Medicine, Faculty of
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UBCV
Li-Leger, Erica
Master of Science - MSc
University of British Columbia
It has been estimated that 15-30% of the ~20,000 genes in C. elegans are essential, yet many of these genes remain to be identified or characterized. With the goal of identifying unknown essential genes, we performed whole genome sequencing on complementation pairs from legacy collections of maternal-effect lethal and sterile mutants. This approach uncovered maternal genes required for embryonic development and genes with sperm-specific functions. In total, 59 essential genes were identified on chromosomes III, IV, and V, and 20 of these genes (42 alleles) were selected for further bioinformatic and functional characterization. The terminal phenotypes of embryos were examined, revealing defects in cell division, morphogenesis, and osmotic integrity of the eggshell. Mating assays with wild-type males revealed previously unknown sperm-expressed genes required for fertilization and embryonic development. The result of this study is a catalogue of mutant alleles in essential genes that will serve as a resource to guide further study toward a more complete understanding of this important model organism. As many genes and developmental pathways in C. elegans are conserved and essential genes are often linked to human disease, uncovering the function of these genes may also provide insight to further our understanding of human biology.
IDENTIFICATION OF ESSENTIAL GENES IN C. ELEGANS THROUGH WHOLE GENOME SEQUENCING OF LEGACY MUTANT COLLECTIONS by Erica Li-Leger B.Sc., The University of British Columbia, 2016 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Cell and Developmental Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) June 2020 © Erica Li-Leger, 2020 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, a thesis entitled: Identification of essential genes in C. elegans through whole genome sequencing of legacy mutant collections submitted by Erica Li-Leger in partial fulfillment of the requirements for the degree of Master of Science in Cell and Developmental Biology Examining Committee: Dr. Don Moerman, Professor, Department of Zoology, UBC Supervisor Dr. Kota Mizumoto, Assistant Professor, Department of Zoology, UBC Supervisory Committee Member Dr. Harald Hutter, Professor, Department of Biological Sciences, Simon Fraser University Supervisory Committee Member Dr. Elizabeth Rideout, Assistant Professor, Department of Cellular and Physiological Sciences, UBC Additional Examiner iii Abstract It has been estimated that 15-30% of the ~20,000 genes in C. elegans are essential, yet many of these genes remain to be identified or characterized. With the goal of identifying unknown essential genes, we performed whole genome sequencing on complementation pairs from legacy collections of maternal-effect lethal and sterile mutants. This approach uncovered maternal genes required for embryonic development and genes with sperm-specific functions. In total, 59 essential genes were identified on chromosomes III, IV, and V, and 20 of these genes (42 alleles) were selected for further bioinformatic and functional characterization. The terminal phenotypes of embryos were examined, revealing defects in cell division, morphogenesis, and osmotic integrity of the eggshell. Mating assays with wild-type males revealed previously unknown sperm-expressed genes required for fertilization and embryonic development. The result of this study is a catalogue of mutant alleles in essential genes that will serve as a resource to guide further study toward a more complete understanding of this important model organism. As many genes and developmental pathways in C. elegans are conserved and essential genes are often linked to human disease, uncovering the function of these genes may also provide insight to further our understanding of human biology. iv Lay Summary An organismâs genes contain the molecular instructions for life. Some genes are absolutely necessary for the organism to develop, survive, and reproduce; these are termed essential genes. Caenorhabditis elegans is a simple roundworm that is widely studied in developmental biology. While this organism has been investigated for decades, there are still essential genes in C. elegans that remain unidentified. This study brings together DNA sequencing technology with decades-old genetic studies in order to identify new essential genes in C. elegans. By identifying these genes and examining their roles in reproduction and development, this work offers a resource for further research aimed at fully understanding the genetic basis of life in this organism. As many genes in C. elegans are closely related to human genes, this will not only help answer fundamental questions in roundworm biology but also has implications for understanding human development and disease. v Preface This study was carried out in the laboratory of Dr. Don Moerman, who provided direction for the project. Input was also provided by committee members Dr. Kota Mizumoto and Dr. Harald Hutter. Laboratory work, including DNA extractions, genome engineering, nematode assays, genetic crosses, and microscopy were carried out by Erica Li-Leger. Analysis of sequencing data was conducted with the help of Dr. Stephane Flibotte. The mutant strains examined in this study were originally isolated and mapped by our collaborators, Dr. Richard Feichtinger, Dr. Heinke Holzkamp, and Dr. Ralf Schnabel. A publication of this work is currently being prepared for journal submission. Special Note: Some of the work pertaining to this thesis was cut short due to curtailment of research at the University of British Columbia during the COVID-19 pandemic. Experiments that are in progress will be completed when the lab reopens but were necessarily omitted from the thesis. vi Table of Contents Abstract .............................................................................................................................................. iii Lay Summary ...................................................................................................................................... iv Preface ................................................................................................................................................ v Table of Contents ............................................................................................................................... vi List of Tables ...................................................................................................................................... ix List of Figures ...................................................................................................................................... x List of Abbreviations .......................................................................................................................... xi Acknowledgements .......................................................................................................................... xiii Chapter 1: Introduction ..................................................................................................................... 1 1.1 Essential Genes and Model Organisms .............................................................................. 1 1.2 C. elegans: A Powerful Model for Developmental Biology ................................................ 4 1.2.1 Investigating Essential Genes in C. elegans ............................................................... 5 1.2.2 Essential Gene Discovery in C. elegans ...................................................................... 7 1.2.3 Whole Genome Sequencing as a Tool for Gene Identification ................................. 9 1.3 Thesis Objectives ............................................................................................................. 10 Chapter 2: Materials and Methods ................................................................................................. 12 2.1 Mutant Collections .......................................................................................................... 12 2.1.1 List of Strains ........................................................................................................... 13 2.2 DNA Extraction ................................................................................................................ 14 vii 2.3 Whole Genome Sequencing and Analysis ....................................................................... 15 2.4 Validation of Gene Identities ........................................................................................... 17 2.4.1 Corroboration with Mapping Data .......................................................................... 17 2.4.2 Complementation Tests .......................................................................................... 17 2.4.3 CRISPR-Cas9 Deletions ............................................................................................ 18 2.5 Bioinformatic Analysis ..................................................................................................... 20 2.6 Temperature Sensitivity and Mating Assays ................................................................... 21 2.7 Microscopy ...................................................................................................................... 22 Chapter 3: Results ............................................................................................................................ 23 3.1 Identification of 59 Essential Genes through Whole Genome Sequencing ................... 23 3.2 Validation of Candidate Gene Identities ......................................................................... 37 3.2.1 Inadvertent Inclusion of a Test Set Validates the Analysis Approach .................... 41 3.3 Bioinformatic Analysis: Human Orthologs, Gene Function and Expression Patterns .... 42 3.4 Temperature Sensitivity & Mating Assays ...................................................................... 46 3.5 Terminal Phenotypes of Maternal-Effect Lethal Embryos ............................................. 48 Chapter 4: Discussion ...................................................................................................................... 50 4.1 Revisiting Legacy Mutant Collections with Whole Genome Sequencing ....................... 50 4.2 Complications with Gene Identification .......................................................................... 51 4.3 Insights from Bioinformatic Analyses .............................................................................. 53 4.3.1 Molecular Nature of the Genetic Variants ............................................................. 53 4.3.2 Gene Ontology Analysis Reveals Common Themes and Gaps in Our Knowledge . 53 viii 4.3.3 Expression Patterns Offer Insight into Gene Function ........................................... 54 4.4 Relevance Beyond C. elegans .......................................................................................... 55 4.5 Discovery of Genes Involved in Male Fertility ................................................................. 57 4.6 Interpreting Terminal Phenotypes of Maternal-Effect Lethal Mutants ......................... 59 Chapter 5: Conclusion ...................................................................................................................... 62 References ....................................................................................................................................... 63 Appendices ....................................................................................................................................... 77 Appendix A Flanking sequences for genetic variants .................................................................. 77 Appendix B Gene Ontology terms and associated genes ........................................................... 82 Appendix C Life stage-specific expression patterns .................................................................... 84 Appendix D Terminal phenotypes of maternal-effect lethal embryos ....................................... 91 ix List of Tables Table 2.1 Summary of mutant collections ...................................................................................... 12 Table 3.1 List of 59 essential genes with associated maternal-effect lethal or sterile alleles ....... 27 Table 3.2 Genes of interest and associated phenotypes ................................................................ 34 Table 3.3 Complementation tests for conflicting groups ............................................................... 40 Table 3.4 Putative sperm-expressed genes .................................................................................... 47 x List of Figures Figure 1.1 Genetics of maternal-effect lethal mutants ..................................................................... 6 Figure 2.1 Schematic of CRISPR-Cas9 gene deletion strategy ........................................................ 19 Figure 3.1 Gene identification in complementation groups from two collections ........................ 24 Figure 3.2 Schematic of gene assignments and deficiency mapping ............................................. 38 Figure 3.3 Complementation tests for the strains in Table 3.3 ...................................................... 40 Figure 3.4 Top GO terms represented in the set of 59 genes ........................................................ 44 Figure 3.5 Examples of characteristic expression patterns ............................................................ 46 Figure 3.6 Embryonic arrest visualized with DIC microscopy for select maternal-effect lethal mutants ............................................................................................................................................ 49 xi List of Abbreviations ATP Adenosine triphosphate BLASTp Basic Local Alignment Search Tool (proteins) BP Biological process (GO term) BWA Burrows-Wheeler Aligner CRISPR Clustered regularly interspaced short palindromic repeats DIC Differential interference contrast Dpy Dumpy dsDNA Double-stranded DNA EDTA Ethylenediaminetetraacetic acid EMS Ethyl methanesulfonate GFP Green fluorescent protein GO Gene ontology GOI Genes of interest IDT Integrated DNA Technologies IEM Inborn errors of metabolism indel Small insertion or deletion M9+TX M9 buffer with 0.5% Triton X-100 MF Molecular function (GO term) modENCODE Model Organism Encyclopedia of DNA Elements NCBI National Center for Biotechnology Information NGM Nematode growth medium NGS Next-generation sequencing OID Osmotic integrity defective OMIM Online Mendelian Inheritance in Man PCR Polymerase chain reaction RNAi RNA interference RNP Ribonucleoprotein SNV Single nucleotide variant xii tracrRNA Trans-activating CRISPR RNA ts Temperature-sensitive Unc Uncoordinated WGS Whole genome sequencing WT Wild type xiii Acknowledgements I offer my sincere gratitude to Dr. Don Moerman for motivating me to pursue this project, guiding me through it, and providing me with opportunities to grow and learn. I also thank Dr. Ralf Schnabel for inspiring this study and offering materials for collaboration, along with Dr. Heinke Holzkamp and Dr. Richard Feichtinger. I am grateful to my committee members, Dr. Kota Mizumoto and Dr. Harald Hutter, for their guidance and encouragement throughout this process. I also owe many thanks to Dr. Stephane Flibotte, whose contribution of data analysis and guidance throughout the course of the project was indispensable. I would like to acknowledge the support of my fellow lab members; in particular, Mark Edgley, whose aptitude for problem solving and expertise in worm genetics was a remarkable resource to have on hand; Vinci Au, for talking through problems with me and offering advice; Lisa Fernando, for her diligent work in supplying the reagents and materials necessary for this project; Kiana Martin and Claudia Doell for their assistance and encouragement; and finally, Negin Khosravi, whose enthusiastic and inquisitive nature kept me motivated and energized to complete this work. xiv Two financial awards that made this work possible were graciously provided by the Canadian Institutes of Health Research and the UBC Faculty of Medicine. Special thanks are owed to my family for their enduring support and encouragement, and to Dr. Robin van der Lee for his sage advice, emotional support, and persistent motivation that kept me moving forward.1 Chapter 1: Introduction 1.1 Essential Genes and Model Organisms Essential genes are those required for the survival or reproduction of an organism, and therefore encode elements that are foundational to life. This class of genes has been widely studied for a number of reasons. Essential genes are often well conserved and can offer insight into the principles that govern biological processes that are common across species (Chervitz et al. 1998; Hughes 2002; Jordan et al. 2002; Georgi et al. 2013). Researching these genes and their functions has important implications in understanding the cellular and developmental processes that form complex organisms, including humans. Additionally, identifying genes that are lethal when mutated opens up new avenues through which drug development approaches can target parasites, pathogens, and cancer cells (for example, Doyle et al. 2010; Shi et al. 2015; Vyas et al. 2015; Zhang et al. 2018). Finally, the concept of a minimal gene set that is comprised of all genes necessary for life has been the subject of much investigation and has recently been of particular interest in the field of synthetic biology (reviewed in Ausländer et al. 2017). While valuable insights can be gained from investigating essential gene function, studying this class of genes in higher organisms such as humans is complicated by a number of factors. In addition to the practical limitations of longer generation times and complex developmental progression, there are obvious ethical considerations that preclude the possibility of doing most experiments on essential genes in humans. For these reasons, model organisms such as yeast (Saccharomyces cerevisiae), nematodes (Caenorhabditis elegans), fruit flies (Drosophila 2 melanogaster), zebrafish (Danio rerio) and mice (Mus musculus) have played key roles in identifying essential genes and understanding the biological processes they control. D. melanogaster was adopted early on as a model system in developmental biology (Poulson 1937; Hadorn 1951). Several decades later, classical genetic screens in flies uncovered genes required for oogenesis and embryonic development (for example, Gans et al. 1975; Mohler 1977; Nüsslein-Volhard and Wieschaus 1980; Jürgens et al. 1984; Nüsslein-Volhard et al. 1984, 1987; Wieschaus et al. 1984; Schüpbach and Wieschaus 1986, 1989). This led to breakthrough discoveries about how genes control embryonic patterning and differentiation (Lewis 1978; Nüsslein-Volhard and Wieschaus 1980). Later, large-scale studies using transposable elements for insertional mutagenesis uncovered hundreds of essential genes by screening for lethal mutations on particular chromosomes (Deák et al. 1997; Salzberg et al. 1997; Bourbon et al. 2002; Peter et al. 2002; Oh et al. 2003). Importantly, many of the genes that were found to be critical for fly development are conserved across eukaryotes (Rubin et al. 2000) and have informed our understanding of human development and disease (Reiter et al. 2001); for example, Notch (Wharton et al. 1985; Louvi and Artavanis-Tsakonas 2012), Wnt (Nüsslein-Volhard et al. 1984; Baker 1987; Clevers and Nusse 2012), and Hox genes (Lewis 1978; Kaufman et al. 1980; McGinnis et al. 1984; Krumlauf 1994; Shah and Sukumar 2010). The biological importance of essential genes has inspired many efforts to identify all such genes in a given model organism. The first systematic genome-wide studies of gene function were done in S. cerevisiae (Winzeler et al. 1999; Giaever et al. 2002). These studies and subsequent 3 experiments have uncovered more than 1,100 essential genes, many of which have conserved roles in fundamental biological processes such as cell division, protein synthesis and metabolism (Winzeler et al. 1999; Giaever et al. 2002; Yu et al. 2006; Li et al. 2011). Due to efficient homologous recombination and a relatively small genome size (~6,000 genes), S. cerevisiae researchers have been able to use a targeted, systematic approach to analyze deletion mutants for every gene in the genome. In more complex model organisms, identifying all essential genes in the genome has not been as straightforward. The discovery of RNA interference (RNAi; Fire et al. 1998), however, accelerated this work by enabling researchers to employ genome-wide reverse genetic screens to examine the phenotypic effects of gene knockdown. For example, an RNAi experiment targeting 86% of genes in C. elegans found 1,170 genes that resulted in a lethal or sterile phenotype when knocked down (Kamath et al. 2003). A similar screen in D. melanogaster found that approximately 30% of the tested genes were essential for viability (Dietzl et al. 2007). In general, this has been an effective, high-throughput method for identifying many genes with essential functions, including gene paralogs with redundant functions that are masked in single gene mutants. However, there are some limitations to using RNAi to screen for all essential genes. Firstly, it is possible to observe off-target effects when double-stranded RNAs contain short stretches of sequence similarity with other genes in the genome (Jackson et al. 2003; Birmingham et al. 2006; Kulkarni et al. 2006; Ma et al. 2006). Furthermore, RNAi can knock down the function of a gene but does not necessarily recapitulate the phenotype of a complete gene knockout, or null allele (for example, Chen et al. 2006). Finally, genes expressed in certain cell 4 types, including neurons, sperm, pharyngeal and vulval muscle, can be resistant to RNAi (Fire et al. 1998; Fraser et al. 2000; Tavernarakis et al. 2000; Timmons et al. 2001; Kamath et al. 2003; Kennedy et al. 2004; del Castillo-Olivares et al. 2009; Kumsta and Hansen 2012; Shiu and Hunter 2017); thus, many genes of biological importance will escape identification in high-throughput RNAi screens. This highlights the importance of striving to obtain null alleles for every gene in the genome, which has been the goal of several model organism consortia (C. elegans Deletion Mutant Consortium 2012; Bradley et al. 2012; Varshney et al. 2013), though it has not yet been achieved for any metazoan. 1.2 C. elegans: A Powerful Model for Developmental Biology C. elegans is one of the most commonly used model organisms in developmental biology. This simple, genetically tractable nematode is well suited to developmental biology for a number of reasons, including its short generation time, transparent body, and fully mapped invariant cell lineage (Brenner 1974; Sulston and Horvitz 1977; Sulston et al. 1983). Importantly, the ability to freeze and store populations of these animals indefinitely has helped foster a collaborative research community in which investigators share strains through a centralized stock center (Caenorhabditis Genetics Center; cgc.umn.edu). In addition, WormBase (wormbase.org; Harris et al. 2020) serves as an invaluable online repository for information of all kinds to be made freely available to the community. As the first metazoan to have its entire genome sequenced (The C. elegans Sequencing Consortium 1998), C. elegans has become a valuable system in which to interrogate gene 5 function in animals. This has led to important contributions to developmental biology, including the discovery of the first axon guidance genes (Hedgecock et al. 1987; 1990; Ishii et al. 1992); the elucidation of the role of Notch signaling in embryonic patterning (Greenwald et al. 1983; Priess et al. 1987; Hutter and Schnabel, 1994; Mango et al. 1994; Mello et al. 1994; Moskowitz et al. 1994); the discovery of the par genes, required for partitioning cellular components in embryos (Kemphues et al. 1988a; Kirby et al. 1990; Morton et al. 1992; Cheng et al. 1995); and the identification of genes controlling programmed cell death (Hedgecock et al. 1983; Ellis and Horvitz 1986; Yuan and Horvitz 1992; Yuan et al. 1993). While much has been learned about gene function in C. elegans over decades of research, there are still many unknowns about the ~20,000 genes in this organism. 1.2.1 Investigating Essential Genes in C. elegans In C. elegans, essential genes can be defined as genes required for growth and development into fertile adulthood. These genes have roles in basic cellular functions, such as metabolism and cell division, as well as developmental and reproductive processes, such as gonadogenesis, fertilization, and embryogenesis. Loss-of-function mutants in essential genes block survival or reproduction and can fall into three broad phenotypic categories: zygotic lethal mutants, in which homozygous animals do not reach adulthood; sterile mutants, in which homozygous animals do not produce fertilized eggs; and maternal-effect lethal mutants, in which homozygous animals survive to adulthood but produce inviable progeny (Kemphues 2005). In this last category, gene products that are supplied to the egg by the hermaphrodite parent are required for viability and mutant phenotypes are not apparent until the F2 generation (Figure 6 1.1). For this reason, it is particularly challenging to study maternal-effect genes in more complex organisms in which generation times are long and large-scale mutant screens are impractical. Figure 1.1 Genetics of maternal-effect lethal mutants, in which gene products inherited from the hermaphrodite parent are required for viability. Homozygous mutants (mut/mut) from heterozygous parents survive to reproductive adulthood, but their progeny are inviable. There are numerous motivations behind the study of essential genes in C. elegans. Firstly, these genes are integral to understanding nematode biology, as they are required in fundamental processes during development and reproduction. In addition, this field of study has implications in parasitic nematology, where C. elegans is an important model system from which functional information can be inferred about genetically intractable parasitic species (Bürglin et al. 1998; Hashmi et al. 2001). In this regard, essential genes are promising candidates for anthelmintic drug targets (Kumar et al. 2007; Doyle et al. 2010), which are the focus of an increasingly important area of research as widespread drug resistance to nematode infection treatments P0F1F2+/mut+/mut mut/mutinviableviable+/++/mutmut/mutmut/mut+/+viable+/+7 emerges (Jabbar et al. 2006). Finally, as mentioned above, essential genes in model organisms are generally well conserved (Hughes 2002; Jordan et al. 2002; Georgi et al. 2013) and are often associated with human diseases (Culetto and Sattelle 2000; Dickerson et al. 2011; Qin et al. 2018). Studying orthologous genes in C. elegans provides an experimental system in which it is possible to discover genes and interrogate gene function in ways that are not practical or ethical in humans. This can lead to a better understanding of the molecular mechanisms that underlie disease phenotypes and, in some cases, has provided the basis for future experimental treatments (Culetto and Sattelle 2000; Silverman et al. 2009). 1.2.2 Essential Gene Discovery in C. elegans In the first few decades of C. elegans research, dozens of forward genetic screens were performed in which mutants of interest were isolated for further study. Large screens for lethal and sterile alleles led to the isolation of mutants for hundreds of different essential genes (for example, Herman 1978; Meneely and Herman 1979; Rogalski et al. 1982; Howell et al. 1987; Clark et al. 1988; Johnsen and Baillie 1988; Kemphues et al. 1988b; McKim et al. 1988; Howell and Rose 1990; Johnsen and Baillie 1991; McKim et al. 1992; Stewart et al. 1998; Gönczy et al. 1999). These early studies generated what I refer to here as legacy mutant collections. Many of these screens were designed to achieve saturation, i.e. to uncover mutants for every essential gene in the targeted genomic interval. Their scope was limited by the availability of genetic balancers that are used to maintain stable heterozygous populations (Edgley et al. 1995), which have only recently been expanded to cover 90% of protein-coding genes (Dejima et al. 2018). The alleles obtained from these genetic screens were often mapped to a region of the genome 8 through deficiency or linkage mapping; however, the process of identifying the molecular nature of genetic mutations one-by-one using traditional methods was slow and laborious. RNAi has also been a useful tool for identifying essential genes in C. elegans and has been the basis of several genome-wide screens (for example, Fraser et al. 2000; Gönczy et al. 2000; Piano et al. 2000; Maeda et al. 2001; Kamath et al. 2003; Simmer et al. 2003), including the first RNAi profiling of early embryogenesis (Sönnichsen et al. 2005). There are currently 3,755 C. elegans genes that have been annotated with lethal or sterile phenotypes from RNAi knockdown studies (data from WormBase version WS275). In comparison, the number of genes currently represented by lethal or sterile mutant alleles is 1,885 (data from WormBase version WS275). These numbers should be considered minimums, as the database annotations are not necessarily up to date. The discrepancy in these numbers could be illustrative of the comparatively time-consuming and laborious nature of isolating and identifying mutants. Additionally, some of the genes identified as essential in RNAi screens may belong to paralogous gene families. All members of a closely related gene family can be targeted simultaneously with RNAi knockdown, while the phenotype of a single loss-of-function mutant for one of the genes can be masked by compensation from its paralogs. The total number of essential genes in C. elegans is unknown, but extrapolation from saturation mutagenesis screens has led to estimates that approximately 15-30% of the ~20,000 genes in this organism are essential (Clark et al. 1988; Howell and Rose 1990; Feichtinger 1995; Johnsen and Baillie 1997; The C. elegans Deletion Mutant Consortium 2012). This suggests the possibility 9 that there are many essential genes in C. elegans that remain unidentified and/or lack representation by a null allele. 1.2.3 Whole Genome Sequencing as a Tool for Gene Identification The emergence of high-throughput sequencing technologies (reviewed in Metzker 2010; Goodwin et al. 2016) has introduced new possibilities for identifying the causal mutations in alleles isolated from genetic screens. As whole genome sequencing (WGS) has become widely adopted, methods for identifying mutant alleles have evolved to take advantage of these technological advances (Sarin et al. 2008; Smith et al. 2008; Srivatsan et al. 2008; Blumenstiel et al. 2009; Schneeberger et al. 2009; Doitsidou et al. 2010; Flibotte et al. 2010; Zuryn et al. 2010; Smith et al. 2016). With WGS becoming increasingly affordable over time (Wetterstrand 2019), mutant collections can now be mined for data in efficient ways that were not possible two decades ago. Performing WGS on a single mutant genome is often insufficient to identify a causal variant due to the abundance of background mutations in any given strain, particularly one that has been subjected to random mutagenesis (Denver et al. 2004; Hillier et al. 2008; Sarin et al. 2008; Flibotte et al. 2010). However, when paired with additional strategies such as deletion or SNP-based mapping, extensive backcrossing and sequencing of the parental strain, or bulk segregant analysis, WGS becomes a valuable tool to expedite gene identification. Furthermore, if multiple independently derived allelic mutants exist, an even simpler approach can be taken. By sequencing two or more mutants within a complementation group and looking for mutations in 10 the same gene, the need for additional mapping or crossing schemes is greatly reduced (Schneeberger et al. 2011; Nordström et al. 2013). In the legacy mutant collections described above, where large numbers of mutants are isolated, it is feasible to obtain complementation groups with multiple alleles for many loci. In addition, the abundance of mutants obtained in these large-scale genetic screens suggests that some existing frozen mutant collections may harbor strains for which the mutations remain unidentified. If such collections are coupled with thorough annotations, they are valuable resources that can be mined with WGS. Indeed, some investigators have recently used such WGS-based approaches to uncover novel essential genes from legacy mutant collections (Jaramillo-Lambert et al. 2015; Qin et al. 2018). These projects bring us closer to identifying all essential genes in C. elegans, and also contribute to the ongoing efforts to obtain null mutants for every gene in the genome (C. elegans Deletion Mutant Consortium 2012). 1.3 Thesis Objectives In this study, we used WGS to revisit two C. elegans legacy mutant collections isolated more than 25 years ago by Dr. Richard Feichtinger, Dr. Heinke Holzkamp and Dr. Ralf Schnabel (Feichtinger 1995; Holzkamp and Schnabel, unpublished data). These collections are a rich resource for essential gene discovery; they comprise 75 complementation groups containing alleles with sterile or maternal-effect lethal phenotypes. The complementation groups each consist of at least two alleles and have been mapped under deficiencies on chromosomes III, IV, and V, which facilitates the efficient identification of causal variants using WGS. With these collections, I 11 sought to identify novel essential genes and to conduct a preliminary characterization of their roles in fertilization and development. By performing wild-type male rescue assays, I was able to attribute some mutant phenotypes to sperm-specific genetic defects. In addition, I examined arrested embryos using differential interference contrast (DIC) microscopy and documented their terminal phenotypes. This thesis comprises a catalogue of 125 alleles with mutations in 59 essential genes. I present several genes which are reported here for the first time as essential genes and mutant alleles for genes that have only previously been studied with RNAi knockdown. The aim of this work is to help accelerate research efforts by identifying essential genes and providing an entry point into further investigations of gene function. Advancing our understanding of essential genes is imperative to reaching a more comprehensive knowledge of gene function in C. elegans and may provide insight into conserved processes in developmental biology, parasitic nematology, and human disease. 12 Chapter 2: Materials and Methods 2.1 Mutant Collections The legacy mutant collections used for this project were a gift from Dr. Ralf Schnabel (Technical University of Braunschweig, Germany). The strains were originally isolated in screens for maternal-effect lethal and sterile mutations using ethyl methanesulfonate (EMS) mutagenesis of two parental balancer strains, qC1(III) and nT1(IV; V). The balancer strains were marked with visible markers to facilitate the identification of homozygotes: unc-32(e189) Coilers for qC1 and unc-24(e138); dpy-11(e224) Dpy-Uncs for nT1. Additionally, him-3(e1147) or him-9(e1487) mutations were introduced to facilitate the generation of males for crossing. Following mutagenesis, animals with homozygous markers in the F2 generation were screened for the production of dead eggs, unfertilized oocytes, or no eggs laid. The two mutant collections are summarized in Table 2.1. Collection A was generated by Dr. Heinke Holzkamp and Dr. Ralf Schnabel in the early to mid 1990s (unpublished data), while collection B was generated by Dr. Richard Feichtinger (Feichtinger 1995). Table 2.1 Summary of mutant collections Collection Number of Complementation Groups Chromosome Genotype A 32 III unc-32(e189) let(tâ¦)/qC1 III; him-3(e1147) IV B 25 IV him-9(e1487) II; unc-24(e138) mat(tâ¦)/nT1 [let(m435)] IV; dpy-11(e224)/nT1 [let(m435)] V 18 V him-9(e1487) II; unc-24(e138)/nT1 [let(m435)] IV; dpy-11(e224) mat(tâ¦)/nT1 [let(m435)] V 13 Together, these collections comprise 75 complementation groups representing essential genes on three chromosomes. A minimum of two alleles for each complementation group (157 strains in total) were examined in this study. At the time of their isolation, each strain was outcrossed at least once, and the alleles were localized to a chromosomal region using deficiency mapping. 2.1.1 List of Strains The Bristol N2 strain PD1074 and the following mutant strains were used in this study: GE1713(t1433), GE1715(t1436), GE1718(t1441), GE1720(t1446), GE1722(t1435), GE1725(t1437), GE1731(t1456), GE1734(t1438, t1477), GE1735(t1470), GE1742(t1461), GE1744(t1463), GE1745(t1464), GE1929(t1729), GE1932(t1732), GE1936(t1738), GE1937(t2189), GE1938(t1742), GE1939(t1745), GE1958(t1726), GE1998(t1790), GE1999(t1793), GE2000(t1795), GE2003(t1817), GE2028(t1801), GE2034(t1808), GE2035(t1809), GE2046(t2015), GE2047(t2021), GE2056(t2016), GE2059(t2025), GE2063(t2042), GE2077(t1757), GE2078(t2033), GE2082(t2053), GE2086(t2093), GE2091(t1772), GE2093(t1800), GE2101(t2080), GE2103(t2092), GE2112(t2037), GE2122(t2007), GE2130(t1765), GE2134(t2079), GE2135(t2043), GE2142(t2074), GE2153(t1773), GE2189(t1750), GE2200(t1480), GE2206(t1514), GE2207(t1539, t1541), GE2211(t1476), GE2219(t1501), GE2220(t1516), GE2237(t1614), GE2242(t1618), GE2264(t1628), GE2275(t1517), GE2277(t1496), GE2278(t1502), GE2288(t1835), GE2289(t1836), GE2292(t2114), GE2304(t2129), GE2305(t1867), GE2314(t1724), GE2317(t2098), GE2326(t2106), GE2335(t2056), GE2337(t2095), GE2343(t1521), GE2345(t1525), GE2347(t1519), GE2348(t1518), GE2352(t1535), GE2355(t1528), GE2357(t1527), GE2359(t1478), GE2360(t1481), GE2362(t1547), GE2366(t1561), GE2367(t1563), GE2373(t1811), GE2379(t1886), GE2386(t2165), GE2387(t1913), GE2391(t1932), GE2393(t2183), GE2399(t1559), GE2402(t1940), GE2407(t1906), GE2430(t2135), GE2433(t1885), GE2442(t1920), GE2445(t1935), GE2447(t1879), GE2449(t2143), GE2451(t2144), GE2452(t1897), GE2453(t1900), GE2455(t1914), GE2467(t2155), GE2469(t2173), GE2476(t2147), GE2487(t2149), GE2497(t2137), GE2499(t1877), GE2503(t1888), GE2511(t2162), GE2512(t1909), GE2514(t1912), GE2517(t2175), GE2523(t2180), GE2526(t1937), GE2533(t1638), GE2541(t2035), GE2545(t1577), GE2551(t1792), 14 GE2577(t1642), GE2579(t1644), GE2583(t1654), GE2590(t1657), GE2595(t1662, t1718), GE2596(t1663), GE2597(t1664), GE2605(t1674), GE2611(t1675), GE2612(t1676), GE2613(t1677), GE2621(t1587), GE2627(t1603), GE2629(t1595), GE2644(t1594), GE2646(t1600), GE2653(t1698), GE2656(t1701), GE2657(t1704), GE2666(t1693), GE2692(t1717), GE2730(t1550), GE2733(t2008), GE2734(t2029), GE2738(t1833), GE2740(t1576), GE2824(t1758), GE2826(t2052), GE2827(t1786), GE2836(t1780), GE2837(t1791), GE2840(t1860), GE2881(t1744), GE2884(t1755), GE2886(t2055), GE2890(t1821), GE2895(t1866), GE2958(t1464, t1484), GE3128(t2177), GE3129(t2181). 2.2 DNA Extraction Balanced heterozygous strains were grown on 100 mm nematode growth medium (NGM) agar plates (standard recipe with 3 times concentration of peptone) seeded with OP50 and harvested at starvation. Worms were washed from the plates with 7 ml of M9 buffer with Triton X-100 (0.5%; M9+TX) into 15 ml tubes and pelleted by centrifugation. The remaining buffer was exchanged with sterile distilled water during 3 subsequent wash steps and the final pellet of worms was frozen at -80°C. Approximately 75 µl of each worm pellet was thawed and treated with Proteinase K (50 µg/ml in 600 µl of reaction buffer: 0.01M Tris pH 7.8, 0.005M EDTA, 0.5% SDS) at 55°C for 2 hours, then with RNAse A (37.5 µg/ml) at 37°C for 2 hours. The resulting lysate was subjected to protein precipitation with 200 µl of NaCl (6 M), followed by cold isopropanol DNA precipitation and a 70% ethanol wash. DNA pellets were dried and resuspended in TE buffer (10 mM Tris, 0.1 mM EDTA; pH 7.5). DNA quality was assessed with a NanoDrop 2000c Spectrophotometer (Thermo Scientific) to ensure A260/280 ratios were between 1.8 and 2.0, and DNA concentration was measured using a Qubit 2.0 Fluorometer and dsDNA Broad Range Assay kit (Life Technologies). 15 2.3 Whole Genome Sequencing and Analysis DNA library preparation and whole genome sequencing were carried out by The Centre for Applied Genomics (The Hospital for Sick Children, Toronto, Canada). Up to 33 C. elegans mutant strains were run together on a single lane of an Illumina HiSeq X to generate 150-bp paired-end reads. Sequencing analysis was performed by Dr. Stephane Flibotte using a modified version of a previously designed custom pipeline (Flibotte et al. 2010; Thompson et al. 2013). Reads were aligned to the C. elegans reference genome (WS263; wormbase.org) using the short-read aligner BWA version 0.7.16 (Li and Durbin 2009). Single nucleotide variants (SNVs) and small insertions or deletions (indels) were called using SAMtools toolbox version 1.6 (Li et al. 2009). To eliminate unreliable calls, variants at genomic locations for which the canonical N2 strain has historically had low read depth or poor quality (Thompson et al. 2013) were removed as potential candidates. The variant calls were annotated with a custom Perl script and labeled heterozygous if represented by 20-80% of the reads at that location. The remaining candidates were then subjected to a series of custom filters. Any calls that appeared in more than three strains from the same collection were removed, since these likely belonged to the genotype of the parent from which the mutant strains were derived. The remaining list was filtered to only include mutations affecting coding exons (indels, missense and nonsense mutations) and splice sites (defined as the first two and last two base pairs in an intron), as these are more simply attributed 16 to a single gene than mutations in noncoding regions of the genome. Finally, the list of candidate mutations was trimmed to include only mutations on the chromosome to which the mutation had originally been mapped. For each pair of strains belonging to a complementation group, the final list of candidate mutations was compared and the gene or genes in common were identified. In cases where there was only one gene in common on both lists, this gene was designated the candidate essential gene. For complementation groups with multiple candidate genes in common, the deficiency mapping data from the original study were used to narrow down the region of interest. Additional information such as the nature of the mutations and existing knowledge about the genes was used to select a single candidate gene, when possible. Gene candidates with identical variants in each strain were removed from consideration*. In several cases, the analysis pipeline described above did not produce any gene candidates for a complementation group, as there was no gene in common within the pair of strains. In such cases, the list of variants was reanalyzed with less stringent filters. This allowed larger deletions and rearrangements to be called in addition to SNVs and indels. For groups with no candidate genes in common after this reanalysis, two additional alleles (if available) were sequenced to help identify the gene of interest. * With the exception of t2033 and t2079, both of which are published alleles with identical mutations in sas-5 (see Section 3.2.1). 17 2.4 Validation of Gene Identities 2.4.1 Corroboration with Mapping Data To corroborate the candidate gene identities with the deficiency mapping data collected when the complementation groups were originally isolated, I compared the genomic location of the candidate gene with the approximate physical map position of the corresponding deficiency. To determine the position of each deficiency, I searched for genes which were known to lie either internal or external to the deletion and obtained their physical map coordinates (when available) to resolve approximate boundaries for the deficiencies. Since the exact boundaries of the deficiencies could not be determined, the mapping data were assumed to be in agreement with the assigned gene identities unless contradicted by information found during this process. 2.4.2 Complementation Tests To verify the complementation groups of a small number of alleles, cross plates were set up at 25°C with 10 heterozygous males from one strain and four heterozygous L4 hermaphrodites of another strain. Successful crosses were determined by the abundance of male progeny on a cross plate. Ten homozygous hermaphrodites (Coilers for Collection A, Dpy-Uncs for Collection B) were selected from the successful cross plates and transferred to new plates at 25°C. After three days in isolation, the plates were screened for the presence of viable progeny, indicating that complementation had occurred and the alleles belong to distinct groups. When none of the homozygous cross progeny exhibited phenotypic rescue, the alleles were designated as members of the same complementation group. 18 2.4.3 CRISPR-Cas9 Deletions Deletion mutants were generated using a CRISPR-Cas9 genome editing strategy previously described (Norris et al. 2015; Au and Li-Leger et al. 2019). Two guide RNAs were used to excise the gene of interest and replace it with a selection cassette expressing G418 drug resistance and pharyngeal GFP (loxP + Pmyo-2::GFP::unc-54 3Ê¹UTR + Prps-27::neoR::unc-543Ê¹UTR + loxP vector, provided by Dr. John Calarco, University of Toronto, Canada), as shown in Figure 2.1. Guide RNAs were designed using the C. elegans Guide Selection Tool (genome.sfu.ca/crispr) and synthesized by Integrated DNA Technologies (IDT). Repair templates were generated by assembling homology arms (450-bp gBlocks synthesized by IDT) and the selection cassette using the NEBuilder Hifi DNA Assembly Kit (New England Biolabs). CRISPR-Cas9 deletions were designed to induce homology-directed repair that would eliminate all or most of the coding region for the gene of interest. 19 Figure 2.1 Schematic of CRISPR-Cas9 gene deletion strategy. (1) Two guide RNAs bind to cut sites at either end of the gene, where Cas9 creates double-strand breaks. (2) Homology arms adjacent to the cut sites flank a dual-marker selection cassette in the repair plasmid. (3) Homology-directed repair causes the sequence between the cut sites to be replaced with the selection markers. The GFP and drug resistance markers are flanked by loxP sites to enable excision, if desired. Figure adapted from Norris et al. (2015). Cas9 protein (generously gifted from Dr. Geraldine Seydoux) was assembled into a ribonucleoprotein (RNP) complex with the guide RNAs and tracrRNA (IDT) following the manufacturerâs recommendations. PD1074 (a sequenced derivative of the original N2 wild-type strain) animals were injected using standard microinjection techniques (Mello et al. 1991; Kadandale et al. 2009) with an injection mix consisting of: 50 ng/µl repair template, 0.5 µM RNP complex, 5 ng/µl pCFJ104 (Pmyo-3::mCherry), and 2.5 ng/µl pCFJ90 (Pmyo-2::mCherry). Injected animals were screened according to the protocol described in Norris et al. (2015) and genomic edits were validated using the PCR protocol described in Au and Li-Leger et al. (2019). Pmyo-2::GFP Prps-27::NeoRPmyo-2::GFP Prps-27::NeoRcut sitecut site5â homology arm3â homology armSelection CassettePlasmidloxPloxPloxPloxPATG5â 3â5â 3â5â 3âCas9 induces double-strand breaksGenomic DNA is replaced by the dual-marker selection cassette via homology-directed repair12320 2.5 Bioinformatic Analysis Previously reported phenotypes from RNAi experiments or mutant alleles were retrieved from WormBase (version WS275; wormbase.org) and GExplore (genome.sfu.ca/gexplore; Hutter et al. 2009; Hutter and Suh 2016). Life stage-specific gene expression data from the modENCODE project (Hillier et al. 2009; Gerstein et al. 2010, 2014; Boeck et al. 2016) were also accessed through GExplore. Visual inspection of these data revealed genes with maternal expression patterns (high levels of expression in the early embryo and hermaphrodite gonad) as well as those predominantly expressed in males. Human orthologs of C. elegans genes were determined using Ortholist 2 (ortholist.shaye-lab.org; Kim et al. 2018), a database which compiles worm-human ortholog predictions from six different programs: Ensembl Compara (Vilella et al. 2009), HomoloGene (Wheeler et al. 2007), InParanoid (Remm et al. 2001; Sonnhammer and Ãstlund 2015), OMA (Roth et al. 2008), OrthoMCL (Li et al. 2003), and OrthoInspector (Linard et al. 2011, 2015). For maximum sensitivity, the minimum number of programs predicting a given ortholog was set to one, as recommended by the authors of the meta-analysis. NCBI BLASTp (blast.ncbi.nlm.nih.gov; Altschul et al. 1990) was used to examine distributions of homologs across species and potential nematode-specificity in genes with no human orthologs. Protein sequences from the longest transcript of each gene were used to query the non-redundant protein sequences (nr) database, with default parameters and a maximum of 1,000 target sequences. The results were filtered with an E-value threshold of 10-5. Genes were 21 designated nematode-specific if all resulting homologous sequences belonged only to nematode species. When only Caenorhabditis species appeared in the results, the gene was designated Caenorhabditis-specific. Functional clustering of genes was performed using DAVID version 6.8 (david.ncifcrf.gov; Huang et al. 2009a, 2009b). The list of candidate genes was used to retrieve gene ontology (GO) terms and group genes with similar GO annotations. For GO term enrichment analysis, the set of all C. elegans genes was used as a background list. The minimum number of genes per category was set to three for biological process (BP) terms, and two for molecular function (MF) terms, with the EASE threshold (modified Fisherâs Exact P-Value) set to the default value, 0.1. 2.6 Temperature Sensitivity and Mating Assays To assay temperature sensitivity, heterozygous strains were propagated at the permissive temperature (15°C) and homozygous L4 animals were isolated on new 60 mm NGM plates (2 x 6/plate or 3 x 3/plate). After one week at 15°C, plates were screened for the presence of viable homozygous progeny. If present, L4 homozygotes were re-plated and transferred to the restrictive temperature (25°C) for three days, then screened again to confirm lethality or sterility. Mating assays were carried out using PD1074 males and mutant hermaphrodites. Three L4-stage homozygous mutant hermaphrodites were isolated and plated with ten N2 males on each of three 60 mm NGM plates. Control plates consisted of three L4 hermaphrodite mutants without 22 males. Mating assays were carried out at 25°C and observations were taken after three days, noting the absence or presence of viable cross progeny. 2.7 Microscopy The terminal phenotypes of dead eggs from maternal-effect lethal mutants were observed using DIC microscopy. Young adult homozygous mutants were dissected to release their eggs in either M9+TX or distilled water and embryos were left to develop at 25°C overnight (~16 hours). Embryos were mounted on 2% agarose pads and visualized using a Zeiss Axioplan 2 equipped with DIC optics. Images of representative embryos were captured using a Zeiss Axiocam 105 Color camera and ZEN 2.6 imaging software (Carl Zeiss Microscopy). For embryos incubated in distilled water, an osmotic integrity defective (OID) phenotype was noted for embryos that burst or swelled and filled the eggshell, as described by Sönnichsen et al. (2005). 23 Chapter 3: Results 3.1 Identification of 59 Essential Genes through Whole Genome Sequencing Whole genome sequencing was performed on a total of 157 strains with Illumina HiSeq X 150-bp paired-end reads. The average mapping rate was 98% and depth of coverage ranged between 21x and 65x (average = 38x). The raw sequence data from this study have been deposited in the NCBI Sequence Read Archive (SRA; ncbi.nlm.nih.gov/sra) under accession number PRJNA628853. Two alleles for each of 75 complementation groups were sequenced and the data were analyzed according to the custom pipeline described in Section 2.3. In the majority of cases, this resulted in a single gene candidate for each pair of alleles (Figure 3.1). When multiple gene candidates arose from the initial analysis, the list was narrowed down using the deficiency mapping data from our collaborators who originally isolated the alleles (see Section 3.2). Phenotype data from existing mutant alleles or RNAi experiments, as well as life stage-specific expression patterns provided additional evidence with which to assign a single gene identity to a pair of strains with multiple hits in common. 24 Figure 3.1 Gene identification in complementation groups from two collections. Most of the complementation groups in Collection A (23 of 32 complementation groups) and Collection B (34 of 43 complementation groups) were assigned gene identities from one pair of alleles. When possible, others were elucidated with the help of additional alleles (4 groups in Collection B). Some complementation groups could not be assigned gene identities because they shared no mutant gene hits in common, or they had multiple hits that could not be resolved. The rate of gene identification was comparable in the two collections (Figure 3.1). For Collection A, the initial analysis resulted in a single gene candidate for 23 out of 32 complementation groups. There were seven groups with no gene hits in common, therefore no gene identity could be assigned. Finally, two complementation groups had candidate gene hits but remain unresolved. One of these groups only had identical variants in common, which was not considered a meaningful result because these are unlikely to be independent alleles. For the other group, a single gene candidate arose from the pair of alleles; however, this gene (top-3) does not reside in the mapped region from the original study and was later shown to be independent of other top-3 alleles (see Section 3.2). Collection A (Chr. III)Collection B (Chr. IV/V)Gene identified from allele pairIdentified from additional allelesUnresolvedNo common hits23347244125 Initial analysis of Collection B produced a single gene candidate for 34 of the 43 complementation groups. There were four groups in this collection for which additional alleles were obtained and sequenced; this led to candidate gene designations that were not apparent from the first two alleles. Four other complementation groups remained without any gene hits in common and no additional alleles could be obtained. One final group was removed from consideration because it had two gene candidates which could not be resolved. The two genes in question, sart-3 and swsn-4, are both located in the region covered by sDf2, yet these two alleles did not map under any of the tested deficiencies (Feichtinger 1995). As these genes are both known essential genes that have been studied with existing null alleles (Sawa et al. 2000; Mathies et al. 2015; Rüegger et al. 2015), we decided not to pursue this complementation group any further. The final set of essential genes identified in this study includes 59 genes on three chromosomes (Table 3.1); Appendix A contains the flanking sequences of the corresponding mutations. Literature searches revealed that 43 of these genes have been previously annotated with lethal or sterile phenotypes from either mutant alleles or RNAi studies. While the list includes many known essential genes, our collection comprises alleles with novel genetic variants. Furthermore, there are 20 genes in this collection which have not been characterized and/or are not yet represented by lethal or sterile mutants*. These 20 genes, represented by 42 alleles, were * One gene, atg-7, was included in this set despite having null alleles that have been previously studied. 26 designated Genes of Interest (GOI; Table 3.2), which were further characterized as part of this study (see Sections 3.3, 3.4 and 3.5). They include 15 genes (30 alleles) with a maternal-effect lethal phenotype (homozygotes lay dead eggs) and 5 genes (12 alleles) with a sterile phenotype (homozygotes lay unfertilized oocytes).27 Table 3.1 List of 59 essential genes with associated maternal-effect lethal or sterile alleles Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ Y GE2430 t2135 air-1 V 8221773 C T SNV missense R62C 326 AURKA, AURKB, AURKC, STK36 Colorectal cancer, susceptibility to [114500]; Spermatogenic failure 5 [243060] GE2337 t2095 air-1 V 8223169 CAT C deletion frameshift - x GE2314 t1724 aptf-2 IV 13414105 A G SNV missense L244P 367 TFAP2A, TFAP2B, TFAP2C, TFAP2D, TFAP2E Char syndrome [169100]; Patent ductus arteriosus 2 [617035]; Branchiooculofacial syndrome [113620] GE2289 t1836 aptf-2 IV 13414263 G T SNV nonsense C191* H GE1958 t1726 atg-7 IV 11079764 G A SNV nonsense Q367* 647 ATG7 (none) GE1936 t1738 atg-7 IV 11079973 C T SNV nonsense W311* T GE2449 t2143 atl-1 V 9635587 C T SNV nonsense W2346* 2531 ATR, PRKDC Cutaneous telangiectasia and cancer syndrome, familial [614564]; Seckel syndrome 1 [210600]; Immunodeficiency 26 with or without neurologic abnormalities [615966] GE2467 t2155 atl-1 V 9637978 C T SNV missense E1710K gene-28 GE2200 t1480 bckd-1A III 12969933 G A SNV nonsense Q174* 432 BCKDHA, TMEM91, AC011462.1 Maple syrup urine disease [248600] GE1742 t1461 bckd-1A III 12971429 G A SNV nonsense Q109* gene-17 GE2206 t1514 bckd-1A III 12971273 G A SNV nonsense Q161* GE2627 t1603 bckd-1A III 12971305 C T SNV nonsense W150* vz GE2890 t1821 C34D4.4 IV 7150054 G A SNV nonsense W101* 205 TVP23A, TVP23B, TVP23C, TVP23C-CDRT4 (none) GE2840 t1860 C34D4.4 IV 7150143 G A SNV nonsense W131* a GE2734 t2029 C56A3.8 V 13560728 G A SNV missense G62E 402 PI4K2A, PI4K2B (none) GE2886 t2055 C56A3.8 V 13560787 G A SNV missense E243K GE2487 t2149 C56A3.8 V 13561369 C T SNV missense P82L 28 Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ V GE2142 t2074 ccz-1 V 13679756 T A SNV nonsense Y248* 528 CCZ1, CCZ1B (none) GE2304 t2129 ccz-1 V 13680792 C T SNV nonsense Q361* b GE2047 t2021 cept-2 V 14349388 G A SNV nonsense W128* 424 CEPT1, CHPT1, SELENOI Spastic paraplegia 81, autosomal recessive [618768] GE2122 t2007 cept-2 V 14349747 G A SNV splice site - gene-4 GE2275 t1517 cls-2 III 9055405 G A SNV missense R102Q 1023 CLASP1, CLASP2 (none) GE2357 t1527 cls-2 III 9055440 G A SNV missense G114R R GE2082 t2053 cpl-1 V 16593886 G A SNV missense S148F 337 CTSF, CTSK, CTSL, CTSS, CTSV Pycnodysostosis [265800]; Ceroid lipofuscinosis, neuronal, 13 [615362] GE2451 t2144 cpl-1 V 16595201 G A SNV nonsense Q49* A GE2447 t1879 cpt-2 IV 11180120 C T SNV nonsense Q141* 646 CPT2 Carnitine palmitoyltransferase II deficiency [600649, 608836, 255110]; Encephalopathy, acute, infection-induced, susceptibility to, 4 [614212] GE1938 t1742 cpt-2 IV 11180603 G A SNV nonsense W194* gene-24 GE2657 t1704 cra-1 III 6867181 G A SNV nonsense Q525* 958 NAA25 (none) GE2242 t1618 cra-1 III 6868737 C T SNV nonsense W149* D GE1929 t1729 csr-1 IV 7960467 T A SNV missense N708K 1030 (none) (none) GE1929 t1729 csr-1 IV 7961246 G A SNV missense G922E GE2452 t1897 csr-1 IV 7959252 G A SNV splice site - gene-25 GE2595 t1662 t1718 cup-5 III 7585568 C T SNV nonsense R263* 668 MCOLN1, MCOLN2, MCOLN3 Mucolipidosis IV [252650] GE2355 t1528 cup-5 III 7590536 G A SNV splice site - gene-30 GE2345 t1525 cyk-3 III 6020590 C T SNV nonsense Q98* 1178 USP15, USP32, USP6 (none) GE2352 t1535 cyk-3 III 6022863 G A SNV nonsense W723* J GE2499 t1877 D2096.12 IV 8363937 C T SNV nonsense Q126* 763 (none) (none) GE2407 t1906 D2096.12 IV 8365654 T A SNV nonsense L638* 29 Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ O GE2135 t2043 dgtr-1 V 6497335 G A SNV splice site - 359 AWAT1, AWAT2, DGAT2, DGAT2L6, MOGAT1, MOGAT2, MOGAT3 (none) GE2063 t2042 dgtr-1 V 6498186 G A SNV missense G310R C GE2028 t1801 dif-1 IV 7552230 A C SNV nonsense Y187* 312 SLC25A20 Carnitine-acylcarnitine translocase deficiency [212138] GE1932 t1732 dif-1 IV 7552641 C T SNV missense G75D gene-13 GE2612 t1676 div-1 III 10245480 G A SNV nonsense Q489* 581 POLA2 (none) GE2577 t1642 div-1 III 10248544 C T SNV start ATG M1I d GE2335 t2056 dlat-1 V 14445907 G A SNV nonsense Q419* 507 DLAT Pyruvate dehydrogenase E2 deficiency [245348] GE2541 t2035 dlat-1 V 14446981 G A SNV missense P83L u GE2402 t1940 F21D5.1 IV 8727315 C T SNV missense A436V 550 PGM3 Immunodeficiency 23 [615816] GE2445 t1935 F21D5.1 IV 8727668 C T SNV missense L539F t GE2837 t1791 F56D5.2 IV 9397791 G A SNV nonsense Q214* 385 (none) (none) GE2881 t1744 F56D5.2 IV 9398158 G A SNV missense S107F gene-26 GE1715 t1436 gsp-2 III 7337087 C T SNV nonsense R95* 333 PPP1CA, PPP1CB, PPP1CC Noonan syndrome-like disorder with loose anagen hair 2 [617506] GE2360 t1481 gsp-2 III 7337383 G A SNV missense G174E gene-32 GE2545 t1577 gsr-1 III 3652401 G A SNV missense G335R 473 GSR, TXNRD1, TXNRD2, TXNRD3 Hemolytic anemia due to glutathione reductase deficiency [618660]; Glucocorticoid deficiency 5 [617825] GE2644 t1594 gsr-1 III 3652407 C T SNV nonsense R337* gene-31 GE2583 t1654 hcp-3 III 9615498 G A SNV missense R269C 288 CENPA (none) GE2692 t1717 hcp-3 III 9615555 C T SNV missense E250K B GE2034 t1808 K08D8.6 IV 12887810 G A SNV missense T418I 491 (none) (none) GE2035 t1809 K08D8.6 IV 12889703 T A SNV missense Q93H 30 Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ G GE2455 t1914 klp-18 IV 7040335 T C SNV missense Y42H 932 KIF15 (none) GE2000 t1795 klp-18 IV 7041203 G A SNV missense E316K gene-6 GE2367 t1563 klp-19 III 13306451 A T SNV missense L230H 1083 KIF4A, KIF4B Mental retardation, X-linked 100 [300923] GE2367 t1563 klp-19 III 13306457 G A SNV missense A228V GE2264 t1628 klp-19 III 13306872 C T SNV missense G90R I GE2003 t1817 let-99 IV 12569291 C T SNV nonsense Q447* 698 (none) (none) GE2514 t1912 let-99 IV 12570199 C T SNV missense L617F gene-22 GE2730 t1550 lis-1 III 13375376 C T SNV nonsense W92* 404 PAFAH1B1 Lissencephaly 1; Subcortical laminar heterotopia [607432] GE2653 t1698 lis-1 III 13375401 C T SNV splice site - z GE2130 t1765 mbk-2 IV 13033086 C T SNV missense R533C 817 DYRK2, DYRK3, DYRK4 (none) GE2503 t1888 mbk-2 IV 13033644 C T SNV missense P701L gene-10 GE2740 t1576 mel-32 III 6440655 C T SNV missense G395R 507 SHMT1, SHMT2 (none) GE1731 t1456 mel-32 III 6440831 C T SNV missense G336E M GE1999 t1793 mex-5 IV 13354014 T G SNV nonsense Y79* 468 (none) (none) GE2093 t1800 mex-5 IV 13354478 T A SNV nonsense L219* S GE2511 t2162 mom-2 V 8356808 T G SNV missense C80G 362 WNT11, WNT9A, WNT9B (none) GE2523 t2180 mom-2 V 8357121 T C SNV missense C139R W GE2497 t2137 mre-11 V 10735712 G A SNV missense H269Y 728 MRE11 Ataxia-telangiectasia-like disorder 1 [604391] GE2103 t2092 mre-11 V 10736080 A G SNV missense F146S v GE2091 t1772 nstp-2 IV 6604731 A T SNV missense L277H 324 SLC35B4 (none) GE2288 t1835 nstp-2 IV 6605266 C T SNV missense G131R F GE2391 t1932 perm-5 IV 5696931 A T SNV missense C454S 518 (none) (none) GE2453 t1900 perm-5 IV 5698096 A G SNV missense S323P 31 Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ gene-21 GE2237 t1614 pod-1 III 13518266 G A SNV missense A912V 1136 CORO7, CORO7-PAM16 (none) GE2605 t1674 pod-1 III 13518357 G A SNV nonsense R882* U GE3128 t2177 pos-1 V 8414544 G A SNV splice site - 264 (none) (none) GE2101 t2080 pos-1 V 8414579 T A SNV missense V145D Z GE2517 t2175 rad-50 V 12247914 T A SNV nonsense L350* 1312 RAD5, AC116366.3 Nijmegen breakage syndrome-like disorder [613078] GE2476 t2147 rad-50 V 12250324 T A SNV missense I1101N E GE2189 t1750 rad-51 IV 10282013 A T SNV missense I384N 395 DMC1, RAD51, RAD51B, RAD51C, RAD51D Fanconi anemia, complementation group R, group O [617244, 613390]; Mirror movements 2 [614508]; Breast-ovarian cancer, familial, susceptibility to, 3 [613399] GE2433 t1885 rad-51 IV 10282328 C T SNV missense V323I gene-11 GE2347 t1519 rmd-1 III 9759805 G A SNV missense G89R 226 RMDN2, RMDN3 (none) GE2219 t1501 rmd-1 III 9759929 G A SNV missense R130H gene-18 GE2211 t1476 sas-1 III 12710102 C T SNV missense P419S 570 (none) (none) GE2343 t1521 sas-1 III 12710202 G A SNV missense G452E f GE2078 t2033 sas-5 V 11612449 C T SNV missense R397C 404 (none) (none) GE2134 t2079 sas-5 V 11612449 C T SNV missense R397C P GE2469 t2173 spn-4 V 6783986 A T SNV nonsense L259* 351 RBFOX1, RBFOX2, RBFOX3 (none) GE2317 t2098 spn-4 V 6784646 A T SNV missense V55D g GE2386 t2165 sqv-4 V 10660827 G A SNV missense P182L 481 UGDH Epileptic encephalopathy, early infantile, 84 [618792] GE2059 t2025 sqv-4 V 10661143 G A SNV missense S93L gene-5 GE2277 t1496 such-1 III 11515520 G A SNV missense L686F 798 ANAPC5 (none) GE2277 t1496 such-1 III 11515883 G A SNV missense H565Y GE2666 t1693 such-1 III 11515540 C T SNV missense R679K 32 Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ q GE2827 t1786 T22B11.1 IV 4692945 G A SNV nonsense W35* 468 (none) (none) GE2895 t1866 T22B11.1 IV 4696017 G A SNV nonsense W356* gene-12 GE1734 t1438 t1477 tlk-1 III 9707175 C T SNV nonsense Q412* 965 TLK1, TLK2, TLK2PS1 Mental retardation, autosomal dominant 57 [618050] GE2613 t1677 tlk-1 III 9708080 G A SNV missense A694T gene-15 GE2399 t1559 top-3 III 11951381 G A SNV nonsense Q602* 759 TOP3A Progressive external ophthalmoplegia with mitochondrial DNA deletions, autosomal recessive 5 [618098]; Microcephaly, growth restriction, and increased sister chromatid exchange 2 [618097] GE2220 t1516 top-3 III 11958680 C T SNV missense G59R gene-35 GE1735 t1470 top-3 III 11957525 C T SNV nonsense W114* GE2958 t1464 t1484 top-3 III 11951669 C T SNV missense G506R L GE2512 t1909 trcs-1 IV 9587541 C T SNV missense E373K 428 AADAC, AADACL2, AADACL3, AADACL4, NCEH1 (none) GE1939 t1745 trcs-1 IV 9587985 G A SNV nonsense Q242* c GE2112 t2037 unc-112 V 14692219 C T SNV missense R669Q 720 FERMT1, FERMT2, FERMT3 Kindler syndrome [173650]; Leukocyte adhesion deficiency, type III [612840] GE2326 t2106 unc-112 V 14696546 C T SNV splice site - gene-27 GE1722 t1435 vps-33.1 III 8701605 C T SNV nonsense R159* 603 VPS33A, VPS33B, AC048338.1 Mucopolysaccharidosis-plus syndrome [617303]; Arthrogryposis, renal dysfunction [208085] GE2366 t1561 vps-33.1 III 8702923 G A SNV nonsense W536* Q GE2292 t2114 vps-39 V 14035713 G A SNV nonsense Q754* 926 VPS39 (none) GE1937 t2189 vps-39 V 14036143 C T SNV nonsense W626* GE2056 t2016 vps-39 V 14037839 G C SNV nonsense Y122* N GE2153 t1773 wapl-1 IV 4444464 C T SNV nonsense W348* 748 WAPL (none) GE2305 t1867 wapl-1 IV 4442749-4442872 - - 122-bp deletion deletion - 33 Group Strain Allele(s) Gene Chr. Position Base Change Mutation Mutation Type Amino Acid Changeâ Protein Size (Amino Acids)â Human Ortholog(s) Associated OMIM phenotype(s)â¡ p GE2738 t1833 Y54G2A.73 IV 3000662 A T SNV nonsense L341* 380 (none) (none) GE2387 t1913 Y54G2A.73 IV 3001767 G A SNV nonsense R252* GE2884 t1755 Y54G2A.73 IV 3008481 C T SNV splice site - gene-23 GE1713 t1433 ZK688.9 III 7882477 C T SNV nonsense W135* 281 TIPRL (none) GE2621 t1587 ZK688.9 III 7882717 C T SNV splice site - gene-14 GE2348 t1518 zyg-8 III 12063671 C T SNV nonsense R312* 802 DCLK1, DCLK2, DCLK3, DCX Lissencephaly, X-linked, 1; Subcortical laminar heterotopia, X-linked [300067] GE2362 t1547 zyg-8 III 12063832 G A SNV splice site - â Amino acid position and size derived from the longest transcript (wormbase.org, version WS275) â¡Phenotypes retrieved from omim.org 34 Table 3.2 Genes of interest and associated phenotypes Strain Allele Gene Name Protein Functionâ Amino Acid Changeâ RNAi Phenotypeâ¡ Mutant Phenotype Embryonic Osmotic Integrity Defect GE1936 t1738 atg-7 E1 ubiquitin-activating-like enzyme orthologous to the autophagic budding yeast protein Apg7p W311* growth variant; dauer body morphology variant; pathogen induced death increased; P granule localization defective; dauer development variant; protein aggregation variant; shortened life span; transgene subcellular localization variant; transgene expression variant; necrotic cell death variant; autophagy variant; antibody staining reduced dead embryos no GE1958 t1726 Q367* dead embryos no GE2627 t1603 bckd-1A Predicted mitochondrial protein with alpha-ketoacid dehydrogenase activity W150* shortened life span; small dead embryos yes GE2206 t1514 Q161* dead embryos yes GE2840 t1860 C34D4.4 Predicted to have the following domain: Golgi apparatus membrane protein TVP23-like W131* -- unfertilized oocytes N/A GE2890 t1821 W101* unfertilized oocytes N/A GE2734 t2029 C56A3.8 Predicted to have 1-phosphatidylinositol 4-kinase activity G62E larval lethal; accumulated germline cell corpses; germ cell compartment morphology variant; germline nuclear positioning variant; larval arrest; cell membrane organization biogenesis variant; embryonic lethal; rachis narrow; apoptosis variant; maternal sterile; reduced brood size unfertilized oocytes N/A GE2487 t2149 P82L unfertilized oocytes N/A GE2886 t2055 E243K unfertilized oocytes N/A GE2122 t2007 cept-2 Predicted to have diacylglycerol cholinephosphotransferase activity and ethanolaminephospho-transferase activity splice site fat content reduced; embryonic lethal; long dead embryos no GE2047 t2021 W128* no eggs laid (dead embryos) [ts] some GE2275 t1517 cls-2 Member of the CLASP family of microtubule-binding proteins R102Q locomotion variant; mitosis variant; univalent meiotic chromosomes; no polar body formation; chromosome segregation variant karyomeres early emb; mitotic chromosome segregation variant; mitotic spindle defective early emb; chromosome segregation variant; embryonic lethal; meiotic spindle defective; meiotic progression during oogenesis variant; exploded through vulva; reduced brood size; antibody subcellular localization variant; meiotic chromosome segregation variant dead embryos N/T GE2357 t1527 G114R dead embryos no 35 Strain Allele Gene Name Protein Functionâ Amino Acid Changeâ RNAi Phenotypeâ¡ Mutant Phenotype Embryonic Osmotic Integrity Defect GE1938 t1742 cpt-2 Carnitine palmitoyl transferase W194* embryonic lethal dead embryos no GE2447 t1879 Q141* dead embryos no GE2407 t1906 D2096.12 -- L638* locomotion variant dead embryos some GE2499 t1877 Q126* dead embryos yes GE2063 t2042 dgtr-1 Acyl chain transfer enzyme G310R sterile; sick; oocyte number decreased; germline nuclear positioning variant; oocyte septum formation variant; embryonic lethal; embryo osmotic integrity defective early emb; oocyte morphology variant; pachytene region organization variant; reduced brood size; germ cell compartment expansion variant; oogenesis variant dead embryos some GE2135 t2043 splice site dead embryos yes GE2541 t2035 dlat-1 Predicted to have dihydrolipoyllysine-residue acetyltransferase activity P83L embryonic lethal; slow growth; receptor mediated endocytosis defective; pattern of transgene expression variant; sterile progeny; transgene expression increased; general pace of development defective early emb dead embryos no GE2335 t2056 Q419* dead embryos no GE2402 t1940 F21D5.1 Predicted to have phosphoacetyl-glucosamine mutase activity A436V sterile; germ cell compartment size variant; rachis wide; rachis morphology variant; accumulated germline cell corpses; germ cell compartment morphology variant; germline nuclear positioning variant; embryonic lethal; embryo osmotic integrity defective early emb; apoptosis variant; reduced brood size; oogenesis variant dead embryos yes GE2445 t1935 L539F dead embryos yes GE2881 t1744 F56D5.2 -- S107F -- unfertilized oocytes N/A GE2837 t1791 Q214* unfertilized oocytes N/A GE2034 t1808 K08D8.6 Ortholog of human epoxide hydrolase 1§ T418I -- dead embryos some GE2035 t1809 Q93H dead embryos (some unfertilized oocytes) no GE2091 t1772 nstp-2 Predicted to have UDP-N-acetylglucosamine and UDP-xylose transmembrane transporter activity L277H lysosome-related organelle morphology variant; transgene subcellular localization variant; RAB-11 recycling endosome localization variant; RAB-11 recycling endosome morphology variant dead embryos no GE2288 t1835 G131R dead embryos no 36 Strain Allele Gene Name Protein Functionâ Amino Acid Changeâ RNAi Phenotypeâ¡ Mutant Phenotype Embryonic Osmotic Integrity Defect GE2391 t1932 perm-5 Predicted to have lipid binding activity C454S sterile; apoptosis reduced; oocytes lack nucleus; oocyte number decreased; germ cell compartment morphology variant; germline nuclear positioning variant; germ cell compartment anucleate; oocyte septum formation variant; cell membrane organization biogenesis variant; embryonic lethal; embryo osmotic integrity defective early emb; oogenesis variant; diplotene region organization variant dead embryos yes GE2453 t1900 S323P dead embryos yes GE2827 t1786 T22B11.1 -- W35* -- unfertilized oocytes [ts] N/A GE2895 t1866 W356* unfertilized oocytes [ts] N/A GE2399 t1559 top-3 Exhibits DNA topoisomerase type I (single strand cut, ATP-independent) activity G59R chromosome morphology variant; hermaphrodite germline proliferation variant; antibody staining increased; somatic gonad development variant; gonad degenerate; chromosome instability; germ cell mitosis variant; gonad arm morphology variant; meiosis variant; oocyte morphology variant; nuclear appearance variant; fewer germ cells; oogenesis variant dead embryos N/T GE2220 t1516 Q602* dead embryos N/T GE2512 t1909 trcs-1 Putative arylacetamide deacetylase and microsomal lipase E373K apoptosis reduced; diplotene absent during oogenesis; oocyte number decreased; embryo osmotic integrity defective early emb; rachis narrow; chromosome condensation variant; pachytene region organization variant; membrane trafficking variant; pachytene progression during oogenesis variant; apoptosis fails to occur; egg laying variant; germ cell compartment expansion absent; embryonic lethal; cell membrane organization biogenesis variant; no oocytes; germ cell compartment expansion variant dead embryos [leaky ts] yes GE1939 t1745 Q242* no eggs laid (dead embryos) [ts] yes GE2884 t1755 Y54G2A.73 -- splice site -- unfertilized oocytes N/A GE2387 t1913 R252* unfertilized oocytes N/A GE2738 t1833 L341* unfertilized oocytes N/A GE1713 t1433 ZK688.9 Predicted to have the following domain: TIP41-like protein (TOR signaling pathway regulator) W135* egg laying variant; locomotion variant dead embryos no GE2621 t1587 splice site dead embryos no [ts] = temperature sensitive. â From WormBase (WS275; wormbase.org); amino acid position derived from the longest transcript. N/A = not applicable. â¡ Phenotypes retrieved from GExplore (genome.sfu.ca/gexplore). N/T = not tested. § No human orthologs according to Ortholist 2 (ortholist.shaye-lab.org). -- = no information available. 37 3.2 Validation of Candidate Gene Identities The candidate gene identities were corroborated with the deficiency mapping data generated by our collaborators, Dr. Richard Feichtinger, Dr. Heinke Holzkamp and Dr. Ralf Schnabel (Feichtinger 1995; Holzkamp and Schnabel, unpublished data), as shown in Figure 3.2. In most cases, the mapping data were in agreement with the positions of the candidate gene loci, providing validation for the assigned gene identities. For example, there were four complementation groups that had been mapped under the deletion eDf19 on chromosome IV. The genes that we assigned to these groups in our initial analysis (D2096.12, F21D5.1, F56D5.2, and trcs-1) each have physical map coordinates that fall into the region covered by eDf19, as shown in Figure 3.2. When the initial analysis resulted in more than one possible gene candidate, the deficiency mapping was used to narrow down the list. For example, the strains GE1713(t1433) and GE2621(t1587) both had mutations in two genes on chromosome III, nft-1 and ZK688.9; however, nft-1 resides outside of the deletion (nDf16) under which these mutations were mapped so it was eliminated from consideration. 38 Figure 3.2 Schematic of gene assignments and deficiency mapping. Genes and deficiencies are shown with their relative positions on chromosomes III, IV, and V. Approximate boundaries of each deficiency were determined by the coordinates of the closest gene known to lie outside of the deletion, when possible (indicated by a faded edge). If no such genes with physical coordinates are known, the outermost gene known to lie inside the deletion was used as the boundary (indicated by a sharp edge). Gene names are coloured according to the deficiency to which the alleles were mapped. Genes names assigned to alleles that did not map under any of the tested deficiencies are highlighted in grey. top-3 and bckd-1A on chromosome III are represented by multiple complementation groups with conflicting results from deficiency mapping. Chr. IIItop-3mel-32rmd-1tlk-1div-1 zyg-8bckd-1Asas-1pod-1lis-1ZK688.9cra-1cup-5gsp-2 vps-33.1cyk-3hcp-3gsr-1cls-2such-1klp-19sDf125nDf40 tDf5tDf6nDf16sDf110tDf7Chr. Vdlat-1spn-4 cpl-1mom-2atl-1rad-50dgtr-1air-1sqv-4mre-11ccz-1 cept-2unc-112C56A3.8vps-39pos-1sas-5nDf32sDf35ctDf1itDf2D2096.12Chr. IVdif-1csr-1perm-5 rad-51F56D5.2F21D5.1nstp-2trcs-1atg-7C34D4.4T22B11.1 klp-18cpt-2K08D8.6mbk-2aptf-2wapl-1let-99Y54G2A.73mex-5nDf41stDf7eDf19sDf2sDf2139 There were some conflicts between the legacy mapping data and the gene candidates proposed through sequencing analysis. Four complementation groups that did not map under any of the tested deficiencies (Feichtinger 1995; Dr. Ralf Schnabel, personal communication) were assigned gene identities whose genomic coordinates fall into regions covered by the tested deficiencies (e.g. K08D8.6 falls under sDf21). In addition, two of these groups were assigned the same gene identity as another, supposedly distinct, complementation group (Table 3.3). From our analysis, bckd-1A was the initial gene candidate for two different complementation groups, yet only one of these groups had been mapped to a deletion (tDf5) that covers the bckd-1A locus. Similarly, top-3 was the assigned gene candidate for three different complementation groups, only one of which was mapped under a deficiency (tDf5) encompassing that gene. By performing complementation tests with select alleles, I concluded that the two bckd-1A groups are not distinct, and indeed they share the same gene identity. One of the groups (gene-35) originally identified as top-3 is a double mutant which fails to complement gene-15 (top-3) and gene-34 (unknown gene). The results of the complementation tests are summarized in Table 3.3 and Figure 3.3. 40 Table 3.3 Complementation tests for conflicting groups Original Group Strain Allele Preliminary Gene Candidate Amino Acid Change Mapped Under Final Gene Assignment gene-28 GE1742 t1461 bckd-1A Q109* None of tested deficiencies bckd-1A GE2200 t1480 Q174* None of tested deficiencies gene-17 GE2627 t1603 bckd-1A W150* tDf5 bckd-1A GE2206 t1514 Q161* tDf5 gene-15 GE2220 t1516 top-3 G59R tDf5 top-3 GE2399 t1559 Q602* tDf5 gene-34 GE1745 t1464 top-3 G506R None of tested deficiencies (double mutant gene-34 and gene-35) unknown gene GE2278 t1502 P555L None of tested deficiencies gene-35 GE1735 t1470 top-3 W114* None of tested deficiencies double mutant: top-3 + unknown gene GE2958 t1464 t1484 G506R None of tested deficiencies Figure 3.3 Complementation tests for the strains in Table 3.3. Strain names are coloured by their original complementation pairs and grouped by the preliminary gene identities assigned to them (bckd-1A and top-3). The results of each cross reveal which groups are in fact allelic. bckd-1Atop-3Preliminary gene candidateâGE2220GE2278GE1735GE2220GE2399GE2220ââGE1742 GE2627GE1742 GE2206âGE2627GE2206ââ=x=x=x=x=x=xGE2278GE1735 =x âComplementation testsâ - failure to complement | â - complementationGE1745GE2958GE2220GE2399GE2278GE1735GE1742GE2200GE2627GE220641 For four candidate genes, their identities were confirmed by generating a deletion of the gene in a wild-type background using CRISPR-Cas9 genome editing. The genes subjected to this additional validation step were chosen because they had conflicting mapping data (K08D8.6, see Figure 3.2), the phenotyping results were inconsistent between alleles (nstp-2; see Table 3.4), or they were expected to be of interest to the broader research community (C34D4.4 and F56D5.2). The deletion alleles for C34D4.4 and F56D5.2 have been verified with PCR and their phenotypes corroborate those of their legacy mutant counterparts (unfertilized oocytes laid). PCR validation and phenotyping for the K08D8.6 and nstp-2 deletion alleles are currently in progress, as is complementation testing between the newly generated CRISPR-Cas9 mutants and the legacy mutant strains. 3.2.1 Inadvertent Inclusion of a Test Set Validates the Analysis Approach During the course of this project it became known to us that 17 previously studied strains were unknowingly included in Collections A and B. These 17 strains were put through our analysis pipeline before we discovered they were not novel alleles and therefore served as a blind validation set. This set contained previously sequenced alleles representing 9 genes with documented maternal-effect lethal phenotypes (Vatcher et al. 1998; Gönczy et al. 2001; Kaitna et al. 2002; Brauchle et al. 2003; Cockell et al. 2004; Delattre et al. 2004; Sonneville et al. 2004; Bischoff and Schnabel 2006; Nieto et al. 2010). Eight of these nine genes were correctly identified upon our initial analysis, and one was not resolved. For this last gene (cul-2), one of its alleles was mutated in an intronic region and the variant was therefore eliminated by our 42 filtering criteria, leaving no gene hits in common between the pair of alleles (see Section 4.2). Having the results of this test set corroborated with published data was considered an important validation of our analysis approach. 3.3 Bioinformatic Analysis: Human Orthologs, Gene Function and Expression Patterns Of the 59 essential genes identified, 47 genes have predicted human orthologs (Table 3.1). Many of these human genes have been implicated in disease and are associated with OMIM disease phenotypes (Online Mendelian Inheritance in Man; omim.org). Conversely, six genes in the GOI collection are either nematode-specific or Caenorhabditis-specific, based on BLASTp searches. F56D5.2, perm-5, and T22B11.1 are nematode-specific genes that have homologs in parasitic species, such as Loa loa, Brugia malayi, Haemonchus contortus, and Onchocerca volvulus. Furthermore, there are three uncharacterized genes (D2096.12, K08D8.6, and Y54G2A.73) that do not have significant homology outside the Caenorhabditis genus. To gain insight into the functions of the identified essential genes, DAVID functional annotation (Huang et al. 2009a, 2009b) was used to elucidate the most prominent gene ontology (GO) terms associated with them. Diverse biological process (BP) and molecular function (MF) terms were represented in the set of 59 essential genes, with the 20 GOI being distributed among them (Figure 3.4). While the majority of genes (45 of 59), were unsurprisingly annotated with âembryonic development ending in birth or egg hatchingâ, they were broken down into smaller groups by association with other BP terms including chromosome segregation (8 genes), oogenesis (7 genes), DNA repair (5 genes), and vesicle-mediated transport (4 genes) (Figure 3.4). 43 Among the most common MF annotations were ATP binding (10 genes), nucleotide binding (10 genes), transferase activity (9 genes), and kinase activity (6 genes) (Figure 3.4). The MF annotations were more sparse than the BP annotations for this set of genes: 7 genes were not included in the BP output, whereas 33 genes were not included in the MF output due to either a lack of annotation or enrichment levels below the threshold. A full list of genes associated with each GO term is included in Appendix B. 44 Figure 3.4 Top GO terms represented in the set of 59 genes. (A) Biological process terms represented by at least three genes in the set, with 7 genes omitted from the results. (B) Molecular function terms represented by at least 2 genes in the set, with 33 genes omitted from the results. Genes belonging to the set of 20 GOI are coloured in blue. All GO terms are organized from left to right in order of increasing p-value. 05101520253035404550emb. dev. ending in birth or egg hatchingcell cyclecell divisionmeiotic cell cyclemitotic nuclear divisionmeiotic chromosome segregationmulticellular organism developmentmeiotic nuclear divisionapoptotic processcellular response to DNA damage stimulusoogenesisDNA repairmitotic spindle organizationvesicle-mediated transportmicrotubule-based movementestablishment of mitotic spindle orientationreproductionphosphorylationpronuclear migrationovipositiondetermination of adult lifespanNumber of GenesGenes of InterestOther Genes0246810121416microtubule bindingprotein bindingATPase activitykinase activityATP bindingtransferase activityprotein serine/threonine kinaseâ¦nucleotide binding3'-5' exonuclease activityNumber of GenesGenes of InterestOther GenesAB45 To examine the timing of gene expression throughout the life cycle, gene expression data from the modENCODE project (Hillier et al. 2009; Gerstein et al. 2010, 2014; Boeck et al. 2016) was retrieved from GExplore (genome.sfu.ca/gexplore; Hutter et al. 2009; Hutter and Suh 2016) for the 20 genes of interest (Appendix C). These data show a U-shaped expression pattern for ten of the GOI, with high expression occurring in the early embryonic stages as well as in adulthood, and particularly in the hermaphrodite gonad (for example, F21D5.1; Figure 3.5). This U-shaped pattern is characteristic of a maternal-effect gene, for which gene products are passed on to the embryo from the parent. Five genes had a maternal gene expression pattern as well as expression throughout other stages of the life cycle, indicating an additional, zygotic role for the gene. Another common pattern seen among the GOI was gene expression predominantly in males and L4-stage hermaphrodites (for example, F56D5.2; Figure 3.5). These genes were suspected to be involved in sperm production or fertilization. The associated strains were subjected to mating assays, and indeed seven genes were found to be required in male mating (Section 3.4). 46 Figure 3.5 Examples of characteristic expression patterns for maternal-effect and sperm-specific genes. A) F21D5.1 has a characteristic maternal expression pattern, with a high level of gene expression in the early embryo and hermaphrodite gonad. (B) F56D5.2 is a putative sperm-specific gene which is predominantly expressed in males and L4 hermaphrodites. Expression patterns were generated by the modENCODE project (Hillier et al. 2009; Gerstein et al. 2010, 2014; Boeck et al. 2016) and retrieved from GExplore (genome.sfu.ca/gexplore). 3.4 Temperature Sensitivity & Mating Assays The 42 alleles associated with the 20 GOI were further examined to gain insight into the phenotypic consequences of their mutations. Each allele was assayed for temperature sensitivity, as some of the original mutant screening was carried out at 25°C. Five alleles (marked with a [ts] phenotype in Table 3.2) were deemed temperature sensitive and could proliferate as homozygotes at a permissive temperature of 15°C, while being maternal-effect lethal or sterile at a restrictive temperature of 25°C. Four of these alleles had stringent temperature sensitivity at 25°C, while the phenotype of trcs-1(t1909) was leaky at the restrictive temperature. A BF21D5.1 F56D5.247 Eight candidate genes (18 alleles) were hypothesized to be involved in male fertility, based on the production of unfertilized oocytes by hermaphrodites and/or predominantly male-expressed gene expression patterns. These 18 strains were assayed for their ability to be rescued through mating with wild-type males. 14 of the strains were rescued by the mating assay, while four strains failed to rescue (Table 3.4). Phenotypic rescue through mating was consistent among alleles of the same gene in six of the eight genes, while two genes had conflicting results among the pair of alleles in their complementation groups (F56D5.2 and nstp-2). Table 3.4 Putative sperm-expressed genes Strain Allele Gene Observed Mutant Phenotype Successful WT Male Rescue GE2627 t1603 bckd-1A dead embryos yes GE2206 t1514 dead embryos yes GE2840 t1860 C34D4.4 unfertilized oocytes yes GE2890 t1821 unfertilized oocytes yes GE2734 t2029 C56A3.8 unfertilized oocytes yes GE2487 t2149 unfertilized oocytes yes GE2886 t2055 unfertilized oocytes yes GE2881 t1744 F56D5.2 unfertilized oocytes no GE2837 t1791 unfertilized oocytes yes GE2034 t1808 K08D8.6 dead embryos no GE2035 t1809 dead embryos (some unfertilized oocytes) no GE2091 t1772 nstp-2 dead embryos no GE2288 t1835 dead embryos yes GE2827 t1786 T22B11.1 unfertilized oocytes [ts] yes GE2895 t1866 unfertilized oocytes [ts] yes GE2884 t1755 Y54G2A.73 unfertilized oocytes yes GE2387 t1913 unfertilized oocytes yes GE2738 t1833 unfertilized oocytes yes [ts] = temperature sensitive 48 3.5 Terminal Phenotypes of Maternal-Effect Lethal Embryos Using DIC microscopy, the terminal phenotypes of 28 maternal-effect lethal strains (a subset of the 42 GOI strains) were observed. Representative images were selected and compiled into a catalogue of terminal phenotypes (Appendix D). Ten strains showed an osmotic integrity defective (OID) phenotype (as described in Sönnichsen et al. 2005) in nearly all embryos after incubation in distilled water, while four additional strains had only some embryos that exhibited this phenotype (Table 3.2). The OID phenotype was evident in embryos that filled the eggshell completely (for example, dgtr-1(t2043), Figure 3.6A) and eggs that burst in their hypotonic surroundings. Seven mutant strains exhibited early embryonic arrest; four of these did not appear to complete the first cell division and were also associated with an OID phenotype when incubated in distilled water. Alternatively, the two dlat-1 mutant strains (t2035 and t2056) maintained their osmotic integrity but often arrested with only one to four cells (for example, Figure 3.6B). 12 strains had embryos that terminated with approximately 100-200 cells (for example, ZK688.9(t1433), Figure 3.6C); while four strains developed into two- or three-fold stage embryos that did not hatch and exhibited clear morphological defects, such as nstp-2(t1772) with a lumpy body wall and constricted nose tip (Figure 3.6D). 49 Figure 3.6 Embryonic arrest visualized with DIC microscopy for select maternal-effect lethal mutants. Eggs were dissected from homozygous mutants and imaged immediately (A) or incubated in distilled water overnight before imaging (B, C, and D). (A) Eggs dissected from dgtr-1(t2043) homozygotes exhibit signs of an osmotic integrity defect, by filling the eggshell completely. (B) dlat-1(t2035) embryos exhibit early embryonic arrest, with most embryos consisting of four cells or less. (C) ZK688.9(t1433) embryos arrested with approximately 100 cells. (D) Terminal embryos of nstp-2(t1772) have a lumpy body wall morphology and constricted nose; most animals were moving inside the eggshell but did not hatch. All scale bars represent 10 μm. A BC D50 Chapter 4: Discussion 4.1 Revisiting Legacy Mutant Collections with Whole Genome Sequencing This thesis focused on reexamining legacy collections of C. elegans mutants which were isolated before the genome sequence was published (The C. elegans Sequencing Consortium 1998) and before massively parallel sequencing was available and widely used. With major advances in sequencing technology in the past 30 years (reviewed in Goodwin et al. 2016), WGS has become affordable and accessible, enabling us to revisit past projects with new approaches and advanced capabilities. In this study, we sequenced paired alleles from 75 complementation groups on chromosomes III, IV, and V and identified 59 essential genes (Table 3.1). While WGS is a powerful tool, it does not stand alone as a solution to identifying mutant alleles. This study has shown the power of having multiple alleles in a complementation group when faced with the abundance of genomic variants found in WGS analysis. Indeed, when we sequenced four single alleles which had no complementation pairs, we were unable to designate a single mutation as the variant responsible for maternal-effect lethality (data not shown). Our approach to gene identification proved to be effective and was validated by a combination of different methods. The test set of 17 sequenced alleles that was unintentionally included in this study serves as a powerful control and gives confidence in the novel results we obtained. Having eight of the nine genes in this blind test set readily identified was an important validation of our analysis pipeline. The mapping data from our collaborators, gene expression patterns 51 from the modENCODE project, GO term analysis, and phenotypes previously documented from mutants or RNAi experiments also provided evidence to support the gene identities we assigned in these collections. To obtain further confirmation for selected gene candidates, I generated deletion alleles using CRISPR-Cas9 genome editing. In addition to providing validation of gene identities, these alleles will be made available to the research community and may serve as useful tools in future studies. While the mutant alleles from the original study have been outcrossed, the genetic balancer background and additional mutations that persist can make them difficult to work with. In contrast, these new CRISPR-Cas9 deletion strains were made in a wild-type (N2) background which makes it much easier to handle them and interpret their mutant phenotypes. Furthermore, the pharyngeal GFP expression introduced by the gene editing approach is a straightforward marker for tracking the alleles in a heterozygous population. 4.2 Complications with Gene Identification Filtering steps in genome analysis pipelines are necessary for narrowing down long lists of genomic variants to a number that is manageable to study; however, this comes at the cost of potentially excluding functionally important mutations. While the analysis pipeline we used was efficient, it was not perfect. For example, by eliminating non-coding regions of the genome from consideration, variants affecting regulatory regions and introns were omitted from the results. Doing so yielded cleaner results, making it less likely to have multiple gene hits that would 52 require resolution in other ways; however, this approach omitted calls that could have led to gene identification. Two such examples in the collection are known and are described below. One gene from the test set of 17 alleles was not identified in our analysis due to an intronic mutation in one of the two alleles. This mutation, a SNV at the 5th base pair of an intron in the gene cul-2, is reported to alter a splice donor site (Sonneville and Gönczy 2004) but was eliminated by our filtering criteria. In our analysis, we used a stricter definition of splice sites and only looked for splicing defects in the first two and last two base pairs of an intron. Another intronic mutation was found in the gene Y54G2A.73. Four alleles from this complementation group were sequenced, and three of these had a common gene hit in Y54G2A.73. Upon manual inspection of this geneâs sequence in the fourth allele, we found a point mutation in the middle of a 3.2 kb intron. In our analysis, we did not look for these types of deep intronic mutations; however, there is evidence that they can disrupt splicing and some are known to cause disease (reviewed in Vaz-Drago et al. 2017). These examples suggest that the complementation groups for which we found no gene hits in common could be reexamined by relaxing the filtering criteria. However, doing so would produce an abundance of variant calls from which it can be difficult to extract meaningful results in the absence of more precise mapping data. Furthermore, there are several other possible explanations for the pairs which produced no results. Poor coverage or sequence quality at a particular locus could have eliminated relevant calls from our analysis. Additionally, complex rearrangements may have gone undetected, as structural variants can be difficult to detect with 53 short-read sequencing (Skovgaard et al. 2011; Huddleston et al. 2017). Finally, we found one example of an apparent strain mix-up in these collections, which is unsurprising given the number of strains being handled, their transport between labs, and the amount of time passed since the original data were documented. To track down the genes we were unable to identify, repeating the complementation tests and re-tooling the analysis approach is likely required. 4.3 Insights from Bioinformatic Analyses 4.3.1 Molecular Nature of the Genetic Variants In total, we identified 128 variants in 59 essential genes that lead to maternal-effect lethality or sterility. 80% of the variants we found are caused by a G/C to A/T transition, which reflects a known bias of EMS mutagenesis reported by others (Flibotte et al. 2010). The majority of these genetic variants cause missense mutations (49%) or nonsense mutations (41%). Approximately 75% of the nonsense mutations introduce a premature stop codon in the first two thirds of the protein sequence. These are likely null alleles, as transcripts with premature stop codons are targeted for degradation by the nonsense-mediated decay pathway, a process that is most effective when stop codons are distant from the 3â end of the gene (Longman et al. 2007). 4.3.2 Gene Ontology Analysis Reveals Common Themes and Gaps in Our Knowledge The underlying biological themes of the 59 essential genes were revealed by examining their GO terms. The most common BP annotation (embryonic development ending in birth or egg hatching) confirms the nature of the set, as a collection of genes that are required for embryonic development. Many of the essential genes that came out of this study are enzymes, which is 54 appropriate since metabolism is a basic requirement for life. The microscopy of terminal embryos in Appendix D demonstrates how the disruption of some of these vital enzymes affects embryonic development. The 20 GOI were associated with BP and MF annotations shared by other genes in the set, indicating that, while many of these genes were not previously known to be essential, they are likely playing roles in similar processes as known essential genes. Performing GO-term analysis also revealed that a number of the genes in this collection lacked sufficient annotation to be interpreted this way. Seven genes were omitted from the BP analysis and 33 genes were omitted from the MF analysis due either to a lack of annotation for those ontology categories or insufficient enrichment of the GO term relative to the threshold. Moreover, we found five genes about which there is little to nothing known (D2096.12, F56D5.2, K08D8.6, T22B11.1, and Y54G2A.73). For example, F56D5.2 is a gene with no associated GO terms, no known protein domains, and no orthologs in other model organisms. These wholly uncharacterized genes are intriguing candidates which may help uncover new biological processes and biochemical pathways that are evidently fundamental to life for this organism. 4.3.3 Expression Patterns Offer Insight into Gene Function The life stage-specific expression patterns (Appendix C) also provide some insight into the roles the genes in this collection play in development. 15 of the 20 GOI are highly expressed in the early embryo and hermaphrodite gonad. This is a strong indication that the gene product is passed on to the embryo from the parent and is therefore required in early development. Five of these maternal genes also have noticeable spikes in their expression during late embryonic and 55 larval stages which suggests they are pleiotropic, having zygotic roles in addition to the maternal ones. The zygotic functions of these genes must be non-essential or else a zygotic lethal, rather than maternal-effect lethal, phenotype would be observed. We also identified five genes that are most highly expressed in males and L4 hermaphrodites, as well as two genes that have prominent male expression in addition to characteristic maternal expression patterns. This led to the hypothesis that these genes are required in sperm, and indeed all but one of these genes were confirmed to have a role in male fertility. With the availability of RNA-seq data across different life stages for nearly every gene in the C. elegans genome (Hillier et al. 2009; Gerstein et al. 2010, 2014; Boeck et al. 2016), screening through these data for characteristic expression patterns may be a useful approach for identifying sterile and maternal-effect lethal genes that remain to be discovered. 4.4 Relevance Beyond C. elegans In this collection of 59 essential genes, there are 47 genes (80%) with human orthologs; a two-fold enrichment when compared to all C. elegans genes, 41% of which have human orthologs (Kim et al. 2018). This is in line with previous findings that essential genes are more often conserved than non-essential genes (Hughes 2002; Jordan et al. 2002; Georgi et al. 2013). Essential genes in model organisms are often associated with human diseases (Culetto and Sattelle 2000; Silverman et al. 2009; Dickerson et al. 2011; Qin et al. 2018), making the alleles identified in this study potentially relevant to understanding human health. Indeed, there are OMIM disease phenotypes associated with a number of the human orthologs identified in Table 3.1. Notably, bckd-1A, cpt-2, dlat-1, and F21D5.1 are genes in the GOI set whose human 56 orthologs are associated with inborn errors of metabolism (IEM; iembase.org). IEM are a group of approximately 1,000 congenital metabolic disorders which result from enzyme deficiencies and disruptions to metabolic pathways and cause a wide range of phenotypes including neurodevelopmental defects (Ferreira et al. 2019). Collectively, these disorders account for 0.4% of childhood deaths worldwide (Waters et al. 2018). Novel mutant alleles in C. elegans may help us better understand genetic disorders by providing new opportunities to interrogate gene function, explore genetic interactions, and screen prospective therapeutics. As reproductive genes are known to evolve quickly (Wyckoff et al. 2000; Swanson and Vacquier, 2002; Haerty et al. 2007), it is unsurprising that a number of genes in this collection do not share sequence similarity with human genes. These genes represent important gaps in our understanding of C. elegans biology; they are essential for reproduction and development, yet in many cases, their function remains a mystery. Studying these genes will be key to gaining a more complete understanding of the molecular basis of form and function in this model organism and may provide information about similar processes in other species. Indeed, genes with little sequence similarity can still perform conserved roles in different species. For example, some long intervening noncoding RNAs (lincRNAs) in zebrafish and mammals have limited sequence similarity but conserved functionality (Ulitsky et al. 2011), and sperm-specific transmembrane proteins encoded by C. elegans spe-45 and mouse Izumo1 perform analogous functions to enable fertilization in their respective species (Nishimura et al. 2015; Singaravelu et al. 2015). 57 Finally, essential genes that are nematode-specific are important to nematode biology in general and are particularly relevant in parasitic nematology. We found three genes in our GOI list (F56D5.2, perm-5, and T22B11.1) that have orthologs in parasitic nematode species and not in other phyla. With growing anthelminthic drug resistance around the world (Jabbar et al. 2006), novel management strategies are needed to combat parasitic nematodes which infect crops, livestock, and people worldwide (Nicol et al. 2011; Wolstenholme et al. 2004; Hotez et al. 2008). Essential genes are desirable targets for drug development, yet identifying such genes in parasites experimentally is difficult (Kumar et al. 2007; Doyle et al. 2010). As a free-living nematode, C. elegans is a widely used model for genetically intractable parasitic species (Bürglin et al. 1998; Hashmi et al. 2001). Our identification of novel lethal and sterile alleles with orthologs in parasitic nematodes may provide new opportunities to explore management strategies. 4.5 Discovery of Genes Involved in Male Fertility Fertilization requires specialized interactions of sperm and egg, but the molecular underpinnings of this process are still largely unknown (Krauchunas et al. 2016). C. elegans has emerged as an important model system in which to study fertility and several mutants have been isolated, though many genes remain to be identified (Singaravelu and Singson 2011). While there is much diversity in the fertilization process across different species, common themes exist. Indeed, C. elegans genes and mammalian genes with conserved roles have been identified (reviewed in Nishimura and LâHernault 2017). Furthermore, the genetic pathways that regulate C. elegans 58 fertilization may have relevance to human fertility as well as various human diseases (Han et al. 2010). Mutants defective in male fertility have been isolated through forward genetic screens in C. elegans (for example, Singson et al. 1998; Xu and Sternberg 2003; Chatterjee et al. 2005). The genetic screens by our collaborators and the analysis in this study have also proven successful in uncovering genes with sperm-specific defects, as discussed below. As several studies have shown that genes expressed in sperm are largely insensitive to RNAi (Fraser et al. 2000; Gönczy et al. 2000; Reinke et al. 2004; del Castillo-Olivares et al. 2009; Zhu et al. 2009; Ma et al. 2014), these types of genes are particularly difficult to identify in high-throughput RNAi screens. As suggested above, searching for genes with male-dominated expression patterns in existing RNA-seq data sets may be an effective way to identify the remaining sperm-expressed genes that have yet to be uncovered. There are seven genes in the GOI set which appear to be involved in sperm production and/or function (bckd-1A, C34D4.4, C56A3.8, F56D5.2, nstp-2, T22B11.1, and Y54G2A.73). These genes, which have pronounced expression levels in males and L4 hermaphrodites, were confirmed to have roles in male fertility. While the mutant hermaphrodites lay unfertilized oocytes (5 genes) or dead eggs (2 genes), this phenotype was rescued in 14 of the 16 alleles by the introduction of wild-type sperm through mating. The two alleles that were not rescued through mating disagreed with their allele pairs within the same complementation group. In a separate project carried out by an undergraduate student under my supervision (Negin Khosravi), one of these 59 discrepancies, F56D5.2(t1744) and F56D5.2(t1791), was resolved when she showed that a second mutation in a nearby essential gene was likely responsible for the inability of one strain to be rescued (data not shown). It is possible that the other such discrepancy, nstp-2(t1772) and nstp-2(t1835), could be explained by a similar circumstance. Examination of the whole genome sequencing data for GE2091(t1772) revealed missense mutations in several genes within the mapped region that are annotated with lethal or sterile phenotypes. While it is not known whether any one of these mutations confers maternal-effect lethality, this is one possible explanation for the conflicting results between the pair of nstp-2 alleles. Mating assays for the nstp-2 deletion mutant generated by CRISPR-Cas9 genome editing will be conducted when the allele has been validated by PCR. Most of these sperm-expressed genes are uncharacterized and this is the first reporting of their involvement in male fertility. With a keen interest from members of the research community, we have already begun working with collaborators who are interested in studying these mutants in more depth. For Negin Khosraviâs undergraduate research project, we created a transcriptional GFP reporter strain for F56D5.2 and confirmed its expression in sperm. We are also working on an endogenously tagged GFP strain to share with our collaborators. 4.6 Interpreting Terminal Phenotypes of Maternal-Effect Lethal Mutants The catalogue of terminal phenotypes (Appendix D) created in this study provides a window into the roles these genes play in C. elegans development. Some of the phenotypes I observed corroborate previously observed phenotypes from RNAi studies. For example, RNAi knockdown 60 has shown that DLAT-1 is an enzyme involved in metabolic processes that are required for cell division in one-cell C. elegans embryos (Rahman et al. 2014). We uncovered two alleles of dlat-1 in this study (t2035 and t2056) in which most embryos arrest at the one- to four-cell stage (Figure 3.6A and supplemental Figures D.15 and D.16). The mutant alleles presented here can confirm previously reported phenotypes and serve as new genetic tools for continuing the study of essential gene function. I also found alleles for six genes that exhibited an osmotic integrity defective (OID) phenotype, resulting in embryos that filled the eggshell completely or burst in distilled water. The C. elegans eggshell consists of multiple layers, including a chitin layer that maintains the ovoid shape of the egg and a lipid-rich permeability layer that protects the embryo from the external environment, maintaining its osmotic integrity (Olson et al. 2012). More than 100 genes have been identified in RNAi screens as important for the osmotic integrity of developing embryos (reviewed in Stein and Golden 2018). Some of these genes have roles in lipid metabolism (Rappleye et al. 2003; Benenati et al. 2009), cellular trafficking (Rappleye et al. 1999), and chitin synthesis (Johnston et al. 2006). Four of the six genes identified with OID mutants in this study have been previously implicated in osmotic sensitivity: dgtr-1 is involved in lipid biosynthesis (Carvalho et al. 2011; Olson et al. 2012), trcs-1 is involved in lipid metabolism and membrane trafficking (Green et al. 2011); perm-5 is predicted to have lipid binding activity; and F21D5.1 is an ortholog of human PGM3, an enzyme involved in the hexosamine pathway which generates substrates for chitin synthase. I found OID mutants for two additional genes that were not previously characterized with this phenotype, bckd-1A and D2096.12. bckd-1A is a component of the branched-chain 61 alpha-keto dehydrogenase complex, which is involved in fatty acid biosynthesis (Kniazeva et al. 2004); this may be indicative of a role in generating or maintaining the lipid-rich permeability barrier. D2096.12 is a Caenorhabditis-specific gene with no known protein domains. Elucidating the function of this uncharacterized gene may lead to new insights about the biochemistry of eggshell formation and permeability in C. elegans embryos. Most of the mutant strains I examined arrested around the 100- to 200-cell stage as a seemingly disorganized group of cells (for example, Figure 3.6C). Others developed into two-fold or later stage embryos that moved inside the eggshell but did not hatch (for example, Figure 3.6B). The terminal phenotypes reveal how long the embryo can persist without the maternal contribution of gene products, and the developmental defects that ensue. Further investigation of these genes might include using a fluorescent marker or antibody staining to determine whether specific tissues fail to differentiate in the mutant embryos. Multifocal, time-lapse (4D) microscopy could also be used to perform automated tracking of the cell lineage in mutant embryos (Thomas et al. 1996; Schnabel et al. 1997; Bao et al. 2006). This approach may show alterations in cell division, embryonic patterning, and organogenesis that are not immediately apparent in the snapshots of terminal phenotypes presented here. The wealth of single-cell transcriptome data that has recently been made available for C. elegans embryos (Tintori et al. 2016; Packer et al. 2019) could also be useful in determining gene function by providing clues about where and when gene products appear and what other genes may interact with them. It is our hope that the alleles and phenotypes presented here will serve as a starting point and guide future studies to help elucidate the specific roles these genes play in embryogenesis. 62 Chapter 5: Conclusion We brought together legacy mutant collections from decades-old forward genetic screens and modern, high-throughput sequencing technology to look for essential genes in C. elegans. With this approach, we identified 59 genes represented by 125 mutant alleles with sterile and maternal-effect lethal phenotypes. Many of these genes have only previously been studied with RNAi knockdown. These alleles, along with a preliminary characterization of the mutants, offer an entry point to study gene function in greater detail. Notably, several genes in this collection are completely uncharacterized and are being reported here as essential genes for the first time. By identifying novel essential genes and providing mutant alleles for further study, we are contributing to ongoing efforts to achieve a more complete understanding of gene function in C. elegans. This work will provide insight into fundamental processes in nematode biology and may inform our understanding of human development and disease. All of the alleles examined in this study will be made available to the research community through the Caenorhabditis Genetics Center (cgc.umn.edu) and are anticipated to serve as a valuable resource in the years to come. The wealth of material uncovered in these legacy collections may inspire similar explorations of frozen mutant collections which might harbour interesting alleles that await identification. 63 References Altschul, S. F., W. Gish, W. Miller, E. W. Myers and D. J. Lipman, 1990 Basic local alignment search tool. J. Mol. Biol. 215: 403-410. Au, V., E. Li-Leger, G. Raymant, S. Flibotte, G. Chen et al., 2019 CRISPR/Cas9 methodology for the generation of knockout deletions in Caenorhabditis elegans. G3: Genes, Genomes, Genet. 9: 135-144. Ausländer, S., D. Ausländer and M. Fussenegger, 2017 Synthetic biologyâthe synthesis of biology. Angew. Chem. Int. Ed. 56: 6396-6419. Baker, N. E., 1987 Molecular cloning of sequences from wingless, a segment polarity gene in Drosophila: The spatial distribution of a transcript in embryos. EMBO J. 6: 1765-1773. Bao, Z., J. I. Murray, T. Boyle, S. L. Ooi, M. J. Sandel et al., 2006 Automated cell lineage tracing in Caenorhabditis elegans. Proc. Natl. Acad. Sci. U.S.A. 103: 2707-2712. Benenati, G., S. Penkov, T. Müller-Reichert, E. V. Entchev and T. V. Kurzchalia, 2009 Two cytochrome P450s in Caenorhabditis elegans are essential for the organization of eggshell, correct execution of meiosis and the polarization of embryo. Mech. Dev. 126: 382-393. Birmingham, A., E. M. Anderson, A. Reynolds, D. Ilsley-Tyree, D. Leake et al., 2006 3Ê¹ UTR seed matches, but not overall identity, are associated with RNAi off-targets. Nat. Methods 3: 199-204. Bischoff, M., and R. Schnabel, 2006 A posterior centre establishes and maintains polarity of the Caenorhabditis elegans embryo by a Wnt-dependent relay mechanism. PLoS Biol. 4: e396. Blumenstiel, J. P., A. C. Noll, J. A. Griffiths, A. G. Perera, K. N. Walton et al., 2009 Identification of EMS-induced mutations in Drosophila melanogaster by whole-genome sequencing. Genetics 182: 25-32. Boeck, M. E., C. Huynh, L. Gevirtzman, O. A. Thompson, G. Wang et al., 2016 The time-resolved transcriptome of C. elegans. Genome Res. 26: 1441-1450. Bourbon, H., G. Gonzy-Treboul, F. Peronnet, M. Alin, C. Ardourel et al., 2002 A P-insertion screen identifying novel X-linked essential genes in Drosophila. Mech. Dev. 110: 71-83. Bradley, A., K. Anastassiadis, A. Ayadi, J. F. Battey, C. Bell et al., 2012 The mammalian gene function resource: The international knockout mouse consortium. Mammalian genome 23: 580-586. Brauchle, M., K. Baumer and P. Gönczy, 2003 Differential activation of the DNA replication checkpoint contributes to asynchrony of cell division in C. elegans embryos. Curr. Biol. 13: 819-827. Brenner, S., 1974 The genetics of Caenorhabditis elegans. Genetics 77: 71-94. 64 Bürglin, T. R., E. Lobos and M. L. Blaxter, 1998 Caenorhabditis elegans as a model for parasitic nematodes. Int. J. Parasitol. 28: 395-411. C. elegans Deletion Mutant Consortium, 2012 Large-scale screening for targeted knockouts in the Caenorhabditis elegans genome. G3: Genes, Genomes, Genet. 2: 1415-1425. Carvalho, A., S. K. Olson, E. Gutierrez, K. Zhang, L. B. Noble et al., 2011 Acute drug treatment in the early C. elegans embryo. PloS one 6: e24656. Chatterjee, I., A. Richmond, E. Putiri, D. C. Shakes and A. Singson, 2005 The Caenorhabditis elegans spe-38 gene encodes a novel four-pass integral membrane protein required for sperm function at fertilization. Development 132: 2795-2808. Chen, Z., J. Stockton, D. Mathis and C. Benoist, 2006 Modeling CTLA4-linked autoimmunity with RNA interference in mice. Proc. Natl. Acad. Sci. U.S.A. 103: 16400-16405. Cheng, N. N., C. M. Kirby and K. J. Kemphues, 1995 Control of cleavage spindle orientation in Caenorhabditis elegans: The role of the genes par-2 and par-3. Genetics 139: 549-559. Chervitz, S. A., L. Aravind, G. Sherlock, C. A. Ball, E. V. Koonin et al., 1998 Comparison of the complete protein sets of worm and yeast: Orthology and divergence. Science 282: 2022-2028. Clark, D. V., T. M. Rogalski, L. M. Donati and D. L. Baillie, 1988 The unc-22 (IV) region of Caenorhabditis elegans: Genetic analysis of lethal mutations. Genetics 119: 345-353. Clevers, H., and R. Nusse, 2012 Wnt/β-catenin signaling and disease. Cell 149: 1192-1205. Cockell, M. M., K. Baumer and P. Gönczy, 2004 Lis-1 is required for dynein-dependent cell division processes in C. elegans embryos. J. Cell. Sci. 117: 4571-4582. Culetto, E., and D. B. Sattelle, 2000 A role for Caenorhabditis elegans in understanding the function and interactions of human disease genes. Hum. Mol. Genet. 9: 869-877. Deák, P., M. M. Omar, R. D. Saunders, M. Pál, O. Komonyi et al., 1997 P-element insertion alleles of essential genes on the third chromosome of Drosophila melanogaster correlation of physical and cytogenetic maps in chromosomal region 86E-87F. Genetics 147: 1697-1722. Dejima, K., S. Hori, S. Iwata, Y. Suehiro, S. Yoshina et al., 2018 An aneuploidy-free and structurally defined balancer chromosome toolkit for Caenorhabditis elegans. Cell Rep. 22: 232-241. del Castillo-Olivares, A., M. Kulkarni and H. E. Smith, 2009 Regulation of sperm gene expression by the GATA factor ELT-1. Dev. Biol. 333: 397-408. Delattre, M., S. Leidel, K. Wani, K. Baumer, J. Bamat et al., 2004 Centriolar SAS-5 is required for centrosome duplication in C. elegans. Nat. Cell Biol. 6: 656-664. Denver, D. R., K. Morris, M. Lynch and W. K. Thomas, 2004 High mutation rate and predominance of insertions in the Caenorhabditis elegans nuclear genome. Nature 430: 679-682. Dickerson, J. E., A. Zhu, D. L. Robertson and K. E. Hentges, 2011 Defining the role of essential genes in human disease. PloS one 6: e27368. 65 Dietzl, G., D. Chen, F. Schnorrer, K. Su, Y. Barinova et al., 2007 A genome-wide transgenic RNAi library for conditional gene inactivation in Drosophila. Nature 448: 151-156. Doitsidou, M., R. J. Poole, S. Sarin, H. Bigelow and O. Hobert, 2010 C. elegans mutant identification with a one-step whole-genome-sequencing and SNP mapping strategy. PloS one 5: e15435. Doyle, M. A., R. B. Gasser, B. J. Woodcroft, R. S. Hall and S. A. Ralph, 2010 Drug target prediction and prioritization: Using orthology to predict essentiality in parasite genomes. BMC Genomics 11: 222. Edgley, M. L., D. L. Baillie, D. L. Riddle and A. M. Rose, 1995 Genetic balancers, pp. 147-184 in Methods in Cell Biology. Elsevier. Ellis, H. M., and H. R. Horvitz, 1986 Genetic control of programmed cell death in the nematode C. elegans. Cell 44: 817-829. Feichtinger, R. E., 1995 Quantitative Analysis of Maternal Gene Functions of Caenorhabditis Elegans. Ph.D. Thesis, University of Vienna, Austria. Ferreira, C. R., C. D. van Karnebeek, J. Vockley and N. Blau, 2019 A proposed nosology of inborn errors of metabolism. Genet. Med. 21: 102-106. Fire, A., S. Xu, M. K. Montgomery, S. A. Kostas, S. E. Driver et al., 1998 Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391: 806-811. Flibotte, S., M. L. Edgley, I. Chaudhry, J. Taylor, S. E. Neil et al., 2010 Whole-genome profiling of mutagenesis in Caenorhabditis elegans. Genetics 185: 431-441. Fraser, A. G., R. S. Kamath, P. Zipperlen, M. Martinez-Campos, M. Sohrmann et al., 2000 Functional genomic analysis of C. elegans chromosome I by systematic RNA interference. Nature 408: 325-330. Gans, M., C. Audit and M. Masson, 1975 Isolation and characterization of sex-linked female-sterile mutants in Drosophila melanogaster. Genetics 81: 683-704. Georgi, B., B. F. Voight and M. BuÄan, 2013 From mouse to human: Evolutionary genomics analysis of human orthologs of essential genes. PLoS genet. 9: e1003484. Gerstein, M. B., J. Rozowsky, K. Yan, D. Wang, C. Cheng et al., 2014 Comparative analysis of the transcriptome across distant species. 512: 445-448. Gerstein, M. B., Z. J. Lu, E. L. Van Nostrand, C. Cheng, B. I. Arshinoff et al., 2010 Integrative analysis of the Caenorhabditis elegans genome by the modENCODE project. Science 330: 1775-1787. Giaever, G., A. M. Chu, L. Ni, C. Connelly, L. Riles et al., 2002 Functional profiling of the Saccharomyces cerevisiae genome. Nature 418: 387-391. 66 Gönczy, P., H. Schnabel, T. Kaletta, A. D. Amores, T. Hyman et al., 1999 Dissection of cell division processes in the one cell stage Caenorhabditis elegans embryo by mutational analysis. J. Cell Biol. 144: 927-946. Gönczy, P., J. Bellanger, M. Kirkham, A. Pozniakowski, K. Baumer et al., 2001 Zyg-8, a gene required for spindle positioning in C. elegans, encodes a doublecortin-related kinase that promotes microtubule assembly. Dev. Cell. 1: 363-375. Gönczy, P., C. Echeverri, K. Oegema, A. Coulson, S. J. Jones et al., 2000 Functional genomic analysis of cell division in C. elegans using RNAi of genes on chromosome III. Nature 408: 331-336. Goodwin, S., J. D. McPherson and W. R. McCombie, 2016 Coming of age: Ten years of next-generation sequencing technologies. Nat. Rev. Genet. 17: 333. Green, R. A., H. Kao, A. Audhya, S. Arur, J. R. Mayers et al., 2011 A high-resolution C. elegans essential gene network based on phenotypic profiling of a complex tissue. Cell 145: 470-482. Greenwald, I. S., P. W. Sternberg and H. R. Horvitz, 1983 The lin-12 locus specifies cell fates in Caenorhabditis elegans. Cell 34: 435-444. Hadorn, E., 1951 Developmental action of lethal factors in Drosophila, pp. 53-85 in Advances in Genetics. Elsevier. Haerty, W., S. Jagadeeshan, R. J. Kulathinal, A. Wong, K. R. Ram et al., 2007 Evolution in the fast lane: Rapidly evolving sex-related genes in Drosophila. Genetics 177: 1321-1335. Han, S. M., P. A. Cottee and M. A. Miller, 2010 Sperm and oocyte communication mechanisms controlling C. elegans fertility. Dev. Dyn. 239: 1265-1281. Harris, T. W., V. Arnaboldi, S. Cain, J. Chan, W. J. Chen et al., 2020 WormBase: A modern model organism information resource. Nucleic Acids Res. 48: D762-D767. Hashmi, S., W. Tawe and S. Lustigman, 2001 Caenorhabditis elegans and the study of gene function in parasites. Trends Parasitol. 17: 387-393. Hedgecock, E. M., J. G. Culotti and D. H. Hall, 1990 The unc-5, unc-6, and unc-40 genes guide circumferential migrations of pioneer axons and mesodermal cells on the epidermis in C. elegans. Neuron 4: 61-85. Hedgecock, E. M., J. E. Sulston and J. N. Thomson, 1983 Mutations affecting programmed cell deaths in the nematode Caenorhabditis elegans. Science 220: 1277-1279. Hedgecock, E. M., J. G. Culotti, D. H. Hall and B. D. Stern, 1987 Genetics of cell and axon migrations in Caenorhabditis elegans. Development 100: 365-382. Herman, R. K., 1978 Crossover suppressors and balanced recessive lethals in Caenorhabditis elegans. Genetics 88: 49-65. 67 Hillier, L. W., V. Reinke, P. Green, M. Hirst, M. A. Marra et al., 2009 Massively parallel sequencing of the polyadenylated transcriptome of C. elegans. Genome Res. 19: 657-666. Hillier, L. W., G. T. Marth, A. R. Quinlan, D. Dooling, G. Fewell et al., 2008 Whole-genome sequencing and variant discovery in C. elegans. Nat. Methods 5: 183. Hotez, P. J., P. J. Brindley, J. M. Bethony, C. H. King, E. J. Pearce et al., 2008 Helminth infections: The great neglected tropical diseases. J. Clin. Invest. 118: 1311-1321. Howell, A. M., S. G. Gilmour, R. A. Mancebo and A. M. Rose, 1987 Genetic analysis of a large autosomal region in Caenorhabditis elegans by the use of a free duplication. Genetics Research 49: 207-213. Howell, A. M., and A. M. Rose, 1990 Essential genes in the hDf6 region of chromosome I in Caenorhabditis elegans. Genetics 126: 583-592. Huang, D. W., B. T. Sherman and R. A. Lempicki, 2009a Bioinformatics enrichment tools: Paths toward the comprehensive functional analysis of large gene lists. Nucleic Acids Res. 37: 1-13. Huang, D. W., B. T. Sherman, X. Zheng, J. Yang, T. Imamichi et al., 2009b Extracting biological meaning from large gene lists with DAVID. Curr. Protoc. Bioinformatics 27: 13.11. 1-13.11. 13. Huddleston, J., M. J. Chaisson, K. M. Steinberg, W. Warren, K. Hoekzema et al., 2017 Discovery and genotyping of structural variation from long-read haploid genome sequence data. Genome Res. 27: 677-685. Hughes, T. R., 2002 Yeast and drug discovery. Funct. Integr. Genomics 2: 199-211. Hutter, H., and J. Suh, 2016 GExplore 1.4: An expanded web interface for queries on Caenorhabditis elegans protein and gene function. Worm 5: e1234659. Hutter, H., and R. Schnabel, 1994 Glp-1 and inductions establishing embryonic axes in C. elegans. Development 120: 2051-2064. Hutter, H., M. Ng and N. Chen, 2009 GExplore: A web server for integrated queries of protein domains, gene expression and mutant phenotypes. BMC Genomics 10: 529. Ishii, N., W. G. Wadsworth, B. D. Stern, J. G. Culotti and E. M. Hedgecock, 1992 UNC-6, a laminin-related protein, guides cell and pioneer axon migrations in C. elegans. Neuron 9: 873-881. Jabbar, A., Z. Iqbal, D. Kerboeuf, G. Muhammad, M. N. Khan et al., 2006 Anthelmintic resistance: The state of play revisited. Life Sci. 79: 2413-2431. Jackson, A. L., S. R. Bartz, J. Schelter, S. V. Kobayashi, J. Burchard et al., 2003 Expression profiling reveals off-target gene regulation by RNAi. Nat. Biotechnol. 21: 635-637. Jaramillo-Lambert, A., A. S. Fuchsman, A. S. Fabritius, H. E. Smith and A. Golden, 2015 Rapid and efficient identification of Caenorhabditis elegans legacy mutations using Hawaiian SNP-based mapping and whole-genome sequencing. G3: Genes, Genomes, Genet. 5: 1007-1019. 68 Johnsen, R. C., and D. L. Baillie, 1997 Mutation, pp. 79-95 in C. Elegans II, edited by Riddle, D.L., Blumenthal, T., Meyer, B.J., Priess, J.R. Cold Spring Harbor Laboratory Press, Cold Spring Harbor (NY). Johnsen, R. C., and D. L. Baillie, 1991 Genetic analysis of a major segment [LGV (left)] of the genome of Caenorhabditis elegans. Genetics 129: 735-752. Johnsen, R. C., and D. L. Baillie, 1988 Formaldehyde mutagenesis of the eT1 balanced region in Caenorhabditis elegans: Doseâresponse curve and the analysis of mutational events. Mutat. Res. 201: 137-147. Johnston, W. L., A. Krizus and J. W. Dennis, 2006 The eggshell is required for meiotic fidelity, polar-body extrusion and polarization of the C. elegans embryo. BMC biology 4: 35. Jordan, I. K., I. B. Rogozin, Y. I. Wolf and E. V. Koonin, 2002 Essential genes are more evolutionarily conserved than are nonessential genes in bacteria. Genome Res. 12: 962-968. Jürgens, G., E. Wieschaus, C. Nüsslein-Volhard and H. Kluding, 1984 Mutations affecting the pattern of the larval cuticle in Drosophila melanogaster. Roux Arch. Dev. Biol. 193: 283-295. Kadandale, P., I. Chatterjee and A. Singson, 2009 Germline transformation of Caenorhabditis elegans by injection, pp. 123-133 in Microinjection. Springer. Kaitna, S., H. Schnabel, R. Schnabel, A. A. Hyman and M. Glotzer, 2002 A ubiquitin C-terminal hydrolase is required to maintain osmotic balance and execute actin-dependent processes in the early C. elegans embryo. J. Cell. Sci. 115: 2293-2302. Kamath, R. S., A. G. Fraser, Y. Dong, G. Poulin, R. Durbin et al., 2003 Systematic functional analysis of the Caenorhabditis elegans genome using RNAi. Nature 421: 231-237. Kaufman, T. C., R. Lewis and B. Wakimoto, 1980 Cytogenetic analysis of chromosome 3 in Drosophila melanogaster: The homoeotic gene complex in polytene chromosome interval 84a-B. Genetics 94: 115-133. Kemphues, K., 2005 Essential Genes in WormBook. The C. elegans Research Community. Kemphues, K. J., M. Kusch and N. Wolf, 1988b Maternal-effect lethal mutations on linkage group II of Caenorhabditis elegans. Genetics 120: 977-986. Kemphues, K. J., J. R. Priess, D. G. Morton and N. Cheng, 1988a Identification of genes required for cytoplasmic localization in early C. elegans embryos. Cell 52: 311-320. Kennedy, S., D. Wang and G. Ruvkun, 2004 A conserved siRNA-degrading RNase negatively regulates RNA interference in C. elegans. Nature 427: 645-649. Kim, W., R. S. Underwood, I. Greenwald and D. D. Shaye, 2018 OrthoList 2: A new comparative genomic analysis of human and Caenorhabditis elegans genes. Genetics 210: 445-461. Kirby, C., M. Kusch and K. Kemphues, 1990 Mutations in the par genes of Caenorhabditis elegans affect cytoplasmic reorganization during the first cell cycle. Dev. Biol. 142: 203-215. 69 Kniazeva, M., Q. T. Crawford, M. Seiber, C. Wang and M. Han, 2004 Monomethyl branched-chain fatty acids play an essential role in Caenorhabditis elegans development. PLoS biology 2: e257. Krauchunas, A. R., M. R. Marcello and A. Singson, 2016 The molecular complexity of fertilization: Introducing the concept of a fertilization synapse. Mol. Reprod. Dev. 83: 376-386. Krumlauf, R., 1994 Hox genes in vertebrate development. Cell 78: 191-201. Kulkarni, M. M., M. Booker, S. J. Silver, A. Friedman, P. Hong et al., 2006 Evidence of off-target effects associated with long dsRNAs in Drosophila melanogaster cell-based assays. Nat. Methods 3: 833-838. Kumar, S., K. Chaudhary, J. M. Foster, J. F. Novelli, Y. Zhang et al., 2007 Mining predicted essential genes of Brugia malayi for nematode drug targets. PloS one 2: e1189. Kumsta, C., and M. Hansen, 2012 C. elegans rrf-1 mutations maintain RNAi efficiency in the soma in addition to the germline. PloS one 7: e35428. Lewis, E. B., 1978 A gene complex controlling segmentation in Drosophila, pp. 205-217 in Genes, Development and Cancer. Springer. Li, H., and R. Durbin, 2009 Fast and accurate short read alignment with BurrowsâWheeler transform. Bioinformatics 25: 1754-1760. Li, H., B. Handsaker, A. Wysoker, T. Fennell, J. Ruan et al., 2009 The sequence alignment/map format and SAMtools. Bioinformatics 25: 2078-2079. Li, L., C. J. Stoeckert and D. S. Roos, 2003 OrthoMCL: Identification of ortholog groups for eukaryotic genomes. Genome Res. 13: 2178-2189. Li, Z., F. J. Vizeacoumar, S. Bahr, J. Li, J. Warringer et al., 2011 Systematic exploration of essential yeast gene function with temperature-sensitive mutants. Nat. Biotechnol. 29: 361. Linard, B., J. D. Thompson, O. Poch and O. Lecompte, 2011 OrthoInspector: Comprehensive orthology analysis and visual exploration. BMC Bioinformatics 12: 11. Linard, B., A. Allot, R. Schneider, C. Morel, R. Ripp et al., 2015 OrthoInspector 2.0: Software and database updates. Bioinformatics 31: 447-448. Longman, D., R. H. Plasterk, I. L. Johnstone and J. F. Cáceres, 2007 Mechanistic insights and identification of two novel factors in the C. elegans NMD pathway. Genes Dev. 21: 1075-1085. Louvi, A., and S. Artavanis-Tsakonas, 2012 Notch and disease: A growing field. Semin. Cell Dev. Biol. 23: 473-480. Ma, X., Y. Zhu, C. Li, P. Xue, Y. Zhao et al., 2014 Characterisation of Caenorhabditis elegans sperm transcriptome and proteome. BMC Genomics 15: 168. Ma, Y., A. Creanga, L. Lum and P. A. Beachy, 2006 Prevalence of off-target effects in Drosophila RNA interference screens. Nature 443: 359-363. 70 Maeda, I., Y. Kohara, M. Yamamoto and A. Sugimoto, 2001 Large-scale analysis of gene function in Caenorhabditis elegans by high-throughput RNAi. Curr. Biol. 11: 171-176. Mango, S. E., C. J. Thorpe, P. R. Martin, S. H. Chamberlain and B. Bowerman, 1994 Two maternal genes, apx-1 and pie-1, are required to distinguish the fates of equivalent blastomeres in the early Caenorhabditis elegans embryo. Development 120: 2305-2315. Mathies, L. D., G. G. Blackwell, M. K. Austin, A. C. Edwards, B. P. Riley et al., 2015 SWI/SNF chromatin remodeling regulates alcohol response behaviors in Caenorhabditis elegans and is associated with alcohol dependence in humans. Proc. Natl. Acad. Sci. U S A. 112: 3032-3037. McGinnis, W., R. L. Garber, J. Wirz, A. Kuroiwa and W. J. Gehring, 1984 A homologous protein-coding sequence in Drosophila homeotic genes and its conservation in other metazoans. Cell 37: 403-408. McKim, K. S., M. F. Heschl, R. E. Rosenbluth and D. L. Baillie, 1988 Genetic organization of the unc-60 region in Caenorhabditis elegans. Genetics 118: 49-59. McKim, K. S., T. Starr and A. M. Rose, 1992 Genetic and molecular analysis of the dpy-14 region in Caenorhabditis elegans. Mol. Gen. Genet. 233: 241-251. Mello, C. C., J. M. Kramer, D. Stinchcomb and V. Ambros, 1991 Efficient gene transfer in C. elegans: Extrachromosomal maintenance and integration of transforming sequences. EMBO J. 10: 3959-3970. Mello, C. C., B. W. Draper and J. R. Prless, 1994 The maternal genes apx-1 and glp-1 and establishment of dorsal-ventral polarity in the early C. elegans embryo. Cell 77: 95-106. Meneely, P. M., and R. K. Herman, 1979 Lethals, steriles and deficiencies in a region of the X chromosome of Caenorhabditis elegans. Genetics 92: 99-115. Metzker, M. L., 2010 Sequencing technologiesâthe next generation. Nat. Rev. Genet. 11: 31-46. Mohler, J. D., 1977 Developmental genetics of the Drosophila egg I. identification of 59 sex-linked cistrons with maternal effects on embryonic development. Genetics 85: 259-272. Morton, D. G., J. M. Roos and K. J. Kemphues, 1992 Par-4, a gene required for cytoplasmic localization and determination of specific cell types in Caenorhabditis elegans embryogenesis. Genetics 130: 771-790. Moskowitz, I. P., S. B. Gendreau and J. H. Rothman, 1994 Combinatorial specification of blastomere identity by glp-1-dependent cellular interactions in the nematode Caenorhabditis elegans. Development 120: 3325-3338. Nicol, J. M., S. J. Turner, D. L. Coyne, L. Den Nijs, S. Hockland et al., 2011 Current nematode threats to world agriculture, pp. 21-43 in Genomics and Molecular Genetics of Plant-Nematode Interactions. Springer. 71 Nieto, C., J. Almendinger, S. Gysi, E. Gómez-Orte, A. Kaech et al., 2010 Ccz-1 mediates the digestion of apoptotic corpses in C. elegans. J. Cell. Sci. 123: 2001-2007. Nishimura, H., and S. W. LâHernault, 2017 Spermatogenesis. Curr. Biol. 27: R988-R994. Nishimura, H., T. Tajima, H. S. Comstra, E. J. Gleason and S. W. LâHernault, 2015 The immunoglobulin-like gene spe-45 acts during fertilization in Caenorhabditis elegans like the mouse Izumo1 gene. Curr. Biol. 25: 3225-3231. Nordström, K. J., M. C. Albani, G. V. James, C. Gutjahr, B. Hartwig et al., 2013 Mutation identification by direct comparison of whole-genome sequencing data from mutant and wild-type individuals using k-mers. Nat. Biotechnol. 31: 325. Norris, A. D., H. Kim, M. P. Colaiacovo and J. A. Calarco, 2015 Efficient genome editing in Caenorhabditis elegans with a toolkit of dual-marker selection cassettes. Genetics 201: 449-58. Nüsslein-Volhard, C., and E. Wieschaus, 1980 Mutations affecting segment number and polarity in Drosophila. Nature 287: 795-801. Nüsslein-Volhard, C., H. G. Frohnhofer and R. Lehmann, 1987 Determination of anteroposterior polarity in Drosophila. Science 238: 1675-1681. Nüsslein-Volhard, C., E. Wieschaus and H. Kluding, 1984 Mutations affecting the pattern of the larval cuticle in Drosophila melanogaster. Roux Arch. Dev. Biol. 193: 267-282. Oh, S., T. Kingsley, H. Shin, Z. Zheng, H. Chen et al., 2003 A P-element insertion screen identified mutations in 455 novel essential genes in Drosophila. Genetics 163: 195-201. Olson, S. K., G. Greenan, A. Desai, T. Müller-Reichert and K. Oegema, 2012 Hierarchical assembly of the eggshell and permeability barrier in C. elegans. J. Cell Biol. 198: 731-748. Packer, J. S., Q. Zhu, C. Huynh, P. Sivaramakrishnan, E. Preston et al., 2019 A lineage-resolved molecular atlas of C. elegans embryogenesis at single-cell resolution. Science 365: eaax1971. Peter, A., P. Schöttler, M. Werner, N. Beinert, G. Dowe et al., 2002 Mapping and identification of essential gene functions on the X chromosome of Drosophila. EMBO Rep. 3: 34-38. Piano, F., A. J. Schetter, M. Mangone, L. Stein and K. J. Kemphues, 2000 RNAi analysis of genes expressed in the ovary of Caenorhabditis elegans. Curr. Biol. 10: 1619-1622. Poulson, D. F., 1937 Chromosomal deficiencies and the embryonic development of Drosophila melanogaster. Proc. Natl. Acad. Sci. U.S.A. 23: 133-137. Priess, J. R., H. Schnabel and R. Schnabel, 1987 The glp-1 locus and cellular interactions in early C. elegans embryos. Cell 51: 601-611. Qin, Z., R. Johnsen, S. Yu, J. S. Chu, D. L. Baillie et al., 2018 Genomic identification and functional characterization of essential genes in Caenorhabditis elegans. G3: Genes, Genomes, Genet. 8: 981-997. 72 Rahman, M. M., S. Rosu, D. Joseph-Strauss and O. Cohen-Fix, 2014 Down-regulation of tricarboxylic acid (TCA) cycle genes blocks progression through the first mitotic division in Caenorhabditis elegans embryos. Proc. Natl. Acad. Sci. U.S.A. 111: 2602-2607. Rappleye, C. A., A. Tagawa, N. Le Bot, J. Ahringer and R. V. Aroian, 2003 Involvement of fatty acid pathways and cortical interaction of the pronuclear complex in Caenorhabditis elegans embryonic polarity. BMC developmental biology 3: 8. Rappleye, C. A., A. R. Paredez, C. W. Smith, K. L. McDonald and R. V. Aroian, 1999 The coronin-like protein POD-1 is required for anteriorâposterior axis formation and cellular architecture in the nematode Caenorhabditis elegans. Genes Dev. 13: 2838-2851. Reinke, V., I. San Gil, S. Ward and K. Kazmer, 2004 Genome-wide germline-enriched and sex-biased expression profiles in Caenorhabditis elegans. Development 131: 311-323. Reiter, L. T., L. Potocki, S. Chien, M. Gribskov and E. Bier, 2001 A systematic analysis of human disease-associated gene sequences in Drosophila melanogaster. Genome Res. 11: 1114-1125. Remm, M., C. E. Storm and E. L. Sonnhammer, 2001 Automatic clustering of orthologs and in-paralogs from pairwise species comparisons. J. Mol. Biol. 314: 1041-1052. Rogalski, T. M., D. G. Moerman and D. L. Baillie, 1982 Essential genes and deficiencies in the unc-22 IV region of Caenorhabditis elegans. Genetics 102: 725-736. Roth, A. C., G. H. Gonnet and C. Dessimoz, 2008 Algorithm of OMA for large-scale orthology inference. BMC Bioinformatics 9: 518. Rubin, G. M., M. D. Yandell, J. R. Wortman, G. L. Gabor, C. R. Nelson et al., 2000 Comparative genomics of the eukaryotes. Science 287: 2204-2215. Rüegger, S., T. S. Miki, D. Hess and H. GroÃhans, 2015 The ribonucleotidyl transferase USIP-1 acts with SART3 to promote U6 snRNA recycling. Nucleic Acids Res. 43: 3344-3357. Salzberg, A., S. N. Prokopenko, Y. He, P. Tsai, M. Pál et al., 1997 P-element insertion alleles of essential genes on the third chromosome of Drosophila melanogaster: Mutations affecting embryonic PNS development. Genetics 147: 1723-1741. Sarin, S., S. Prabhu, M. M. O'meara, I. Pe'er and O. Hobert, 2008 Caenorhabditis elegans mutant allele identification by whole-genome sequencing. Nat. Methods 5: 865-867. Sawa, H., H. Kouike and H. Okano, 2000 Components of the SWI/SNF complex are required for asymmetric cell division in C. elegans. Mol. Cell 6: 617-624. Schnabel, R., H. Hutter, D. Moerman and H. Schnabel, 1997 Assessing normal embryogenesis in Caenorhabditis elegans using a 4D microscope: Variability of development and regional specification. Dev. Biol. 184: 234-265. Schneeberger, K., and D. Weigel, 2011 Fast-forward genetics enabled by new sequencing technologies. Trends Plant Sci. 16: 282-288. 73 Schneeberger, K., S. Ossowski, C. Lanz, T. Juul, A. H. Petersen et al., 2009 SHOREmap: Simultaneous mapping and mutation identification by deep sequencing. Nat. Methods 6: 550-551. Schüpbach, T., and E. Wieschaus, 1989 Female sterile mutations on the second chromosome of Drosophila melanogaster. I. maternal effect mutations. Genetics 121: 101-117. Schüpbach, T., and E. Wieschaus, 1986 Maternal-effect mutations altering the anterior-posterior pattern of the Drosophila embryo. Roux Arch. Dev. Biol. 195: 302-317. Shah, N., and S. Sukumar, 2010 The Hox genes and their roles in oncogenesis. Nat. Rev. Cancer 10: 361-371. Shi, J., E. Wang, J. P. Milazzo, Z. Wang, J. B. Kinney et al., 2015 Discovery of cancer drug targets by CRISPR-Cas9 screening of protein domains. Nat. Biotechnol. 33: 661-667. Shiu, P. K., and C. P. Hunter, 2017 Early developmental exposure to dsRNA is critical for initiating efficient nuclear RNAi in C. elegans. Cell reports 18: 2969-2978. Silverman, G. A., C. J. Luke, S. R. Bhatia, O. S. Long, A. C. Vetica et al., 2009 Modeling molecular and cellular aspects of human disease using the nematode Caenorhabditis elegans. Pediatr. Res. 65: 10-18. Simmer, F., C. Moorman, Van Der Linden, Alexander M, E. Kuijk, Van Den Berghe, Peter VE et al., 2003 Genome-wide RNAi of C. elegans using the hypersensitive rrf-3 strain reveals novel gene functions. PLoS biol. 1: E12. Singaravelu, G., and A. Singson, 2011 New insights into the mechanism of fertilization in nematodes. Int. Rev. Cell Mol. Biol. 289: 211-238. Singaravelu, G., S. Rahimi, A. Krauchunas, A. Rizvi, S. Dharia et al., 2015 Forward genetics identifies a requirement for the izumo-like immunoglobulin superfamily spe-45 gene in Caenorhabditis elegans fertilization. Curr. Biol. 25: 3220-3224. Singson, A., K. B. Mercer and S. W. L'Hernault, 1998 The C. elegans spe-9 gene encodes a sperm transmembrane protein that contains EGF-like repeats and is required for fertilization. Cell 93: 71-79. Skovgaard, O., M. Bak, A. Løbner-Olesen and N. Tommerup, 2011 Genome-wide detection of chromosomal rearrangements, indels, and mutations in circular chromosomes by short read sequencing. Genome Res. 21: 1388-1393. Smith, D. R., A. R. Quinlan, H. E. Peckham, K. Makowsky, W. Tao et al., 2008 Rapid whole-genome mutational profiling using next-generation sequencing technologies. Genome Res. 18: 1638-1642. Smith, H. E., A. S. Fabritius, A. Jaramillo-Lambert and A. Golden, 2016 Mapping challenging mutations by whole-genome sequencing. G3: Genes, Genomes, Genet. 6: 1297-1304. 74 Sonneville, R., and P. Gönczy, 2004 Zyg-11 and cul-2 regulate progression through meiosis II and polarity establishment in C. elegans. Development 131: 3527-3543. Sonnhammer, E. L., and G. Ãstlund, 2015 InParanoid 8: Orthology analysis between 273 proteomes, mostly eukaryotic. Nucleic Acids Res. 43: D234-D239. Sönnichsen, B., L. B. Koski, A. Walsh, P. Marschall, B. Neumann et al., 2005 Full-genome RNAi profiling of early embryogenesis in Caenorhabditis elegans. Nature 434: 462-469. Srivatsan, A., Y. Han, J. Peng, A. K. Tehranchi, R. Gibbs et al., 2008 High-precision, whole-genome sequencing of laboratory strains facilitates genetic studies. PLoS Genet. 4: e1000139. Stein, K. K., and A. Golden, 2018 The C. elegans eggshell in WormBook: The Online Review of C. Elegans Biology. The C. elegans Research Community. Stewart, H. I., N. J. O'Neil, D. L. Janke, N. W. Franz, H. M. Chamberlin et al., 1998 Lethal mutations defining 112 complementation groups in a 4.5 mb sequenced region of Caenorhabditis elegans chromosome III. MGG 260: 280-288. Sulston, J. E., and H. R. Horvitz, 1977 Post-embryonic cell lineages of the nematode, Caenorhabditis elegans. Dev. Biol. 56: 110-156. Sulston, J. E., E. Schierenberg, J. G. White and J. N. Thomson, 1983 The embryonic cell lineage of the nematode Caenorhabditis elegans. Dev. Biol. 100: 64-119. Swanson, W. J., and V. D. Vacquier, 2002 The rapid evolution of reproductive proteins. Nat. Rev. Genet. 3: 137-144. Tavernarakis, N., S. L. Wang, M. Dorovkov, A. Ryazanov and M. Driscoll, 2000 Heritable and inducible genetic interference by double-stranded RNA encoded by transgenes. Nat. Genet. 24: 180-183. The C. elegans Sequencing Consortium, 1998 Genome sequence of the nematode C. elegans: A platform for investigating biology. Science 2012-2018. Thomas, C., P. DeVries, J. Hardin and J. White, 1996 Four-dimensional imaging: Computer visualization of 3D movements in living specimens. Science 273: 603-607. Thompson, O., M. Edgley, P. Strasbourger, S. Flibotte, B. Ewing et al., 2013 The million mutation project: A new approach to genetics in Caenorhabditis elegans. Genome Res. 23: 1749-1762. Timmons, L., D. L. Court and A. Fire, 2001 Ingestion of bacterially expressed dsRNAs can produce specific and potent genetic interference in Caenorhabditis elegans. Gene 263: 103-112. Tintori, S. C., E. O. Nishimura, P. Golden, J. D. Lieb and B. Goldstein, 2016 A transcriptional lineage of the early C. elegans embryo. Dev. Cell 38: 430-444. Ulitsky, I., A. Shkumatava, C. H. Jan, H. Sive and D. P. Bartel, 2011 Conserved function of lincRNAs in vertebrate embryonic development despite rapid sequence evolution. Cell 147: 1537-1550. 75 Varshney, G. K., J. Lu, D. E. Gildea, H. Huang, W. Pei et al., 2013 A large-scale zebrafish gene knockout resource for the genome-wide study of gene function. Genome Res. 23: 727-735. Vatcher, G. P., C. M. Thacker, T. Kaletta, H. Schnabel, R. Schnabel et al., 1998 Serine hydroxymethyltransferase is maternally essential in Caenorhabditis elegans. J. Biol. Chem. 273: 6066-6073. Vaz-Drago, R., N. Custódio and M. Carmo-Fonseca, 2017 Deep intronic mutations and human disease. Hum. Genet. 136: 1093-1111. Vilella, A. J., J. Severin, A. Ureta-Vidal, L. Heng, R. Durbin et al., 2009 EnsemblCompara GeneTrees: Complete, duplication-aware phylogenetic trees in vertebrates. Genome Res. 19: 327-335. Vyas, V. K., M. I. Barrasa and G. R. Fink, 2015 A Candida albicans CRISPR system permits genetic engineering of essential genes and gene families. Sci. Adv. 1: e1500248. Waters, D., D. Adeloye, D. Woolham, E. Wastnedge, S. Patel et al., 2018 Global birth prevalence and mortality from inborn errors of metabolism: A systematic analysis of the evidence. J. Glob. Health 8: 021102. Wetterstrand, K. A., 2019 DNA Sequencing Costs: Data from the NHGRI Genome Sequencing Program (GSP). Wharton, K. A., K. M. Johansen, T. Xu and S. Artavanis-Tsakonas, 1985 Nucleotide sequence from the neurogenic locus notch implies a gene product that shares homology with proteins containing EGF-like repeats. Cell 43: 567-581. Wheeler, D. L., T. Barrett, D. A. Benson, S. H. Bryant, K. Canese et al., 2007 Database resources of the national center for biotechnology information. Nucleic Acids Res. 36: D13-D21. Wieschaus, E., C. Nüsslein-Volhard and G. Jürgens, 1984 Mutations affecting the pattern of the larval cuticle in Drosophila melanogaster. Roux Arch. Dev. Biol. 193: 296-307. Winzeler, E. A., D. D. Shoemaker, A. Astromoff, H. Liang, K. Anderson et al., 1999 Functional characterization of the S. cerevisiae genome by gene deletion and parallel analysis. Science 285: 901-906. Wolstenholme, A. J., I. Fairweather, R. Prichard, G. von Samson-Himmelstjerna and N. C. Sangster, 2004 Drug resistance in veterinary helminths. Trends Parasitol. 20: 469-476. Wyckoff, G. J., W. Wang and C. Wu, 2000 Rapid evolution of male reproductive genes in the descent of man. Nature 403: 304-309. Xu, X. S., and P. W. Sternberg, 2003 A C. elegans sperm TRP protein required for sperm-egg interactions during fertilization. Cell 114: 285-297. Yu, L., L. P. Castillo, S. Mnaimneh, T. R. Hughes and G. W. Brown, 2006 A survey of essential gene function in the yeast cell division cycle. Mol. Biol. Cell 17: 4736-4747. 76 Yuan, J., and H. R. Horvitz, 1992 The Caenorhabditis elegans cell death gene ced-4 encodes a novel protein and is expressed during the period of extensive programmed cell death. Development 116: 309-320. Yuan, J., S. Shaham, S. Ledoux, H. M. Ellis and H. R. Horvitz, 1993 The C. elegans cell death gene ced-3 encodes a protein similar to mammalian interleukin-1β-converting enzyme. Cell 75: 641-652. Zhang, M., C. Wang, T. D. Otto, J. Oberstaller, X. Liao et al., 2018 Uncovering the essential genes of the human malaria parasite Plasmodium falciparum by saturation mutagenesis. Science 360: eaap7847. Zhu, G., G. Salazar, S. A. Zlatic, B. Fiza, M. M. Doucette et al., 2009 SPE-39 family proteins interact with the HOPS complex and function in lysosomal delivery. Mol. Biol. Cell 20: 1223-1240. Zuryn, S., S. Le Gras, K. Jamet and S. Jarriault, 2010 A strategy for direct mapping and identification of mutations by whole-genome sequencing. Genetics 186: 427-430.77 Appendices Appendix A Flanking sequences for genetic variants Table A.1 Mutant alleles and flanking sequences for genetic variants Strain Allele(s) Gene Chr. Mutation Position Base Change Left Flanking Sequence (30 bp) Right Flanking Sequence (30 bp) GE2430 t2135 air-1 V SNV 8221773 C T AAAGGAAAGTTTGGAAACGTTTTCATCTCT GTGAAAAAAAGACCAAGCGAATCATAGCTT GE2337 t2095 air-1 V deletion 8223169 CAT C GTGGATTAAGGAAATGAAGCAACGCGAAGA TGAAGTTCCACTCTTCATTTCGACGCTCAC GE2314 t1724 aptf-2 IV SNV 13414105 A G ACTGGATAATCATCTCGTACAGCTGTTCCA GATCAAGTCCCAAATGAACAGCTTCCGCTT GE2289 t1836 aptf-2 IV SNV 13414263 G T TCCATTCTTCAGTTTTGGCCTTTTAAGATT CATGCGATACTCGAACTATTGAGCTTCTCC GE1958 t1726 atg-7 IV SNV 11079764 G A GGCGTGCATCTTCAAATTCTGAGAGACTCT GCGAACGGGGTTATTGTAGCTGACAGTTGA GE1936 t1738 atg-7 IV SNV 11079973 C T TCTTTCCAATTGAATATCTGGGTGCAATCT CATTTGATTAATGATAAATTAAGATCCACA GE2449 t2143 atl-1 V SNV 9635587 C T GATTTTGCAAACAATTTCCGAGAGGCAAAC ATTGCCCTGCTTCTGGATATTGGATACGGA GE2467 t2155 atl-1 V SNV 9637978 C T CGTTGCTGACATTCTTTAATTCGACAATTT ATTCAATTTCTGAAGTTCAACCAAAAATGG GE2200 t1480 bckd-1A III SNV 12969933 G A GCTCTTTTGTTCCGAAATGCATTGGCATTT GCGGCCTGAAAATGGAATTTTTGGTTTTTG GE1742 t1461 bckd-1A III SNV 12971429 G A AGCTTGTCATATAAAACGAGATGCGTCCTT ACGCTGCGAATCATACAGGATCCGATCCAT GE2206 t1514 bckd-1A III SNV 12971273 G A TTCCCAAGTCATCCGCATTTCCATAGCACT ATTCATGAAATTCTCCATAGTATATCCTCT GE2627 t1603 bckd-1A III SNV 12971305 C T TTCATGAAATTCTCCATAGTATATCCTCTC ACAGCAGGACCCCGGCTTCACGATATTGTC GE2890 t1821 C34D4.4 IV SNV 7150054 G A TTTCGTTGATGCTGATGGAAATAATCACTG AAGTTTGAAAGTGCCAAAGATATGACTCGT GE2840 t1860 C34D4.4 IV SNV 7150143 G A GGCTTGGACTTGTTGTTGGGCCTGCGGCTT GATCTTCTTTGTTGTCACTGCGTTTTTGAC GE2734 t2029 C56A3.8 V SNV 13560728 G A AACCAAGTTTGATCCCCGAAGGCAGTTCTG ATCTTATTTTGTCTATGGATTGGAAGGAGA GE2886 t2055 C56A3.8 V SNV 13560787 G A GAAATTGTTGGAGTTTTCAAACCAAAAGAC AAGAGCCATTCGCTTCTCTCAACCCAAAAT GE2487 t2149 C56A3.8 V SNV 13561369 C T TGGCTGCTATCGACAATGGATTGGCATTTC AGTTAGACATCCGGAATGCACTTCAAGATT GE2142 t2074 ccz-1 V SNV 13679756 T A GTTCTTGTATCAAGATAAACTTCTCTACTA CAACTTTCGAAACGCGATCTTCCATCACTA GE2304 t2129 ccz-1 V SNV 13680792 C T TCCGACTTATTTTCTTTTCCAGAATCATCT AATCATCTCGCGCCACTCCTTCACAGAATG GE2047 t2021 cept-2 V SNV 14349388 G A GTCCAACAGCTACTGAAACTGCACCCGCGT GGCCTATTTTCTGGCCGCATTCGGGCTTTT GE2122 t2007 cept-2 V SNV 14349747 G A TTGGAGCACATATTGTACTGGCCAGCTCAG TAAACTTAAAAAAAAACAAAAAGATTAATG GE2275 t1517 cls-2 III SNV 9055405 G A ACTGCCTGGACCTCTCAGTGAAAGATCTCC ATCGCAAATCCTGCGAGAAGCTGCGATCAC GE2357 t1527 cls-2 III SNV 9055440 G A CAAATCCTGCGAGAAGCTGCGATCACCTGT GATTCCTATTCAAACGGTTCGGGACGGATG GE2082 t2053 cpl-1 V SNV 16593886 G A GCGTGTTGTCCTTCGAGGGCTCCGGTGGCG AGAAGGCCCAGCACGATCCGCACATTCCTT GE2451 t2144 cpl-1 V SNV 16595201 G A TATTCTTGACAAATGCCTCCATATAGGTCT CTCCTCGCTCTCCGAGTACTCCTTATCAAA 78 Strain Allele(s) Gene Chr. Mutation Position Base Change Left Flanking Sequence (30 bp) Right Flanking Sequence (30 bp) GE2447 t1879 cpt-2 IV SNV 11180120 C T GCTCCAGATCCAAATCCGAAATTCAATGAT AGGTAAAATACGTTTCAGAGAAATAATGTT GE1938 t1742 cpt-2 IV SNV 11180603 G A TTTTCAGAACTCTTCCACCAAGTCTCTCAT GTTCGGTGCAGTTGCATTCAAAGCTTTCCC GE2657 t1704 cra-1 III SNV 6867181 G A GGAATGTCAAATGACCAAGTGTATCCATTT AATTACTTTTATGTCGAGCGTCTTCACGAG GE2242 t1618 cra-1 III SNV 6868737 C T GTAACCTGCATAATTAATGACATGACGCTC AAAAATAGTACGGAGCATTTCCGAAATCCT GE1929 t1729 csr-1 IV SNV 7960467 T A CATTCCAACGCAGCACGTACTTCTCAAAAA GTCAAAAAGATGCGTGACAACATCGAAAAG GE1929 t1729 csr-1 IV SNV 7961246 G A CATCTGTTGGTGGTGATATTGCCGAATATG AAAAAGACAAATCTTCATCCAAGCTTTCCG GE2452 t1897 csr-1 IV SNV 7959252 G A CTTACTTATTGCTTTCCAAATGTATCTTCA CGAAAGATATTTGTCGCAGTCGAAAGGTCT GE2595 t1662 t1718 cup-5 III SNV 7585568 C T ATATCGACGGTCCAATTCAAATTTCGACTA GAACAATTCATTATTCACCAACAGCCGGAG GE2355 t1528 cup-5 III SNV 7590536 G A AAAGTTAATAATTATATCTCATTCATTCCA GATCGCTACTCAGACGGTCTTCGAGCCATC GE2345 t1525 cyk-3 III SNV 6020590 C T CTTGTTGGATTGTGTAGAGTAGAAGAAGTA AATCGAGTCAGTCAAGAAACATTTCAGAAA GE2352 t1535 cyk-3 III SNV 6022863 G A AAGCTGATGAAGAAAAAGCTGAAAGATCAT GACGGAGTACGAAAAACAAAATGAGTCACT GE2499 t1877 D2096.12 IV SNV 8363937 C T CCTCAAATAAATACAGTTCAATCTGTACAA AGAAACCATCAGCGAGTGCCACTGGCAGAG GE2407 t1906 D2096.12 IV SNV 8365654 T A TGGATGAACTGACTGAGAAAGGAGAGTTAT GGCGTTTTTTGAATTAACAAGTGACGGTGG GE2135 t2043 dgtr-1 V SNV 6497335 G A ATCAAGTAGAAAAATTTTAAAAATATTGCA GTTCCATTCCACGTCTTGTTCAATACTCGA GE2063 t2042 dgtr-1 V SNV 6498186 G A CCGTTCCGAAAACCAGTTACAACAGTCATG GAAGGCCAATTCGGTAAGAAATTTTTTTAG GE2028 t1801 dif-1 IV SNV 7552230 A C TTCTCCAGAGAATTTCTTTTTTAGATATTC TATACTGAAAGATAAGCGGCTGAAGCTGGG GE1932 t1732 dif-1 IV SNV 7552641 C T CCGAAAAAAACTGCAAAAAGTGGAGAAACA CAACAAGTGGAGCTGCCATTCCTTTGTAGA GE2612 t1676 div-1 III SNV 10245480 G A CCGTTGGCTCCAATGGATACATGCACTGCT AGTGAGAAGATGACTGCTGAGACGGGCGAC GE2577 t1642 div-1 III SNV 10248544 C T GTTAAAATCTTCCAAATCGAACTCGTATGC ATTTTATGACGGGAAAAAGGAAGGAAATTG GE2335 t2056 dlat-1 V SNV 14445907 G A CGAGGTTCGAGACGGTGAAAGTACCTCCTT GAACTCATGAGGCTGCAATTTTCCTTCGCG GE2541 t2035 dlat-1 V SNV 14446981 G A GTTCCGAGTTCCATAGTTGGAGAAAGTGCT GGAGAGCTACCCGATTGTGCTTTGGCAGAT GE2402 t1940 F21D5.1 IV SNV 8727315 C T TCATCAATGAAACTGTTGGGGATGCTTTTG AGATTTGCTTGCCGTGGAAGCTGTTCTCCG GE2445 t1935 F21D5.1 IV SNV 8727668 C T GTTGAAAATACATTGCAACTCGGAAAATCT TCGAACAAGTCGTTCTCAACCTTTGCAACT GE2837 t1791 F56D5.2 IV SNV 9397791 G A ATTCAATGTGTTACCTGAAGTAATTAATTT ACTCAAAAGCTGTGGGCAATACTTTTCAGC GE2881 t1744 F56D5.2 IV SNV 9398158 G A TTTGTAAGAGTTTTGAGGCGGTAAAGTGCA ATAGCTCACCAAGAGAGACAGGCTGAAAAA GE1715 t1436 gsp-2 III SNV 7337087 C T AACTACTTGTTCCTCGGCGATTATGTGGAC GAGGGAAACAATCGTTGGAAACTATCTGTC GE2360 t1481 gsp-2 III SNV 7337383 G A TCGACGAAAAGATCTTCTGCTGTCACGGAG ACTTTCTCCAGATCTTCAGTCTATGGAGCA GE2545 t1577 gsr-1 III SNV 3652401 G A TTCCTTCTCACACCAGTCGCAATTGCTGCC GACGTCGACTCTCTCATCGTCTGTTCAATG GE2644 t1594 gsr-1 III SNV 3652407 C T CTCACACCAGTCGCAATTGCTGCCGGACGT GACTCTCTCATCGTCTGTTCAATGGTGAAA GE2583 t1654 hcp-3 III SNV 9615498 G A ACTGAATATCCGTTGTCATGAGTGTGACAC TTTCGCATGGGTGGATATAAGAGACGATCC GE2692 t1717 hcp-3 III SNV 9615555 C T ATCCTTCGAACATTTCGACCAAAAATGCTT CGCCGCTTCTTGAAGAGCACTGATGGCGTC GE2034 t1808 K08D8.6 IV SNV 12887810 G A ACATTGTTCAGCACAGTTTTCTGTTCAGAA TAATCTTAACTGTAAGAGATGCACCTCCAA GE2035 t1809 K08D8.6 IV SNV 12889703 T A AACGAACTTAGTAACGTAATCCACTGAATC TGCATTGTCAAAAGCGATGACACATCTGTA 79 Strain Allele(s) Gene Chr. Mutation Position Base Change Left Flanking Sequence (30 bp) Right Flanking Sequence (30 bp) GE2455 t1914 klp-18 IV SNV 7040335 T C ACGATTGAAACTGATAATGGATCCAAAAGA ATGAGCTCCATCATATCTTTGATAGCTCTT GE2000 t1795 klp-18 IV SNV 7041203 G A AATATGCACCCGGATCGTGATTTTGCACAG AAAGCAACTCCACCTTGATGTTCGCTCAGT GE2367 t1563 klp-19 III SNV 13306451 A T CTCTCGGATCCAGCCAAATCGACCAATTGG GCTTCGCGCTGAATGCGCTATCACACTCTG GE2367 t1563 klp-19 III SNV 13306457 G A GATCCAGCCAAATCGACCAATTGGAGCTTC CGCTGAATGCGCTATCACACTCTGCAGTGG GE2264 t1628 klp-19 III SNV 13306872 C T TGTCTTCAGTGCCCATGGTGTAGGTCTTTC ACTTCCAGTTTGTCCATAAGCCAACACTGT GE2003 t1817 let-99 IV SNV 12569291 C T AATCGTCTCGAAGACTTGAATGCTCCTCTT AATTCGATGAATGCTCTCTTATTGTTAATC GE2514 t1912 let-99 IV SNV 12570199 C T CAACAAGGAATGCTGATAACCGCTGTCCTC TCGCAAACTTTCAATCTCTTTTTGCAGTAC GE2730 t1550 lis-1 III SNV 13375376 C T CAGTTTCTGTGTCTCTGGCGGGCGAGGGAT CAGTCTGCGGCTTGACGCTTGTCTCTGAAA GE2653 t1698 lis-1 III SNV 13375401 C T GGGATCCAGTCTGCGGCTTGACGCTTGTCT TGAAAATTAATTTGTTGGAGGGGGCGAGTT GE2130 t1765 mbk-2 IV SNV 13033086 C T CGAGATCAAGCACATGCTCAGCACGAGCTC TTCAACTCCGATTGGGCGCAAGTCAACAAG GE2503 t1888 mbk-2 IV SNV 13033644 C T GAAAGGCATTGTCTCAAAGCAACGGCAATT CAGCAATAGTGTGATTGTGTGGTCCACCTT GE2740 t1576 mel-32 III SNV 6440655 C T ATTATCCATATGCTCGACTATTTCAATTTC GTAATCACAAGTGCATCACGTTTGAGCTTC GE1731 t1456 mel-32 III SNV 6440831 C T CGCGTACTTTCATCACATCGAAAGGATATC ACGTTATTGCACGGCCACATCGATGCCAGA GE1999 t1793 mex-5 IV SNV 13354014 T G TCCATATTATCAAGATGCCAACGGACAGTA GTGCAGGTTCCCGCGAACGGATCAATGGCT GE2093 t1800 mex-5 IV SNV 13354478 T A TGTTTTTCAGTGTTATGGAGCATGACGAGT GCCGATCGATGAGATCTCAAAGATCACCCT GE2511 t2162 mom-2 V SNV 8356808 T G ATTTCTGGACTTCGAAGTGCTCTTCACACT GCGAGTACACATTTCAACGAGAAGCATGGA GE2523 t2180 mom-2 V SNV 8357121 T C CCTTTGCAGGGCTTAATCGACGATTGTGGA GCGGAGAAACACCACAAGGATCTGGAAGTG GE2497 t2137 mre-11 V SNV 10735712 G A GGTGCATGGCTCTCCGTGGACGATTTTGAT GAGGACAAACAAATTAAACCAATCCTCGGC GE2103 t2092 mre-11 V SNV 10736080 A G TGATTAAAATTGACAGAAGCATCTGAGAGA ACTCCAACGCGATTGGATTCCCATTCAAGC GE2091 t1772 nstp-2 IV SNV 6604731 A T GGATTCTTGAAATAGACAATGGAAACGATA GTGAGAGGAATTTCCGAAGAGTAACAACCA GE2288 t1835 nstp-2 IV SNV 6605266 C T TTGACGTGGCAAGTGTGCAAATGACAATTC GATGGTAATAGCGAATACGGAAATGTATTT GE2391 t1932 perm-5 IV SNV 5696931 A T CAATCGCCACATTATTAGGATCAATTCCAC GAATTTGCTCAATTTCAGGCCTTTTCCATA GE2453 t1900 perm-5 IV SNV 5698096 A G CTTGGAAATAGCACTCGGAGCAGAGAGTTG GAGAAATTCACGAGAATCTGGAGGAATAAT GE2237 t1614 pod-1 III SNV 13518266 G A TCAAAAACTCACTTCAGTTTCTGATGGAAC CGATTCCTTGACTTCCAACAGGCGACATAA GE2605 t1674 pod-1 III SNV 13518357 G A CATAGATCACCTCGAACATGTTCACAAATC ATCACCTTTTCCGCTAAGGAAAAGCACATT GE3128 t2177 pos-1 V SNV 8414544 G A AAATTTCTTATTTTATGACTGGAATTTTCA CCTCGTGGGAGAAATCATCCCAAGTACAAG GE2101 t2080 pos-1 V SNV 8414579 T A GTGGGAGAAATCATCCCAAGTACAAGACGG TCTTTGCGACAAGTTCTCGATGACTGGAAA GE2517 t2175 rad-50 V SNV 12247914 T A TTGGTAAAAATAATCAGGAACGACAGGAGT GTCGCAAAAGAAAACTGAATTCGAGAACAG GE2476 t2147 rad-50 V SNV 12250324 T A ATTATCGTGATGCTATCATAGAACTGGCAA TACTAAAGAATCTATCTCTGATTTAACGAA GE2189 t1750 rad-51 IV SNV 10282013 A T TCGCGTGCGTCCTCAATACCATGATTCGTG TTGAGTAGGTCGCTTCGGCTTCTGGTAAAT GE2433 t1885 rad-51 IV SNV 10282328 C T TCGAGGCTCCTCCGTCGACTTGTGCTACAA TTGATTGGTAATAATTACAGCGACACCATA GE2347 t1519 rmd-1 III SNV 9759805 G A CTTGCCGCTAAATGGACTGCCATCATGTTC GACTTGTTGTCGATCAACTGCCAACTAAGG GE2219 t1501 rmd-1 III SNV 9759929 G A CCGACTTTGCTCTTCTCCATCTTCGTGCTC CTTCTCTTACACCATCGCGAACCTTTCATG 80 Strain Allele(s) Gene Chr. Mutation Position Base Change Left Flanking Sequence (30 bp) Right Flanking Sequence (30 bp) GE2211 t1476 sas-1 III SNV 12710102 C T CGCTCGCCGGTATTCCGCAACTCTCGAAAC CGAACTGGAATTTTATGGCCCGATTCGCTC GE2343 t1521 sas-1 III SNV 12710202 G A ATCGAGGCGCCCTCCTTGATCTACCACTGG ATTCGTCTCAATTCCGCTTCCAACACCGAA GE2078 t2033 sas-5 V SNV 11612449 C T GAGACTCCAGCTGAACGAGAACGCCGTATT GTGAAAAATACGCTCGCAGGAAATGATATC GE2134 t2079 sas-5 V SNV 11612449 C T GAGACTCCAGCTGAACGAGAACGCCGTATT GTGAAAAATACGCTCGCAGGAAATGATATC GE2469 t2173 spn-4 V SNV 6783986 A T TCACGAAGATGATGTGGGAATATAGAGTTC ACTCCACATCAGACAGCTCGTGATTCGAAG GE2317 t2098 spn-4 V SNV 6784646 A T TGTTCACGAAACGAGAAGGGAATATTGCTG CATATATCCTAAAAGTTTTTTGATGAATTC GE2386 t2165 sqv-4 V SNV 10660827 G A GAGGATTCTCCTCCAATGAGCACACGATCT GATTGGCAAGATCCTTCATAGCTGTTCCTG GE2059 t2025 sqv-4 V SNV 10661143 G A CGTCCGTACATTTTTGTGGGAGTGTTCACC AGATAAAAATAAGATCGGCTTCAGCAATCG GE2277 t1496 such-1 III SNV 11515520 G A GATAAGCCTCCCGAGCCATAAACCGGCCGA CATCAAGCTTCCAACTCGTCTAACAACCAG GE2277 t1496 such-1 III SNV 11515883 G A AATTGACGATTTTCGAGCAAATATCCACGT GCGAGCTGCCATCCAATTGATATTCTCGGG GE2666 t1693 such-1 III SNV 11515540 C T AACCGGCCGAGCATCAAGCTTCCAACTCGT TAACAACCAGCAAATTTGCATCCAAAAAAT GE2827 t1786 T22B11.1 IV SNV 4692945 G A ACGTTGGGTGCCCCCGCAGGCACATGAATG AAGAATGCGGATATTCCAAAAAGTGTTAAG GE2895 t1866 T22B11.1 IV SNV 4696017 G A TGAAGTCCTCAGACAAGAAACTGTACTCGT GATCAAGAATCTGGAGACTGTGTTCGATCG GE1734 t1438 t1477 tlk-1 III SNV 9707175 C T AGGATGGTGAGAACAATCGATGAAGAGACT AGACGGATTCTTCGCTATCTCAGGTTTGTT GE2613 t1677 tlk-1 III SNV 9708080 G A GTTAACAAAGACTGGAAGGAGGAGAAGAAG CGAACTACGTCAAGCACGCGATGCGTGAAA GE2399 t1559 top-3 III SNV 11951381 G A TATTCTTGTCCAAATATCGTTGCAAACTTT CGAGAGCACGCCAATTTTGCGTTCAGACTC GE2220 t1516 top-3 III SNV 11958680 C T TCTCGTGAAATTGGAAATTCATCATATGTC CGAAACACTTGTCACCGAAATTGCGGTTTG GE1735 t1470 top-3 III SNV 11957525 C T ATTGCTTCACCTTCTCGATCGCAATCTGTC AGACTACCAGGATATCGTGTCGTTGGGCTT GE2958 t1464 t1484 top-3 III SNV 11951669 C T CAATGTGTTCGGCGTGTGTTGCGTCGGTTC AATTCCGTATTTGTCCATTAGTGAGATTAA GE2512 t1909 trcs-1 IV SNV 9587541 C T CCTTCAGTCGCTCTCCATATATAAGACCCT GTCACGAAGAATATCAAATTCGCAAGTGAT GE1939 t1745 trcs-1 IV SNV 9587985 G A GAGAGTGGAAGTAACGGTATGAAACTGTTT CATGTCAGCCATTTGCAAAAGTGGGTAGAT GE2112 t2037 unc-112 V SNV 14692219 C T TCATCAAGATTTTGAGAATGTTCTTTGCTT GCATTGACAGGAAAATGTATCCACCGATGA GE2326 t2106 unc-112 V SNV 14696546 C T ATTTTCTAAATATATAATTTTAAATCCTCA CTTTGAATATTCAAATCGGTGACCAAAATT GE1722 t1435 vps-33.1 III SNV 8701605 C T GTAAAAGAAATTCCATTGTGCTGGTTGCCA GAGACGGCGAATGTCTATCACTCTCCAGTC GE2366 t1561 vps-33.1 III SNV 8702923 G A AATGCTTGAAGAAGGAGATCGAGTGAAATG GTCGGCTGGCCAAAAACTGTAATTGGAGAT GE2292 t2114 vps-39 V SNV 14035713 G A CCAGAGTTCGGAAGAGCATCAGATACACCT TGAATTGGTCTCATCGTGTGCATTGTAGTA GE1937 t2189 vps-39 V SNV 14036143 C T GTCTCGTGAAAATATGAAGAAGGTTCTTCC ATTTGAAAATCACGTGCTCCAAGTACGGAA GE2056 t2016 vps-39 V SNV 14037839 G C ATATAACTTCTTTCGTGACGAAACAATTAC TACAGGTCTCCAGATGTCTGAAAACAAAAA GE2153 t1773 wapl-1 IV SNV 4444464 C T TGCTTTTCGATGTACGAATTGAACACTTCC AGACCATATTCACAACTTTATCCTTGTCTT GE2305 t1867 wapl-1 IV 122-bp deletion 4442749-4442872 TATTGAAAATTCAAAATAACTCAGACTCCT TCCGACGAGAAGAGCGTGGTAAGAAGCGAT 81 Strain Allele(s) Gene Chr. Mutation Position Base Change Left Flanking Sequence (30 bp) Right Flanking Sequence (30 bp) GE2738 t1833 Y54G2A.73 IV SNV 3000662 A T TTTTTGAGCGGTTGCTGAAGAAGCTCCTCT ATTGAGTCTTCGGACGTTTTGTACCAGCCG GE2387 t1913 Y54G2A.73 IV SNV 3001767 G A TCGGTACTATCTCGTCTTGATGGCTCTGTC TTCGTCGTTCTACAAGATTCTCATGAAATA GE2884 t1755 Y54G2A.73 IV SNV 3008481 C T CGCGATTTTTCAAGCATTTTCTGGTGCTCA TCTGAAAATGATTGTATTGTTGCAAAGTAG GE1713 t1433 ZK688.9 III SNV 7882477 C T CACAGTTCCTTTGTAATGTGTAGTATAGGT CAGTCAAACGGCTTCTGATGTTCCGATATT GE2621 t1587 ZK688.9 III SNV 7882717 C T AAGTTTCAACTCCCGGTCATATTTGCATTT TGTTGAATGAACACATTTAGGCAAGAGCAT GE2348 t1518 zyg-8 III SNV 12063671 C T GAGCCAAGTTGGCGTCTGACACTTGCCAAT GATACAATCGACACTTGGAGACGAAGAAGC GE2362 t1547 zyg-8 III SNV 12063832 G A TAGGCATTTGTTGAACAAGAAGACGGCGAG TGAGAGCAAAAAATCGCCCCGACCAATCAG 82 Appendix B Gene Ontology terms and associated genes Table B.1 Gene Ontology terms (Biological Process category) and associated genes Biological Process Term Description Associated Genes GO:0009792 Embryo development ending in birth or egg hatching perm-5, tlk-1, hcp-3, sas-5, cra-1, csr-1, dgtr-1, pod-1, lis-1, atl-1, mel-32, pos-1, unc-112, div-1, mre-11, sas-1, C56A3.8, such-1, atg-7, mom-2, zyg-8, rmd-1, cept-2, spn-4, cpl-1, rad-51, mbk-2, mex-5, trcs-1, F21D5.1, air-1, let-99, cyk-3, dlat-1, wapl-1, rad-50, cpt-2, aptf-2, sqv-4, vps-39, dif-1, gsp-2, cls-2, klp-18, klp-19 GO:0007049 Cell cycle zyg-8, lis-1, rmd-1, gsp-2, cls-2, sas-5, such-1, air-1, let-99, rad-50 GO:0051301 Cell division lis-1, rmd-1, gsp-2, cls-2, div-1, sas-5, such-1, let-99, cyk-3 GO:0051321 Meiotic cell cycle gsp-2, cls-2, cra-1, mre-11, such-1, wapl-1, rad-50 GO:0007067 Mitotic nuclear division lis-1, rmd-1, gsp-2, cls-2, hcp-3, such-1, klp-19, let-99 GO:0045132 Meiotic chromosome segregation atl-1, gsp-2, mbk-2, rad-51, mre-11, klp-18, wapl-1, rad-50 GO:0007275 Multicellular organism development zyg-8, lis-1, dif-1, gsp-2, mbk-2, tlk-1, sas-5, mex-5, such-1, let-99, mom-2 GO:0007126 Meiotic nuclear division gsp-2, top-3, mbk-2, rad-51, klp-18, rad-50 GO:0006915 Apoptotic process pod-1, rmd-1, perm-5, rad-51, C56A3.8, trcs-1, F21D5.1, air-1, cyk-3, rad-50 GO:0006974 Cellular response to DNA damage stimulus atl-1, rad-51, cra-1, mre-11, wapl-1, rad-50 GO:0048477 Oogenesis pod-1, lis-1, rmd-1, perm-5, cra-1, F21D5.1, dgtr-1 GO:0006281 DNA repair atl-1, cra-1, mre-11, wapl-1, rad-50 GO:0007052 Mitotic spindle organization sas-5, hcp-3, such-1, klp-19 GO:0016192 Vesicle-mediated transport vps-39, ccz-1, C34D4.4, vps-33.1 GO:0007018 Microtubule-based movement lis-1, klp-18, klp-19 GO:0000132 Establishment of mitotic spindle orientation lis-1, let-99, mom-2 GO:0000003 Reproduction perm-5, mbk-2, tlk-1, cra-1, hcp-3, sas-5, csr-1, dgtr-1, F21D5.1, air-1, let-99, dlat-1, sqv-4, pod-1, unc-112, C56A3.8, klp-19, mom-2 GO:0016310 Phosphorylation zyg-8, atl-1, mbk-2, tlk-1, C56A3.8, air-1 GO:0035046 Pronuclear migration lis-1, mbk-2, let-99 GO:0018991 Oviposition sqv-4, gsp-2, ZK688.9, trcs-1, wapl-1 GO:0008340 Determination of adult lifespan pod-1, atl-1, pos-1, cup-5, vps-33.1, sas-5, bckd-1A, atg-7, mom-2 83 Table B.2 Gene Ontology terms (Molecular Function category) and associated genes Molecular Function Term Description Associated Genes GO:0008017 Microtubule binding zyg-8, lis-1, cls-2, klp-18, klp-19 GO:0005515 Protein binding zyg-8, ccz-1, cpl-1, rad-51, tlk-1, mbk-2, vps-33.1, sas-5, air-1, let-99, dlat-1, lis-1, gsp-2, cls-2, unc-112 GO:0016887 ATPase activity rad-51, klp-18, klp-19, rad-50 GO:0016301 Kinase activity zyg-8, atl-1, mbk-2, tlk-1, C56A3.8, air-1 GO:0005524 ATP binding zyg-8, atl-1, mbk-2, tlk-1, rad-51, klp-18, klp-19, air-1, rad-50 GO:0016740 Transferase activity zyg-8, cept-2, atl-1, mel-32, mbk-2, tlk-1, dgtr-1, dlat-1, cpt-2 GO:0004674 Protein serine/threonine kinase activity zyg-8, atl-1, mbk-2, tlk-1, air-1 GO:0000166 Nucleotide binding zyg-8, atl-1, spn-4, mbk-2, tlk-1, rad-51, klp-18, klp-19, air-1, rad-50 GO:0008408 3'-5' exonuclease activity mre-11, rad-50 84 Appendix C Life stage-specific expression patterns These plots show gene expression patterns across embryonic and larval life stages for each of the 20 essential genes of interest identified in this study. The data originate from RNA-seq experiments from the modENCODE project (Hillier et al. 2009; Gerstein et al. 2010, 2014; Boeck et al. 2016) and the figures were retrieved from GExplore (genome.sfu.ca/gexplore; Hutter et al. 2009; Hutter and Suh 2016). Short descriptions of the gene expression patterns indicate genes with characteristic maternal gene expression (high expression in the early embryo and hermaphrodite gonad), zygotic expression (additional peaks of gene expression at developmental time points other than the maternal ones), and high expression in males.85 Figure C.1 atg-7 Early zygotic expression. Figure C.2 bckd-1A Characteristic maternal, zygotic, and male expression. Figure C.3 C34D4.4 Characteristic maternal and some male expression. Figure C.4 C56A3.8 Characteristic maternal and high male expression. 86 Figure C.5 cept-2 Characteristic maternal expression. Figure C.6 cls-2 Characteristic maternal and zygotic expression. Figure C.7 cpt-2 Characteristic maternal and zygotic as well as some male expression. Figure C.8 D2096.12 Characteristic maternal expression. 87 Figure C.9 dgtr-1 Characteristic maternal expression. Figure C.10 dlat-1 Characteristic maternal and zygotic as well as some male expression. Figure C.11 F21D5.1 Characteristic maternal expression. Figure C.12 F56D5.2 High expression in males. 88 Figure C.13 K08D8.6 High expression in males and zygotic expression. Figure C.14 nstp-2 High expression in males and zygotic expression. Figure C.15 perm-5 Characteristic maternal and zygotic expression. Figure C.16 T22B11.1 High expression in males. 89 Figure C.17 top-3 Characteristic maternal expression. Figure C.18 trcs-1 Characteristic maternal expression. Figure C.19 Y54G2A.73 (Figure from wormbase.org; not available from GExplore) High expression in males. 90 Figure C.20 ZK688.9 Characteristic maternal expression. 91 Appendix D Terminal phenotypes of maternal-effect lethal embryos This appendix contains representative images of the terminal phenotypes observed in 27 maternal-effect lethal strains. Eggs were dissected from young homozygous mutant adults and incubated overnight in distilled water (unless otherwise indicated). Corresponding gene and strain names are indicated in each panel with a short description of the terminal phenotypes observed. Multiple images per allele are included where appropriate, if more than one phenotype was common in a particular strain. All scale bars represent 10 μm.92 Figure D.1 A atg-7 GE1936(t1738) Embryos arrest at the 2-fold stage. Figure D.1 B atg-7 GE1936(t1738) Some embryos have invaginations along the body wall. Most embryos are moving inside the eggshell but do not hatch. 93 Figure D.2 atg-7 GE1958(t1726) Most embryos arrest around the 2-fold stage, moving, with a lumpy body wall. Figure D.3 A bckd-1A GE2627(t1603) (Embryo incubated in M9 buffer.) Some embryos appear undivided with invaginations along the cell membrane. 94 Figure D.3 B bckd-1A GE2627(t1603) (Embryos incubated in M9 buffer.) Some embryos fill the eggshell and exhibit Brownian motion in large vacuoles. Figure D.4 A bckd-1A GE2206(t1514) Embryos appear undivided; some eggs are more round than oval. 95 Figure D.4 B bckd-1A GE2206(t1514) (Embryos imaged immediately, without overnight incubation.) Undivided embryos fill the eggshell completely. Figure D.5 cept-2 GE2122(t2007) Embryos appear disorganized with approximately 100 cells. Some have large vacuoles with Brownian motion. Few exhibit a contraction movement. 96 Figure D.6 cept-2 GE2047(t2021) Most embryos appear disorganized with approximately 100 cells and large vacuoles with Brownian motion. Some embryos fill the eggshell completely. Figure D.7 cls-2 GE2275(t1517) (Embryo incubated in M9 buffer.) Embryos arrest with approximately 100 cells. Some exhibit a contraction movement. 97 Figure D.8 cls-2 GE2357(t1527) Embryos appear undivided and shrunken within the eggshell. Figure D.9 cpt-2 GE1938(t1742) (Embryo incubated in M9 buffer.) Embryos arrest with approximately 100 cells. 98 Figure D.10 cpt-2 GE2447(t1879) (Embryo incubated in M9 buffer.) Embryos arrest at the bean stage. Figure D.11 D2096.12 GE2407(t1906) Embryos arrest with approximately 100 cells. Most exhibit a contraction movement. Some show signs of osmotic integrity defect. 99 Figure D.12 D2096.12 GE2499(t1877) Embryos appear disorganized and some exhibit a contraction movement. Most have large vacuoles exhibiting Brownian motion and some eggs burst after incubation in water. Figure D.13 A dgtr-1 GE2063(t2042) Some embryos fill the eggshell completely and appear roundish; others burst after incubation in water. 100 Figure D.13 B dgtr-1 GE2063(t2042) Most embryos arrest with approximately 100 cells and have misshapen eggs. Figure D.14 dgtr-1 GE2135(t2043) (Image taken immediately after dissection.) Embryos fill the eggshell completely. Many eggs burst after incubation in water. 101 Figure D.15 A dlat-1 GE2541(t2035) Some embryos arrest at the one-cell stage. Figure D.15 B dlat-1 GE2541(t2035) Some embryos arrest with four cells or less. 102 Figure D.15 C dlat-1 GE2541(t2035) Some embryos arrest with approximately 20 cells and others arrest during the first cell division. Figure D.16 A dlat-1 GE2335(t2056) Some embryos arrest at the one-cell stage. 103 Figure D.16 B dlat-1 GE2335(t2056) Some embryos arrest during the first cell division. Figure D.16 C dlat-1 GE2335(t2056) Some embryos arrest with approximately 20 cells. 104 Figure D.17 F21D5.1 GE2402(t1940) (Embryos shown were incubated in M9 buffer.) Embryos appear undivided with a severe osmotic integrity defect. Many eggs burst after incubation in water. Figure D.18 F21D5.1 GE2445(t1935) Embryos appear undivided and shrunken within the eggshell; many eggs burst after incubation in water. 105 Figure D.19 A K08D8.6 GE2034(t1808) Some embryos appear disorganized with approximately 100 cells. Figure D.19 B K08D8.6 GE2034(t1808) Most embryos fill the eggshell completely and have large vacuoles. 106 Figure D.20 K08D8.6 GE2035(t1809) Embryos arrest with approximately 100 cells. Figure D.21 nstp-2 GE2091(t1772) Embryos develop to approximately the 3-fold stage, with a lumpy body morphology and constricted nose tip. 107 Figure D.22 nstp-2 GE2288(t1835) Embryos develop to the 2-fold or 3-fold stage, with a lumpy body morphology and constricted nose tip. Figure D.23 perm-5 GE2453(t1900) (Image taken immediately after dissection.) Embryos fill the eggshell completely. Many eggs burst after incubation in water, as did eggs from GE2391(t1932) (not shown). 108 Figure D.24 trcs-1 GE2512(t1909) Embryos fill the eggshell completely; some have invaginations along the cell membrane. Many eggs burst after incubation in water. Figure D.25 trcs-1 GE1939(t1745) (Embryo shown was incubated in M9 buffer.) Undivided embryos have a severe osmotic integrity defect. Many eggs burst after incubation in water. 109 Figure D.26 ZK688.9 GE1713(t1433) Embryos arrest with approximately 100 cells. Figure D.27 ZK688.9 GE2621(t1587) Embryos arrest with approximately 100 cells.
Thesis/Dissertation
eng
Cell and Developmental Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Identification of essential genes in C. elegans through whole genome sequencing of legacy mutant collections
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