Functional Annotation of Uncharacterized Enzymes in Yeast

Functional Annotation of Uncharacterized Enzymes in Saccharomyces cerevisiae

Julia Ann Hanchard

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Molecular Genetics University of Toronto

Abstract Functional Annotation of Uncharacterized Enzymes in Saccharomyces cerevisiae Julia Hanchard Doctor of Philosophy, 2019 Department of Molecular Genetics University of Toronto In the post-genomic era, clinicians and scientists are increasingly reliant on interpretation of variants in metabolic genes for determining pathogenicity. These interpretations depend on functional annotation of the roles genes provide in metabolism, an annotation that is far from complete. I embarked on a journey of enzyme discovery to fill gaps in our knowledge of metabolism in the budding yeast, Saccharomyces cerevisiae. I carried out a genetic and metabolomic screen of 120 uncharacterized candidate enzyme encoding genes that comprised my master’s thesis. This dissertation describes my work in ascribing function to two distinct enzymes, Das2 and Tda5. Throughout my study I have found that Das2 is a novel uridine/cytidine kinase that functions in concert with a second minor uridine kinase, Urk1.

These two enzymes are interdependent and in turn depend on a third enzyme, the major uracil phosphoribosyl transferase, Fur1 for stability. These three enzymes form a complex that is essential to wild-type pyrimidine salvage. As I aimed to elucidate the function of Tda5, I discovered that this uncharacterized enzyme is essential to growth. Loss of function mutations in TDA5 are alleviated by de-repression of its sporulation specific paralog,

Ydl114w, and when YDL114W is deleted, tda5Δ is rescued by hypomorphic mutations in the ergosterol biosynthetic pathway. Analysis of metabolite levels in hand with nutritional sensitivities and genetic interaction data support the essential role of Tda5 in lipid homeostasis. Together, these studies assign new functions to previously uncharacterized genes and reveal functional interdependence of enzymes that sheds light on metabolic interpretations of genetic data.

Acknowledgements I’m grateful to my supervisor, Dr. Amy Caudy, for her leadership at the helm of a multi- faceted and complex metabolomics lab. Her support and insight were crucial to the projects that comprise this dissertation. Thanks to my graduate advisory committee members Jim Dennis and Andy Bognar for their expert guidance through experiments and refining the trajectory of my thesis.

I’d like to thank the Caudy lab tribe: Dr. Olga Zaslaver, Dr. Soumaya Zlitni, Yutong Ma and Yoomi Oh for their steady provision of motivation, support and friendship. I am especially grateful to my awesome team of undergraduate students who I’ve had the pleasure and privilege of working with: Muhammad Bin Munim, Connie Liu, Mashiat Khan, Sheena Faye Garcia, Lily Chen, Angela Dennisova, Sara Allarhakia and Jonathan Ward. Each of these students has provided technical work critical to this thesis, and it’s been a lot of fun in training them.

I’ll always be indebted to my family for their unwavering love and support. Above all, I am thankful to Kasra Zokaei for his love, companionship and wise advice throughout my PhD.

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Table of Contents Acknowledgments...... ii List of Tables...... ix List of Figures ...... x List of Abbreviations...... xii

1 Introduction ...... 1

1.1 Our knowledge of metabolism is incomplete ...... 1

1.1.1 Early study of metabolism ...... 1 1.1.2 Clinical interpretation of genomes requires more study of metabolism ...... 1 1.1.3 Selective pressures lead to evolution of metabolic pathways and genome expansion ...... 2 1.1.4 Prediction of enzyme function ...... 4 1.1.5 Prediction of enzyme substrates ...... 5 1.1.6 Domains of unknown function...... 6 1.1.7 Metabolic networks contain missing enzymes ...... 6 1.1.8 Metabolic networks are missing metabolites ...... 7 1.1.9 New metabolic pathway information from fluxomics ...... 7 1.1.10 Enzyme discovery through metabolomic screening in yeast ...... 8 1.2 Pyrimidine metabolism of Saccharomyces cerevisiae ...... 10

1.2.1 UMP is the central pyrimidine metabolite ...... 10 1.2.2 de novo biosynthesis of UMP ...... 10 1.2.3 Regulation of UMP biosynthesis ...... 12 1.2.4 Multi-phosphorylated pyrimidine nucleotides ...... 13 1.2.5 Catabolism as a source of 5’ pyrimidine nucleotide monophosphates ...... 13 1.2.6 During autophagy 3’ pyrimidine nucleotides are hydrolyzed to nucleosides .... 14 1.2.7 Pyrimidine salvage from an uptake perspective ...... 15 1.2.8 Pyrimidine salvage enzymes- Early studies reveal pathway architecture and activities ...... 15 1.2.9 Pyrimidine salvage - Purification of enzymes and cloning of genes ...... 16 1.2.10 Project rationale: Resolving pyrimidine salvage in yeast ...... 17 1.3 Background for Tda5: a novel short chain dehydrogenase essential for respiration 17

1.3.1 Oxidoreductases and short chain dehydrogenases ...... 18

1.3.2 SDR activity pervades lipid homeostasis ...... 18 1.3.3 Membrane fluidity and structural integrity ...... 23 1.3.4 Yeast growth is supported by fermentation and respiration ...... 24 1.3.5 Compartmentalization of metabolism in yeast ...... 25 1.3.6 Compartmental proteomes are distinct ...... 26 1.3.7 Project rationale: Elucidating the functions of Tda5 ...... 28 2 Methods ...... 29

2.1 Methods relating to Chapter 3: Uncovering the function of Das2 ...... 29

2.1.1 Growth and maintenance of yeast cultures ...... 29 2.1.2 Construction of Mutants ...... 29 2.1.3 Preparation of point mutants by in vivo site directed mutagenesis ...... 29 2.1.4 HA fusion tagged alleles ...... 30 2.1.5 Heterologous expression of enzymes...... 30 2.1.6 Preparation of Das2 G17E ...... 31 2.1.7 Radiometric enzyme assays ...... 31 2.1.8 Pyrimidine analogue resistance assays ...... 32 2.1.9 Metabolite extracts ...... 32 2.1.10 In vivo tracking of uridine kinase activity...... 32 2.1.11 Western Blot ...... 32 2.1.12 Preparation of Ribose-1-phosphate ...... 33 2.2 Methods for Chapter 4: Phenotyping and mapping suppressors of Tda5 ...... 33

2.2.1 Growth and maintenance of yeast cultures ...... 33 2.2.2 Construction of Mutants ...... 33 2.2.3 Spot assays ...... 35 2.2.4 Metabolite extracts ...... 35 2.2.5 Purification of short and medium chain acyl-CoA metabolites ...... 35 2.3 Strains and Plasmids used in this study ...... 35

3 Discovery of Das2 a Novel Uridine Kinase and Obligate Member of a Novel UMP Salvage Complex ...... 40

3.1 Rationale and summary ...... 40

3.2 Motivational phenotypes for study of Das2 ...... 41

3.2.1 Metabolomic phenotyping reveals das2Δ accumulates uridine ...... 41

3.2.2 Literature studies implicate DAS2 in response to 5-FOA and 6-azauracil ...... 43 3.2.3 Deletion of DAS2 confers resistance to 6-azauracil ...... 44 3.2.4 Mechanism of action for 6-azapyrimidines ...... 44 3.2.5 Resistance to 6-azapyrimidines...... 45 3.2.6 Mechanism of action of 5-fluoropyrimidines ...... 45 3.2.7 Resistance to 5-fluoropyrimidines ...... 46 3.2.8 Das2 physically interacts with Urk1:implications for a pyrimidine salvage complex ...... 46 3.2.9 Resolving the activities of Das2, Urk1 and Fur1 ...... 47 3.2.10 Null mutations in fur1Δ are synthetic lethal with ura3Δ ...... 48 3.2.11 Genetic interactions point to a role for Das2 in pyrimidine salvage ...... 48 3.2.12 Das2 and Urk1 topology ...... 50 3.2.13 Das2 catalyzes the uridine kinase reaction in vitro ...... 52 3.2.14 Das2 pull-downs of Urk1 reveal UPRTase activity ...... 55 3.3 Using resistance to pyrimidine analogues to determine the topology of pyrimidine salvage pathways in yeast ...... 56

3.3.1 Mutations in das2Δ and urk1Δ are non-additive ...... 56 3.3.2 Activation of 6-azauracil occurs exclusively through the UPRTase reaction ... 58 3.4 Response to 5-fluorouridine is not additive between das2Δ and urk1Δ ...... 60

3.4.1 Uridine accumulation unmasks Urh1 function ...... 61 3.4.2 Residual activities of Urk1 and Das2 are unveiled by double and triple mutant resistance to 5-fluorouridine ...... 61 3.4.3 Blocking UPRTases while Urh1 is active creates a sink for 6-azauracil and uracil 62 3.5 6-azauridine phenotypes match those for 5-fluorouridine ...... 65

3.5.1 Enzyme activity assays indicate Das2 and Urk1 both function as uridine kinases 67 3.5.2 Radiometric assays on cell free lysates ...... 67 3.5.3 UPRTase activity assays on cell free extracts...... 71 3.6 Metabolite levels elucidate bottlenecks and pathway logic ...... 72

3.6.1 Metabolite levels indicate Urk1 is the elusive secondary UPRTase...... 78 3.6.2 Metabolite level changes for point mutants phenocopy deletion mutants and do not pinpoint activities ...... 79

3.7 Das2 and Urk1 protein levels establish interdependence of uridine kinases ...... 81

3.8 Discussion ...... 85

3.8.1 Pyrimidine analogs are crucial drug targets in pathogenic yeasts ...... 87 3.8.2 Saccharomyces cerevisiae salvage complex is conserved to Candida albicans 89 3.8.3 Pyrimidine salvage is a selective target for neoplastic cells ...... 89 3.8.4 6-azauridine is more potent than 6-azauracil ...... 91 4 Elucidation of Tda5, a novel essential short chain dehydrogenase ...... 93

4.1 Rationale and summary ...... 93

4.2 Results ...... 94

4.2.1 Metabolomic phenotypes discriminate TDA5 from other uncharacterized enzymes ...... 94 4.2.2 Growth phenotype of fresh deletion mutant ...... 94 4.2.3 Tda5 is a novel essential dehydrogenase ...... 96 4.2.4 Evolutionary conservation of TDA5 ...... 97 4.2.5 Examination of yeast short chain dehydrogenases ...... 101 4.2.6 Topology and localization of the Tda5 protein ...... 103 4.2.7 How Tda5 transits from the ER to the mitochondria ...... 105 4.2.8 Expression of Ydl114w rescues loss of function in Tda5 ...... 105 4.2.9 HST1 controls expression of middle sporulation genes ...... 108 4.2.10 Metabolomic phenotyping of tda5Δ strains ...... 111 4.2.11 Identification of YDL114W independent suppressors of tda5∆ deletion mutants 114 4.2.12 Mapping suppressors of tda5∆ loss of function independent of YDL114W .... 115 4.2.13 Loss of function mutations in ergosterol biosynthesis suppress tda5Δ ...... 117 4.2.14 Large scale genetic interaction datasets reveal additional candidate positive genetic interactions with tda5Δ ...... 118 4.2.15 Negative genetic interactions of mitochondrial proteins with tda5∆ ...... 120 4.2.16 Chemical suppression of tda5Δ...... 121 4.2.17 Strains deleted for tda5Δ have a membrane defect ...... 124 4.2.18 The growth defect of tda5Δ can be rescued by N-acetyl-cysteine...... 125 4.2.19 Defects in lipid homeostasis and mitochondrial activity cause ROS formation 127

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4.2.20 Measurement of acyl-CoA pools in fresh and suppressed tda5∆ mutants informs on routes of metabolic compensation ...... 127 4.2.21 Initial metabolic response induced by loss of function of Tda5 ...... 130 4.3 The suppressed tda5Δ deletion mutant has negative genetic interactions with deletions in other short chain dehydrogenases...... 137

4.3.1 GO enrichment suggests metabolic process for Tda5 ...... 139 4.3.2 The tda5Δ strain is sensitive to unsaturated fatty acids ...... 140 4.4 Discussion ...... 143

5 Thesis summary and future directions ...... 149

5.1 Conclusions ...... 149

5.2 Future Directions ...... 150

5.2.1 Future directions relating to Das2 ...... 150 5.2.2 Future directions for annotation of Tda5 ...... 152 5.2.3 Resolving functions of clinical variants in metabolism ...... 155 6 References ...... viii

7 Appendix ...... xlvii

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List of Tables

Table 1. Plasmids created for this study ...... 35 Table 2. Yeasts strains used in this study ...... 37 Table 3. Orthologues of S. cerevisiae pyrimidine salvage enzymes in pathogenic yeasts ...... 88 Table 4 Recessive mutations in sterol biosynthesis suppress loss of function in TDA5...... 117 Table 5 Gene ontology terms enriched for negative genetic interactions with tda5Δ ...... 140

List of Figures

Figure 1. Metabolic Pathway for de novo Biosynthesis of Pyrimidines ...... 12 Figure 2. Conventional model of UMP salvage in Saccharomyces cerevisiae ...... 17 Figure 3. Localizations of characterized short chain dehydrogenases in Saccharomyces cerevisiae ...... 20 Figure 4. Full-scan LC-MS metabolomic profiling reveals uridine accumulates in the das2Δ prototrophic deletion mutant...... 42 Figure 5. Enzyme domain topologies for Urk1 and Das2 ...... 48 Figure 6. Multiple sequence alignment shows DAS2 is unique among uridine kinases ...... 51 Figure 7. Das2 catalyzes a uridine kinase reaction ...... 53 Figure 8. Das2 pull-downs of Urk1 identify the UPRTase activity for Urk1...... 55 Figure 9 Growth inhibition of pyrimidine salvage mutants by 6-azauracil ...... 59 Figure 10. Growth inhibition of pyrimidine salvage mutants by 5-fluorouridine ...... 64 Figure 11 Growth inhibition of pyrimidine salvage mutants by 6-azauridine ...... 66 Figure 12 Specific activities of uridine salvage reactions ...... 68 Figure 13 In vivo tracking of uridine kinase activity using 6-azauridine incorporation in growing cultures...... 70 Figure 14. UMP levels are stable across salvage mutants ...... 73 Figure 15 Relative uridine and uracil levels across pyrimidine salvage mutants ...... 74 Figure 16. Uracil and Uridine levels for point mutants phenocopy full deletions...... 80 Figure 17 Das2-3XHA levels in various pyrimidine salvage mutants ...... 82 Figure 18 Urk1-3XHA levels measured across various salvage mutants backgrounds...... 84 Figure 19. Revised model of UMP salvage in Saccharomyces cerevisiae ...... 86 Figure 20. The freshly dissected tda5∆ strains are petite whereas tda5∆ from the prototrophic yeast deletion collection carried a suppressor mutation...... 96 Figure 21. TDA5 orthologs are distributed among eukaryotes and bacteria and conserved to human homologs ...... 99 Figure 22. Reactions catalyzed by human orthologs of TDA5 conserved domain cluster 5539 ...... 100 Figure 23. Yeast short chain dehydrogenases display a conserved nucleotide binding domain and catalytic residues ...... 102 Figure 24. Fluorescence microscopy shows Tda5 is localized to the plasma membrane and endoplasmic reticulum...... 104

Figure 25. Tda5 is an integral membrane short chain dehydrogenases/reductase ...... 104 Figure 26. Inactivation of Hst1-Sum1-Rfm1 results in expression of the SDR YDL114W which rescues the growth of tda5Δ...... 107 Figure 27. Metabolomic analysis of tda5Δ and tda5Δ suppressed strains to wild-type ...... 113 Figure 28 The frequency of suppressors is vastly reduced in the tda5Δydl114wΔ double deletion mutant...... 114 Figure 29 Illustration of process for identifying causative mutations in tda5 suppressor strains...... 116 Figure 30 The tda5∆ mutant has positive genetic interactions with hypomorphs in ergosterol biosynthesis ...... 119 Figure 31 Stronger growth is observed for tda5Δ strains grown in the presence of ergosterol pathway inhibitors...... 121 Figure 32. Cells are sensitive to Fenpropimorph and myriocin...... 123 Figure 33 Detection of a membrane defect for tda5Δ...... 124 Figure 34 N-acetyl cysteine rescues growth defect of tda5Δ mutant...... 126 Figure 35 Acetyl-CoA and HMG-CoA levels across ergosterol hypomorphs in combination with tda5Δ ...... 129 Figure 36 CoA levels for TDA5 and tda5Δstrains due to YDL114W repression ...... 131 Figure 37 Changes to N-acetyl-glutamate cycle due to YDL114W repression in TDA5 and tda5Δ strains...... 133 Figure 38 Changes to Tricarboxylic Acid Cycle due to YDL114W repression in TDA5 and tda5Δ strains...... 135 Figure 39 Branched chain amino acid dynamics in tda5Δ and TDA5 upon conditional knockdown of YDL114W...... 136 Figure 40. tda5Δ has negative genetic interactions with deletions in other short chain dehydrogenases...... 138 Figure 41. tda5Δ strains are sensitive to unsaturated fatty acids ...... 142 Supplemental Figure 42: Ergosterol biosynthetic pathway of Saccharomyces cerevisiae .. xlvii

List of Abbreviations CDP-cytidine diphosphate CDP-DAG-cytidine diphosphate diacylglycerol CMP- cytidine monophosphate CTP-cytidine triphosphate DAS2- gene named for dst1 6-azauracil sensitivity -uridine kinase elucidated in this study DAG- diacylglycerol DNA- deoxyribonucleic acid DUF- domain of unknown function ERG1- ergosterol biosynthetic gene encoding the enzyme squalene epoxidase ERG11-ergosterol biosynthetic gene encoding the enzyme lanosterol demethylase ERG12-ergosterol biosynthetic gene encoding the enzyme mevalonate kinase ERMES- endoplasmic reticulum mitochondria encounter structure ETC- electron transport chain FCY1- fluorocytosine resistance gene encoding the enzyme cytosine deaminase FCY2-fluorocytosine resistance gene encoding the permease for cytosine FUI1-fluorouridine resistance gene encoding the permease for uridine FUR1- fluorouracil resistance gene encoding the major uracil phosphoribosyl transferase FUR4- fluorouracil resistance gene encoding the permease for uracil HMG-CoA- 3-hydroxy-3-methyl-glutaryl-coenzyme A IEM- inborn error of metabolism IPP- isopentyl pyrophosphate IPRP- ion paired reverse phase chromatography LC-MS- liquid chromatography mass spectrometry MDA-5052- autoinduction media named by Studier MDG- non-inducing media named by Studier mGWAS- metabolite genome-wide associate study OTC- ornithine transcarbamylase PPR1- pyrimidine pathway regulation gene encoding transcription factor PtdIns- phosphatidyl inositol PSM-pro-spore membrane q-TOF- quadrupole time of flight mass spectrometer RNA- ribonucleic acid ROS- reactive oxygen species SAM- S-adenosyl-methionine SAH- S-adenosyl-homocysteine SDR- short chain dehydrogenase SNARE- Soluble N-ethylmaleimide-sensitive factor attachment protein receptor SKI- Super Killer referring to genes that convey sensitivity to toxin when inactivated TDA5- topoisomerase damage affected gene encoding dehydrogenase explored in this study TRAMP- Trf-Air2-Mtr4 polyadenylation complex UMP- uridine-5’-monophosphate (uridine-3’-monosphosphates are referred to specifically) UPC2- uptake control gene encoding the transcription factor controlling sterol uptake UPRTase- uracil phosphoribosyl transferase enzyme URA1- uracil requiring gene encoding the enzyme dihydroorotate dehydrogenase URA2- uracil requiring gene encoding the enzyme bifunctional carbamoylphosphate. synthetase/ aspartate transcarbamylase URA3- uracil requiring gene encoding the enzyme orotidine-monophosphate decarboxylase URA4- uracil requiring gene encoding the enzyme dihydroorotase

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URA5- uracil requiring gene encoding the enzyme orotate phosphoribosyl transferase URA6- uracil requiring gene encoding the enzyme uridine monophosphate kinase URA7- uracil requiring gene encoding the enzyme CTP synthase URA8- uracil requiring gene encoding the enzyme CTP synthase URA10- uracil requiring gene encoding the enzyme orotate phosphoribosyl transferase URK1- gene here elucidated as minor uridine kinase and minor UPRTase URH1- uridine hydrolase encoding gene vCLAMP- vacuole- mitochondrion membrane contact site for lipid transfer WGD- whole genome duplicate YDL114W- uncharacterized gene encoding uncharacterised paralog of TDA5 YNB- yeast nitrogen broth containing vitamins and minerals to support yeast growth YPD- yeast peptone dextrose rich media containing yeast hydrolysate and glucose

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1 Introduction 1.1 Our knowledge of metabolism is incomplete 1.1.1 Early study of metabolism Metabolism is the nexus of transformative reactions required to sustain life which is bounded by the availability of elemental resources, capacitance of extant genomes and thermodynamic barriers. Enzymes are the genetically encoded catalysts of metabolism and their study began with the discovery of the transformative action of diastase (Payen and Persoz, 1833). The first observations of metabolism were carried out by Louis Pasteur who found that when sugars were degraded to ethanol and carbonic acid, yeast was always present (reprint of the original 1857 article available as Pasteur, 1995). Not long after, it was shown that cell homogenates could likewise ferment— forming the basis of the enzyme theory of biochemistry (Kresge et al., 2005). By the early 1900’s it was determined by Harden and Young that metabolites were conserved between organisms (Harden and Young, 1906). It follows from this observation that by determining metabolic reactions in one organism, parallel reactions present in distant evolutionary lineages could be revealed. The conservation of reactions between organisms was crucial to the golden age of biochemistry, between the 1920’s to 1960’s, during which many of the enzymes and the reactions they catalyze were determined. With the advent of genetic mapping, in hand with recombinant DNA technologies, forward and reverse genetic studies were carried out to screen for and identify genes encoding enzymes and their cognate reactions. Development of X-ray crystallography led to the determination of three dimensional structures of proteins and their conformational states that aid in our understanding of enzymatic substrate binding and inhibition (Johnson and Phillips, 1965).

1.1.2 Clinical interpretation of genomes requires more study of metabolism In the post-genomic age, low cost whole genome sequencing has endowed researchers with the complete set of instructions encoding endogenous metabolism for any given organism of their choosing. At the same time, more systematic and genome-wide experiments are completed that assist in assigning functions to genes, yet no organism has a fully annotated genome and many metabolic functions remain elusive. Much of the research into metabolism was fueled by its relevance to disease and the use of personal genomic data is expanding in use as a first line diagnostic as well as identification of biomarkers for clinical management (Lionel et al., 2018; Reuter et al., 2018; Stavropoulos et al., 2016). In cancer cells metabolic pathways are altered to support tumor initiation, growth and metastasis (reviewed in Vander Heiden and DeBerardinis,

2017). Metabolic syndrome, diabetes and cardiovascular disease have a metabolic basis and are associated with high incidences of morbidity and significant financial burden (reviewed in O’Neill et al., 2016). Congenital defects in enzyme encoding genes give rise to a group of diseases called inborn errors of metabolism (IEM). Although each disease is individually rare, together IEMs comprise the largest class of inherited diseases (Childs et al., 2001). Understanding the etiology of metabolic diseases is complex since metabolism is integrated with every other cellular process. In order to identify causal pathogenic alleles and novel gene-metabolite relationships, genome-wide association studies on targeted populations in hand with measurements of metabolite levels are used to provide estimates of linkage disequilibrium for metabolic traits (mGWAS) (Illig et al., 2010; Nath et al., 2017; Raffler et al., 2015; Suhre et al., 2016; Yousri et al., 2018). Apart from discovery- based studies, routine clinical testing has expanded the number of clinically relevant loci alongside the number of variants of uncertain significance, with the ClinVar database containing more than 331,000 variants from over 500,000 variant-phenotype associations reported across the genome (Landrum et al., 2018). Although determination of penetrance and validation of variants is required, uncertain significance of pathogenicity arises from insufficient statistical power or lack of orthogonal experimental evidence to establish causality. In order to interpret functional impacts of genomic variants, clinicians and scientists must collaborate to achieve a higher level of functional annotation for eukaryotic genomes.

1.1.3 Selective pressures lead to evolution of metabolic pathways and genome expansion Although broad insight has been gained in understanding extant metabolism, its emergence and evolution remain unknown. The origin of life theory ascribed to Haldane and Oparin posits that on the primordial Earth, basic nutrient monomers of amino acids, sugars and nucleobases that are synthesized by all life were widely available in the environment—known as the primordial soup (Haldane, 1929; Oparin, 1924). Haldane further conjectured that ancient cells were bounded by oily membranes—a now-recognized property of all cells (Haldane, 1929). Membrane partitioning provided early cells with a mechanism for maintaining chemical gradients whereby lumenal uptake and concentration of both energy and nutrients allowed for the macromolecular polymerization reactions needed to support growth and spontaneous division (Deamer, 2016). As the numbers of cells grew, essential nutrients were depleted from the primordial soup and under natural selection, cells adapted to assimilate and transform the leftover nutrients into essential ones (Alves et al., 2002). Under this scenario, cells that could synthesize key molecules increased in fitness while decreasing the organism’s dependence on the soup, giving rise to metabolic pathways which are ordered reactions for conversion of substrate to product (Fani, 2012). Metabolism is old, having 2

evolved before the divergence of the last universal common ancestor (Delaye et al., 2005). The first enzymes are suggested to have had broad substrate specificity and to catalyze multiple reactions (Yčas, 1974). Early metabolic networks contained enzymes that were highly interconnected and if each enzyme was represented as a node with reactions as connections, nodes would be few with multiple links (Alves et al., 2002). Gradually, through mutation, gene fusion, small scale and large- scale duplication events, horizontal gene transfer and transposon insertions followed by neofunctionalization, the metabolic networks expanded capabilities along with genome size. Over time, the number of nodes increased and the independence of nodes grew (Fani, 2012). Horowitz posited the retrograde hypothesis of metabolic evolution, where enzymes evolved in the reverse order beginning with the essential metabolites and evolving out toward the most dispensable nutrient (Horowitz, 1945). This is in contrast with the recruitment hypothesis or patchwork hypothesis of enzyme evolution where the first enzymes encoding multiple functions were duplicated and evolved increasing specificity and narrower substrate range in concert with evolution of metabolic pathways (Jensen, 1976; Yčas, 1974). In my work, both novel enzymes are examples of the recruitment hypothesis: Tda5 and Das2 belong to enzyme classes containing multiple homologs in the same species.

After gene duplication, sub-functionalization also occurs where the activity of an enzyme is spread over two genome duplicates, so that both copies are retained (Force et al., 1999; Stoltzfus, 1999). One would expect that complete functional overlap would lead to robustness. This is not the case, as functional redundancy leads to reduced selective coefficients on both genes, which is evolutionarily disadvantageous. Gene duplicates are instead specialized and full backups are not maintained (Brookfield, 1992). Where one enzyme duplicate becomes more efficient, pseudogenization of the second copy can occur since the function of the second copy is no longer strongly selected.

Duplications lead to new enzyme functions as well as to expansion of metabolic pathways and it follows that larger extant genomes tend to encode a larger number of metabolic capabilities. Analysis of fatty acid biosynthesis and β-oxidation pathways in Escherichia coli K12 has suggested that these two pathways emerged from gene duplication as there is much functional overlap between the metabolites and enzymes involved and reactions essentially only differ in acyl carrier protein coenzyme and direction of reaction (Wakil and Esfahani, 1971; Díaz-Mejía et al., 2007). In yeast 16% of the genome encodes paralog pairs, however, 44% of all metabolic genes are paralogs displaying clear enrichment for evolutionary backups (Kuepfer et al., 2005). In contrast, this dissertation highlights examples of interdependent isozymes, which presents an unusual case as 3

most paralogs have retained their function and deleting one does not have a large fitness cost (DeLuna et al., 2008). Metabolism is scattered with enzyme complexes composed of non- paralogous members that are noted for interdependence such as the mitochondrial inner membrane complex required for ubiquinone biosynthesis, where if any single member is deleted, none of the enzymes execute catalytic steps independently, leading to accumulation of precursor (González- Mariscal et al., 2014). In the coming results chapters, I will discuss the case of interdependent soluble enzymes Das2, Urk1 and Fur1 that overlap in biochemical functions. The most famous example of interdependent isozymes in yeast are Pfk1 and Pfk2 which form a hetero-octameric complex (Kopperschläger et al., 1970). Neither of these paralogues can be deleted without a substantial decrease in specific growth rate, providing a rare example of mutual dependency of paralogs. In most cases, one paralog is dispensable and genetic redundancy is exemplified within the glycolysis pathway as Saccharomyces cerevisiae encodes duplicates for all 12 reactions from glucose to ethanol except fructose-1,6-bisphosphatase. All paralogs have retained function with the exception of the GPM2 and GPM3 pseudogenes of the functional phosphoglycerate mutase GPM1. Solis-Escante and colleagues prepared a yeast strain with minor paralogues deleted and found no major changes in metabolic fluxes or specific growth rates on glucose during fermentation/diauxic shift and under anaerobic conditions (Solis-Escalante et al., 2015). Although reactions under high flux are more likely to have a backup paralog (Papp et al., 2004), deletion of the minor paralogues appears to have a neutral effect on specific growth rate. These results do not support the backup theory of fixation for duplicated genes by increasing an organism’s ability to buffer mutations and increasing its evolvability, and instead demonstrates distinct expression of duplicates (Solis- Escalante et al., 2015). Consistent with this, most synthetic negative genetic interactions were observed when nutritional conditions were altered, and only ~15% occurring across all conditions tested, showing lack of compensation under very specific conditions indicating a specialized function (Harrison et al., 2007). In addition to providing nutritional robustness, duplicates may be retained to compensate for stress (Musso et al., 2007). Genes with whole genome duplicates tend to have low numbers of genetic interactions due to genetic buffering and have asymmetric interaction partners due to conditional specialization (VanderSluis et al., 2010).

1.1.4 Prediction of enzyme function We can now predict the classes of functions encoded in genomes by identification of conserved domains and phylogenetic analyses. Prediction of protein function is classically carried out by pairwise sequence analysis using BLAST, and with the wealth of genomic data as training sets subtle sequence similarities can be detected using hidden Markov models for protein family 4

classification (Yoon, 2009). One can now search functional databases such as CDD (Marchler- Bauer et al., 2017), PFAM (Finn et al., 2014) and CATH (Sillitoe et al., 2015) for the inferred evolutionary relationship and function for a sequence of interest. We can thereby discern the fraction of genomes and fraction of metabolism that is devoted to synthesis of amino acids, breakdown of carbohydrates and detoxification of xenobiotics. Not all enzyme classes have been elucidated and as more functional experiments are carried out, researchers find more cases of convergent evolution. As of 2010, there were 185 known enzyme commissions for which empirical evidence has shown similar reaction, with no detectible sequence homology (Omelchenko et al., 2010). For example, functional enzyme discovery I helped carry out during my undergraduate studies led to the discovery of a novel β-galactosidase that was not predicted based on conserved domains from sequence information (Cheng et al., 2017). Furthermore, there are cases of enzyme families with no known structures or functional characterization discussed below. With the advancement of bioinformatics tools for phylogenetic analysis and homology detection, new functional relationships are continually uncovered.

1.1.5 Prediction of enzyme substrates A major barrier to predicting the metabolism of an organism is not determination of the type of reaction catalyzed, but instead the prediction of substrates based on conservation of functional domains. A wealth of structural data has been collected annotating the folding of proteins and their interactions with molecular ligands and methods for prediction of substrates based on the structures of binding surfaces have also been developed, but challenges remain in substrate prediction as structures having less than 2 Å backbone deviation even when sequence identity drops below 25% (Berman et al., 2000; Lichtarge et al., 1996). Indeed, for some enzyme classes, it is not possible to predict substrate from amino acid sequence alone. This may result from a large number of mis- annotations resulting in inclusion of multiple structures for reactions that are chemically dissimilar. For pairwise amino acid sequence alignments having over 50% similarity, less than 30% are classified to the same enzyme commission (Rost, 2002). Functional predictions for uncharacterized proteins are enhanced by inclusion of gene ontology term annotations into predictive models alongside sequence identity (Clark and Radivojac, 2011; Jiang et al., 2016).

A major challenge in predicting the function of enzymes is the lack of experimental data supporting associations. In prokaryotes the genomic context gives clues on encoded functions due to the ordered arrangement of genes in operons, however this is not generally conserved to eukaryotes and functional insights cannot be gleaned from chromosomal context (Fondi et al., 2009; Jacob et al., 5

1960). Modern tools for the identification of homologues are effective, however detecting deviations in function from the reference sequence has proven difficult alongside interpretation of concerted shifts in amino acid preference. For large enzyme super-families where genomic data is rapidly generated, it is still difficult to predict ligand and mechanistic structure-function relationships due to lack of empirical evidence of activities (Brown and Babbitt, 2014). Indeed, the ability to predict functional relationships has outpaced our ability to test new functions and outliers can only be determined once mechanisms are experimentally supported.

1.1.6 Domains of unknown function With the wealth of annotated proteins a functional paradox exists as 20% of domains in PFAM are hitherto uncharacterized conserved “domains of unknown function” (DUF) (Punta et al., 2012). These domains are especially relevant because prokaryotic DUFs predominantly encode essential functions (Goodacre et al., 2014). Between 2013 and 2016 there were significant advances in computational methods for predicting the biological processes that proteins are annotated to as predictive algorithms had improved alongside advances in functional data (Jiang et al., 2016). These developments expedite researchers’ ability to develop hypotheses and design experiments to more rapidly deduce gene function. In 2017 the PFAM database contained nearly 4000 DUF families, for which 50 inferences of structure could be made based on similarity to known folds and catalytic residues and thereby ascribed tentative functions (Mudgal et al., 2015). Threading algorithms go as far as suggesting the primary enzyme commission, but it remains a very high bar to determine substrate. Furthermore, knowing the type of reaction does not define the biological process or role the enzyme is carrying out. However, the prevalence of essential DUFs again underscores the need for experimental validation of enzymatic mechanism.

1.1.7 Metabolic networks contain missing enzymes Understanding the complex biological interactions that govern the way cells respond to internal or external changes at a systems biology level requires a high level of annotation and completeness of genome scale metabolic networks (Palsson, 2002). It has been established that there are uncharacterized genes that are not yet added to the network. At the same time, there is empirical evidence for reactions that occur that have not yet been linked to the enzymes that catalyze them, known as orphan reactions. This is a longstanding research question as in 2004, 38% of experimentally observed reactions had no annotated protein sequence (Karp, 2004). There are also reactions that are completely missing, creating dead-ends in the metabolic network (Orth and Palsson, 2010). Depending on the model used, at least 30% of reactions used in genome scale 6

metabolic network reconstructions are not yet assigned to enzymes (Heavner and Price, 2015). Indeed, a portion of these may be spontaneous or metal catalyzed, however most transformations are associated with barriers, such as low natural abundance of non-enzymatic catalysts and the high frequency of inefficient off-target reactions, that impose a requirement for enzymatic catalysis (Keller et al., 2015). These missing enzymes represent knowledge gaps for an organism’s metabolism. In yeast, examples of orphan reactions include aromatic amino acid breakdown and cytidine activation to CMP which is addressed in this work. Furthermore, I have studied two enzymes that are not currently added to the yeast metabolic network. In one case, it was missed because it is an obligate part of a complex although the reaction was included in the network and the other was missed due to genetic suppression and its reaction remains unknown and unconnected.

1.1.8 Metabolic networks are missing metabolites In addition to missing enzymes, gaps in metabolic networks also occur due to missing metabolites. Dead ends in metabolic networks manifest in two ways: root no consumption metabolites or root no production metabolites, where the following or preceding metabolite and reaction is unknown (Satish Kumar et al., 2007). Gap-filling algorithms can be used to predict missing metabolite- enzyme pair functions leading to annotations in cases homologs are readily identifiable (Reed et al., 2006). A third type of dead-end metabolite lacks both production and consumption reactions and is described as an “island”. In the case of island metabolites, the unknown metabolic pathway the island metabolite functions in can be suggested by algorithmic prediction of intermediary reactions (Kumar and Maranas, 2009). Experimental validation of the biochemical predictions used to resolve gaps in genome scale metabolic network reconstructions has led to the discovery of new enzymes and transporters (Kumar and Maranas, 2009; Reed et al., 2006). Full-scan untargeted liquid chromatography-mass spectrometry metabolomics, as detailed below in genetic screening for enzyme discovery, has the capability of identifying these missing metabolites as an unbiased and global approach is taken to identification of mass spectral peaks.

1.1.9 New metabolic pathway information from fluxomics Throughout this study, steady state measurements of metabolite levels are used to identify cases of metabolite accumulation due to enzymatic loss of function. Steady state measurements of enzyme levels and metabolite levels do not capture the magnitude of metabolic flow through a given metabolic pathway. Also, the necessity of measuring large populations of cells means that measurement of metabolite levels may fail to capture dynamics of these changes as cells grow and 7

divide. In contrast, stable isotope tracing—measuring abundances of positionally labeled 2H, 13C, and 15N compounds compared to 1H, 12C and 14N counterparts as the compound passes through a metabolic pathway—allows for time-resolved metabolite level measurements and calculation of metabolic fluxes (Fiehn et al., 2000; Yuan et al., 2008). In order to quantify metabolism, it is necessary to measure and model multiple metabolic reaction rates in parallel using the help of stable isotope tracers in fluxomics studies where the relative distribution of isotopically labeled and unlabeled metabolite is determined over time (Wiechert, 2001). Recent measurements of global metabolic fluxes in synchronized cells have shed light on the temporal significance of metabolic reactions and predominance of one pathway over another a within metabolic network (Ahn et al., 2017). Fluxomic studies are crucial to understanding metabolic network connectivity and determining mechanisms of metabolic compensation in nutritionally, chemically or mutationally perturbed systems (Chokkathukalam et al., 2014; Jang et al., 2018). Fluxomics is useful in capturing temporal changes in reaction rates that are previously unknown, further resolving annotations of known metabolites and enzymes (Ahn et al., 2017). An additional benefit of flux estimation in hand with matched temporal proteomic datasets is predictions of how metabolite ligands act allosterically to activate or inhibit enzymes as uncovered by a systematic study of regulatory ligands in yeast (Hackett et al., 2016). As more global and temporal comparisons of metabolite levels are carried out and more relationships between genes, proteins and metabolite levels are uncovered, ability to predict missing reactions and enzymes in the metabolic network will improve.

1.1.10 Enzyme discovery through metabolomic screening in yeast Functional metabolomics combines genetics with multi-parallel analysis of metabolite levels and enables predictions of metabolic functions of uncharacterized genes and discovery of underexposed aspects of metabolism. In 2014, I set out to assign function to uncharacterized candidate enzyme- encoding genes (Caudy et al., 2018). A list of 141 non-essential uncharacterized genes encoding conserved enzymatic domains or conserved domains of unknown function was prepared by my supervisor. Deletion mutants for 120 candidate enzymes were obtained from the prototrophic yeast deletion collection (Gibney et al., 2013; VanderSluis et al., 2010). I back-crossed and selected six independent haploid isolates representing each deletion mutant to mitigate the presence of second site mutations causing metabolomic phenotypes. Freshly back-crossed candidate enzyme deletion mutant isolates were grown over several rounds of exponential growth and sub-culture so that cells could habituate to minimal media and extracted metabolites while cells were in mid-log phase. Metabolites were separated and quantified by reversed phase ion paired chromatography hyphenated to accurate mass q-TOF mass spectrometry. In order to capture metabolic phenotypes in 8

metabolites not captured by a set of commercially available metabolite standards, I took an untargeted approach to identification of peaks in ion intensity by using a peak picking algorithm available within the Agilent Profinder software. Identified peaks were integrated across mass spectral datasets derived from wild-type and candidate enzyme controls. I developed a pipeline in R for automated data visualization and statistical comparison of changes in mean ion intensity between wild-type and mutant by unpaired t-test. I identified 26 candidate enzyme deletion mutants that had greater than 2-fold changes in metabolite levels, with a student’s t-test p-value of less than 0.001 (Caudy et al., 2018). These metabolite level changes were present for known metabolites in pathways across the metabolic network from purine and pyrimidine nucleotide metabolism, amino acid breakdown and long chain fatty acid biosynthesis. In comparison, a recent targeted study of amino acid levels measured across the entire nonessential yeast deletion collection with restored prototrophy found that most uncharacterized proteins clustered with known proteins within a network of phenotypes but did not identify any new enzymatic activities (Mülleder et al., 2016). My studies of two of the candidate enzymes uncovered from my screen comprise the body of this thesis.

1.2 Pyrimidine metabolism of Saccharomyces cerevisiae In Chapter 3, I will describe the uncharacterized enzyme, Das2 and show its role in pyrimidine salvage.

1.2.1 UMP is the central pyrimidine metabolite A key metabolite in pyrimidine metabolism is uridine monophosphate (UMP), which gives rise to all cellular pyrimidine metabolites and allows for multiple fundamental processes and functions. In Saccharomyces cerevisiae, UMP is phosphorylated and aminated to form the essential nucleic acids uridine 5’ triphosphate (UTP) and cytidine 5’ triphosphate (CTP) which are polymerized into RNA chains. Alternately, ribonucleotides are reduced to deoxy forms and dUMP is methylated and phosphorylated to dTTP for incorporation into DNA chains. Phosphorolytic addition of UTP to the sugar amine N-acetyl-glucosamine-6-phosphate gives rise to UDP-N-acetyl-D-glucosamine (UDP- GlcNAc), which is the precursor to cell and spore wall polymers chitin and chitosan. Likewise, UTP addition to glucose-1-phosphate provides UDP-glucose for lipid-linked oligosaccharide biosynthesis (Burda and Aebi, 1999), trehalose synthesis (Bell et al., 1998), and glycogen biosynthesis (François and Parrou, 2006). Whereas the pyrimidine nucleotides UTP and TTP do not act as phosphate group donors except in activating nucleoside diphosphates (Jong and Ma, 1991), CTP is noted as an energy source for activation of intermediary lipid metabolites functioning as the primary phosphate donor for synthesis of phospholipids in formation of CDP-diacylglycerol (Chang and Carman, 2008; Shen et al., 1996). These comprise the well-known functions of pyrimidines produced from UMP.

1.2.2 de novo biosynthesis of UMP As all cellular pyrimidines are derived from UMP, its biosynthesis is well studied with initial elucidation in Escherichia coli where the genes encoding enzymes and their regulatory elements are ordered within an operon (Beckwith et al., 1962). Parallel enzymes and metabolites comprising the UMP biosynthetic pathway in Saccharomyces cerevisiae were later identified and are conserved across all branches of life (Delaye et al., 2005; Lacroute, 1968). As demonstrated in Figure 1, the yeast pathway begins with action of the bifunctional Ura2 enzyme which not only encodes domains for aspartate transcarbamylase (with glutamine amidotransferase) and carbamyl phosphate synthase activities, but also has a non-functional dihydroorotase domain (Berman et al., 2000; Lacroute, 1968). During the initiating step, Ura2 carries out the carbamyl phosphate synthetase reaction, which aminates carbonate to carbamoyl phosphate at the expense of 2 ATP. In the second step, the energetically favorable release of phosphate is coupled to aspartate addition to deliver carbamoyl 10

aspartate. The third step is carried out by the essential dihydroorotate amidohydrolase, Ura4, where the amide group attacks the C4 carbonyl to cyclize carbamoyl aspartate to release 4,5- dihydroorotate and water. The six-member ring is aromatized by oxidation carried out by fumarate dependent dihydroorotate dehydrogenase (DHOase), encoded by URA1 (Guerry-Kopecko and Wickner, 1980). Budding yeast DHOase is distinct from that found in fission yeast, Schizosaccarhomyces pombe, where 80% of the ubiquinone dependent dihydroorotase dehydrogenase is mitochondrially localized and activity is restricted to aerobic growth as electrons are transferred to molecular oxygen via the electron transport chain, while in S. cerevisiae, the enzyme is localized to the cytosol and appears to be more similar to eukaryotic homologues (Nagy et al., 1992; Voříšek J. et al., 2002). The S. pombe DHOase is similar to the E. coli homologs, where the electrons are donated to the electron transport chain for aerobic growth, while S. cerevisiae uses fumarate as the terminal electron acceptor from dihydroorotate to form succinate and orotate (Nagy et al., 1992). In the fourth step of de novo UMP production, orotate is phosphorylated to 5’ orotidine monophosphate by the actions of two orotate phosphoribosyl transferases encoded by the paralogues URA5 and URA10 (de Montigny et al., 1989)(de Montigny et al., 1990). The final step in de novo biosynthesis of UMP is catalyzed by Ura3, which decarboxylates orotidine-5’-monophosphate to uridine-5-monophosphate (Lacroute, 1968). As described below, Ura3 is widely used as an auxotrophic marker for positive and negative selection as it is the target of 5-fluoroorotic acid (Boeke et al., 1984).

Figure 1. Metabolic Pathway for de novo Biosynthesis of Pyrimidines Structures of intermediates are given on the left, while pathway intermediates and metabolic reactions are shown through the centre and enzymes catalyzing each step are listed on the right. Broken lines indicate regulatory relationships between metabolites and targets. 1.2.3 Regulation of UMP biosynthesis The de novo biosynthetic pathway for pyrimidine biosynthesis is subject to regulation at several levels. The entire pathway is subject to negative feedback control as expression of URA2 encoding the enzyme catalyzing initial steps of the pathway is inhibited by the final metabolite in the pathway, UTP, and by the salvage intermediate uracil indicating that de novo biosynthesis and salvage of pyrimidines are balanced (Potier et al., 1990). Furthermore, inhibitors of Ura3, such as 6- azauracil lead to induction of URA2 transcription (Potier et al., 2006). The dihydroorotase is encoded by URA4, and is subject to positive regulation by its substrate, dihydroorotate, and the zinc finger transcription factor Ppr1 (reviewed in Denis-Duphil, 1989). Ppr1 directly senses dihydroorotate and orotic acid on its carboxy terminal to induce expression of URA1, URA3, URA4 12

and URA10 (Flynn and Reece, 1999). PPR1 is not necessary for induction of its targets as ppr1- mutants have a wild-type basal expression of URA1 and URA3 (Liljelund et al., 1984). URA1 and URA3 contain nonsense mutations in the 5’ UTR that reduce steady state levels of the transcripts without reducing the rate of transcription during repressive conditions, while the URA2 gene encodes two alternative start sites producing a cryptic unstable transcript under repressive conditions (Losson and Lacroute, 1979; Pelsy and Lacroute, 1984; Thiebaut et al., 2008).

1.2.4 Multi-phosphorylated pyrimidine nucleotides The URA6 gene encodes the essential UMP kinase which activates UMP to UDP using ATP as a cofactor (Liljelund et al., 1989) (Liljelund and Lacroute, 1986). The YNK1 gene encodes the promiscuous nucleoside diphosphate kinase (NDP kinase) known for converting UDP to UTP, CDP to CTP, dTDP to dTTP and all nucleoside diphosphates to triphosphates except ADP (Jong and Ma, 1991). Surprisingly, ynk1 mutants are viable and display normal growth rates, indicating that there must be several isozymes as the ynk1Δ strain does not appear to have pairwise synthetic lethal interactions and steps before and after the Ynk1 reaction are both essential. Ynk1 is not an abundant enzyme and, unlike the S. pombe orthologue, is not cell cycle regulated in S. cerevisiae. Ynk1 is predominantly cell membrane associated, interacts with the dTMP kinase, encoded by CDC8, which may indicate channeling (Zhang et al., 1995). CTP is produced from UTP via the CTP synthetase enzymes encoded by the synthetic lethal pair, URA7 and URA8 (Ozier-Kalogeropoulos et al., 1994).

1.2.5 Catabolism as a source of 5’ pyrimidine nucleotide monophosphates RNA catabolism provides the intermediary pools of pyrimidines involved in the breakdown and recycling of ribonucleic acids. Yeast cells are continuously turning over their transcriptome of non- coding RNAs, transfer RNAs and messenger RNAs along with ribosomal RNAs in response to cell cycle position and adaptation to environmental conditions. RNA decay is the process of enzymatic degradation of RNA molecules to produce RNA oligomers and monomers. The RNA decay mechanism allows for quality control of mRNA to ensure production of functional proteins. RNA decay is responsible for degrading excised intronic RNAs as well as routine turnover carried out by mRNA surveillance pathways in the cytoplasm and nucleus. Cells employ several mechanisms for RNA decay in two differentially localized exosome complexes, notably the SKI complex in the cytoplasm and the TRAMP complex in the nucleus (LaCava et al., 2005). Each compartment and complex is equipped with nucleases catalyzing breakdown of RNA. In the cytoplasm, decapped and deadenylated mRNAs are degraded by the 5’ to 3’ exonuclease encoded by XRN1 (Dykstra et al., 13

1991) and by exosomal catalysis via the highly processive 3’to 5’ exoribonuclease/weak endonuclease Dis3 (Chlebowski et al., 2013). In the nuclear exosome, 3’to 5’exonuclease activity is carried out by Rrp6 (Chlebowski et al., 2013). Xrn1, Dis3 and Rrp6 ribonuclease activities produce 5’ mononucleotides (Stevens, 1980) (Dziembowski et al., 2007).

S. cerevisiae encodes a repertoire of endoribonucleases with specialized roles that include: unfolded protein response site specific ribonuclease Ire1 (Gonzalez et al., 1999), Ysh1 functioning in 3’ processing of mRNAs and snoRNAs (Garas et al., 2008), Ngl1,2,3 functioning in processing of ribosomal RNAs (Chen et al., 2002), Dom34 catalyzing RNA cleavage in no-go decay (Lee et al., 2007), Rcl1 for cleaving pre-rRNA (Billy et al., 2000), Nob1 for cleaving ribosomal RNAs (Fatica et al., 2003) and Swt1 for processing anomalous pre-mRNAs (Skružný et al., 2009). Specialized exoribonucleases include Ccr4 involved in the Ccr4-NOT transcriptional complex for modulating gene expression in response to catabolite repression (Tucker et al., 2001), Usb1 for cleaving poly(U) tracts in splicing (Mroczek et al., 2012) and the Rat1 5’ to 3’ nuclear exoribonuclease having roles in regulating RNA Pol II transcript elongation (Amberg et al., 1992).

1.2.6 During autophagy 3’ pyrimidine nucleotides are hydrolyzed to nucleosides In contrast to production of 5’ ribonucleotide monophosphates by Xrn1, Dis3 and Rrp6, the formation of 3’mononuceotides has not been thoroughly studied in yeast. The enzymatic production of 3’ nucleotide monophosphates was identified in 1973, however the genes encoding these enzymes and their regulation was, until recently, poorly understood (Dauber, 1973). Nitrogen starvation induces non-selective autophagy (macroautophagy), a process whereby swaths of cytoplasmic components are engulfed into autophagosomes for degradation in the vacuole where ribosomal RNAs are catabolized to nucleotide components (Takeshige et al., 1992). Recently, Huang and colleagues found that during nitrogen starvation induced autophagy, cellular levels of nucleosides increased up to one hour post nitrogen starvation, and this was extended up to 1.5 hours for uridine (Huang et al., 2015). Upon dissecting the origin of these nucleosides, the authors found that vacuolar RNA is degraded to 3’-nucleotides by the acid ribonuclease, Rny1, and further hydrolyzed to nucleosides by Pho8 (Huang et al., 2015). Pyrimidine nucleosides are released to the cytoplasm, likely by the vacuolar equilibrative nucleoside transporter encoded by ScENT1, and are hydrolyzed by the uridine/cytidine hydrolase, Urh1, before excretion as uracil (Boswell-Casteel et al., 2018; Huang et al., 2015; Jund and Lacroute, 1970a). Excretion of nucleoside bases predominates as yeast do not encode enzymes for degradation of the pyrimidine ring.

1.2.7 Pyrimidine salvage from an uptake perspective In addition to reactivation of nucleotides from degradation through salvage, organisms can take up nucleobases and nucleosides from their environment. The transport of nucleobases across cell membranes is carried out by selective carriers called permeases. Activity of high affinity permeases is rapid as cells equilibrate with their environment within seconds of exposure (Marz et al., 1979). In yeast, uracil uptake activity is carried out via the high affinity uracil permease, Fur4 (Séron et al., 1999). Negative feedback of the Fur4 permease is caused by cytoplasmic binding of uracil to the permease which alternatively sorts Fur4 to the vacuole for degradation instead of the plasma membrane (Blondel et al., 2004) and increases the rate of its turnover. Under uracil binding, the FUR4 transcript is additionally subject to negative feedback as it is turned over more rapidly. The uridine permease, encoded by FUI1 (Jund and Lacroute, 1970a; Zhang et al., 2006), is similarly sorted to the vacuole during exposure to high levels of uridine (Blondel et al., 2004). Uridine is also transported to the vacuole by the equilibrative nucleoside transporter ScENT1 (Boswell-Casteel et al., 2018; Vickers et al., 2000). Cytosine is transported through the plasma membrane by Fcy2 (Jund and Lacroute, 1970a). There is no salvage pathway for the dNTP components thymine or thymidine in Saccharomyces cerevisiae (Bisson and Thorner, 1982).

1.2.8 Pyrimidine salvage enzymes- Early studies reveal pathway architecture and activities Whether nucleobases and nucleosides originate from degradation of RNA and DNA or from uptake from the environment, they must be activated to UMP before carrying out subsequent roles. In a landmark study, while investigating uptake of exogenous pyrimidines in yeast, Grenson aimed to identify the pyrimidine permeases to show that pyrimidines are actively transported by saturable, specific and regulated processes (Grenson, 1969). In identifying permease candidates among other potential mutants in enzymatic activation of cytosine, the author aimed to show that downstream salvage to UMP was normal and therefore analyzed candidate mutants for known reactions occurring in E. coli: cytosine deaminase, uridine kinase, uridine phosphorylase and uridine hydrolase activities. Activities identified included transport of uracil, uridine and cytosine, mapped to Fur4, Fui1 and Fcy2, respectively (Grenson, 1969; Jund & Lacroute, 1970; Jund et al., 1988; Kurtz et al., 2002; Vickers et al., 2000). In addition to transporters, three salvage activities were identified including cytosine deaminase activity for conversion of cytosine to uracil. Uridine kinase (UKase) activity, which activates uridine to UMP in a single step was identified, however, this reaction was not essential for activation of uridine to UMP. In the absence of uridine kinase, uridine is instead enzymatically hydrolyzed to uracil, followed by activation to UMP by an uracil 15

phosphoribosyl transferase (UPRTase) reaction requiring PRPP. UKase mutants “udk” were not impaired for salvage from uracil to UMP, however, UPRTase mutants “ups” showed reduced UKase activity. This is the first example showing that uridine kinase activity is dependent on a second salvage activity in yeast (Grenson, 1969). It is also demonstrated that mutation of the uridine hydrolase suppresses secretion of uracil and uridine is excreted instead, indicating that uracil is predominantly formed by hydrolysis of uridine (Grenson, 1969).

Pyrimidine analogs represent an important class of antineoplastic, antiviral and antifungal agents that require intracellular activation to nucleotide forms that inhibit nucleic acid synthesis (De Clercq, 2011; Galmarini et al., 2003; Vermes et al., 2000). Since mutations in pyrimidine salvage reactions lead to resistance to the analogue, Jund and Lacroute examined resistance to the 5- fluoropyrimidines: 5-fluorouracil, 5-fluorouridine and 5-fluorocytosine to determine pyrimidine salvage reactions in Saccharomyces cerevisiae and identified UPRTase, uridine kinase, uridine phosphorylase, uridine hydrolase cytidine deaminase and cytosine deaminase activities (Jund and Lacroute, 1970). Mutations in FCY1 were deficient in cytosine deaminase activity while FCY2 was identified as encoding the cytosine permease (Jund and Lacroute, 1970). Jund and Lacroute also provided the second observation that across various Fur1 UPRTase mutants, strains displayed a decrease in uridine kinase activity. Between the work of Grenson, Jund and Lacroute, the topology of the UMP salvage pathway was defined in these early studies and is illustrated in Figure 2.

1.2.9 Pyrimidine salvage - Purification of enzymes and cloning of genes Isolating proteins from complex cellular homogenates is crucial to understanding properties of the enzyme at study. The first salvage enzyme for which purification was achieved was the uridine hydrolase (Carter, 1951; Magni et al., 1975). This was followed by purification of a UPRTase complex in showing that yeast produced two UPRTases, one of 27,000 Da and one of 58,000 Da (Natalini et al., 1979). By 1990, mutations in the FUR1 locus had been physically mapped and led to cloning of the FUR1 gene (Kern et al., 1990b). Distinct point mutants in Fur1 were sequenced and either showed diminished UPRTase activity or diminished expression of Fur1 in response to exogenous uracil (Kern et al., 1991). Uridine kinase activity was annotated to the URK1 gene as a clone harbouring the Urk1 coding sequence complemented a uridine kinase deficient mutant (Kern, 1990a). Hitherto, no purification of the yeast uridine kinase has been carried out and the uridine kinase mutant involved in the study has not been mapped or cloned.

Figure 2. Conventional model of UMP salvage in Saccharomyces cerevisiae 1.2.10 Project rationale: Resolving pyrimidine salvage in yeast Pyrimidine salvage pathways are essential to activation to important classes of antineoplastic and antifungal inhibitors and are critical to understanding the balance between nucleotide catabolism and synthesis. Up to now, the roles Das2 and its homolog, Urk1, provide in pyrimidine salvage have not been described. The topology of the pyrimidine salvage pathways suggests that uridine is salvaged directly through the uridine kinase reaction, forgoing the need for the hydrolase and UPRTase pathway, except when uracil is salvaged from the media. Furthermore, the synthetic lethality between fur1Δ and ura3Δ suggests that salvage is dependent on the presence of Fur1, however the basis of this dependency has not been determined. My studies aim to resolve the role of Das2 in pyrimidine salvage. I will describe the contributions of Fur1, Urk1, Urh1 and Das2 to pyrimidine salvage, resolve the relative uridine kinase and UPRTase activities catalyzed by these enzymes and determine the basis of interdependence of these enzymes for stability and activity.

1.3 Background for Tda5: a novel short chain dehydrogenase essential for respiration In Chapter 4, I will describe studies toward delineation of the function of the uncharacterized short chain dehydrogenase, Tda5. I will describe phenotypes associated with genetic knock-out including perturbed mitochondrial metabolism as well as mechanisms of suppression of tda5Δ: by de-

repression of a compensatory paralog, Ydl114w, and by recessive mutations in ergosterol biosynthesis.

1.3.1 Oxidoreductases and short chain dehydrogenases Transfer of electrons through enzymatic catalysis of oxidation/reduction reactions require cofactors such as the pyridine nucleotide derivatives NAD+ and NADPH or porphyrin-based heme. Short chain dehydrogenases (SDR) comprise a large class of single domain enzymes that catalyze NAD(P)(H) dependent reactions distinct from the multidomain medium and long chain dehydrogenase/reductases (Ghosh et al., 1991). These enzymes act on a wide array of substrates including sugars, alcohols, steroids and aromatic compounds and catalyze a variety of reactions. The family is highly divergent as 15-30% amino acid identity is observed in pairwise analyses compared to a median of 35% for pairwise analysis within all protein superfamilies (Atkinson et al., 2009; Jörnvall et al., 1995; Kallberg et al., 2002). NAD(P)(H) binding occurs in a conserved Rossman fold sequence motif GXXXGXG (Jörnvall et al., 1995). SDRs contain a highly conserved triad of serine, tyrosine and lysine residues where the single ubiquitously conserved residue homologous to Tyr152 of the human 3α/20β-hydroxysteroid dehydrogenase, the first SDR for which a crystal structure was obtained (Ghosh et al., 1991; Jörnvall et al., 1995). Determination of the structure revealed that the folding of the protein was much different than long chain dehydrogenases, defining the “long chain” and “short chain” designations (Ghosh et al., 1991). Since then, structures have been determined that show a consistent folding backbone for SDRs. There are several classical SDRs which are ~250 amino acids in length with a conserved Rossman fold such as carbonyl reductases and steroid dehydrogenases, versus extended SDRs that are ~350 with a variation on a Rossman fold isomerases, epimerases, lyases (Persson et al., 2003). The NADP+ dependent 17β-hydroxysteroid dehydrogenase was isolated in 1972 by Mulder and colleagues and had a broad substrate specificity, catalyzing oxidation of androstanes, and estradiol to estradione (Mulder et al., 1972). Chapter 4 concerns the biology of novel short chain dehydrogenases Tda5 and Ydl114w in yeast.

1.3.2 SDR activity pervades lipid homeostasis Analyses detailed in Chapter 4 indicate that Tda5 and Ydl114w alongside all of their characterised yeast homologs act in lipid metabolism. The major classes of lipids in yeast are phospholipids, fatty acids, sphingolipids and heterocyclic sterols (Rattray et al., 1975). Long chain fatty acids are esterified with alcohols such as glycerol to form neutral lipid triacyl-glycerol or glycerol-3- phosphate to form charged phospholipids, which are in turn esterified to serine or inositol (reviewed 18

in Kohlwein, 2010). Ergosterol is the main sterol in yeast, the functional equivalent of cholesterol in humans (reviewed in Sturley, 2000). Ergosterol is found esterified to fatty acids, forming neutral sterols while free sterols contain a hydroxyl group and are amphipathic. Together, free ergosterol and phospholipids form hydrophobic membrane bilayers such as the plasma membrane and also provide barriers for excluding polar materials from organelles (reviewed in Klug and Daum, 2014). Although phospholipids comprise one half of total lipids, there is twice as much ergosterol in the plasma membrane as phospholipid (reviewed in Fraenkel, 2011). Sphingolipids are unsaturated long chain fatty acids with a serine-derived amino alcohol base, are low in abundance and enriched in the yeast plasma membrane (reviewed in Dickson, 2010).

The localizations and activities of all of the characterized SDRs in Saccharomyces cerevisiae are shown in Figure 3. Reductive activities localized to the endoplasmic reticulum include 3- ketosphinganine reductase (Tsc10), acyl-DHAP reductase (Ayr1) and long chain β-keto reductase (Ifa38) (Athenstaedt and Daum, 2000; Beeler et al., 1998; Huh et al., 2003). The β-keto-acyl reductase (Oar1) resides within the mitochondrion (Schneider et al., 1997). The latter steps of the ergosterol biosynthetic pathway including the 5-cholesta-dieneone reductase (Erg27) are localized to lipid droplets (Huh et al., 2003). The oxidative reactions catalyzed by β-keto acyl dehydrogenase- enoyl-CoA hydratase (Fox2) and dienoyl-CoA reductase (Sps19) are localized to peroxisomes.

Figure 3. Localizations of characterized short chain dehydrogenases in Saccharomyces cerevisiae

1.3.2.1 Fatty acids

Free fatty acids in yeast are usually found in C16 and C18 lengths, namely palmitic and stearic acids, and can be thought of as a polymerized and saturated form of acetyl-CoA. De novo biosynthesis is catalyzed by the fatty acid synthase Fas1/Fas2 holoenzyme containing NADPH dependent SDR activity of keto-acyl reductase in the cytosol from acetyl-CoA and malonyl-CoA (reviewed in Tehlivets et al., 2007). A suite of isozymes act in the mitochondrion where 2-keto-acyl reductase activity is carried out by Oar1, a short chain dehydrogenase (Venkatesan et al., 2014). Fatty acid biosynthesis is distinct in yeast from the human pathway which relies on ATP citrate lyase for production of acetyl-CoA whereas the yeast fatty acid biosynthesis pathway relies on pyruvate dehydrogenase. Malonyl-CoA is formed from acetyl-CoA and bicarbonate via catalysis by acetyl- CoA carboxylase Acc1 and its mitochondrial counterpart Hfa1 (reviewed in Hiltunen et al., 2005).

Fatty acids having chains longer than C16 and C18 up to C24 in length are extended using the elongase enzymes Elo2 and Elo3 along with malonyl-CoA as a precursor and takes place in the endoplasmic reticulum, whereas salvaged fatty acids are extended by Elo1 (reviewed in Klug and Daum, 2014). Around 80% of yeast fatty acids are monounsaturated (Tehlivets et al., 2007). In contrast to human cells that encode a repertoire of lipid desaturases, Ole1 is the sole lipid desaturase in yeast, introducing a double bond between carbons 9 and 10 on palmitoyl and stearyl-CoAs (Stukey et al., 1989).

1.3.2.2 Sphingolipids Sphingolipid biosynthesis is an important target of antifungal drugs as phosphoinositol-containing sphingolipids are not present in humans (Rollin-Pinheiro et al., 2016). Sphingolipids are a class of lipids derived from condensation of mainly palmitoyl-CoA and serine to form the short lived intermediate 3-ketodehydrosphingosine for which sphingolipids are named in reference to the elusive sphinx (Dickson, 2010). This long chain base can be N-acylated, or hydroxylated and these bases can be further modified with polar head groups to inositolphosphoceramide, mannose inositol phosphoceramide (M-IP2-C) 75%, and mannose-(inositol-P)2-ceramide added in the Golgi. Sphingolipids are predominantly localized to the plasma membrane (Kitagaki et al., 2007). Ergosterol and sphingolipids must interact to form stoichiometric complexes called lipid rafts that function in bringing together proteins required for wildtype levels of pheromone signalling (Jin et al., 2008).

1.3.2.3 Lipids from Phosphatidic acid The pathways for glycerophospholipids, phosphatidylinositol (PtdIns) and neutral triacylglycerol each begin with condensation of DHAP or glycerol with long chain fatty acid acyl CoAs to form 1,2-diacylglycerol-3-phosphate which is either acylated to form neutral lipid triacylglycerol or activated to CDP-diacylglycerol (CDP-DAG) for incorporation into PtdIns synthesis. The PtdIns synthase catalyzes an essential role in forming PtdIns from CDP-DAG and inositol (Nikawa and Yamashita, 1984). PtdIns is the substrate for polyphosphorylation by phosphatidylinositol kinases where these downstream polyphosphorylated metabolites are involved in morphogenesis and regulation of metabolic processes (Odorizzi et al., 2000). PtdIns are essential for synthesis of glycosylphosphatidylinositol (GPI) anchors (Lykidis, 2007).

Several metabolic pathways for phospholipid biosynthesis are localized to the mitochondrion. After CDP-DAG is condensed with serine to give phosphatidylserine, it is decarboxylated to phosphatidylethanolamine within the mitochondrion before methylation to produce phosphatidylcholine (Daum et al., 1998). Phosphatidylserine decarboxylase coats the inner mitochondrial membrane (Kuchler et al., 1986). The inner mitochondrial membrane is low in sterols but rich in cardiolipin, a heavily alkylated diphosphatidylglycerol lipid formed from CDP-DAG. Cardiolipin is synthesized within the mitochondrion and is required for maintenance of mitochondrial membrane potential (reviewed in Fraenkel, 2011).

1.3.2.4 Sterols and isoprenoids from the mevalonate pathway Fungi were the some of the earliest eukaryotes to produce sterols (Parks and Casey, 1995). Sterols are vital membrane components allowing organisms to adapt to changes in temperature, however come at a high metabolic cost. Sterols are derived from a series of condensations of acetyl-CoA to HMG-CoA followed by reduction to mevalonate (reviewed in Karst and Lacroute, 1977), and the pathway is shown in Appendix I. Mevalonate is phosphorylated by Erg12, the mevalonate kinase, and then by Erg8, the mevalonate-5-phosphate kinase, which is then carboxylated and dehydrated to form the activated isoprene subunit isopentyl-pyrophosphate. Six isopentyl-pyrophosphate subunits are isomerized and condensed to squalene in the mevalonate pathway. Squalene is cyclized and oxidized to form lanosterol and ergosterol, the major sterols in yeast. Only a single methyl group is added beyond this point. Erg1 is the squalene epoxidase and is the first enzyme in the lower ergosterol biosynthetic pathway (reviewed in Karst and Lacroute, 1977). The final modifications of lanosterol to ergosterol involves reduction, removal of methyl groups including the action of a short chain dehydrogenase Erg27, addition of a methyl group from S-adenosylmethionine, isomerization 22

of double bonds and desaturations by oxidoreductases. The lower ergosterol pathway involves use of 14 NAD(P)H, 1 S-adenosylmethionine and 11 molecules of O2 to form ergosterol from 1 molecule of squalene. The high energetic cost of ergosterol biosynthesis is offset by storage of sterol intermediates in lipid particles in an esterified form by condensation with fatty acid acyl CoA . Ergosterol depletion inhibits lipid raft formation which is required for signalling events, and decreases cell fusion and cell membrane remodelling during mating (Jin et al., 2008).

In Chapter 4, recessive mutations in the mevalonate pathway will be discussed alongside lesions occurring downstream in the lower portion of the ergosterol biosynthetic pathway. In addition to sterol biosynthesis, the mevalonate pathway contributes isoprenoids to other metabolic pathways. The polyisoprenyl chain on Coenzyme Q is formed from prenyl subunits derived in the mevalonate pathway (Tran and Clarke, 2007). Farnesyl-pyrophosphate is used to form dolichyl-phosphate necessary for glycosylation of proteins in the endoplasmic reticulum and for formation of glycophosphatidylinositol (GPI) anchors necessary for docking membrane proteins (GrabiÅska and Palamarczyk, 2002). Protein modification by prenylation is crucial to localization of signal transducers such as Ras to the plasma membrane (Dimster-Denk et al., 1999).

1.3.2.5 Sterol biosynthesis is targeted by anti-mycotics The most commonly used antifungals are azoles which target the ergosterol biosynthetic pathway by binding the porphyrin ring of the cytochrome P450 enzyme 14α-lanosterol demethylase, encoded by ERG11. When cells are treated with azoles, 14α-methyl-3,6-diol, accumulates where the β-3-hydroxy group interferes with the native interaction between sterol and phospholipid within membranes, leading to toxicity (Kelly et al., 1995). Erg11 is the essential target of azole anti- mycotics, although hypomorphs can be rescued by a mixture of ergosterol and fatty acid under anaerobic conditions (Kelly et al., 1995). Saccharomyces cerevisiae does not take up ergosterol under aerobic conditions. Sterol uptake under low oxygen conditions is dependent on the zinc finger transcription factor, UPC2, which is named for uptake control (Zavrel et al., 2013). Mutations in UPC2 enable cells to take up sterol under aerobic conditions.

1.3.3 Membrane fluidity and structural integrity Lipid homeostasis contributes to two major physiological aspects of membranes: structural integrity and membrane fluidity. The introduction of cis double bonds to lipid acyl chains reduces the interactions between lipid molecules within membrane bilayers and results in reduced packing and increased membrane fluidity. Membrane fluidity is regulated by cells in response to temperature

and solvent exposure. Cells with defects in membrane fluidity are sensitive to changes in temperature and solvent. Lipid unsaturation was recently found to regulate cell adhesion and flocculation as mutants having reduced lipid unsaturation highly induced expression of FLO1, a facilitator of cell wall fusion (Degrief et al., 2017). Cells with defects in membrane structural integrity often exhibit endocytic defects leading to alterations in cell polarity alongside sensitivity to detergents, salts and drugs—phenotypes examined in Chapter 4.

1.3.4 Yeast growth is supported by fermentation and respiration The appearance of substantial concentrations of molecular oxygen on the earth provided an abundant and advantageous terminal electron acceptor due to its high redox potential, permitting evolution of respiratory metabolic pathways (Castresana and Saraste, 1995; Walker, 1978). The budding yeast, Saccharomyces cerevisiae, can both ferment and respire. During aerobic exponential growth in batch culture, strains belonging to the Saccharomyces genus are efficient fermenters of glucose to the mixed products acetate, glycerol and ethanol via glycolysis and reductive metabolism. After glucose is exhausted, acetate and ethanol are oxidized by aerobic respiration. Reflecting the metabolism of ancestral organisms, Saccharomyces is also well adapted to anaerobiosis by forming ATP via glycolysis (reviewed in Fraenkel, 2011).

S. cerevisiae grows most rapidly when glucose is present as a carbon source, however once glucose is exhausted, yeast grow slowly by reducing ethanol to form acetate and undergo gluconeogenesis, where CO2 is the main by-product formed. During fermentation, carbon is both assimilated into metabolites required for growth and repair and released as ethanol. As fermentation is famously inefficient for releasing relatively high energy by-products, a cost apparently defrayed by rapid ATP formation (reviewed in Fraenkel, 2011). Significantly more ATP is produced when all pyruvate is oxidized through the tricarboxylic acid (TCA) cycle releasing CO2, however for yeast growing on glucose the amount of ATP produced through respiration is minimal, even in abundant molecular oxygen (Swanson and Clifton, 1948). Alternately, when mannose or galactose are the primary carbon and energy source, respiratory enzymes are de-repressed and fermentation and respiration proceed simultaneously. These phenomena are known as Crabtree effects, which describe the logic of the preference for fermentation over respiration for a given carbon source during abundance of molecular oxygen. The Pasteur effect describes the increase in cell mass achieved by yeast cultures grown in the presence of oxygen due to the efficiency of ATP per mole of glucose generated via respiration over anaerobic fermentation (de Deken, 1966). Crabtree effects are also known as “contre-effet Pasteur” where in the presence of high concentrations of glucose and oxygen 24

respiration is mainly repressed. Diauxic shift is the transition between glucose fermentation to aerobic respiration where ethanol is the carbon source after glucose exhaustion (reviewed in Fraenkel, 2011).

Reducing equivalents of NADH are vital to glucose oxidation and energy absorption in both fermentation and respiration. NAD+ is the major oxidative cofactor in the cell and is present in millimolar concentrations (Bennett et al., 2009). During fermentative growth, NADH is re-oxidized to NAD+ by pyruvate decarboxylation to CO2 and acetaldehyde and the ensuing reduction to ethanol by alcohol dehydrogenase. During aerobic respiration, NADH is oxidized by the electron transport chain so that NAD+ is reformed and electrons are used to drive oxidative phosphorylation of ATP (reviewed in Fraenkel, 2011). NADPH is NADH phosphorylated on the 2’hydroxyl group, forms the major reductive cofactor in the cell which is produced by the oxidative branch of the pentose phosphate pathway. Having two pools of redox cofactors allows for differential positioning toward NADPH preferred anabolic and NAD+ preferred catabolic reactions (reviewed Fraenkel, 2011). Over 200 metabolic reactions in yeast require NAD(P)H as a cofactor and roughly one sixth of all reactions listed in the BRaunshweig ENzyme DAtabase (BRENDA) are NAD(P)H dependent (Förster et al., 2003; Schomburg et al., 2002).

1.3.5 Compartmentalization of metabolism in yeast The cytoplasmic milieu is the major nexus for influxed nutrients and transformative enzymes. Metabolism is not restricted to the cytoplasm as specialized pathways are partitioned within yeast’s fifteen compartments including membrane compartments and many reactions catalyzed by isozymes occur simultaneously across several organelles (Herrgård et al., 2008). In budding yeast, the cytoplasm comprises 50% of cell volume and the vacuole is the next largest organelle functioning in provision of a large sink for free amino acids and other storage reserves and is also a major site of catabolism. Proteins destined for membranes are translated on membrane bound ribosomes of the endoplasmic reticulum and are trafficked to their final destinations within vesicles. Membrane bound vesicles comprise the major site of lipid synthesis and are inherently formed upon fragmentation of phospholipid membranes (Paltauf et al., 1992). Respiration occurs within the cell’s mitochondria. When cells are grown on glucose, mitochondria make up 3-5% of cell volume and are increased to 10% of cell volume on glycerol (Stevens, 1977). When fatty acids are used as a carbon source, the peroxisomal β-oxidation pathway is used to produce acetyl-CoA and NADH, in contrast to higher eukaryotes which employ mitochondrial β-oxidation (Hiltunen et al., 2003).

1.3.6 Compartmental proteomes are distinct Evolution of organelles can occur through autogenous subdivisions of existing compartments or through xenogenous incorporation of a parallel compartment from another organism following endosymbiosis—the accepted model for mitochondria and plastids (Wallin, 1927; Gray et al., 1999). These organelles contain genomes that encode a small portion of the resident proteome, while the majority of the proteins are nuclear encoded. Although peroxisomes do not encode their own genomes, there is an astonishing amount of overlap for subcellular proteomes where 64% of peroxisomal proteins have homologs expressed in the mitochondria and 50% have homologs expressed in the nucleus (Gabaldón and Pittis, 2015). Localizations of proteins are not fixed as at least 32% of eukaryotic protein families contain homologs that are expressed in different subcellular compartments emphasizing retargeting of localization throughout evolutionary history (Gabaldón and Pittis, 2015).

Throughout Chapter 4, I will discuss how loss of function in Tda5 leads to perturbed mitochondrial metabolism. The phenotypes involve significant changes in metabolite levels within differentially compartmentalized portions of metabolic pathways including the tricarboxylic acid (TCA) cycle, N- acetyl-glutamate cycle of arginine biosynthesis, lysine biosynthesis and branched chain amino acid biosynthesis. As reference, the metabolic pathways are described below.

1.3.6.1 Compartmentalization of Respiration The oxidative nature of respiratory reactions as well as maintenance of a stable proton gradient for driving ATP generation requires sequestration of these reactions within the double membrane bound mitochondrion. Respiration involves oxidation of carbon sources by the electron transport chain where electrons are carried through reducing equivalents of NADH and FADH2. The mitochondrial outer membrane is permeable to both NAD+ and NADH. NADH also forms two distinct pools where cytoplasmic NADH is respired in the mitochondrion by Nde1 and Nde2 and mitochondrial NADH by Ndi1(reviewed Fraenkel, 2011). Transport of pyruvate derived from glycolysis into the mitochondrion is facilitated by the recently discovered mitochondrial pyruvate carrier Mpc1/Mpc2 on glucose and Mpc1/Mpc3 on lactate (Herzig et al., 2012). Pyruvate and acetyl-CoA also form two distinct pools where pyruvate derived acetate is used to form acetyl-CoA via acetyl-CoA synthetase in the cytoplasm and nucleus where these pools are required for fatty acid biosynthesis and genome acetylation, respectively (Van den Berg and Steensma, 1995). The matrix localized pyruvate dehydrogenase is needed for production of acetyl-CoA in the mitochondrion where it is oxidized to oxaloacetate in the TCA cycle. Several TCA intermediates 26

are exchanged with the cytoplasm, as fumarate and succinate are exchanged by the Sfc1 transporter while the malate dehydrogenase (Mdh1) and aspartate aminotransferase (Aat1) form the malate- aspartate shuttle in yeast (Palmieri et al., 2000; Bakker et al., 2001).

1.3.6.2 Arginine biosynthesis A classic example of biosynthetic pathways spanning multiple milieus is arginine biosynthesis, a biosynthetic pathway that will be discussed in Chapter 4. The pathway to arginine formation begins with glutamate in the matrix of the mitochondrion and is called the N-acetyl-glutamate cycle. Glutamate is channelled into the mitochondrion by the aspartate/glutamate transporter Agc1 where it is N-acylated by Arg7 using acetyl CoA (Jauniaux et al., 1978). Subsequent phosphorylation, reduction and amination reactions results in release of ornithine. Ornithine provides an important source of polyamines and is transported to the cytoplasm by the ornithine transporter (Arg11) (Palmieri et al., 2000). The biosynthesis of arginine in the Saccharomyces genera diverges here from higher eukaryotes where ornithine is retained in the mitochondrion. Production of arginine from ornithine occurs through citrulline and argininosuccinate intermediates. The amino acids ornithine, citrulline and arginine are each taken up by the vacuole (Jauniaux et al., 1978).

1.3.6.3 Lysine biosynthesis Unlike higher eukaryotes, fungi are lysine prototrophs. Lysine biosynthesis is unusual in yeasts as it is carried out in the mitochondrion by enzymes in the 2-amino-adipate pathway comprising reactions that parallel the TCA cycle (reviewed in Zabriskie and Jackson, 2000). Mitochondrial 2- ketoglutarate is acetylated to homocitrate by homocitrate synthase in an acetyl-CoA dependent reaction, and is isomerised to homoisocitrate by Lys4 and oxidized by Lys12 consuming NAD+ and

CO2 while releasing 2-ketoadipate in the last mitochondrial step. Transamination occurs in the cytoplasm followed by condensation of 2-amino-adipate semialdehyde with glutamate to produce saccharopine in the cytoplasm where it is oxidized to lysine (reviewed in Zabriskie and Jackson, 2000). Since humans are lysine auxotrophs while fungal pathogens are lysine prototrophs, the lysine biosynthetic pathway is a key target for anti-mycotic therapies as mutants in the 2-amino-adipate pathway exhibit attenuated virulence (Jastrzębowska and Gabriel, 2015).

1.3.6.4 Branched chain amino acid biosynthesis The pyruvate family of branched chain amino acids leucine, isoleucine and valine are essential amino acids for humans, while fungi are prototrophs for these and all other proteogenic amino acids. The biosynthesis of isoleucine and valine shares five enzymatic steps, where 2-ketobutyrate

is substituted for pyruvate in isoleucine biosynthesis, and 2-ketobutyrate additionally forms the substrate for threonine synthesis (Holmberg and Petersen, 1988). Valine begins with condensation of two molecules of the three carbon 2-keto-carboxylic acid pyruvate to form 2-acetolactate, followed by reduction to 2,3-dihodroxy-isovalerate, hydrolysis to 2-ketoisovalerate and transamination to valine. Leucine diverges at 2-ketoisovalerate, which is condensed with acetyl- CoA to form 2-isopropylmalate and transported to the cytosol via Oac1, isomerized to 3- isopropylmalate, oxidation to 2-keotisocaproate followed by transamination to leucine (Marobbio et al., 2008). Notably, isoleucine biosynthesis shares all enzymes with valine production, but instead begins with the 4 carbon 2-keto carboxylic acid 2-ketobutyrate, which is derived by deamination of threonine. All reactions of isoleucine and valine biosynthesis appear to be mitochondrial up to transamination and involve sharing of all enzymes, whereas synthesis from 2-isopropylmalate to leucine appears to be cytoplasmic and uses the Leu2 enzyme. The final transaminase reaction is carried out by Bat1, Bat2 enzymes which have mitochondrial and cytosolic localizations (Colón et al., 2011).

1.3.7 Project rationale: Elucidating the functions of Tda5 It is clear that short chain dehydrogenases provide essential roles in yeast metabolism, yet six of fifteen enzymes in this class have no assigned activity in yeast. Even the essential short chain dehydrogenase, Pbr1, has no assigned reaction. Genetic screening for metabolomic phenotypes for uncharacterized candidate enzymes revealed numerous phenotypes for the tda5Δ deletion mutant. In order to determine the contribution of Tda5 to metabolism, I aimed to determine its biochemical functions. Studies outlined in Chapter 4 will describe new aspects of Tda5 function alongside its paralog, Ydl114w, in their contributions to cytosolic and mitochondrial metabolism and mechanisms for compensation when these dehydrogenases are knocked out.

2 Methods 2.1 Methods relating to Chapter 3: Uncovering the function of Das2 2.1.1 Growth and maintenance of yeast cultures Yeast strains were streaked to YP-dextrose (YPD) unless a nutritional marker was used for plasmid selection, and incubated at 30 °C for two nights and stored for up to two weeks at 4 °C. All liquid culture ahead of assays was carried out in modified YNB-glucose to avoid sulfate suppression in negative mode LC-MS (Per litre: 0.67 g YNB salts w/o amino acids and ammonium sulfate, 20 g dextrose, 0.5 g ammonium sulfate, 2.62 g ammonium acetate). Transformations were carried out according to Gietz and Woods (Gietz and Woods, 2006).

2.1.2 Construction of Mutants FY4 is a MATa haploid prototrophic yeast isogenic to Saccharomyces cerevisiae BY4742 was used as wild-type across all studies. The das2Δ::kanMX, urk1Δ::kanMX strains were obtained from the prototrophic yeast deletion collection. Deletion alleles were amplified by PCR and transformed into the FY4/FY5 prototrophic diploid (Winston et al., 1995), sporulated and dissected to obtain newly prepared both MATa and MATα prototrophic haploid strains. As the fur1Δ deletion allele is not present in the prototrophic yeast deletion collection, I amplified the natMX selection cassette using primers homologous to the 5’ and 3’ UTRs (Longtine et al., 1998), transformed FY4/FY5 and selected fur1Δ::natMX/FUR1 diploids, sporulated and dissected to obtain both MATa and MATα fur1Δ::natMX prototrophic haploids. The urh1Δ::kanMX strain was obtained from the prototrophic yeast deletion collection. Double and triple mutants were prepared by crossing MATa and MATα strains, picking zygotes by micromanipulation on a yeast dissection microscope, sporulating diploids, dissection, phenotyping of markers followed by confirmation of deletion and wild- type alleles by PCR genotyping.

2.1.3 Preparation of point mutants by in vivo site directed mutagenesis Fragments of each gene were PCR amplified in two to three sections using oligonucleotide primers with 20 bp overlaps and used as substrates for Gibson assembly(Gibson et al., 2009). The full length Gibson product was amplified with inset primers and used as an integrative recombinant fragment for allele swap by delitto perfetto technique (Storici and Resnick, 2006). Haploid yeast landing pad strains were prepared by amplification of a Kluyveromyces lactis Ura3 linked to kanMX cassette counter-selectable reporter (CORE),) flanked with 40 bp homology to the DAS2 and URK1 genomic loci. CORE strains were grown overnight in SD-

URA, followed by transformation using the integrative recombinant fragment prepared above, and plating on YPD overnight at 30 °C. Lawns were replica plated to SC-ura+0.3% FOA (since das2 and urk1 strains are resistant to FOA) and incubated overnight at 30°C. For the third round of replica plating, strains were transferred to both SC-ura+0.3% FOA and YPD+G418 and incubated overnight at 30 °C. Colonies that were resistant to 0.3% FOA and sensitive to G418 were chosen for genotyping. Genotypes of resultant haploid point mutants were determined by Sanger sequencing. We verified presence of point mutations and that loci were otherwise wildtype sequence from 5’ to 3’UTRs. Haploid point mutant strains were crossed to various deletion mutant backgrounds, sporulated, dissected, phenotyped and genotyped by Sanger sequencing to confirm presence of deletion and point mutant alleles.

2.1.4 HA fusion tagged alleles The C-termini of Das2 and Urk1 were fused to Human influenza hemagglutinin (3XHA) tags by amplification of cassettes using custom primers with 40 bp homology to protein coding sequences and 3’UTR using HygMX selection marker (Goldstein and McCusker, 1999; Longtine et al., 1998). The 3XHA::HygMX cassettes were transformed into FY4 MATa prototrophic haploid, PCR amplified in a single piece and transformed into various pyrimidine salvage deletion mutant backgrounds. The function of the DAS2-3XHA::HygMX and URK1-3XHA::HygMX alleles were determined by measuring resistance to 6-azauracil and by measuring uridine and uracil levels between tagged and untagged strains.

2.1.5 Heterologous expression of enzymes DAS2 and URK1 were cloned using Gateway site-specific recombination (ThermoFisher Scientific) into pDEST17 for N-terminal 6x-histidine tagging alongside high copy inducible expression and transformed into the BL21(DE3)pLysS Escherichia coli strain. Strains were pre-grown in MGD non-inducing media supplemented with ampicillin and chloramphenicol (Amp/Chlor) and auto-induced at room temperature in MDA-5052 (Amp/Chlor) media (Studier, 2005). Once optical density had plateaued, 500 mL cells was harvested by centrifugation at 4000 rpm in a SLC4000 rotor, washed in PBS and frozen at -80 °C. Cells were thawed on ice and resuspended in 50 mM phosphate buffer pH 7.5, 5mM imidazole with PMSF, dispersed by sonication for 10 minutes, and lysis carried out by nitrogen decompression on ice (Simpson, 2010). Cellular debris was separated from soluble proteins by centrifugation of homogenates for 15 minutes at 12,000 rpm in a SW-28 rotor. Supernatants were filtered using 0.22 µM filters and filtrate was loaded into a sample

column. 6x-His tagged proteins were purified using an AKTA purifier system FPLC equipped with TALON metal affinity resin column. For 6xHis-Das2, fractions containing protein of interest were further purified by ion exchange against NaCl in 20 mM Phosphate pH 7.5 using MonoQ 10/100 resin and stored at -20 °C in 50% glycerol. 6xHis-Urk1 was stored in DMSO and MgCl2 according to (Natalini et al., 1979). Fur1 was cloned into NdeI/XhoI sites of the pET24a (+) plasmid (Novagen) which provides a tag on the C terminus and purified using methods above. For dual expression of 6xHis-Das2 was cloned into the BamHI/PstI sites and Urk1 was cloned with stop codon intact into the NdeI/XhoI sites. 6xHis-Das2 pull-down of Urk1 was carried according to conditions for 6xHis-Urk1.

2.1.6 Preparation of Das2 G17E The phosphate binding loop was modified for Das2 to prevent uridine binding and prepared by Quikchange site-directed mutagenesis method (Agilent).

2.1.7 Radiometric enzyme assays Radioisotopes were purchased from Moravek Inc. 3H-Uridine kinase assays were carried out as previously described, except that GTP was used as a phosphate donor, and samples were removed at 0, 12 and 24 minutes at 24 °C (Cheng et al., 1986). 3H-UPRTase assays were carried out as previously described, except that 200 µM uracil and 1 mM PRPP was used, and samples were removed at 0, 12 and 24 minutes at 24 °C (McIvor et al., 1983). Reactions were quenched in 40 µL 1M urea and spotted on DEAE-cellulose ion exchange filters and dried (Perkin-Elmer, 1450-522). For washes, six filters at a time were washed in 50 mL 10 mM Tris pH 7.4 for 2 minutes of agitation, for three cycles. Filters were eluted in 0.1N HCl 1M NaCl in plastic 20 mL scintillation vials, 9 mL scintillation fluid was added, mixed and left to stand overnight before activity was measured by scintillation counting. Assays using purified protein contained 0.5 mg/mL. For assays containing homogenates, 500 mL of yeast cultures at an optical density of 1 grown in YNB-glucose were harvested by centrifugation, washed in 100 mM Tris, concentrated in 2 mL screw top vials, 200 µL of zirconium beads were added and volume adjusted to 2.0 mL. Homogenates were prepared by bead-beating in a Geno- Grinder 2000 at 1700 strokes per minute for 40 minutes. Homogenates were separated by centrifugation at 20,000 x g for 30 minutes at 4 °C. The top 300 µL was removed, mixed and total protein content was measured by Micro BCA kit (ThermoFisher Scientific).

2.1.8 Pyrimidine analogue resistance assays Cultures were grown overnight in YNB-glucose and culture density was adjusted to 0.2 OD/mL in YNB-glucose. Inhibitors were prepared at 2X concentration in YNB-glucose. 100 µL cells were added to replicate 96-well optically clear plates, and 100 µL diluted inhibitor added on top. Plates were sealed with breathable fabric seals and incubated at room temperature for 20 hours. Optical density at 600 nm was measured using a Perkin-Elmer microplate-reader.

2.1.9 Metabolite extracts Cells were grown over 3 rounds of sub-culture in YNB-glucose at room temperature to mid- log phase, metabolites extracted, separated and detected by LC-MS (Costanzo et al., 2016; Rosebrock and Caudy, 2017). Chromatographic peaks were extracted and integrated using a custom R package, ChromXtractorPro.

2.1.10 In vivo tracking of uridine kinase activity Pyrimidine salvage mutants were grown over 3 rounds of sub-culture to mid-log phase in 100 mL YNB-glucose at room temperature. With continuous shaking, 6-azauridine was added to a final concentration of 100 µg/mL. and replicate metabolite extracts were prepared over time. 6-aza-UMP was quantified by LC-MS.

2.1.11 Western Blot Strains containing the Das2-3XHA allele in the wild-type and pyrimidine salvage mutant contexts were grown in YNB-glucose through several rounds of growth and sub-culture and protein was extracted according to (Zhang et al., 2011) were extracted at an OD600 of ~0.5. Protein homogenates were separated under denaturing conditions on a 12.5% poly- acrylamide gel. Proteins were transferred to PVDF by electrophoretic transfer for 2 hours at 70 V at 4 °C, blocked overnight in Millipore FL-Blok fluorescent block buffer. Primary hybridization was carried out overnight in Blok with anti-HA 3F10 (Roche) at 1:200, 100 µL 10% azide, 100 µL 25% Tween-20. Membranes were washed 5X in 100 mL TBST, hybridized with anti-tubulin 1:2500 Abcam EPR13799 anti-α-tubulin, 100 µL 10% azide, 100 µL 25% Tween-20 1 hour at room temperature, followed by washing 5 times in 100 mL TBST. For Das2-3XHA and tubulin detection, hybridization of secondary antibodies was carried out for one hour at room temperature with the following antibodies and reagents: Li- COR IRDye® 680RD Goat anti-Rat IgG at 1:1000, IRDye® 800RD Donkey anti-Rabbit IgG at 1:3300 in 10 mL Blok with 10 µL 10% SDS, 100 µL azide, 100 µL Tween-20. Secondary

antibody hybridized blots were washed 5 times in 100 mL TBST. Images were captured on a LI-COR Odyssey Imaging System 10-minute exposure at 700 nm and 2-minute exposure at 800 nm. Signal was quantified in ImageStudio Lite Version 5.2 and linear regression was applied to Das2-3XHA dilution curves to determine the relationship between signal and input. Urk1-3XHA levels were measured as above, except, a 10% polyacrylamide gel was used, secondary hybridization with Li-COR IRDye® 800 RD Goat anti-Rat IgG at 1:5000, IRDye® 680RD Donkey anti-Rabbit IgG at 1:3300.

2.1.12 Preparation of Ribose-1-phosphate Preparation of ribose-1-phosphate was carried out by phosphorolytic cleavage of inosine with reaction scaled to 53 mL (Klenow and Emberland, 1955). Reaction buffer contained 0.1 M Tris-Formic acid buffer, pH 7.6, 7.5 units of nucleoside phosphorylase, 10 units of xanthine oxidase, 20 units of catalase and 1430 uM inosine. Reaction was initiated by addition of 1800 uM phosphate buffer, pH 7.4 and rocked for 6 hours at room temperature, and uric acid precipitate was enhanced by storage at 4 °C. I developed a flash chromatography method to clean up ribose-1-phosphate from reaction components using an Agela amino bonded silica column (CN140012-0) eluted under the following HILIC chromatographic condition: 70 mM ammonium acetate, 44 mM ammonium hydroxide in 30% ethanol. The IntelliFlash 310 system was run at 30 mL/min using 34 5 mL fractions of buffer. Elution of ribose-1- phosphate was monitored by LC-MS. Ribose-1-phosphate eluted in fractions 17 to 34. Ribose-1-phosphate containing fractions were dried overnight under nitrogen flow at 5 PSI.

2.2 Methods for Chapter 4: Phenotyping and mapping suppressors of Tda5 2.2.1 Growth and maintenance of yeast cultures As above, the Saccharomyces cerevisiae FY4 MATa prototroph was used as wild-type across all studies. All strains were grown as listed above and appropriately supplemented to support growth of auxotrophs according to (Amberg et al., 2005).

2.2.2 Construction of Mutants The tda5Δ::kanMX, strain was obtained from the prototrophic yeast deletion collection. Deletion alleles were amplified by PCR and transformed into the FY4/FY5 prototrophic diploid, sporulated and dissected to obtain newly prepared both MATa and MATα prototrophic haploid strains. I aimed to generate a tda5Δ strain that was compatible with SGA

and complementation via URA3 and HIS3 marked plasmids. I PCR amplified fragments of the tda5Δ::HygMX, deletion allele and transformed into a MATa/MATα diploid, sporulated and dissected to obtain a tda5Δ::HygMX MATα ura3Δ his3Δ can1Δ LYP1 isolate. The ydl114wΔ::kanMX strain was obtained from the auxotrophic yeast deletion collection, marker switched to ClonNat resistance, transformed into the FY4/FY5 diploid, sporulated and dissected. The tda5Δ ydl114wΔ double deletion strain was generated by crossing tda5Δ::kanMX to ydl114wΔ::natMX, sporulated, dissected and phenotyped.

In selecting second site suppressors of tda5Δ, the freshly dissected tda5Δ::HygMX MATα ura3Δ his3Δ can1Δ LYP1 isolate was streaked onto YP-glycerol and incubated at 30 °C and incubated 4 nights. For determination of whether strains were allelic to hst1Δ, sum1Δ or rfm1Δ, suppressors were crossed to hst1Δ::kanMX MATa tda5Δ::kanMX can1Δ::STE2pr_Schz.pombeHIS5 his3Δ TDH2-tagRFP::natMX, and isogenic sum1Δ and rfm1Δ strains, diploids selected by streak purification on YPD+ClonNAT+hygromycin and tested for growth on respiratory carbon sources. For generating pools ahead of sequencing, suppressors were crossed to ydl114wΔ::natMX MATa, sporulated, dissected, phenotyped and phenotyped for small, ClonNAT and hygromycin resistant colonies. Six to 12 colonies for suppressed and unsuppressed groups were grown overnight in YPD and pooled according to equal numbers of cells, genomic DNA prepared.

The YDL114W glucose repressible allele was generated by Gateway site specific recombination into pYES-DEST52 (ThermoFisher Scientific) containing a high copy 2µ origin, URA3 selection, and YDL114W containing a stop codon under the control of the GAL1 promoter. The pYES-DEST52-YDL114W plasmid was transformed into the tda5Δ::kanMX/TDA5 ydl114wΔ::natMX /YDL114W ura3/ura3 his3/his3 diploid, sporulated in media lacking uracil supplemented with histidine, dissected on YP-galactose, and phenotyped to obtain pYES-DEST52-YDL114W in the tda5Δ and TDA5 backgrounds. This method was carried out for pYES-DEST52-TDA5 in parallel, however, the YDL114W repressible allele gave an immediate decrease in growth rate when switched to glucose, which may indicate a less stable protein, consistent with its higher instability index.

The pAG426GPD-TDA5-dsRED strain for determination of localization of Tda5 was prepared by Gateway site specific recombination of TDA5 lacking a stop codon ingot the pAG426GPD-dsRED vector (Alberti et al., 2007) and transformed into a his3/his3 diploid strain.

2.2.3 Spot assays Wild-type and mutant cultures were grown in YNB-glucose overnight, OD was adjusted to 1.0 and 10-fold serial dilutions were prepared in water. A multi-channel pipette was used to deliver 3 µL to a dry YPD plates with supplement or inhibitor as indicated. Plates were imaged every 24 hours up to 72 hours on an AlphaImager gel documentation system using white light transillumination.

2.2.4 Metabolite extracts Metabolite extracts for tda5Δ mutants were prepared according to standard procedures listed above. Metabolite extracts analyzed for TCA cycle intermediates were analyzed by LC-MS as previously described (Soloveychik et al., 2016).

2.2.5 Purification of short and medium chain acyl-CoA metabolites Strains were grown in YNB-glucose+histidine over several rounds of pre-growth and sub- culture until an OD of 1.0 was reached. 100 mL cultures were harvested by centrifugation in SLA-600TC rotor at 1000 rpm for 5 minutes, supernatants decanted, to which 1.2 mL ACN/IPA (3:1, v/v) extraction solvent was added and samples were freeze-thawed for three cycles. Solid phase extraction columns containing 2-(2-Pyridyl)ethyl silica gel (Supelco) were used to purify short to long chain acyl-CoA metabolites as previously described (Snyder et al., 2015) with the omission of phosphate, as phosphate addition suppressed metabolites up to three rounds of chromatography after addition. Preparation of 15N Acyl-CoA extract was prepared as above, except that 15N Ammonium Sulfate was substituted instead of 14N ammonium acetate and 14N ammonium sulfate. Samples were resuspended in 75 µL 15 ACN:H2O 3:7 per 100 OD of cells and N extract added ahead of LC-MS. LC-MS was carried out as previously described, except a steeper gradient was used so that CoA metabolites were not obscured by two successive broad suppressive peaks having an m/z of 159.1027.

2.3 Strains and Plasmids used in this study Table 1. Plasmids created for this study

Plasmid Description pJH0010 pDONR221-URK1-Stop pJH0012 pDONR221-DAS2-Stop pJH0016 pDONR221-TDA5-No_stop pJH0020 pDEST17-URK1-Stop pJH0022 pDEST17-DAS2-Stop

pJH0044 pDEST17-YDL114W-Stop pJH0045 pVV214-HygMX-ura3-DAS2-Stop pJH0057 pET-DUET-6xHis-DAS2-Stop (Pst1BamHI) URK1-Stop (Nde1/EcoRV) pJH0079 pDEST17-das2G17E-Stop pJH0111 pET24a-FUR1-No_stop-6xHis (NdeI/XhoI) pJH0150 pDONR221-TDA5-Stop pJH0153 pDONR221-YDL114W-No_stop pJH0160 pAG426-GPDpr-TDA5-No_stop-dsRED pJH0161 pYES-DEST52-YDL114W-Stop pJH0162 pYES-DEST52-TDA5-Stop

Table 2. Yeasts strains used in this study

Strain Genotype Source JH0013 FY4/FY5 MATa/MATα (Winston, 1995) JH0014 FY4 MATa (Winston, 1995) JH0045 erg12 G392D ura3Δ MATa Hanchard, 2015 JH0059 RCY604 ura3Δ MATα Amy Caudy JH0108 urk1Δ::kanMX MATa This study. JH0109 urk1Δ::kanMX, MATα This study. JH0110 fur1Δ::natMX MATa This study. JH0111 fur1Δ::natMX, MATα This study. JH0112 das2∆::kanMX MATα This study. JH0113 das2∆::kanMX MATa This study. JH0114 urh1∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014) JH0115 urh1∆::kanMX das2∆::kanMX MATa This study. JH0116 urh1∆::kanMX das2∆::kanMX MATα This study. JH0117 urh1∆::kanMX urk1∆::kanMX MATa This study. JH0118 urk1∆::kanMX fur1∆::natMX MATα This study. JH0119 urk1∆::kanMX fur1∆::natMX MATa This study. JH0120 das2∆::kanMX fur1∆::natMX MATα This study. JH0121 das2∆::kanMX fur1∆::natMX MATa This study. JH0122 urh1∆::kanMX ura3∆::kanMX MATα This study. JH0124 tda5∆::kanMX fresh 1D isolate MATa This study. JH0125 tda5∆::kanMX fresh 2C isolate MATa This study. JH0126 tda5∆::kanMX fresh 2D isolate MATa This study. JH0127 tda5∆::kanMX fresh 4D isolate MATa This study. JH0128 tda5∆::kanMX fresh 6C isolate MATa This study. JH0129 tda5∆::kanMX fresh 6D isolate MATa This study. JH0131 tda5∆::kanMX suppressor 1D MATa This study. JH0132 tda5∆::kanMX suppressor 2C MATa This study. JH0133 tda5∆::kanMX suppressor 2D MATa This study. JH0137 tda5∆::kanMX suppressor 4D MATa This study. JH0140 tda5∆::kanMX suppressor 6C MATa This study. JH0141 tda5∆::kanMX suppressor 6D MATa This study. JH0142 urk1∆::kanMX das2∆::kanMX MATa This study. JH0143 urk1∆::kanMX das2∆::kanMX MATα This study. JH0144 fur4∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014) JH0145 fui1∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014) JH0146 hst1∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014)

JH0177 rfm1∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014) JH0178 sum1∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014) JH0179 ydl114w∆::kanMX ura3Δ met15Δ leu2Δ his3Δ1 (Winzeler, 1999; Giaever, can1Δ::STE2pr_Schz.pombeHIS5 lyp1Δ MATa 2002) JH0182 tda5∆::kanMX hst1 D406V MATa This study. JH0217 urh1∆::kanMX fur1∆::natMX MATa This study. JH0218 urh1∆::kanMX fur1∆::natMX MATα This study. JH0219 urh1∆::kanMX das2∆::kanMX urk1∆::kanMX MATa This study. JH0220 urh1∆::kanMX das2∆::kanMX fur1∆::natMX This study. can1∆::STE2pr_Schz.pombe_HIS5 MATa JH0241 ura3∆/URA3 fur1∆::natMX/FUR1 MATa/MATα This study. JH0254 tda5∆::kanMX ydl114w∆::natMX MATα This study. JH0252 tda5∆::hygMX ura3∆ MATα This study. JH0253 tda5∆::hygMX ura3∆ MATa This study. JH0256 ydl114w∆::natMX MATα This study. JH0257 ydl114w∆::natMX MATa This study. JH0282 sum1∆::kanMX tda5∆::kanMX TDH2-TagRFP::natMX This study. MATa JH0283 rfm1Δ::kanMX tda5∆::kanMX TDH2-TagRFP::natMX This study. MATa JH0284 hst1∆::kanMX tda5∆::kanMX TDH2-TagRFP::natMX This study. MATa JH0293 das2∆::kanMX fur1∆::natMX urk1∆::kanMX MATa This study. JH0335 upc2∆::kanMX his3Δ1 can1Δ::STE2pr_Schz.pombeHIS5 (Gibney, 2013; van der Sluis, lyp1Δ MATa 2014) JH0337 das2Δukase::KlURA3-kanMX ura3∆ This study. JH0338 urk1Δukase::KlURA3-kanMX ura3∆ This study. JH0339 urk1Δuprtase::KlURA3-kanMX ura3∆ This study. JH0340 das2 G17E ura3∆ MATa This study. JH0342 urk1 G424E ura3∆ MATa This study. JH0343 urk1 G424E ura3∆ MATα This study. JH0364 das2 G17E URA3 MATα This study. JH0365 das2 G17E URA3 MATa This study. JH0366 urk1 G424E URA3 MATa This study. JH0368 urk1 G63E URA3 MATα This study. JH0369 urk1 G63E URA3 MATa This study. JH0370 das2 G17E urh1Δ::kanMX MATa This study. JH0372 urk1 G63E fur1∆::natMX MATa This study. JH0380 urk1 g63e urh1∆::kanMX MATa This study. JH0381 urk1 g63e urh1∆::kanMX MATα This study. JH0382 urk1 UPRTase fur1::natMX MATα This study.

JH0383 urk1 UPRTase fur1::natMX MATa This study. JH0388 TDA5 erg12 G392D This study. JH0389 tda5∆::HygMX erg12 G392D This study. JH0390 tda5∆::HygMX erg12 G392D This study. JH0391 TDA5 erg12 G392D This study. JH0392 TDA5 YDL114W upc2 R855L This study. JH0393 TDA5 YDL114W upc2 R855L This study. JH0394 tda5∆::HygMX ydl114w∆::natMX This study. JH0395 tda5∆::HygMX ydl114w∆::natMX upc2 This study. JH0396 tda5∆::HygMX ydl114w∆::natMX erg11 upc2 This study. JH0397 TDA5 YDL114W erg11 S382F This study. JH0398 TDA5 YDL114W erg11 S382F This study. JH0399 tda5∆::HygMX ydl114w∆::natMX erg11 S382F This study. JH0400 urk1 G63E fur1∆::natMX MATa This study. JH0401 urk1 G63E fur1∆::natMX MATα This study. JH0402 das2 G17E fur1∆::natMX MATa This study. JH0403 das2 G17E fur1∆:: MATα This study. JH0405 tda5∆::kanMX/TDA5 ydl114w::natMX/YDL114W This study. ura3∆/ura3∆ his3Δ/his3Δ MATa/MATα JH0437 TDA5 his3Δ ura3Δ pYES-DEST52-YDL114W This study. JH0438 tda5Δ::kanMX his3Δ ura3Δ pYES-DEST52-YDL114W This study. JH0436 TDA5 his3Δ ura3Δ pAG426GPD-TDA5-dsRED This study. JH0409 URK1-3XHA::HygMX MATa This study. JH0412 URK1-3XHA::HygMX das2Δ::kanMX MATa This study. JH0423 URK1-3XHA::HygMX fur1Δ::natMX MATa This study. JH0420 URK1-3XHA::HygMX das2 G17E MATa This study. JH0422 urk1G63E-3XHA::HygMX MATa This study. JH0421 urk1G424E-3XHA::HygMX MATa This study. JH0410 DAS2-3XHA::HygMX MATa This study. JH0417 DAS2-3XHA::HygMX urkΔ::kanMX MATa This study. JH0442 DAS2-3XHA::HygMX fur1Δ::natMX MATa This study. JH0414 das2G17E-3XHA::HygMX MATa This study. JH0416 DAS2-3XHA::HygMX urk1 G63E MATa This study. JH0415 DAS2-3XHA::HygMX urk1 G424E MATa This study. JH0439 tda5Δ::hygMX ura3Δ his3Δ This study. can1Δ::STE2pr_Schz.pombeHIS5 erg11 S382F MATα JH0440 tda5Δ::hygMX ura3Δ his3Δ This study. can1Δ::STE2pr_Schz.pombeHIS5 upc2 R855L MATα JH0441 tda5Δ::kanMX ydl114wΔ::natMX erg1 Δ1469 MATa This study. JH0439 tda5Δ::kanMX/TDA5 ydl114wΔ::natMX/YDL114W This study. pVV214-hyg-YDL114W

3 Discovery of Das2 an Novel Uridine Kinase and Obligate Member of a Novel UMP Salvage Complex 3.1 Rationale and summary In line with a role in catalyzing an enzymatic reaction using uridine as a substrate, I observed uridine accumulation in metabolite extracts derived from the das2Δ candidate enzyme mutant. Therefore, I aimed to determine the enzymatic function of Das2 alongside its contribution to pyrimidine salvage relative to known pyrimidine salvage pathways and determine the architecture of the pyrimidine salvage pathways.

My experiments show that Das2 is the major uridine kinase in yeast, and this novel enzyme depends on the bifunctional uridine kinase/UPRTase Urk1 for stability. Das2 and Urk1, in turn, depend on the major UPRTase Fur1 for stability, and together Das2, Urk1 and Fur1 form a pyrimidine salvage complex. For yeast grown in minimal media, cellular pyrimidine pools are generated by RNA catabolism and Das2 uridine kinase activity provides the major route of re-activation of uridine to UMP.

3.2 Motivational phenotypes for study of Das2 3.2.1 Metabolomic phenotyping reveals das2Δ accumulates uridine In order to systematically assign metabolic functions to uncharacterized enzymes, I screened 120 candidate gene deletion mutants for metabolomic phenotypes. As shown in Figure 4a across ~3000 mass spectral features measured in wild-type and das2Δ mutant, uridine is the most significantly changed metabolite as shown in Figure 4b, a 20-fold increase in uridine levels over wildtype was identified, where the difference in the means carried a Student’s t- test p-value of 1.9 x 10-5. When retention times, shifts in 13C and 15N labelling data were assessed, it was determined that other significantly upregulated mass spectral features were fragments of uridine and ions that change in response to uridine accumulation. This specific uridine accumulation phenotype motivated the exploration into the function of DAS2. The protein encoded by DAS2 contains a predicted uridine kinase domain, belonging to the enzyme commission assignment 2.7.1.48, which is widely distributed among Bacteria and Eukarya and only narrowly distributed in Archaea (Armenta-Medina et al., 2014). Among the three other known pyrimidine salvage activities, uridine kinases are more widely distributed than uridine phosphorylases, however are more narrowly distributed than uracil phosphoribosyl transferases (Armenta-Medina et al., 2014). Hitherto, no activity has been reported for the Das2 predicted enzyme. Furthermore, no studies have implicated the involvement of the DAS2 gene in the UMP salvage pathway. The function of Das2 has remained elusive due to the widespread adoption of auxotrophic markers as genetic tools for positive and negative selection, especially ura3Δ in combination with 5-fluoro-orotic acid (5- FOA) (Boeke et al., 1984, 1987). The use of uracil supplementation to support growth of ura3Δ mutants masks the appearance of phenotypes in uridine salvage and inhibited discovery of novel salvage components. My metabolomic screen for enzyme function was carried out on minimal media without exogenous supplementation. These conditions in combination with unbiased detection of phenotypes in metabolite levels led to identification of the change in uridine level.

As further discussed below, uridine kinases are critical to activation of pyrimidine analogues used as anti-cancer therapeutics in humans and, most relevantly, in treatment of fungal infection. As pyrimidine analogues represent an important class of anti-fungal therapeutics to which resistance rapidly evolves (Hospenthal and Bennett, 1998), it is of high priority to elucidate the role of the Das2 enzyme in its contribution to pyrimidine salvage.

***

Figure 4. Full-scan LC-MS metabolomic profiling reveals uridine accumulates in the das2Δ prototrophic deletion mutant A) Volcano plot for das2Δ metabolite levels relative to wild-type for ~3000 mass spectral features. Candidate enzyme deletion mutants and wild-type controls were grown in YNB-glucose through several rounds of growth and sub-culture and metabolites were extracted at an optical density of ~0.5. Extract supernatants were dried under N2, reconstituted in a volume of water according to optical density at time of extraction. Metabolites were separated by reverse-phase ion-paired chromatography and detected by Q-TOF. Untargeted analysis of chromatographic peaks was carried out in Agilent Profinder, significantly altered mass spectral features were determined using custom scripts R scripts. B) Dot and whisker plot showing 20-fold increase in uridine levels was observed for das2Δ mutant metabolite extracts relative to wild-type. Uridine was identified by m/z and chromatographic retention time matching a neat standard. Dots represent means of 6 independent biological replicates, error bars represent +/- standard error of the mean.

3.2.2 Literature studies implicate DAS2 in response to 5-FOA and 6-azauracil Although the enzymatic function of Das2 has not yet been determined, two studies have identified phenotypes involving the DAS2 gene thus far. The first study observed das2Δ partially suppresses dst1Δ sensitivity to low doses of 6-azauracil (Gómez-Herreros et al., 2012). The authors did not measure das2Δ mutant resistance on its own and my own studies below will show that resistance of das2Δ is not specific to the dst1Δ background. DST1 encodes the DNA strand transfer subunit of the transcription elongation factor TFIIS (Clark et al., 1991). Deletion mutants in DST1 are compromised in their ability to upregulate transcription of IMD2, encoding the inosine monophosphate dehydrogenase, in response to NTP depletion (Gómez-Herreros et al., 2012). As 6-azauracil causes NTP depletion, the combination of the dst1Δ mutation and 6-azauracil leads to growth inhibition. This study named DAS2 for Dst1 6-azauracil sensitivity and identified a role for Dst1 in transcription of rDNA during transcriptional stress. The phenotype of das2Δ conferring resistance to 6- azauracil suggests an involvement for Das2 in the UMP salvage pathway.

In a separate study by Hontz and colleagues, researchers aimed to identify factors necessary for rDNA transcription using a modified URA3 reporter driven by the TRP1 promoter in the non-transcribed spacer region 1 of the rDNA locus (Hontz et al., 2009). This screen involved selection of an insertion mutant library on various media including synthetic media with 0.2% 5-FOA added to select for mutants that silenced the URA3 reporter. Among inactivated genes that are identified for growth on 5-FOA are notable genes involved in pyrimidine metabolism: URA5, one of the orotate phosphoribosyl transferases, URA6 the uridine monophosphate kinase, FUR4 the uracil permease and YPR021C which is homologous to URC2 involved in uracil catabolism in the senso lato yeast, Saccharomyces kluyveri- the ability to catabolize uracil as a sole nitrogen source is an activity unseen in S. cerevisiae (Andersen et al., 2008). It is expected that ura5 mutants would have decreased UMP leading to reduced Ura3 steady state levels and reduced activity. Since Ura3 is the target of 5-FOA, mutations in URA5 would lead to increased growth on 5-FOA. Inactivation of FUR4 would preclude the entry of 5-FOA in the cell, likewise leading to resistance. Lesions in URA6 would lead to inability to activate UMP to UDP, leading to an increased pool of UMP rendering strains less sensitive to 5-FOA. It’s interesting that the Urc2 homologue was a hit in this screen as S. cerevisiae is unable to use uracil as a sole nitrogen source (Andersen et al., 2008). However, YDR520C inactivation results in increased growth on 5-FOA, which is consistent with its expression and a potential role in pyrimidine metabolism (Hontz et al.,

2009). Since this screen was heavily enriched for pyrimidine biosynthesis and uptake genes, I propose DAS2 was identified due to its role in affecting steady state levels of pyrimidines, and not for an activity related to rDNA transcription (Hontz et al., 2009). This, however, cannot be ruled out, as Das2 may be involved in a feedback response to selective degradation of ribosomes during autophagy, also known as ribophagy. In this model, loss of Das2 may reduce the rate of ribophagy, thereby stabilizing ribosomes and leading to decreased rDNA transcription to balance ribosome biogenesis and turnover. This would lead to increased growth on 5-FOA, consistent with the phenotype observed for das2 in the screen.

3.2.3 Deletion of DAS2 confers resistance to 6-azauracil The DAS2 gene is named for the ability of its deletion mutant to resist 6-azauracil (6-AU) in the dst1Δ background (Gómez-Herreros et al., 2012). Below, I will demonstrate that Das2 has uridine kinase activity suggesting that the resistance to 6-AU seen in the null mutant is due to diminished ability to activate 6-AU to 6-azauridine-monophosphate (6-AUMP). Salvage pathways for activation of 6-AU to 6-azaUMP are distinct to each organism (Brockman and Anderson, 1963). In humans 6-AU is activated first to the ribonucleoside via the action of a uridine phosphorylase, followed by activation from 6-azauridine (6-AUR) to 6-azaUMP by uridine kinases reaction although it is not activated to di- and triphosphate nucleotides. Notably, in human cells, hydrolysis of azauridine to azauracil does not occur. In contrast with the uridine salvage mechanism of activation, trypanosomes use a distinct route to activate 6AU by UPRTase directly to 6-AUMP (Brockman and Anderson, 1963). In yeast, 6-azauracil similarly is activated by UPRTase to 6-AUMP.

3.2.4 Mechanism of action for 6-azapyrimidines There are two mechanisms of action for 6-azapyrimidines, where the first relies on conformational and electrostatic mimicry to OMP. The 6-AUMP molecule contains a phosphate positioned 4.25 Å away from the phosphate on UMP, giving it distinct biological properties from UMP (Rada and Doskocil, 1980). Furthermore, at pH 7 the negatively charged triazine ring on 6-AUMP closely resembles the ionized carboxyl group on orotidine- 5-phosphate (OMP) (Rada and Doskocil, 1980). This conformation is not allowed for UMP, and thereby allows competitive inhibition of OMP decarboxylase by 6-AUMP in human cells at physiological pH (Rada and Doskocil, 1980). Secondly, 6-azauracil inhibits inosine monophosphate dehydrogenase, leading to 10-fold reduction in GTP pools and ensuing transcriptional stress (Exinger and Lacroute, 1992; Riles et al., 2004). Indeed, treatment with

5 uM 6-AU leads to a reduction in IMP dehydrogenase levels to 3.5% of wild-type and SchizoSaccharomyces pombe and Escherichia coli IMP dehydrogenases are likewise sensitive (Exinger and Lacroute, 1992).

3.2.5 Resistance to 6-azapyrimidines Resistance to 6-AU arises by several mechanisms and has been classically used to decipher primary pathways for activation of salvage pyrimidine metabolites to UMP and CMP. Lactobacillus bulgaricus has a natural resistance to 6-AU as the organism lacks UPRTase enzymes for conversion to 6-AUMP but does encode a uridine kinase. Likewise, Streptococcus faecalis mutants in uridine phosphorylase are resistant to 6-AU but not 6-AUR (Brockman and Anderson, 1963). In human cells, resistance to 6-AUR is attributed to loss of uridine kinase activity, as seen in L178Y lines. When resistance to 6-AUR is attributed to loss of uridine kinase activity, one can expect cross-resistance to fluorouridine and fluorocytidine (Sköld, 1960) as the enzyme is active on all three compounds. In treatment of humans with 6-AUR, a tolerance develops over time as the result of an increasing ability to metabolize orotic acid to UMP. In general, mechanisms of resistance to azapyrimidines are used to elucidate the pathways for pyrimidine activation. In this way, one can measure resistance to 6-AU and 6-AUR to determine topology of activation pathways in yeast and thus determine the contribution of various enzymes to these pathways by using deletion mutants.

Yeast are sensitive to 6-azauracil and resistance to azapyrimidines has been studied. Mutants lacking UPRTase or uracil permease are completely resistant to 6-azauracil (Loison et al., 1980). 6-AU resistance has been used to identify a regulator of de novo pyrimidine biosynthesis, PPR1 (Loison et al., 1980). Mutants in PPR1, named for pyrimidine pathway regulator, constitutively induce the expression of URA3 and URA1 in the absence of dihydroorotate (Loison et al., 1980). PPR1 encodes a zinc finger transcription factor (Todd and Andrianopoulos, 1997).

3.2.6 Mechanism of action of 5-fluoropyrimidines As for fluoropyrimidine antimetabolites, these molecular species are activated to FUMP and then to F-dUMP, which is a suicide inhibitor of dTMP synthetase (thymidylate synthase) (Heidelberger et al., 1957; Sköld, 1960). F-dUMP inhibition of thymidylate synthase blocks formation of dTMP, reducing the pool of TTP available for DNA synthesis. Thus, fluoropyrimidines effectively block DNA synthesis without blocking RNA and protein

synthesis. Initial work uncovering the mechanistic target of fluoropyrimidines was carried out by Sköld on cell free yeast extracts (Sköld, 1960). In addition to FdUMP, anti-mycotic action of 5-fluorocytosine also depends on production of 5-fluorouridine triphosphate, which is incorporated into RNA chains causing disruptions in protein synthesis (Polak, 1974). Fluoropyrimidines are suitable for use as anti-neoplastics and blocking adenovirus infection. Early work on 5-FU resistant mutants showed decreases in uridine kinase activity (Brockman and Anderson, 1963). Consequently, work with 5-FU resistant mutants has provided insight into pyrimidine metabolism.

3.2.7 Resistance to 5-fluoropyrimidines Resistance to 5-FU is accomplished by stimulation of de novo pyrimidine biosynthesis and by blocking incorporation of 5-FU into nascent RNAs (Carlsson et al., 2013). Classically, mutants blocked in uptake and salvage of uracil are resistant to 5-FU (Jund and Lacroute, 1970) and was used to identify FUR1 encoding the uracil phosphoribosyl transferase (UPRTase), FUR4 encoding the uracil permease, and FUR2 and FUR3 which have not been identified. The FUR1 gene was cloned by complementation of a ura5 fur1 leu4 strain growth on uracil (Kern et al., 1990a). The authors map the Fur1 point mutants that have diminished UPRTase activity and slightly lower (77-90%) uridine kinase activity. Based on these results, the author suggests a uridine kinase complex where the three Fur1 sites are residues that play a role in maintenance of uridine kinase activity. Interestingly, a subsequent study of 5- flurorocytidine resistant mutants found five complementation groups: FCY1, FCY2, FUR1, URK1 and URH1 while das2Δ mutants were not identified (Kurtz et al., 1999).

3.2.8 Das2 physically interacts with Urk1:implications for a pyrimidine salvage complex Beyond shared implications in pyrimidine salvage based on phenotypic studies, the physical interaction between Das2 and Urk1 is well documented through multiple genome-wide studies. One study overexpressed tagged Urk1 and identified an interaction with Das2 by affinity capture mass spectrometry (Breitkreutz et al., 2010), whereas three independent studies detected an interaction between this pair by yeast two-hybrid assay (Ito et al., 2001; Uetz et al., 2000; Yu et al., 2008). Remarkably, three of these studies also identified an interaction between the Urk1 and Fur1 proteins (Breitkreutz et al., 2010; Ito et al., 2001; Yu et al., 2008). Urk1 also interacts with itself by yeast two-hybrid (Ito et al., 2001; Yu et al., 2008). These data suggest that Urk1 participates in at least one pyrimidine salvage complex

with Das2 and Fur1 as members. The presence of a salvage complex would imply that mutations causing loss of function or destabilization of one component enzyme could lead to partial loss of a second activity. This is, indeed, the case as Grenson found that uridine kinase activity is controlled by two unlinked genes -uridk and ups, ups is allelic to FUR1 (Grenson, 1969; Jund and Lacroute, 1970). Furthermore, reduced uridine kinase activity is observed for UPRTase point mutants (Kern et al., 1991). Overall, these results indicate uridine kinase activity depends on an intact UPRTase.

3.2.9 Resolving the activities of Das2, Urk1 and Fur1 Grenson’s work provides foundational evidence for the existence of a complex of enzymes responsible for salvage that is further supported by modern proteomic data providing evidence for protein-protein interactions between Das2, Fur1 and Urk1. Uridine kinase activity was first reported in yeast by Grenson in 1969 (Grenson, 1969). In 1990, Kern determined that an URK1 clone restored uridine kinase activity in a deficient mutant (Kern, 1990). The nature of the uridine kinase mutation was not described, and no in vitro studies have been carried out on purified Urk1. As shown below, my finding of uridine kinase activity for Das2 is the first in vitro assignment of uridine kinase activity in yeast and the first direct association between a gene and uridine kinase activity.

The novel activity for Das2 was not mystifying because although uridine kinase activity had previously been annotated to Urk1 in yeast, as previously mentioned, no in vitro assays had been carried out verifying the catalytic reaction for Urk1. In fact, the DAS2 and URK1 genes both encode predicted uridine kinase domains, raising the question: which protein is more highly conserved among uridine kinases? As shown in Figure 5, Das2 and Urk1 both encode uridine kinase domains and, yet, there is no clear evidence as to whether both enzymes are functional, the nature of their interdependence (if any) or which one is carrying out the major role in activating uridine to UMP. Since pyrimidine salvage is critical to activation of antifungal drugs and crucial for maintaining nucleotide levels during stress, it is important to characterize the role of Das2 relative to other salvage enzymes and to determine its contribution to metabolic homeostasis.

aa 1 5 247 291 474 501

aa 1 33 179 232

Figure 5. Enzyme domain topologies for Urk1 and Das2 Protein sequences for Das2 and Urk1 were downloaded from SGD (Cherry et al., 1998) and domain boundaries were determined by PFAM (Punta et al., 2012).

3.2.10 Null mutations in fur1Δ are synthetic lethal with ura3Δ Cellular pyrimidine levels are maintained by a balance of UMP biosynthesis and salvage. When UMP biosynthesis is knocked out via mutation of ura3Δ, additional null mutations in fur1Δ are expected to cause severely compromised ability to salvage due to the dependence of uridine kinase activity(s) on the presence of Fur1. This is, in fact, the case as fur1Δ and ura3Δ mutations are synthetically lethal as the double mutants are inviable (Koren et al., 2003). However, a subset of mutations in the UPRTase enzyme do allow uridine kinase activity as Grenson found that when mutations causing 5-fluorouracil resistance were crossed to ura-2, growth of the double mutant was slow on uracil but not on uridine (Grenson, 1969). One reason why uridine kinase activity may be abolished in the absence of Fur1 is that the uridine kinase is unstable in the absence of Fur1. In line with this hypothesis, URK1 encodes six PEST sequence sites which can promote ubiquitylation and degradation when exposed (Rogers et al., 1986). PEST sites may be buried within the stable Urk1/Fur1 and Urk1/Das2 complexes and preclude surface exposure of the PEST domains, preventing degradation. From studies in mammalian systems, uridine kinases have been difficult to purify in vitro due to stability of sulfhydryl groups (Peck et al., 1971). Below, I present data consistent with the requirement for Urk1 association with Fur1 and Das2 for stability.

3.2.11 Genetic interactions point to a role for Das2 in pyrimidine salvage In addition to protein-protein interactions, DAS2 and URK1 share a common genetic interaction. On rich media, the das2Δ and urk1Δ deletion strains each have eqally strong negative genetic interactions with ura6 temperature sensitive mutants (Costanzo et al., 2016). In the study identifying this interaction, the strains are deleted for URA3, encoding the rate controlling step of de novo pyrimidine biosynthesis and therefore fully rely on salvage for uptake of pyrimidines required for growth. Strains that cannot efficiently salvage may be sensitized to blocking the formation of UDP from UMP by decreased function in the ura6

temperature sensitive mutant. This is likely a trigenic interaction between uridine kinase mutants, ura6 and ura3Δ. Likewise, the das2Δ deletion also has a negative genetic interaction with ura8Δ, encoding the minor CTP synthetase, again showing that efficient pyrimidine salvage is critical to strains compromised for de novo pyrimidine biosynthesis. It’s interesting to note that urk1Δ has a positive genetic interaction with dst1Δ, the DNA strand transferase protein noted for its role in maintaining transcription of rDNA genes (Clark et al., 1991), as Das2 is named for its positive genetic interaction in suppressing sensitivity to 6-azauracil in dst1Δ mutants (Gómez-Herreros et al., 2012). The shared positive genetic interaction between urk1Δ and das2Δ with dst1Δ may indicate a role for these proteins in balancing pyrimidine salvage and nucleoside availability for ribosome biogenesis. The das2Δ mutant has a weak genetic interaction with fcy21Δ, which encodes a purine/cytosine permease, named for fluorocytosine resistance (Paluszynski et al., 2006). Since FUR1 is essential in the ura3Δ background (Koren et al., 2003) the fur1 temperature sensitive allele was used in screening and as expected has negative interactions with ura4Δ and ura1Δ alleles, encoding lesions in the de novo UMP biosynthetic pathway.

3.2.12 Das2 and Urk1 topology By comparing the conservation of amino acid sequences between Das2 and orthologous uridine kinases, one can infer the amount of natural selection on each of the domains giving rise to a conserved structure and function. There are several crystal structures available for a single characterized human uridine kinase, UCK2 (Suzuki et al., 2004). I carried out multiple local sequence alignments to compare the conservation between Das2, Urk1, UCK2 and other uridine kinases spanning the Archaea, Bacteria and Eukarya kingdoms, we notice that Das2 stands out from other enzymes in its class because of its higher degree of non-synonymous amino acid substitution. As indicated by arrows in Figure 6, these substitutions are present even for highly conserved residues crucial to activities such as substrate binding. The topology of the Das2 protein begins with the phosphate donor binding domain (P-loop), which functions to attract the negatively charged phosphates on GTP or ATP and is rich in glycine, we see that the pattern of glycines is not conserved and the conserved lysine (position 112) that coordinates the gamma phosphate is substituted to valine. The active site aspartate (position 146) is conserved across all predicted uridine kinases as well as Das2, although the two conserved aromatic residues tyrosine and phenylalanine (positions 148, 149) are substituted to methionine and isoleucine. The histidine necessary for conferring uridine specificity over cytidine (position 201) is conserved from UDK to Das2 although the arginine two residues away is noticeably absent (position 203) (Tomoike et al., 2017). These changes may convey differences in substrate specificity between Das2 and other uridine kinases. When one examines a cladogram of the sequences representing the evolutionary relationship between Das2, Urk1 and other uridine kinases, one can see that Das2 has the longest branch length, indicating the highest amount of change. The cladogram also indicates single domain homologues cluster together in higher eukaryotes, double domain sequences also cluster together, while Urk1 and Das2 sequences cluster with archaeal and bacterial sequences. Although few sequences were used as input, there is clear distinction between yeast uridine kinases, from those of higher eukaryotes and other unicellular organisms.

Figure 6. Multiple sequence alignment shows DAS2 is unique among uridine kinases A) Multiple Sequence Alignment of uridine kinases spanning all kingdoms of life. Arrows indicate residues where the Das2 sequence diverges from the conserved amino acid. Sequences downloaded from UniProt (UniProt Consortium, 2018), UKase-UPRTase multi-domain sequences were truncated to remove UPRTase domains, and multiple sequence alignment carried out by MAFFT(Katoh et al., 2002) 6mer pairwise alignment algorithm, unreliably aligned positions were down-weighted using the GUIDANCE algorithm (Penn et al., 2010). Over 100 bootstraps were performed to generate alignment confidence scores, and the alignment was viewed in EMBL-MView (Chojnacki et al., 2017) B) Cladogram generated in EMBL-MView multiple sequence alignment in (A) representing hypothetical evolutionary relationships between DAS2, URK1 and other uridine kinases where Das2 has thee longest branch length, indicating the highest amount of character change. Distance values are calculated as the proportion of substituted amino acids over the alignment length.

3.2.13 Das2 catalyzes the uridine kinase reaction in vitro In order to assign an enzymatic reaction to Das2, I overexpressed Das2 in Escherichia coli and purified the recombinant protein using immobilized metal affinity chromatography (IMAC) and ion exchange chromatography. As shown in Figure 7, when the enzyme was incubated in the presence of uridine and phosphate donors ATP or GTP at pH 7.4 and 24 ºC, formation of uridine-5’-monophosphate was observed. I then measured activity of Das2 in uridine kinase assays involving activation of 3H-uridine to 3H-UMP. The velocities over a range of substrate concentrations catalytic constants were calculated. As shown in Figure 7a, the Km for uridine is 154 µM and the maximal specific activity is 243 µmol/ min /mg. These assays unambiguously reveal Das2 is a novel uridine kinase. The Das2 Km is in the range of the two human uridine/cytidine kinases at 268 and 50 µM for UCK1 and UCK2 respectively, which have a maximal specific activity of 0.6 and 2.7 µmol/mg/min respectively at 37 °C and pH 7.6 (Van Rompay et al., 2001). The E. coli uridine/cytidine kinase has a Km of 130 and 350 µM for uridine and cytidine, respectively at pH 7.8 and 37 °C (Valentin-Hansen, 1978). As to preference of co-substrate, we discovered that 100 times more UMP was produced in the presence of GTP than ATP. In preliminary cytidine kinase assays, Das2 was also active toward cytidine in vitro as cytidine-5-monophsophate was produced over a range of substrate concentrations.

A B

Figure 7. Das2 catalyzes a uridine kinase reaction A) Uridine kinase activity as measured by radiometric assay using various concentrations of 3H- uridine and 10 mM GTP as substrates pH 7.4, 24 °C. At 0, 6, and 12 minutes 3 technical replicates were taken, spotted onto DEAE filters, washed, eluted and measured by scintillation counting. Points represent means of two independent biological assays, error bars represent +/- standard error. B) Uridine kinase reaction showing preference for GTP as phosphate donor in yeast

In order to rule out the presence of a co-purified uridine kinase from Escherichia coli, I prepared a catalytically inactive version of Das2 G17E by mutating the glycine-rich phosphate binding loop in the GTP binding domain to aspartate. The negatively charged aspartate in the point mutant repels the γ-phosphate of the GTP co-substrate and thereby increases the Michaelis constant for GTP (Saraste et al., 1990). When catalytically inactive Das2 was challenged in radiometric uridine kinase assays, production of UMP was not observed. In contrast, an equimolar Das2 was catalytically active. This assay rules out the possibility of a co-purified uridine kinase from the heterologous E. coli host.

In order to determine kinetic constants for Urk1 activities and resolve its role relative to Das2, I purified the protein by co-expressing an untagged Urk1 alongside his-tagged Das2 and his-tagged Das2 G17E and measured UPRTase activity in radiometric and mass- spectrometry based assays. As shown in Figure 8, both assays identified UPRTase activity in Das2 pull downs of Urk1, but not in the context of Das2 alone. The trend observed in this in vitro assay confirms a UPRTase activity for Urk1, which establishes Urk1 as a bifunctional uridine kinase and UPRTase. Under the conditions used for the purification, the amount of Urk1 relative to Das2 was very small as assessed by gel electrophoresis, so it was not possible to determine precise kinetic parameters for the UPRTase assay.

3.2.14 Das2 pull-downs of Urk1 reveal UPRTase activity

Figure 8. Das2 pull-downs of Urk1 identify the UPRTase activity for Urk1 6xHis-tagged Fur1 and 6xHis-Das2 were heterologously expressed in E. coli with and without co- expression of untagged Urk1. Proteins were purified by immobilized metal affinity chromatography and eluted with imidazole. Presence of co-purified Urk1 was confirmed by appearance of 60 kDa protein band by silver-staining. Purified proteins were challenged in UPRTase and uridine kinase assays initiated by the addition of enzyme and quenched with extraction solvent at 0, 5, 20, 60 and 120 minutes. The presence of UMP was detected by LC-MS. Enzyme-containing reactions were compared to no-enzyme control reactions containing an equal volume of elution buffer. Uridine kinase assays contained 2mM uridine, 2mM GTP, 10 mM MgCl2,100 mM Tris pH 8. UPRTase assays contained 400 µM uracil, 500 µM PRPP, 10 mM MgCl2. Bar-plots represent integrated intensity of UMP at 120 minutes. I assayed the purified Das2 protein for uridine phosphorylase and uridine kinase activities in the forward direction by incubating with saturating quantities of substrates according to (Reichard and Sköld, 1958) and product formation of uridine was observed by LC-MS. Ribose-1-phosphate was enzymatically synthesized from phosphorolytic cleavage of inosine (Klenow and Emberland, 1955) with clean-up by custom flash chromatography I developed. Incubation of purified recombinant Das2 in the presence of ribose-1-phosphate and uracil did not lead to production of uridine, uridine kinase reactions were used as a positive control. This is consistent with the uridine low degree of sequence similarity for Das2 compared to uridine phosphorylases.

3.3 Using resistance to pyrimidine analogues to determine the topology of pyrimidine salvage pathways in yeast The targets of azapyrimidines vary across organisms in the tree of life. Only 6-AU and 6- AUR among many pyrimidine analogues were found to be competitive inhibitors of OMP decarboxylase in yeast (Handschumacher, 1960). The inhibition of OMP decarboxylase has the growth inhibitory effect of reducing UTP levels by 3-fold (Exinger and Lacroute, 1992). 6-AUMP is known to inhibit OMP decarboxylase in human cells and is correlated with anti- tumour activity (Fallon et al., 1962).

As discussed in section 3.2.3, mutants in pyrimidine salvage enzymes are known to display varying degrees of resistance to nucleobase and nucleoside analogues, depending on the position of the enzyme within the pathway for activating the pyrimidine analogue. Therefore, topology of pathways can be determined by measuring relative resistance of pyrimidine salvage mutants. Accordingly, as a first step in determining the role of Das2 in the pyrimidine salvage pathway I aimed to measure the response of the das2Δ mutant compared to null mutations in genes encoding known pyrimidine salvage enzymes. In order to mitigate the chance of second site mutations leading to spurious conclusions, I ensured the 20-fold accumulation of uridine was linked to das2Δ by transforming the deletion allele into a diploid prototrophic wild-type strain, sporulating and phenotyping spore-derived haploid strains. Once the phenotype was confirmed as linked, I compared the sensitivity to 6-AU of the das2Δ to other freshly prepared mutants in pyrimidine salvage. As shown in Figure 9a, the wild-type strain is sensitive to 2ug/mL 6-azauracil whereas fur1∆ has strong resistance across all dosages tested from 0 to 500 µg/mL. In comparison, the deletion mutants for das2∆ and urk1∆ have overlapping intermediate resistance profiles for 6-AU. This confirms that the 6- AU resistance associated with das2Δ shown by Gómez-Herreros is present in the DST1 wildtype and is due to deletion of the DAS2 locus alone and that the grown benefit conveyed by das2Δ is not specific to dst1Δ suppression (Gómez-Herreros et al., 2012). This is also the first report of 6-AU resistance for the urk1Δ mutant.

3.3.1 Mutations in das2Δ and urk1Δ are non-additive Interestingly, as shown in Figure 9a, the das2∆ urk1∆ double deletion mutant did not show an additive resistance compared to the das2∆ and urk1∆ single mutants. There are three possible explanations for the resistance of the das2Δ mutants to 6-AU. According to the literature, we know that resistance stems from blocking an activation step of which there are two possible

choices for uracil: first, via a two-step pathway involving uridine phosphorylase and uridine kinase activities or second, via direct activation by UPRTase activity. Das2 could act in a linear pathway with Urk1, where one enzyme could act as a uridine kinase and the other could act as a uridine phosphorylase, an enzyme which catalyzes the ribosylation of uracil using ribose-1-phosphate as a ribose donor. Previous attempts to measure uridine phosphorylase in yeast have found no detectable activity (Grenson, 1969; Magni et al., 1975). Similarly, I did not detect uridine phosphorylase activity in vitro using purified Das2 nor copurified Das2 and Urk1. Thus, I rule out the possibility that Das2 is acting as a uridine phosphorylase. The second possible explanation relies on the physical interaction between Das2 and Urk1 and suggests a role for Urk1 in activating 6-AU, where the non-additivity of resistance observed in the das2Δurk1Δ double mutant as compared to the single mutants could result from formation of a protein complex between Das2 and Urk1. Hence in this model, Das2 is required for stabilizing UPRTase activity of Urk1. When UPRTase complexes purified from yeast are analyzed by gel filtration, the active fraction contains two co-eluting proteins, one consistent with the molecular weight of Fur1, the annotated UPRTase, and the other consistent with the molecular weight of Urk1 (Natalini, et al., 1979). Both proteins were shown to possess UPRTase activity (Natalini et al., 1979). Indeed, the 6-AU resistance profiles here determined are consistent with das2Δ and urk1Δ mutants lacking a minor UPRTase reaction.

The superimposable resistance profile of the das2∆ urk1∆ double mutant indicates that Das2 and Urk1 could be required for stabilization of one another. An alternate explanation involves Urk1 and Das2 working together to positively affect Fur1 stability in catalyzing the UPRTase reaction. Hence, in the absence of Das2 or Urk1, Fur1 would be unable to catalyze the UPRTase reaction to its highest capacity. These data are consistent with either model: the contribution of minor UPRTase activity from a Das2/Urk1 complex or with the stabilization of Fur1 by Das2 and Urk1. Alternately, there could be an indirect effect on 6-azauracil uptake or activation due to elevations in uridine in the das2Δ and urk1Δ strains. This indirect effect is ruled out as the Fur4 uracil permease is not affected by uridine (Séron et al., 1999) while steady state levels of transcript and protein are diminished when uracil accumulates (Blondel et al., 2004; Séron et al., 1999). Furthermore, when nucleobases and nucleosides were tested for inhibitory effects on UPRTase complexes, dUMP, UMP, UDP, TTP and dCMP inhibited activity, while an inhibitory effect of uridine on Fur1 was not observed (Natalini et al., 1979).

3.3.2 Activation of 6-azauracil occurs exclusively through the UPRTase reaction In order to assess the functional relatedness among these three enzymes and the uridine hydrolase encoded by URH1 and determine the cumulative effects of their deletion, I prepared all possible double and triple deletion mutant combinations and measured sensitivity to 6-AU. As shown in Figure 9b, when das2∆ and urk1∆ mutations are combined with mutation in the uridine hydrolase urh1Δ, there is no additive effect on 6-azauracil sensitivity. This is consistent with resistance to 6-azauracil emerging solely from the UPRTase reaction and not from a parallel activation pathway involving uridine phosphorylase/uridine kinase reactions. If the uridine phosphorylase/uridine kinase reactions were significant, 6-azauridine formed could no longer be hydrolyzed back to 6-azauracil and would cause a further increase in drug resistance in the das2Δ and urk1Δ mutations. Since all combinations of mutants with urh1Δ overlap in 6-azauracil resistance with the URH1 counterparts, Urh1 does not contribute to 6-azauracil resistance, and therefore all activation of 6-azauracil is through the UPRTase reaction. Accordingly, the urh1Δ single mutant is equivalent to wild type cells in terms of 6- azauracil sensitivity. The fact that the 6-azauracil resistance represents activities of the UPRTase arm of 6-azauracil activation reflects the unidirectional action of uridine hydrolase, the lack of a uridine phosphorylase and supports the role for Urk1 functioning as a minor UPRTase leading to intermediate resistance profiles.

As shown in Figure 9c, double and triple deletion mutants of pyrimidine salvage enzymes with fur1∆ phenocopy the resistance profile of the fur1∆ single mutant, indicating a complete block in 6-azauracil activation. Notably, these cells have increased intracellular uracil levels (Grenson, 1969; Jund and Lacroute, 1970), which act to repress the FUR4 transcript encoding the uracil permease by which 6-azauracil gains entry to the cell (Séron et al., 1999).

As shown in Figure 9d, we examined the 6-AU resistance of other components of pyrimidine salvage metabolism. The fui1Δ uridine permease deletion mutant shows a wild-type phenotype consistent with its lack of affinity and cross-membrane transport for uracil and 6- azauracil. In contrast, the fur4Δ uracil permease deletion, which is the major transporter of uracil and 6-azauracil, shows an intermediate resistance to 6-azauracil. The fact that the fur4Δ deletion strain is not fully resistant, indicates that 6-azauracil entry into the cell is not saturable, and uptake may occur by a non-specific process, or that another transporter is involved, such as Tpn1, Fcy2, Fcy22, and Fcy21, which are implicated in the Candida albicans resistance to 5-fluorocytosine (Hope et al., 2004).

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Figure 9 Growth inhibition of pyrimidine salvage mutants by 6-azauracil Wild-type and pyrimidine salvage strains were pre-grown in minimal media to an optical density of 1.0 and sub-cultured to an optical density of 0.1 in YNB-glucose with addition of 6-azauracil in a 96-well plate. Cells were shaken for 20 hours at room temperature and the optical density was measured by absorbance at 600 nm at the end of this time period. % growth was calculated in Excel as OD for each strain was normalized to untreated control. Plotted values represent the means of 2 biological replicates. Error bars represent +/- standard error.

Sensitivity to 5-fluorouridine identifies uridine metabolizing enzymes The 6-azauracil results indicate differences in activation of nucleobases via the UPRTase reaction for pyrimidine salvage mutants. In contrast, since uridine analogues can be activated in two ways: via hydrolysis to uracil or via phosphorylation to UMP, measuring responses to the nucleosides 5-fluorouridine (5-FUR) and 6-azauridine (6-AUR) reveals quantifiable differences in nucleoside activation across uridine kinase, uridine hydrolase and UPRTase mutants. The response to pyrimidine nucleoside analogues provides an overall output for activation through the pyrimidine salvage pathways and one can thereby associate relative activities between various mutants. I therefore measured sensitivity to 5-FUR and 6-AUR for single, double and triple knockout mutants compared to wild-type, for which the responses largely overlap. I will discuss the 5-FUR results as the phenotypes show greater differences in magnitude due to the potency of the drug and are therefore easier to interpret. When assaying 5-FUR sensitivity, it is important to note that the commercially available compound is hydrolyzed to 5-FU, both endogenously and exogenously as the N-glycosidic bond is sensitive to hydrolysis under acidic conditions and is enzymatically cleaved by hydrolases. As shown in Figure 10. Growth inhibition of pyrimidine salvage mutants by 5-fluorouridine extracellular hydrolysis is evident as the deletion mutant for the uridine permease fui1, has an intermediate resistance to the nucleobase analogue. It is important to note that the fui1Δ and urh1Δdas2Δurk1Δ mutants represent complete blocks in 5-FUR activation, precluding entry and activation, and therefore phenocopy each other’s resistance profile (Figures 10b, 10d). Further resistance (Figure 10c) is due to resisting endogenous and exogenous formation of 5- FU in addition to 5-FUR and their combined activation to 5-FUMP.

Amongst strains analyzed over 5-fluorouridine concentrations from 0 to 5 mg/mL, wild-type yeast cells and urh1Δ mutant cells are the most sensitive to 5-FUR as they are inhibited by 0.3 µg/mL 5-FUR. All other strains fall into three major resistance phenotypes, described below.

3.4 Response to 5-fluorouridine is not additive between das2Δ and urk1Δ The first set of 5-FUR resistant mutants includes the das2Δ deletion mutants and share an expected phenocopy of urk1Δ and urk1Δdas2Δ deletion mutant strains as these mutants are blocked in uridine kinase activity. As shown in Figure 10a, each mutant is equally resistant to 8.0 µg/mL 5-FUR. In this class of phenotypes, the cells are blocked for a single major 5-FUR

activation pathway provided by uridine kinase. From our quantitation of uridine and uracil described below, we know these strains accumulate uridine to an extent where Urh1 can then contribute to hydrolysis. Urh1-dependent hydrolysis of 5-fluorouridine to 5-fluorouracil causes activation of 5-fluorouracil to 5-fluoro-UMP via the UPRTase activity of Fur1. Since uridine kinase mutants are not impaired for Fur1, they only show partial resistance to 5-FUR. This result also indicates that Fur1 is stable and is not dependent on uridine kinase activity.

3.4.1 Uridine accumulation unmasks Urh1 function In the second class of 5-FUR resistant mutants with resistance up to 1.6 µg/mL, strains are substantially, yet incompletely, blocked in direct activation of 5-FUR to 5-FUMP and cannot hydrolyze 5-FUR to 5-FU. This class consists of urh1Δdas2Δ and urh1Δdas2Δ fur1Δ. These two strains overlap in profile with the fur1Δ strain which is blocked in activation of residual endogenous and exogenous 5-fluorouracil and is deficient in uridine kinase activity though destabilization and reduction of study state enzyme levels. This is consistent with loss of Fur1 leading to a destabilization in Das2 activity. This phenotype happens to overlap with urh1Δfur1Δ as it cannot activate residual exogenously hydrolyzed 5-FU as urh1Δ is dominant over fur1Δ since Fur1 is only able to activate uracil produced from hydrolysis of uridine by the action of Urh1. It is important to note that the contribution of Urh1 to the pathway is initially masked by the low levels of uridine. In a urh1Δ strain, uridine kinases provide the major mode of activation of 5-FUR. Urh1 is not implicated in the response to 5-FUR until uridine accumulates, in the case of insufficient uridine kinase activity, due to the high Michaelis constant of Urh1 toward uridine at 0.86 mM (Magni et al., 1975). After this accumulation, Urh1 is active in hydrolyzing uridine to uracil and leads to activation of 5- fluorouracil to 5-fluoro-UMP by UPRTase activity.

3.4.2 Residual activities of Urk1 and Das2 are unveiled by double and triple mutant resistance to 5-fluorouridine As shown in Figure 10c, the third class is resistant to 5-fluorouracil concentrations below 0.03 mg/mL, and consists of das2Δfur1Δ, urk1Δdas2Δfur1Δ and urk1Δfur1Δ. Notably, the das2Δfur1Δ mutant is less resistant than the urk1Δfur1Δ mutant and the urk1Δdas2Δfur1Δ triple mutant. The increased resistance of urk1Δfur1Δ and urk1Δdas2Δfur1Δ relative to das2Δfur1Δ is due to the loss of the UKase and/or UPRTase activity of Urk1 that otherwise would activate both 5-FU and 5-FUR product to 5-FUMP. The major differences in resistance profiles are shown in Figure 10c between the fur1Δdas2Δ double mutant, and the

fur1Δurk1Δ double mutants. The difference in sensitivities could be due to the uridine kinase activity of Urk1 or the UPRTase activity of Urk1, or both. The most resistant strains are urk1Δdas2Δfur1Δ and urk1Δfur1Δ, which are completely blocked in both UPRTase and uridine kinase action. When comparing these two mutants, it is important to note that when urk1Δ and das2Δ mutations are combined, loss of Das2 in an urk1Δ background does not affect 5-FUR resistance, pointing again to compromised activity of Das2 when Urk1 and Fur1 are deleted. This is consistent with measurements of Urk1 levels in the das2Δ mutants where Urk1 is maintained at 88% of wild-type levels as shown in Figure 18. These data exemplify the dependence of Das2 on a functional Urk1 and indicate that Urk1 is not dependent on a functional Das2. Furthermore, the das2Δfur1Δ double mutant is more resistant than both the das2Δ and fur1Δ single mutants, due to the combined blockage in uridine kinase and UPRTase activities in the double mutant. The additive effect of fur1Δ deletion in combination with das2Δ or urk1Δ shows that deletion of Fur1 is not dependent on Das2 for stability.

3.4.3 Blocking UPRTases while Urh1 is active creates a sink for 6-azauracil and uracil Another interesting result of this analysis is shown in Figure 10b, where the non-overlapping phenotypes for the urh1Δurk1Δfur1Δ mutant as compared to urk1Δfur1Δ where the strain carrying wild-type Urh1 is more resistant. This difference in phenotypes indicates that when Urh1 is active, and uridine levels are elevated through loss of URK1, the Urh1 enzyme hydrolyzes 5-FUR to 5-FU, and thereby provides additional resistance especially when the two UPRTases are absent. A possible explanation is that active Urh1 provides a sink for 6- azauracil as there is no route to activation of 6-AU in the absence of UPRTase activity leading to increased resistance in URH1 strains compared to urh1Δ. Further indication that wild-type Urh1 provides a sink for 6-AU in UPRTase mutants is demonstrated by the fact that uracil only accumulates in UPRTase mutants when Urh1 is present, which will be shown by quantitation of uracil in Figure 15. A less likely mechanism is when Urh1 is active, in combination with elevations in uracil levels, the more hydrophobic uracil and 5-fluoroauracil may produce an indirect cytotoxic effect on yeast cells leading to slowed growth. However, there is no direct evidence for this in diseases where uracil accumulates as the etiology is complex (Berger et al., 1984). The rate limiting step of 5-fluorouracil administration in humans is first pass metabolism in the liver where it is cleared by dihydropyrimidine dehydrogenase to dihydrouracil using NADPH as a reducing cofactor and subsequently to β-

alanine and CO2. Individuals with mutations in dihydropyrimidine dehydrogenase have elevated circulating uracil (van Staveren et al., 2011). I could not find evidence of accumulations of uracil leading to secondary cytotoxicity and I do not observe growth defects in mutants that accumulate high levels of uracil, although it does not rule out a secondary effect specific to 5-fluorouracil.

Of important note is that UPRTases are weakly expressed in mammalian cells while OPRTases have, instead, been shown to catalyze activation of 5-FU using PRPP as a co- substrate (Houghton and Houghton, 1983). In yeast, this alternate 5-FU activating activity could possibly be carried out by two closely related enzymes, the OPRTases Ura5 and Ura10. However, we see weaker resistance in the urh1Δurk1Δfur1Δ mutant than the urk1Δfur1Δ mutant, suggesting that 5-FU hydrolyzed by the wild type Urh1 in the urk1Δfur1Δ mutant is not activated by OPRTases to provide additional toxicity. We know that the OPRTases are active in these prototrophic strains growing on minimal medium as the strains display wild- type levels of de novo pyrimidine metabolites.

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Figure 10. Growth inhibition of pyrimidine salvage mutants by 5-fluorouridine Wild-type and pyrimidine salvage mutant strains were pre-grown in minimal media to an optical density of 1.0 and sub-cultured to and optical density of 0.1 in YNB-glucose with addition of 5- fluorouridine in a 96-well plate. Cells were shaken for 20 hours at room temperature and the optical density was measured by absorbance at 600 nm. % Growth was calculated in Excel as OD for each strain was normalized to untreated control. Plotted values represent the means of 2 biological replicates each containing two technical replicates. Error bars represent +/- standard error.

3.5 6-azauridine phenotypes match those for 5-fluorouridine The resistance profiles obtained for pyrimidine salvage mutants in response to 6-azauridine (6-AUR) are given in Figure 11. These results agree with the 5-FUR profiles shown in Figure 10. As demonstrated with 5-FUR, commercially available 6-AUR contains a significant fraction of 6-AU, as determined by differential chromatographic retention times and appearance of 6-AU by LC-MS (data not shown). When compared to commercial 5-FUR, 6- AUR is in a different proportional mixture of nucleobase and nucleoside, 6-FU and 6-AU, respectively. As shown in Figure 11d, the fui1Δ strains and das2Δurk1Δurh1Δ strains overlap in resistance profiles and this is due to sensitivity to the 6-AU component and overlapping inactivation of the 6-AUR component. The das2Δurk1Δurh1Δ and fui1Δ have overlapping profiles as these two mutants represent two different blocks for 6-AUR activation. In the das2Δurk1Δurh1Δ mutant, 6-AUR is taken up, but not further activated. In the fui1Δ mutant, 6-AUR cannot enter the cell. In both cases, sensitivity is due to background hydrolysis to 6- AU (either pre-existing or caused by cells) and activation of this 6-AU by an active Fur1 to 6- aza-UMP.

Across strains analyzed for 6-AUR sensitivity from 0 to 25 mg/mL, the most sensitive strains are the wild-type (Figure 11a-d) and urh1Δ mutant (Figure 11b). Both of these strains have fully functional UPRTase and UKase activities. As shown in Figure 11a, the fur1Δ mutant is less resistant than the das2Δ, urk1Δ and das2Δurk1Δ mutants. This likely reflects a distinct ratio of 6-AUR to 6-AU compared to 5-FUR to 5-FU. As demonstrated by western blotting in Figure 18, this lower resistance shows Fur1 is required for stability of Das2 and Urk1 in activating 6-AUR. As previously demonstrated in response to 6-AU and 5-FUR, these uridine kinase deletion mutants are overlapping in 6-AUR sensitivity profiles. As shown in Figure 11b, in contrast to the 5-FUR data set, the resistance profiles for fur1Δ and urh1Δfur1Δ are overlapping—indicating that Urh1 may have a higher Km for 6-AUR than 5-FUR.

As shown in Figure 11c, the das2Δfur1Δ mutant has a slightly lower resistance than the urk1Δfur1Δ and urk1Δdas2Δfur1Δ mutants as the presence of a functional Urk1 may maintain a UPRTase activity conveying sensitivity to 6-AU. Similar to the 5-FUR results, the most resistant mutants to 6-AUR are urk1Δfur1Δ and urk1Δdas2Δfur1Δ. These mutants are completely blocked in uridine kinase and UPRTase activities and the matched resistance over the urk1Δfur1Δ mutant indicates that Das2 is reliant on Urk1 for activity.

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Figure 11 Growth inhibition of pyrimidine salvage mutants by 6-azauridine Wild-type and pyrimidine salvage strains were pre-grown in minimal media to an optical density of 1.0 and sub-cultured to and optical density of 0.1 in YNB-glucose with addition of 6-azauridine in a 96- well plate. Cells were shaken for 20 hours at room temperature and the optical density was measured by absorbance at 600 nm. % Growth was calculated in Excel as OD for each strain was normalized to untreated control. Plotted values represent the means of 2 biological replicates error bars are +/- standard error of the mean.

A wealth of metabolic information can be drawn from the resistance profiles of the pyrimidine salvage mutants. In many cases, these data indicate activities associated with each enzyme and how enzyme activities, enzyme levels and metabolite levels are integrated into the overall use of the pathway. For 6-AU resistance profiling, we observed phenotypes indicating UPRTase activities for Fur1 and Urk1 and functions for their binding partners. There is a major contrast between the resistance profiles for nucleobase and nucleoside analogs, reflecting the major differences in metabolism of nucleobases as compared to nucleosides. There were three primary resistance phenotypes that indicate Urk1 and Fur1 supply UPRTase activities, but their interaction is non-additive as Urk1 is dependent on Fur1, while Das2 is dependent on both Urk1 and Fur1.

3.5.1 Enzyme activity assays indicate Das2 and Urk1 both function as uridine kinases 3.5.2 Radiometric assays on cell free lysates In order to measure kinetics of uridine activation to UMP in various genetic backgrounds, we carried out uridine kinase assays on cell free lysates shown in Figure 12, where we measured incorporation of 3H-uridine to 3H-UMP (Cheng et al., 1986). We normalized activities to total protein levels quantified by BCA assay and specific activities are given relative to wild-type. As shown in Figure 12a, the das2Δ mutant had a severe reduction in uridine kinase activity to 15% of wild-type which were further reduced to 4% in the urk1Δ background. The das2Δ mutant retains a 15% of wild-type uridine kinase activity which is likely the result of Urk1 acting as a uridine kinase. The fur1Δ mutant uridine kinase activity phenocopies the urk1Δ and urk1Δdas2Δ double deletion mutant, indicating that both uridine kinase activities are dependent on Fur1. As shown in Figure 12b, the urk1 ukase and urk1 UPRTase point mutants do not substantially differ in levels of uridine kinase activity, which is likely due to the less severe destabilizing effect of the point mutations on Das2 and Urk1 protein levels that I demonstrate below by Western analysis.

As shown in Figure 12b, the das2Δ activity was not phenocopied by the das2 ukase point mutant, where activity was reduced even further to 6% of wild-type. As discussed below, the das2 ukase point mutant has reduced steady state levels of Das2 and Urk1. Indeed, all point mutations were detrimental to both Das2 and Urk1 steady state levels as discussed below, leading to a complete block in the uridine kinase reaction.

Figure 12 Specific activities of uridine salvage enzyme reactions A) 3H-Uridine kinase specific activity measured for deletion mutants relative to wild-type; dotted line is 1% of Wild-type B) 3H-Uridine kinase specific activity measured for point mutants relative to wild-type; dotted line is 1% of wild-type C) 3H-UPRTase activity measured for deletion mutants relative to wild- type; dotted line is 5% of wild-type D) 3H-UPRTase activity measured for point mutants relative to wild-type; dotted line is 5% of wild-type For A) Wild-type and pyrimidine salvage mutant strains were pre-grown in minimal media to an optical density of 1.0. 500 mL cultures were collected by centrifugation, concentrated to 1.6 mL in 100 mM Tris pH 7.4 to which 400 µL of zirconium beads were added. Cells were lysed by bead-beating at 500

rpm for 40 minutes. Lysed homogenates were centrifuged at 30, 000 x g for 30 minutes at 4 °C, 300 µL of the top-most supernatant was removed and assayed for protein content by micro-BCA assay (Thermo-Fisher Scientific). Uridine kinase reactions were carried out in the presence of 100 mM Tris, pH 7.4, 10 mM GTP, 10 mM MgCl2, [5,6-3H] uridine 10 µCi/mL, 40 µM uridine, 1 µL homogenate /10 µL. For B) Wild-type and pyrimidine salvage mutant strains were grown and lysed as above. UPRTase reactions were carried out in the presence of 100 mM Tris, pH 7.4, 1 mM PRPP, 5 mM MgCl2, [5,6- 3H]-uracil 10 µCi/mL, 200 µM uracil, 1 µL homogenate /10 µL. Reactions were initiated by the addition of homogenate, after 0, 12 and 24 minutes had elapsed two replicate 10 µL samples were quenched in 40 µL 1M urea. Quenched reactions were spotted to DEAE filters and washed 3 times in 50 mL 10 mM Tris pH 7.4. 3H-UMP was eluted from filters in 1 mL 0.5 M NaCl, 1N HCl shaken for 30 minutes, to which 9 mL scintillation fluid was added. Samples were shaken to mix and left to stand overnight before measuring radioactivity by liquid scintillation counting. CPM was converted to DPM and µmoles 3H-UMP, divided by time to calculate production rates. Values were divided by total protein levels determined by BCA assay to give specific activity. Dots represent the means of 3 independent biological assays each containing two technical replicates. Error bars represent +/- standard error.

Figure 13 In vivo tracking of uridine kinase activity using 6-azauridine incorporation in growing cultures Wild-type and pyrimidine salvage mutants were pre-grown in YNB-glucose and sub-cultured to an optical density of 0.1 and grown at room temperature to reach an optical density of 0.4. Metabolite extract samples were taken at 1 minute post-6-azuridine addition. 6-aza-UMP was separated and detected by LC-MS. Values represent intensities of 6-aza-UMP peak for each sample. Barplots represent values for a single replicate. In advance of adopting radiometric activity assays for uridine kinase activity, I developed an orthogonal assay for measuring uridine kinase activity in dividing cells. The assay results phenocopy the results from radiometric assays and are included as an orthogonal methodology I devised leveraging pyrimidine salvage mutants, pyrmidine analogues and LC- MS. I monitored the incorporation of 6-AUR to 6-aza-UMP in live cultures across various mutants by quenching metabolism at various time-points followed by measuring 6-aza-UMP by LC-MS. In order to minimize hydrolysis of 6-AUR to 6-AU by Urh1 when uridine levels accumulate in the various mutants, all mutants used were in the context of the urh1Δ background. As shown in Figure 13, in this experiment I observed that wild-type cells incorporated 6-AUR over time and the kinetics were matched by the urh1Δ mutant. When the 6-AUR incorporation was measured for the urh1Δdas2Δ, urh1Δurk1Δ and urh1Δdas2Δurk1Δ deletion mutants, I observed the following trends that overlap with the radiometric assays: 1) reduced production of 6-aza-UMP in uridine kinase mutants and 2) slightly higher production

of 6-aza-UMP in the urh1Δdas2Δ mutant, which is consistent with Urk1 acting as a minor uridine kinase or a UPRTase (as there is background 6-AU present).

3.5.3 UPRTase activity assays on cell free extracts When we measure UPRTase activity in various mutants by radiometric assay two patterns emerge as shown in Figure 12c. Firstly, the das2Δ and urk1Δ deletion mutants each harbour wild-type levels of UPRTase activity and these were phenocopied by point mutants shown in 12d. Surprisingly, a ~40% reduction in UPRTase levels was observed for the urk1Δdas2Δ mutant, which is the sole case of divergence from the non-additivity of urk1Δ and das2Δ single mutants in the other assays described in this work. All mutants in combination with fur1Δ have decreased UPRTase activity as compared to wild type cells, however, as shown in Figure 12d, the activity of point mutants in combination with fur1Δ is unexpectedly not as low as the fur1Δ single mutant alone. In this assay, Urk1 is apparently a negative regulator of a third UPRTase activity. In this case Fur1 would acting as the major UPRTase, and supporting Urk1 steady state levels, however, more UPRTase activity is revealed upon loss of Fur1 in combination with loss of function in Urk1 or Das2. Ura5 and Ura10 are the OPRTases whose human homologues are known to carry out the UPRTase side-reaction (Houghton and Houghton, 1983). Urk1 may be working to suppress UPRTase side-reaction activity of OPRTases. For example, combined loss of FUR1 and URK1 may induce a transcriptional response that upregulates Ura5 and Ura10 for de novo synthesis. Although pyrimidine analogue sensitivity profiles and work by Natalini and colleagues (Natalini et al., 1979) suggest and UPRTase activity for Urk1, it has not been captured by radiometric assay.

3.6 Metabolite levels elucidate bottlenecks and pathway logic An alternate method for determination of metabolic pathway topology is to measure changes in metabolite levels that result from knocking out a particular enzymatic step. Enzyme knockout leads to accumulation of metabolites upstream of the enzymatic step, depletion of downstream metabolites or both. Both of these phenotypes can be captured by LC-MS measurements of extracted metabolite levels. In the case of pyrimidine salvage mutants, UMP levels are not expected to decrease as UMP is produced by the de novo UMP biosynthesis pathway. In harmonizing de novo inputs when salvage is impaired, no growth defect is observed for salvage mutant yeast strains, and we find that UMP levels do not vary across pyrimidine salvage mutants as shown in Figure 14 where mean UMP levels range from 0.80 to 1.05 of wild-type for each deletion mutant.

Figure 14. UMP levels are stable across salvage mutants quantified by LC-MS Wild-type and pyrimidine salvage mutants were grown in YNB-glucose through several rounds of growth and sub-culture and metabolites were extracted at an optical density of ~0.5. Extract supernatants were dried under N2, reconstituted in a volume of water according to optical density at time of extraction. Metabolites were separated and measured by reverse-phase ion-paired chromatography and detected by Q-TOF. Chromatographic peaks were extracted and integrated using custom R software. Dots represent a single integrated intensity for UMP. Dots represent the means of 3 independent biological replicates. Error bars represent +/- standard error..

Figure 15 Relative uridine and uracil levels across pyrimidine salvage mutants

A-D) Uracil levels across pyrimidine salvage mutants compared to wild-type E-H) Uridine levels across pyrimidine salvage mutants compared to wild-type Wild-type and pyrimidine salvage mutants were grown in YNB-glucose through several rounds of growth and sub-culture and metabolites were extracted at an optical density of ~0.5. Extract supernatants were dried under N2, reconstituted in a volume of water according to optical density at time of extraction. Metabolites were separated and measured by reverse-phase ion-paired chromatography and detected by Q-TOF. Chromatographic peaks were extracted and integrated using custom R software. Dots represent the means of integrated intensities for four independent biological replicates, error bars represent +/- standard error of the mean.

The initial phenotype that motivated this study was the accumulation of uridine as compared to wild-type for the das2Δ mutant. In order to measure the contribution of Das2 to pyrimidine salvage relative to other pathway members, I prepared double and triple mutants of the deletion strains. Mutants are grown on minimal medium where biosynthesis and salvage are both active. Relative uridine and uracil levels were measured by LC-MS. Figure 15 shows uracil and uridine levels normalized to wild-type for various combinations of deletion mutants where metabolite extract volume was normalized to cell density at time of extraction. As shown in Figure 15a and 15d, strains with mutations in the uridine kinases, das2Δ or urk1Δ mutants, show equivalent elevations in uridine levels over wild-type, without elevations in uracil levels. The lack of uracil accumulation indicates Fur1 function is not compromised in the uridine kinase deletion mutants, the pathway through Fur1 is sufficient to activate hydrolyzed uracil to UMP, and that joint action of Urh1 and Fur1 is able to partially make up for loss of uridine kinase activity by activating uracil derived from uridine hydrolysis to UMP. Mutations in urk1Δ and das2Δ are non-additive as the urk1Δdas2Δ double mutant does not show a further increase in uridine levels. This phenotype parallels the non-additive resistance to 6-azauracil and 6-azauridine and further demonstrates the interdependence of Urk1 and Das2 enzymes for function. The fur1Δ deletion mutant shows a slight 5-fold accumulation of uridine, alongside a 10-fold accumulation of uracil which could be due to two causes: 1) destabilization of the Das2/Urk1/Fur1 complex to reduce the levels of the uridine kinases or 2) blocking direct uracil activation to UMP, uracil accumulates and causes Urh1 to work in reverse culminating in a slight accumulation of uridine. The second cause can be ruled out as uridine hydrolases are unidirectional due to water functioning as a reactant whose high concentrations effectively decrease the rate of the reverse reaction to nil. In Figures 15b and 15e we see that urk1Δ fur1Δ and das2Δ fur1Δ mutants do not show a further increase in uridine levels, but instead show 16-fold increases in uracil levels over wild-type indicating that Urh1 is active and is hydrolyzing uridine to uracil, where the uracil formed is blocked from activation to UMP by the loss of FUR1. The accumulation of uracil in urk1Δfur1Δ and das2Δ fur1Δ mutants is increased over mutations in fur1Δ, and we do not observe a further increase in uracil levels when mutations in das2Δ fur1Δurk1Δ mutant are compared to the urk1Δfur1Δ mutant. These results mirror the phenotypes captured by 6-AUR and 5-FUR resistance, where fur1Δ and das2/Δurk1Δ have additive genetic interactions.

The role of Urh1 is shown in Figures 15c and 15f where urh1Δ single mutants compromised for the uridine/cytidine nucleosidase have unaltered uracil and uridine levels. This is expected

as Urh1 status is not expected to affect the stability of other uridine salvage enzymes and uridine is not hydrolyzed to uracil at appreciable rates under the conditions used. The wild type levels of uridine and uracil in urh1Δ mutants further suggest that the uridine level in each of these strains is below the Km of Urh1, as deleting URH1 bears no effect on uridine or uracil levels. This supports the finding that during vegetative growth, the majority of uridine is directly activated to UMP by the action of uridine kinases (Grenson, 1969). A notable phenotype shown in Figure15f is the significant 1.5-fold (Student’s t-test p-value <0.001) uridine accumulation in the urh1Δ urk1Δ mutant as compared to the urh1Δdas2Δ mutant. The additional increase in uridine in the context of a uridine hydrolase block and uridine kinase mutant can be explained in two ways. Clearly the function of Urk1 is not completely diminished in the absence of Das2. This can be explained by either 1) Urk1 is the major uridine kinase whose function is not completely abolished when DAS2 is deleted or that 2) Das2 is the major uridine kinase and requires Urk1, but Urk1 carries out a minor uridine kinase activity independent of Das2. When the mutations in das2Δurk1Δurh1Δ are combined, we do not observe a significant increase in uridine levels over the urk1Δurh1Δ mutant, which further supports a model where Das2 is not active in the absence of Urk1. A second notable phenotype these data clearly show is that when Urh1 is active in uridine kinase mutants, the accumulated uridine is hydrolyzed to uracil. When Urh1 is blocked in these uridine kinase mutants, uridine accumulates instead of uracil, showing that uracil is formed from uridine.

From Figures 15d and 15h we can see that transporter mutants fur4Δ and fui1Δ are unaltered for uridine and uracil levels. This is expected as transporter status bears no impact on the stability of uridine kinases and uridine kinase activity is fully functional in each of these mutants.

When point mutants in Das2 and Urk1 were measured for uridine levels, the single mutants phenocopied the single deletions and when combined with the urh1Δ deletion, we observed that each of the point mutants phenocopied the full deletions. These phenotypes for the point mutants are consistent either with the expected catalytic dead phenotype for the point mutant, or with diminished steady state protein levels for the point mutant, which was quantified for tagged Das2 and Urk1 in Figures 16 and 17.

Overall these metabolite level data show: 1) Urh1 is the gatekeeper for uridine flow to uracil and its activity is not significant under vegetative growth conditions. It follows that uridine kinases are the major route of activation

of uridine to UMP. 2) Das2 and Urk1 are both required to maintain uridine kinase activity. 3) Loss of Fur1 reduces uridine kinase activity. 4) Differences in metabolite levels in the urh1Δ and fur1Δ contexts show that Urk1 likely possesses minor uridine kinase and UPRTase activities, as indicated by the non-overlapping phenotype for uracil accumulation in the urk1Δfur1Δ mutant as compared to the das2Δfur1Δ mutant. The minor uridine kinase activity of Urk1 is also supported by the background and uridine accumulation in the urh1Δdas2Δ and urh1Δurk1Δ backgrounds that are mirrored by the phenotypes on 5-fluorouridine. 5) Urk1 retains function in the absence of Das2. This is demonstrated by the increase in uridine in the urh1Δdas2Δurk1Δ mutant as compared to the urh1Δdas2Δ mutant, indicating a loss of the uridine kinase function following URK1 deletion and the increase in uracil seen in the das2Δfur1Δ and urk1Δfur1Δ mutants that is not further increased in the das2Δurk1Δfur1Δ mutant.

3.6.1 Metabolite levels indicate Urk1 is the elusive secondary UPRTase The URK1 gene encodes a 56 kDa protein with two predicted enzymatic domains; an N- terminal uridine kinase domain followed by a C-terminal uracil phosphoribosyl transferase domain as shown in Figure 5. UPRTase activity has not been annotated to Urk1 but is consistent with a previous study showing that a complex of UPRTase proteins was purified from yeast by gel filtration (Natalini, 1979). One protein had a similar molecular weight as Urk1, while a smaller protein had a molecular weight similar to Fur1 and both proteins showed UPRTase activity (Natalini, 1979). As previously mentioned, Urk1 and Fur1 have been previously shown by these and other data to physically interact. Further evidence for Urk1 acting as a UPRTase is that UPRTase mutants are known to accumulate uracil (Jund, 1970). As shown in Figure 15b, when I measured uracil levels for the fur1∆ deletion mutant, a 10-fold increase in uracil is seen over wildtype. Consistent with Urk1 acting as a UPRTase, the das2∆fur1∆ deletion mutant shows an additive increase in uracil levels to a 17-fold increase over wildtype, as Urk1 is destabilized by the loss of Das2. The largest increase in uracil levels was measured for the urk1∆fur1∆ deletion mutant showing a 20-fold increase over wildtype, which is consistent with a complete block in UPRTase activity through the loss of URK1. The urk1∆ single mutant does not show an increase in uracil because of minor activity it carries and due to the presence of active Fur1, which is not compromised in a uridine kinase mutant (Kern et al., 1990).

3.6.2 Metabolite level changes for point mutants phenocopy deletion mutants and do not pinpoint activities When the point mutants are examined for uracil and uridine levels, as shown in Figure 16, all three point mutants phenocopied das2Δ in terms of the magnitude of the uridine accumulations observed for the urk1Δdas2Δ and urk1Δdas2Δfur1Δ mutants, while the das2 ukase urh1Δ mutant phenocopied the uridine accumulation of the das2Δurh1Δ mutant. Furthermore, all three of the point mutants in combination with fur1Δ (das2 ukase fur1Δ, urk1 ukase fur1Δ and urk1 UPRTase fur1Δ mutant) phenocopy the uracil accumulation and uridine accumulations observed for the urk1Δdas2Δfur1Δ mutant. The result for the das2 ukase fur1Δ mutant is surprising, as the das2Δ null mutant shows lower uridine accumulation, emphasizing a negative effect on Urk1 stability in the das2 point mutant background. An explanation for the magnitude of the phenotypes is that each of the das2 and urk1 point mutations destabilize Urk1. Indeed, this is the case as when Urk1 steady state levels are measured by Western blot, Urk1 levels are reduced to 50% or less of wild-type as shown below in Figure 18 Urk1-3XHA levels. Measurement of Urk1 levels in this mutant shows that each mutation severely disrupts protein stability of Urk1 and thereby diminishes UPRTase function. Each of these point mutant alleles were fully sequenced from their 5’ untranslated region through their 3’ untranslated region to confirm the sequences were otherwise wild-type; they are untagged and expressed from their wild type promoter. The fact that introduction of a single glycine residue to aspartate mutation in the cofactor binding domain could result in phenocopying the null mutant suggests that stability of Das2 and Urk1 are easily compromised. Recent analysis into the basis of recessive nature of the majority of mutations shows that loss of function mainly results from negative effects on stability and folding that reduce the steady state level of the soluble and functional protein form (Tokuriki and Tawfik, 2009). In particular, mutating a substrate binding pocket places a greater burden on a protein’s thermodynamic stability, destabilizing the protein and leading to diminished steady state levels (Tokuriki et al., 2008). These studies suggest that the thermodynamic stability of Das2 and Urk1 are severely compromised by the point mutations leading to the reductions in steady state levels observed.

Overall, it appears that all three of the point mutants are equivalent. This suggests that the glycine to aspartate substitutions are too destabilizing for Urk1. URK1 encodes two hydrophobic domains, from 154 to 171 amino acids, and another from 410 to 425 amino acids. Since Urk1 is not known to be membrane-bound, these domains might serve as anchors

for protein-protein interactions with Das2 and Fur1. Mutations in the UKase domain may abolish the interaction with Das2 and lead to complete loss of uridine kinase activity. Across senso lato yeasts, these two predicted transmembrane domains are conserved, however the position within the polypeptide is not conserved. If I were to construct two additional point mutants, I would mutate the active site aspartate to alanine, D49A in Das2 and D91A in Urk1. In a structural study of uridine kinase, authors suggest mutating residues equivalent to K68D on Urk1, however, there is no conserved lysine in Das2 (Suzuki et al., 2004). A B

Figure 16. Uracil and Uridine levels for point mutants phenocopy full deletions A) Uracil levels across pyrimidine salvage mutants compared to wild-type B) Uridine levels across pyrimidine salvage mutants compared to wild-type Wildtype and pyrimidine salvage mutants were grown in YNB-glucose through several rounds of growth and sub-culture and metabolites were extracted at an optical density of ~0.5. Extract supernatants were dried under N2, reconstituted in a volume of water according to optical density at time of extraction. Metabolites were separated and measured by reverse-phase ion-paired chromatography and detected by Q-TOF. Chromatographic peaks were extracted and integrated using custom R software. Dots represent the means of integrated intensities for four independent biological replicates, error bars represent +/- standard error of the mean.

3.7 Das2 and Urk1 protein levels establish interdependence of uridine kinases Both pyrimidine analogue resistance profiles and metabolite level changes suggest the interdependence of Urk1 and Das2 for maintaining uridine kinase function. In order to determine whether Das2 and Urk1 are mutually required to maintain steady state levels of both enzymes, Das2 and Urk1 were tagged with the haemagglutinin (3XHA) small epitope tag. I determined the steady state levels of Das2-3XHA in various genetic contexts. Data are four replicate Western blots carried out on independent yeast culture samples. As shown in Figure 17, we found that, indeed, in the urk1Δ strain, the Das2-3XHA levels dropped to below 20% of wild-type. Also, the Das2 UKase domain point mutation leads to destabilized Das2 protein, decreasing Das2-G17E-HA levels to 25% wildtype levels. Notably, as shown in Figure 17c, the fur1Δ mutation also had a destabilizing effect on Das2-3XHA, decreasing steady state levels to 30% wild-type levels and in agreement with our uridine measurements and resistance assays that suggest only a partial dependence of uridine kinase activity on Fur1. These results are also consistent with previous observations that most UPRTase mutations led to decreased uridine kinase activity, as discussed above (Kern et al., 1991). Both the urk1 ukase and urk1 UPRTase point mutations in Urk1 were destabilizing, diminishing Das2-3XHA levels to 35% and 45% of wild-type levels, respectively. I went on to measure Urk1 protein levels to determine whether the destabilization in Das2 was due to loss of Urk1, and in Urk1-3XHA quantitation experiments shown below confirm that these destabilizations are due to loss of Urk1. All of these results establish the dependence of Das2 on a wild-type Urk1 and demonstrate that destabilizing substrate binding sites with glycine to aspartate substitutions in Das2 and Urk1 compromise both steady state levels and function for these enzymes.

A B

Figure 17 Das2-3XHA levels in various pyrimidine salvage mutants A) Representative immunoblot of Das2-3XHA and tubulin levels across salvage mutants B) Das2-3XHA levels in wild-type and fur1Δ strains Strains containing the Das2-3XHA allele in the wild-type and pyrimidine salvage mutant contexts were grown in YNB-glucose through several rounds of growth and sub-culture and protein was extracted and separated under denaturing conditions on a 12.5% poly-acrylamide gel. Transfer to PVDF 2 hours at 70 V at 4 °C, blocked overnight in Millipore FL-Blok fluorescent block buffer, primary hybridization overnight using anti-HA 3F10 (Roche) at 1:200, followed by hybridization with anti- tubulin 1:2500 Abcam EPR13799 for 1 hour at room temperature, followed by Li-COR IRDye® 680RD Goat anti-Rat IgG at 1:1000, IRDye® 800RD Donkey anti-Rabbit IgG at 1:3300 for 1 hour. Images were captured on a LI-COR Odyssey Imaging System using 10 minute exposure at 700 nm and 2 minute exposure at 800 nm. Signal was quantified in ImageStudio Lite Version 5.2 and linear regression was applied to Das2-3XHA dilution curves to determine signal. Dots represent the means Das2-3XHA signal intensities normalized to tubulin levels for three independent biological replicates, error bars represent +/- standard error of the mean.

As shown in Figure 18, Urk1 protein levels were also measured by detection of Urk1-3XHA levels in various genetic backgrounds. In accordance with our observations of decreased uridine kinase activity, we found that Urk1 protein levels are only slightly reduced in the das2Δ mutant but are substantially reduced to 60% of wild-type in the fur1Δ deletion background. The das2 ukase point mutation reduced Urk1-3XHA levels to 60% of wild-type, and the urk1 ukase point mutation reduced Urk1 to 50% of wild-type levels, while the urk1 UPRTase mutation conveyed a decrease to 40% wild-type.

When comparing the correspondence between effect of DAS2 and URK1 point mutations on Das2 levels and Urk1 levels, the levels of Urk1-3XHA are diminished across all mutant backgrounds except das2Δ. Each of the mutations appear to have a greater effect on reducing the steady state level of Das2 relative to Urk1. Overall, Das2 levels showed a greater decrease in protein stability when binding partners were mutated, suggesting that Urk1 is acting as a scaffold for Das2.

Figure 18 Urk1-3XHA levels measured across various salvage mutants backgrounds Strains containing the Urk1-3XHA allele in the wild-type and pyrimidine salvage mutant contexts were grown in YNB-glucose through several rounds of growth and sub-culture and protein was extracted according to (Zhang et al., 2011) when cultures had reached an optical density of ~0.5. Protein homogenates were separated under denaturing conditions on a 10% poly-acrylamide gel. Transfer to PVDF 2 hours at 70 V at 4 °C, blocked overnight in Millipore FL-Blok fluorescent block buffer. Primary hybridization overnight in Blok with anti-HA 3F10 IgG1 from rat (Roche) at 1:200, 100 µL 10% azide, 100 µL 25% Tween-20. Washed 5X in 100 mL TBST, hybridized with anti-α-tubulin IgG from rabbit Abcam EPR13799 1:2500, 100 µL 10% azide, 100 µL 25% Tween-20 1 hour at room temperature. Washed 5X in 100 mL TBST. Li-COR IRDye® 800 RD Goat anti-Rat IgG at 1:5000, IRDye® 680RD Donkey anti-Rabbit IgG at 1:3300 in 10 mL Blok with 10 µL 10% SDS, 100 µL azide, 100 µL Tween- 20. Washed 5X in 100 mL TBST. Images were captured on a LI-COR Odyssey Imaging System 2 minute exposure at 700 nm and 10 minute exposure at 800 nm. Signal was quantified in ImageStudio Lite Ver 5.2 and linear regression was applied to Urk1-3XHA dilution curves. Dots represent the means Urk1-3XHA signal intensities normalized to tubulin levels for three independent biological replicates, error bars represent +/- standard error of the mean..

3.8 Discussion I aimed to identify the roles of undiscovered enzymes by measuring metabolomic phenotypes by untargeted LC-MS metabolomics on prototrophic yeast deletion mutants. The identification of an accumulation in uridine over wild-type led to discovery of the novel pyrimidine salvage enzyme Das2. I determined that Das2 is the major uridine kinase responsible for 88% of activity in yeast, where Urk1 is responsible for the remaining activity. By measuring protein levels in various pyrimidine mutant backgrounds, I was able to show that Das2 function is dependent on wild-type Urk1 and Fur1. I also identified a novel function for Urk1 in catalyzing a minor UPRTase reaction and novel roles for Fur1 in supporting the stability of Das2 and participation in the novel UMP salvage complex. My study of the interaction between Das2, Urk1 and Fur1 demystifies the mechanistic basis of fur1Δ ura3Δ synthetic lethality where both UPRTase and Ukase activities are compromised along with essential de novo UMP biosynthesis by Ura3. This is the first pyrimidine salvage complex identified. As shown in Figure 19, this completely revises our understanding of pyrimidine salvage in yeast. The novel UMP salvage complex is analogous to the purinosome multi-component enzyme complex known for channeling of substrates from PRPP to inosine- 5-monophosphate formation and is responsive to cellular demand for purines (French et al., 2016).

Figure 19. Revised model of UMP salvage in Saccharomyces cerevisiae A) Model of UMP salvage with revisions to include Das2 as the major uridine kinase with Urk1 having a minor activity. Fur1 as the major UPRTase with Urk1 catalyzing a minor role. B) Model for contributions of enzyme complex members to uridine kinase activity with relative uridine kinase activity on the y axis and various mutant backgrounds compared to wild-type on the y-axis. Wild-type shows the highest activity, with the fur1Δ mutant showing tapered activity and further reductions in the das2Δ mutant. Das2 is unable to catalyze the uridine kinase reaction in the absence of Urk1 and Fur1.

The successful assignment of function of Das2 relied on the use of prototrophic strains. These mutants were grown in minimal media where the metabolic pathway for de novo pyrimidine biosynthesis was fully active to support growth. Strikingly, mutations in salvage do not have a growth defect under standard laboratory conditions and Urk1 and Das2 levels appeared largely unchanged in response to various environmental conditions such as induction of non- selective autophagy which is known to alter uridine levels (Huang et al., 2015), uridine or uracil supplementation, hydroxyurea treatment which is known to induce dNTP levels, imidazole treatment, 5-FOA treatment or 6-azauracil treatment. Western analysis of Das2 and Urk1 levels in response to environmental treatments are a single replicate and are unshown. Since these enzymes are lowly expressed and steady state levels are not altered in response to changing conditions, identification of phenotypes for these enzymes relied on the breadth and sensitivity of measurements offered by full-scan untargeted LC-MS metabolomics.

3.8.1 Pyrimidine analogs are crucial drug targets in pathogenic yeasts Pathogenic yeasts represent a major global burden as severe infection leads to 1.5 million deaths each year (Bongomin et al., 2017). In the arms race against pathogenic yeasts, antifungal inhibitors in order of clinical prevalence include amphotericin B, fluconazole and 5-fluorocytosine. Amphotericin B causes nephrotic toxicity and widespread resistance has emerged to fluconazole. Pyrimidine analogues like 5-fluorocytosine represent an important class of anti-fungal therapeutics to which resistance rapidly evolves (Hospenthal and Bennett, 1998). Natural 5-fluorocytosine resistance is reserved to a single clade of natural isolates of Candida albicans (Pujol et al., 2004) which was traced to a single nucleotide change in FUR1 (Dodgson et al., 2004). Resistance to 5-fluorocytosine has been studied in the lab. When a systematic screen was carried out in Candida glabrata to identify genes conveying 5- fluorocytosine resistance, no pyrimidine salvage enzymes were identified (Costa et al., 2015) as the authors supplemented with uracil. Uracil supplementation obscures 5-fluorocytosine sensitivity results as salvage mutants would be unable to activate uracil and would appear sensitive. These results suggest pyrimidine salvage may be an under-explored route to activation of pyrimidine analogue inhibitors. In order to determine whether the here- identified salvage mechanisms were conserved in pathogenic yeasts, I aimed to identify homologues of DAS2, URK1 and FUR1 in pathogenic yeasts Candida albicans and Cryptococcus neoformans. I carried out a sequence comparison using the tBLASTn algorithm and Blosum62 similarity matrix using S. cerevisiae sequences as queries of the Candida Genome Database and JGI MycoCosm database (Nordberg et al., 2014; Skrzypek et al.,

2018). This search revealed orthologs of FUR1 and URK1 in both strains, with a clear orthologue of DAS2 present in Candida albicans, as the closest orthologue of DAS2 in Cryptococcus neoformans is the URK1 dual uridine kinase/ UPRTase homologue.

Table 3. Orthologues of S. cerevisiae pyrimidine salvage enzymes in pathogenic yeasts

Candida albicans Cryptococcus neoformans

DAS2 CAWG_00840 27.2% identity, expect = 1.0 e-19 CNG02150 26% identity, expect = 6.1 e-08 URK1 CAWG_02735 52.4% identity, expect = 6.0 e-134 CNG02150 39% identity, expect = 6.8 e-72 FUR1 CAWG_04715 63.5% identity, expect = 7.0 e-40 CNE02100 67% identity, expect = 4.7 e-73 URH1 CAWG_05839 39.8% identity, expect = 2.0 e-55 CNE04590 28% identity, expect = 1.6 e -09

When ura3Δ is used as a marker for positive and negative selection in C. albicans, ura3Δ auxotrophic mutants are supplemented with uridine instead of uracil. The first case of ura3Δ use in Candida uses uridine instead of uracil, where the basis of this change is not addressed, but it's likely motivated by a difference in growth rate between the two supplements (Kelly et al., 1987). If uridine does endow a growth benefit over uracil, it suggests that Das2/Urk1 activity is highly active in C. albicans and is required to restore wild-type growth in absence of URA3.

As it appeared that UMP salvage complex members were conserved to Candida albicans, I aimed to determine whether pyrimidine salvage is crucial to survival of pathogenic yeast in the host by studying the conservation of URA1 encoding dihydroorotate dehydrogenase orthologues. Since mitochondrial localized URA1 orthologues require respiration using oxygen as a terminal electron acceptor, these yeasts would be rendered auxotrophic when exposed to the low oxygen tension within the host. Candida albicans is a facultative aerobe commensal fungus that colonizes both the aerobic oral cavity and anaerobic environment of the human gut. Biofilms produced by laboratory strains and clinical isolates of Candida albicans are also hypoxic (Fox et al., 2014) and cells that encode a mitochondrial dihydroorotate dehydrogenase for de novo synthesis of pyrimidines would rely on salvage to support macromolecular synthesis during growth under biofilm conditions. I compared URA1 homologues from Candida albicans to the mitochondrial dihydroorotate dehydrogenase sequence from S. pombe and cytosolic dihydroorotate dehydrogenase sequence from S. cerevisiae. Candida albicans dihydroorotate dehydrogenase has 50% identity to the mitochondrial quinone dihydroorotate dehydrogenase and only 25% identity to the cytoplasmic dihydroorotate dehydrogenase. These results suggest that Candida albicans

rely on respiration for de novo synthesis of pyrimidines and during hypoxic biofilm conditions would fully rely on salvage to support macromolecular synthesis. When one examines differences in the Candida albicans transcriptome during planktonic and biofilm conditions, there URK1 and DAS2 orthologues are expressed at higher levels during biofilm conditions. These inductions of UMP salvage complex components are consistent with our model of C. albicans relying on salvage during biofilm formation due to the dependence of Ura1 on respiration.

3.8.2 Saccharomyces cerevisiae salvage complex is conserved to Candida albicans In my hands, as well as Lacroute and Jund’s, pyrimidine nucleoside analogs are more potent than their nucleobase counterparts. Candida albicans takes up pyrimidine nucleosides through the uridine permease (Fasoli and Kerridge, 1990). UPRTase mutants are resistant to 5-fluorouracil and have cross resistance to 5-fluorouridine (Fasoli et al., 1990). This cross- resistance result is similar to the cross-resistance observed for fur1Δ mutants to pyrimidine analogues due to that is due to diminished uridine kinase activity as shown in this work. The finding in C. albicans further predicts that the S. cerevisiae uridine salvage complex elucidated in this work is present and functional in Candida albicans. The authors also found 5-fluoropyrimidines to be more potent than equivalent treatments with 5-fluorouracil. Together, these results suggest that pyrimidine nucleoside analogs may provide more potent therapies against pathogenic C. albicans infections over conventional nucleobase analogs. The resistance profiles from my work show that due to the multiple avenues for activation of nucleoside analogues over nucleobase analogues, full resistance is not achieved indicating that this class of pyrimidine analogues is an under-explored therapeutic tool. Overall, my results suggest that targeting the UMP salvage complex of C. albicans using pyrimidine nucleoside analogues will provide superior potency and reduced resistance compared to nucleosides analogues such as the commonly used 5-fluorocytosine.

3.8.3 Pyrimidine salvage is a selective target for neoplastic cells Pyrimidine nucleobase and nucleoside analogues are commonly used in the clinic and are dependent on phosphorylation for chemotherapeutic activity (Cihák and Rada, 1976). As human uridine phosphorylase and uridine kinases are expressed in solid tumours, they provide a selective mode of activation of nucleoside analogues to their inhibitory forms. Humans encode two single subunit uridine kinases, UCK1 and UCK2, and one potentially bifunctional uridine kinase and UPRTase, UCKL1 (Kashuba et al., 2002). Humans also

encode two uridine phosphorylases UPP1 and UPP2 (Johansson, 2003; Watanabe and Uchida, 1995), and one single domain UPRTase, UPRT (Li et al., 2007). Notably, the human UPRTase has a mutated uracil binding site and no measurable UPRTase activity was found when the recombinant enzyme was purified, suggesting it functions in a role that is independent of catalyzing the UPRTase reaction. Due to the absence of a UPRTase activity, activation of 5-fluorouracil is predominantly carried out by UMP synthase (Cantor et al., 2017; Houghton and Houghton, 1983). The UMP synthase carries out the final two steps of de novo UMP production in human cells by catalyzing the orotate phosphoribosyl transferase and orotidine-5-phosphate decarboxylase reactions (Suchi et al., 1997). The finding that UMP synthase activates 5-fluorouracil suggests that it may also catalyze the UPRTase reaction, or that 5-fluorouracil is acting as a mimic of orotic acid as shown for 6-azauracil (Rada and Doskocil, 1980).

Human cells are predominantly quiescent and require little macromolecular synthesis. In contrast, mitotic tumours both de-repress pathways for nucleotide biosynthesis and increase salvage to meet cellular demand. When uridine phosphorylase activity was measured in dividing cells a lower activity was found in gut mucosa and bone marrow compared to breast cancer tumours (Pizzorno et al., 2002; Pritchard et al., 1997). These UPase positive tumours are prime cases for selective activation of 5-fluorouracil and pyrimidine nucleobase analogs (Pritchard et al., 1997; Yan et al., 2006). Uridine kinase UCK2 is also a prominent activator of pyrimidine nucleosides as a primary step in their anti-tumour trajectory (Murata et al., 2004; Shimamoto et al., 2002). Both uridine/cytidine kinases are selectively expressed in neuroblastoma cell lines and are now recognized as a potential clinical target (Meinsma and van Kuilenburg, 2016). UCK1 is widely expressed in human tissues while UCK2 appears to have a developmental role as its expression is limited to embryonic tissues (Van Rompay et al., 2001). Both human uridine/cytidine kinases catalyze the activation of a broad range of nucleoside analogues (Van Rompay et al., 2001). The historical and active use of pyrimidine nucleoside analogues in the clinic indicates that current therapies can be repurposed for treatment of pathogenic yeasts.

The presence of a conserved uridine salvage complex in Candida albicans that is homologous to the here-elucidated Das2/Urk1/Fur1 complex of Saccharomyces cerevisiae, in hand with the low uridine kinase salvage activity in mitotic and quiescent human cells presents a prime opportunity for use of pyrimidine nucleoside analogs as antifungal agents.

3.8.4 6-azauridine is more potent than 6-azauracil In this system, we are not able to truly measure Ki for 6-AUR without quantitation of the large amount of 6-AU present in the 6-AUR—which can be quantified by LC-MS as 6- azauridine and 6-azauracil retain differentially in reverse phase chromatography. However, we can compare the sensitivity of wild-type to 6-AU and 6-AUR. Wild-type yeast cells are sensitive to 0.001 ug/mL 6-azauridine and sensitive to 1 ug/mL 6-azauracil. Even with hydrolysis of 6-AUR taken into account, 6-azauridine is much more potent, arguing that the uridine kinase reaction is responsible for salvaging more pyrimidines than UPRTase activity. This result is in agreement with findings by Jund and Lacroute, who found that the upper limit of sensitivity is 10 times lower for 5-FUR than 5-FU (Jund and Lacroute, 1970). The reduced solubility of 6-AU and 5-FU may compromise their potency as these two nucleobases are much less polar than their nucleoside counterparts. As discussed below, the higher potency of nucleoside analogs over nucleobase analogues has implications for treatment of pathogenic yeasts.

The potency of anti-metabolites is intrinsically linked to pool size. Activities of enzymes are also modulated by the concentrations of their substrates and products. Concentrations of uridine are 2 mM in E. coli (Park et al., 2016) (Bennett et al., 2009). However, yeast concentrate uridine in the vacuole (Boswell-Casteel et al., 2018). The same is true for humans, where plasma concentrations below 10 uM while tissues concentrate uridine to 10 times higher uridine levels with spleen having 70 times higher uridine levels over plasma (Darnowski and Handschumacher, 1986). The levels of pyrimidine intermediates in yeast are: UMP at 14.5 µM, UDP GlcNAc at 1 mM, UDP-glucose at 268 µM, and UDP at 37 µM, UTP at 500 µM (Bennett et al., 2009). In contrast, CMP levels are much lower than UMP at 5 µM, CTP levels at 250 µM in yeast (Bennett et al., 2009). PRPP levels are 47 µM, GTP is 250 µM, which is much lower than ATP at 2 mM (Bennett et al., 2009). A second reference that quantified uridine observed less than 20 µM in wild-type prototrophic yeast cells, which increases to 60 µM during nitrogen starvation (Huang et al., 2015). Huang also shows that uracil levels are 8 µM and double after nitrogen starvation and accumulate to 3 µM extracellularly. This is a 100-fold difference in uridine levels between E. coli and yeast cells.

The human homologues of DAS2, UCK1 and UCK2, are highly expressed in solid tumors alongside uridine phosphorylases, UPP1 and UPP2 which use ribose-1-phosphate to ribosylate uracil to uridine in a route to UMP formation distinct from fungi (Reichard and

Sköld, 1958). When the uridine phosphorylases are knocked out in mouse embryonic stem cells, an 10-fold increase in IC50 for 5-FU is seen, showing that these cells are reliant on dual action of uridine phosphorylase and uridine kinase which together provide a chemotherapeutic window for targeted for activation of pyrimidine analogue anti-neoplastic therapeutics (Cao et al., 2002).

New functional adaptations spring from point mutations. If a new function is beneficial it may become fixed. Whether it does or not depends on other enzymes encoded, environmental conditions and where the reaction is positioned within the metabolic network whether a new route can be made or another blocked (Copley, 2012). Although much of the sequence change in Das2 relative to other uridine kinases may be attributed to neutral drift, many of the changes occur in conserved residues and likely convey a fitness consequence or alter stability. Gene fusion generally occurs between proteins that act in a concerted fashion, such as catalyzing subsequent steps in a pathway, however Urk1 is unusual as its putative complex members are acting in parallel reactions to produce UMP from two different precursor molecules. Fusion proteins tend to have higher fitness due to tighter co-expression, such as the multi-functional enzymes involved in de novo UMP biosynthesis, however, this appears not to be an apparent advantage for Urk1 as Das2 is the major uridine kinase and Fur1 is the major UPRTase, operating in parallel reactions.

4 Elucidation of Tda5, a novel essential short chain dehydrogenase 4.1 Rationale and summary A major knowledge gap in eukaryotic metabolism is assignment of functional roles of short chain dehydrogenases (SDR). In yeast, this enzyme family contains 16 members, of which only 7 have been annotated. As a result of my screen for assignment of enzyme function to uncharacterized genes, I found that the SDR deletion mutant tda5Δ had vastly more metabolomic phenotypes than any other strain. As I show, many of these phenotypes were due to a second site suppressor mutation. I go on to show that the freshly dissected tda5Δ strains have a severe growth defect and are respiratory incompetent. These strains rapidly suppress by derepressing the sporulation specific SDR paralog of TDA5, YDL114W. This is not the only way to compensate for loss of TDA5, as I identified lesions in the ergosterol biosynthetic pathway that also alleviate loss of function in TDA5, and I find that antioxidant treatment rescues the growth of tda5Δ mutants. I present evidence for a role for Tda5 in lipid homeostasis and present a repressible expression system that allows for analysis of the initial metabolic changes induced by loss of function in TDA5 and mechanisms of genetic and chemical suppression. Ultimately, I find that Tda5 function is closely related to other yeast SDRs in lipid metabolism.

4.2 Results 4.2.1 Metabolomic phenotypes discriminate TDA5 from other uncharacterized enzymes The metabolome of the tda5∆ gene deletion mutant strain was first profiled among 120 candidate enzyme deletion mutants in a genetic screen I carried out to assign metabolic function to uncharacterized genes (Caudy et al., 2018). TDA5 encodes an uncharacterized ORF homologous to short chain dehydrogenase enzymes, containing a Rossman fold and conserved serine, tryptophan and lysine catalytic residues. Out of all the strains profiled across my metabolomic screen for enzyme discovery, the tda5∆ deletion mutant proved to have the highest number of metabolomic phenotypes observed spanning multiple metabolic pathways (Hanchard, 2015). The average number of mutant phenotypes in observed per strain was 1, whereas the tda5∆ deletion mutant harbored over 20 changes in 13C 15N-confirmed metabolite levels with a fold-change greater than 2 and a p-value of less than 0.001. When altered peaks were matched to known standards, I identified elevations in aromatic amino acid levels in the tda5∆ mutant compared to wild-type, alongside kynurenine pathway intermediates, a 10-fold increase in mevalonate over wild-type and elevations in many uncharacterized metabolites whose empirical formulas were confirmed to contain carbon and, in some cases, nitrogen by matching peaks to corresponding peaks in 13C 15N stable isotope tracing.

4.2.2 Growth phenotype of fresh deletion mutant Even more striking than the high number of metabolic phenotypes was the phenotype I observed when the freshly prepared tda5∆/TDA5 mutant was transformed, selected, sporulated and dissected. As shown in Figure 20 a and b, the tda5∆ mutant spores had a severe growth defect, taking four days to form a colony on YPD rich media at 30 °C, a full two days longer than wild- type. The freshly dissected strain was not able to grow on media containing respiratory carbon sources, such as glycerol, acetate and ethanol in contrast to the suppressed strains that grew at wildtype rate. The freshly dissected strain rapidly generated strains with growth rates near wild type, which are suppressed through mutation, as I will show. The growth defect of the freshly dissected tda5∆ strain was surprising, as the strain from the deletion collection was back-crossed and grown alongside wild-type ahead of metabolite profiling in my screen and a growth defect was not observed. As shown in the tetrad dissection of the tda5Δ strain from the prototrophic

yeast deletion collection back-crossed to wildtype displayed in Figure 20 c, where the segregation pattern of wild-type growth is consistent with the presence of a suppressor mutation, the reason that we did not observe a growth defect in this screen was that a second site suppressor mutation exists in the deletion set strain, and the growth benefit conveyed by the second site mutation would have been co-selected in the random spore selection used in generating strains used in my screen.

The growth defect and inability to grow on carbon sources requiring respiratory metabolism suggests that TDA5 is required for mitochondrial function. The petite phenotype—defects in mitochondrial function rendering cells unable to grow on non-fermentable carbon sources and leads to small colony size when grown on a fermentable carbon source—is in stark contrast to the suppressed strains, which have restored ability to grow on respiratory carbon sources. Since the freshly dissected tda5Δ mutant suppresses rapidly, throughout the course of experiments, I monitored for increases in growth rate during sub-culturing cells. I also streaked cultures used in experiments onto YP-glycerol to ensure that a new second site suppressor had not swept the population.

Figure 20. The freshly dissected tda5∆ strains are petite whereas tda5∆ from the prototrophic yeast deletion collection carried a suppressor mutation A) Freshly dissected tda5∆/TDA5strain dissected onto YPD incubated 2 nights at 30 °C B) Freshly dissected tda5∆/TDA5strain dissected onto YPD incubated 4 nights at 30 °C C) Dissected spore progeny of tda5∆ strain to back-crossed to wild-type shows segregation pattern of a single second site suppressor For A and B, the tda5∆::kanMX allele was amplified by PCR and transformed into FY4/FY5 prototrophic diploid and transformants were selected on YPD G418. Transformants were sporulated by shifting cultures from YPD growth to YPA followed by incubation in sporulation medium for 3 nights at 30 °C with shaking. Sporulation cultures were rinsed in water, digested with β-glucuronidase and dissected using by micromanipulator using a yeast dissection microscope onto a YPD Plate. The dissected haplo-spores were incubated at 30 °C. For C, the MATa haploid tda5∆::kanMX strain from the prototrophic yeast deletion collection was mated to FY5 MATα prototrophic wild-type, zygotes were picked by micromanipulation using a yeast dissection microscope, sporulated and dissected as above. 4.2.3 Tda5 is a novel essential dehydrogenase In order to discern the function of Tda5, I studied the protein superfamily that Tda5 belongs to. Tda5 is an uncharacterized enzyme belonging to the short chain dehydrogenase superfamily. Superfamily classifications indicate residue conservation patterns and domain boundaries for protein sequences and provide non-redundant organization of conserved domains across evolutionary history (Marchler-Bauer et al., 2017). The SDR superfamily of enzymes catalyzes a diverse set of NAD(P)(H) dependent reactions such as isomerization, epimerization, double bond reduction, dehydration, dehalogenation and aldehyde/alcohol reduction (Kallberg et al., 2010). SDR co-substrates and products span metabolic pathways including fatty acid biosynthesis, sterol biosynthesis, and ethanol fermentation. For the α-proteobacterium root nodule symbiont Sinorhizobium meliloti, the only organism for which SDR activity has been systematically studied, SDR functions provide ability to grow on a wide range of carbon sources (Jacob et al., 2008). The classical SDR domain contains an N-terminal NAD(P)(H) binding domain within the

Rossman fold, and a C-terminal substrate binding domain with three active site residues forming a catalytic triad: serine, tyrosine and lysine. When reactions proceed in the oxidative direction producing carbonyl products from hydroxyl substrates, the active site tyrosine abstracts a proton from the substrate, while serine polarizes the substrate C-O bond to facilitate electrons moving from the O-H bond to form the C-O double bond. Double bond formation simultaneously ejects a hydride which is donated to the NAD(P)+, which is stabilized by lysine coordination of a water molecule (Kavanagh et al., 2008). The NAD+/NADPH cofactor specificity for the SDR family can be predicted by the presence of an acidic residue conveying specificity to NADP(H) or two basic residues conveying specificity to NAD(H) at the end of the βαβ motif within the Rossman fold (Persson et al., 2003). Tda5 encodes acidic aspartate residues at positions 116 and 120, which would specify NADP(H). The substrate binding pocket is not well conserved, and it is not possible to predict the substrate from the primary amino acid sequence. The diversity of primary structure reflects the wide variety of substrates SDRs can act on.

4.2.4 Evolutionary conservation of TDA5 Since SDRs catalyze a wide range of reactions, and substrate cannot be predicted from sequence information, insights into SDR function can be captured from evolutionary relationships. Evolutionary relationships between proteins can be carried out by analysis of SDR sequences, where pairwise sequence alignments are used to generate a protein distance matrix followed by clustering least squares to determine genetic distance. Evaluations of genetic distance visualized in cladograms are available from NCBI conserved domains (Marchler-Bauer et al., 2017), where Tda5 is a member of the cd05339 sequence cluster named for human 17-beta-hydroxysteroid dehydrogenase XI-like, classical type C SDRs, shown in Figure 21a. The reaction catalyzed by the enzyme that the cd05339 cluster is named for is catalysis of 5α-androstan-3-α,17-β-diol (3-α- diol) to androsterone a human steroid hormone shown in Figure 22 b (Chai et al., 2003). Conserved domain models are organized into hierarchies based on inferred common descent . As shown in Figure 21 a there are 26 members of this short chain dehydrogenase conserved domain cluster with YDL114W as the outgroup. Among fungi, Tda5 is conserved all the way back past Ascomycota and into Basidomycota, and is present in a high number of eukaryotic taxa including Tetrahymenia, Drosophila, Xenopus, Danio and Mus. Cd05339 is also present across kingdoms, found in both Bacteria and Eukarya. There are two human short chain dehydrogenases in the cd05339 cluster, where the closest human homologues of TDA5 are the HSD17B11 and 97

SDR16C5 genes encoding two distinct activities, where the DHB11 protein encoded by HSD17B11 catalyzes the aforementioned steroid hormone reactions and the epidermal retinol dehydrogenase II (RDHE2) protein encoded by SDR16C5 converts retinol to retinal as shown in Figure 22b (Lee et al., 2009). Retinol dehydrogenase is the first and rate limiting step of activation of retinol (Vitamin A) to retinoic acid (Sandell et al., 2007). Unlike DHB11 which is cytoplasmic, the RDH2 enzyme is localized to the endoplasmic reticulum and the localization is conserved among retinol dehydrogenases, which are often cross-reactive on 3-alpha- hydroxysterols (Belyaeva and Kedishvili, 2006; Biswas and Russell,1997; Chetyrkin et al., 2001). In higher eukaryotes, steroid hormones and retinoic acid are important signalling molecules regulating growth and metabolic responses (di Masi et al., 2015). The structure of hRDH2 has been determined however it has not been compared to retinol dehydrogenases as few SDR structures have been determined (Berman et al., 2000). Pairwise sequence alignments of TDA5 and its human orthologs are displayed in Figure 21 showing low similarity over the active site. Tda5 must have a distinct role in yeast from oxidation of retinol as retinol is neither produced nor activated to retinoic acid. Tda5 instead may display a preference for sterols in line with the DHB11 substrate and known promiscuous substrates for retinol dehydrogenases (Heery et al., 1993).

A Ydl114w Tda5

RDHE2

DHB11

Figure 21. TDA5 orthologs are distributed among eukaryotes and bacteria and are conserved to human homologs C B A) Phylogenetic tree for NCBI Conserved Domain 5339 sequence cluster where Gene ID, conservation over range of amino acids and organism are annotated in purple boxes B) Protein sequence alignment for Tda5 and closest human homologs RDH2and DHB11, active site catalytic residues are indicated in purple For A, the cd5339 cluster data was downloaded and viewed in NCBI protein domain hierarchy viewer CDTree (Marchler-Bauer et al., 2007) and cladogram prepared. For B, RDHE2 and DBE11 FASTA sequences were downloaded from UniProt (UniProt Consortium, 2018) and TDA5 sequence downloaded from SGD (Cherry et al., 1998) and pairwise alignments were carried out in LALIGN Server from Swiss Institute of Bioinformatics.

Figure 21. TDA5 orthologs are distributed among eukaryotes and bacteria and conserved to human homologs

Figure 22. Reactions catalyzed by human orthologs of TDA5 conserved domain cluster 5539 A) Human 17β-hydroxysteroid dehydrogenase type XI reaction: oxidation of androstan-3-α,17-β -diol to androsterone using a nicotinamide adenine(phosphate) dinucleotide cofactor B) Human epidermal retinol dehydrogenase II reaction: oxidation of retinol to retinal using a nicotinamide adenine dinucleotide cofactor

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As previously mentioned, yeast do not biosynthesize retinol. However, low doses of retinol do stimulate growth in rich media on fermentable and respiratory carbon sources and truncate lag phase in glycerol whereas high doses of retinol slow growth on glucose and completely inhibit growth on glycerol—consistent with conveying a petite phenotype (Cheng and Wilkie, 1991). The growth-promoting effect of low dose retinol was attributed to promotion of membrane function, as the prenyl tail of retinol and other metabolites can insert into phospholipid membranes to protect against stress, whereas high doses of retinol may destabilize membranes. Treatment with antioxidants and glutathione rescued the respiration defect of high dose retinol, suggesting that retinol toxicity occurs through production of free radicals. The tda5Δ mutant from the prototrophic yeast deletion collection is sensitive to DNA alkylating agents streptozotocin and mechlorethamine (Lee et al., 2005). We confirmed sensitivity of tda5Δydl114wΔ to streptozoticin (data not shown). Treatment of fermenting yeast with DNA alkylating agents induces the production of reactive oxygen species (ROS) superoxide, hydrogen peroxide and hydroxyl radicals that inhibit respiration (Kitanovic et al., 2009). The sensitivity of the deletion mutant to the alkylating agent streptozoticin suggests that the precursors for the reaction catalyzed by Tda5 may convey an oxidative effect when accumulated in the tda5Δ mutant or that the product of the Tda5 reaction is necessary for mounting a stress response or antioxidant activity. In mammalian cells, streptozoticin treatment similarly is associated with production of ROS, oxidative stress and loss of mitochondrial membrane potential leading to apoptosis (Raza and John, 2012). Although the complete mechanism of streptozoticin cytotoxicity remains to be elucidated, glutathione redox enzymes such as glutathione-s- transferase are de-repressed as well as oxidative stress enzymes CYP1A1 and CYP1A2 during streptozoticin treatment (Nahdi et al., 2017).

4.2.5 Examination of yeast short chain dehydrogenases When yeast short chain dehydrogenases are compared by multiple sequence alignment as shown in Figure 23a, one can see that there is a high degree of divergence between the sequences, with only a handful of residues being fully conserved. These residues fall within the nucleotide binding site and active site and the otherwise low conservation implies the broad number of substrates accommodated by SDRs.

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Figure 23. Yeast short chain dehydrogenases display a conserved nucleotide binding domain and catalytic residues A) Multiple Sequence Alignment of S. cerevisiae SDRs. Sequences were downloaded from UniProt (UniProt Consortium, 2018), multiple sequence alignment carried out by MAFFT(Katoh et al., 2002) 6mer pairwise alignment algorithm, unreliably aligned positions were down-weighted using the GUIDANCE algorithm (Penn et al., 2010) over 100 bootstraps to generate alignment confidence scores, and the alignment was viewed in EMBL-MView (Chojnacki et al., 2017) B) Cladogram generated in EMBL-MView Simple Phylogeny multiple sequence alignment in (A) representing hypothetical evolutionary relationships between TDA5 and other yeast SDRs 102

4.2.6 Topology and localization of the Tda5 protein The primary sequence of Tda5 encodes a signal peptide. This signal peptide is needed for co- translational targeting to the endoplasmic reticulum, where it is folded and processed ahead of sorting. Hydropathy plotting shows Tda5 also encodes two N-terminal transmembrane domains for anchoring within cellular membranes. Several proteomic studies have reported that Tda5 is localized to the mitochondria (Sickmann, 2003) (Reinders, 2005), and the respiratory defect I identified for tda5Δ is consistent with mitochondrial dysfunction. In order to visualize localization of Tda5, I C-terminally tagged TDA5 with the dsRed fluorescent protein under the control of the GPD promoter using Gateway cloning (Alberti et al., 2007) and viewed by fluorescence microscopy as shown in Figure 24. In contrast to previous reports of mitochondrial localization, the fusion protein localizes largely to the endoplasmic reticulum and punctate structures within the plasma membrane. This discrepancy between the localization datasets is likely due to minor localization of Tda5 to the mitochondrial membrane through interaction of mitochondria and ER within the endomembrane system. This ER localization is consistent with a 2018 study that identified a co-translational interaction for the TDA5 transcript with an ER membrane protein complex that functions in promoting biogenesis of multipass proteins (Shurtleff and Weissman, 2018). There is a third transmembrane domain from 198-220 amino acids, overlapping with the proton acceptor Tda5 active site, implying a hydrophobic substrate. As shown in Figure 25 the transmembrane domains convey an integral membrane protein status with an overall topology of helix-loop-helix-loop, with the first loop containing the Rossman fold in the cytosolic position followed by a transmembrane helix containing active site residues followed by the second loop within the lumen, suggesting that Tda5 modifies lipids on the outside of the ER membrane. This positions the majority of the short chain dehydrogenase domain on the cytosolic side of the endomembrane system. The homologue of TDA5, AYR1 which functions in phospholipid biosynthesis as the acyl DHAP reductase is likewise localized to the ER lumen and plasma membrane and has the same membrane topology with active site residues overlapping with the transmembrane helix (Athenstaedt and Daum, 2000). The localization is also reminiscent of many of the ergosterol biosynthetic enzymes, Erg1, Erg11, Erg3, Erg4, Erg9, Erg26, Erg5 and Erg25 (Pathway Appendix I) (Huh et al., 2003). This localization must be confirmed by colocalization studies with nuclear ER, cortical ER and plasma membrane markers.

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Figure 24. Fluorescence microscopy shows Tda5 is localized to the plasma membrane and endoplasmic reticulum Gateway site specific recombination was used to clone TDA5 lacking a stop codon into pAG426GPD- dsRED vector to generate a dsRED fusion protein under the control of the constitutive strong promoter GPD, on a low copy CEN ARS plasmid backbone (Alberti et al., 2007).TDA5-dsRED plasmids were transformed into a wild-type his3Δ auxotrophic strain and selected for histidine prototrophy. Transformants were grown overnight in Synthetic Complete media lacking histidine, sub-cultured to an optical density of 0.1 and grown to an OD of 0.4. Cultures were mounted on glass slides and viewed under 100X magnification by spinning disk confocal fluorescence microscope set to 553 nm excitation wavelength and emission at 583 nm.

cytosol

lumen

Figure 25. Tda5 is an integral membrane short chain dehydrogenases/reductase

Topology of Tda5 within the endomembrane system was determined using Phobius and TMHMM webservers. (Käll et al., 2007; Krogh et al., 2001)

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4.2.7 How Tda5 transits from the ER to the mitochondria Mitochondria do not interact with the endomembrane system via vesicular fusion, but through the membrane contact sites (MCS) between the endoplasmic reticulum and mitochondria. Lipids and phospholipids along with integral membrane cargo are exchanged this way. There are two alternate contact sites between the ER and mitochondria provided by the ERMES complex and vCLAMP (Elbaz-Alon et al., 2014). Deleting vCLAMP leads to an increase in ERMES contact sites, and loss of ERMES leads to an expansion of vCLAMP contact sites, establishing that both mitochondrial contact sites are able to compensate for loss of one another (Elbaz-Alon et al., 2014). Deletion of vCLAMP in combination with ERMES is lethal. Tda5 is known to physically interact by pulldown with the Vam6/Vps39 core component of the vCLAMP (Elbaz-Alon et al., 2014). TDA5 has a synthetic negative genetic interaction with MDM10, which encodes the core component of the ERMES complex (Hoppins et al., 2011). Lipids are shared between the ER and mitochondria and are required for respiratory function. Mitochondrial-ER fusion is critical for maintaining mitochondrial lipid pools, although there is no known role for short chain dehydrogenases in this process. For example, phosphatidylserine is synthesized in the ER and transported to the mitochondria where it is converted to phosphatidylethanolamine and transported back to the ER for conversion to phosphatidylcholine. When vCLAMP and ERMES are hypomorphic, 3H-serine labelling shows accumulation of 3H-phosphatidylserine, which is restricted to the ER where it cannot be converted to phosphatidylethanolamine or phosphatidylcholine (Elbaz-Alon et al., 2014). Genetic interactions suggest that Tda5 may be exchanged at a MCS alongside lipids required for respiration.

4.2.8 Expression of Ydl114w rescues loss of function in Tda5 As previously discussed, the tda5Δ strain profiled in my screen did not display a growth defect and I subsequently observed that the freshly dissected strain suppressed very rapidly. A wealth of information on suppressors present in the yeast deletion collection has been generated by systematic analysis of secondary linkage groups identified by Synthetic Genetic Array studies carried out in the Boone and Andrews labs. These analyses identified the presence of a second linkage group in the tda5Δ strain that mapped to the HST1 locus (van Leeuwen et al., 2016). HST1 encodes the homologue of SIR2 protein, a histone deacetylase required for repression of middle-sporulation genes (Derbyshire, 1996). Hst1 functions within a well characterized complex with two other known proteins: the Rfm1 adaptor protein and the Sum1 DNA binding 105

protein shown in Figure 26a (Sutton, 2001) (Weber, 2008). The Hst1-Rfm1-Sum1 complex functions to repress middle-sporulation genes during vegetative growth. Since the strain from the prototrophic yeast deletion collection (Gibney et al., 2013; VanderSluis et al., 2014) was derived by mating and selection from the auxotrophic yeast deletion collection (Giaever et al., 2002; Winzeler et al., 1999) used in SGA studies (Tong et al., 2001) and due to the large growth defect of unsuppressed strains, the strain I profiled likely inherited the suppressor mutation that was present in the SGA suppressor studies. I verified this by sequencing the HST1 locus of the tda5∆ strain used in my metabolomic screen. As shown in Figure 26 d, the HST1 gene in the suppressed tda5Δ strain encodes a glycine to valine substitution that is predicted deleterious by SIFT (Ng and Henikoff, 2001). This loss of function mutation is consistent with destabilization of the Hst1-Sum1-Rfm1 histone deacetylase complex.

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Figure 26. Inactivation of Hst1-Sum1-Rfm1 results in expression of the SDR YDL114W which rescues the growth of tda5Δ A) Illustration of Sum1/Rfm1/Hst1 histone deacetylase complex role in control of meiotic transcription B) Induction of YDL114W transcript by deletion of HST1, SUM1 or RFM1 subunits (Lenstra et al., 2011) C) High copy expression of YDL114W rescues growth of tda5Δ on media containing glycerol D) HST1 locus of tda5Δ mutant carries D406V suppressor mutation For C, YDL114W was cloned into pVV214 high copy 2µ plasmid (van Mullem et al., 2003) modified with HygMX cassette encoding hygromycin resistance. The plasmid was transformed into tda5Δ/TDA5 heterozygous diploid and positive transformants were sporulated by shifting cultures from YPD to YPA to sporulation media without selection for plasmid while shaking at 30 °C. Tetrads were dissected onto YPD petri pates by micromanipulation using a yeast dissection microscope and allowed to germinate by incubating at 30 °C for 4 nights. Haplo-spore derived colonies were picked to YPD in a 96 well plate, grown overnight and spotted onto media containing YPD + hygromycin, YPD+G418, YPD and YP- glycerol. For D, The HST1 locus +/-500 bp from the suspected causal variant was amplified by PCR, and sequenced by Sanger sequencing. Sequences were downloaded, and translated to reveal that the HST1D406V mutation was present in the tda5Δ strain from the prototrophic yeast deletion collection.

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4.2.9 HST1 controls expression of middle sporulation genes In order to identify the mechanism of loss of function of HST1 in genetic suppression of tda5Δ, I investigated the role of HST1 in middle sporulation. To provide necessary context for HST1 function, I’ll first review the physiological changes that occur during stages of sporulation. In response to nutritional cues, absence of nitrogen source and presence of a non-fermentable carbon source, diploid yeast replicate DNA and undergo a sporulation developmental process whereby DNA is segregated into haploid nuclei within protective spores (Freese et al., 1982). Resultant spores are quiescent and poised for survival until conditions become favorable to germinate and begin mitosis. During sporulation, four daughter cells are partitioned within the mother cell. This process involves formation of entirely new cell membranes called pro-spore membranes (PSM) and cell walls surrounding each daughter cell while the mother cell is remodeled to form the ascus (reviewed in Neiman, 2011). Sporulation occurs in three developmental phases. Synthesis of chromosomes occurs in the first stage when cells exit mitotic G1 to enter the pre-meiotic S phase followed by recombination of homologs. Middle sporulation involves vast changes in the yeast cell as chromosomes are segregated and the cell prepares to form the PSM to contain all segregated cellular components from cytoplasmic swaths surrounding the four nuclei. Trafficking is also altered as vesicles are directed by Sec9/Spo20 SNAREs to the PSM rather than to the plasma membrane. In addition to the expression of the sporulation-specific SNARE Spo20, the lipid composition of the Golgi-derived vesicles is altered to prevent binding to the plasma membrane as expression of the PI4P kinase Mss4 for production of PI4,5P2 is necessary for Spo20 function (Coluccio et al., 2004). As yeast progress toward quiescence, from 6.5-8 hours post nutritional switch, sporulating cells maximize catabolism of carbohydrates, from 8-9 hours gluconeogenesis to glycogen is dampened and lipid synthesis is maintained as the only site of macromolecular biosynthesis (Ray and Ye, 2013). There is a second burst of lipid synthesis at 20 hours (Miller, 1963). Lipid synthesis supports growth of the PSM as it transitions from a flattened pouch shape as a function of the leading-edge protein complex and develops into a horseshoe-like ring around the spindle pole body and expands as a function of vesicle fusion to form cylinders. The space between the bell-shape layers is where spore wall is formed from β-glucan, chitosan and dityrosine coatings (Coluccio et al., 2004). The leading edge complex consists of Ssp1, Don1 and Ady3 proteins, where Ssp1 binds inositol phospholipids and the complex is localized to the growing edge of the pro-spore membrane

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(Moreno-Borchart et al., 2001). The bell-shape membrane curvature maintained during sporulation is governed by GPI-anchored proteins such as Spo19 in the spore membrane bending pathway (Maier et al., 2008). GPI-anchored proteins are incorporated into the cell wall by covalent interaction with glycosyl chains. Several GPI-linked membrane proteins and cell wall proteins are expressed during sporulation. In the second stage of sporulation, chromosomes are segregated by karyokinesis and the nuclei are enveloped by PSM cylinders that ultimately enclose the nuclei. In the third stage of sporulation, a thick protective coat is formed around the ascus. Metabolic changes induced during yeast sporulation have been studied, however questions surrounding re-localization of enzymes during sporulation remain as cortical ER is absorbed into the nuclear envelope (Miller, 1963; Ray and Ye, 2013; Weidberg et al., 2016). Overall sporulation represents an alternate developmental state where major physiological changes occur, particularly in the membrane component of the cell.

The first phase of sporulation is governed by expression of Ime1, the master regulator of meiosis, which binds the Ume6 transcriptional regulator, preventing its interaction with a histone deacetylase complex, leading to de-repression and expression of sporulation phase 1 genomic loci (Mitchell, 1994) (Washburn and Esposito, 2001). Genes that allow for expression of middle sporulation genes, such as NDT80 encoding the meiosis specific transcription factor, are induced alongside genes required for premeiotic S phase, recombination and prophase (Primig et al., 2000). The NDT80 locus is repressed by the Hst1- Sum1-Rfm1 histone deacetylase complex and Ime1 expression weakens the Ume6-Sum1 interaction leading to NDT80 expression (Xie et al., 1999). Expression of the Ndt80 transcription factor initiates the second phase of sporulation. This phase of sporulation involves formation of the spindle pole body, microtubule nucleation, and formation of the meiosis II outer plaque. One enzyme essential for middle sporulation is phospholipase D, which hydrolyses phosphatidylcholine to choline and phosphatidic acid (Rose et al., 1995). The newly formed phosphatidic acid is required for the Spo20 SNARE protein selectivity during membrane fusion at the spindle pole body (Nakanishi et al., 2004).

Temporal control of the sporulation process involves initiating several transcriptional programs including proteins for meiotic recombination and proteins that comprise sporulation specific complexes alongside induction and repression of hundreds of genes (Chu et al., 1998; Primig et al., 2000). In the context of hst1Δ, rfm1Δ, or sum1Δ histone deacetylase single mutation mutants,

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hundreds of transcripts coherently change in abundance. When hst1Δ, rfm1Δ or sum1Δ is deleted, NDT80 targets are upregulated including an uncharacterized SDR enzyme YDL114W, as shown in Figure 26 b. This target is of particular interest as YDL114W is the uncharacterized paralogue of TDA5. According to the Yeast Gene Orthology Browser for examination of syntenic relationships due to whole genome duplication, TDA5 and its sporulation specific paralog YDL114W are not whole genome duplicates (Byrne and Wolfe, 2005). The YDL114W gene contains a Ndt80 middle sporulation element (5′- DNCRCAAAW-3′) in its promoter located 112 bp upstream of the start codon (Chu and Herskowitz, 1998). Ydl114w shares 68% similarity with Tda5 over 258 amino acids, 25.8% identity and is also a member of the conserved domain 05339 cluster of short chain dehydrogenases. YDL114W does not contain a signal peptide or an N- terminal transmembrane domain, which is explained by re-localization of ER enzymes and metabolic processes during meiosis. The LoQate database lists observations of cytosolic expression for C terminally tagged Ydl114w-GFP and N terminally tagged Ydl114w-mCherry (Breker et al., 2014). Since YDL114W does not encode transmembrane helices or a signal peptide, it is likely not an integral membrane protein and not targeted to the in the endomembrane system. During vegetative growth, YDL114W expression is repressed. When the fresh ydl114wΔ/YDL114W heterozygous diploid was sporulated and dissected, I observed that the ydl114wΔ deletion strains display a wild-type growth rate and no deviations from wild-type metabolite levels were identified by comparison of ydl114wΔ to wild-type by untargeted metabolomics.

In order to determine whether YDL114W could compensate for TDA5, I expressed YDL114W on a high copy plasmid pVV214-HygMX (van Mullem et al., 2003, modified to contain HygMX by Peter Xu), transformed the plasmid into a back-crossed suppressed tda5∆/TDA5 heterozygous diploid, sporulated and dissected spores. I monitored growth patterns and patterns of inheritance of the plasmid, which is marked by hygromycin resistance. As shown in Figure 26 b, I found that 100% of tda5∆ spores carrying the YDL114W plasmid were able to grow on glycerol, consistent with the TDA5 control, whereas only 60% of tda5Δ strains without the plasmid were able to grow on glycerol. The high proportion of tda5Δ strains without the plasmid that have intact respiratory metabolism (60%) verifies the presence of a suppressor mutation segregating in the cross, and the frequency of greater than 50% suggests the presence of additional newly selected

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second site suppressors. Remarkably, the ability of YDL114W to increase the fraction of haplo- spores’ ability to grow on glycerol to 100% indicates that YDL114W is able to rescue the growth defect and respiratory capacity of tda5Δ mutants and is providing a compensatory function. Since YDL114W is de-repressed in the hst1 loss of function mutant, the mechanism of suppression of tda5∆ is through upregulation of its sporulation-specific paralog, YDL114W, which functionally complements Tda5.

4.2.10 Metabolomic phenotyping of tda5Δ strains In an attempt to identify the reaction catalyzed by Tda5, I compared the metabolomes of wild- type cells to cells from freshly dissected tda5Δ strains and suppressed tda5Δ strains by LC-MS metabolomic analysis. When untargeted metabolomic analysis was carried out on replicate freshly dissected tda5Δ mutant strains compared to wild-type, over 100 mass spectral features showed greater than 2-fold difference in levels as shown in Figure 27 d. When targeted metabolite analysis was carried out, it was determined that multiple metabolic pathways were affected such as nucleotide levels, pentose-phosphate pathway intermediates and amino acid biosynthesis. As shown in Figure 27 a, I observed an accumulation of homocitrate in the freshly dissected tda5Δ strains compared to wild-type, which is the last metabolite in the lysine biosynthetic pathway that is produced within the cytoplasm and this accumulation is diminished in the tda5Δ suppressed strain. Accumulations of metabolites upstream of mitochondrial metabolic pathway steps in hand with depletions of downstream mitochondrially produced metabolites indicate an effect of compartmentalization due to diminished mitochondrial function. The large number of changes in metabolite levels reflects the difference in growth rate of the cultures and pleiotropic effects of the tda5Δ lesion. This analysis did not pinpoint a substrate for Tda5.

In order to identify changes in metabolite levels accompanied by suppression of tda5Δ, I carried out targeted metabolomics on suppressed strains. I determined levels of more than 100 known metabolites, normalized to wild-type to determine fold-change and carried out hierarchical clustering to determine patterns in observed metabolite level changes. Amongst genes and metabolites upregulated during middle sporulation are the BNA2, BNA4, BNA5, BNA6 and BNA1 genes involved in de novo production of NAD+ from tryptophan as displayed in the kynurenine biosynthetic pathway in Figure 27 c. These genes (highlighted in pink) are Ndt80 targets and

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induction of these five genes explains the accumulations of kynurenine pathway intermediates observed for the tda5∆ mutant in my original metabolomic screen for enzyme discovery. When the tda5∆ suppressed strains are compared to the hst1∆, rfm1∆, and sum1∆ single deletion mutants, the accumulations across kynurenine metabolism and aromatic amino acids are shared, indicating that Ndt80 is activated in these suppressed strains. Although the magnitude of the increases in kynurenine pathway intermediates were consistent across hst1∆, rfm1∆ and sum1∆, the magnitudes of phenotypes in the suppressed tda5∆ mutants were lower. This likely indicates only partial rescue for loss of function in TDA5 can be achieved by de-repression of YDL114W and although the growth rate is restored, there are residual functions distinct to TDA5 that manifest in altered levels of other metabolites only in tda5∆ mutant strains. An alternate explanation is that hst1 is only partial loss of function and that YDL114W expression is not fully able to compensate for loss of TDA5. Specifically, I observed decreases in steady state levels for amino acids requiring biosynthesis within the mitochondrial compartment: arginine, lysine, and their precursors. When these metabolites are observed in suppressed strains there is an apparent change in these mitochondrially synthesized metabolites as compared to wild type cells. These phenotypes demonstrate a clear effect of compartmentalization of amino acid biosynthesis is observed due to diminished mitochondrial function. These tda5Δ suppressed strains have a wild- type growth rate on fermentable media, indicating that although the respiratory growth defect is rescued, suppressed strains harbor major defects in mitochondrial compartmentalized metabolism. Although the metabolic profiling of suppressed strains is valuable in identifying the origin of the numerous metabolic phenotypes identified in the tda5Δ strain in the screen, the changes in metabolite levels do not explain the mechanism(s) of suppression of rescuing tda5Δ.

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C D

value)

log10(p -

log2(fold-change)

Figure 27. Metabolomic analysis of tda5Δ and tda5Δ suppressed strains to wild-type A) Homocitrate accumulates in fresh tda5Δ mutants to a much greater extent (blue line) than suppressed mutants (red lines) and wild-type cells (black lines). Chromatographic traces are shown. B) Snapshot of metabolite level changes in suppressed tda5Δ strains compared to histone deacetylase complex mutants relative to wild-type. C) Transcripts repressed by Hst1/Rfm1/Sum1 include kynurenine pathway enzymes (shown in pink), which are upregulated in deletion mutants leading to metabolite level accumulations, D) Volcano plot displaying fold-change and t-test p-values for differences in wildtype and tda5Δ mutant means for 4000 mass spectral features For A, suppressed tda5Δ strains, wild-type controls and 5 newly dissected tda5Δ strains that were unable to grow on glycerol were grown in YNB-glucose and extracted at an OD600 of ~0.5. Metabolites were separated and measured by LC-MS.Targeted analysis of chromatographic peaks corresponding to retention times and m/z for chemical standards were extracted and integrated in Agilent MassHunter Profinder. Metabolite levels are normalized to corresponding peaks derived 13C 15N internal reference. For B, means of five independent biological replicates where reference normalized integrated intensities were normalized to Wild-type levels, values log2 transformed, clustered by centroid linkage, displayed as a heatmap. For D, wildtype and newly dissected tda5Δ strain metabolite extracts were prepared as in A. Untargeted analysis of chromatographic peaks was carried out in Agilent Profinder, significantly altered mass spectral features were determined using custom R scripts.

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4.2.11 Identification of YDL114W independent suppressors of tda5∆ deletion mutants In order to determine whether the sole mechanism of suppression of tda5Δ was through de- repression of YDL114W, I crossed tda5∆ and ydl114w∆ single mutants and, sporulated and dissected the resulting diploid to obtain the tda5∆ ydl114w∆ double deletion mutant. When a newly dissected double mutant culture was plated on rich media, I observed that the tda5∆ ydl114w∆ double deletion mutant has a vastly reduced suppressor frequency of 0.39%, which is 3.5% of tda5Δ single mutant cultures that generate ~11% suppressors. This high frequency may be due to increased levels of oxidative-stress induced DNA damage that surpasses DNA repair mechanisms as has been observed in screens for suppressors of mutation accumulation (Huang et al., 2003). Together with my metabolite analysis of tda5Δ suppressed mutants, the results show that the majority of suppressors rely on a Ydl114w-dependent mechanism. Loss of function mutations in HST1, RFM1, or SUM1 will provide this type of suppression, providing a wide mutational target. This result also shows that there are mechanisms of suppression of loss of function in TDA5 that are independent of YDL114W, and I have mapped several such suppressors (described below).

Figure 28 The frequency of suppressors is vastly reduced in the tda5Δydl114wΔ double deletion mutant A) Colonies derived from freshly dissected tda5Δ spore-derived culture is spread onto YPD B) Colonies derived from freshly dissected tda5Δydl114wΔ spore-derived culture is spread onto YPD For A and B, 4 independent colonies from newly dissected tda5Δ and tda5Δydl114wΔ strains were grown in YPD to mid-log phase and serial dilutions were plated onto YPD, incubated at 30 °C for 2 nights, suppressed and unsuppressed colonies were enumerated and averaged to calculate rates of suppression. Representative images of equal density tda5Δ and tda5Δydl114wΔ strains are shown.

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4.2.12 Mapping suppressors of tda5∆ loss of function independent of YDL114W In order to shed light on mechanisms through which cells can compensate for loss of function of TDA5, I selected second site haploid suppressors of the tda5∆ growth defect on YP-glycerol that were independent of YDL114W. I took two approaches to generating these suppressors, firstly by isolating suppressors of the tda5Δ single mutant and checking for allelism to rfm1Δ, sum1Δ and hst1Δ by crossing to the tda5Δ/rfm1Δ, tda5Δ/sum1Δ and tda5Δ/hst1Δ double deletion mutants and measuring ability to grow on YP-glycerol. In order to determine dominance of the trait, each suppressed mutant was backcrossed to form a tda5Δ/tda5Δ diploid heterozygous for the suppressor mutation and assessed for growth on glycerol, all suppressors were recessive. Since loss of function mutations are recessive, only allelic inactivation would result in a diploid able to respire. I also directly selected second site suppressors of the tda5∆ ydl114w∆ double deletion mutant strain on YP-glycerol.

In order to identify the causal locus of YDL114W independent suppressor mutations, I used a bulk-segregant approach where I measured allele frequency of variants in pools of segregants selected for the phenotype of interest to enrich for the causal variant (Brauer et al., 2006; Dunham, 2012). I back-crossed the suppressed strains to ydl114w∆, sporulated, dissected progeny, ensured the segregation of the tda5Δ growth defect was consistent with the presence of a single unlinked suppressor mutation, and phenotyped for each of the markers and ability to grow on respiratory carbon sources. I identified tda5∆ ydl114w∆ spores that were able to grow on glycerol as positive for the suppressor mutation and tda5∆ ydl114w∆ spores that were unable to grow on glycerol as the negative control group. I pooled cultures by equal numbers of cells, extracted genomic DNA and sequenced by Illumina Hi-Seq to identify variant alleles and determine their allele frequency. Variants that were consistent with linkage to the suppressor were considered for follow-up if the variant was present in ~100% of mutants able to grow on glycerol and the wild type allele was present in ~100% of the mutants unable to grow on glycerol as exemplified in Figure 29. Furthermore, the candidate causal suppressor variants were verified as not present in the ydl114w∆ parent used for back-crossing. Three genomic variants were identified as linked to suppression of tda5Δ where two resulted in non-synonymous coding changes to the amino acid sequence of the encoded protein, whereas one variant caused a frameshift near the 3’ terminus of the open reading frame. All three of these variants are

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predicted deleterious by SIFT (Ng and Henikoff, 2001) and thereby suggest loss of function, which agrees with the recessive nature of the phenotype.

Wildtype ERG11 tda5Δ erg11

Figure 29 Illustration of process for identifying causative mutations in tda5 suppressor strains. Erg11 S382F variant identified as causal allele in suppression of tda5∆ Whole genome sequencing read pileups for ERG11/YHR007C showing 89% allele frequency of C->T transition encoding S->F substitution Each individual suppressed tda5Δ::hygMX strain was back-crossed to MATa TDA5 ydl114wΔ::natMX his3Δ ura3Δ, sporulated, dissected, phenotyped for ClonNAT and hygromycin resistance, and ability to grow on glycerol. Haplospore derived colonies that were ClonNAT resistant, hygromycin resistant, able to grow on glycerol, and wild type size on YPD were considered tda5Δ suppressed, while haplospore derived colonies that were ClonNAT resistant, hygromycin resistant, unable to grow on glycerol and slow growing on YPD were considered tda5Δ unsuppressed. Six to twelve independent spore-derived colonies were selected for each suppressed and unsuppressed group, and individually grown in YPD, equal numbers of cells for each spore-derived colony were pooled and genomic DNA extracted. Sequencing libraries were prepared using transposase-mediated tagmentation according to Nextera, Illumina indexing barcodes and sequences were added for compatibility with Illumina Hi-Seq and libraries were normalized on beads. Illumina Hi-Seq short read platform was used to sequence libraries. Raw reads were processed and mapped to Saccharomyces cerevisiae reference genome and variants were called. Variant alleles were viewed in Interactive Genome Viewer (Broad). As shown in Table 4, two of the causal variants identified occur within enzymes from the ergosterol biosynthetic pathway. These are a frameshift leading to truncation of Erg1, the squalene epoxidase (Jandrositz et al., 1991), and a S382F substitution in Erg11, the lanosterol demethylase (Karst and Lacroute, 1977). Both of these enzymes are essential on media lacking sterols. The recessive nature of the suppressor mutations in hand with the essentiality of these two genes indicates that these strains are hypomorphic for erg1 and erg11.

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Table 4 Recessive mutations in sterol biosynthesis suppress loss of function in TDA5 LOCUS FUNCTION VARIANT ERG1 squalene epoxidase Deletion ΔT1469, Frameshift causing truncation ERG11 lanosterol demethylase Substitution, Non-synonymous coding S382F ERG12 mevalonate kinase Substitution, Non-synonymous coding G392D UPC2 transcriptional activator of ergosterol Substitution, Non-synonymous coding biosynthetic genes R855L 4.2.13 Loss of function mutations in ergosterol biosynthesis suppress tda5Δ To test whether mutations upstream of squalene could suppress loss of function in TDA5, I checked to see if the Erg12 G392D mevalonate kinase hypomorph I had previously identified and characterized could suppress tda5Δ. Indeed, the segregation pattern of growth for spores derived from the tda5Δ/TDA5 ERG12/erg12 heterozygous diploid indicated the presence of a single suppressive mutation. When suppressed tda5Δ strains were pooled and sequenced at the ERG12 locus, the erg12G392D mutation was present at 100% allele frequency, consistent with linkage to the suppressive phenotype. Together, these results indicate that Erg12 hypomorphs could suppress loss of function in Tda5, showing that lesions in ergosterol biosynthesis upstream of squalene synthesis in the mevalonate pathway were capable of suppressing loss of function in TDA5.

Whole genome sequencing on suppressed tda5Δ strains led to the identification of another suppressor that affects the ergosterol biosynthetic pathway. The causal variant was a mutation in the UPC2 gene encoding R855L that had 0% allele frequency in unsuppressed pools and 100% allele frequency in suppressed pools. UPC2 is a sterol regulatory binding protein that acts as a negative transcriptional regulator of ergosterol biosynthetic genes in response to exogenous sterols (Davies et al., 2005). The R855L variant lies within the sterol ligand binding domain of Upc2 and the variant is expected to be destabilizing by SIFT (Ng and Henikoff, 2001), consistent with the recessive nature of the allele. Accordingly, when changes in expression due to alteration of Upc2 function are mined using Yeastract, Upc2 is identified as a positive regulator of 13 ergosterol biosynthetic enzymes which are highlighted in blue in Figure 30 (Teixeira et al., 2018). The R855L variant likely leads to the partial loss of function of Upc2 and to disruption of correct ergosterol pathway regulation resulting in compromised synthesis and attenuated levels

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of sterols. Upc2 is also a negative regulator of ICT1 encoding the lysophosphatidic acid acyltransferase (Reimand et al., 2010) and a negative regulator of INO1, encoding the inositol-3- phosphate synthase (Chua et al., 2006) involved in phospholipid biosynthesis. Since the suppressor mutants were selected on rich media that was not supplemented with sterols, the upc2 mutant is hypomorphic for de novo biosynthesis of sterols and also experiences defects in phospholipid biosynthesis.

4.2.14 Large scale genetic interaction datasets reveal additional candidate positive genetic interactions with tda5Δ A window into the landscape of mutations benefitting tda5Δ mutants has been opened since the publication of genetic interaction data across essential and nonessential yeast genes from the Boone and Andrews’ labs (Costanzo, 2016). Positive genetic interaction data generated using temperature sensitive alleles has provided an additional list of essential genes involved in ergosterol biosynthesis that when compromised for function at the restrictive temperature are able to contribute fitness benefits in combination with lesions in tda5Δ. As shown in Figure 30, the list of positive genetic interactors with tda5Δ includes: erg8, erg9, erg13, and mvd1. Since the suppressed tda5Δ strain is used in this case, the enrichment for positive interactions with lesions in ergosterol biosynthesis indicates that although YDL114W is de-repressed in the suppressed tda5Δ mutant in the deletion set, a further fitness boost is seen when combined with lesions in ergosterol biosynthesis. Furthermore, fitness is determined on synthetic media, which precludes uptake of sterols as part of the suppressive mechanism. It is important to note that these genetic interactions are private to the tda5Δ deletion strain; these interactions are not observed in hst1Δ, rfm1Δ, or sum1Δ deletions. The growth benefit attributed to decreased function in ERG13 is of particular importance, because it is upstream of HMG-CoA in the mevalonate pathway. ERG13 encodes the 3-hydroxy-methylglutaryl coenzyme A synthase, which produces HMG-CoA from acetyl-CoA and acetoacetyl-CoA (Servouse, 1984). The ability of these temperature-sensitive alleles to rescue loss of function of the tda5∆ strain indicates that metabolites upstream of Erg13 such as acetyl-CoA may be involved in mediating metabolic compensation for loss of Tda5.

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Figure 30 The tda5∆ mutant has positive genetic interactions with hypomorphs in ergosterol biosynthesis Ergosterol biosynthetic pathway where mapped suppressors identified in this study are highlighted in green, deletion mutants identified as having positive negative genetic interactions with tda5Δ by synthetic genetic array studies are indicated in pink interactions with stringent cut-off score>0.16 and p-value<0.05 (Costanzo et al., 2016), genes that are positively regulated by Upc2 are in blue.

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Defects in ergosterol biosynthesis cause deficiencies across a wide range of cellular systems. Ergosterol is essential for membrane organization of proteins throughout the secretory pathway and for membrane recognition during vacuolar fusion. Ergosterol biosynthesis mutants also have fragmented vacuoles and reduced vacuolar fusion due to defects in Sec17 release in the priming step which is rescued by ergosterol supplementation (Kato and Wickner, 2001). During homotypic vacuolar fusion, ergosterol diacylglycerol, 3-phosphoinositides and 4- phosphoinositides are interdependently required as regulatory lipids.

In addition to vacuolar fusion, ergosterol homeostasis is further required for maintenance of mitochondrial morphology. Temperature sensitive mutants in ergosterol biosynthesis (Erg1, Erg7, Erg8, Erg10, Erg12, Erg13, Erg25, Erg26, Erg27, Mvd1, Ncp1) have a swollen and clumped mitochondrial morphological defect (Altmann and Westermann, 2005). Integration of mitochondrial outer membrane proteins is dependent on membrane fluidity and low ergosterol content, underscoring that correct localization of lipids is critical for mitochondrial function (Kato and Wickner, 2001).

4.2.15 Negative genetic interactions of mitochondrial proteins with tda5∆ In order to shed light on the role of Tda5 within the mitochondria, I analyzed a mitochondria- specific genetic interaction dataset. This array included 1482 strains comprising ~600 proteins with annotated localizations to the mitochondria, ~400 in the early secretory pathway, ~200 hypomorphic DAmP alleles and genes across diverse cellular processes including metabolic functions (Hoppins et al., 2011). The tda5Δ strain had synthetic negative genetic interactions with deletions in genes that affect Golgi transport and morphology: vps52Δ, rgp1Δ, dop1Δ and pcp1Δ. The cho2Δ mutant was also identified as negatively affecting growth rate in combination with tda5Δ, CHO2 encodes the phosphatidylethanolamine methyltransferase for phosphatidylethanolamine to phosphatidylcholine production (Summers et al., 1988). As previously mentioned, the mdm10Δ mutant was also identified for its negative genetic interaction with tda5Δ. Mdm10 is a ERMES complex subunit. The mitochondrial lipid profile of mdm10Δ single deletion mutant showed that mitochondrial lipids phosphatidylethanolamine and cardiolipin levels were diminished while phosphatidylserine accumulated along with a 2-fold increase in ergosterol (Tan et al., 2013). This shows that the balance of ergosterol and other

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lipids are affected by the loss of the ERMES complex and suggests loss of Tda5 exacerbates lipid defects leading to a diminished growth rate.

4.2.16 Chemical suppression of tda5Δ Just as second site mutations in the ergosterol biosynthetic pathway are capable of rescuing the growth defect of tda5Δ, I aimed to determine whether rescue could be effected by chemical suppression. Therefore, I grew strains in the presence of sub-inhibitory 5 ng/mL miconazole targeting the Erg11 lanosterol demethylase and 10 µg/mL mevinolin targeting HMG-CoA reductases Hmg1 and Hmg2. From spot assays shown in Figure 31, a slight growth benefit is conveyed in the presence of inhibitors for the tda5Δ strain compared to wild-type, tda5Δhst1Δ and hst1Δ controls. In contrast, cells that expressed YDL114W from a galactose inducible plasmid during pre-growth are fully rescued by 5 ng/mL miconazole when plated on the YDL114W repressible condition (YPD). Separate experiments with a growth inhibitory concentration of 10 ng/mL miconazole led to inhibition of growth for all strains except the tda5Δydl114wΔ strain. These results clearly indicate that inhibiting Erg11 rescues tda5Δ.

Figure 31 Stronger growth is observed for tda5Δ strains grown in the presence of ergosterol pathway inhibitors Overnight cultures of strains grown in YPD or YP-galactose were normalized to an optical density of 1 and 10-fold serial dilutions were prepared in a 96 well plate in 200 µL volume. For spotting, 3 µL was transferred using a Rainin P20 multi-channel pipette. Cells were mixed using a P200 multi-channel between spotting. Spot plates were allowed to dry at room temperature before incubating 2 nights at 30 °C. Images were captured using AlphaImager gel documentation system with white light transillumination.

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Toxicity of miconazole is due to depletion of sterols, accumulation of toxic sterol analogues and ensuing deleterious effects on cellular membranes, such as inhibition of vacuolar ATPases (Martel et al., 2010). In contrast to the clear rescue achieved by mevinolin and miconazole, the tda5Δydl114wΔ strain is sensitive to fenpropimorph, the morpholine inhibitor that targets two steps of the ergosterol pathway: Erg24 the C-14 sterol reductase which is the subsequent step after lanosterol demethylase and Erg2 sterol isomerase which converts fecosterol to episterol (Marcireau et al., 1990). Fenpropimorph inhibition leads to accumulation of ergosta-8,22-dienol , 4,4-dimethyl-cholesta8,14-dienol, 4,4-dimethyl-cholesta-8,24-dienol, and ignosterol (Baloch and Mercer, 1987). Although fenpropimorph, miconazole and mevinolin target the same biosynthetic pathway, these drugs have non-overlapping profiles in genome wide drug sensitivity screens and act synergistically when combined (Smith, 2011).

In order to assess whether inhibition of other lipid pathways could lead to suppression of tda5Δ, I tested for resistance to myriocin, the serine palmitoyl transferase inhibitor. As shown in Figure 32, the strain was sensitive to 1.25 mM myriocin indicating that Tda5 does not act downstream of Tsc10 in production of long chain ceramides.

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Figure 32. Cells are sensitive to fenpropimorph and myriocin Overnight cultures of strains grown in YPD or YP-galactose were normalized to an optical density of 1 and 10-fold serial dilutions were prepared in a 96 well plate in 200 µL volume. For spotting, 3 µL was transferred using a Rainin P20 multi-channel pipette. Cells were mixed using a P200 multi-channel between spotting. Spot plates were allowed to dry at room temperature before incubating 2 nights at 30 °C. Images were captured using AlphaImager gel documentation system with white light transillumination.

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4.2.17 Strains deleted for tda5Δ have a membrane defect In dissecting the tda5Δ/TDA5 tetrads, I found that the asci were sensitive to digestion with β- glucuronidase which cleaves cell wall sugars, which may indicate a cell wall or membrane structural integrity defect. In addition, I found that when either TDA5 or YDL114W is overexpressed, these cells became resistant to β-glucuronidase digestion. As suppressor mutations in ergosterol biosynthesis indicate a likely role in membrane function, I tested the tda5Δ mutant response to osmotic and detergent stress. Spot assays shown in Figure 33 reveal a sensitivity to osmotic stress and a slight sensitivity to SDS. Together these phenotypes indicate a membrane defect for tda5Δ. Membrane defects are surprisingly rare with various single mutants in lipid metabolism as compensation for lesions in fatty acid, sterol, sphingolipid, phospholipid and triacyl-glycerides by upregulation of the remaining pathways (Daum et al., 1999).

Figure 33 Detection of a membrane defect for tda5Δ Overnight cultures of strains grown in YPD or YP-galactose were normalized to an optical density of 1 and 10-fold serial dilutions were prepared in a 96 well plate in 200 µL volume. For spotting, 3 µL was transferred using a Rainin P20 multi-channel pipette. Cells were mixed using a P200 multi-channel between spotting. Spot plates were allowed to dry at room temperature before incubating 2 nights at 30 °C. Images were captured using AlphaImager gel documentation system with white light transillumination.

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4.2.18 The growth defect of tda5Δ can be rescued by N-acetyl-cysteine. As membrane structural integrity defects are often due to oxidative damage, we spotted strains onto N-acetyl-cysteine to determine if we could reverse the growth defect due to the tda5Δ lesion. As shown in Figure 34a, to our surprise, N-acetyl-cysteine restored a wild-type growth level for the tda5Δydl114wΔ strain as well as the tda5Δ strain conditionally repressed for YDL114W. These data indicate that tda5Δ may be slow growing due to increased oxidative stress which may have many sources: the oxidative effect of accumulating lipid precursor and ensuing ROS production, ROS production due to lack of Tda5 function indicating the product of Tda5 reaction acts to protect against ROS, or compromised membrane integrity due to ROS production that cannot be rescued by endogenous levels of antioxidants. As shown in Figure 34b, alleviation of mitochondrial debilitation by antioxidants would rescue the petite phenotype observed for tda5Δ as discussed below.

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Figure 34 N-acetyl cysteine rescues growth defect of tda5Δ mutant A) Spot assays for tda5Δydl114wΔ and control strains on YPD and N-acetyl-cysteine B) Model for attenuation of lipotoxicity by N-acetyl-cysteine Overnight cultures of strains grown in YPD or YP-galactose were normalized to an optical density of 1 and 10-fold serial dilutions were prepared in a 96 well plate in 200 µL volume. For spotting, 3 µL was transferred using a Rainin P20 multi-channel pipette. Cells were mixed using a P200 multi-channel between spotting. Spot plates were allowed to dry at room temperature before incubating 2 nights at 30 °C. Images were captured using AlphaImager gel documentation system with white light transillumination.

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4.2.19 Defects in lipid homeostasis and mitochondrial activity cause ROS formation Lipid homeostasis is critical in the response to oxidative stress. Deletion of ISC1, the enzyme responsible for hydrolysis of phosphatidylcholine to phosphorylcholine and diacylglycerol, leads to an increase in oxidative stress that is mediated by lipid hydroperoxide (Dickson, 2010). Mitochondrial ETC complex III is the major origin of ROS in the cell as free radical oxygen or superoxide ions are formed and lead to formation of hydroxyl radicals, which can damage lipids and other cellular components. Mitochondrial activity is essential to protection against oxidative stress as petite cells cannot mount an adaptive response to peroxide likely by not affording the extra energy required for upregulating superoxide dismutase and peroxidases (Grant et al., 1997). Defects in mitochondrial lipid homeostasis, such as deletion of CRD1 encoding cardiolipin synthase or TAZ1 encoding lyso-phosphatidylcholine acyltransferase results in inability to synthesize or remodel cardiolipin alongside increased protein carbonylation, a hallmark of increased ROS. The inner mitochondrial membrane component cardiolipin supports electron transport chain stabilization by promoting interaction between respiratory complexes and prevents excess ROS production (Chen et al., 2008). Exogenous oleic acid incorporation into lipids can rescue phosphatidylcholine and phosphatidylethanolamine levels in taz1Δ mutants (Chen et al., 2008).

4.2.20 Measurement of acyl-CoA pools in fresh and suppressed tda5∆ mutants informs on routes of metabolic compensation Mapped suppressors of tda5Δ include various mutations expected to compromise ergosterol and lipid biosynthesis that rescue a growth defect that is associated with oxidative stress in the tda5Δ mutant, thereby implicating the involvement of lipid homeostasis in compensation for the loss of tda5Δ. Since acetyl-CoA is a central input for both sterols and other lipid biosynthetic pathways, I hypothesize that alterations in CoA pools alleviate loss of function in Tda5, and that suppressed tda5∆ will harbor alterations in acetyl-CoA and malonyl-CoA pools where malonyl-CoA would serve as a sentinel for fatty acid biosynthesis. Measurement of CoA pools will allow me to pinpoint blockages in fatty acid biosynthesis and ergosterol pathways in the unsuppressed mutant and determine compensation via other metabolic pathways.

In order to achieve this, I optimized a method to purify acyl-CoA metabolites in yeast by solid phase extraction (SPE) for downstream measurement by LC-MS (Snyder et al,. 2015). When

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strains carrying erg12, upc2 and erg11 suppressor mutations in the TDA5 context and in the tda5Δ context were compared to wild-type, several patterns emerge shown in Figure 35. We observe reduced HMG-CoA in the tda5Δerg12 mutant and elevated HMG-CoA levels in TDA5 erg12 mutants. The increase observed in the TDA5 erg12 mutant is expected as the strains have 100-fold accumulation of mevalonate over wild-type and as mevalonate is directly downstream of HMG-CoA. The decrease in HMG-CoA in the tda5Δerg12 may indicate increased demand for upstream precursors by pathways that compensate for tda5Δ. A decrease in acetyl-CoA is likewise observed the tda5Δ erg12 strain, and may accordingly reflect increased demand for the metabolite. Malonyl-CoA is at a lower level in the TDA5 strain than tda5Δ, which may reflect an increased demand for fatty acid biosynthesis when erg12 is hypomorphic. When we observe CoA levels for the upc2 mutant in the context of tda5Δ, acetyl-CoA levels are slightly lower than wild-type, however HMG-CoA levels are reduced to 25% of wild-type in the tda5Δ deletion strains. The change in HMG-CoA is likely due to lowered expression of enzymes upstream of HMG-CoA that are controlled by Upc2, such as Erg10, the acetoacetyl-CoA thiolase (Davies and Rine, 2006). When CoA metabolites are measured for the erg11 variant, we observe also lower acetyl-CoA levels in the tda5Δ context compared to wild-type and TDA5erg11, while HMG-CoA levels are lower across both TDA5erg11 and tda5Δerg11 contexts as compared to wild-type. From this analysis, acetyl-CoA, HMG-CoA and malonyl-CoA levels alone do not fully explain metabolic compensation for loss of function in TDA5. In both TDA5erg11 and tda5Δ erg11 suppressed mutants, both HMG-CoA and acetyl-CoA levels are much lower than wild-type and may indicate re-routing of acetyl-CoA into another metabolic pathway. Alternatively, acetyl- CoA levels could be lower due to loss function in erg11. Stable isotope tracing using 13C-glucose could resolve dynamics using flux measurements to resolve these two models. I also carried out targeted analysis of over 100 known compounds and did not observe coherent changes in metabolite levels in soluble metabolites between mutants that would indicate a common mode of rescue.

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A Figure 35 Acetyl-CoA, HMG-CoA and Malonyl- CoA levels across ergosterol hypomorphs in combination with tda5Δ

A) Acetyl-CoA levels normalized to wild-type B) HMG-CoA levels normalized to wild-type C) Malonyl-CoA levels normalized to wild-type

Back-crossed strains carrying each suppressor mutation in both TDA5 and tda5Δ contexts were identified by phenotyping dissected tetrads. Cells were grown in YNB-glucose + histidine through B several rounds of growth and sub-culture to a final optical density of 0.7. For each biological replicate, cells were harvested by centrifugation and extracted by adding 1.2 mL ACN/IPA 3:1, followed by three rounds of freeze-thaw. Samples were centrifuged, and 125 µL acetic acid was added and separated and eluted according to (Snyder et al., 2015). Eluates were dried under N2, reconstituted in a volume of water according to optical density at time of extraction. 15N purified CoA reference from yeast was added. Metabolites were separated and measured by reverse-phase ion-paired chromatography and detected by Q-TOF. Targeted analysis of chromatographic peaks corresponding to C retention times and m/z for chemical standards were extracted and integrated in ChromXtractorPro. Metabolite levels are normalized to corresponding peaks in the 15N internal reference and normalized to wild-type. Dots represent the ratios of integrated intensity of metabolite to integrated intensity of its 15N reference peak. Centre lines show the medians of 4 biological replicates. Upper box limit is the 25th percentile and Lower box limit is the 75th percentile as determined by R. The extent of the whiskers is 1.5 times the box limits (interquartile range from the 25th and 75th percentiles). Asterisks indicate a Student’s t-test p-value of less than 0.001.

Figure 35 Acetyl-CoA and HMG-CoA levels across ergosterol hypomorphs in combination with tda5Δ

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I observe markedly lower acetyl-CoA in each of the tda5Δ suppressed strains whereas TDA5 strains harbouring the same mutations showed wild-type levels. These data appear to support the hypothesis that reduced acetyl CoA levels are involved in the metabolic response to loss of function in TDA5. In order to identify the reaction Tda5 functions in and to understand how metabolic phenotypes emerge due to loss of function of Tda5, I set out to characterize the metabolome of yeast carrying conditional expression of Tda5.

4.2.21 Initial metabolic response induced by loss of function of Tda5 In order to measure initial metabolite level changes induced by loss of function of TDA5, I cloned TDA5 and YDL114W into a high copy plasmid under the control of the GAL1 promoter for a high level of induction when galactose is used as a carbon source. The GAL1 promoter is strongly repressed through catabolite repression by growth on glucose. The plasmids were transformed into wild-type ura3Δ strains and crossed to tda5Δ ydl114wΔ unsuppressed strain, confirmed unsuppressed by streaking to YP-glycerol. The heterozygous diploids were sporulated and dissected to give rise to fully complemented haploids displaying wild-type growth. The haploid strains were phenotyped for markers and growth on glucose and raffinose, both of which repress expression of the gene under the GAL1 promoter. The pGAL1-YDL114W tda5Δ ydl114wΔ strain showed a noticeable growth defect on synthetic media plates containing glucose and a growth defect was measured in liquid culture microplate assay. Furthermore, tda5Δ strains could not grow without the YDL114W plasmid, as demonstrated by streaking TDA5 and tda5Δ strains carrying the Ura3-selectible plasmid onto 5-FOA, further indicating a dependence on YDL114W supplying function in place of TDA5. pGAL1-YDL114W tda5Δ ydl114wΔ cells were grown in YNB-galactose to ensure high level expression of YDL114W and switched to YNB- glucose in order to repress further expression of Ydl114w. Metabolite extracts were prepared from the galactose grown cultures as well as 5 minutes, 30 minutes, 1 hour, 2 hours and 3 hours post-carbon source switch. Metabolite levels were measured by LC-MS using chromatography for detection of organic acid as well as reverse-phase ion paired chromatography for detection of nucleobases, amino acids and central carbon metabolites.

As shown in Figure 36, CoA metabolites were analyzed pre-shift and 3 hours post-shift, I found a trend toward increased levels of acetyl-CoA 3 hours after shift to glucose for the TDA5 strain, while the tda5Δ strain appears to maintain pre-shift levels of acetyl-CoA. This is consistent with

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the suppressed strains, which likewise show decreased acetyl-CoA. Accordingly, HMG-CoA levels were slightly lower in the tda5Δ strain compared to the TDA5 control. Malonyl-CoA appeared to increase in the TDA5 strain, while pre-shift malonyl-CoA levels are observed in the tda5Δ strain.

Figure 36 CoA levels for TDA5 and tda5Δ strains due to YDL114W repression Strains carrying the GAL1pr_YDL114W plasmid under TDA5 and tda5Δ contexts were grown in YNB- galactose + histidine through several rounds of growth and sub-culture to a final optical density of 0.7. 100 mL cultures were harvested for galactose pre-grown cells, and cells were collected onto filters by vacuum filtration and transferred to an equal volume YNB-glucose + histidine. After three hours incubation in YNB-glucose CoA metabolites were extracted. For each biological replicate, CoA metabolites were extracted by harvested by centrifugation and adding 1.2 mL extraction solvent ACN/IPA 3:1, followed by three rounds of freeze-thaw. Samples were centrifuged, and 125 µL acetic acid was added. Acidified extracts were separated and eluted according to (Snyder et al., 2015). Eluates were dried under N2, reconstituted in a volume of water according to optical density at time of extraction and 15N purified CoA reference from yeast was added. Metabolites were separated and measured by reverse- phase ion-paired chromatography and detected by Q-TOF. Targeted analysis of chromatographic peaks corresponding to retention times and m/z for chemical standards were extracted and integrated in ChromXtractorPro. Metabolite levels are normalized to corresponding peaks derived 15N internal reference and normalized to wild-type. Dots represent the ratios of integrated intensity of metabolite to integrated intensity of its 15N reference peak. Centre lines show the medians of 3 biological replicates. Upper box limit is the 25th percentile and Lower box limit is the 75th percentile as determined by R.

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When the dynamics of metabolite levels were assessed, I found that unlike the subtle changes observed for CoA metabolites, a dramatic shift in N-acetyl-glutamate levels appears over time and this effect is slower or less prominent in the tda5Δ strain compared to TDA5 as shown in Figure 37. N-acetyl-glutamate is part of the mitochondrially localized cycle for production of ornithine and its formation is dependent on acetyl-CoA. The slower increase in N-acetyl- glutamate in the tda5Δ compared to the TDA5 strain may be due to insufficient relative acetyl- CoA input through this pathway. Ornithine and citrulline levels, on the other hand, appear to be static.

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Figure 37 Changes to N-acetyl-glutamate cycle due to YDL114W repression in TDA5 and tda5Δ strains Strains carrying the GAL1pr_YDL114W plasmid under TDA5 and tda5Δ contexts were grown in YNB- galactose + histidine through several rounds of growth and sub-culture to a final optical density of 0.7. During metabolite extraction, 5 mL of cultures were harvested for galactose pre-grown cells, cells were collected onto filters by vacuum filtration and transferred to an equal volume YNB-glucose + histidine. Metabolite samples were extracted at 5, 30, 60, 120 and 180 minutes post shift to YNB-glucose. CoA metabolites were extracted by harvested by centrifugation and adding 1.2 mL extraction solvent ACN/IPA 3:1, followed by three rounds of freeze-thaw. Samples were centrifuged, and 125 µL acetic acid was added. Acidified extracts were separated and eluted according to (Snyder et al., 2015). Eluates were dried under N2, reconstituted in a volume of water according to optical density at time of extraction and 15N purified CoA reference from yeast was added. Metabolites were separated and measured by reverse- phase ion-paired chromatography and detected by Q-TOF. Targeted analysis of chromatographic peaks corresponding to retention times and m/z for chemical standards were extracted and integrated in ChromXtractorPro. Metabolite levels are normalized to corresponding peaks derived 15N internal reference and normalized to wild-type. Dots represent the ratios of integrated intensity of metabolite to integrated intensity of its 15N reference peak. Centre lines show the medians of 3 biological replicates. Upper box limit is the 25th percentile and Lower box limit is the 75th percentile as determined by R.

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The trends in metabolite level changes due to YDL11W4 depletion in the tda5Δ strain observed for the N-acetyl-glutamate cycle were also observed in the TCA cycle where immediate changes were observed and the dynamics were altered between tda5Δ and TDA5. As shown in Figure 38, the most prominent change is the decrease in the left hemisphere of the TCA cycle including intermediates succinate, fumarate and malate. Where tda5Δ and TDA5 are initially well-matched for levels, the return to pre-shift levels is asynchronous for the two strains. The asynchrony may be due to a difference in growth rate in response to loss of Ydl114w in the tda5Δ strain. Furthermore, malate levels are fully restored after 2 hours in TDA5 and 3 hours in tda5Δ, whereas succinate is elevated but not fully restored. There appears to be a block from 2- ketoglutarate to succinate as citrate, isocitrate and 2-ketoglutarate appear to increase over time. Again, we observe an asynchrony between TDA5 and tda5Δ, which could be due to slower adaptation of metabolism to changing nutritional requirements or due to cells becoming respiratory deficient.

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malate citrate

fumarate isocitrate

succinate 2-ketoglutarate

Figure 38 Changes to Tricarboxylic Acid Cycle due to YDL114W repression in TDA5 and tda5Δ strains Samples were prepared and plotted as in Figure 37 with alternating TDA5 and tda5Δ strains and hours post-shift to glucose plotted on the x axis.

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As shown in Figure 39, for both TDA5 and tda5Δ strains, the initial decreases in metabolite levels seen for the left arm of the TCA cycle are also observed for branched chain amino acids where 2-isopropylmalate accumulates over time and leucine, isoleucine and valine levels have diminished and are not restored to levels achieved on galactose. This could be an adaptation to growth on glucose where branched chain amino acids are consumed more rapidly to match demand for amino acid synthesis in the faster growing strain (Godard et al., 2007). Expression of mitochondrial BCAA enzymes are increased upon conditions that increase intracellular level of AMP, such as carbon source switch to glucose (reviewed in Kohlhaw, 2003).

Figure 39 Branched chain amino acid dynamics in tda5Δ and TDA5 upon conditional knockdown of YDL114W Samples were prepared and plotted as in Figure 37. In addition to targeted metabolomics, full scan untargeted analyses were carried out to identify changes in metabolites not directly captured by our set of chemical standards. I took an unbiased approach to peak-finding and integrated peaks across all metabolite samples for all strains at each timepoint. There were no peaks that were significantly altered between 0 and 3 hours that showed a coherent change in levels due to YDL114W repression.

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4.3 The suppressed tda5Δ deletion mutant has negative genetic interactions with deletions in other short chain dehydrogenases In order to determine which pathways may be upregulated during response to loss of TDA5, I hypothesized that deletion of compensating metabolic pathways would produce a deleterious effect leading to negative genetic interactions with the tda5Δ strain. Since tda5Δ had been screened twice in global screens of genetic interactions in yeast, I examined the overlap between these two screens (Costanzo et al., 2016). Since these two tda5Δ strains had been suppressed by loss of function mutations in HST1 and SUM1 (van Leeuwen et al., 2016), I determined the set of negative genetic interactions that were exclusive to the tda5Δ screens and not present in the sum1Δ, rfm1Δ and hst1Δ deletion screens as shown in Figure 40b. Remarkably, there was a small window of overlap yielding 8 deletion mutants with strong negative genetic interactions. These interacting genes were ldo16Δ, the recently annotated component of the seipin complex responsible for maintaining contacts between lipid droplets and the ER (Teixeira et al., 2018), the dehydrodolichyl diphosphate synthase nus1Δ (Park et al., 2014), the Glc-NAc acetyltransferase component alg14Δ required for dolichyl-linked oligosaccharide synthesis in the ER membrane, the mitochondrial t-RNA-modifying enzyme slm3Δ, and a set of genes 4 encoding short chain dehydrogenases homologous to TDA5 highlighted in Figure 40a: env9Δ, an uncharacterised SDR found in lipid droplets and the endomembrane system; srl4Δ, a second uncharacterised SDR; ydl114wΔ the sporulation specific SDR that I showed is a suppressor of tda5Δ; and oar1Δ encoding the characterized SDR that is mitochondrial 3-keto-acyl-ACP- reductase involved in mitochondrial fatty acid biosynthesis (Venkatesan et al., 2014). Functional descriptions, strength of genetic interaction (GI score) and similarity are shown in Figure 40a. These negative genetic interactions suggest a function for Tda5 in providing fatty acids between ER/vacuole and mitochondria and potentially suggests a role in dolichol metabolism. Dolichol is exchanged for very long chain fatty acids during GPI anchor formation within the ER.

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Figure 40. tda5Δ has negative genetic interactions with deletions in other short chain dehydrogenases Genetic interaction data was downloaded from thecellmap.org (Costanzo et al., 2016) for hst1Δ, sum1Δ, rfm1Δ and two tda5Δ screens. Deletion mutants having SGA scores less than -0.1 in both tda5Δ screens were included for analysis and deletion mutants having negative genetic interactions in hst1Δ, sum1Δ or rfm1Δ screens were removed. As previously mentioned, the majority of short chain dehydrogenases in yeast remain uncharacterized. As shown in Table 2 , 9 of 16 have no assigned reaction. They represent an important functional class of enzymes as 2 are known to be essential and as shown in this study, Tda5 is essential, a phenotype that was missed due to suppressors. Negative genetic interactions indicate parallel activities between SDRs and functional analysis may necessitate the use of double and triple knockouts.

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Table 2. Most short chain dehydrogenases remain uncharacterized

4.3.1 GO enrichment suggests metabolic process for Tda5 In order to determine which biological processes Tda5 interacts with, I carried out gene ontology enrichment on all genes with negative genetic interactions with an SGA score of less than -0.1 that were private to the tda5Δ screens, not requiring such hits to be present in both tda5Δ screens. As shown in Table 5, the highest enrichment was for lipid metabolic processes, protein lipidation, vesicle organization, and regulation of protein modification process. These results are consistent with a role for Tda5 in supporting lipid production in the endoplasmic reticulum and supply of lipids between compartments.

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Table 5 Gene ontology terms enriched for negative genetic interactions with tda5Δ Genetic interaction data were downloaded from thecellmap.org (Costanzo et al., 2016) for hst1Δ, sum1Δ, rfm1Δ and two tda5Δ screens. Deletion mutants having SGA scores less than -0.1 in either tda5Δ screens were included for analysis and deletion mutants having negative genetic interactions in hst1Δ, sum1Δ or rfm1Δ screens were removed. GO enrichment for the resultant gene list was determined using SGD Gene Ontology Slim Mapper for the Process ontology set (Cherry et al., 1998).

4.3.2 The tda5Δ strain is sensitive to unsaturated fatty acids As GO enrichment analysis suggested that Tda5 genetically interacts with proteins required for lipid homeostasis, I assessed tda5Δ growth in the presence of multiple long, medium and short chain fatty acids in order to measure defects in lipid metabolism,. The freshly dissected strain demonstrates lipo-sensitivity as it is slower growing in the presence of long chain monounsaturated fatty acid oleate (C18) and as shown in Figure 41a, the tda5Δ strain is completely unable to grow in the presence of polyunsaturated lineoleic acid (C18). There are several explanations for the growth defect observed in the presence of these fatty acids which may be the result of inability to store or breakdown unsaturated fatty acids, leading to an accumulation of precursors. Since respiration is required to metabolize acetyl-CoA from fatty acid breakdown, I tested whether the respiratory deficient mutant of the mitochondrial DNA polymerase mip1Δ strain was likewise sensitive to oleate. The mip1Δ strain grew well in the presence of oleate, ruling out reduced mitochondrial function as the cause of tda5Δ sensitivity to oleate. As modeled in Figure 41b, the sensitivity observed toward unsaturated fatty acids for the tda5Δ strain could also be the result of the inability to withstand oxidative stress caused by breakdown of unsaturated fatty acids. Many deletion mutants of genes with roles in the peroxisome are known to be oleate sensitive due to their inability to break down oleate through β-oxidation (Lockshon et al., 2007). Mutants defective in lipid transport such as the mdm10Δ deletion strain are likewise sensitive to oleate alongside mutants defective in lipid particle

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synthesis (Lockshon et al., 2007). Lipid particles provide stores for neutral lipids: steryl esters and triglycerides which poise the cell for rapid responding to increased lipid demand. Steryl esters are produced by Are1/Are2 the acyl-CoA: sterol acyltransferases (Czabany et al., 2008). These two enzymes are inhibited in the presence of oleate (Connerth et al., 2010). The are1Δare2Δ double mutant is viable but does not produce lipid particles, which are necessary for sequestering toxic lipophilic molecules and maintaining low levels of free fatty acids (Sandager et al., 2002). The are1Δare2Δ double mutant is synthetic lethal with arv1Δ, where Arv1 acts in GPI anchor insertion in the endoplasmic reticulum and sterol insertion into the plasma membrane and the arv1Δ deletion mutant has altered sterol distribution (Ikeda et al., 2016; Kajiwara et al., 2008; Tinkelenberg et al., 2000). The fact that tda5Δ cells are slow growing in the presence of oleate suggests that tda5Δ might have disrupted lipid flow and diminished GPI anchor synthesis alongside reduced lipid particle synthesis in the presence of oleate analogous to the are1Δare2Δarv1Δ triple mutant synthetic lethality. The GO enrichment data for tda5Δ and the sensitivity to oleate for the tda5Δ strain implicate Tda5 function in lipid metabolism supporting biogenesis of the ER and plasma membrane.

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Figure 41. tda5Δ strains are sensitive to unsaturated fatty acids A) Spot assays for tda5Δydl114wΔ and control strains on YPD and unsaturated fatty acids B) Model for exacerbation of lipotoxicity by unsaturated fatty acids Overnight cultures of strains grown in YPD or YP-galactose were normalized to an optical density of 1 and 10-fold serial dilutions were prepared in a 96 well plate in 200 µL volume. Cells were mixed using a P200 multi-channel between spotting. For spotting, 3 µL was transferred using a Rainin P20 multi- channel pipette. Spot plates were allowed to dry at room temperature before incubating 2 nights at 30 °C. Images were captured using AlphaImager gel documentation system with white light transillumination.

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4.4 Discussion When I set out to uncover the function of TDA5, the literature surrounding this gene had been muddied by the use of a suppressed strain available from the systematic deletion set. TDA5 is named for topoisomerase I damage affected for its sensitivity in response to DNA damage (Reid et al., 2011). In this screen, a dominant negative topoisomerase mutant was overexpressed in the yeast deletion collection inducing DNA breaks and apoptosis identifying sensitivity for tda5Δ alongside over 100 other mutants including 10 additional mutants in uncharacterized genes including TDA5. Seven years after the study was carried out, only one of these ten uncharacterized genes, TDA2, has been characterized. TDA2 is now known for its role in regulating actin assembly, a role incongruent with its naming (Shin et al., 2018) and the remaining 8 have proven recalcitrant to functional study. Three of the TDA genes in addition to TDA5 have paralogues, and analogous to YDL114W compensation for loss of TDA5, the paralogue may compensate in the absence of its TDA counterpart. Building on suppressor mapping by van Leeuwen and colleagues, this work demonstrates the tda5Δ strain used in this study was suppressed by loss of function in HST1 causing de-repression and compensation via YDL114W (van Leeuwen et al., 2016). Knowing the strain was suppressed, it would now seem unlikely that the true function of TDA5 is in maintaining response to DNA breaks. This could be tested by measuring DNA damage foci dynamics in response to DNA double strand breaks in tda5Δ and wild-type yeast strains by fluorescence microscopy (Styles et al., 2016). Knowing that tda5Δ mutants have increased levels of oxidative stress may account for sensitivity to the top1 allele, and the identification of TDA5 in this screen may reflect its role in maintaining oxidative stress levels.

Throughout this work, we have revealed that yeast has a severe growth defect upon loss of tda5Δ. Tda5 can be described as essential in several contexts. The most prominent essential role for Tda5 is providing growth on respiratory carbon sources as the strains are petite. We know that a minor localization of Tda5 is within the mitochondria, which is consistent with the observed respiratory defect and metabolomic phenotypes which display defects in metabolic reactions compartmentalized within the mitochondria. Metabolomic analysis of suppressed strains as well as metabolic phenotypes due to repression of YDL114W indicate tda5Δ harbours defects in mitochondrial metabolic pathways including branched chain amino acid synthesis, arginine biosynthesis, as well as altered dynamics of changes in TCA cycle intermediates in response to shift from galactose to glucose. Experiments carried out in this study have not revealed whether the major role of Tda5 is in its mitochondrial context, or

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whether the product of the reaction catalyzed by Tda5 is essential for respiratory function. The product of the Tda5 reaction or pathway it acts in may be necessary for maintenance of mitochondrial membrane integrity and membrane potential. Analysis of protein localization carried out in this study suggest Tda5 is present in the endoplasmic reticulum and plasma membrane. Blocking the fusion of the mitochondrial associated membrane with the endoplasmic reticulum may result in blocked trafficking of Tda5 to the mitochondrion. This can be tested using mutants defective in ERMES or vCLAMP complexes and measuring Tda5 localization by immunoblotting fractionated cells. We have cloned a truncated version of Tda5 in yeast that lacks an ER localization sequence. Loss of localization to the ER would yield a cytoplasmic Tda5. It is important to determine phenotypes of ectopically expressed Tda5 in a tda5Δ background and measure metabolite level changes in order to assess whether these mutants are able to fully restore Tda5 function, both on fermentative and respiratory carbon sources. Alternately, a mitochondrial localization sequence could be added in place of the secretion signal to direct Tda5 to the mitochondrion in order to assess whether the primary function of Tda5 is within the mitochondrion. Since Ydl114w is localized to the cytoplasm and can rescue loss of function mutations in tda5Δ, it suggests that activity carried out in the cytosol is able to compensate and implicates soluble metabolites in compensating for the loss of Tda5.

If localization of Tda5 to the mitochondrion is not required, the product of the Tda5 catalyzed reaction may be required for respiration. Lesions in enzymes that alter the makeup of mitochondrial membrane lead to accumulation of ROS. Oxidative damage induced by ROS can be reversed by addition of antioxidants—consistent with the rescue of tda5Δ growth on N-acetyl-cysteine. This is analogous to the previously mentioned deletion of cardiolipin synthase Crd1 or lyso-phosphatidylcholine acyltransferase Taz1 which result in inability to synthesize or remodel cardiolipin leading to accumulation of ROS (Chen et al., 2008). Elevated ROS results in oxidant sensitivity for crd1 mutants that can be rescued by oleic acid, the mechanism of the rescue is not understood. In contrast, tda5Δ cells are sensitive to oleic acid. The sensitivity to oleic acid may result from inability to breakdown unsaturated fatty acids. This is unlikely as mutations in TDA5 have no genetic interactions with peroxisomal β- oxidation, and the peroxisomal compartment has not been implicated at all throughout this study. An alternate explanation is that incorporation of unsaturated fatty acids into membranes in the tda5Δ mutant exacerbates the observed membrane defect leading to growth suppressing levels of oxidative stress. This is consistent with antioxidant N-acetyl-cysteine

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rescuing the growth defect of tda5Δ. A third explanation is that oleic acid repression of steryl ester synthesis in combination with mutation in tda5Δ leads to improper localization and composition of membrane lipids to support insertion and wild-type function of integral membrane proteins such as GPI anchor proteins, analogous to the arv1Δare1Δare2Δ synthetic lethality. The human ARV1 gene complements the yeast arv1Δ deletion and causes neurodevelopmental defects and epileptic encephalopathy (Palmer et al., 2016).

Among the most striking findings of this work was the identification of multiple lesions in the ergosterol biosynthetic pathway were able to suppress loss of function in tda5Δ and restore the growth defect on respiratory media. Since the vast majority of enzymes in the ergosterol biosynthesis pathway are essential, it emphasizes the crucial role of Tda5 in the cell and may indicate that compensation for loss of TDA5 takes precedence over cellular demand for sterols or that reducing sterol biosynthesis is a way to rescue the lipid defect induced by loss of tda5Δ. From mapping suppressors independent of YDL114W expression, alongside high throughput positive genetic interactions in combination with the chemical- genetic interaction identified for miconazole and mevinolin in rescuing the tda5Δ growth defect, these orthogonal studies have underscored a connection between Tda5 and ergosterol. However, the connection between the two remains unclear. It could be that the product of Tda5 is required for negative regulation of ergosterol biosynthesis. In the absence of Tda5, cells may have elevated ergosterol biosynthesis which would lead to loss of membrane fluidity and a membrane defect. It would also lead to incorporation of ergosterol at a higher proportion into mitochondrial membrane, leading to loss in the assembly of cytochrome complexes and ultimately loss of respiratory capacity. Metabolomics analysis of the initial changes induced upon loss of YDL114W expression did not reveal elevations in HMG-CoA, which suggests that a model invoking elevated sterol production is inconsistent with data. Instead, the data indicate that metabolites upstream of ergosterol are involved in compensation for loss of tda5Δ. Measurements of CoA levels in tda5Δ strains undergoing loss of complementation by YDL114W, indicate that a lower acetyl-CoA and malonyl-CoA compared to TDA5, which may indicate increased demand for these metabolites in the tda5Δ context. Examination of lipid dynamics via stable isotope labelling followed by analysis of hydrophobic compounds could resolve the question of increased demand for CoA metabolites.

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The GO enrichment data, localization of Tda5 to the ER and plasma membrane, and the nature of the known functions of other short chain dehydrogenases in reducing acyl hydroxyl groups to ketones in fatty acid synthesis (Oar1, Fas2, Ifa38), sphingolipid synthesis (Tsc10) and triglyceride synthesis (Ayr1), implicate a role for Tda5 in lipid metabolism. As described in the results section, there are several known connections between ergosterol synthesis and other lipid metabolic pathways. Fatty acid elongase mutants are sensitive to the polyene amphotericin B, a drug that binds ergosterol to form pores in the membrane (Sharma et al., 2014). This finding emphasizes the link between fatty acid and ergosterol lipid pools in maintenance of membrane integrity. Of note is the fatty acid elongase Elo2, which was previously named Fen1 for its resistance to fenpropimorph, the morpholine Erg24 inhibitor. Mutants in elo2Δ displayed an increased ergosterol level (Ladevèze et al., 1993). Elo2 mutants are not cross-resistant to amphotericin B indicating that the fatty acid elongase resistance to amphotericin B may not be a result of a membrane defect. Ergosterol serves two purposes, firstly to provide neutral lipid for incorporation into membranes and secondly for promoting G1 exit also known as sparking function (Ramgopal and Bloch, 1983). The fen1 (elo2) elongase mutant is known to relieve the need for sparking ergosterol. We do not know if tda5Δ lesions have a parallel role, although the strains are sensitive to fenpropimorph. When I measured fenpropimorph sensitivity in Ifa38, the SDR directly downstream of fatty acid elongase, the mutants demonstrated wild-type levels of resistance. The second site suppressors of tda5Δ strains were selected on media without sterol supplementation, thus the upc2, erg11 and erg1 mutants in combination with tda5Δ are able to grow on media without sterols. The yeast requirement for sterols depends on the composition and availability of other membrane lipids. Triple mutants of erg25, erg11, and hem2 or hem4 are able to grow exclusively on lanosterol. The erg6 hypomorphic mutants have altered permeability and inability to use respiratory carbon sources. Erg3 is only essential in heme deficient cells (Davies and Rine, 2006). Mutants in ERG25 can be rescued by low levels of heme as the P450 enzyme Erg11 is dependent on heme and it prevents the accumulation of toxic sterols (Gachotte et al., 1999). When yeast are deficient in heme production, they are sterol and unsaturated fatty acid auxotrophs which are dependent on accumulation of short chain fatty acids and their incorporation into PtdIns to support growth (Ferreira et al., 2004). Together, these studies demonstrate compensation between lipid metabolic pathways, however the nature of the compensation varies between mutants and the mechanisms are notoriously difficult to predict (Daum et al., 1999).

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As Tda5 is closely related to other short chain dehydrogenases, it may overlap in its biochemical reaction. One important reaction to consider is the 3-ketoacyl-reductase step of very long chain fatty acid biosynthesis, for which only one enzyme has been annotated, Ifa38 (Denic and Weissman, 2007). A noted discrepancy identified within the yeast genome scale metabolic network is that very long chain fatty acids are essential for their roles in GPI anchor biosynthesis, however, the ifa38Δ strain is viable (Aung et al, 2013). It is reasonable that Tda5 or an alternate uncharacterized SDR catalyzes a parallel role in very long chain amino acid synthesis and is compensating for Ifa38 in the ifa38Δ mutant.

The function of Tda5 may be within the ergosterol pathway itself (shown in Appendix I). There are two steps of ergosterol biosynthesis that require SDR activity. The 3-keto-sterol reductase Erg27 apparently acts on both of these steps, performing reduction of the C3 ketone 3-keto-methyl zymosterol to the alcohol methyl zymosterol so that the C4 methyl can be activated and removed by the dual action of Erg25 and Erg26, which reforms the ketone, subsequently, Erg27 acts again to reduce the ketone to the alcohol zymosterol (Gachotte et al., 1999). The yeast erg27Δ strain is viable in the upc2 background on rich media and the reaction is complemented by the human SDR catalyzing reduction of zymosterone (cholesta- dienone) to zymosterol encoded by HSD17B7 (Marijanovic et al., 2003). In the characterization of Erg27, the erg27Δ deletion mutant was prepared and fed lanosterol and sterols were measured by GC-MS to find that zymosterone accumulated (Gachotte et al., 1999). I find it surprising that the erg27Δ mutant produces zymosterol at all if Erg27 is truly acting upstream of zymosterone. This incongruency may indicate the presence of a second 3- keto-sterol reductase that acts upstream of Erg27 and may indicate a function provided by Tda5 or another uncharacterised SDR. Analogous to miconazole suppressing loss of function in TDA5, azoles also suppress erg25Δ and are likewise suppressed by erg11 hypomorphs (Gachotte et al., 1997). This overlap between the erg25Δ and tda5Δ mutants is striking and further implicates Tda5 in the first 3-keto-sterol reductase reaction. The human cholesterol biosynthetic pathway has a distinct sterol reductase at this step and no in vitro activity assays have assigned Erg27 to this reaction. Furthermore, it’s surprising that HSD17B7 could complement Erg27 (Marijanovic et al., 2003), as it is not shown to carry out the first reduction step, which implies that the wild-type function for the first step is intact in the erg27Δ mutant.

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The resistance of tda5Δ mutants to ergosterol pathway inhibitors underscores a role for Tda5 in antifungal resistance. Multi-drug resistance in fungi is dependent on membrane lipid environment, status of efflux pumps and uptake mechanisms (Mukhopadhyay et al., 2002). Lowered expression of Erg11 is the main mechanism of resistance to azole antifungals (reviewed in White et al., 1998). At the same time, lipid homeostasis also contributes to drug sensitivity. Efflux pumps localize to lipid rafts and lowered expression of Erg1 from the ergosterol biosynthetic pathway and Ipt1 from the sphingolipid pathway lowers plasma membrane surface localization of the major efflux pumps, leading to drug sensitivity and decreased efflux of rhodamine 6G (Pasrija et al., 2005). Further studies of lipid homeostasis in fungi will elucidate connections between resistance and sensitivity and the complement of lipids involved.

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5 Thesis summary and future directions 5.1 Conclusions In summary, the overall accomplishments of this dissertation are in understanding the functions of two previously uncharacterized enzymes in yeast, and expanding the understanding of related metabolic pathways. As yeast is the best annotated eukaryotic organism, refinement of gene functions is critical to achieving a high-level knowledge of eukaryotic biology.

The Das2 enzyme has been completely missed from studies concerning pyrimidine metabolism. My experiments unambiguously identify Das2 as a uridine kinase and its role as an obligatory member of the Das2-Urk1-Fur1 pyrimidine salvage complex. These data revise the accepted model of pyrimidine salvage in yeast and indicate the presence of a previously unrecognized pyrimidine salvage complex. The presence of orthologues of Das2, Urk1 and Fur1 in the commensal pathogenic yeast Candida albicans as well as the dependence of uridine kinase activity on a functional UPRTase in this organism suggests that the composition and function of this complex is conserved from S. cerevisiae to the pathogen C. albicans. The minimal amount of pyrimidine nucleoside salvage present in human cells, in hand with the efficacy of pyrimidine analogue treatment, and the increased potency of nucleoside analogs over nucleobase inhibitors demonstrates the potential selectivity of this class of drugs in treatment of fungal infections in mammals.

These studies also have established Tda5 as a short chain dehydrogenase that is mainly localized to the endoplasmic reticulum and plasma membrane. The severe growth defect of the tda5Δ mutant demonstrates the crucial function conveyed by Tda5. In addition, the enzyme is essential for growth on respiratory carbon sources, response to oxidative stress and growth in the presence of oleate. High copy expression of the sporulation specific paralog of TDA5, YDL114W, rescues the growth of tda5Δ on respiratory carbon sources. The tda5Δ mutants suppress readily, through lesions in the Hst1/Rfm1/Sum1 histone deacetylase complex that lead to expression of YDL114W, which complements tda5Δ. This meiotic histone deacetylase complex comprises a large mutational target, with over 5.5 kilobases of protein coding sequence. I also identified suppressors of tda5Δ that are independent of YDL114W expression and mapped them to hypomorphic mutations in ERG1, ERG11, and UPC2. All of these suppressors, as well as a hypomorphic mutant in ERG12 function in ergosterol biosynthesis and are able to suppress loss of function in tda5Δ. Although the

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mechanism of suppression is not yet clear, acetyl-CoA levels are altered in the suppressed mutants. Strains deleted for TDA5 have a membrane defect, are sensitive to growth on unsaturated fatty acids, and the tda5Δ growth defect can be relieved by growth in the presence of N-acetyl-cysteine and the ergosterol biosynthetic pathway inhibitors miconazole and mevinolin. GO enrichment of negative genetic interactions reveal further links to lipid homeostasis. Overall, the compromised membrane function and mechanisms of suppression suggest that Tda5 is acting in lipid metabolism.

5.2 Future Directions

5.2.1 Future directions relating to Das2 In order to establish the homolog of Das2 as an effective target of nucleoside analogues in pathogenic Candida albicans, one must show that the function of Das2 is conserved. One would begin by preparing the homozygous deletion of the putative single domain uridine kinase and measuring changes in uridine concentrations by LC-MS. Since the activity appears to be dependent on the UPRTase, one would need to prepare mutants for the Urk1 and Fur1 enzymes as well and establish whether there are conserved genetic interactions between Fur1, Urk1 and Das2, by measuring pyrimidine metabolite levels. The presence of a mitochondrial dihydroorotate dehydrogenase in Candida albicans suggests that these cells are pyrimidine auxotrophs under anaerobic conditions. One ought to test the sensitivity to pyrimidine nucleoside analogues under biofilm conditions and under hypoxic conditions. Since there is natural resistance to 5-fluorocytosine among a single clade of natural isolates due to natural variation in FUR1, and the fact that nucleosides are more potent than nucleobases, one should establish the efficacy of pyrimidine nucleosides over their nucleobase counterparts in antifungal treatment.

A formal comparison of function between Das2 and other uridine kinases has not been carried out. My analysis of the amino acid conservation of Das2 in Figure 6 indicates a high degree of non-synonymous amino acid substitutions compared to other uridine kinases, even for highly conserved residues of Das2. Although new mutations rarely convey enhanced function, 80% of mutations decrease the thermodynamic stability of the protein (Tokuriki and Tawfik, 2009; Tokuriki et al., 2008). Consequently, a high evolutionary speed is indicative of high selective pressure for functional enhancement. A high degree of non-synonymous amino acid substitutions is also associated with loss of selective pressure leading to complete loss of function and pseudogenization. To determine whether these changes in Das2 are associated

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with a gradual diminished function leading toward pseudogenization versus enhanced activity or a specialized role, one must directly compare uridine kinase activities within divergent yeasts to determine structure-function relationships that were present in the ancestral strain. A comparison of the activities of Das2 to other uridine kinases would reveal whether Das2 has enhanced uridine kinase activity. Das2 catalyzes the uridine kinase reaction at a maximal velocity of 243 µmol/min/mg protein which is close to 200 µmol/min/mg protein for uridine kinase from E. coli.

Although this study suggests Fur1 does not rely on Urk1 and Das2 to function, this has not been directly measured. Immunoblotting to measure Fur1 levels needs to be carried out to determine effects of mutations in URK1 and DAS2. We also have not addressed the stoichiometry of Das2/Urk1/Fur1 complexes. Each protein can be fluorescently tagged and complex membership can be determined by fluorescence resonance energy transfer (Damelin and Silver, 2002; Wilson et al., 2002) and localization by super-resolution microscopy (Szymborska et al., 2013). These experiments would also reveal the localization of complexes within cells and determine the ratio of Fur1 free versus in complex. Alternately, size exclusion chromatography on extracts of tagged strains will allow us to quantify relative stoichiometric ratios of Das2/Urk1/Fur1, but will not reveal the absolute nature of higher order oligomers or patterning within the complex. Additional proteomics studies remain to be carried out to determine novel complex members not identified in this study. These studies may reveal the relationship between the UMP salvage complex and other metabolic processes such as de novo pyrimidine and purine biosynthesis, as well as ribosome biogenesis.

Our ultimate goal is to understand how cells alter metabolism to support growth under dynamic environmental conditions. Several key questions have merged from this study in how cells coordinate pyrimidine salvage and de novo synthesis. The fur1Δ mutant strains used in this study in the context of wild-type URH1 show severe accumulations of uracil. The presence of uracil is expected to repress URA2 transcription (Potier et al., 2006), which would dampen UMP biosynthesis. Yet, there is no growth defect observed for these strains. This lack of growth defect indicates the presence of a mechanism for expression of biosynthetic enzymes to maintain growth even in the presence of high uracil.

My literature search surrounding the identification of DAS2 revealed a potential role for another uncharacterized enzyme in pyrimidine metabolism. Saccharomyces kluyveri is a yeast strain that can use uracil as a sole nitrogen source. The nitrogen catabolism pathway

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involves URC1, which is the homolog of the uncharacterized YDR520C gene in S. cerevisiae. Recent studies have shown uridine kinases and UPRTases are the first steps in utilization of pyrimidines as nitrogen sources. In senso lato yeasts, a reductive pathway is required for nitrogen catabolism. Since standard laboratory conditions use fermentative carbon sources where mitochondrial activity is low, previous experiments may have missed uracil catabolism due to limiting amounts of reducing equivalents. A first step toward determination of whether Saccharomyces cerevisiae can catabolize uracil would be to challenge yeast with uracil in the presence of a more respiratory carbon source. I have previously profiled the ydr520cΔ mutant by LC-MS metabolomics and identified 5 compounds that are significantly (Student’s t-test p-value <0.001) altered compared to wild-type. The phenotypes from the literature support an enzymatic function for YDR520C in pyrimidine metabolism, warranting further examination of the nature of these compounds as well as further experiments involving additional carbon and nitrogen sources.

5.2.2 Future directions for annotation of Tda5 Acetyl-CoA is the metabolite at the base of the ergosterol and fatty acid biosynthetic pathways, which in turn leads to production of other lipid metabolites. Metabolomics analysis of suppressed tda5Δ strains show lowered acetyl-CoA levels which may reflect altered demand for the metabolite, or production of it. The nature of a metabolic block or increased demand cannot be determined without measuring acetyl-CoA flux in wild-type and suppressed mutants. This can be carried out by stable isotope labelling using uniformly labeled 13C-glucose and quantitation by LC-MS using the methodology and mutants developed in this study. These experiments will suggest the cause of altered levels of acetyl- CoA in the tda5Δ deleted cells and will implicate pathways involved in relieving demand for the Tda5 reaction.

Likewise, we must determine the relative amount of sterol biosynthesis carried out by suppressed tda5Δydl114wΔ strains and whether there are any blocks in this pathway that result from the suppressor mutations. Sterol extracts have been prepared for suppressor mutations in the TDA5 and tda5Δ contexts, as well as loss of complementation by YDL114W in experiments analogous to those carried out above for measuring the soluble metabolome. Once analyzed, these data will provide steady state measurements for sterol metabolites, from which we can indicate activation or repression of pathways and indicate blocks. In contrast, pathway flux can be measured by incorporation of 13C-mevalonate and measured by GC-MS

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in suppressed strains or after loss of expression of inducible TDA5. Together, measuring the complement of sterols and their dynamics will reveal whether Tda5 acts in ergosterol biosynthesis as well as determining the nature of each of the suppressors and their roles in compensation for tda5Δ.

A full lipidomic analysis of the balance of sterols, phospholipids, sphingolipids and fatty acids is required to determine the proportions of lipids harboured by suppressed tda5Δ strains. There may be a lipid that is missing or accumulating in the tda5Δ strains that would be missed by the electrospray ionization LC-MS techniques used thus far. Furthermore, changes in lipid homeostasis in suppressed mutants will reveal modes of compensation and lipidomics of TDA5 repression would reveal the dynamics of changes in lipid pools in response to loss of Tda5. In order to determine the relationship between Tda5 and other SDRs, the lipid profiles of these deletion strains warrant characterization.

We have predicted that Tda5 may have a role in GPI anchor assembly from gene ontology analysis and have also determined that the tda5Δ deletion strain is sensitive to oleate. Together, these data support a possible role for Tda5 in GPI anchor assembly and membrane insertion in yeast analogous to the synthetic lethal interaction between are1Δare2Δ and arv1Δ, where Are1 and Are2 are suppressed by oleate. First the negative genetic interactions must be confirmed, and work is underway to confirm whether these interactions are private to the tda5Δhst1 and tda5Δsum1 contexts used in the SGA screens. If tda5Δ does indeed have negative genetic interactions in the SUM1 and HST1 contexts, we can determine whether Tda5 has a role in GPI anchor assembly and membrane support, analogous to Arv1. GPI anchor defects are identified by measuring accumulation of mannosylated GPI anchor components using 3H-mannose and visualizing by thin layer chromatography according to (Ikeda et al., 2016). The arv1Δ deletion strain is sensitive to aureobasidin A, an inhibitor of inositol phosphorylceramide synthase Aur1, and we can determine whether tda5Δ is likewise sensitive (Zhong et al., 1999).

As we have seen in the study of Das2, pyrimidine salvage represents a relatively isolated node on the metabolic network where we were able to rapidly dissect and establish the relative roles of Das2, Urk1, Fur1 and Urh1. We were also able to establish rules for metabolic flow on how the metabolic network reacts due to loss of function in these enzymes. In contrast, lipid homeostasis comprises a relatively vast network of genes, pathways and metabolites. The lipid metabolic network is buffered against genetic deficiencies as single

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lesions in lipid metabolism are compensated for and it is difficult to predict changes to the complement of lipids involved in compensation for a given mutation (Daum et al., 1999). Furthermore, many single mutants in lipid metabolism do not demonstrate membrane structural integrity or fluidity defects or strain growth defects (Daum et al., 1999) and it is therefore difficult to design systematic genetic screens for annotation of lipid functions. As I have outlined in this study, we can measure initial metabolic changes due to loss of function using conditional mutants. In measuring modes of compensation for loss of function mutations in lipid homeostasis, we can determine patterns in which lipids can compensate for one another in order to establish a set of rules for lipid compensation. For example, even without establishing direct functionalities, temporal lipidomic analysis following repression of tda5Δ cells rescued with Ydl114w expression will pinpoint where in lipid metabolism lesions are induced and how this lesion is compensated for. Common modes of compensation likely prevail that are conserved from yeast to humans even though the signalling mechanisms and cohorts of genes and metabolites differ. In this way, we can begin to understand how the balance of lipids is altered in genetically and nutritionally perturbed cells. I have outlined the methodologies for revealing functions of uncharacterized genes with predicted roles in lipid metabolism, and for explaining unresolved mechanisms of lipid compensation, as we can induce a genetic lesion to alter an activity from wild-type to mutant and subsequently measure initial metabolic changes. If there is a suppressible growth defect, we can select second site suppressors that will indicate modes of suppression. Ultimately, we aim to predict the dynamic response of the lipid metabolic network and how it contributes to homeostasis. This will additionally require study of specific organelle requirements, and methodologies can be expanded to fractionated cells.

There are many human diseases related to lipid homeostasis. Wolman disease and cholesterol ester storage disease are examples of inborn errors of metabolism where early onset lysosomal acid lipase deficiency causes hypercholesteremia and pathology related to appearance of foam cells across tissues including bone marrow and these diseases are mediated with treatment recombinant lipase and statins (Du et al., 1998). Niemann-Pick Disorder (NPD) is another inborn error of metabolism associated with mutations in the NPC1 and NPC2 genes which encode the intracellular cholesterol transporters that lead to inability to esterify cholesterol and is also associated with appearance of foam cells in tissues, elevated sphingomyelin and white matter reduction leading to neurologic abnormalities (Rao and Spence, 1977; Walterfang et al., 2010). Tangier’s disease is an inborn error of metabolism

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caused by mutations in the ABCA1 gene leading to incorrect egress of cholesteryl esters and HDL deficiency and is associated with cerebral lipidosis alongside loss of heat and pain sensation (Pietrini et al., 1985; Suarez et al., 1982). Each of these diseases have forms of late onset clinical manifestation, indicating that at some point compensation for these recessive mutations is lost or overwhelmed. Each of these IEMs is associated with neurological defects. The balance of lipids is also implicated in Alzheimer’s disease (AD), where synthesis of steryl esters is associated with enhanced Amyloid β-peptide formation and hypercholesteremia is a risk factor for AD (Huttunen et al., 2010). By understanding dynamic changes in the lipidome due to genetic perturbation, we will likely reveal rules for lipid compensation that are conserved from yeast to humans and broaden our understanding of loss of compensation that leads to disease pathology.

5.2.3 Resolving functions of clinical variants in metabolism Over the course of my thesis, the field of clinical metabolomics has expanded and we are now capable of measuring a wide range of metabolites from a diverse set of tissues and other human biological samples. At the same time, whole genome sequencing has emerged as a routine procedure in clinical analysis of pathogenicity and response to drugs. Together, we are now better equipped in establishing genotype-phenotype relationships between variants and their beneficial, neutral or pathogenic effects on metabolism. Although there are a wide range of single analyte assays available for measurement of specific metabolite levels, these, by nature, are limited in scope. By developing and standardizing expanded clinical testing to global comparisons of human metabolomes, scientists and clinicians will be provided unprecedented insight into metabolic variation within populations and their impacts on disease and developing optimal practices for personal therapeutic intervention.

Causal relationships between variants and metabolic disease cannot be established without functional assays. Metabolomic profiles can be used readouts of gene function. A major, but not insurmountable, roadblock in understanding genotype-phenotype relationships is the large amount of genomic variation between humans. Functional assays can be carried out in vitro, assessing impacts of genomic variants one disease gene at a time. One strategy is to triage functional analysis of metabolic genes, beginning with prioritization of metabolomic functional assays for pathological variants in clinically actionable genes followed by variants with conflicting reports of pathogenicity (Starita et al., 2017). Functional assays measuring metabolomic phenotypes for variants relative to unaffected controls will help to develop a

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probabilistic model of variant interpretation beginning with a few metabolic disease genes which can be expanded to causal loci used as biomarkers and diagnostics, and then to understanding loci implicated in mGWAS and mQTL studies. The studies of actionable genes can also be expanded to understanding intergenic epistasis. In this way we can convert variants of uncertain significance to pathological or benign variants in a metabolic disease context.

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7 Appendix

Supplemental 42: Ergosterol biosynthetic pathway of Saccharomyces cerevisiae

Erg10 acetoacetyl-CoA thiolase H-S-CoA

Erg13 Acetyl-CoA +H O 2 HMG-CoA synthase H-S-CoA

2 NADPH Hmg1, Hmg2 HMG-CoA reductase NADP(+)+ HS-CoA ATP Erg12 Mevalonate kinase ADP

ATP Erg8 Phosphomevalonate kinase ADP

Mvd1 ATP Mevalonate pyrophosphate decarboxylase ADP + Pi + H2O

Idi1 IPP:DMAPP isomerase

IPP Erg20 Farnesyl diphosphate PPi synthetase

IPP Erg20 Farnesyl diphosphate synthetase

PPi

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NADPH Erg9 Squalene synthase + NADP + PPi

Erg1 O 2 Squalene epoxidase H O 2

Erg7 Oxidosqualene cyclase

3O2 + 3NADPH Erg11 Lanosterol demethylase

+ 4H2O + 3NADP + formate

NADPH Erg24 C14 sterol reductase NADP+

Erg25 O2 + NADPH Methyl sterol monooxygenase + 2H2O + NADP

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O2 + Erg25 NADPH Methyl sterol monooxygenase 2H2O +

NADP+

O2 + Erg25 NADPH Methyl sterol 2 H2O + monooxygenase

NADP+

O2 + NADPH Erg26 H2O + C3 Sterol dehydrogenase NADP+

NADPH Erg27 3-Keto-sterol reductase NADP +

SAM Erg25 Methylsterol SAH monooxygenase

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NADPH

+ O2 Erg25

H2O + Methylsterol NADP+ monooxygenase

NADPH

+ O2 Erg25 Methylsterol 2 H2O + monooxygenase NADP+

NADPH

+ O2 Erg26 H2O + C13 sterol NADP+ dehydrogenase

NADPH Erg27

NADP+ 3-Keto-sterol reductase

SAM Erg6 C24 sterol SAH methyltransferase

Erg2 C8 sterol isomerase

NADPH Erg3 C5 sterol desaturase + O2 NADP+

+ 2H2O

NADPH

+ O2 Erg5 NADP+ C22 sterol desaturase

+ 2H2O

NADPH Erg4 C24 sterol reductase

NADP+