1 and Figure 2.5), as did mutants for the VAB-2 receptor VAB-1 (data not shown). My analysis suggests that the ?notched head? class of mutants is associated with the ventral anterior muscle process extension.  These results suggested SAX-3/Robo as another potential candidate, as certain alleles of sax-3 show a ?notched head? phenotype as well. Testing sax-3 mutants revealed they are defective for ventral anterior muscle process extension (Table 2.1). These sax-3 mutants result in the same posterior displacement of the ventral anterior-most cells seen in the other mutants (Figure 2.5), though it, as with the ina-1, vab-1 and vab-2 mutants, displays none of the dorsal-ventral migration defects seen in laminin RNAi embryos. The mutant alleles of the canonical SAX-3 ligand SLT-1 show no body morphology defects suggesting that SAX-3 does not act as a receptor for SLT-1 in this case. Live imaging showed that ventral muscle cells in sax-3 mutants retain the ability to extend the processes to the anterior, as some cells still extend processes, but they are diminished in thickness and are misguided. Examination of both the ina-1 and vab-2 mutants and the laminin RNAi embryos also showed the same phenomenon, suggesting that the loss of these genes is affecting the ventral anterior muscle processes in a similar fashion.  Since ROBO, integrin and ephrin signaling seemed to be involved in a subset of muscle cell migrations I wanted to determine whether other signaling pathways used in cell migrations were involved. I examined mutants in the UNC-   35 6/netrin, the LIN-44, EGL-20, CWN-1 and CWN-2 Wnt proteins, the FGF receptor EGL-15 and the semaphorins MAB-20, SMP-1 and SMP-2. In addition I targeted with RNAi knockdown members of the Ig family of receptors, cadherins and the Notch signaling pathway. Even after this extensive survey of genes I was unable to identify any muscle defects in mutants or RNAi knockdowns. (Table 2.1 and Appendix A: Supplementary Table A.1). Previously published phenotypic characterizations of the genes that I tested with RNAi also revealed no phenotypes consistent with mislocalized muscle. 2.4.4 Improper ventral muscle process extension is not due to defects in ventral hypodermal morphogenesis  As these mutations also affect hypodermal morphogenesis, I next observed hypodermal and muscle development in the mutant strains to determine if there were any hypodermal defects associated with the improper extension of ventral muscle processes. As muscle cells adhere to the hypodermis via the ECM later in development (Reviewed in Kramer, 2005; Labouesse, 2006; Moerman and Williams, 2006), the possibility exists that the anterior muscle extensions are a consequence of muscle cells adhering to migrating hypodermal cells, rather than independent migrations of the muscle cells themselves. Using an adherens junction marker, DLG-1::RFP (a gift  from M. Labouesse), I was able to visualize the boundaries of the hypodermal cells alongside the muscle cells (Firestein and Rongo, 2001). This allowed me to visualize how the extension of the anterior muscle processes relates to anterior hypodermal enclosure.  I found that the hypodermal anterior epiboly coincides    36 with anterior muscle migration. The leading edge of the anterior muscle processes is closely associated with the leading edge of the hypodermal sheet (Figure 2.7). Following enclosure by the hypodermis, the cell bodies of anterior-most muscle cells are located adjacent to the H0 and H1 hypodermal seam cells, with their anterior processes extending across the anterior edge of hyp7, the entirety of hyp6 and hyp5 and into hyp4 (Figure 2.7). Though I lack the resolution to determine the anterior endpoint of these processes in the embryo, transmission electron microscopy of the anterior regions of adults has shown that body wall muscle does not extend all the way to the anterior tip of the animal, but terminates before the hyp4-hyp3 boundary (Altun and Hall, 2009). This may also be the case for the anterior processes in the embryo. Altogether, muscle migration and hypodermal enclosure appear to coincide and defects in muscle migration may be a reflection of improper hypodermal enclosure. To test whether this was the case, I visualized the hypodermis in the mutant strains by using a GFP-tagged adherens junction marker, AJM-1, in combination with the muscle membrane marker. While muscle process extension defects occurred when the mutant or RNAi treated embryos exhibited ventral enclosure defects, improper ventral muscle process extension also occurs when hypodermal enclosure is normal (Figure 2.7). I found no correlation between hypodermal closure and muscle migratory defects, which lead me to conclude that defects in the hypodermis are not the cause of the muscle defects found in these mutant strains. The persistence of improper anterior muscle process extension even when hypodermal epiboly is normal in these mutants suggests    37 that the anterior muscle membrane processes are not simply the result of physical towing by the hypodermal sheet.  2.4.5 SAX-3 is required in the nervous system for proper extension of ventral anterior muscle processes  It was previously shown that the posterior displacement of ventral anterior muscle cells in INA-1/a integrin and VAB-2/VAB-1/ephrin/ephrin receptor could be rescued by restoring expression in the hypodermis and nervous system respectively (Tucker and Han, 2008). As sax-3 is expressed in both neurons and the hypodermis I wished to determine in which tissue it was functioning to affect muscle processes extension. To do this I assessed the ability of SAX-3 protein to rescue the muscle phenotype using either a pan-neuronal or hypodermal promoter in a sax-3 mutant background. I used the rgef-1 promoter, which drives expression in all post mitotic neurons (Altun-Gultekin et al., 2001), or the ajm-1 promoter, which drives hypodermal expression (Mohler et al., 1998). I found that hypodermal expression of sax-3 failed to rescue the ventral muscle migration defects, but expression in the nervous system did rescue the defects (Table 2.1). This indicates that sax-3 is required in neurons for proper ventral muscle migration.    38   Figure 2.7: Anterior muscle processes extend during of hypodermal anterior enclosure and converge with the hypodermal leading edge at the sensory depression. Panels A and C are Nomarski images of developing embryos. Panels B and D are fluorescent images of a strain showing the membrane-bound GFP labeled muscle cells and the DLG-1::RFP labeled hypodermal adherens junctions. E-H are fluorescent images of the mutant and RNAi treated embryos showing the membrane-bound GFP labeled muscle cells and the AJM::GFP labeled hypodermal adherens junctions. In panel B the white arrow indicates the leading edge of the hypodermis and the white triangle an    39 extending muscle process. The arrow in panel D indicates the sensory depression. Brackets in E-H mark missing or defective anterior ventral muscle processes. In all panels anterior is to the left and in A and B the right side of the embryo is to the bottom and in C and D ventral is to the bottom. Scale bar is 10?m.  2.4.6 Mislocalized neurons are associated with ventral anterior muscle defects  Both sax-3 and vab-2/vab-1 have been implicated in hypodermal ventral enclosure and neuronal migration (George et al., 1998; Mohler et al., 1998; Zallen et al., 1998). Since both are required in neurons for proper ventral muscle migration, I was interested to observe the organization of the nervous system in these animals. To visualize the neurons a strain expressing a mCherry tagged histone marker in neuronal precursors and postmitotic neurons was crossed into muscle marker containing strains. Examining the relationship between the muscle cells and neurons in wild type embryos, I observed a neuron free zone between the neurons lining the ventral surface of the embryo and a more dorsally situated cluster of neurons that the ventral muscle cells pass through (Figure 2.8). In the sax-3 and vab-2 mutant embryos, where the ventral muscle fails to extend normally, neurons are disorganized and appear to be in the migratory path of the muscle cells. This neuronal disorganization is also apparent in the laminin depleted and ina-1 mutant embryos (Figure 2.8). Proper neuronal organization appears to be a key factor in promoting muscle process extension.    40     Fig 2.8: Displaced neurons are associated with improper extension of the ventral anterior muscle processes. Panels A, C, E, G and I are fluorescent images of GFP labeled muscle membranes and mCherry labeled neuronal nuclei. B, D, F, H and J show the neurons alone, with the outline of the embryo indicated in white. Arrows indicate the region of the cluster of sensory neurons present in the wild type.  In all panels anterior is to the left and the ventral side of the embryo is to the bottom. Scale bar is 10?m.     41 2.5 Discussion 2.5.1 C. elegans muscle as a model to study muscle cell migration in vivo.   C. elegans has proven to be an excellent model for the  study of cell migration (For review see, Chen and Stern, 1998; Lehmann, 2001; Simske and Hardin, 2001; Killeen and Sybingco, 2008). Embryonic muscle migration provides a model for coordinated and simultaneous movement of a large number of cells in the very dynamic environment of a developing embryo as a prerequisite for proper tissue organization itself. While the basic movements of muscle cells have been described in the past (Sulston et al., 1983; Hresko et al., 1994; Schnabel et al., 1997), none of these earlier studies determined whether muscle migrations are the result of directed cell migration, or occur via some other means of organization such as differential cell adhesion or cell focusing (Schnabel et al., 2006; Steinberg, 2007). In this study I provide evidence of lamellipodia and filopodia, hallmarks of directional cell migration, in migrating C. elegans embryonic muscle cells. Directed cell migration appears to be necessary only in the later organization stages, since the formation of the initial lateral muscle rows occurs in the absence of any telltale membrane extensions, although I cannot rule out the involvement of short-range migrations in this initial organization. Taken altogether, this makes embryonic muscle cells in C. elegans an excellent system for the study of tissue organization and cell migration.  The high degree of cell contact maintained between dorsal and ventral muscle quadrants while they migrate in opposing directions is intriguing. This is    42 especially striking when one considers that some muscle cells maintain a connection with their dorsal or ventral contralateral counterparts, while other anterior and posterior ipsilateral neighbours do not. These cell-cell contacts between the contralateral cells may simply be the result of homophilic adhesion between the muscle cells so that the contacts are broken by tearing forces, as the cells separate. However, if the connections are only due to adhesion, I would expect that both cells contribute equally and this is not what I observe. Some contacts are due to one cell process extending across to the opposite side, with no apparent contribution from the target cell. I also observe that, after the dissolution of these contacts, the processes can re-extend and re-establish contact across the intervening space between the contralateral quadrants. While the cause of this phenomenon remains unknown, the active membrane dynamics of these cells at their lagging edge suggests the possibility of continued communication between cells of the dorsal and ventral quadrants as they migrate apart. 2.5.2 Embryonic muscle migrations in C. elegans require laminin.  In mammals and Drosophila, laminin is a well-characterized substrate for muscle migration (For review see, Ocalan et al., 1988; Yarnitzky and Volk, 1995; Ryan et al., 1996). While previous studies have shown that loss of laminin in C. elegans results in defects in muscle attachment and sarcomere organization, (Huang et al., 2003; Kao et al., 2006), they did not identify any requirement for laminin in muscle cell migration. I demonstrated that laminin is indeed required for late muscle migration in C. elegans.    43  The best characterized receptor involved in mediating muscle migration across a laminin substrate is the heterodimeric receptor integrin (Buck and Horwitz, 1987; Crawley et al., 1997; Belkin and Stepp, 2000). C. elegans has one ? subunit and two ? subunits, producing two integrin receptors (Gettner et al., 1995; Baum and Garriga, 1997). Interestingly, loss of the either of the subunits of the primary muscle integrin receptor, PAT-3/ ?-integrin or PAT-2/ ?-integrin, results in embryonic lethality without affecting muscle migration (Gettner et al., 1995; Hresko et al., 1994; Viveiros, Pers. Obs.; Williams and Waterston, 1994), suggesting that integrins may not mediate binding of muscle cells to laminin during migration or that laminin indirectly affects migration. The lack of muscle migration defects in the only ?-integrin (pat-3) is particularly surprising considering that ina-1/ ?-integrin is required for the proper migration of some of the muscle cells. This incongruity supports the idea that ina-1 mediates these migrations without a ?-integrin. Though an unconventional explanation, it is consistent with the data and this would be the first example of an a-integrin with a function independent of a ?-integrin partner. Alternatively there may be another unidentified ?-integrin present in the genome.   Laminins can also interact with non-integrin receptors, some of which have C. elegans homologues, including dystroglycan, the 32/67 kDa laminin receptor, galactoside-binding lectin, heparin sulfate proteoglycans, (Reviewed in Patarroyo et al., 2002). Most of these genes were included in my initial RNAi screen and exhibited no discernable muscle phenotype. RNAi does not always results in a complete loss of gene function, so that negative RNAi results need to    44 be interpreted with caution. For a number of these genes mutants are available and have previously been characterized: there is evidence that dystroglycan functions as a receptor for laminin in C. elegans, but this does not appear to be relevant for muscle development (Johnson et al., 2006). Of the heparin sulfate proteoglycans in C. elegans, agrin, perlecan, glypican and syndecan (Rogalski et al., 1993; Minniti et al., 2004; Gumienny et al., 2007; Hrus et al., 2007), only perlecan has been found to a play a role in muscle function and it has no effect on muscle migration. Although these laminin receptors appear not to be required for muscle migrations, it is possible some act redundantly, or that additional receptors remain to be identified.  Defects in overall ECM organization could potentially impede the muscle cells' ability to migrate. This seems unlikely as loss of function studies on other ECM components in C. elegans, including basement membrane collagens, heparin sulphate proteoglycans and nidogen, have not revealed any defects in muscle cell migration (Guo et al., 1991; Rogalski et al., 1993; Kang and Kramer, 2000; Minniti et al., 2004; Gumienny et al., 2007; Hrus et al., 2007). Looking at the distribution of perlecan in relationship to the migrating muscle cells, while considering its known role in mediating muscle adhesion and non-involvement in muscle migration (Hresko et al., 1994; Rogalski et al., 1993), it is easy to surmise that perlecan and these other ECM proteins may act more as glue to lock the cell in place rather than as substrates or guidance cues for muscle migration. While it is possible that loss of a single ECM component may not be sufficient to disrupt the muscle cells ability to migrate, it seems more likely that embryonic muscle    45 migration is facilitated in the presence of laminin and may be independent of all other currently identified ECM components. 2.5.3 Multiple signaling pathways affect ventral anterior muscle processes Axon guidance and cell migration share many similarities and over the past several years a number of signaling pathways have been implicated in axon guidance, including the ephrin, slit, wnt and netrin pathways (Reviewed in Quinn and Wadsworth, 2006; Killeen and Sybingco, 2008). The pathways have also been shown to mediate cell migration outside the nervous system, including hypodermal ventral enclosure and distal tip cell migration (Su et al., 2000; Ghenea et al., 2005). Here I identified integrin, ephrin and Robo signaling pathways as required for proper migration of muscle cells in the anterior ventral region of the developing embryo. I did not find any evidence implicating Wnt, Netrin or semaphorin signaling in this process. The disrupted integrin, ephrin and slit signaling pathways do not seem to act cell autonomously in displaced muscle cells.  Tucker and Han (2008) showed that the notched head phenotype of ina-1 and vab-2 animals could be rescued by expression in the hypodermis or nervous system, respectively, and I have shown that neuronal sax-3 is also required for proper anterior muscle morphogenesis. As ROBO and ephrin signaling are both required for ventral enclosure of the hypodermis (Ghenea et al., 2005), I considered whether the muscle defects I observe might be the result of defects in hypodermal epiboly. I did find that ventral enclosure defects affected the    46 underlying muscle cells, but I also observed examples of normal hypodermal closure and improper muscle process extension As ina-1, sax-3 and vab-2 are all known to be involved in mediating neuronal migration (Baum and Garriga, 1997; Zallen et al., 1998; Chin-Sang et al., 1999) I examined the distribution of neurons in the embryo in these mutants. Most evident in these mutants was the disorganization, shared by the laminin depleted embryos, of a the anterior neurons that results in the mislocalization of neurons into the path of the migrating muscle cells, displacing the cells that would normally form the muscle processes migratory environment. . This disruption may be due to an anterior shift of the sensory cells, since amphid neurons are anteriorly displaced in ephrin and ROBO deficient adult animals (Ghenea et al., 2005). These neuronal defects offer an explanation for why disruption of anterior muscle processes is limited to ventral muscle cells in these mutants, since at this developmental stage the majority of the neurons are ventrally located and very few are near the anterior dorsal muscle cells (Sulston et al., 1983; Hallam et al., 2000). These defects in organization of neurons in the head region may affect the pathfinding ability of muscle cells, since these cells can still extend processes, but these processes cannot find their target. This may be due to conflicting signals originating from the neurons or defects in the ECM resulting from the presence of the mislocalized cells. Another attractive model is that of steric hindrance: it may be that in the anterior ventral quadrant muscle extensions are simply blocked from reaching their proper destination by the shift in neural cell bodies. This latter model explains why only ventral anterior    47 processes are affected and not dorsal muscle extension as well. This obviates the need to postulate separate signaling mechanisms for the dorsal and ventral anterior processes. What is required now, if understanding how anterior muscle extensions find their targets is to be elucidated, is to identify mutants that affect both the ventral and dorsal anterior muscle membrane process extensions.  2.5.4 Anterior muscle processes are required for anterior migration  Previous studies of embryonic muscle migration in C. elegans proposed that flattening of muscle cells was responsible for the extension of the muscle quadrants as the embryo elongated (Hresko et al., 1994; Sulston et al., 1983). While flattening most likely plays a role, I have identified an anterior migration event that is mediated by processes extending from the anterior-most pair of cells in each muscle quadrant. Anterior migration of muscle cells is dependent upon the extension of these processes, as loss of the ventral processes in laminin knockdown embryos and ina-1, vab-2 and sax-3 mutant embryos prevents anterior migration of the ventral muscle cells, while the dorsal cells, whose processes are unaffected, migrate normally. What role these processes are playing in anterior muscle migration is currently unknown. It seems likely that they provide a tether for the anterior extension of the muscle quadrant, but this remains to be proven. Though not a certainty, they appear to be required for proper muscle morphogenesis.   The extension of these anterior processes coincides with the anterior enclosure of the embryo by the hypodermal sheet and muscle cells fail to extend processes when hypodermal enclosure is defective, as in the sax-3 and vab-2    48 mutants, suggesting a potential connection between the two processes. Though anterior epiboly appears to be necessary for anterior muscle migration, as muscle cells fail to migrate properly in animals with defective hypodermal enclosure, it is unlikely that the anteriorly migrating hypodermis is contributing to the anterior extension of these muscle processes by physically towing them along. The anterior muscle cells show lamellipodia and filopodia directed towards the anterior, implying that they play an active role in the membrane extension and in the mutant embryos that extend defective processes, these processes continue to wander even after epiboly has concluded. Moreover, the improper extension of the ventral membrane processes in the laminin RNAi and Robo, ephrin and integrin mutants occurs even when anterior hypodermal enclosure occurs normally, indicating that anterior hypodermal enclosure is not sufficient for anterior muscle process extension. The hypodermis is most likely providing a substrate for the extending anterior processes with muscle and hypodermal crosstalk resulting in the deposition of ECM to reinforce the adhesion between the tissues once the muscle cells have migrated to their final location. This is supported by the observation that perlecan is deposited beneath the processes as they extend and that it is only present beneath the area of the muscle cells that have reached their final location. The perlecan isoform responsible for muscle attachment at this stage in development is made in the hypodermis and its secretion is dependent upon the presence of the underlying muscle (Moerman et al., 1996; Spike et al., 2002). While there is no evidence that direct contact is required, it seems probable considering the extremely short distances involved.     49 Proper organization of the nascent nervous system appears necessary for the extension of the ventral muscle processes. In the ina-1, vab-2 and sax-3 mutants, as well as the laminin knockdown embryos, the loss of the ventral muscle processes is associated with disorganized neurons that appear to be in the direct migratory path of the ventral anterior muscle processes. This could be a case of steric hindrance, whereby the mislocalized neurons physically block the migration of muscle cells. I favor this model as it readily explains the differences I observe in muscle cell extensions between the dorsal and ventral muscle quadrants. While possible, it seems unlikely that each quadrant would use unique guidance cues.  If steric hindrance is the explanation for these observed alterations in anterior muscle migrations, then I have identified only one protein that directly affects this migration process, laminin, which appears to be required for the initial formation of the first two muscle clusters and the later dorsal-ventral migrations, as well as indirectly affecting the ventral anterior muscle processes.  Considering the number of ECM protein components and receptors I screened by RNAi or mutant analysis this is a surprising observation. These results may represent genetic redundancy or buffering, but I suspect the paucity of genes detected by my screen is the result of my focus on defects in the older, post epiboly embryos. It is highly probable that early organization of the two lateral muscle cell clusters and the later migrations that form the final four quadrants require the same components. If that is the case I was limited to detecting only those genes that behave like laminin, where a weak allele proceeds past epiboly and does not    50 show morphological defects until later in development. Using automated 4-D lineaging (Bao et al., 2006; Murray et al., 2008) it should be possible to identify such genes by examining early embryos and determining whether muscles cells are the only cell type disorganized in a mutant embryo.      51 3 UNC-54 and MEM-1 Work in Concert to Inhibit Ectopic Membrane Extensions Away From the Nerve Cords in C. elegans Body Wall Muscle.  3.1 Synopsis  In Drosophila and mice, muscle membrane extensions play an important role in the establishment of contact between neurons and muscle so that neuromuscular junctions can form. In C. elegans, muscle cells also extend membrane processes to facilitate neuromuscular junction formation. Disruption of certain genes, such as those involved in the FGF signaling pathway and components of integrin adhesion complexes have previously been shown to result in the formation of ectopic muscle membrane extension distinct from those normally required for proper innervation. In this study, I have identified two more genes whose loss also results in the production of these membrane processes, unc-54, a myosin heavy chain B and mem-1 (Y54E5B.2), a previously uncharacterized gene. These two genes appear to work in a synergistic fashion to inhibit improper membrane extensions. In the course of my analysis, I have also identified that the ?-integrin derived, plasma membrane localized fluorescent markers used to visualize muscle membrane extension actually sensitize muscle cells for their production. This sensitization is also conferred by the expression of full length, fluorescently tagged PAT-3/ ?-integrin, providing a useful tool for screening for new genes involved in the negative regulation of membrane extensions.    52 3.2 Introduction   The regulation of membrane protrusions plays an important role in a number of developmental processes. It is required for epiboly and ingression during gastrulation, for neurite extension during nervous system development and all forms of cell migration (For review see Solnica-Krezel, 2005; O'Donnell et al., 2009; Petrie et al., 2009). These processes also play a central role in cancer progression, as the invasion of cancerous cells in to the surrounding tissue and their migration away from the primary tumor are the hallmarks of metastasis, which severely reduces the survival chances of cancer sufferers (Reviewed by Nurnberg et al., 2011). While many of the guidance cues, extracellular matrix proteins, cell surface receptors and cytoplasmic effectors responsible for the cytoskeleton remodeling that drives these processes are well characterized, we still do not have a complete picture of how these events unfold (Mattila and Lappalainen, 2008; Petrie et al., 2009). Muscle development provides an excellent model to study membrane processes. During embryogenesis, nascent myoblasts must migrate from the somites, to the sites of muscle formation (Buckingham, 2001) and once at their destination, these myoblasts must interact with each other and organize themselves before fusing into myotubes (Snow et al., 2008). Muscle cells have also been shown extend filopodia to interact with approaching neurons to facilitate the formation of neuromuscular junctions (NMJ) and these structures are called myopodia (Uhm et al., 2001; Ritzenthaler and Chiba, 2003; Madhavan et al., 2006).  These myopodia are induced via the basement membrane    53 proteoglycan agrin (Uhm et al., 2001) and it has been shown that this agrin induction is dependent on p120 catenin, a component of cadherin adhesion complexes (Madhavan et al., 2006). Though some progress has been made, the proteins involved in regulating these processes are still largely unknown. In C. elegans, muscle cells also extend membrane processes, termed muscle arms, to help facilitate NMJ formation. Since C. elegans neurons don?t extend axons directly to the muscle cells, these muscle membrane processes are necessary to bring the two cell types into contact so NMJs can form and they persist for the life of the animal (White et al., 1986; Dixon and Roy, 2005). Previous studies have identified a number of proteins that are involved in mediating these muscle arms, including cell adhesion components and the fibroblast growth factor (FGF), insulin and UNC-40/DCC signaling pathways (Dixon and Roy, 2005; Dixon et al., 2006; Dixon et al., 2008; Alexander et al., 2009). Interestingly, the disruption, such as in the case of loss of FGF signaling, or over expression, as with UNC-40/DCC, of some of these components can also result in the generation of ectopic muscle membrane extensions away from the nerve cords (Dixon et al., 2006; Alexander et al., 2009), making it an exciting model for the identification of factors involved in promoting defective membrane behavior. In this study, I have identified two more genes that appear to be involved in the inhibition of these ectopic membrane extensions. The first, unc-54, a C. elegans muscle specific myosin heavy chain B homologue (MacLeod et al., 1981), has been previously shown be required for proper muscle arm extension    54 (Dixon and Roy, 2005), but has not previously been reported to be involved in EMEs. The other is mem-1, a previously uncharacterized gene of unknown function. The loss of either of these genes results in the generation of ectopic muscle membrane extensions. While the loss of unc-54 maybe be promoting EMEs simply due to the destabilization of the muscle adhesion complexes, whose components have previously been shown to be required to inhibit these processes (Dixon et al., 2006), mem-1?s role in inhibiting these processes remains the be determined.   3.3 Materials and methods  3.3.1 C. elegans strains  Animals were maintained as described (Brenner, 1974). The N2 Bristol strain was used as reference wild type. The strains used in this study were: GE6583 unc-54(t3197), mem-1(t3198); PD7963 ccIs7963[phlh-1::GFP];; VH95, hdIs61[phlh-1::memGFP, rol-6(su1006)]; RP247 trIs30[him-4p::MB::YFP + hmr-1b::DsRed2 + unc-129nsp::DsRed2], VG147 zdIs5[pmec-4::gfp]. The Hawaiian strain used for SNP mapping was CB4856. Quantification of EMEs was done using the trls30 marker. 3.3.2 RNAi  Feeding RNAi was done as per Kamath et al (2003). Feeding was done at the non-permissive temperature in the double mutant using the trls30 marker. L1 animals were grown on E. coli expressing the RNAi vector until they reached    55 adulthood. Young adults were then harvested and assayed for muscle membrane defects. Each RNAi experiment was performed in triplicate and the results from the most penetrant RNAi replicate were used for the comparison. 3.3.3 Microscopy Imaging was done using either a Zeiss Axioplan 2 imaging compound microscope with 2.5x Optivar and a Q imaging Retiga Exi digital camera and inverted a Zeiss Axiovert LSM 5 confocal microscope equipped with epifluorescence, Nomarski optics, and LSM 5 Pascal software. Image acquisition and analysis was done with the LSM 5 Pascal volume-rendering software 3.3.4 Microinjection procedure  The generation of transgenic animals was performed via microinjection.  To generate the rescue lines fosmids carrying genomic fragments contain either unc-54 or Y54E5B.2 were injected into GE6583. Three co-injection plasmids were also used: pRF4 [rol-6(su1006dm)], which contains a copy of rol-6 and confers a dominant Rol phenotype, pBx [pha-1::pha-1(+)], which was used as filled DNA and pCFJ90 [pmyo-2::RFP], which contains a pharyngeally expressed RFP. Injection mixes were prepared containing 40ng/ml pRF4, 40ng/ml pBx [pha-1::pha-1(+)], 10ng/ml pCFJ90 and 10ng/ml of fosmid. All injections were carried out using a microinjection setup featuring a Zeiss inverted compound microscope (IM35) by conventional methods (Mello and Fire, 1995).    56  3.3.5 Isolation of temperature sensitive mutants using EMS mutagenesis  N2 worms were mutagenized in 50 mM ethyl methanesulfonate (EMS) for 4 hr at room temperature. Mutagenized worms were grown at 15?C on NGM plates seeded with E. coli OP50. Animals were allowed to propagate for two generations and then individual F2 animals were transferred onto fresh 60mm NGM plates and allowed to establish clonal populations. Samples of these populations were then transferred onto separate plates and grown on at 25?C and scored for mutant phenotypes. Mutants suspected to have phenotypes associated with muscle defects at the non-permissive temperature, such as embryos arresting during elongation or animals with uncoordinated movement, were crossed into a muscle cell membrane marker and then assayed for muscle morphology defects.  3.3.6 Sequencing of GE6583 and identification of candidate mutations  Sequencing of GE6583 was performed at the Oklahoma Medical Research Foundation sequencing center and it along with the sequence analysis was done as per (Flibotte et al., 2010), with the following modification to the genomic DNA isolation protocol. GE6583 worms were grown to starvation and harvested by washing into 15-mL centrifuge tubes with 10 mL of M9 buffer containing 0.01% Triton X-100. They were then washed seven times by centrifugation, removal of supernatant by aspiration, and resuspension and vortexing in fresh M9. After the final wash ~400ml pellets were frozen at -80?C.    57 Thawed aliquots of pelleted worms were transferred to 1.5-mL microcentrifuge tubes containing lysis buffer (50 mM KCl, 10 mM Tris-HCl at pH 8.3, 2.5 mM MgCl2, 0.45% NP-40 [Igepal], 0.45% Tween-20, 0.01% gelatin, 300 ?g/mL Proteinase K), frozen at ?20?C, and incubated at 55?C?60?C for 4-5 h, followed by RNase A treatment. Proteins were then precipitated using 6M NaCl and DNA isolated by isopropanol precipitation. DNA was washed in 70% ethanol, air dried and resuspended in TE (10 mM Tris-HCl, 1 mM EDTA at pH 7.0?8.0). DNA concentrations were determined using a Thermo Fisher Nanodrop spectrophotometer, and quality was checked by electrophoresis on 1% agarose gels.  3.3.7 Mapping the GE6583 causative allele  SNP-Chip mapping was done as per (Flibotte et al., 2009) with the following modifications. To generate the DNA sample, GE6583 hermaphrodites were mated to CB4856 males at 15?C. F1 heterozygotes were then plated on 60mm agar plates seeded with OP50 E. coli and grown at 25?C. A 1000 F2 animals displaying the GE6583 paralyzed phenotype were then collected and plated in groups of 10 on fresh 60mm plates and grown to starvation. Worms were then collected and DNA was prepared in the same manner as described in section 3.2.4. Two biological replicated were performed and the data was combined to map the causative allele.     58 3.4 Results  3.4.1  Identification of t3197, a new temperature sensitive allele of the C. elegans myosin, unc-54, that results in ectopic muscle membrane extensions.  t3197 was isolated in a screen to identify new temperature sensitive lethal mutations. At the non-permissive temperature the strain displays low level larval lethality (~15%) and those animals that survive are severely uncoordinated, with most appearing to be completely paralyzed and failing to lay eggs. These animals also exhibit ectopic muscle membrane extensions similar to those previously reported in animals defective in FGF signaling and muscle attachment (Dixon et al., 2006) (Figure 3.1) and this muscle morphology defect was the focus of this study. These membrane extensions appear to travel along the inner surface of the hypodermis and can cross the area between muscle quadrants, resulting in ectopic membrane sheets covering the area between the dorsal and ventral muscle cells (Figure 3.1). Similar to the EMEs reported by Dixon et al. (2006), these ectopic extensions arose during larval development and shifting worms to the non-permissive temperature after they had reached adulthood did not significantly increase the muscle membrane defects (data not shown). To identify the location of the mutation responsible for the membrane extension defects, I performed single nucleotide polymorphism mapping using a microarray ship (SNP-CHIP) (Flibotte et al., 2009), using the Hawaiian wild type isolate strain for comparison. Results of the SNP-CHIP mapping indicated that the causative mutation was located on the end of the right arm of chromosome I    59 (Appendix B, Figure B1).  Whole genome sequencing revealed a point mutation (an A-to-T transversion) in unc-54, at position 1873 in the coding region that results in a tyrosine(Y)-to-asparagine(N) amino acid change at position 625. The Y625N mutation resides in the myosin head domain of the protein (Figure 3.2). A complementation test was performed at the non-permissive temperature between t3197 and e190, an unc-54 null allele, and the paralyzed phenotype failed to complement, indicating that t3197 is a new temperature sensitive unc-54 allele. At the non-permissive temperature t3197 appears to phenocopy the e190 allele, suggesting that t3197 behaves as a null at the non-permissive temperature. Two other temperature sensitive alleles of unc-54 have previously been isolated, e1175 and e1301, though neither have been sequenced (Waterston et al., 1980), making t3197 the first temperature sensitive allele of unc-54 whose causative mutation is known.  3.4.2 The genes, unc-54 and mem-1 are both involved in mediating the ectopic muscle membrane extension phenotype.  While the paralyzed phenotype failed to complement with the unc-54 null allele, the ectopic membrane extension phenotype showed a marked reduction in the unc-54 t3197/e190 heterozygote animals, suggesting the involvement of a second gene (Table 3.1). Revisiting the sequence data I found a point mutation, t3198, (a G-A transition) in the coding region of a tightly linked gene, 36kb downstream of unc-54, Y54E5B.2, which I have renamed mem-1 (membrane morphology gene 1) that results in a leucine(L)-to-phenylalanine(F) amino acid     60    Figure 3.1: Ectopic membrane extensions in the unc-54(t3197) mem-1(t3198) double mutant strain. Panel A depicts wild type muscle. Panels B-D depict ectopic membrane extensions in the unc-54(t3197) mem-1(t3198) double mutant strain at the non-permissive temperature. All panels depict left lateral views of the animal with the exception of panel C with is a right lateral view. All panels depict fluorescent images of muscle membrane. Panels A and C are of phim-4::YPF animals, panel B is of a pmyo-3::GFP animals and panel C is of a PAT-3/?-integrin::GFP animal.  Arrows indicate EMEs. Scale bars are 20mm.    61   Figure 3.2: The t3197 temperature sensitive mutation resides in the myosin head domain of UNC-54/MHCB. Panel A is a diagram illustrating the conserved domains in the unc-54 protein sequence. The unlabeled green box represents a conserved myosin SH2 domain. Panel B is a sequence alignment of UNC-54 myosin homologues in other species. The was done using the Multalign sequence alignment tool (Corpet, 1988). Red highlights indicate amino acid positions with 100% identity between all the sequences and blue highlights mark positions with at least three sequences have identical or similar amino acids. In both panels the red arrow indicates the position of the t3197 Y625N allele.  change at position 699. MEM-1 contains two clusters of WD motifs, approximately 40-amino acid long stretches typically ending in Trp-Asp, but otherwise exhibiting only limited amino acid sequence conservation. WD motifs are often involved in mediating protein-protein interactions (Neer et al., 1994). The L699F mutation resides in the C-terminal WD motif cluster of the protein (Figure 3.3). MEM-1 is predicted to be the C. elegans homologue of WDR6, a human WD motif containing protein (Li et al., 2000). WDR6 has previously been shown to interact via its C-terminal WD motif with the serine/threonine kinase    62 LKB1 (also known as STK11) (Xie et al., 2007), which when mutated leads to Peutz?Jeghers syndrome that is associated with increased incidence of malignant cancers (Marignani, 2005; Alessi et al., 2006). An alignment of the two proteins using ClustalW2 (Chenna et al., 2003) reveals substantial similarity between C. elegans MEM-1 and human WDR6 throughout the protein, with MEM-1 sharing 17% identity and 54% similarity with the human protein and both proteins contain a WD motif cluster at both the amino and carboxyl terminal ends (Figure 3.3).  To test if mem-1 was indeed involved, I generated a transgenic rescue line carrying a fosmid for mem-1. The rescue construct did not rescue the paralyzed phenotype, but was able to rescue the membrane extension defects to the levels observed in the unc-54 t3197/e190 heterozygote animals (Table 3.1), suggesting that mem-1 is also involved in the process. That a certain amount of membrane extension defects were still observed in both the mem-1 rescue line and the unc-54 t3197/e190 heterozygote animals, which are also heterozygous for the mem-1 mutation, suggested that loss of unc-54 alone was sufficient to result in EMEs. To test this I generated a line carrying the marker in an unc-54(e190) null background and found that loss of unc-54 alone was sufficient to generate EMEs, but these were not as frequent as observed in the double mutant line (Table 3.1). Knocking down mem-1 by RNAi    63      64 Figure 3.3: Comparison of MEM-1 protein sequence with human WDR6.   MEM-1 aligns strongly with WDR6 (Ensembl:ENSP00000413432) along the length of the protein and shares a similar arrangement of WD motif domains. Shaded regions indicated the predicted positions of the WD motif clusters in both sequences and the black box indicates the location of the t3198 L699F mutation in the terminal WD motif cluster. Amino acid alignments were marked using ClustalW2 conventions (Larkin et al., 2007): asterisks (*), identical amino acids; colons (:), conserved substitutions; periods (.), semi-conserved substitutions.  in the unc-54(e190) background resulted in an increase in the severity of the of the EME phenotype (Table 3.1), though knocking down mem-1 alone was unable to generate EMEs (data not shown). 3.4.3 Loss of mem-1 has no effect on muscle arm extension.  As both unc-54 and mem-1, appear to negatively regulate membrane extensions, and unc-54 had previously been found to also be required for normal muscle arm development, with unc-54 mutants failing to generate post-embryonic muscle arms, (Dixon and Roy, 2005), I wanted to determine whether the loss of mem-1 would also have an effect on muscle arm extensions. Using the same strain as Dixon and Roy (2005) to visualize the muscle arms, I determined that there is no difference in the muscle arm phenotype between the temperature sensitive double mutant and the unc-54(e190) null mutant, which suggests that mem-1 is not required for embryonic muscle arm extension (Table3.2). As the loss of unc-54 is sufficient to inhibit post-embryonic muscle    65 arms, I also observed the muscle arms in the unc-54 fosmid rescue strain and found that loss of mem-1 alone does not have any effect on normal muscle arm extensions (Table 3.2).   Table 3.1: Rescue of ectopic membrane extension by unc-54 or mem-1  #Severe refers to EMEs that extend the from one muscle quadrant to another e.g. Figure 3.1 panel D     n=100, n.s.=not significant         66 Table 3.2: Analysis of muscle arm extensions in the unc-54(t3197) mem-1(t3198) double mutant. Strain # of arms/muscle cell (n=100) p value Wild type 3.4 ? .8  unc-54(e190) 2.4 ? 1.1 <.0001 unc-54(t3197) mem-1(t3198) 2.2 ? 1.2 <.0001 unc-54(t3197) mem-1(t3198)  unc-54 rescue 3.3  ? 1.0 n.s.   3.4.4 GFP or YFP bound to the membrane via the ?-integrin transmembrane domain increases ectopic membrane extensions.  Over the course of my analysis of the ectopic membrane projections I used two different membrane marker strains, VH95, which has membrane-bound GFP expressed in all body wall muscle cells driven by the hlh-1/MyoD promoter (Viveiros et al., 2011) and RP247, which has membrane-bound YFP expressed in a subset of distal body wall muscle cells driven by the him-4 promoter (Dixon and Roy, 2005). In both of these constructs the transmembrane domain is derived from PAT-3/?-integrin. I observed that the mutant phenotype was more severe when visualized by the him-4 marker, even though expressed in fewer muscle cells, than when visualized by the hlh-1 driven marker, which is expressed in al the muscle cells (Figure 3.3). I also observed that, while the markers are distributed throughout the membrane, I do see some enrichment at    67 the muscle adhesion complexes, suggesting that the transmembrane domain of PAT-3/?-integrin is sufficient for localize the reporter protein to the muscle adhesion complexes. I also observed clumping of the marker in the membrane (Figure 3.3). Concerned that this localization of the markers might be affecting my results, I crossed a muscle specific, cytoplasmically located GFP marker into the double mutant. Using this method, I found that the severity of the EMEs was substantially reduced compared to that observed using either of the membrane bound markers (Table 3.4). To determine whether this sensitization was due to the PAT-3/?-integrin transmembrane domain GFP fusion protein interacting with PAT-3/?-integrin binding partners and thus preventing wild type protein complexes from forming, I visualized the EMEs using a PAT-3/?-integrin::GFP functional fusion, which rescues pat-3/?-integrin loss of function mutations. This is possible as the adhesion complexes become disorganized when unc-54 is disrupted and the normally tightly localized integrin marker becomes diffuse in the membrane (Figure 3.4). When using this marker, I also saw an increase in the penetrance of the EMEs, suggesting that the effect observed with the PAT-3/?-integrin transmembrane domain GFP fusion may not solely be due to it interfering with ?-integrin?s normal binding partners and that the functional fusion might not be behaving as wild type (Figure 3.5). Whether this sensitization is specific to PAT-3/?-integrin derived membrane markers, or a general effect of membrane-bound GFP still remains to be determined. Since the membrane markers appeared to sensitize animals for the production of EMEs, I wanted to see if the membrane extension defects    68 observed in the weIll characterized unc-54(e190) null background were due to this sensitization. When the cytoplasmic marker was used, the unc-54(e190) null mutants appeared wild type, indicating that loss of unc-54 does not normally result in EMEs (data not shown). This, along with the observation that RNAi targeting mem-1 fails to generate EMEs even in a sensitized background, the membrane defects observed using the cytoplasmic localized GFP in the mem-1, unc-54 double mutant are possibly the result of a synthetic effect between the two genes.   3.4.5 RNAi depletion of actin remodeling or cell adhesion components reduces the severity of the ectopic membrane extensions.  To determine whether EMEs might prove useful for identifying factors that can regulate membrane extensions, I used RNAi to target a small subset of genes known to be required for membrane extension, such as components of the Arp2/3 complex, the Rho family GTPase members Rac1 and Cdc42 required for cortical actin remodeling and components of the integrin adhesion complex required for proper attachment to the ECM (Reviewed in Le Clainche and Carlier, 2008) (Table 3.3). I found that by depleting genes involved in any of these processes I was able to reduce the severity of the EMEs. Two of the most effective RNAi targets were Cdc42 and UNC-112/kindlin. Since it has been previously been shown that Cdc42 interacts with UNC-112/Kindlin via the guanine nucleotide exchange factor (GEF) UIG-1 (Hikita et al., 2005), I tested to see if knocking down UIG-1 would also result in a rescue of the EME phenotype, but my RNAi experiments failed to show any effect, possibly due to inefficient    69 knockdown of the UIG-1 by the RNAi vector (Appendix B, Supplementary Table B.1).    Figure 3.4: Loss of unc-54 results in disruption of the muscle adhesion complexes in C. elegans body wall muscle. All panels are fluorescent images of body wall muscles expressing PAT-3/?-integrin::GFP. Panels A and B are images at the cell surface of the muscle cells showing the arrangement of integrin adhesion complexes. Panel C is an image taken at a focal plane in the middle of the cells and shows the diffuse localization of PAT-3/?-integrin::GFP throughout the plasma membrane. Scale bar is 20mm    70  Figure 3.5: PAT-3/?-integrin based membrane bound fluorophore markers sensitize muscle cells for the production of ectopic membrane extensions. Panels A, B and C are false coloured fluorescent images showing the wild type localization of PAT-3/?-integrin::GFP, hlh-1::memGFP(Mb::GFP) and him-4::memYFP (Mb::YFP) respectively. Scale bar is 20 mm Panel D is a table showing the proportion of animals exibiting EMEs in wild type and mutant backgrounds with each of the different flurorescent markers.  3.4.6 The unc-54(t3197) mem-1(t3198) double mutants exhibit neuronal defects.  As the muscle cells in the double mutant at the non-permissive temperature had severe defects in membrane extension, I examined other tissues to determine if they also displayed membrane extensions or other abnormalities. Touch neurons, specifically the AVM neuron, were chosen as an alternative tissue type    71 Table 3.3: Ectopic membrane extension defects can be rescued by targeted depletion of actin remodeling and cell adhesion components.      #Severe refers to EMEs that extend the from one muscle quadrant to another  e.g. Figure 3.1 panel D. Panel D: n=100.   to examine. Touch neurons have long axons that extend for a large part of the length of the worm, making them well suited to screen for neurite extension defects. The AVM neuron arises and migrates during larval development, allowing me to focus solely on the post-embryonic effects of the mutant (Chalfie et al., 1985). To that end, I used a pmec-4::GFP reporter to visualize the touch neurons in the mutant strain (Hong et al., 2000). What I observed was that while there was no affect on AVM morphology at the permissive temperature, at the non-permissive temperature, there was a significant increase in the number of axon migration defects compared to the wild type AVM neuron (Figure 3.6). These axon migration defects can be rescued by the expression of wildtype unc-54, suggesting that mem-1 does not play a role in axon guidance (Figure 3.6). As    72 unc-54 is a muscle specific myosin, these AVM defects must be an indirect effect of the mutation.  3.5 Discussion  3.5.1 Both unc-54 and mem-1 are required to prevent ectopic membrane extensions Membrane protrusions plays an important role in a number of developmental processes and mutations in genes involved in the regulation of cellular membrane dynamics are associated with human diseases. C. elegans muscle has been shown to be a good model for the study of both wild type and ectopic membrane projections (Dixon and Roy, 2005; Dixon et al., 2006). Here, I have presented a new temperature sensitive allele of unc-54, which encodes a myosin heavy chain B protein that is required for post-embryonic myofilament organization in C. elegans (MacLeod et al., 1981; Waterston et al., 1982; Bejsovec and Anderson, 1988) and an allele of mem-1, a WD motif containing homologous to mammalian WDR6, that both appear to be involved in modulating normal muscle membrane behaviour. While unc-54 has previously been     73   Figure 3.6: AVM guidance defects in the unc-54(t3197) mem-1(t3198) double mutant. Panels A-C are fluorescent images showing the mechanosensory neurons. A is wildtype and B-C are of the unc-54(t3197) mem-1(t3198) double mutant at the non-permissive temperature. The arrows indicate the AVM cell body. Scale bar is 20mm. Panel D is a graph of the percentage of animals exhibiting AVM guidance defects at different temperatures in either the mutant or wild type background. Statistical analysis was done using the Fisher exact test.   shown to be required for the generation of post-embryonic muscle arms (Dixon and Roy, 2005), it has not previous been implicated in ectopic membrane extensions. These unc-54 dependant EMEs are only observed when using a sensitized background (discussed below). Disruption of actin-myosin interactions has been shown to inhibit focal adhesion assembly (Folsom and Sakaguchi,    74 1999; Griffin et al., 2004) and the same phenomenon can be seen for muscle adhesion complexes in unc-54 mutants (Figure 3.4). As integrin adhesions have been shown to be required for the inhibition of ectopic membrane extensions in C. elegans muscle, it is possible that the EMEs observed in the unc-54(e190) null mutants are due to the disruption of the muscle adhesion complexes rather than due to a direct effect of the loss of myosin. I also identified a second gene, mem-1, involved in inhibiting EMEs. This effect of mem-1 is possibly dependant on the loss of unc-54, as RNAi target depletion of mem-1 in a wild type background fails to produce ectopic membrane extension, while depletion in an unc-54(e190) mutant background significantly increases the penetrance of the EME phenotype. In the gene specific rescue experiments in the double mutant, the combined effect of either gene alone was less that what was observed in the double mutant, suggesting that the two genes behave synergistically. A previous study of ectopic membrane extensions in C. elegans found that FGF signaling and adhesion to the ECM were both involved in preventing EME from forming (Dixon et al., 2006). As the predicted human homologue of MEM-1, WDR6, has been shown to be up-regulated in response to the activation of the FGF signalling pathway (Hinsby et al., 2004), it is possible that mem-1 is playing a role downstream of FGF signalling in the worm, but further study is needed. WDR6 has also been shown to interact with the serine/threonine kinase LKB1, which when mutated leads to Peutz?Jeghers syndrome that is associated with increased incidence of malignant cancers (Marignani, 2005; Alessi et al.,    75 2006), and expression of WDR6 enhances the growth suppressing behaviour of LKB1 in tissue culture (Xie et al., 2007). The C. elegans LKB1 homologue, PAR-4, is required for asymmetrical cell division (Watts et al., 2000) and also has been shown to be involved in UNC-40/DCC mediated dendrite outgrowth (Teichmann and Shen, 2011). UNC-40/DCC has been shown to be required for the proper extension of muscle arms in C. elegans and overexpression of UNC-40/DCC results in the generation of ectopic myopodia (Alexander et al., 2009), so there is a possibility that mem-1 plays a role in regulating UNC-40/DCC function. Lastly, WDR6 has been implicated in the insulin-signalling pathway, as its expression is upregulated in response to Insulin-like growth factor-1 and insulin treatment in tissue culture and interacted with IRS-4, an important insulin receptor substrate (Chiba et al., 2009). Insulin-like signalling has been shown to negatively regulate muscle arm extension in C. elegans and mutations that disrupt the insulin-like signalling pathway result in the generation of extra muscle arms, though not EMEs (Dixon et al., 2008). As loss of mem-1 does not generate supernumerary muscle arms, it can?t be a critical component of the pathways, but perhaps its minor perturbation of the Insulin-like signalling pathway is sufficient to generate the phenotype.  It is worth noting that these are not mutually exclusive possibilities and loss of mem-1 might result in disruption of more than one, or possibly all of these signalling pathways. Determining whether mutations in other genes involved in these pathways are also able to exacerbate the unc-54 EME phenotype will go a long in determining which pathway, if any, mem-1 plays a role in.    76 3.5.2 Depletion of components involved in actin remodeling or cell adhesion inhibit ectopic membrane extensions.   In the course of my study, I found that the extension of ectopic muscle membrane processes requires actin remodeling proteins and integrin adhesion complex components. This is un-surprising as both actin remodeling and cell adhesion are well-documented requirements for membrane protrusion formation (For review see Le Clainche and Carlier, 2008; Nurnberg et al., 2011). As both integrin independent (Lammermann et al., 2008; Kardash et al., 2010) and actin independent (Haaf et al., 1998) forms of cell migrations do exist, the observation in this study confirm muscle membrane processes are behaving as traditional migrating membrane protrusions. Interestingly, it has previously been shown that depletion of integrin adhesion components generates EMEs (Dixon et al., 2006), while in my study I found that depleting them reduced the severity of the membrane extension defects. These results suggest that integrin plays a role in preventing the initiation of ectopic membrane extensions, but once they are initiated, it helps to promote membrane extension. Integrin adhesion has been shown to be involved in contact inhibition of migration (Huttenlocher et al., 1998), it is possible that the proper adhesion of muscle cells to the basement membrane prevents ectopic membrane extensions and reduction of that adhesion allows for the processes to form. Such a model is consistent with my observation of unc-54 being involved in inhibit membrane extensions. Loss of unc-54 leads to the disruption of the integrin adhesion complexes, which would then reduce adhesion to the ECM, allowing for EMEs to form.     77 3.5.3 AVM neurons display pathfinding defects in the unc-54(t3197) mem-1(t3198) double mutants.   During my analysis of the unc-54(t3197) mem-1(t3198) double mutant, I observed defects in AVM axon migration. As UNC-54 is a muscle specific myosin (MacLeod et al., 1981; Waterston et al., 1982; Bejsovec and Anderson, 1988), I thought it unlikely that it was responsible for these defect, but expression of wild type UNC-54 was able to rescue the AVM defects, indicating that the mem-1(t3198) mutation is not responsible for this phenotype. Loss of unc-54 results in animals that are substantially thinner than wild type animals (MacLeod et al., 1981; Waterston et al., 1982; Bejsovec and Anderson, 1988, Viveiros Pers. Obs.) and this change in body morphology may be adversely affecting post-embyonically migrating neurons such as AVM (Chalfie et al., 1985). Another possibility is that the EMEs themselves, either physically impede AVM migration, or change the extracellular environment preventing proper pathfinding. Both hypotheses could account for the defects in AVM process guidance, but further study is required to determine which model, if either, is correct. 3.5.4 Expression of a PAT-3/?-integrin transmembrane domain::GFP fusion protein predisposes muscle cells for ectopic membrane extension defects  As a part of my study I found that the expression of the PAT-3/?-integrin transmembrane domain::GFP membrane marker sensitized muscle cells for the generation of EMEs and was sufficient for the production of EMEs (Figure 3.5). While this effect might be a general response of muscle to the presence of a    78 plasma membrane bound marker, ?-integrin?s well know involvement in regulating migratory behaviour suggests that this effect may be specific to PAT-3/?-integrin derived membrane markers. PAT-3/?-integrin is the sole ?-integrin present in the C. elegans (Gettner et al., 1995) and forms heterodimers with one of the two ?-integrins, INA-1, predominantly required for neuronal migrations, or PAT-2, which is necessary for proper muscle adhesion complex formation (Williams and Waterston, 1994; Baum and Garriga, 1997). Mutations that disrupt PAT-3/?-integrin function result in an embryonic arrest phenotype, since proper embryonic elongation requires muscle attachment to the basement membrane (Gettner et al., 1995; Moerman and Williams, 2006). Increased expression of ?-integrins in cancer have been shown to promote membrane protrusions and cell spreading in fibroid cancer cells (Chen et al., 2013) and in ovarian cancer, ectopic expression of ?-integrin has been shown to induce cell invasion (Lau et al., 2012).  In this study I found increased expression of PAT-3/?-integrin in muscle cells is not sufficient to induce membrane protrusions, but it does lead to a predisposition for pro-migratory behavior. Previous studies have shown that in the absence of the extracellular domain, overexpression of the transmembrane domain and cytoplasmic tail of ?-integrin disrupts the normal function of the endogenous ?-integrin in myocytes (Ross et al., 1998; Cheng et al., 2004).  In C. elegans, similar results have been observed with PAT-3/?-integrin truncations. The expression of PAT-3/?-integrin without its extracellular domain acts as a dominant negative and worms    79 expressing it exhibit a variety of PAT-3/?-integrin associated defects (Lee et al., 2001). Lee et al. (2001) also found that deleting the cytoplasmic domain of the truncated PAT-3/?-integrin removed the protein?s dominant negative behaviour and animals were able to express the truncated proteins without any observable effects.  In my study, I found that use of the PAT-3/?-integrin transmembrane domain was sufficient for partial localization of GFP to the adhesion complexes. This construct does not induce the uncoordinated or embryonic lethal phenotypes expected from loss of pat-3 (Gettner et al., 1995; Lee et al., 2001), but can generate ectopic muscle membrane extensions and predisposes the cells for the production of the EMEs. Interactions between the integrin transmembrane domains are important for the regulation of integrin signaling activation and clustering (Luo et al., 2004; Kim et al., 2011; Mehrbod and Mofrad, 2013). As it has been previously shown that the loss of cell adhesion complex proteins can induce ectopic membrane extension defects in muscle cells (Dixon et al., 2006), the presence of the extra PAT-3/?-integrin transmembrane domain may be interacting with the endogenous integrin and inhibiting its wild type function, resulting in the EME formation and sensitization. Since both the functional PAT-3/?-integrin::GFP functional fusion protein and the PAT-3/?-integrin transmembrane domain GFP fusion protein both result in a similar EME phenotype, its possible that the fluorescent protein tags themselves are responsible. The fluorescently tagged muscle proteins have previously been shown to be more sensitive to disruption than their endogenous    80 untagged counterparts (Meissner et al., 2009) and it is possible that the presence of the fluorescent tag in the adhesion complexes might be subtlety destabilizing it, making it more susceptible to disruption. Conversely, this sensitization may not involve integrin at all and simply be a common effect of GFP localized to the muscle cell membrane and as such further analysis using non-integrin derived plasma membrane targeting methods are needed. 3.5.5 Summary  In my study I have identified two genes, unc-54 and mem-1, that work to inhibit EMEs in C. elegans muscle cells and the EMEs that result from the loss of these genes results in have shown that they can be rescued by inhibiting cortical actin remodeling or cell adhesion. My data suggests that the pro-extension behavior of mem-1 may depend on the loss of unc-54 and either be enhancing the EME phenotype of the unc-54 mutation alone, or function in a synergistic way with unc-54. I have also identified a pro-EME sensitization in muscle cells that is conferred by the overexpression of ?-integrin derived membrane localized GFP. The exact cause of sensitivity is unknown, but may be due to destabilization of the integrin adhesion complex. The sensitized nature of these fluorescent markers provides a useful tool for the future identification of other genes involved in mediating muscle membrane extensions.     81 4. Conclusion  In this thesis, C. elegans body wall muscle was investigated at both the embryonic and post-embryonic stages of development and I have identified defects in muscle membrane extensions at both developmental time points. In the embryo I characterized the membrane dynamics of migrating muscle cells and identified a previously unknown anterior migration event that requires long membrane extensions (Viveiros et al., 2011). I have shown that these membrane extensions can be disrupted by mislocalized neurons and that they are required for proper body wall muscle organization. In post-embryonic muscle, I have identified two genes, unc-54/MHCB and mem-1 that are necessary to inhibit ectopic membrane extensions. In addition, I have shown that expression of PAT-3/ ?-integrin bound to GFP confers upon the muscle cells a predisposition for ectopic membrane extensions and that its transmembrane domain is sufficient to induce this sensitization.  4.1 A novel anterior muscle migration event is necessary for proper embryonic muscle morphogenesis in C. elegans.   Previous characterization of C. elegans embryonic muscle migration depended on following single muscle cell nuclei in vivo or using anti-MyoD antibodies to visualize all muscle cells in fixed samples. In my study my use of an embryonically expressed, muscle specific, membrane bound fluorophore allowed for the first time the visualization of these dynamic migrations. Using this marker    82 facilitated my identification of the long membrane extensions that are sent out from the anterior-most pair of cells in each muscle quadrant. These membrane processes are required for anterior muscle migrations that culminate when the muscle quadrants extend the entire length of the developing embryo. Defects in the extension of these muscle processes result in a posterior displacement of the body wall muscle cells leaving no muscle cells at the anterior end of the animal.  4.2 Proper neuron organization is crucial for anterior muscle migration.  In characterizing these membrane processes I identified several signaling pathways that are required for proper extension of the membrane processes from the two ventral embryonic muscle quadrants. Using mutant analysis, I have shown that Ephrin, SAX-3/Robo, and integrin all are required for the proper anterior migration of ventral muscle. I have demonstrated that these genes are not directly required for the muscle migrations.  Rather they are necessary to maintain proper neural embryonic organization, which is essential to maintain the muscle migration pathway intact. My analysis has shown that the absence of these pathways results in a ventral displacement of the embryonic neurons, apparently into the path of the migrating ventral muscle cells and results in the inhibition of the ventral anterior muscle processes. To my knowledge this is the first documentation of the mis-localization of cells of another tissue affecting muscle cell migration and organization in C. elegans.     83 4.3 Embryonic muscle migration in C. elegans is a laminin dependent, but integrin independent process.  Laminin is a well-known substrate for muscle cell migration in a number of biological systems, but until my study, it had not been shown in C. elegans. Using both mutant and RNAi analysis, I have shown that laminin plays a role in all aspects of embryonic muscle migration in C. elegans. Despite the many years of extensive study of C. elegans muscle, this is the first time a gene has been shown to affect muscle cell migration. I have also shown that PAT-3:: ?-integrin, the sole C. elegans ?-integrin, plays no role in embryonic muscle migrations, consistent with previous characterization of pat-3 mutants. While there is some evidence to suggest there might be a second ?-integrin in the worm, null mutations of the downstream components of the C. elegans integrin adhesion complex also fail to disrupt embryonic muscle migrations, which suggests that embryonic muscle cells migrate in an integrin independent manner. While integrin independent migration is not unheard of, to my knowledge this would be the first reported instance of such in muscle cells.  4.4 UNC-54/MHCB and MEM-1 in inhibit ectopic muscle membrane extensions.  The suppression of ectopic membrane extensions in C. elegans has previously been shown to involve the FGF signaling pathway and the integrin adhesion complex. I have identified two other genes that are also necessary for the suppression of these processes, unc-54 and mem-1. I have identified a new    84 temperature sensitive allele of the myosin heavy chain B, UNC-54, and have demonstrated that UNC-54 is was necessary for the prevention of inappropriate membrane extensions. I have also identified a new gene, mem-1 that when mutated also affects the production of these processes. Loss of unc-54/MHCB and mem-1 appear to behave synergistically in the generation of these ectopic processes and that the effect of the loss of mem-1 may be completely dependent on the loss of unc-54.   4.5 Membrane-bound GFP or YFP predisposes muscle cells to ectopic membrane extensions. The ability for ?-integrin expression to affect cell migration is a well-established phenomenon. I have shown that the increased expression of a ?-integrin GFP translational fusion predisposes muscle cells to send out ectopic membrane extensions. I have also shown that expression of the transmembrane domain tagged with a fluorophore is sufficient to confer this sensitivity. These ?-integrin derived markers still show localization to the adhesion complexes and may serve to destabilize the complexes, making them more sensitive to further disruption. Strains carrying these markers are thus sensitized and provide us with a useful tool for the identification of genes involved in suppressing ectopic membrane extension that would otherwise not be detectable.  4.6 Future directions In my analysis of embryonic muscle cell migration in C. elegans I have shown that laminin is required for proper organization of embryonic muscle.    85 While this is the first reported protein to be involved in mediating embryonic muscle migration in C. elegans how muscle cells are interacting with this ECM component remains unknown. The classical receptor for laminin is integrin, but since the loss of C. elegans sole ?-integrin, PAT-3, does not result defects in embryonic muscle migration, a different receptor must be involved. Non-integrin laminin receptors are known and some, such as dystroglycan and the 32/67 kDa laminin receptor, have homologues in C. elegans and are good candidates for further study. I have also identified an anterior migration event that is dependent on the extension of long membrane processes. While I have identified a number of genes that can disrupt the extension of these processes, no genes have yet been identified that are required cell-autonomously for these processes. One of these genes, ina-1, is one of the two ?-integrins found in the worm. Interestingly, while its loss results in defects in ventral muscle membrane extensions, loss of PAT-3/ ?-integrin, as mentioned above, has no defects in muscle membrane extensions. This indicates that there must be a second ?-integrin in the C. elegans genome. The gene C05D9.3 has some sequence similarity to ?-integrins and may prove to be such, but has not been studied in depth. I have identified two genes, unc-54/MHCB and mem-1, involved in inhibiting ectopic muscle membrane extensions. While the loss of UNC-54/MHCB most likely results in ectopic membrane extensions due to destabilization of the integrin adhesions, the function of mem-1 remains unknown. Based on the function of its human homologue, WDR6, mem-1 may play a part in FGF, insulin-   86 like or UNC-40/DCC signalling. All three pathways have been implicated in mediating muscle membrane extensions in C. elegans, but more work is required to determine if mem-1 is involved in one or more of these pathways. WDR6 is expressed ubiquitously and has been shown be present in both the cytoplasm and nucleus. Determining whether MEM-1 shares a similar expression pattern and subcellular localization would lend support that MEM-1 and WDR6 share conserved functions. Also, as mem-1 displays a synergistic effect with UNC-54/MHCB, it would be useful to determine whether this synergism is gene specific, or whether loss of mem-1 is able to exacerbate the effect of other EME generating mutants. Such information would inform the function of the gene and further our understanding of how muscle cells prevent EMEs from forming. In my analysis of muscle plasma membranes, I have identified that ectopic expression of PAT-3/ ??integrin sensitizes muscles for the production ectopic membrane extensions. This sensitization is not dependant on the functionality of the protein, but whether the presence of an attached fluorescent protein is required for this phenomenon still remains to be determined. This type of sensitization has previously been reported for myosin GFP fusions and may represent a common feature of fluorescently tagged proteins in C. elegans (Meissner et al., 2009). 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Mutants Screened Description Embryonic muscle migration emb-9 (hc70) type IV collagen wt let-2 (g25) type IV collagen wt lon-2 (e678) glypican wt nid-1 (cg119) nidogen wt pat-2 (st538) a integrin wt pat-3 (st564) b integrin wt plx-1 (nc36) plexin wt plx-2 (ev773) plexin wt unc-40 (e271) netrin receptor wt clr-1(e1745) FGF negative regulator wt unc-5(e53) netrin receptor wt epi-1 (gm57) laminin a subunit wt gpn-1 (ok377) glypican wt sdn-1(ok449) syndecan wt unc-52(gk3) perlecan wt Legend  bmd body morphology defect ste sterile rup ruptured vulva unc uncoordinated emb embryonic lethality lva larval arrest egl egg laying defective dpy dumpy (body length shorter than wildtype) let lethal n/a wild muscle in larvae and adults early arrest embryos arrest before elongation, muscle not obs.      105 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) H19M22.2 let-805 emb/unc muscle detachment unc/lva muscle detachment F56B3.2  unc muscle detachment molting defect muscle detachment K08C7.3 epi-1 let/emb/bmd muscle disoganized in bmd and late arrest embs bmd/emb muscle disorganized in bmd and late arrest embs W02B9.1 hmr-1 bmd n/a bmd n/a C31G12.2  ste n/a bmd n/a F47C12.2 clec-78 ste n/a dpy n/a F15B9.8  bmd n/a egl n/a C40H5.7  wt n/a  n/a R07G3.9  egl n/a egl n/a R02D3.6 grl-19 ste n/a egl n/a R13F6.4 ten-1 unc n/a egl n/a C08G9.2  emb early arrest emb early arrest K07A12.2  ste n/a emb early arrest C08B11.1 zyg-11 emb early arrest emb early arrest Y49G5A.1  fer n/a fer n/a ZK783.1 fbn-1 lva n/a lva n/a F53G12.3  lva n/a lva n/a M88.6 pan-1 lva n/a lva n/a C49C3.4  lva n/a lva n/a F56C11.1 bli-3 lva n/a lva n/a ZK1058.2 pat-3 lva n/a lva n/a T27E4.5  Po rup n/a Po egl n/a K04B12.1 plx-2 bmd n/a rup n/a B0228.4 tag-308 egl n/a rup n/a B0412.2 daf-7 egl n/a rup n/a AC7.2 soc-2 egl n/a rup n/a F25B4.9 clec-1 lva n/a rup n/a M57.2  rup n/a rup n/a Y71D11A.1  rup n/a rup n/a F32E10.3  rup n/a rup n/a C18E9.7  rup n/a rup n/a Y70G10A.3  ste n/a rup n/a Y73C8B.4 lag-2 ste n/a rup n/a F25H8.3 gon-1 ste n/a rup n/a Y18D10A.12  ste n/a rup n/a    106 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) ZC101.2 unc-52 unc n/a rup n/a T19B4.7 unc-40 unc n/a slow growth n/a K11B4.1  Po dead n/a ste n/a ZK39.7  ste n/a ste n/a F08B4.1 dic-1 ste n/a ste n/a C37C3.6 ppn-1 ste n/a ste n/a C54D1.5  ste n/a ste n/a B0432.12  ste n/a ste n/a H20J04.8 sap-1 ste n/a ste n/a ZK39.5  bmd n/a wt n/a F47A4.2 dpy-22 dpy n/a wt n/a Y38F1A.9 oig-2 dpy n/a wt n/a M01E10.2  dpy n/a wt n/a ZC15.6  dpy/egl n/a wt n/a M03F4.6  egl n/a wt n/a F15G9.4 him-4 egl n/a wt n/a Y69H2.2  egl n/a wt n/a F46B3.9  egl n/a wt n/a ZC84.1  egl n/a wt n/a F49F1.9  egl n/a wt n/a C33D9.2  egl n/a wt n/a F23B2.3  egl n/a wt n/a R09E10.5  egl n/a wt n/a T12A7.2  egl n/a wt n/a C32H11.4  egl n/a wt n/a CD4.9  egl n/a wt n/a F26D11.5  egl n/a wt n/a ZK287.4  egl n/a wt n/a F58E6.3  egl n/a wt n/a F32D8.7  egl n/a wt n/a F57B7.4 mig-17 egl n/a wt n/a C29A12.6  egl n/a wt n/a F58H1.7  egl n/a wt n/a F08H9.6 clec-57 egl n/a wt n/a F40G12.3  egl n/a wt n/a F35E12.7  egl n/a wt n/a F53B1.8  egl n/a wt n/a C25B8.4  egl n/a wt n/a F41D9.3 wrk-1 egl n/a wt n/a F19C6.3  egl n/a wt n/a K09C8.5 pxn-2 egl n/a wt n/a F40E10.1 hch-1 egl n/a wt n/a F52E1.2  egl n/a wt n/a F32D8.13  egl n/a wt n/a    107 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F20B10.2  egl/rup n/a wt n/a K08D8.5  egl/rup n/a wt n/a C32H11.9  egl/rup n/a wt n/a F08G5.7  egl/rup  wt n/a C15H11.6 nxf-2 elg/rup  wt n/a T21E3.3 lrp-2 emb early arrest wt n/a F33G12.4  emb early arrest wt n/a K10D2.1  emb early arrest wt n/a Y55F3BR.2  emb early arrest wt n/a C17D12.6 spe-9 emb early arrest wt n/a Y69H2.11  emb early arrest wt n/a ZC15.2  emb early arrest wt n/a F54E2.2  emb early arrest wt n/a B0205.3 rpn-10 emb early arrest wt n/a T23F4.4 nas-27 emb early arrest wt n/a F44G4.8 dep-1 emb early arrest wt n/a K01C8.8  emb early arrest wt n/a C18H7.1  emb early arrest wt n/a E03H12.2  emb early arrest wt n/a Y9C9A.13  emb early arrest wt n/a T23B12.8  emb early arrest wt n/a T02C5.3 igcm-3 emb early arrest wt n/a K04E7.3 nas-33 emb early arrest wt n/a C32H11.12 dod-24 emb early arrest wt n/a F54D1.6  emb early arrest wt n/a ZK682.5  emb early arrest wt n/a Y47H9C.4 ced-1 emb early arrest wt n/a F28B3.1  emb early arrest wt n/a F32D8.3  fer n/a wt n/a C41H7.7 clec-3 let n/a wt n/a C18F3.2 sax-7 let n/a wt n/a F58G4.4 sdz-23 let n/a wt n/a T09F5.9 clec-47 let n/a wt n/a D1009.3  let n/a wt n/a T01C8.7 mec-4 let n/a wt n/a F54E2.3 ketn-1 let n/a wt n/a C04H5.2  lva n/a wt n/a C25G4.1  lva n/a wt n/a F56A4.1 nas-2 lva/rup n/a wt n/a K06G5.1  let n/a wt n/a E02H4.1 del-1 ste n/a wt n/a F49E12.1  ste n/a wt n/a M162.1  rup n/a wt n/a Y69H2.13  rup n/a wt n/a DC2.3  rup n/a wt n/a    108 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) Y50D4B.5  rup n/a wt n/a Y73C8C.2  rup n/a wt n/a F55H12.3  rup n/a wt n/a Y105E8A.7 lev-10 rup n/a wt n/a F10F2.5  rup n/a wt n/a F48E8.7  rup n/a wt n/a H20E11.1  rup n/a wt n/a T20B3.12 clec-26 rup n/a wt n/a W01A11.4 lec-10 rup n/a wt n/a T07H8.4 mec-1 rup n/a wt n/a F58G4.3  rup n/a wt n/a VC5.2  rup n/a wt n/a C25E10.9 swm-1 rup n/a wt n/a K03B8.5 nas-19 rup n/a wt n/a F22E12.1  rup n/a wt n/a D1054.7  rup n/a wt n/a T04F3.2  rup n/a wt n/a W09D12.1  rup n/a wt n/a C54G7.3 lgx-1 rup n/a wt n/a T07H6.4  rup n/a wt n/a C09C7.1 zig-4 rup n/a wt n/a ZC374.2  rup n/a wt n/a C04A11.4 adm-2 rup n/a wt n/a C11H1.5  rup n/a wt n/a F10G2.3 clec-7 rup n/a wt n/a R151.5 dpy-31 rup n/a wt n/a F36F12.6  rup n/a wt n/a C37H5.9 nas-9 rup n/a wt n/a T22A3.8 lam-3 rup n/a wt n/a T01D3.3  rup n/a wt n/a F41C6.1 unc-6 rup n/a wt n/a F33C8.1 tag-53 rup n/a wt n/a C44H4.2 sym-5 rup n/a wt n/a F33H2.3  rup n/a wt n/a F11D11.1  rup/egl n/a wt n/a R02F11.4  rup/egl n/a wt n/a C10G8.2  rup/egl n/a wt n/a F28D1.8  rup/egl n/a wt n/a F46E10.11  rup/egl n/a wt n/a R107.8 lin-12 ste n/a wt n/a K08E5.3 mua-3 ste n/a wt n/a T11F8.3 rme-2 ste n/a wt n/a B0024.14 crm-1 ste n/a wt n/a F29G6.1  ste n/a wt n/a F15B9.7 cdh-6 ste n/a wt n/a    109 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F56A4.2  ste n/a wt n/a F58E1.8  ste n/a wt n/a W10G11.7  ste n/a wt n/a ZC84.6  ste n/a wt n/a T21D12.7  ste n/a wt n/a W02C12.1  ste n/a wt n/a F38A1.7  ste n/a wt n/a F49F1.10  ste n/a wt n/a C02B10.3  ste n/a wt n/a F56H11.1 fbl-1 ste n/a wt n/a F20B10.1 nlr-1 ste n/a wt n/a C32H11.1  ste n/a wt n/a ZK896.4  ste n/a wt n/a Y46C8AL.4 clec-71 ste n/a wt n/a Y55F3AL.1 plx-1 ste n/a wt n/a Y102A5B.1 clec-27 ste n/a wt n/a F39G3.8 tig-2 ste n/a wt n/a B0222.5  ste n/a wt n/a F58B4.1 nas-31 ste n/a wt n/a F35B12.6 tag-290 ste n/a wt n/a B0365.5  ste n/a wt n/a C54D10.10  ste n/a wt n/a H42K12.3  ste n/a wt n/a R07E4.1  ste n/a wt n/a C52B9.3  ste n/a wt n/a T22E5.3  ste n/a wt n/a F16F9.5 mec-10 ste n/a wt n/a T25C12.3  ste n/a wt n/a C18A11.7 dim-1 ste n/a wt n/a K09E2.1  ste n/a wt n/a M02D8.5  ste n/a wt n/a F38B2.3  ste n/a wt n/a F59F3.1 ver-3 ste n/a wt n/a F16B12.1  ste n/a wt n/a F16B12.2 dsl-4 ste n/a wt n/a F20D1.7  ste n/a wt n/a C27C12.5 tag-324 ste n/a wt n/a T04H1.6 lrx-1 ste n/a wt n/a F09F9.4  ste n/a wt n/a C32H11.10 dod-21 ste n/a wt n/a T07D10.4 clec-15 ste n/a wt n/a T13C2.6  ste n/a wt n/a F45G2.5 bli-5 ste n/a wt n/a T01D3.1  ste n/a wt n/a ZK377.2 sax-3 ste n/a wt n/a    110 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F40F4.6  ste n/a wt n/a F02D10.5 flr-1 ste n/a wt n/a B0034.3 cdh-11 wt n/a   B0207.12 avr-14 wt n/a   B0218.6 clec-51 wt n/a   B0218.8 clec-52 wt n/a   B0244.8 egg-1 wt n/a   B0273.4 unc-5 wt n/a   B0286.2 lat-2 wt n/a   B0365.6 clec-41 wt n/a   B0393.3  wt n/a   B0393.5  wt n/a   B0393.7  wt n/a   B0454.7 clec-2 wt n/a   B0457.1  wt n/a   B0495.10  wt n/a   B0523.5  wt n/a   B0564.9  wt n/a   C01F1.5  wt n/a   C01F6.1  wt n/a   C01G6.8 cam-1 wt n/a   C02B4.1 adt-1 wt n/a   C02F12.5  wt n/a   C02F5.7  wt n/a   C02H7.3 aex-3 wt n/a   C03H5.1 clec-10 wt n/a   C04F6.1 vit-5 wt n/a   C04G6.10  wt n/a   C04G6.7  wt n/a   C05D11.6 nas-4 wt n/a   C05D12.2  wt n/a   C05D9.3  wt n/a   C06A1.6  wt n/a   C06A8.6  wt n/a   C06B8.2  wt n/a   C06B8.7  wt n/a   C07A9.1  wt n/a   C07D10.4 nas-7 wt n/a   C07G1.2  wt n/a   C09D1.1 unc-89 wt n/a   C09D1.2  wt n/a   C09D8.1 ptp-3 wt n/a   C09F9.2  wt n/a   C09G9.3  wt n/a   C10G8.3  wt n/a      111 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) C10G8.4  wt n/a   C11E4.3 tag-263 wt n/a   C11E4.4  wt n/a   C14A6.1 clec-48 wt n/a   C14C11.4  wt n/a   C14F11.2  wt n/a   C14F5.2 zig-3 wt n/a   C15A11.3 sol-1 wt n/a   C16A11.8  wt n/a   C16A3.6  wt n/a   C16B8.4  wt n/a   C16C8.2  wt n/a   C16D9.2 rol-3 wt n/a   C16E9.1  wt n/a   C17G1.6 nas-37 wt n/a   C17H12.6 spe-9 wt n/a   C17H12.8  wt n/a   C18B2.6  wt n/a   C24B9.3  wt n/a   C24F3.3 nas-12 wt n/a   C24G6.2  wt n/a   C24G7.1  wt n/a   C24G7.2  wt n/a   C24G7.4  wt n/a   C25F6.4  wt n/a   C26C6.3 nas-36 wt n/a   C26G2.1 syg-2 wt n/a   C27A2.5  wt n/a   C27B7.7  wt n/a   C27C7.5  wt n/a   C29A12.4 nrx-1 wt n/a   C29F3.4  wt n/a   C29F3.5  wt n/a   C29F3.7  wt n/a   C30H6.1  wt n/a   C30H6.3  wt n/a   C30H6.4  wt n/a   C31B8.8  wt n/a   C32H11.13  wt n/a   C32H11.3  wt n/a   C33A12.12  wt n/a   C33F10.5 rig-6 wt n/a   C34C6.3  wt n/a   C34F6.1  wt n/a   C34F6.10  wt n/a      112 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) C34H3.1 tag-275 wt n/a   C35D10.15 clec-6 wt n/a   C36B1.1 cle-1 wt n/a   C36B7.5  wt n/a   C36B7.7 hen-1 wt n/a   C36F7.4 rig-5 wt n/a   C37C3.7  wt n/a   C39F7.2  wt n/a   C41C4.3  wt n/a   C41C4.5 unc-105 wt n/a   C42D4.11  wt n/a   C43H6.6  wt n/a   C44F1.3 lec-4 wt n/a   C44H4.1  wt n/a   C44H4.3 sym-1 wt n/a   C45G7.5 cdh-10 wt n/a   C46A5.2  wt n/a   C46A5.4  wt n/a   C46A5.9 hcf-1 wt n/a   C47C12.6 deg-1 wt n/a   C48D1.1  wt n/a   C49A1.6  wt n/a   C49C3.11  wt n/a   C49C3.12  wt n/a   C49C3.13  wt n/a   C49C8.5  wt n/a   C50C3.9 unc-36 wt n/a   C50F2.1  wt n/a   C50H2.1 fshr-1 wt n/a   C50H2.1 fshr-1 wt n/a   C50H2.3 mec-9 wt n/a   C53A5.13  wt n/a   C53B7.1 rig-3 wt n/a   C53D6.7  wt n/a   C54C8.7 clec-11 wt n/a   C54D1.2 clec-86 wt n/a   C54E4.2  wt n/a   C54G4.4  wt n/a   C56E6.6  wt n/a   D1044.2  wt n/a   E01G6.1  wt n/a   E03A3.5  wt n/a   E03H4.10 clec-17 wt n/a   EEED8.11  wt n/a   EGAP1.3 zmp-1 wt n/a      113 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F01F1.13  wt n/a   F02A9.6 glp-1 wt n/a   F02G3.1 ncam-1 wt n/a   F02H6.6  wt n/a   F02H6.7  wt n/a   F07A5.1 inx-14 wt n/a   F07C4.2 clec-45 wt n/a   F08B4.2 cdh-5 wt n/a   F08C6.1 adt-2 wt n/a   F08D12.4  wt n/a   F08G5.6  wt n/a   F08H9.5  wt n/a   F08H9.7 clec-56 wt n/a   F08H9.8 clec-54 wt n/a   F08H9.9 clec-55 wt n/a   F09B12.3  wt n/a   F09E8.6 nas-14 wt n/a   F09F3.5  wt n/a   F09G8.5  wt n/a   F09G8.8  wt n/a   F10E7.4  wt n/a   F10F2.4  wt n/a   F10F2.6  wt n/a   F10F2.7  wt n/a   F10F2.8  wt n/a   F11C1.5  wt n/a   F11D11.5  wt n/a   F11D5.3  wt n/a   F13B12.2  wt n/a   F13E9.1  wt n/a   F14B4.1  wt n/a   F14H12.3  wt n/a   F15B9.9 dsl-7 wt n/a   F15D3.2  wt n/a   F16H6.1 clec-42 wt n/a   F17B5.2  wt n/a   F17B5.3  wt n/a   F17B5.5  wt n/a   F17C11.5  wt n/a   F18F11.3  wt n/a   F19B10.7 srx-97 wt n/a   F19F10.6  wt n/a   F19F10.7 srx-97 wt n/a   F20G2.4  wt n/a   F20G2.5  wt n/a      114 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F21C10.7  wt n/a   F21H7.4  wt n/a   F22D3.6  wt n/a   F23H12.5  wt n/a   F25D1.4  wt n/a   F25D7.5  wt n/a   F25F2.2 cdh-4 wt n/a   F26A1.11  wt n/a   F26A1.12  wt n/a   F26A1.8  wt n/a   F26C11.3  wt n/a   F26D10.12  wt n/a   F26D11.11 let-413 wt n/a   F26D11.9  wt n/a   F26D2.12  wt n/a   F26F2.6  wt n/a   F27D9.7  wt n/a   F28A12.1  wt n/a   F28B4.3  wt n/a   F28C1.3  wt n/a   F28E10.2  wt n/a   F28H6.7  wt n/a   F29D10.2  wt n/a   F29D11.1 lrp-1 wt n/a   F30H5.3  wt n/a   F31A9.3 arg-1 wt n/a   F31D4.4  wt n/a   F31D5.3 tag-149 wt n/a   F31F7.2  wt n/a   F32A5.2  wt n/a   F32A7.3 tag-271 wt n/a   F33E2.3  wt n/a   F35A5.3 abu-10 wt n/a   F35B12.4  wt n/a   F35C5.6 clec-62 wt n/a   F35C5.7 clec-64 wt n/a   F35C5.8 clec-65 wt n/a   F35C5.9 clec-66 wt n/a   F35D11.10  wt n/a   F35D11.8  wt n/a   F35D2.3  wt n/a   F35E12.10  wt n/a   F35E12.6  wt n/a   F35E12.8  wt n/a   F36A2.4 twk-30 wt n/a      115 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F36D3.10  wt n/a   F36F12.5  wt n/a   F36G9.11  wt n/a   F36H1.4 lin-3 wt n/a   F36H2.3  wt n/a   F36H5.6 clec-19 wt n/a   F37E3.2  wt n/a   F38A1.1  wt n/a   F38A1.10  wt n/a   F38A1.4  wt n/a   F38A1.5  wt n/a   F38A5.3 lec-11 wt n/a   F38C2.6  wt n/a   F40E10.4 slt-1 wt n/a   F40G9.10  wt n/a   F40H6.5  wt n/a   F41D3.6  wt n/a   F41E6.6  wt n/a   F41G3.12  wt n/a   F42A10.8 nas-28 wt n/a   F42F12.2 zig-2 wt n/a   F43C1.1  wt n/a   F43G6.3  wt n/a   F44E2.4  wt n/a   F44E5.2  wt n/a   F44E5.3  wt n/a   F46A8.3  wt n/a   F46A8.5  wt n/a   F46B3.14  wt n/a   F46C5.3 nas-25 wt n/a   F46C8.1  wt n/a   F47B3.8 tag-162 wt n/a   F47C12.1  wt n/a   F47D12.5  wt n/a   F47F2.3  wt n/a   F47F6.5  wt n/a   F47G4.1  wt n/a   F48C11.2  wt n/a   F48C5.1  wt n/a   F48E3.8  wt n/a   F49A5.2 clec-31 wt n/a   F49A5.3 clec-22 wt n/a   F49A5.4 clec-24 wt n/a   F49A5.5 clec-28 wt n/a   F49A5.7 clec-33 wt n/a      116 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) F49A5.9 clec-32 wt n/a   F49F1.11  wt n/a   F49H6.1  wt n/a   F52C6.14  wt n/a   F52C9.5  wt n/a   F52H3.7 lec-2 wt n/a   F53B6.2  wt n/a   F53B7.4  wt n/a   F53B7.5  wt n/a   F53C11.6 twk-32 wt n/a   F53F4.8  wt n/a   F53G2.1  wt n/a   F54D7.4 zig-7 wt n/a   F54F3.1 nid-1 wt n/a   F55C7.7 unc-73 wt n/a   F55D12.5  wt n/a   F55F10.1  wt n/a   F55G1.12  wt n/a   F55G1.13  wt n/a   F55G11.5 dod-22 wt n/a   F56A8.3  wt n/a   F56D1.4 clr-1 wt n/a   F56D6.1 clec-68 wt n/a   F56F10.4  wt n/a   F56H6.8 clec-18 wt n/a   F57C12.1 nas-38 wt n/a   F58A3.2 egl-15 wt n/a   F58A4.5  wt n/a   F58B3.8 dsl-5 wt n/a   F58D2.1  wt n/a   F58F9.6  wt n/a   F59A6.3  wt n/a   F59D8.2 vit-4 wt n/a   F59F3.4  wt n/a   F59F3.5 ver-4 wt n/a   H02I12.4 dsl-6 wt n/a   H02K04.1  wt n/a   H02K04.2  wt n/a   H16D19.1 clec-13 wt n/a   H20E11.2  wt n/a   H36L18.1  wt n/a   H43E16.1  wt n/a   K02E10.8  wt n/a   K02F3.5  wt n/a   K03A1.2  wt n/a      117 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) K03B8.1 nas-16 wt n/a   K03B8.2 nas-17 wt n/a   K03B8.3 nas-18 wt n/a   K03B8.6  wt n/a   K03D10.1 kal-1 wt n/a   K03E5.1  wt n/a   K03H1.5  wt n/a   K03H6.4  wt n/a   K04D7.4  wt n/a   K04H8.1  wt n/a   K05C4.11  wt n/a   K05C4.9  wt n/a   K06A9.3  wt n/a   K07D8.1 mup-4 wt n/a   K07E12.1  wt n/a   K07G5.1  wt n/a   K07H8.1 vit-6 wt n/a   K08B4.6 cli-2 wt n/a   K08D8.3  wt n/a   K08D8.6  wt n/a   K08E7.8  wt n/a   K09C8.3 nas-10 wt n/a   K09F5.2 vit-1 wt n/a   K10B2.3  wt n/a   K10B4.1  wt n/a   K10C3.3 zig-1 wt n/a   K10D11.3  wt n/a   K10D11.5  wt n/a   K10D3.4  wt n/a   K11D12.6  wt n/a   K11G12.1 nas-11 wt n/a   M02D8.1  wt n/a   M02F4.7  wt n/a   M02G9.3  wt n/a   M03A1.1 vab-1 wt n/a   M142.2 cut-6 wt n/a   M199.4  wt n/a   R01B10.1 cli-1 wt n/a   R01H2.3 egg-2 wt n/a   R02F11.2  wt n/a   R05A10.5  wt n/a   R05F9.12  wt n/a   R05H10.6 cdh-7 wt n/a   R06C7.4  wt n/a   R07A4.4  wt n/a      118 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) R07B1.10 lec-8 wt n/a   R07B1.2 lec-7 wt n/a   R07C3.1 clec-43 wt n/a   R07C3.12 clec-44 wt n/a   R08C7.6  wt n/a   R08F11.7  wt n/a   R10E8.2  wt n/a   R10F2.1  wt n/a   R10H10.3  wt n/a   R11G1.1  wt n/a   R12A1.3  wt n/a   R13A1.4 unc-8 wt n/a   R13F6.8  wt n/a   R193.2  wt n/a   T01B7.8  wt n/a   T01D3.6  wt n/a   T01E8.1  wt n/a   T01G9.3  wt n/a   T02B11.7 nas-32 wt n/a   T02G6.7  wt n/a   T03F1.10 clec-53 wt n/a   T03G11.8 zig-6 wt n/a   T04A11.3  wt n/a   T04A8.3  wt n/a   T04G9.2 nas-15 wt n/a   T05A7.3  wt n/a   T05E12.6  wt n/a   T05G5.1  wt n/a   T05H4.3  wt n/a   T06D8.10  wt n/a   T06E6.10  wt n/a   T07H3.4 clec-21 wt n/a   T07H6.5  wt n/a   T08G3.4  wt n/a   T09A5.9  wt n/a   T09D3.3  wt n/a   T10H9.2 scd-2 wt n/a   T11F9.3 nas-20 wt n/a   T11F9.5 nas-21 wt n/a   T11F9.8  wt n/a   T14E8.2  wt n/a   T15B7.1  wt n/a   T16H12.4  wt n/a   T17A3.1 ver-1 wt n/a   T17A3.10  wt n/a      119 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) T19C9.6  wt n/a   T19D12.4  wt n/a   T19D12.6  wt n/a   T19D12.7  wt n/a   T19D2.1  wt n/a   T19E7.1  wt n/a   T19H12.2  wt n/a   T20B3.13 clec-40 wt n/a   T20D3.1  wt n/a   T21B6.3  wt n/a   T21D11.1  wt n/a   T21D12.12  wt n/a   T21D12.9  wt n/a   T22A3.6  wt n/a   T22E7.1  wt n/a   T22F7.3  wt n/a   T22H2.6  wt n/a   T23G11.6  wt n/a   T23H4.3 nas-5 wt n/a   T24A11.3 toh-1 wt n/a   T24F1.6 tag-180 wt n/a   T25D10.2  wt n/a   T25E12.10 clec-38 wt n/a   T25E12.7 clec-39 wt n/a   T25E12.8 clec-34 wt n/a   T25E12.9 clec-29 wt n/a   T25F10.2 dbl-1 wt n/a   T26E3.1  wt n/a   T27A1.4  wt n/a   T27C10.6 lrk-1 wt n/a   T27D12.3  wt n/a   T27F6.2 clec-12 wt n/a   T28B8.5  wt n/a   T28C12.6  wt n/a   T28D9.7  wt n/a   T28F3.1  wt n/a   T28F4.2 asic-2 wt n/a   W01F3.3 mit-11 wt n/a   W03D8.6 itx-1 wt n/a   W04E12.6 clec-49 wt n/a   W04H10.4  wt n/a   W05B2.2  wt n/a   W06H8.8 tag-58 wt n/a   W09D10.5  wt n/a   W09G10.5  wt n/a      120 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) W09G10.6  wt n/a   W09G12.4 dsl-1 wt n/a   W10G11.11  wt n/a   W10G11.12  wt n/a   W10G11.13  wt n/a   W10G11.14  wt n/a   W10G11.15  wt n/a   W10G11.6  wt n/a   Y102A5B.2 clec-35 wt n/a   Y102A5B.3 clec-37 wt n/a   Y102A5C.16  wt n/a   Y102A5C.17  wt n/a   Y113G7A.5  wt n/a   Y116A8A.1  wt n/a   Y116A8A.3  wt n/a   Y116A8A.8  wt n/a   Y116A8C.21  wt n/a   Y116F11B.7  wt n/a   Y18D10A.10  wt n/a   Y25C1A.1  wt n/a   Y25C1A.4  wt n/a   Y26D4A.12  wt n/a   Y26D4A.3  wt n/a   Y26D4A.4  wt n/a   Y26D4A.6  wt n/a   Y37D8A.13 unc-71 wt n/a   Y37D8A.2  wt n/a   Y37E11AL.6  wt n/a   Y38E10A.4 clec-8 wt n/a   Y38E10A.5 clec-4 wt n/a   Y38H6C.8  wt n/a   Y39A1B.1  wt n/a   Y39A3CL.6 pvf-1 wt n/a   Y39B6A.8  wt n/a   Y39E4B.8 zig-8 wt n/a   Y39G10AR.5  wt n/a   Y41D4B.10 dsl-3 wt n/a   Y42G9A.3  wt n/a   Y43C5A.2  wt n/a   Y43F8B.3  wt n/a   Y46C8AL.1 clec-73 wt n/a   Y46C8AL.5 clec-72 wt n/a   Y46C8AL.8 clec-74 wt n/a   Y46C8AL.9 clec-75 wt n/a   Y46C8AR.1 clec-76 wt n/a      121 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) Y46C8AR.3 clec-77 wt n/a   Y46G5A.29  wt n/a   Y46H3B.2  wt n/a   Y48A6A.1 zig-5 wt n/a   Y48G8AL.1  wt n/a   Y50D4C.2  wt n/a   Y50E8A.3 oig-3 wt n/a   Y51A2A.1  wt n/a   Y51A2A.11  wt n/a   Y52B11A.5  wt n/a   Y53H1A.3  wt n/a   Y54E10A.6  wt n/a   Y54E5B.1 smp-1 wt n/a   Y55B1AR.1 lec-6 wt n/a   Y55D5A.5 daf-2 wt n/a   Y55F3C.5  wt n/a   Y64G10A.1  wt n/a   Y64G10A.7  wt n/a   Y68A4B.1  wt n/a   Y68A4B.2  wt n/a   Y69H2.10  wt n/a   Y69H2.12 tag-336 wt n/a   Y69H2.3  wt n/a   Y70C5C.2 clec-9 wt n/a   Y70C5C.5  wt n/a   Y70G10A.2  wt n/a   Y71A12B.12  wt n/a   Y71F9B.8  wt n/a   Y87G2A.2  wt n/a   Y8A9A.2  wt n/a   Y95B8A.1 nas-30 wt n/a   ZC101.1  wt n/a   ZC13.4 mab-7 wt n/a   ZC262.3  wt n/a   ZC317.3 glc-3 wt n/a   ZC518.3 ccr-4 wt n/a   ZK1037.6  wt n/a   ZK112.7 cdh-3 wt n/a   ZK1193.2  wt n/a   ZK1248.16 lec-5 wt n/a   ZK154.7 adm-4 wt n/a   ZK39.2  wt n/a   ZK39.3  wt n/a   ZK39.6  wt n/a   ZK39.8  wt n/a      122 Gene locus Initial Screen (gross morphology) Muscle defect (GFP Marked) Rescreen (gross morphology) Muscle (GFP Marked) ZK430.8  wt n/a   ZK546.2  wt n/a   ZK617.1 unc-22 wt n/a   ZK666.3 clec-58 wt n/a   ZK666.6 clec-60 wt n/a   ZK673.9  wt n/a   ZK770.1 asic-1 wt n/a   ZK892.1 lec-3 wt n/a   ZK896.3  wt n/a   ZK896.5  wt n/a   ZK896.6  wt n/a   ZK896.7  wt n/a   ZK994.3 pxn-1 wt n/a   ZK994.4  wt n/a       123 Appendix B: Chapter 3 supplementary data   Figure B.1: SNP mapping of the causative allele of GE6583 indicates it is located on the right arm of chromosome 1. Figure represents the relative signal of N2 vs the Hawaiian wild type isolate strain. Each circle represents an individual SNP. Each colour represents a different chromosome. The positive spike at the end of the red section indicates a region linked to the causative alleles of GE6583 and represents the right arm of chromosome 1.         0 1000 2000 3000 40000.00.51.01.5Average of 4 hybridisations with Hawaiian as referenceSNP indexMapping Signal   124 Table B.1: RNAi targets that failed to rescue ectopic membrane extensions in the unc-54(t3187) mem-1(t3198) double mutant. RNAi Target Percentage with severe ectopic membrane extension# p value L4440 no insert control 89  ced-10/Rac1 89 n.s uig-1 80 n.s. pat-3/?-integrin 79 n.s unc-73/Trio 78 n.s unc-53/NAV1 91 n.s wsp-1/WASP 78 n.s arx-5/ARP2/3 protein 83 n.s #Severe refers to EMEs that extend the from one muscle quadrant to another e.g. Figure 3.1 panel D    n=100, n.s.=not significant  

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