Regulation of Neutral Lipid Metabolism Through Phosphorylation of the Yeast Acyltransferase Gpt2

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2018-10-04 Regulation of neutral lipid metabolism through phosphorylation of the yeast acyltransferase Gpt2

Tavassoli, Marjan

Tavassoli, M. (2018). Regulation of neutral lipid metabolism through phosphorylation of the yeast acyltransferase Gpt2 (Unpublished doctoral thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/33158 http://hdl.handle.net/1880/108818 doctoral thesis

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Regulation of neutral lipid metabolism through phosphorylation of the yeast acyltransferase Gpt2

Marjan Tavassoli

A THESIS

SUBMITTED TO THE FACULTY OF GRADUATE STUDIES

IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE

DEGREE OF DOCTOR OF PHILOSOPHY

GRADUATE PROGRAM IN BIOLOGICAL SCIENCES

CALGARY, ALBERTA

SEPTEMBER, 2018

Abstract

Glycerol-3-phosphate acyltransferases (GPATs) catalyze the first step of glycerol-3-phosphate acylation at the sn-1 position producing lyso-phosphatidic acid. This is the committed and rate limiting step in de-novo synthesis of phosphatidic acid, the key intermediate in the glycerophospholipids and triacylglycerols (TAG) biosynthetic pathways. Two GPATs have been identified in S cerevisiae, Gpt2p and Sct1p. The role of phosphorylation in the serine-rich C- terminal tail of Gpt2p is unknown. In this work is shown that lack of phosphorylation on three conserved phosphorylation sites (S664, S668, S671) of Gpt2p alters protein stability, activity and neutral lipid metabolism. Specifically, a triple Gpt2p mutant (Gpt2-3A) where all these three residues were converted to alanine to mimic dephosphorylation induce a rise in the protein abundance, displayed more activity and resultedi in higher accumulation of DAG and TAG during exponential phase of growth. Lack of phosphorylation on Gpt2p delayed TAG lipolysis upon growth resumption from stationary phase which might be due to a futile TAG cycle that slows down mobilization of produced acyl-chains channeled for the formation of phospholipids.

Unregulated Gpt2p probably remains constitutively active and displaces Sct1p in exponential phase. Notoriously, Sct1p is found associated with lipid droplets in cells carrying Gpt2-3A mutant.

Considering Sct1p has never been identified in lipid droplets proteomes this consequence of having a constitutively de-phosphorylated Gpt2p might explain the alterations seen in neutral lipid metabolism of this mutant cells.

Preface

Chapter 3 of this thesis has been published as: Smart, Heather C., Fred D. Mast, Maxwell F.J. Chilije, Marjan Tavassoli, Joel B. Dacks, and Vanina Zaremberg. 2014. “Phylogenetic Analysis of Glycerol 3-Phosphate Acyltransferases in Opisthokonts Reveals Unexpected Ancestral Complexity and Novel Modern Biosynthetic Components.” PLoS ONE 9 (10). https://doi.org/10.1371/journal.pone.0110684.

Acknowledgements

I would like to thank my supervisor, Dr. Vanina Zaremberg for giving me the opportunity to peruse my goals in her research lab and for showing me how to elaborate my ideas and go beyond the limits. She gave me freedom to explore my scientific interests. I am very grateful for the scientific training I received in her lab. I would like to thank my committee members, Dr.

Gordon Chua and Dr. Elmar Prenner for their support and advices. I would also like to thank our research collaborator, Dr. Karin Athenstaedt in university of Graz. I would like to thank the former and present lab members, specially Maxwell Chilije, Suriakarthiga Ganesan, Brittney Shabits and

Laura Sosa for all your support and good memories that we had during my research work. Talking and working with you makes lab work so much more enjoyable. I would also appreciate all undergrad student helping me during my project, specially Anabel Cardenas, Ricky Chang and

Patrick Sipila. I would like to thank Dr. Jana Patton-Vogt, Dr. Shirin Bonni and Dr. Peter Tieleman for taking on the respective roles of external examiner and neutral chair of my defence exam.

Special thanks to my fiancé, Frashid, my best friend, partner, and love of life. Without your support and encouragement, I would not be able to finish this journey successfully. I would like to thank my dear parents and brothers for their love and care. Excited to see what the future has in store for us.

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Table of contents

Abstract ...... i

Preface ...... ii

Acknowledgement ...... iii

Table of Contents ...... iv

List of Abbreviations ...... ix

List of Figures ...... xiii

List of Tables ...... xviii

Chapter 1 – Introduction ...... 1

1.1 Introductory Remarks ...... 1

1.2 Major Lipid Classes in Eukaryotes ...... 2

1.3 Saccharomyces cerevisiae as model organism ...... 8

1.4 De novo Glycerolipid Synthesis in Yeast ...... 11

1.5 Glycerolipid catabolism, remodeling pathways ...... 18

1.5.1 Phospholipid remodeling ...... 18

1.6 Glycerol-3-Phosphate Acyltransferases (GPAT) ...... 20

1.6.1 Gpt2p and Sct1p Topology and Phosphorylation ...... 21

1.6.2 Gpt2p and Sct1p Substrate Specificity and Localization ...... 25

1.7 PA and DAG as Signaling Molecules ...... 28

1.8 TAG biogenesis and formation of lipid droplet ...... 32

1.9 Glycerolipid homeostasis during growth stages ...... 40

1.10 Signalling pathways that regulate lipid metabolism ...... 43

1.11 Hypothesis and Goals ...... 45

Chapter 2 – Materials and Methods ...... 47

2.1 Yeast Strains, Plasmids, Primers, Culture Conditions, and Transformations ...... 47

2.1.2 Gene cloning ...... 53

2.1.3 Site-directed mutagenesis ...... 54

2.1.4 Yeast crossing and tetrad dissection ...... 55

2.2 Cell lysate and microsomal fraction preparation, Protein Determination, SDS-PAGE, and Western Blot ...... 56

2.2.1 Cell Lysate preparation ...... 56

2.2.2 Microsomal Fractionation ...... 56

2.2.3 Protein determination, SDS PAGE and Western blot ...... 57

2.2.4 Phosphatase treatment experiment ...... 58

2.3 GPAT activity assay ...... 58

2.4 Quantitative Real-Time PCR ...... 59

2.4.1 Total RNA isolation ...... 59

2.4.2 Revers transcription ...... 60

2.4.3 Quantitative Real-time PCR ...... 61

2.5 Plating (growing) on different carbon source conditions ...... 62

2.5.1 Oleic acid experiment ...... 62

2.5.2 Plating on fermentable and non-fermentable carbon sources ...... 62

2.5.3 Gradual and acute glucose depletion experiment ...... 63

2.5.4 Different drug treatments ...... 63

2.5.4.1 Cerulenin ...... 63

2.5.4.2 Myriocin ...... 63

2.5.4.3 Aureobasidin A ...... 64

2.6 Microscopy analysis ...... 64

2.6.1 Lipid droplet staining with Nile red ...... 65

2.6.2 Lipid droplet staining with BODIPY 493/503 ...... 65

2.6.3 Lipid droplet analysis with AutoDot ...... 65

2.6.4 Cerulenin treatment ...... 66

2.7 Lipid droplet analysis with Typhoon method ...... 66

2.8 Yeast lipid extraction ...... 67

2.9 Neutral lipid Thin-layer Chromatography (TLC) experiment ...... 68

2.10 Yeast lipid droplet isolation and analysis ...... 69

2.11 Mass spectrometry analysis ...... 70

2.11.1 Clustering Analysis ...... 71

2.12 Comparative Genomic Survey ...... 73

2.13 Statistical analysis ...... 73

Chapter 3 - Phylogenetic study of fungal acyltransferases ...... 73

3.1 Introduction ...... 73

3.2 Goals ...... 74

3.3 Results ...... 75

3.3.1 Searching for Sct1p/Gpt2p identifies two undescribed deep clades of fungal GPATs

...... 75

3.3.2 Comparison of signature motifs between characterized and novel acyltransferases identified in this study ...... 80

3.4 Discussion and Conclusions ...... 85

Chapter 4 - Identification of phosphorylation sites in Gpt2p ...... 89

4.1 Introduction ...... 89

4.2. Goals ...... 90

4.3 Results and Discussions ...... 91

4.3.1 Identification of phosphorylation sites in Gpt2p ...... 91

4.3.2 Site directed mutagenesis of identified phosphorylation sites in Gpt2p ...... 96

4.3.3 Phenotypic characterization of Gpt2-phosphomutants ...... 102

4.4 Conclusion Remarks ...... 106

Chapter 5 - Metabolic impact of unregulated Gpt2p mutants ...... 108

5.1 Introduction ...... 108

5.2 Goals ...... 111

5.3 Results and Discussion ...... 111

5.3.1 Characterization of Gpt2 phospho-mutants expressed at endogenous levels 111

5.3.2 Unregulated Gpt2p- localization and impact on protein levels ...... 112

5.3.3 Unregulated Gpt2p- Impact on the timing of lipid droplet accumulation and morphology ...... 118

5.3.4 Unregulated Gpt2p- Impact on enzyme activity and TAG metabolism ...... 129

5.3.5 Concluding Remarks ...... 141

Chapter 6 - Role of Gpt2p phosphorylation during growth resumption from stationary phase ...... 142

6.1 Introduction ...... 142

6.2 Goals ...... 146

6.3 Results and Discussions ...... 147

6.3.1 Phenotypic characterization of unregulated Gpt2p phospho-mutants during calorie restriction regimes and growth resumption from stationary phase ...... 147

6.3.2 Unregulated Gpt2p alters diacylglycerol metabolism and distribution ...... 151

6.3.3 Proteomic analysis of Gpt2p enriched membranes and lipid droplets ...... 161

6.3.3.1 Assessing the importance of DAG/PA ration and LD formation in Gpt2p stability ...... 161

6.3.3.2 LD purification from cells re-entering growth from stationary phase ...... 165

6.3.3.3 Proteomic analysis of LDs obtained from stationary versus growth re-entry phases ...... 167

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Chapter 7 - Discussions and future directions ...... 182

7.1 Future Directions ...... 188

References ...... 190

Appendix A - Investigation of possible kinases phosphorylating Gpt2p ………………. 225 A.1 Introduction ………………………………………………………………………225 A.2 Goals ……………………………………………………………………………. 226 A.3 Results and discussion ……………………………………………………………226 A.4 Concluding remarks………………………………………………………………236 Appendix B – Copy right permissions ……………………………………………………237

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List of abbreviations

AbA; aureobasidin Acyl-CoA; acyl-coenzyme A Agos; Ashbya gossypii AMPK; AMP-activated protein kinase BLAST; protein-protein Basic Local Alignment Search Tool cAMP; cyclic-AMP CDP-DAG; CDP-diacylglycerol CDS; CDP-DAG synthase Cgla; Candida glabrata CK2; casein kinase 2 CL; cardiolipin DAG; diacylglycerol DGK; DAG kinase DHAP; dihydroxyacetone phosphate DHAPAT; dihydroxyacetone phosphate acyltransferase DTT; dithiothreitol ECL; enhanced chemiluminescence EPT; ethanolamine phosphotransferase ER; endoplasmic reticulum ERMES; ER mitochondria encounter structure EtBr; ethidium bromide FA; fatty acid FFA; free fatty acid FAS; fatty acid synthase G1/S; Gap 1/replication phase G-3-P; glycerol-3-phosphate

GAL; galactose GAM; goat anti-mouse GO; Gene Ontology GPAT; glycerol-3-phosphate acyltransferase GPC; glycerophosphocholine Gpt2p3xA; Gpt2p mutant with three serine residues, S664, S668, S671, mutated to alanine Gpt2pTrunc; Gpt2p mutant which lacks the serine-rich C-terminus (last 133 residues) GR; glucose restriction hPSK; human PSK HR; homologous recombination IPC; inositol phosphorylceramide KEGG; Kyoto Encyclopedia of Genes and Genomes Klac; Kluyveromyces lactis LC; liquid chromatography LC-MS/MS; liquid chromatography tandem mass spectrometry LCAT; lecithin:cholesterol acyltransferase LD; lipid droplet LiAc; lithium acetate LPAAT; lysophosphatidic acid acyltransferase Lyso-PA; lysophosphatidic acid Lyso-PC; lysophosphatidylcholine MAPK; mitogen-activated protein kinase MCS; membrane contact sites MS; mass spectrometry MS/MS; tandem mass spectrometry mTOR; mammalian TOR MUFA; monounsaturated fatty acid MW; molecular weight

Ni-NTA; Ni-nitrilotriacetic acid NP40; Nonidet P-40 NVJ; nuclear ER (nER)–vacuole junction OA; oleic acid ORF; open reading frame PA; phosphatidic acid Palmitic acid; C16:0 PAS; Per-Arnt-Sim domain PBS; phosphate buffered saline PC; phosphatidylcholine PE; phosphatidylethanolamine PG; phosphatidylglycerol PI; phosphatidylinositol PI3K; phosphatidylinositol 3-kinase PI(3)P; phosphatidylinositol 3-phosphate PI(3,4)P2; phosphatidylinositol 3,4-bisphosphate PI(3,5)P2; phosphatidylinositol 3,5-bisphosphate PI(4)P; phosphatidylinositol 4-phosphate PI(4,5)P2; phosphatidylinositol 4,5-bisphosphate PI(5)P; phosphatidylinositol 5-phosphate PKA; protein kinase A PKC; protein kinase C PLD; phospholipase D PM; plasma membrane PP1; protein phosphatase-1 PS; phosphatidylserine PSD; PS decarboxylase PSK; PAS kinase

PTM; post-translational modification PUFA; polyunsaturated fatty acid PVDF; polyvinylidene fluoride RT-qPCR; quantitative real-time PCR SACK; Saccharomyces cerevisiae, Ashbya gossypii, Candida glabrata, and Kluyveromyces lactis Scer; Saccharomyces cerevisiae SD; defined medium (2% glucose) SDS-PAGE; sodium dodecyl sulfate polyacrylamide gel electrophoresis SER; smooth ER SFA; saturated fatty acid Sgal; defined medium (2% glucose) SGD; Saccharomyces Genome Database SPT; serine palmitoyltransferase TAG; triacylglycerol TAP; tandem affinity purification TCE; trichloroethanol TF; transcription factor TLC; thin-layer chromatography TM; transmembrane TGN; trans Golgi network TOR; target of rapamycin TORC1; TOR complex 1 TORC2; TOR complex 2 Ura; uracil YNB; yeast nitrogen base YPD; yeast peptone dextrose

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List of Figures

Figure 1.1 Representative structures of the three eukaryotic lipid classes ...... 4 Figure 1.2 Structures of glycerophospholipids ...... 5 Figure 1.3 Fluctuations in storage and membrane lipids and acyl chain composition during yeast growth ...... 10

Figure 1.4 Reaction of glycerol-3-phosphate acyltransferases (GPATs) ...... 13 Figure 1.5 De novo synthesis of glycerolipids in yeast ...... 14 Figure 1.6 Biosynthesis and acyl chain remodeling of CL in yeast mitochondria ...... 17 Figure 1.7 Metabolism of PC in yeast ...... 19 Figure 1.8 Predicted topology of Gpt2p ...... 22 Figure 1.9 Putative phosphosites identified in global proteome studies ...... 23 Figure 1.10 Characteristic band pattern distinguishing phosphorylation species of Sct1p and Gpt2p ...... 25 Figure 1.11 Hydrogen bonding and hydrophobic interactions of phosphatidic acid (PA) and effector proteins ...... 31 Figure 1.12 Models of lipid droplet biogenesis ...... 34

Figure 1.13 Possible association between the ER membrane and LDs to facilitate the bidirectional exchange of integral membrane proteins between the two compartments ...... 36 Figure 2.1 pYES2.1/V5 His-TOPO cloning vector ...... 52

Figure 3.1 Opisthokont relationships and GPAT evolution ...... 76 Figure 3.2 Phylogeny of fungal GPAT homologues found in opisthokonts ...... 78 Figure 3.3 Phylogenetic tree of GDPAT-related genes in opisthokonts ...... 80 Figure 3.4 Sequence alignments of the catalytic motifs ...... 82 Figure 3.5. Sequence alignments of the catalytic motifs in strongly supported orthologs of fungal GPATs ...... 84 Figure 4.1 Predicted topology of Gpt2p ...... 90 Figure 4.2 Characteristic band pattern distinguishing phosphorylation species of Sct1p and Gpt2p ...... 92 Figure 4.3 Differential phosphorylation pattern in conditions of high vs low GPAT demand 94

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Figure 4.4 Sequence alignments of serine-rich C-terminal of Gpt2p related GPATs ...... 96 Figure 4.5 Characteristic band pattern of overexpressed Gpt2p phosphorylation mutants .... 97 Figure 4.6 Characteristic band pattern of overexpressed Gpt2p triple phosphomutants ...... 98 Figure 4.7 Characteristic band pattern of overexpressed Gpt2p carrying multiple mutations .99 Figure 4.8 Map of phosphorylation sites in the amino acid sequence of Gpt2p ...... 100 Figure 4.9 Overexpression of Gpt2-phosphomutants reduces yeast fitness ...... 102

Figure 4.10 Overexpression of unregulated Gpt2p variants increases LD size in exponential phase ...... 103 Figure 4.11 Migration of endogenously expressed Gpt2p phospho-deficient mutants ...... 104 Figure 4.12 Characteristic band pattern of Gpt2p carrying multiple mutations expressed at endogenous levels ...... 105 Figure 5.1 Gpt2p enriched ER structures interact with lipid droplets ...... 110 Figure 5.2 Endogenous levels of Gpt2p phosphomutants ...... 112 Figure 5.3 Live imaging of Gpt2p phospho-mutants during exponential and stationary phases of growth ...... 114 Figure 5.4 Protein analysis of GFP-tagged proteins in exponential phase ...... 115 Figure 5.5 Lack of Sct1p alleviates degradation of Gpt2 in stationary phase ...... 117 Figure 5.6 Gpt2p is enriched in ER-domains in late exponential growth ...... 118 Figure 5.7 Co-localization analysis of Gpt2p and lipid droplets in log and stationary phases of growth ...... 121 Figure 5.8 Gpt2-3A domains are intimately associated with lipid droplets ...... 122 Figure 5.9 Lack of phosphorylation induces Gpt2p association with lipid droplets ...... 123 Figure 5.10 Gpt2-3A cells have larger LDs in exponential phase of growth ...... 125 Figure 5.11 Analysis of lipid droplet population in Gpt2p phosphomutants ...... 126 Figure 5.12 Size distribution of lipid droplets in Gpt2p phosphomutants ...... 126 Figure 5.13 Analysis of lipid droplet morphology in the absence of Sct1p ...... 129 Figure 5.14 Neutral lipid content in 3A-phosphomutant and wild type cells ...... 130

Figure 5.15 Constitutive dephosphorylation in Gpt2-3A impacts GPAT activity ...... 131 Figure 5.16 Phosphorylation deficiency impacts TAG levels in the absence of Sct1p ...... 133

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Figure 5.17 TAG and DAG levels in strains expressing endogenous levels of wild type or 3A Gpt2p in the presence or absence of Sct1p ...... 134 Figure 5.18 Phosphorylation pattern of wild type and 3A Gpt2 catalytically-dead versions . 135 Figure 5.19 Accumulation of DAG (but not TAG) correlates with the toxic effect of unregulated Gpt2p overproduction ...... 137 Figure 5.20 Lack of phosphorylation on endogenously GPT2 does not change cell response to oleic acid ...... 138 Figure 5.21 Overexpression of unregulated Gpt2p induces sensitivity to oleic acid ...... 139 Figure 5.22 Unregulated Gpt2p induces accumulation of oversized but few LDs ...... 141 Figure 6.1 Cycles of TAG metabolism during yeast growth ...... 144 Figure 6.2 Survival of Gpt2p phospho-mutants expressed at endogenous levels during gradual and acute calorie restriction regimes ...... 148 Figure 6.3 Increased cytosolic lipolysis buffers Gpt2-3A toxicity ...... 148 Figure 6.4 Lack of phosphorylation on Gpt2p shows delayed TAG lipolysis detected by tracking stained lipid droplets morphology with BODIPY 493/503 over time ...... 151 Figure 6.5 DAG rich vacuoles associate with lipid droplets during growth resumption ...... 156 Figure 6.6 DAG rich vacuoles from wild type and Gpt2-3A cells ...... 157 Figure 6.7 DAG enriched puncta are not lipid droplets ...... 158 Figure 6.8 DAG polarization upon growth resumption is delayed in Gpt2-3A cells ...... 161

Figure 6.9 Decreased vacuolar degradation levels of Gpt2 in elevated levels of PA in exponential phase ...... 163 Figure 6.10 Lack of Nem1p phosphatase induces changes in Gpt2p abundance and a shift in the phosphorylation pattern of the unregulated Gpt2p3xA ...... 164 Figure 6.11 LD purification from wildtype and Gpt2-3A cells during growth resumption from stationary phase ...... 167 Figure 6.12 Proteome of LDs from stationary phase wild type cells identified by LC-MS/MS compared to the literature ...... 168 Figure 6.13 Proteome analysis of wild type and Gpt2-3A LDs obtained from stationary phase ...... 169 Figure 6.14 Enrichment categories of unique proteins (57) found in Gpt2-3A LDs obtained from stationary phase...... 173

Figure 6.15 Proteome analysis of wild type LDs obtained from stationary phase versus growth re- entry ...... 174

Figure 6.16 Enrichment categories of unique proteins (167) found in wild type LDs obtained during growth resumption...... 174 Figure 6.17 Comparison between wild type and Gpt2-3A proteins identified in LDs from growth resumption ...... 175 Figure 6.18 Glycerophospholipid metabolism identified as the only lipid metabolic category in unique protein dataset from Gpt-3A compared to wt ...... 175

Figure 6.19 Enrichment categories of proteins (58) found in both Gpt2-wt and Gpt2-3A LDs obtained from growth resumption ...... 176 Figure 6.20 Comparison between LD-enriched proteins found in both Gpt2-wt and Gpt2-3A cells during growth resumption (from figure 6.17) versus proteins found in Gpt2-wt LD during stationary phase ...... 177 Figure 6.21 Enrichment categories of unique proteins to Gpt2-wt LD found in stationary phase and unique proteins enriched with LDs during growth resumption in Gpt2-wt and Gpt2-3A cells ...... 177 Figure 6.22 Overexpression of wild type and phosphomutant Gpt2-3A in cells lacking LPAAT enzymes...... 178 Figure 6.23 Effect of Myriocin on cells expressing different sets of GPATs ...... 180 Figure 6.24 Effect of Aureobasidin A on cells expressing different sets of GPATs ...... 181 Figure 6.25 Model explaining the combined phenotypes of resistance and sensitivity to Myriocin and Aureobasidin A ...... 181

Figure 7.1 Proposed model for contribution of Gpt2p and Sct1p during three different growth phases in regulated and unregulated Gpt2 backgrounds ...... 183 Figure 7.2 Proposed model suggesting the presence of a futile TAG cycle during growth resumption in Gpt2p-3A ...... 184 Figure 7.3 Increased levels of GPT2 gene expression in stationary phase ...... 185 Figure 7.4 Overexpression of Gpt2p lacking phosphorylation increases degradation of Sct1 in exponential phase ...... 186

Figure A.1 Psk1p-TAP associates with microsomes when grown to stationary phase in the presence of galactose or raffinose, but not glucose ...... 228 Figure A.2 Lack of Psk1 kinase did not change phosphorylation pattern in Gpt2p-wt and Gpt2p- 3A ...... 230

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Figure A.3 Phosphorylation pattern and abundance level of Gpt2 in the absence of Snf1 kinase ...... 231 Figure A.4 Signaling pathways responding to glucose levels ...... 232 Figure A.5 Growth pattern of Gpt2p phosphomutants in the presence of Rapamycin ...... 233 Figure A.6 Regulation of sphingolipids biosynthesis through TORC2/Ypk1p signaling ...... 234 Figure A.7 Phosphorylation pattern of Gpt2p-wt and Gpt2p-3A in the absence of Ypk1 ..... 235

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List of Tables

Table 2.1 Yeast strains used in this study ...... 47 Table 2.2 Primers used in this study ...... 50 Table 3.1 Names of species and specific strains used in the proteomic analysis of opisthokont GPATs...... 87 Table 3.2 Acyltransferase motifs and distance between motifs (DBM) in microsomal yeast . 88 Table 4.1 Results from MS analysis ...... 95 Table 4.2 Phosphorylated amino acids identified in Gpt2p ...... 101 Table 6.1 Proteins found in both wt and Gpt2-3A enriched LDs from stationary phase never reported to be associated with LDs before ...... 170 Table 6.2 Proteins found in Gpt2-3A enriched LDs from stationary phase never reported to be associated with LDs before …………………………………………………………………. 170

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Chapter 1 - Introduction

1.1 Introductory Remarks

Lipid metabolism is conserved in eukaryotes, from yeast to humans. The role of lipids as mediators of signaling pathways that control cell proliferation and cell death is also conserved. Impaired lipid homeostasis contributes to the pathogenesis of several neurodegenerative disorders, including Alzheimer’s disease and Parkinson’s disease (Xu et al., 2017) and metabolic disorders, such as obesity, diabetes and atherosclerosis which are associated with excessive exposure of cells to fatty acids (Guo et al. 2009; Beller et al.

2010). Over the last decades, a strong increase in the prevalence of these diseases has led to intense efforts to deepen our knowledge on how cells regulate uptake, biosynthesis, storage and mobilization of lipids. Alterations of these pathways are not only underlying causes of many diseases, they can also be exploited for biotechnological developments like industrial production of oils and biofuels (Sheng et al., 2015). Channeling excess fatty acids into glycerolipids is a cellular strategy to maintain lipid homeostasis. Glycerolipids are a class of lipids that include membrane phospholipids and storage lipids like triacylglycerols (TAGs). Research conducted in the model organism Saccharomyces cerevisiae has been at the vanguard of the identification of the enzymes involved in the metabolism of glycerolipids and its regulation (Henry et al., 2012). The initial two sequential acylations of glycerol 3-phosphate using acyl-CoA as an acyl donor to produce phosphatidic acid (PA) are conserved steps in the de novo synthesis of glycerolipids.

Glycerol 3-phosphate acyltransferases (GPATs) catalyze the first committed and rate limiting step in the de novo synthesis of PA. PA represents a critical branching point in the synthesis of all glycerolipids. PA has surfaced as not only the precursor for all

1 glycerolipids, but also as a potent signaling lipid able to integrate cellular cues to balance synthesis of phospholipids for membrane expansion versus storage. The PA-dependent transcriptional regulatory circuit that controls expression of enzymes involved in glycerolipid synthesis downstream of the PA branching point has been very well characterized in yeast (White et al., 1991; Yamashita et al., 2001). Less is known about the regulation of the upstream initial steps leading to PA synthesis. Two yeast GPATs named

Gpt2/Gat1 and Sct1/Gat2 contribute to the production of different pools of PA in concert with other pathways that produce and consume acyl-CoAs (Zaremberg et al., 2002;

Bratschi et al., 2009; De Smet et al., 2012; Marr et al., 2012). This thesis focuses on the study of the regulation of the yeast GPAT Gpt2p through phosphorylation. More specifically, investigations were directed towards understanding the role that this post- translational modification plays in regulating localization, protein stability and interactions of Gpt2p. The results unveiled a novel contribution of Gpt2p to TAG metabolism, with a central role of this GPAT in favoring anabolic vs catabolic pathways depending on its phosphorylation status.

1.2 Major Lipid Classes in Eukaryotes

Lipids, or fats, are an important source of energy in eukaryotic cells. Lipids are constantly broken down, metabolized and used for energy production and replenished from available nutrients. Lipids are defined as organic substances which are soluble in nonpolar organic solvents, but not in water. They are small biomolecules that vary in their physical properties from highly hydrophobic to amphiphilic. According to their molecular structure and function, lipids are categorized into eight classes which are fatty acids (FA),

2 glycerolipids, glycerophospholipids, sterols and sterol derivatives, sphingolipids, prenol lipids, glycolipids, and polyketides (Dennis et al., 2009; Fahy et al., 2009). Lipids not only serve as building blocks of biological membranes, they also play roles as signaling molecules, energy source or as mediators of membrane fusion and fission in the dynamic behavior of organelles (Escribá et al., 2008; Rego et al., 2014). There are three major classes of lipids in eukaryotes, glycerolipids, sphingolipids (SLs), and sterols (Figure 1.1), whose biochemical and biophysical properties vary considerably and impact upon their function.

Glycerolipids contain a glycerol backbone as they are derived from glycerol-3- phosphate (G-3-P). G-3-P can be esterified with one or two fatty acids to the sn-1 and sn-

2 positions and make up monoacyl- and diacyl- (glycerophospholipids) or can be found with no phosphate headgroup, esterified on all three sn-positions as TAG (a neutral lipid).

Figure 1.1- Representative structures of the three eukaryotic lipid classes. Glycerolipids contain a glycerol backbone (highlighted by the blue box), while sphingolipids contain a sphingosine backbone (red box). Sterols are composed of a four- ring structure. R represents variable headgroups.

Classification of glycerophospholipids is according to the structure of their headgroup, which is zwitterionic in phosphatidylcholine (PC) and phosphatidylethanolamine (PE), and anionic in phosphatidylinositol (PI), phosphatidylserine (PS), phosphatidic acid (PA), phosphatidylglycerol (PG), and cardiolipin (CL) (Figure 1.2).

Figure 1.2- Structures of glycerophospholipids. CxHy, CnHm, CiHj, CkHl corresponding to the acyl chains. Adapted from (De Kroon et al., 2013).

Next, a brief overview of glycerophospholipids with emphasis in their behavior and abundance in budding yeast is provided.

Phosphatidic acid (PA) is the smallest and simplest glycerophospholipid which is negatively charged at physiological pH. A small, highly charged head group whose charge lies very close to the glycerol backbone provides a unique cone shape in PA. This structural specificity allows PA to play a significant role in membrane vesicle fission and fusion events by inducing high membrane curvature. PA can also affect adjacent proteins activity by introducing negative curvature in membranes (Stillwell, 2016). PA is a key intermediate

5 glycerophospholipid in glycerolipid biosynthesis, although its amount is quantitatively negligible at 1 mol% of membrane lipids. PA has an important regulatory function impacting membrane lipid synthesis and composition. It can play a central role in glycerophospholipid synthesis, in addition to diverse functions in membrane dynamics and lipid signaling that highlight the physiological importance of PA (Kooijman et al., 2009).

PC is the most abundant glycerophospholipid in most eukaryotic cells including budding yeast. PC is essential for yeast growth (Kodaki et al., 1989; Zaremberg et al.,

2002; De Kroon et al., 2007). PC with a cylindrical shape is a typical bilayer lipid structurally related to PE as their headgroups carry an aminoalcohol. Both PC and PE are zwitterionic however, PC headgroup is larger than that of PE, with three methyl groups attached to the amine in PC. The role of PC for stabilizing the membrane bilayer is remarkable as depletion of PC in yeast triggers a cellular response that involves an integration of PE with shorter and more saturated acyl chain to increase its bilayer propensity (De Kroon et al., 2013).

PE is another essential phospholipid in yeast (Storey et al. 2001). PE can play an important role in mitochondrial function in yeast, as deletion of PE results in defects in mitochondrial respiration, impaired assembly of mitochondrial protein complexes and loss of mitochondrial DNA (Birner et al. 2001; Koltzscher et al. 2003). PE is also involved in delivery of cytoplasmic proteins to the vacuole by autophagy (Ichimura et al. 2000), and in the transport of the amino acid transporters to the plasma membrane (Opekarová et al.,

2002). PE can induce an inverted hexagonal (HII) phase at physiological conditions. The tendency of PE in formation of non-bilayer structures is related to its smaller head group and acyl chain composition with double bonds and longer acyl chains participating in

6 negative membrane curvature (Cullis et al., 1986; Lindblom et al., 1992; De Kroon et al.,

2013).

PI is another abundant membrane glycerophospholipid which is negatively charged at physiological pH. PI is an essential lipid both as a key membrane constituent and as a participant in essential metabolic processes. PI by serving as a biosynthetic precursor in formation of phosphoinositides plays a critical role in cellular signal transduction pathways, as effector proteins containing phosphoinositide-binding domains are recruited to specific membranes in signaling cascades (Strahl et al., 2007).

PS is a quantitatively minor glycerophospholipid with negative charge, but as a precursor of PE and PC can play an essential role in the cytidine diphosphate-DAG (CDP-

DAG) pathway. Since synthesis of PE and PC is sustained, PS is not an essential glycerophospholipid in yeast (Birner et al., 2001).

PG is a minor mitochondrial glycerophospholipid in yeast which is negatively charged; and is the precursor for cardiolipin (CL) biosynthesis by CL synthase (De Kroon et al., 2013). CL is an anionic glycerophospholipid in yeast which is localized in the mitochondrial inner membrane (Daum, 1985; Janssen et al., 1999). CL, the signature lipid of mitochondria, plays a critical role in mitochondrial function and biogenesis. The unique structure of CL is composed of two 1,2-diacylphosphatidate moieties that are connected to the 1- and 3-hydroxyl groups of a single glycerol headgroup (Gebert et al. 2009). Although

CL is not essential in yeast (Jiang et al., 1997), its function is critical for mitochondrial structure and activity of enzymes imbedded in mitochondrial membranes.

Sphingolipids are a class of lipids containing a backbone of sphingoid bases, a set of aliphatic amino alcohols that includes sphingosine which can bind to a (usually) charged

7 headgroup and a FA (Figure 1.1). Ceramides, the simplest sphingolipids, which contain an acyl chain and no headgroup make up the early products of the sphingolipid biosynthetic pathway. Complex sphingolipids are formed from binding of ceramide to different headgroups, such as sugars or phospho groups. Both glycerolipids and sphingolipids are amphipathic, with hydrophilic headgroups and hydrophobic acyl tails. Complex sphingolipids in yeast are based on phosphoinositol headgroups.

Finally, sterols are composed of a four-ring planar structure, with a hydroxyl group at 3-position of the A-ring (Figure 1.1). In yeast, the principal sterol is ergosterol which is the fungal analog of cholesterol in mammals. Sterols play essential roles in cell physiology, from membrane building blocks to signaling during development (Bidet et al., 2011).

Ergosterol regulates membrane fluidity, plasma membrane biogenesis and function (Yang et al., 2015). Ergosterol homeostasis is critical for fungal cells. Regulatory machinery involves the transcription control of genes encoding ergosterol biosynthetic enzymes and proteins that perform sterol processing and uptake (Vik et al., 2001). Regulatory proteins in some fungi species, such as Schizosaccharomyces pombe, share common features with the mammalian sterol regulatory element-binding proteins (SREBPs) (Hughes et al., 2005;

Porter et al., 2010). However, many other fungal species, including Saccharomyces cerevisiae and Candida species, do not have mammalian homologues of cholesterol regulator proteins (Maguire et al., 2014).

1.3 Saccharomyces cerevisiae as model organism

Lipid metabolism is conserved in both yeast and humans and the pathways of glycerolipid metabolism in Saccharomyces cerevisiae (budding yeast) have been

8 elucidated with few remaining gaps in knowledge. Therefore, the findings discovered by studying lipid metabolism in yeast can be subsequently applied to humans or other eukaryotes. Budding yeast has long been used as a model organism in lipid metabolism research addressing how it impacts regulation of growth, development, and cellular homeostasis (Agmon et al., 2017).

Working with yeast as a model organism has many advantages, including the accessibility of powerful and unique genetic and biochemical approaches. Since its genome was entirely sequenced (Goffeau et al., 1996), S. cerevisiae has been at the forefront of the development of functional genomic, transcriptomic, proteomic, and metabolomic approaches, resulting in vast amounts of high-throughput data and strains collections.

Currently, S. cerevisiae is the organism with the most comprehensive and well-curated experimental dataset available to the research community (Cherry et al., 2012).

Yeast lipid metabolism is actively changing throughout its growth cycle. Re-entry to growth from stationary phase in the presence of glucose as a carbon source requires a transient lag phase for cells to adjust their metabolism and start proliferation (Figure 1.3).

Figure 1.3- Fluctuations in storage and membrane lipids and acyl chain composition during yeast growth. A) Cell density (OD/mL) and extracellular glucose and ethanol concentrations. B) Temporal dynamics of storage and membrane lipids. Shaded areas correspond to phases of proliferation. C) Temporal profile of saturated (SFA) and monounsaturated (MUFA) fatty acid moieties in the yeast lipidome. All results are expressed as the mean ± SD of five biological replicates. Adapted from (Casanovas et al. 2015).

This lag phase is followed by the exponential phase, when cells are actively proliferating by fermenting glucose to ethanol. Once the level of glucose becomes limiting, cells enter a phase identified as the diauxic shift characterized by decreased growth rate, where metabolism is again reprogrammed to support growth in the post-diauxic phase by using cellular respiration of ethanol and fatty acid oxidation. Finally, cells enter the quiescent stationary phase and growth ceases (Galdieri et al., 2010). As cells grow and

10 divide or enter the non-replicative stationary phase, the specific requirements for storage and membrane lipids fluctuate. During these periods, it is essential to control the metabolic flux of fatty acids into either phospholipids (proliferative state) or into TAG (nutrient limitation conditions) (Kohlwein, 2010). The ratio between membrane and storage lipids oscillates during growth. Synthesis of neutral lipids for storage [TAG and sterol esters

(SE)] increases as cells enter stationary phase while are consumed immediately when cells exit the stationary phase, and enter a growth period upon supplementation with glucose

(Figure 1.3) (Kohlwein, 2010). Breakdown of TAG and SE provides fatty acids (FAs) and

DAG to be used for the synthesis of membrane lipids to support rapid initiation of cellular growth and division. (Cherry et al., 2012; Kohlwein et al., 2013). TAG synthesis and turnover is regulated at many levels, including gene expression controlled by phospholipid precursors, compartmentalization, and posttranslational modifications of enzymes (Henry et al., 2012).

1.4 De novo Glycerolipid Synthesis in Yeast

Different lipid classes are modulated by lipid metabolic pathways. Despite the importance of lipids, they only recently came into focus of systematic studies, aiming to understand the contribution of lipid to a fully functional cell (Schützhold et al., 2016). The mechanisms regulating lipid metabolic pathways have been partially clarified, although molecular aspects of lipid partitioning across organelles, the recruitment of effector proteins by signaling lipids, and understanding how lipid levels are spatiotemporally regulated remain major challenges in the field. Lipidomics studies conducted in the last decade have revealed that biological membranes are more heterogeneous than was

11 previously thought, made up of complex of membrane lipids diverse in structure, and the distribution of different lipids and their species (membrane lipid composition) varies at the organism, cell type, organelle, membrane, bilayer-leaflet and membrane subdomain level

(Zhang et al., 2017). Structural/compositional changes of membrane lipids are known to directly affect the cell membrane integrity, which is important for regulating the cell signaling transduction and chemically-isolated intracellular environment (Hendricks et al.,

2014). Biomechanical studies on cancer cells (Lam et al., 2009; Zhang et al., 2002) have shown that various types of cancer cells exhibit lower fluidity due to the structural and compositional alterations in cellular bio-membranes.

Lipid composition might affect transmembrane protein recruitment, conformation, activity and localization through specific lipid–protein interactions or through their effects on membrane properties (Harayama et al., 2018). These lipids are dynamically regulated by phosphorylation and dephosphorylation, which allows them to recruit proteins in a concerted manner (De Craene et al., 2017). PA, DAG and PS are known to recruit proteins

(Lemmon, 2008). Therefore, identifying the cellular pools of these glycerolipids allows us to understand the mechanism regulating this diversity in bio-membranes.

Acyltransferases can play a critical role in channeling of specific FAs into distinct pools of lipids. This thesis focuses on the acyltransferases that initiate the synthesis of PA in yeast.

De novo synthesis of glycerolipid starts with acylation of the sn-1 position of glycerol-3-phosphate (G-3-P) by glycerol-3-phosphate acyltransferases (GPATs) and formation of lysophosphatidic acid (lyso-PA) (Figure 1.4).

Figure 1.4- Reaction of glycerol-3-phosphate acyltransferases (GPATs). The GPATs transfer an acyl group from an acyl-CoA molecule to the sn-1 position of glycerol-3- phosphate (G-3-P) to make lysophosphatidic acid (lyso-PA).

This reaction is the rate-limiting and committed step in the pathway for synthesis of PA and downstream glycerolipids. Gpt2p and Sct1p are the yeast GPATs (Zaremberg et al., 2002; Zheng et al., 2001). Addition of another acyl group to the sn-2 position by lyso-

PA acyltransferases (LPAATs) forms PA (Figure 1.5), the simplest glycerophospholipid

(Han et al., 2007; Miner et al., 2017; Ouahoud, 2018). LPAATs Ale1 and Slc1 are known to catalyze this acylation step in yeast.

Figure 1.5- De novo synthesis of glycerolipids in yeast. De novo synthesis of glycerolipids in yeast is initiated by the sequential acylation of glycerol-3-phosphate (G3- P) or dihydroxyacetone phosphate (DHAP) to produce lysophosphatidic acid (lyso-PA) and then phosphatidic acid (PA). PA represents a branching point as it can be converted to diacylglycerol (DAG) feeding the Kennedy pathways (blue pathways) or to CDP-DAG (green pathways). Mitochondrial pathways are represented within double-membrane- enclosed compartments. Lipid droplet (purple) is the storage compartment for triacylglycerol (TAG). PC, phosphatidylcholine; PI, phosphatidylinositol; PE, phosphatidylethanolamine; PS, phosphatidylserine; PG, phosphatidylglycerol; CL, cardiolipin. Taken from (Zaremberg et al., 2017).

PA is a crucial branching molecule which is used in two glycerophospholipids biosynthesis pathways. It can be used in CDP-DAG pathway or the Kennedy pathway

(Figure 1.5). The CDP-DAG pathway produces major glycerophospholipids PC, PI, PE and PS (Carter et al., 1966; Shen et al., 1996). CDP-DAG is an energy-rich intermediate formed from PA by CDS1-encoded CDP-DAG synthase. The CMP moiety in CDP-DAG can be replaced by inositol to form PI by the PIS1-encoded PI synthase and also can be replaced with serine by the CHO1-encoded PS synthase to form PS (Nikawa et al., 1984;

Gene, 1987). The produced PS can be decarboxylated by the PSD1- and PSD2-encoded PS

14 decarboxylases to form PE, which then gets methylated on its amine group in three-step by

S-adenosyl-l-methionine (Ado-Met) dependent methylation reactions to form PC (Clancey et al., 1993; Trotter et al., 1993). The first methylation on PE is catalyzed by the CHO2- encoded PE methyltransferase to form phosphatidylmonomethylethanolamine (PME) intermediate. The last two steps of methylation produce phosphatidyldimethylethanolamine intermediate and PC, respectively, which both are catalyzed by OPI3-encoded phospholipid methyltransferase (Kodaki et al., 1987; Summers et al., 1988; Mcgraw et al., 1989).

The Kennedy pathway is responsible for de novo synthesis of PC and PE (Smith et al., 1957). The two characteristic high-energy intermediates: CDP-ethanolamine and CDP- choline control production of PE and PC resulting in formation of two branches of Kennedy pathway known as CDP-ethanolamine and CDP-choline, respectively. As the first step, ethanolamine and choline can get phosphorylated by EKI1-encoded ethanolamine kinase and by CKI1-encoded choline kinase to form phosphoethanolamine and phosphocholine, respectively (Hosaka et al., 1989; Kim et al., 1999). In the next step, these formed intermediates are activated by addition of CTP group by the ECT1-encoded phosphoethanolamine cytidylyltransferase and by the PCT1-encoded phosphocholine cytidylyltransferase to form CDP-ethanolamine and CDP-choline, respectively

(Tsukagoshi et al., 1987; Min-seok et al., 1996). Finally, PE and PC are formed from reaction of CDP-ethanolamine and CDP-choline with DAG catalyzed by the EPT1- encoded ethanolamine phosphotransferase and by CPT1-encoded choline phosphotransferase, respectively (Hjelmstad et al., 1988; Hjelmstad et al., 1991; Gene,

1987). PA can be converted to DAG by PA phosphatases (encoded by APP1, DPP1, LPP1

15 and PAH1 genes) (Figure 1.5). The reverse reaction is catalyzed by DAG-kinase (encoded by DGK1) (Han et al., 2008). DAG is the substrate for Lro1p, the yeast orthologue of mammalian lecithin:cholesterol acyltransferase (LCAT), which is associated with ER with its active site exposed to the lumen (Jacquier et al., 2011). Lro1p is a transacylase that produces TAG and lysophospholipids as byproducts. Dga1p, the orthologue of mammalian

DGAT2, is the second major acyltransferase involved in TAG synthesis that localizes mostly to the ER and also associates with LDs (Dahlqvist et al., 2000).

Besides the de novo pathway, there are alternate routes where turnover of other lipids produces pools of PA and DAG, usually functioning as second messenger molecules in signaling (Foster et al., 2014). These pathways can be activated in response to growth factors, changes in nutrients, and stress (Foster et al., 2014; Wang et al., 2006; Testerink et al., 2011; Arisz et al., 2013). PA for example, can be formed from phosphorylation of

DAG by DAG kinases, in which the DAG results from the hydrolysis of phospholipids such as phosphoinositides by phospholipase C (PLC) or from TAG lipolysis (Wang et al.,

2006). Additional pools of DAG are produced from complex sphingolipid biosynthesis

(Cerbon et al., 2005). PA can also be produced from the hydrolysis of PC by phospholipase

D (PLD) (Jenkins et al., 2005).

PA needs to be transported into mitochondria to be used as precursor in the formation of cardiolipin (CL) in the matrix face of the mitochondrial inner membrane

(Figure 1.6). PA produced in the ER or at the interphase with LDs is transported to the inner mitochondria membrane for CL biosynthesis.

Figure 1.6- Biosynthesis and acyl chain remodeling of CL in yeast mitochondria Enzymes highlighted in blue. PA, phosphatidic acid; CDP-DAG, CDP-diacylglycerol; PGP, phosphatidylglycerolphosphate; PG, phosphatidylglycerol; MLCL, monolysocardiolipin; PL, phospholipid; MOM, mitochondrial outer membrane; MIM, mitochondrial inner membrane.

The Ups1p/Mdm35p complex localized to the mitochondrial intermembrane space has been identified as a PA transporter from the mitochondrial outer membrane to the inner one (Connerth et al., 2012; F. Yu et al., 2015; Watanabe et al., 2015; Miliara et al., 2015).

Tam41p catalyzes the first step of the pathway, namely, the synthesis of CDP- diacylglycerol (CDP-DAG) from PA and CTP (Tamura et al., 2013). Phosphatidylglycerol phosphate (PGP) can be formed from CDP-DAG and glycerol 3-phosphate catalyzed by

PGP synthase (Pgs1p) (Chang et al., 1998; Džugasová et al., 1998). PGP becomes dephosphorylated by Gep4p and produces phosphatidylglycerol (PG) (Osman et al., 2010).

Finally, a cardiolipin synthase, Crd1p, catalyzes the CL formation from PG and CDP-DAG

(Jiang et al., 1997; Tuller et al., 1998).

1.5 Glycerolipid catabolism, remodeling pathways

Each glycerophospholipid class has distinct physicochemical features acquired by the composition of acyl chain (Renne et al., 2015). Acyl chain composition of membrane glycerophospholipids determine physical properties of a membrane, such as fluidity, surface charge, intrinsic curvature (De Kroon et al., 2013) reflecting diversity in the functions of organelles which requires regulation of lipid homeostasis. Homeostasis of cellular membrane lipid is controlled by de novo biosynthesis, remodeling, intracellular mobilization, and degradation of glycerolipids.

1.5.1 Phospholipid remodeling

The synthesis and turnover of PC are highly regulated in all eukaryotic cells (Dowd et al., 2001). In yeast, PC is synthesized through the CDP-choline pathway when choline is supplied exogenously. In the absence of exogenous choline, PC is catabolized from the triple methylation process on PE while the CDP-choline pathway is mostly involved in recycling choline from PC turnover. The yeast phospholipase D Spo14p (Sreenivas et al.,

1998) converts PC into choline and PA. Alternatively, choline can be produced by the combined function of the phospholipases B (PLB) Plb1p (Lee et al., 1994), or Nte1p

(Zaccheo et al., 2004) with the phosphodiesterase Gde1p (Fernández-Murray et al., 2005;

Fisher et al., 2005) (Figure 1.7).

Figure 1.7- Metabolism of PC in yeast. Solid arrows showing acyl chain remodeling, degradation pathways. Enzymes are highlighted in blue. PC can be catabolized by (1) phospholipase B (PLB) Plp1p and/or Nte1p; (2) phospholipase D (PLD) Spo14p. Deacylation of PC by PLB forms glycerophosphocholine (GPC) and free fatty acids, while PLD hydrolyzes PC to phosphatidic acid (PA) and choline. GPC phosphodiesterase (Gde1p) can hydrolyze GPC to choline and G-3-P. Lysophosphatidylcholine (lyso-PC) produced by the action of PLA can be reacylated by the acyltransferase Ale1p in an acyl- CoA-dependent manner or can be converted to GPC by PLB. The biosynthesis routes of PC are indicated by the dashed arrows. Modified from (Renne et al., 2015).

Acyl chain remodeling of PC can proceed by deacylation to lyso-PC and/or GPC catalyzed by the PLBs Plb1p and Nte1p and/or by PLA and/or PLB. Subsequently, lyso-

PC can be reacylated by the 1-acyl lyso-PC acyltransferase Ale1p.

The CDP-choline route produces a more diverse PC species profile, while methylation of PE primarily generates di-unsaturated PC (Boumann et al., 2003, 2004).

Regulation of synthesis and turnover of PC is essential in all eukaryotic cells as altered metabolism of PC leads to different diseases. Impaired CDP-choline pathway, either genetically (Cui et al., 1996) or treated by enzyme inhibitors (Anthony et al., 1999),

19 causes apoptosis in several mammalian cell types. In Alzheimer’s disease, brain cells showed elevated levels of glycerophosphocholine (GPC) (Nitsch et al. 1992).

Metabolism studies on glycerophospholipids in yeast indicates a tight relation between synthesis and degradation of different glycerophospholipids, especially PC metabolism that plays a critical role in this metabolic network. For instance, a study for identifying the acyltransferases and phospholipases involved in acyl chain exchange of PC showed that deletion of the SCT1-encoded GPAT exhibited reduced PC remodeling (De

Smet et al., 2012).

1.6 Glycerol-3-Phosphate Acyltransferases (GPAT)

The first step in the synthesis of almost all membrane phospholipids and neutral glycerolipids is catalyzed by glycerol-3-phosphate acyltransferases (GPATs). This step is committed and rate-limiting step and is redundant in Saccharomyces cerevisiae, mammals, and plants (Smart et al., 2014). GPAT transfers a fatty acid from fatty acyl coenzyme A to the sn-1 position of glycerol-3-phosphate (G-3-P) to produce lyso-PA. Subsequently, lyso-

PA can be further acylated at the sn-2 position by a separate acyltransferase to produce PA

(Figure 1.4).

There are four motifs (motif I, II, III, and IV) found in GPATs, which are responsible for catalysis and substrate binding (Smart et al., 2014; Lewin, Wang et al.,

1999). There are four mammalian GPATs; each encoded by a different gene (Bell et al.,

1980; Lewin et al., 2004; Cao et al., 2006). Two of the mammalian GPATs are localized to the mitochondria (GPAT1 and GPAT2) and two are localized to the ER (GPAT3 and

GPAT4) (Bell et al., 1980; Lewin et al., 2004; Cao et al., 2006). Yeast has two GPAT

20 isoforms, Gpt2p and Sct1p, encoded by GPT2 and SCT1, respectively. Simultaneous loss of function of GAT1 and GAT2 is synthetically lethal (Zaremberg et al., 2002; Zheng et al.,

2003). GPATs are integral membrane proteins localized to the ER membrane (Zheng et al., 2001; Bratschi et al., 2009). Interestingly, yeast GPATs are dual specificity acyltransferases, meaning that they can catalyze the addition of a FA to the sn-1 position of both G-3-P and dihydroxyacetone phosphate (DHAP), although Sct1p prefers G-3-P over DHAP (Zheng et al., 2001). In mammals, the addition of a FA to DHAP is done by a unique peroxisomal enzyme, dihydroxyacetone phosphate acyltransferase (DHAPAT)

(Ofman et al., 1994).

1.6.1 Gpt2p and Sct1p Topology and Phosphorylation

Sequencing analysis has revealed that Gpt2p and Sct1p have 31% amino acid sequence identity, and while single deletion of either GPT2 or SCT1 is viable, a double deletion is lethal (Zheng et al., 2001; Zaremberg et al., 2002). Both Gpt2p and Sct1p are integral membrane proteins and have six predicted transmembrane segments, though their exact topology has yet to be elucidated. Of its six hydrophobic stretches, three are uniformly predicted to be transmembrane helices (TMs) (Pagac et al., 2012). The proposed topology for Gpt2p predicts a cytosolic orientation for both N- and C-termini of protein

(Figure 1.8).

Figure 1.8- Predicted topology of Gpt2p. Gpt2p has six predicted transmembrane segments, with the current predicted topology positioning the N- and C- termini facing the cytosol. The serine-rich C-terminus is represented by the orange diamonds. Three of the four catalytic motifs (green boxes) are predicted to face inwards towards the ER lumen, while motif IV is predicted to be on the cytosolic side. Based on data from (Pagac et al., 2012)

Highly phosphorylated serine-rich C-terminal of Gpt2p corresponds with its cytosolic orientation which allows this region being accessible to protein kinases and phosphatases. The proposed topology model supports an ER lumenal location of motifs I,

II, III and cytosolic location for motif IV. This is controversial as it is believed the substrates and activity of Gpt2p are present in the cytosolic side. Currently, there is no protein topology proposed for Sct1p, though it also has several predicted transmembrane segments and multiple phosphorylation sites at the C-terminus, suggesting a cytosolic localization.

Both Gpt2p and Sct1p have been identified as phospho-protein in many global phosphorylation studies (Beh et al. 2001; Helbig et al. 2010; Urban et al. 2007;

Albuquerque et al. 2008; Swaney et al. 2013). Figure 1.9 shows putative phosphosites in these GPATs. This is discussed in depth in Chapter 4.

Gpt2p

Sct1p

Figure 1.9- Putative phosphosites identified in global proteome studies. Amino acid sequences of Gpt2p and Sct1p. Red highlights phosphorylation sites found in global phosphoproteomic studies and blue highlights the four catalytic motifs.

Several kinases for the phosphorylation sites of Gpt2p has been predicted by kinase motif predictions, global phosphorylation analyses or high-throughput kinase deletion studies. A serine/threonine protein kinase Ypk1p, has been identified as a physical interactor of Gpt2p in vitro. Ypk1p is an important kinase for cell survival in response to membrane stress and is a downstream effector of the target of rapamycin complex 2

(TORC2) in yeast. Ypk1p has been linked to lipid homeostasis by inactivating inhibitors of sphingolipid biosynthesis, Orm1p and Orm2p, when sphingolipid levels are low

(Roelants et al. 2011; Fröhlich et al. 2016). Gpt2p was identified to be an in vitro substrate of Ypk1p (Muir et al., 2014), though this Ypk1p-dependent phosphorylation could not be confirmed in vivo (Muir et al. 2015).

It has been shown that phosphorylation level of Gpt2p undergoes changes responding to the growth phase, different carbon sources, and cellular GPAT activity, indicating that it may be regulated through (Bratschi et al. 2009c; Marr et al. 2012). Mass spectrometry (MS) analysis reveals that Gpt2p is a phospho-protein with at least two different phosphorylation states, P1 and P2 (Figure 1.10).

Gpt2p appears as a double band in the SDS/PAGE analysis, lower band represents

P1 and upper band represents P2. Over-expressed Gpt2p shows thicker P2 band, while P1 has been detected either in the absence of Sct1p or in presence of oleic acid (C18:1 Δ9) as the only carbon source (Bratschi et al., 2009; Marr et al., 2012; Pagac et al., 2012).

Treated protein with phosphatase results in a single band, P0, runs slightly lower than P1 (Figure 1.10).

Figure 1.10- Characteristic band pattern distinguishing phosphorylation species of Sct1p and Gpt2p. Red and blue asterisks identify P2 and P1 phosphorylated states, respectively. Green asterisk identifies the P0 state, the resulting band after phosphatase treatment. Sct1p-V5 and Gpt2p-V5 were visualized by western blot using anti-V5 antibody. Modified from (Bratschi et al. 2009).

1.6.2 Gpt2p and Sct1p Substrate Specificity and Localization

It is well known that growth in (C18:1) oleic acid-containing medium induces high degree of TAG accumulation which results in formation of super-sized LDs. Cells regulate the lipid homeostasis by incorporating FAs into TAG and channeling TAG in LDs. Cells lacking Gpt2p, but not Sct1p, are sensitive to oleic acid indicating that they fail to accumulate neutral lipids (Marr et al. 2012). This suggests that Gpt2p, but not Sct1p, is responsible for utilizing C18:1 and channeling it into LDs in the form of TAG. This is consistent with earlier results on the substrate specificity of the yeast GPATs, which showed that Sct1p prefers palmitic acid (C16:0), while Gpt2p had a broader range of substrates (Zheng et al., 2001). Other study shows that Sct1p is a novel regulator of fatty

25 acid desaturation by competing with the desaturase Ole1p for C16:0-CoA and sequestering

C16:0 in lipids (De Smet et al., 2012). Lack of Sct1p induces number of unsaturated FAs, while the overexpression of SCT1 decreases number of unsaturated FAs, by incorporating

C16:0 into phosphatidylcholine (PC), and therefore making it unavailable to Ole1p (De

Smet et al., 2012; De Smet et al., 2013). These results confirm that C16:0 is the preferred substrate of Sct1p.

Furthermore, contribution of Gpt2p and Sct1p in glycerolipid biosynthetic pathways displays degree of differences (Zaremberg et al., 2002). Surprisingly, cells lacking Gpt2p show 50% increase in TAG synthesis while cells lacking Sct1p show 50% decrease in TAG synthesis during log phase. On the other hand, absence of Gpt2p induces a 5-fold production of glycerophosphocholine (GPC), a product resulting from the deacylation of PC synthesized through the Kennedy pathway, while this metabolite was not even detected in the absence of Sct1p (Zaremberg et al., 2002). Taken together, it seems that Gpt2p is mostly involved in utilization of unsaturated acyl-CoA, specifically oleic acid, while Sct1p is mostly involved in incorporation of saturated FAs into DAG to be used to make PC and TAG (Oelkers et al., 2016). These evidences can explain the role of GPATs in controlling distinct pools of PA, which vary in chain length and degree of saturation. PA pools control the contribution of FAs for storage into TAG or for membrane biogenesis responding to the growth phase and cellular conditions (Zaremberg et al., 2017).

Yeast GPATs overlap mostly in their localization and are in fact microsomal

GPATs, localized to both perinuclear and cortical ER in actively proliferating cells. It seems that their localization is differentially enriched in distinct subdomains of ER

(Bratschi et al., 2009). ER is a continuous membranous network made up from

26 interconnected tubules and flat sheets, peripheral ER and the nuclear envelope. ER has distinct regions with specific morphology which allows them to involve in different cellular processes (Christodoulou et al. 2016). ER sheets covered with ribosomes are responsible for the synthesis and folding of membrane, luminal and secreted proteins. ER tubules contain fewer ribosomes are therefore considered 'smooth' ER. They are branched and spread throughout the cytosol and compose approximately 30% of the ER in S. cerevisiae

(Shibata et al., 2006; West et al. 2011). The tubular ER network forms abundant membrane contact sites (MCSs) with other organelles and with the plasma membrane. MCS are sites where two organelles are in close touch, usually tethered together via proteins, but are not fused. Typically, the ER network form MCS with an individual organelle. ER-organelle

MCSs are responsible for organelles dynamics and trafficking within the cell, and are becoming known as sites of lipid biosynthesis and exchange. Recent studies indicate that inter-organelle MCSs are important centers for spatially compartmentalizing lipid metabolic reactions (Hariri et al. 2017). MCSs formed between the ER and the plasma membrane (PM) coordinate phospholipid trafficking to maintain organelle identity and homeostasis. It has been shown that nuclear ER (nER)–vacuole junction, or the NVJ plays a key role in regulation of LDs biogenesis and lipid homeostasis (Hariri et al. 2017). Gpt2p and Sct1p probably regulate lipid homeostasis by associating with these MCS where the transport of lipids occurs between the ER and other organelles.

Intriguingly, Gpt2p has been identified in LD isolation of cells grown in the presence of glucose as a carbon source but not when cells are grown with oleic acid (Marr et al. 2012), while Sct1p has never been reported to be associated with LDs (Binns et al.

2006; Grillitsch et al. 2011; Currie et al. 2014). It has been shown that growing cells in the

27 presence of oleic acid induces a rise in Gpt2p levels and changes in its phosphorylation status. Most importantly, this study revealed the formation of unique Gpt2p-enriched ER structures associated with LDs (Marr et al. 2012), supporting recent models of LD biogenesis in which the ER bilayer juxtaposes the monolayer of LDs (Mishra et al. 2016a).

In addition, a recent study has shown that in the presence of oleic acid, both Gpt2p and

Sct1p were co-immunoprecipitated with Fld1p (Pagac et al. 2016). The ‘few lipid droplet protein 1’ (Fld1p) is a transmembrane ER protein which is enriched at the junctions between ER and LDs. Fld1p participates in coupling neutral lipid synthesis with LD assembly by forming a collar (Wang et al., 2014). Fld1p is the ortholog of the human protein seipin which, when mutated, causes Berardinelli-Seip congenital lipodystrophy, where patients are almost completely lacking adipose tissue (Magré et al. 2001). It seems that Fld1p modulates the activity of GPATs. Cells lacking Fld1p show elevated activity of

Gpt2p and Sct1p, followed by increased level of PA and super-sized LDs formation (Pagac et al. 2016). Finally, Gpt2p, but not Sct1p, has been found to be localized to the tubular network component of the ER (Wang et al. 2017).

1.7 PA and DAG as Signaling Molecules

In S. cerevisiae, PA is the branching precursor that can be used either for making all membrane glycerophospholipids through both Kennedy and CDP-DAG pathways or for making storage lipid (TAG) through the Kennedy pathway. Understanding the genetic mechanism controlling the enzymes involved in glycerolipids (glycerophospholipids +

TAG) biosynthesis pathways is crucial. Regulation of genes expressing these enzymes can be affected by multiple factors, including carbon source, nutrient availability and growth

28 stage. Genes encoding enzymes in the CDP-DAG (e.g., CDS1, CHO1, PSD1, CHO2, and

OPI3) and Kennedy (e.g., EKI1, EPT1, CKI1, CPT1) pathways, and in the synthesis of PI

(e.g., INO1), as well as the genes encoding the inositol (ITR1) and choline/ethanolamine

(HNM1) permeases are coordinately regulated via a inositol-sensitive upstream activating sequence (UASINO) located in their promoter (Lopes et al. 1991).

UASINO is the inositol-responsive cis-acting regulatory element of the mentioned genes. The trans-acting regulatory elements are Ino2p, Ino4p and Opi1p which Ino2p-

Ino4p could bind to UASINO and activate gene expression, while Opi1p represses gene expression by binding to Ino2p (White et al., 1991; Nikoloff et al., 1992; Ambroziak et al.,

1994; Loewen et al., 2003). INO2 and INO4 regulatory genes are responsible for the transcriptional activation of glycerophospholipid biosynthetic enzymes. The regulated genes are repressed in response to the glycerophospholipid precursors, inositol and choline.

In absence of inositol (derepression condition), opi1 mutants overexpress INO1 gene and excrete inositol. Either ino2Δ or ino4Δ can exhibit inositol auxotrophy (ino- phenotype) which is not recovered by mutations at OPI1 locus. The repressing function of Opi1p depends on its cellular location. It has been observed that PA levels control Opi1p localization. Opi1p has ER localization through interaction with Scs2 and PA. When PA levels is high Opi1p remains in the ER membrane but when PA level is decreased, Opi1p gets released and translocates from the ER into the nucleus and results in repression of glycerophospholipid gene transcription. This fact highlights the role of PA availability in regulation of glycerophospholipids biosynthesis at the gene level (White et al., 1991).

The level of PA can be affected directly by the activities of lyso-PA acyltransferase,

CDP-DAG synthase, PA phosphatase, DAG kinase, and phospholipase D. Among these

29 enzymes, PA phosphatase (Pah1p) and DAG kinase (Dgk1p) have the major regulatory role on PA consumption and production, respectively. DAG level can be changed directly by PA phosphatase, DAG kinase, phospholipase C, synthesis of complex sphingolipids, and TAG lipolysis. In great contrast to DAG, the phosphomonoester of the PA headgroup is a key characteristic of PA that participates in protein-binding specificity (Figure 1.11)

(Kooijman et al., 2009). PA can form a hydrogen bond to an effector protein via its phosphomonoester, giving it a greater negative charge. PA acts as a hydrogen bond acceptor, while the effector proteins containing lysine or arginine residues acts as hydrogen bond donors that explain the theory of “electrostatic/hydrogen bond switch” mechanism for PA-specific effector binding (Kooijman et al. 2007).

Both PA and DAG are a cone shaped lipid that induce negative curvature when present in membranes, however it is not clear how changes in the PA:DAG ratio would affect the intrinsic curvature. Variation in PA:DAG balance might impact the negative surface potential of the membrane and the recruitment of effector proteins (Ganesan et al.,

2015).

Figure 1.11- Hydrogen bonding and hydrophobic interactions of phosphatidic acid (PA) and effector proteins. PA (light orange) is the only cone-shaped, anionic lipid in the bilayer. Diacylglycerol (DAG, dark orange) is also cone-shaped inducing negative curvature, and can be produced by de-phosphorylation of PA, but unlike PA, can flip-flop between the leaflets of the membrane without protein assistance. Inverted cone shaped lipids (blue) such as lysophosphatidylcholine (lyso-PC), cause the membrane to have positive curvature. Cylindrical-shaped lipids such as phosphatidylcholine (PC) and phosphatidylinositol (PI) (light and dark purple) are part of the bulk of the bilayer. The cone-shape of PA can expose the hydrophobic environment of the membrane and may facilitate the insertion of hydrophobic portions of PA binding proteins (green) into the membrane. In addition, hydrogen bonding of primary amines from either lysine or arginine residues increases the negative charge of PA by lowering its pKa, causing the proton to dissociate, as proposed by the electrostatic/hydrogen bond switch hypothesis. (Shabits, 2017).

Different studies have shown that the PA:DAG ratio regulates cellular processes.

Impaired PA:DAG balance leads to broad cellular defects including the enlargement of the endoplasmic reticulum and nuclear membranes, blocked lipid droplet synthesis and inhibited endosomal (Han et al., 2007; Adeyo et al., 2011; Sasser et al., 2012; Lawrence

31 et al., 2014). The ratio of DAG to PA is a critical component of regulating vacuole fusion.

Elevated PA:DAG ratios show increased vacuole fusion (Miner et al. 2017).

It has been shown that defects in vacuolar homeostasis increases the PA:DAG ratio through TORC1 signaling and prevent LD and DAG utilization during growth resumption

(Ouahoud, 2018). DAG levels in the ER bilayer membrane can mediate LDs biogenesis due to its ability to generate high negative curvature of membrane monolayers (Choudhary et al., 2018).

1.8 TAG biogenesis and formation of lipid droplet

The ability of eukaryotic organisms to store excess energy from nutrients in the form of non-polar or “neutral” lipids plays a critical role in their survival. At the cellular level, this storage takes place in specialized organelles called lipid droplets (LDs). Other terms for these structures include lipid bodies and adiposomes in animals and oil bodies in plants. LDs are dynamic organelles that are actively engaged in multiple functions. They serve as a storage of neutral lipids, TAG and steryl esters (SE), in all eukaryotic organisms

(Guo et al. 2009; Beller et al. 2010). Lipid metabolism play a critical role for the survival of the organism in conditions in which nutrient supplies are limited. LDs are not only important for lipid storage, but also for maintaining lipid homeostasis (Goodman, 2008;

Fahy et al., 2009; Sheng et al., 2015). In addition, LDs serve a protective role in lipotoxicity. Free fatty acids inside the cells are involved in micelle formation that disrupt cellular membranes and create pores. LDs by storing the excess free fatty acids such as

TAG eliminate the risk of micelle formation (Beller et al. 2010). Noteworthy, the initial pathway of TAG biosynthesis up to PA is shared with the de novo branch of

32 glycerophospholipid biosynthesis, which leads to all major glycerophospholipid classes under normal growth conditions (Figure 1.5) (Henry et al., 2012). In addition, DAG, which is the major precursor of TAG, also serves as a precursor of other glycerophospholipids.

Therefore, it is not surprising that processes controlling TAG homeostasis also affect glycerophospholipid metabolism. Impaired glycerophospholipid synthesis and turnover lead to TAG accumulation indicating the close relationship between glycerophospholipids and TAG formation (Radulovic et al. 2013). Toward the end of the logarithmic phase of growth when the neutral lipid content is particularly high, neutral lipids accumulate between ER bilayer membrane and form lipid droplets.

LDs biogenesis happens in the smooth endoplasmic reticulum. Despite lack of adequate knowledge and direct visualization for the initial steps of the formation process, there is much evidence supporting a model whereby lipid droplets are derived from the endoplasmic reticulum (ER) (Jacquier et al. 2011). For instance, most of the enzymes involved in TAG or SE synthesis are localized to the ER (in the absence of LDs) (Thiam et al., 2013). Electron microscopy analysis also shows association of LDs with the ER

(Arrese et al., 2014). There are certain proteins with dual localization between LDs and ER like Dga1p which is one of the enzymes responsible for TAG synthesis (Thiam et al., 2013;

Jacquier et al., 2011).

The initial steps of lipid droplet formation are not fully understood. There are three proposed models for the LDs biogenesis: (1) lensing model, (2) bicelle formation and (3) vesicle model (Figure 1.12) (Kohlwein et al., 2013). The lensing model is the most accepted one compared to the other possible models (Jacquier et al., 2011). Most models posit an initial lens-shaped accumulation of TAG (or SE) within the secreted regions of ER

33 bilayer. At some point, the growing structure (aggregated neutral lipids) is predicted to bud off the ER, forming a lipid droplet (Jacquier et al., 2011; Thiam et al., 2013) (Figure 1.12).

Figure 1.12- Models of lipid droplet biogenesis. (A) Lensing model: neutral lipids are accumulated between the ER leaflet membrane. LD is formed from emergence of neutral lipid core reached to a critical size; the monolayer of LD is derived from the cytosolic leaflet of the ER membrane. LD can separate from the ER membrane, or remain connected, with the surface layer forming a continuum with the ER. (B) Bicelle formation: similar to the lensing model but the LD is cut off from the ER membrane, and both ER membrane leaflets form the LD monolayer. (C) Vesicle formation: inner leaflet of the bilayer is involved for engulfing of neutral lipid core in the membrane vesicle. Adapted from (Radulovic et al. 2013).

In the bicelle model, LDs are first formed in the ER and then bud into the ER lumen rather than into the cytosol. Therefore, the lumenally localized LD would be enclosed by the ER bilayer membrane (Figure 1.12). The bicelle formation will cause a hatch in the ER structure which seems a problem for this model (Olzmann et al., 2011).

Once LDs are formed, the newly synthesized membrane proteins are moved from the ER to pre-existing LDs and label the entire LD population uniformly. Therefore, the integral membrane proteins, which initially are homogenously distributed throughout the

ER membrane, associate with the nascent LDs. The integral membrane proteins do not require the vesicular transport machinery or vesicle fusion for their transportation from the

ER to LDs. The process is also independent of temperature and energy. The protein migration is reversible and results in ER localization of LD-localized membrane proteins based on regression of LDs under lipolytic conditions (Jacquier et al., 2011).

There are several models explaining the LDs associated with the ER and the proteins transportation between the ER bilayer onto an LD monolayer (Figure 1.13)

(Robenek et al., 2006; Zehmer et al., 2009; Jacquier et al., 2011). The two “Stalk-model” and “ER luminal LDs” models are more possible comparing to other proposed ones

(Nicolas Jacquier et al. 2011). Both models suggest the lateral partitioning of membrane proteins with a hairpin-type topology between the ER bilayer and an LD monolayer

(Jacquier et al. 2011). How proteins that have an ER lumenal domain, could cross the interface between the two compartments is unclear.

Figure 1.13- Possible association between the ER membrane and LDs to facilitate the bidirectional exchange of integral membrane proteins between the two compartments. (A) Four models proposing the transient or permanent association between the ER membrane and LDs. Depicted are cytosolic LDs, which transiently interact with the surface of the ER membrane, egg-cup model: ER membrane holding an LD, an extended membrane stalk enclosing the LD, a modified stalk model to account for the high degree of colocalization of LDs with the ER membrane, and a model in which LDs would be located in the ER lumen and enclosed by the ER bilayer membrane (Robenek et al. 2006; Zehmer et al. 2009). (B) Translocation of integral membrane proteins between the ER and LDs. Peripheral membrane protein with a hairpin-type structure is shown in blue. The integral membrane protein consisting an ER-lumenal domain shown in red, needs to transfer its lumenal domain across a membrane monolayer in the stalk-model (red arrow). Thin lines represent a lipid monolayer, thick lines represent a bilayer membrane (Jacquier et al., 2011).

In the stalk-model, the produced monolayer LD remains between the outer and inner ER leaflet. ER luminal domain of the protein crosses the inner leaflet of the ER membrane to allow the protein transfer from the ER bilayer onto an LD monolayer, a

36 process that is unlikely to occur spontaneously. The ER luminal LDs model can resolve these topological problems for the transportation of these specific proteins. In this model, the LD compartment is first formed in the ER and then bud into the ER lumen rather than into the cytosol. Therefore, the lumenally localized LD would be enclosed by the ER bilayer membrane.

This model suggests the formation of an ER lateral domain within which LD- localized membrane proteins can concentrate and segregate laterally (Jacquier et al. 2011).

This model is not in agreement of many morphological studies by electron microscopy analysis which did not detect any bilayer membrane at the periphery of LDs. Therefore novel complementary and functional approaches are required to reevaluate these investigations (Jacquier et al. 2011).

The LD membrane is decorated by specific proteins varying depending on LDs function and cellular condition (Simons et al. 2011). The structural proteins of LDs might regulate lipolysis of newly synthesized lipids to proceed LD maturation and lipid trafficking (Van Meer, 2001). The proteome analysis of budding yeast LDs suggests that the majority of LD-localized proteins are enzymes participating in lipid metabolism. These proteins include Erg1p, Erg6p and Erg7p for ergosterol (yeast sterol) biosynthesis; Faa1,

Faa4 and Fat1p for fatty acid activation; and Tgl1p, Tgl3p, Tgl4p for neutral lipid degradation (K Athenstaedt et al., 1999). It is not clear whether these enzymes also have structural roles or there are additional LD-associated structural proteins that are yet to be identified.

Yeast cells need to keep their LDs size at a certain level, suggesting that cells control the LD size to maintain the balance between lipid usage and storage processes. It

37 has been shown that deletion of nine specific genes lead to the supersized LDs (SLD) phenotype. The nine genes are CDS1, INO2, INO4, CHO2, and OPI3 which are known to regulate glycerophospholipid metabolism, CKB1 and CKB2 which are responsible for encoding of casein kinase II subunits, and MRPS35 and RTC2 of unknown function (Fei et al. 2011). The majority of these SLD genes seem to contribute directly or indirectly in the metabolism of glycerophospholipids, especially PC, the most common and abundant form of glycerophospholipid in yeast. Decreased PC synthesis, and consequently an increased

PE to the total membrane phosphoglycerolipids ratio (or a decrease in PC/TAG), has been associated with SLD formation. Further investigation has revealed that the synthesis of not only PC, but also other glycerophospholipids could be critical for LDs morphology.

There are other mechanisms that would affect the LDs size. The ‘few lipid droplets protein 1’ (Fld1p) encoded by FLD1 is the functional orthologue of the human BSCL2 gene encoding seipin protein. These two evolutionary conserved genes are responsible for LDs size maintenance (Fei et al., 2008). Fld1p is a transmembrane ER protein which is enriched at the junctions between ER and LDs. Thus, this protein might participate in coupling neutral lipid synthesis with LDs assembly by forming a collar. The strains missing FLD1 gene (fld1Δ) with two independent genetic screens show aberrant LDs morphology (Wang et al., 2014; Fei et al., 2011). The other gene involved in LDs morphology is LDB16 coding low dye-binding protein 16 (Ldb16p) which has a potential role in protein glycosylation.

Ldb1 is a Fld1-interacting protein and Fld1p is required for Ldb1p stability (Wang et al.,

2014). The Ldb16p and Fld1p are mutually dependent for the maintenance of LDs.

UBX-domain-containing proteins belong to a class of evolutionarily conserved adaptor proteins for the ubiquitin-selective chaperone Cdc48 (Römisch, 2006). Ubx2p

(encoded by UBX2) is a transmembrane protein that localizes to the ER (Schuberth et al.,

2005). Aberrant LDs morphology is observed in cells either lacking or overproducing

Ubx2p (Wang et al., 2012).

Lro1p is one of the major enzymes that have a redundant role in LDs formation in yeast. Lro1 participates in LD assembly by localizing to the dynamic foci and remodeling glycerophospholipids. It seems that ubx2Δ results in decreased TAG levels, which might be linked to the mislocalized Lro1p enzyme. Thus, the access of mislocalized Lro1p to

DAG might be decreased (Wang et al., 2012).

Lipid droplet hydrolase 1 (Ldh1p) is encoded by LDH1 gene which is associated with LDs. Ldh1p is a membrane-active serine hydrolase with a classical catalytic triad containing a serine. Ldh1p exhibits active esterase plus weak triacylglycerol lipase activities (Thoms et al., 2011). It seems that Ldh1p mediates the mobilization of LD-stored lipids. Cells deficient in Ldh1p (ldh1Δ) are characterized by accumulation of nonpolar lipids leading to the formation of giant LDs.

When budding yeast reaches the late exponential and stationary phase of its growth, all TAG synthesis occurs by Dga1p on LDs (Markgraf et al., 2014). During growth resumption, the process of the net TAG consumption is mediated by lipases, such as Tgl3p, that always associate with LDs. However, since during the growth resumption Dga1p is also initially localized to LDs, a mechanism that suppresses a dispensable cycle between

DAG and TAG is required to allow degradation of net TAG. Inheritance of cortical ER

(Ice2p) protein encode by the ICE2 gene can mediate the suppression of the unnecessary cycling (Markgraf et al., 2014). Ice2p is a transmembrane protein localized in the ER. Ice2p regulates the channeling of DAG from LDs to the ER.

The Ice2p-mediated regulation of a dispensable cycle on LDs thus has an important role in regulatory mechanism of lipid homeostasis during growth resumption. The absence of Ice2p leads to three-fold higher TAG levels in exponentially growing cells resulting in two-fold increase in the number of LDs per cell. Furthermore, deletion of ICE2 decreases the glycerophospholipids production specially PE and PS (Markgraf et al., 2014).

1.9 Glycerolipid homeostasis during growth stages

In eukaryotic cells, hundreds of molecular lipid species produced by a metabolic network make full lipid complement (lipidome). This network connects the metabolism of fatty acids, glycerophospholipids, glycerolipids, sphingolipids, and sterol lipids (Ejsing et al., 2009; Gaspar et al., 2011; Rajakumari et al., 2010). Understanding how cells modulate the lipidome through this network to mediate remodeling of cellular processes during physiological adaptations is critical. Combination of comprehensive quantitative proteomics and lipidomics (proteolipidomics) enables us to analyze dynamic co-regulation of lipid metabolism and cellular processes during physiological adaptations and to compare fermentative and respiratory metabolism during proliferation and quiescence stages.

Analysis of yeast cells shifting from fermentation to respiration reveals an extensive remodeling in their proteome with 3,105 proteins showing significant differences in abundance (Casanovas et al., 2015). Sorted proteins based on gene ontology (GO) categories for cellular components and biochemical pathway annotation were used to determine the mean fold change within each structural or functional category. Resulted data showed i) elevated levels of mitochondrial proteins which is consistent with the increased number of mitochondria during a shift from a fermentable to a non-fermentable

40 carbon source (Egner et al., 2002); ii) increased level of peroxisomal proteins resulted from induced peroxisome proliferation in the post-diauxic shift (Lefevre et al., 2013); and iii) decreased nucleolar and ribosomal proteins showing a down-regulation of protein synthesis.

This analysis showed increased abundance of Gpt2p by 2.5 fold and 4.15 fold in diauxic shift and post-diauxic phase, respectively (Casanovas et al., 2015), highlighting the role of Gpt2p as part of protein machinery programming lipid metabolism as cells undergo different growth stages.

Levels of Sct1p showed same pattern of increase in both diauxic shift and post- diauxic phase by 0.8-fold, while no change was detected in its level from diauxic shift to the post-diauxic phase (Casanovas et al., 2015).

Altogether, it seems that post-diauxic shift can induce glyoxylate cycle, TCA cycle, fatty acid oxidation, glycerol degradation and lipid precursor inositol synthesis. On the contrary, the protein machinery regulating sphingolipid metabolism, sterol metabolism, and the oleic acid synthesis was suppressed. These findings indicate that cellular metabolism including lipid metabolic processes and organelle biogenesis are adapted to diauxic shift and post-diauxic phase.

Further analysis on lipid metabolism at the cellular level by mapping of lipid metabolic network of 138 proteins involved in fatty acid, glycerophospholipid, glycerolipid, sphingolipid, and sterol lipid metabolism pathways showed changes in abundance level of 85% of the proteins (Casanovas et al., 2015). Analyzed data showed (i) a decrease in fatty acid desaturase Ole1p and fatty acid elongase Sur4p (ii) varying expression level of the fatty acyl-CoA synthetases Faa1, Faa2, Faa3, and Faa4 (iii)

41 fluctuations in the levels of glycerophospholipid enzymes (e.g., increased levels of Opi3p and Cki1p) (iv) suppression of several ergosterol biosynthetic enzymes (v) reduced levels of the sphingolipid enzymes Lcb1p and Tsc10p, and increased level of ceramidase Ydc1p

(vi) varying expression level of enzymes involved in the synthesis and degradation of TAG and SE. These findings explain a broad reprogramming of the lipid metabolic network during switch from fermentative to respiratory metabolism.

Analyzing the changes in cellular lipid composition affected by proteome adjustment show that cell lipidomics correlate with proteomics adapting with cellular processes (Casanovas et al., 2015). Quantitative lipidomic analysis of 223 lipid species across different time point of growth stages showed that relative abundance of 221 lipids changed along the growth profile indicating extensive modulation of the lipidome broader than analyzing only the protein complement of the cell. Remarkably, the membrane and storage lipid ratio fluctuates during all growth stages, meaning a dynamic transition between lipids required for membrane biogenesis and storage lipids. When cells enter quiescent phases, membrane lipids become recruited for secretory traffic and plasma membrane expansion and mobilized during proliferation. During a diauxic shift, excessive lipids become stored in form of neutral lipids, TAG and SE. Surprisingly, the total levels of sphingolipids and sterols despite the down-regulation of their de novo biosynthetic machinery, did not fluctuate during all growth phases suggesting presence of additional regulatory mechanisms.

Storage lipids accumulated in LDs can undergo two different cycles of mobilization and accumulation prompted by active growth and quiescence, respectively. During transition into the diauxic shift, accumulated TAG and SE lipids are initially mobilized.

Both TAG and SE exist in similar amounts (Kohlwein et al., 2013) and display parallel temporal profiles.

On the contrary, during the post-diauxic phase, mobilized TAG and SE show distinct dynamics (Casanovas et al., 2015). High expression of the lipases Tgl3p, Tgl4p, and Tgl1p happens at this phase, indicating role of these enzymes in hydrolysis of TAG and SE. Accordingly, proxisomal enzymes, Pox1p, Fox1p, Pot1p, and acyl-CoA synthase

Faa2p, were up-regulated leading to increased β-oxidation which shows that growth resumption is supported by channeling of fatty acids towards β-oxidation (Casanovas et al., 2015). When cells enter to the stationary phase, TAG is re-synthetized, whereas SE pools remains stable. Elevated levels DAG acyltransferase enzymes, Are2p (Kohlwein et al., 2013) and probably Dga1p, show their role in synthesizing TAG. As cells enter the stationary phase, TAG is re-synthesized, whereas the SE pool is kept relatively unchanged

(Casanovas et al., 2015). According to the protemics and lipidomics analyses together,

Gpt2p expression is induced during both diauxic shift and post-diauxic phase to respond to the fluctuated levels of lipid metabolites that support its need for regulation of cellular lipid homeostasis.

1.10 Signalling pathways that regulate lipid metabolism

Glycerophospholipids by serving roles in secretory trafficking, organelle identity, and anchoring of membrane proteins (Van Meer et al., 2008) are major constituents of cellular membranes. Broad-range composition of present glycerophospholipids in cells highlights extensive dynamics across all growth phases. Glycerophospholipids abundance fluctuate responding to the different cell growth stages. It has been shown that their levels

43 increased during proliferation as secretory traffic drives membrane expansion and cell division that are resulted by elevated levels of PI and PE (Casanovas et al. 2015). When cells go through a diauxic shift, a gradual augmentation in PC levels is detected which can be explained by up-regulation of enzymes synthesizing PC through the CDP choline pathway (Cki1p) and the PE methylation pathway (Opi3p). Besides, up-regulation of the

PS synthase enzyme Cho1p and reduced levels of PS and PE, demonstrate that increased level of PC is resulted by sequential conversion of PS. It has been seen that during the diauxic shift and post-diauxic shift, Plb1p phospholipase level which participates in remodeling of PC species, increases. Moreover, systematic changes in the composition of

PC and lyso-PC indicate activation of PC remodeling during the diauxic shift.

Notably, genes encoding subset of enzymes involved in glycerophospholipid synthesis show increased expression level during the post-diauxic shift. The regulated genes include OPI3, CKII and CHO1 and INO1 which are controlled by the transcriptional regulators Ino2p, Ino4p, and Opi1p (Henry et al., 2012). Opi1p can be sequestered and attached to the high PA level in the ER membrane allowing the Ino2-Ino4 transcription factor to induce gene expression (Henry et al., 2012). On the contrary, when PA levels are reduced, Opi1p is released form the ER membrane and bonded to Ino2p in the nucleus which lead to inhibition of gene expression.

Apparently, PA level goes up during the exponential phase when PI synthesis is high (Casanovas et al. 2015). This result seems to be in contrast with the model of transcriptional regulation proposing elevated PC synthesis due to high PA levels and induced biosynthesis of PC.

It has been shown that during post-diauxic phase, most enzymes involved in glycerophospholipids biosynthesis are up-regulated. Although the PA levels does not show an increase, a pronounced alteration of PA species profile takes place indicating that varying PA species play distinct roles in the transcriptional program of glycerophospholipid metabolism (Casanovas et al. 2015).

1.11 Hypothesis and Goals

The main goal of this research was to study the regulation of Gpt2p through phosphorylation. Our findings show that lack of phosphorylation on key phosphorylation sites of Gpt2p induces conformational changes that might affect protein localization, stability, activity, and protein-protein interactions. The conformational change due to lack of phosphorylation probably limits access of regulatory kinase and phosphatase enzymes of Gpt2p involved in glucose depletion signaling and cell-cycle progression. Impaired regulation on the catalyzer of the rate-limiting step of glycerolipids biosynthesis pathway,

Gpt2p, can halt its contribution with downstream effector proteins leading to certain metabolic phenotypes.

By performing well-established protocols for the identification of Gpt2p phosphorylation sites and characterization of different phospho-mutants, we hypothesized that these phosphorylation sites by themselves or in combination with other residues play a critical role in regulation of Gpt2p through phosphorylation. Identifying the function of serine-rich tail of Gpt2p enables us to understand the mechanism regulating Gpt2p through phosphorylation.

The objectives of this thesis were as follow:

1. To identify phosphorylation sites in Gpt2p

2. To investigate the role of key phosphosites in Gpt2p regulation through phosphorylation

3. To study metabolic impact of unregulated Gpt2p variants

4. To investigate the impact of unregulated Gpt2p variants in the proteome of lipid droplets (protein-protein interactions)

Chapter 2 – Materials and Methods

2.1 Yeast Strains, Plasmids, Primers, Culture Conditions, and Transformations

All yeast strains and primers used in this study are listed in Table 2.1 and Table 2.2.

Yeast transformations were performed using the lithium acetate method (Gietz et al. 1992).

Table 2.1 Yeast strains used in this study.

Application Name Genotype Source BY4741wt BY4741; MAT a; his3Δ1; leu2Δ0; Euroscarf

met15Δ0; ura3Δ0

gpt2Δ BY4741; MAT a; his3Δ1; leu2Δ0; Euroscarf met15Δ0; ura3Δ0; gpt2Δ::kanMX4 sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; Euroscarf met15Δ0; ura3Δ0; sct1Δ::kanMX4 Characterization of GPT2wt-V5 BY4741; MAT a; his3Δ1; leu2Δ0; This study GPT2 phospho-mutants met15Δ0; ura3Δ0; GPT2wt-V5- 6xHis::HIS3MX6 GPT2S664A-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23xA-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT2S668A-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23xA-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT2S671A-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23xA-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT23xA-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23xA-V5- collaboration 6xHis::HIS3MX6 with K.

Athenstaedt GPT23xD-V5 BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT23xD-V5- 6xHis::HIS3MX6 GPT2Trunc-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2Trunc-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT2wt-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2wt- collaboration GFP::KanMX6 with K. Athenstaedt

GPT23xA-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23xA- collaboration GFP::KanMX6 with K. Athenstaedt GPT23xD-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23xD- collaboration GFP::KanMX6 with K. Athenstaedt GPT2Trunc-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2Trunc- collaboration GFP::KanMX6 with K. Athenstaedt GPT2S651A-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2S651A-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT2S693A-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2S693A-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT23xA+S693A-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23XA+S693A-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT29xD-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT29D-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT2S651A-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2S651A- collaboration GFP::KanMX6 with K. Athenstaedt GPT2S693A-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2S693A- collaboration GFP::KanMX6 with K. Athenstaedt GPT23xA+S693A-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23XA+S693A- collaboration GFP::KanMX6 with K. Athenstaedt GPT29xD-GFP BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT29D- collaboration GFP::KanMX6 with K. Athenstaedt GPT29xA-GFP BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT29A- GFP::KanMX6 Characterization of GPT2 GPT2wt-V5/sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study phospho-mutants in the met15Δ0; ura3Δ0; GPT2wt-V5- absence of Sct1p 6xHis::HIS3MX6; sct1::URA3-KL GPT23xA-V5/sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT23XA-V5- 6xHis::HIS3MX6; sct1::URA3-KL

GPT2wt-GFP/sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2wt- collaboration GFP::kanMX6; sct1::URA3-KL with K. Athenstaedt GPT23xA-GFP/sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23XA- collaboration GFP::kanMX6; sct1::URA3-KL with K. Athenstaedt GPT23xD-GFP/sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23XD- collaboration GFP::kanMX6; sct1::URA3-KL with K. Athenstaedt GPT2Trunc-GFP/sct1Δ BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT2Trunc- collaboration GFP::kanMX6; sct1::URA3-KL with K. Athenstaedt Lipid droplet GPT2wt-GFP/Erg6p- BY4741; MAT a; his3Δ1; leu2Δ0; This study morphology analysis RFP met15Δ0; ura3Δ0; GPT2wt- using ERG6-RFP LDs GFP::kanMX6; ERG6-RFP::kanMX6 marker GPT23xA- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/ERG6-RFP met15Δ0; ura3Δ0; GPT23XA- GFP::kanMX6; ERG6-RFP::kanMX6 GPT23xD- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/ERG6-RFP met15Δ0; ura3Δ0; GPT23XD- GFP::kanMX6; ERG6-RFP::kanMX6 GPT2Trunc- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/ERG6-RFP met15Δ0; ura3Δ0; GPT2Trunc- GFP::kanMX6; ERG6-RFP::kanMX6 GPT2wt- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/sct1pΔ/ERG6- met15Δ0; ura3Δ0; GPT2wt- RFP GFP::kanMX6; ERG6-RFP::kanMX6; sct1::HIS3-KL GPT23xA- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/sct1pΔ/ERG6- met15Δ0; ura3Δ0; GPT23XA- RFP GFP::kanMX6; ERG6-RFP::kanMX6; sct1::URA3-KL GPT23xD- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/sct1pΔ/ERG6- met15Δ0; ura3Δ0; GPT23XD- RFP GFP::kanMX6; ERG6-RFP::kanMX6; sct1::HIS3-KL GPT2Trunc- BY4741; MAT a; his3Δ1; leu2Δ0; This study GFP/sct1pΔ/ERG6- met15Δ0; ura3Δ0; GPT2Trunc- RFP GFP::kanMX6; ERG6-RFP::kanMX6; sct1::URA3-KL Characterization GPT2G261L-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In of inactive GPT2 mutant met15Δ0; ura3Δ0; GPT2G261L-V5- collaboration 6xHis::HIS3MX6 with K. Athenstaedt GPT23xA+G261L-V5 BY4741; MAT a; his3Δ1; leu2Δ0; In met15Δ0; ura3Δ0; GPT23XA+G261L- collaboration V5-6xHis::HIS3MX6 with K. Athenstaedt SDS-PAGE analysis of GPT2wt-V5/nem1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study GPT2 phospho-mutants met15Δ0; ura3Δ0; GPT2wt-V5- in the absence of 6xHis::HIS3MX6; nem1::kanMX6

49 possible phosphatase or GPT23xA-V5/nem1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study kinase enzymes met15Δ0; ura3Δ0; GPT23XA-V5- 6xHis::HIS3MX6; nem1::kanMX6 GPT2wt-V5/psk1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT2wt-V5- 6xHis::HIS3MX6; psk1::kanMX6 GPT23xA-V5/psk1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT23XA-V5- 6xHis::HIS3MX6; psk1::kanMX6 GPT2wt-V5/ypk1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT2wt-V5- 6xHis::HIS3MX6; ypk1::kanMX6 GPT23xA-V5/ypk1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT23XA-V5- 6xHis::HIS3MX6;ypk1::kanMX6 GPT2wt-V5/snf1Δ BY4741; MAT a; his3Δ1; leu2Δ0; This study met15Δ0; ura3Δ0; GPT2wt-V5- 6xHis::HIS3MX6; snf1::kanMX6

Table 2.2 Primers used in this study.

Application Name of Tm Sequence Primer (˚C) Amplifying GPT2 G1-F 68 5’ATGTCTGCTCCCGCTGCCGAT3’ gene G1-(no- 54 5’TTCTTTCTTTTCGTGTTCTC3’ STOP)-R Site directed G1- 108 5’CGCCCTAGCAAGAGTGAATGATAGAGGCGCGTTG mutagenesis for A668D-F A3’ converting S668A to 668D from a G1- 108 5’TCAACGCGCCTCTATCATTCACTCTTGCTAGGGCG3 S664,668,671A A668D-R ’ template Site directed G1- 118 5’CTAGCTTCTAACGCCCTAGATAGAGTGAATGATAG mutagenesis for A664D66 AGGCG3’ converting S664A 8D-F to 664D from a G1- 118 5’CGCCTCTATCATTCACTCTATCTAGGGCGTTAGAA S664A,668D,671A A664D66 GCTAG3’ template 8D-R Site directed G1-3XD- 142 5’CTAGATAGAGTGAATGATAGAGGCGATTTGACCGA mutagenesis for F TATTCCAATTTTTTCTG3’ converting S671A G1-3XD- 142 5’CAGAAAAAATTGGAATATCGGTCAAATCGCCTCTA to 671D from a R TCATTCACTCTATCTAG3’ S664D,668D,671A template Site directed G1- 108 5’GACGGTTATGATGTCTCTGCTGATGCAGAGTCTTCT mutagenesis for S632A-F A3’ making S632A G1- 108 5’TAGAAGACTCTGCATCAGCAGAGACATCATAACCG S632A-R TC3’ Site directed G1- 132 5’TCTCTAAGAGAGACGGTTATGATGTCTCTGATGAT mutagenesis for S632D-F GCAGAGTCTTC3’ converting S632A G1- 132 5’GAAGACTCTGCATCATCAGAGACATCATAACCGTC to 632D S632D-R TCTCTTAGAGA3’

Site directed G1- 112 5’ATCTAGAAGCCGCTCTTCTGCTATACATTCTATTGG mutagenesis for S651A-F CTC3’ making S651A G1- 112 5’GAGCCAATAGAATGTATAGCAGAAGAGCGGCTTCT S651A-R AGAT3’ Site directed G1- 126 5’CAATCTAGAAGCCGCTCTTCTGATATACATTCTATT mutagenesis for S651D-F GGCTCGCT3’ converting S651A G1- 126 5’AGCGAGCCAATAGAATGTATATCAGAAGAGCGGC to 651D S651D-R TTCTAGATTG3’ Site directed G1- 126 5’GGTCAATGGAAAAGTGAAGGTGAAACTGCTGAGG mutagenesis for S693A-F ATGAGGATG3’ making S693A G1- 126 5’CATCCTCATCCTCAGCAGTTTCACCTTCACTTTTCC S693A-R ATTGACC3’ Site directed G1- 126 5’GGTCAATGGAAAAGTGAAGGTGAAACTGATGAGG mutagenesis for S693D-F ATGAGGATG3’ converting S693A G1- 126 5’CATCCTCATCCTCATCAGTTTCACCTTCACTTTTCC to 693D S693D-R ATTGACC3’ qPCR experiment qPCR- 64 5’CGTGGTGCATATAACGTTCCC3’ GPT2 gene GPT2-F qPCR- 64 5’GCATTCTGGATCGGGACTTTC3’ GPT2-R qPCR experiment qPCR- 54.5 5’AATTCCAGATTCTAGCGCGG3’ TFC1- house- ROO- keeping gene 193-F qPCR- 52 5’AATTCTGGGCAGGTCTAACG3’ ROO- 194-R qPCR experiment qPCR- 54.5 5’GAGGAAGATTACGATGCGGAG3’ TAF1- house- ROO- keeping gene 195-F qPCR- 54.5 5’CTTCATGTTTTCTGCGACAGAG3’ ROO- 196-R qPCR experiment GK073-F 53 5’CGAAAGAGTACAAGTTGATGGTG3’ UBC6 – house- GK074-R 52.5 5’AAGTACCGTGATATTGACCGC3’ keeping gene

Briefly, cells were grown to mid-exponential phase (OD600 ~0.5) and were harvested by centrifugation (Sorvall Legend RT Centrifuge). Pellets were washed with water and 1xTE (50 mM Tris-HCl (pH 7.4) and 2 mM EDTA)/100 mM LiAc and then resuspended in 1xTE/LiAc. The washed cells (50 μL) were added to tubes containing the desired plasmid and salmon sperm DNA (boiled for 15 minutes and then placed on ice). A mixture of 40% polyethylene glycol/TE/100 mM LiAc was added and samples were incubated at room temperature for one hour with shaking, followed by a heat-shock at 42°C for 15 minutes. Cells were collected resuspended in 1xTE and plated onto plates lacking

51 uracil to select for the pYES plasmid. Strains lacking endogenous GPT2 and SCT1 (gpt2Δ and sct1Δ) were used for the yeast transformations with the pYES expression vector

(Invitrogen) (Figure 2.1) containing phospho-mutant strains. Genes cloned in the pYES vector are under the galactose-inducible GAL1 promoter. The vector also possesses a 2µ origin of replication resulting in high copy number (~50) and URA3 gene for selection.

GPATs over-produced using this expression system were fused to a V5 and 6xHis tag in the C-terminus. Plasmids were isolated from DH5α Escherichia coli stocks using the

GeneJET Plasmid mini-prep Kit (Fermentas). DNA concentration was determined spectrophotometrically using an ND-1000 Nanodrop spectrophotometer 30 (Thermo

Scientific™ NanoDrop™).

Figure 2.1 pYES2.1/V5 His-TOPO cloning vector. PGAL1: galactose promoter.

Strains containing the pYES vector were grown in synthetic defined medium lacking uracil (SD-Ura). Once in mid-exponential phase (OD600 ~0.5), cells were

52 collected, washed, and resuspended in SGal-Ura media (same composition as SD-Ura, but with 2% galactose instead of glucose) to induce gene expression from the pYES vector.

When necessary, strains were also grown in rich Yeast Peptone Dextrose (YPD) (1% (w/v) yeast extract, 2% (w/v) bacto-peptone (MP Biomedicals), 2% glucose (Fisher)).

For making C1δ-GFP transformants, cells were transformed with C1δ-GFP plasmid which is the cloned C1 domain of PKCδ (C1δ) into a centromeric yeast expression vector under a constitutive promoter for in vivo visualization of DAG pools. Strains containing the C1δ-GFP vector were grown in defined synthetic minimal medium (SD-Ura) to maintain and express the plasmid.

2.1.2 Gene cloning

The GPT2 gene was cloned into a pYES vector using TOPO TA cloning kit

(Invitrogen). First, GPT2 gene was amplified by PCR using forward primer and reverse primer missing the stop codon followed by PCR clean up step. Cloning reaction with a 6

µL total volume was prepared by addition of 4 µL PCR reaction, 1 µL salt solution, and 1

µL TOPO vector. The mixture was incubated at room temperature for 15 minutes, then was placed on ice for 2-3 minutes. All TOPO reaction was transferred into a vial of One Shot

Chemically Competent E.coli cells and mixed gently avoiding pipetting up and down. The vial was remained on ice for 5 extra minutes followed by heat-shock at 42˚C for 30 seconds without shaking and it was immediately placed on ice. Next, 250 µL of room temperature

S.O.C. medium (2% Tryptone, 0.5% Yeast Extract, 10 mM NaCl, 2.5 mM KCl, 10 mM

MgCl2, 10 mM MgSO4, 20 mM glucose) was added to the vial and was shaken horizontally

(200 rpm) at 37˚C for 1 hour. Volumes of 50 and 250 µL of transformation were spread on

53 pre-warmed LB + Ampicillin plates. Colony PCR and DNA sequencing steps were done to confirm the proper insertion of GPT2 gene into the vector. pYES vector provides a PGAL1 promoter at the upstream and a V5 epitope + 6XHis at the downstream of the inserted gene.

2.1.3 Site-directed mutagenesis

To replace a single amino acid in the GPT2 sequence, QuickChange II XL Site-

Directed Mutagenesis kit (Stratagene, Germany) was used according to the manufacturer’s instructions. Primer sets containing the desired mutation in the middle and flanking by sequences complementary to the template sequence were designed using QuickChange

Primer Design program. The site-directed mutagenesis reaction was performed using a

PfuUltra high-fidelity DNA polymerase for mutagenic primer-directed replication of both plasmid strands. The basic procedure utilizes a supercoiled double-stranded DNA vector with an insert of interest and two synthetic oligonucleotide primers, both containing the desired mutation. The primers used were between 25 and 45 bases in length and were designed so that they annealed to the same sequence on opposite strands of the plasmid.

The primers are extended during temperature cycling by the DNA polymerase, without primer displacement. Extension of the primers generates a mutated plasmid containing staggered nicks. Following temperature cycling, the product is treated with DpnI endonuclease, which is specific for methylated and hemimethylated DNA and is used to digest the parental DNA template and to select for mutation-containing synthesized DNA.

1 µL of the nicked vector DNA incorporating the desired mutation is then transformed into

25 µL of ice pre-cooled DH10B Electrocompetent E. coli cells (Invitrogen) in a 0.1 cm gap electroporation cuvette on ice. Electroporation was done by placing the cuvette in an

54 electroporation apparatus set at 25 µF, 200 Ω, 2.5 kV and holding the keys for 4-5 seconds.

1 mL S.O.C broth was added immediately to the cuvette. The reaction was transferred into a microfuge tube and incubated at 37 ˚C for 1 hour. Cells were spun down and plated onto

LB + Ampicillin plates and grown overnight at 37 ˚C. Multiple mutant strains were prepared by designing new primer sets introducing new mutations in already made mutant strains as template.

2.1.4 Yeast crossing and tetrad dissection

Haploid yeast strains of interest with opposite mating type were mated on a YPD plate. Strains had complimentary auxotrophic genotype for specific amino acids. Cells were incubated at 30 °C overnight to form diploid cells. Diploid cells containing wild type copy of gene for amino acids were selected by transferring cells on a selective plate lacking auxotrophic amino acids using replica plating method. Selected cells were transferred on minimal medium plate for sporulation. Cells were incubated at 25 °C in humid condition for about 4 days and were checked by light microscopy to examine formation of sufficient

4-spore asci (tetrad). Head full of cells were collected with a toothpick and were resuspended in a solution of 0.5 mg/mL zymolyase in 1 M sorbitol. Samples were incubated for 10 minutes at 30 oC and slowly and carefully 800 µL of sterile distilled water was added to the tube and place on ice. 10 µL of the cells were transferred onto one end of a YPD plate and smeared across end with a toothpick. Tetrad dissection was done with dissection microscope. Genotype of grown spores were characterized by plating them on different auxotrophic media to find the interested recombinant strain. For making strains

55 carrying two different fluorophore-tagged proteins, fluorescence microscopy analysis was used to detect the desired recombinant strain having both fluorescence signals.

2.2 Cell lysate and microsomal fraction preparation, Protein Determination, SDS-PAGE, and Western Blot

2.2.1. Cell Lysate preparation

Yeast cells were grown to exponential phase in the synthetic defined medium (SD) media containing 0.67% (w/v) Yeast Nitrogen Base without amino acids (YNB with ammonium sulfate, MP Biomedicals), 2% glucose, 0.002% methionine, arginine, histidine, tryptophan, and adenine sulfate, and 0.003% leucine and lysine (all from Sigma). About

150 OD600 units were collected in Sorval using SC6000 ROTOR at 5000 rpm for 15 minutes. Cells were resuspended in ice cold homogenization buffer I (0.25 M sucrose, 10 mM Tes pH 7.5, 1 mM EDTA). After a short centrifuge step of 2500 rpm at 4˚C, the cell pellet was resuspended in homogenization buffer II (0.25 M sucrose, 10 mM Tes pH 7.5,

1 mM EDTA, ROCHE protease inhibitor, 150 µL aprotinin (1 mg/ml), 90 µL pepstatin (1 mg/ml) followed by vortexing with glass beads five times for 30 seconds with 30 seconds intervals on ice in between. Cell lysates were mixed with equal volume of SDS Laemmli and boiled for 5 minutes and were subjected to Western blot analysis.

2.2.2 Microsomal Fractionation

Cells were grown in SD media to mid-exponential phase (OD600 ~0.5) and ~150

OD600 units were collected and resuspended in GTE lysis buffer containing protease and phosphatase inhibitors (20% glycerol, 50 mM Tris-HCl pH 7.4, 1 mM EDTA, 1 mM PMSF

(Sigma), 1x protease inhibitor (Roche), 3 μg/mL Pepstatin (Sigma), 10 μL/mL phosphatase inhibitor cocktail 2 (Sigma)). Briefly, cells were lysed via vortexing with glass beads five times for 30 seconds with 30 second intervals on ice in between. The sample was spun down at 16,000 x g for 15 minutes at 4 °C, and the resulting supernatant (from here on, called the “lysate”) was then centrifuged at 450,000 x g for 15 min at 4 °C a Beckman benchtop ultracentrifuge using Beckman rotor TLA100.2. Pellets were resuspended in

GTE lysis buffer and homogenized to obtain the microsomal samples.

2.2.3 Protein determination, SDS PAGE and Western blot

Before protein determination, samples were mixed with equal volume of 2x SDS

Laemmli buffer (125 mM Tris-HCl pH 6.8, 4% (w/v) SDS, 5% glycerol, 250 mM EDTA)

(Laemmli, 1970) (1:1 v/v). Samples were then boiled for 2 minutes. Protein concentration was determined using the BCA assay (Thermo Scientific) with bovine serum albumin as a standard. Before being loaded onto an SDS-PAGE gel, all samples in 1x gel loading buffer

(GLB, 0.2 M Tris HCl (pH 6.8), 8% SDS, 0.4% bromophenol blue, and 40% glycerol) were boiled for 1 minute. SDS-PAGE was carried out by the method of Laemmli (Laemmli

1970). Proteins were separated by 8% stacking gel containing trichloroethanol (TCE,

Sigma) to visualize proteins without staining (Ladner et al. 2004). Proteins were transferred to a polyvinylidene fluoride (PVDF) membrane (Millipore) using Bio-Rad transfer system at 100 v for 1 hr, and then stained with red Ponceau (Sigma) to confirm transfer. For the western blot analysis, V5-tagged proteins were detected with an anti-V5 primary antibody

(Invitrogen #R96025). GFP-tagged protein was detected by anti-GFP primary antibody

(Roche). RFP-tagged Erg6 protein was detected by polyclonal anti-Erg6 primary antibody

(G. Daum, University of Graz). Vma2 was detected by monoclonal anti-Vma2 primary antibody (Invitrogen). Dpm1 was detected by anti-Dpm1 primary antibody (Life

Technologies). Pma1 was detected by polyclonal anti-Pma1 primary antibody (University of Politecnica de Valencia). Secondary antibodies used in this study were goat anti-mouse

(GAM) IgG and goat anti-rabbit (GAR) IgG conjugated with horseradish peroxidase (Life

Technologies).

These antibodies were used to visualize the presence of the proteins by chemiluminescent signal detection (Amersham enhanced chemiluminescence (ECL), GE

Healthcare) using Amersham Imager 600 system and analyzed by ImageJ software.

2.2.4 Phosphatase treatment experiment

Phosphatase treatment assay was done using Lambda protein phosphatase enzyme

(New England BioLabs P0753S). Microsomal fraction of samples were prepared in the absence of phosphatase inhibitor. 30 µg of total protein was added in a total volume of 50

µL reaction buffer (50 mM Tris-HCl, pH 7.5, 0.1 mM EDTA, 5 mM dithiothreitol, 0.01%

Brij 35, and 2 mM MnCl2). Samples were split in two and incubated for 30 minutes at 30

°C, with or without the addition of 400 units of λ-phosphatase. The reaction was stopped by addition of SDS-PAGE loading buffer.

2.3 GPAT activity assay

Enzyme reaction was performed by incubating of 40 µL GPAT protein sample

(lysate or microsomal fraction) with oleyl-CoA (0.08 mmol/L) and 0.04 mL of the mixed

14C- radiolabeled and non-radioactive G-3-P (0.42 mmol/L final concentration) as enzyme

58 substrates in Tris-HCL buffer (75 mmol/L, pH 7.5), MgCl2 (8.3 mmol/L), NaF (8.3 mmol/L), fatty acid poor bovine serum albumin (1 mg) at 37 ˚C for 15 minutes aiming for formation of PA as the final product. Reaction was stopped by addition of chloroform/methanol (1:2, v/v). After 5 minutes centrifugation at 2000 rpm, the resulting supernatant was decanted and chloroform was added, followed by 2M KCl in 0.2M H3PO4 to develop separated phases. After mixing and centrifugation for 5 minutes at 2000 rpm, the top layer was discarded and the lower chloroform layer washed twice by mixing with chloroform/methanol/water (1/12/12). The chloroform phase was added to scintillation vials to assess acyl incorporation into G3P. Enzyme activity was measured using a scintillation counter LS 6500 multi-purpose (Beckman Coulter).

2.4 Quantitative Real-Time PCR

2.4.1 Total RNA isolation

Cell cultures were grown in three replicates to mid-exponential (OD = 0.5) and stationary (OD600 = 1.2) phases in SD media. About 70 OD600 units were collected in

50 ml falcon tubes for each phase. Pellets were resuspended in 4 ml pre-warmed AE/SDS buffer (50 mM NaOAc, 10 mM EDTA, 1% SDS, pH 5.2) followed by addition of pre- warmed acid-phenol, pH 4.5-5.5 and was added to the equal volume of glass beads.

Samples were heated at 65˚C water bath for 4 minutes and vortexed for 1 minute, repeating last 2 steps 4 times. Samples were incubated on ice for 10 minutes and spun at 3000 rpm,

5 minutes, 4˚C. The top layer was transferred to the tubes containing 4 ml pre-aliquoted

Phenol:Chloroform:Isoamyl alcohol (phenol is equilibrated to 10 mM Tris-HCl, 1 mM Na2

EDTA, pH 7.6) on ice. Samples were vortexed 3 X 30 seconds with 1 minute interval and

59 were spun at 3000 rpm, 5 minutes, 4˚C. The top layer was transferred to the tube containing

4 ml pre-aliquoted Chloroform:Isoamyl alcohol stored on ice. Samples were spun at 3000 rpm, 5 minutes, 4 ˚C. Top layer was transferred to a tube and 1/10 volume (300 – 400 µL) of 3 M sodium acetate (pH 5.2) was added and vortexed for 1 minute. An equal volume of room temperature isopropanol (3 – 4 ml) was added to the samples and mixed well by gently inverting the tubes 30-40X. Samples were stored at -20 ˚C overnight. On the following day, samples were spun at 4000 rpm, 30 minutes, 4 ˚C and poured out supernatant from each tube. The pellet was washed with 800 µL ice cold 70% ethanol by resuspending the pellet (break up pellet by pipetting up and down with P1000 pipetman) followed by spinning at 3000 rpm, 5 minutes, 4˚C and pouring out the supernatant. Samples were pulse centrifuged to collect remaining ethanol at the bottom and ethanol was removed with P200. Tubes were inverted on absorbent paper to drain all the liquid. Pellet was air- dry for about 8 minutes. RNA was dissolved in 1 ml of DEPC water by pipetting up and down. Samples were heated in 65 ˚C water bath for 5 – 10 minutes to fully dissolve the

RNA pellet. Total RNA was quantified by Nano-Drop and stored at -20 ˚C.

2.4.2 Reverse transcription

Oligo-dT premix containing nuclease free water, oligo-dT, 10 mM dNTP mix and total RNA (1 µg/µL) was prepared in a 200 µL PCR tube per reaction on ice. Oligo-dT mix was heated at 65 °C for 5 minutes; then chilled on ice for 5 minutes. First Strand Buffer mix (5X First Strand Buffer and 0.1 M DTT) was added to the oligo-dT mix. Samples were heated at 42 °C for 2 minutes then, Super Script II Reverse Transcriptase (Invitrogen) was added to the oligo-dT + FS Buffer mix and heated at 42 °C for 50 minutes. RNA samples

60 were extra heated at 72˚C for 15 minutes to inactivate the RNase enzymes. A 1/20 dilution of the cDNA was prepared and used 1 µL per qPCR reaction.

2.4.3 Quantitative Real-time PCR

qPCR reaction was done in in three independent experiments. Each set of experiment included three replicates with triple negative controls. The template of assay was designed in a 96-well plate accordingly. Master mix of 1X Power SYBR Green

(Invitrogen) in nuclease free water was prepared. 16.6 µL of mixture was added into each designated well followed by addition of 2.4 µL of primer mix with a final concentration of

0.6 µM. At the end, 1 µL of diluted cDNA (1:10) was pipetted into each designated well, following the template. 1 µL of nuclease free water was added to the negative control samples. The PCR plate was loaded into the qPCR cycler and the program (95 ˚C / 10 minutes – 95 ˚C / 15 seconds – 60 ˚C / 1 minute) was run.

Three house-keeping genes (TFC1, TAF10, UBC6) with a stable level of gene expression in the mid-exponential and stationary phases (Teste et al. 2009) were used as a background to measure the changes in a gene expression levels of GPT2WT and GPT23xA in the mid-exponential and stationary phases. ΔCt value was calculated by deduction of average of each house-keeping gene reading from the average of GPT2WT and GPT23xA readings. To calculate changes in gene expression of GPT23xA versus GPT2wt, ratio of

GPT23xA ΔCt / GPT2wt ΔCt for each house-keeping gene was calculated separately for the exponential phase and stationary phase.

2.5 Plating (growing) on different carbon source conditions

2.5.1 Oleic acid experiment

Cells were grown in SD media to mid-exponential phase (OD600 ~0.5) or Sgal-

Ura (for transformant cells). Cells were pelleted and washed with synthetic minimal medium lacking glucose (STY) containing 0.05% yeast extract, 0.002% methionine, arginine, histidine, tryptophan, and adenine sulfate, and 0.003% leucine and lysine (all from Sigma). Cells were serially diluted 1:10 starting with an OD600 of 0.5. Cells then were spotted on the STY medium containing 1% Tergitol + 0.1% (3.5 mM) or 0.17% (6 mM) or 0.23% (8 mM) or 0.28% (10 mM) oleic acid plates applying replica plating technique. STY medium + 1% Tergitol used as a control plate. Plates were incubated at 30

°C for 4 days. Plates were imaged using a GelDoc system (Bio-Rad).

2.5.2 Plating on fermentable and non-fermentable carbon sources

Transformant cells carrying cloned GPT2 phospho-mutants in a pYES vector under

GAL-promoter were grown in Sgal-Ura (2% galactose) to mid-exponential phase (OD600

~0.5). Cells were serially diluted 1:10 starting with an OD600 of 0.5 and spotted on SD-

Ura (2% glucose), Sgal-Ura (2% galactose) and STY-Ura medium containing 3% glycerol and 2% ethanol applying replica plating technique. Plates were incubated at 30°C for 4 days. Plates were imaged using a GelDoc system (Bio-Rad).

2.5.3 Gradual and acute glucose depletion experiment

For the gradual glucose depletion experiment, cells were grown in SD media (2% glucose) for 4-5 days. Every day, cells were serially diluted 1:10 starting with an OD600 of 0.5 and were spotted on the same media plates applying replica plating technique. Plates were incubated at 30 °C for 4 days. Plates were imaged using a GelDoc system (Bio-Rad).

For the acute glucose depletion experiment, cells were grown in SD media to mid- exponential phase (OD600 ~0.5). Cells were pelleted and washed with SD media containing 0.4% glucose and were grown for 51 days. Every 4 days, cells were serially diluted 1:10 starting with an OD600 of 0.5 and were spotted on the same media plates applying replica plating technique. Plates were incubated at 30 °C for 4 days. Plates were imaged using a GelDoc system (Bio-Rad).

2.5.4 Different drug treatments

2.5.4.1 Cerulenin

Cells were grown in YPD media for 5 days to induce formation of large LDs

(OD600~15). Cells were serially diluted 1:10 starting with an OD600 of 0.5. Cells then were spotted on YPD medium + ethanol (control) or YPD medium + 5 µg/mL cerulenin

(Sigma) applying replica plating technique. Plates were incubated at 30 °C for 4 days.

Plates were imaged using a GelDoc system (Bio-Rad).

2.5.4.2 Myriocin

Cells were grown in SD media to mid-exponential phase (OD600 ~0.5). Cells were serially diluted 1:10 starting with an OD600 of 0.5. Cells then were spotted on SD medium

+ methanol (control) or SD medium + 2 µM Myriocin (Sigma) applying replica plating

63 technique. Plates were incubated at 30°C for 4 days. Plates were imaged using a GelDoc system (Bio-Rad).

2.5.4.3 Aureobasidin A

Cells were grown in SD media to mid-exponential phase (OD600 ~0.5) and were serially diluted 1:10 starting with an OD600 of 0.5. Cells then were spotted on SD medium

+ ethanol (control) or SD medium + different concentrations of Aureobasidin (0.03, 0.05,

0.06, 0.07, 0.08, and 0.1 µg/mL) applying replica plating technique. Plates were incubated at 30 °C for 4 days. Plates were imaged using a GelDoc system (Bio-Rad).

2.6 Microscopy analysis

For confocal microscopy, a Leica TCS SP5 II confocal laser scanning microscope was used. Images were acquired with HCX PL APO CS 63/1.4 oil UV immersion objective using Leica Application Suite Advanced Fluorescence (LAS AF) software. For image acquisition, samples were excited using visible laser at 488 nm and 555 nm for EGFP and mRFP, respectively, and fluorescence was detected using Leica HyD detector with the following emission settings: EGFP (508–540 nm) and mRFP (563-580 nm). Transmitted light for bright field images were detected using a PMT detector.

Some images were acquired using a Zeiss Imager Z.1 epifluorescence microscope with an AxioCam MRm camera (Zeiss) fitted with EC Plan Neofluar 100/1.3 Oil M27 objective lens using Axiovision Rel. 4.7 software. Cells were excited at 488 nm and 555 nm for GFP and RFP, respectively, and fluorescence was detected in the range of 515-565 nm for GFP signal and 565-605 nm for RFP, 495-503 nm for BODIPY, and 505-553 nm for Nile red. The 3D projections were rendered from deconvolved z-stacks images acquired

64 with a Quorum Technologies WaveFX Spinning Disk Confocal Microscope (Quorum

Technologies Canada) controlled by MetaMorph acquisition software (Molecular Devices,

LLC) and equipped with Hamamatsu Dual EMCCD cameras and an ASI motorized XY stage. Images were acquired using a 63x oil immersion objective, NA 1.4. 3D rendering, deconvolution (90% confidence) and analysis was performed with Volocity 6.3

(PerkinElmer Inc.) and/or ImageJ (v. 1.47 bundled with 64-bit Java). Images were processed with Adobe Photoshop and Illustrator (Adobe Systems Inc.). Quantification of images was completed using cell counter plug-in ImageJ.

2.6.1 Lipid droplet staining with Nile red

2 µL of 1 mg/ml Nile red dye (Sigma) was added to the OD600 of 1 of cells, to have final concentration of 2 µg/µL of dye. Cells were incubated for 30 minutes at 30ºC. Cells were washed 2 times with fresh media and were analyzed with fluorescence microscopy.

2.6.2 Lipid droplet staining with BODIPY 493/503

1 µL of 1 mg/ml BODIPY 493/503 dye was added to the OD600 of 1 of cells, to have final concentration of 1 µg/µL of dye. Cells were incubated for 5 minutes at 30 ºC.

Cells were washed 2 times with fresh media and were analyzed with fluorescence microscopy.

2.6.3 Lipid droplet analysis with AutoDot

Cell were grown for 48 hours (late stationary phase) in SD-Ura media allowing cells to form large LDs. Cells were then incubated with 200 µM AutoDot dye (FroggaBio)

65 for 20 minutes to stain lipid droplets. Cells were spun down and re-suspended in fresh media two times to wash out the dye. Cells were grown for 20 minutes in fresh media to start growth resumption. Samples were object to spinning disk microscopy live imaging using a dual camera. Intensity for both green confocal laser 491 nm and red confocal laser

561 nm was set at 10 in order to capture the GFP (excitation: 488 nm, emission: 509 nm) and RFP (excitation: 555 nm, emission: 584 nm) signals. Intensity for blue confocal laser

406 nm was set at 15.5 to capture blue signal (excitation: 405 nm, emission: 455 nm). Step size for Z-series was 0.15 µm. Microscopy images were analyzed using Volocity and

ImageJ software.

2.6.4 Cerulenin treatment

Cells were grown in YPD for 5 days to form large LDs. Cells were pelleted and resuspended in fresh YPD. For microscopy analysis, cells were loaded on an agar pad containing 5 µg/mL cerulenin + 1 µg/mL BODIPY. To do so, 50 µL of 2% agarose in YPD medium was prepared at 60ºC and 2.5 µL of 100 µg/mL cerulenin and 5 µL of 10 µg/mL

BODIPY was added. Control experiment was done in the absence of cerulenin. Cells were tracked over time for about 3 hours.

2.7 Lipid droplet analysis with Typhoon method

For this experiment lipid droplets were stained with Nile red same as done for the microscopy analysis. Serial dilutions of 1:2 was prepared in a 96-well fluorescence micro- plate in duplicate for the stained cells and unstained cells as a control. Fluorescence intensity of cells was measured at 532 nm using LPG filter - Typhoon FLA 9500 (GE

Healthcare Life Sciences). Results were analyzed with ImageQuant TL and ImageJ programs.

2.8 Yeast lipid extraction

Yeast culture were grown in SD media to late stationary phase for 48 hours. OD600 of 150 of cells was collected by spinning down at 3300 rpm (2500g) for 5 minutes. Cells were grown in fresh media to start growth resumption for 2 hours. Cells were pelleted by spinning down at 3300 rpm (2500g). The cell pellet was washed with 1 mL 1 M sorbitol and re-spun briefly. Sorbitol was removed by Pasteur pipette. This step was repeated. 1 mL

CHCL3/MeOH (1/1) was added, vertex and transferred to a 2 mL bead beater vial 1/8 filled with 0.5 mm acid washed beads. The bead beating step was done for 60 seconds (in

BioSpec Bead Beater apparatus) at 4 ºC. Cells were transferred to a test tube with a Pasteur pipette. Beads were rinsed with 1 mL CHCL3/MeOH (1/1) and combined with the first extract. 0.5 mL CHCL3/MeOH (2/1), 0.5 mL CHCL3, and 1.5 mL ddH2O were added and mixed well by vertexing followed by spinning down at 2500 rpm for 10 minutes to phase separate. The organic fraction was saved by removing the aqueous phase and aspirating off the protein layer. 2.5 mL of artificial aqueous phase (3:48:47 CHCL3/MeOH/ddH2O) was added to the samples and mixed well by vortexing. Samples were spun down at 2500 rpm for 10 minutes to phase separate. This step was repeated. The organic phase was transferred to the pre-weighted vials. Lipid film was dried by evaporating excess of chloroform using liquid nitrogen.

2.9 Neutral lipid Thin-layer Chromatography (TLC) experiment

TLC experiment was done by two methods using different solvents system with

Silica gel on TLC Aluminum foil plate (Sigma-Alderich). One was the FOLCH method in which the plate was washed with 6:4 methanol/ ethyl acetate. 100 µg of lipid extracts were spotted on the plate following the labels at the 10 mm start line. Samples were run to 40 mm in 80:16:4 dichloroform/ethyl acetate/acetone. The plate was dried for 5 minutes.

Samples were run to 55 mm in 80:16:4 dichloromethane/ethyl acetate/acetone. The plate was dried for 5 minutes. Samples were run to 68 mm in 90:10 hexanes/ethyl acetate/acetone. The plate was dried for 5 minutes. Samples were run to 80 mm in 95:5 hexanes/ ethyl acetate. Plate was dried for 5 minutes. Samples were run to 90 mm in hexanes. Lipid spot separation were visualized using iodine vapour.

With the other method, lipids were separated using the solvent system ether/diethyl ether/acetic acid (80:20:1, v/v/v) using TLC silica gel plate (60 A Whatman). Lipid samples were dissolved in CHCl3/MeOH (2:1) and 100 µg of total lipid extracted samples were spotted. Plates were developed until the solvent front reaches the top of the plate. Plates were dried under the fume hood, use a hair dryer at low heat. This step took several minutes.

Separated bands were visualized by iodine vapor or using charring by dipping the plate for

10 seconds into a solution containing 50% ethanol in water, 3.2% H2SO4, and 0.5% MnCl2.

Char the plate for 30 minutes at 120 ˚C.

TAG and DAG concentrations were measured using a standard curve from a gradient concentration of TAG or DAG.

2.10 Yeast lipid droplet isolation and analysis

Yeast cells expressing endogenous GPT2wt-V5-6xHis or GPT3xA-V5-6xHis and

RFP-fused ERG6 and carrying plasmid including C1δ-GFP under constitutive promoter were grown for 24 hours in SD-Ura. Cultures were diluted (10 mL of the pre-culture into

500 mL fresh SD-Ura) and grown for 24 hours. Cells were grown to late-exponential/early- stationary. Total OD600 of 1000 cells were collected using Sorvall SLC-3000 rotor at 4400 g for 15 minutes and re-suspended in 500 mL fresh SD-Ura media and were grown for 30 minutes to start growth resumption. Cells were spun down briefly to remove the media and re-suspended in ddH2O and were spun down again. About 4 grams of cell pellet was collected. Cells were shaken for 10 minutes at 30 ºC in 15 mL solution A (0.1 M Tris/H2SO4 buffer (pH 9.4) + 10 mM DTT). Cells were transferred to 50 mL Falcon tube and spun down at 1950 g for 5 minutes using Sorvall Legend RT benchtop centrifuge. Cells were resuspended in 30 mL of solution B (1.2 M sorbitol + 20 mM KH2PO4 (pH 7.4). Briefly, cells were collected and treated with zymolyase 100T (0.4 mg per gram cell wet weight) to obtain spheroplasts. Samples were shaken at 200 rpm speed at 30 ºC for 20 minutes and then at 80 rpm speed at 30 ºC for 20 minutes. After 40 minutes, turbidity of cultures was checked to ensure if zymolyase treatment worked properly. Spheroplasts were washed twice with solution B and spun down using SLC-3000 rotor at 2800 g for 5 minutes. All following steps were done on ice. Re-suspended spheroplasts were washed in total volume of 4.6 mL buffer A + protease inhibitors (10.4% ficoll + 1 mM PMSF + 1X ROCHE inhibitors + 3 µg/µL pepstatin). Cells were homogenized and cell homogenate was centrifuged using SS-34 rotor at 5000 g for 5 minutes at 4 ºC. Pellet was re-suspended in

2.781 mL of buffer A + protease inhibitors. Samples were centrifuged by swinging type

69 ultracentrifuge using SW-40 rotor at 100,000 g for 60 minutes at 4 ºC. The white floating layer was collected and resuspended very gently in 5 mL buffer A. 5 mL sample was overlayed with 5 mL buffer B (8% ficoll) and centrifuged at 100,1000 g for 30 minutes at

4 ºC. The white floating layer was collected and resuspended gently in buffer C (8% ficoll

+ 0.6 M sorbitol) followed by homogenizing 8x strokes using 30 mL homogenizer. 2.5 mL buffer D (0.25 M sorbitol) was added by syringe on top of 7 mL of samples, ending with a total volume of 10 mL. Samples were centrifuged at 100,000 g at 4 ºC. The white floating fraction representing lipid droplets and also the pellet fraction was collected and saved at -

20 ºC.

2.11 Mass Spectrometry Analysis

Prepared LD and pellet layers from LDs isolation of wildtype GPT2 and 3A cells were subjected to mass spectrometry analysis. Mass spectrometry analysis was done using

GFP-trap beads to enrich associated with C1δ-GFP probe. To do so, 500 µL of LD and 50

µL of pellet samples were used for incubation with GFP-trap beads and agarose beads as the negative control. Samples were prepared by addition of final concentration of 0.3 M

NaCl and 1 mM EDTA. Prepared samples were transferred to the tubes containing 25 µL and 50 µL of GFP-trap and agarose beads, respectively. Samples were rotated at medium speed for 1 hour at 4 ºC. Samples then washed 3 times with washing buffer (10 mM

Tris/HCl, 150 mM NaCl, 0.5 mM EDTA). 20 µL of 4X SDS gel loading buffer and 20 µL of 1X PBS both prepared in milli-Q ddH2O were added to the beads. Samples were boiled for 10 minutes and were spun down at 5200 rpm (2500 g) for 2 minutes. Supernatant was collected and 30 µL of the supernatant were run on a SDS-PSGE gel (5% stacking and

10% resolving) prepared with milli-Q ddH2O. Samples were cut after running 0.5 cm below from the stacking gel’s bottom line (0.5 x 0.5 cm2) and stored in a tube containing 1% acetic acid. Samples were sent to the Southern Alberta Mass Spectrometry Centre at the

University of Calgary for analysis. Scaffold (version Scaffold_4.4.4, Proteome Software

Inc., Portland, OR) was used to validate MS/MS based peptide and protein identifications.

Peptide identifications were accepted if they could be established at greater than 20.0 % probability by the Peptide Prophet algorithm (Keller et al., 2002) with Scaffold delta-mass correction. Protein identifications were accepted if they could be established at greater than

20.0% probability and contained at least one identified peptide. Protein probabilities were assigned by the Protein Prophet algorithm (Nesvizhskii et al., 2003). Proteins that contained similar peptides and could not be differentiated based on MS/MS analysis alone were grouped to satisfy the principles of parsimony. Proteins sharing significant peptide evidence were grouped into clusters.

2.11.1 Clustering Analysis

Clustering analysis was done to find unbiased and significant patterns in the large amounts of data obtained from the MS experiments. Two databases widely used for finding shared traits, themes, and connections between genes/proteins for clustering analysis are the Gene Ontology (GO) Consortium and Kyoto Encyclopedia of Genes and Genomes

(KEGG). GO is a database dedicated to consistently and accurately describing all gene products in a species-independent manner (Ashburner et al., 2000). To do this, three structured ontologies were developed; biological processes, cellular components, and molecular functions. The different terms within three ontologies can be organized into

71 multiple levels; where the first levels are more general and higher levels encompass very specific terms. KEGG is a separate database that uses molecular-level information to understand high level functions and assigns genes to functional pathways, therefore providing more detailed information on overall systems and pathways (Kanehisa, 2002).

FunSpec (Robinson et al., 2002), an online clustering tool provided protein enrichment data in each of the three GO classes. The significance score (p-value) associated with each cluster represents the probability that the respective list and any given functional category occurs by chance (Robinson et al. 2002). The R package clusterProfiler (Yu et al., 2012) was also used to obtain unbiased enrichments of proteins based on the same GO classes, with the additional ability to provide a wide range of visualization methods to accurately depict the clustering results. Despite this, clusterProfiler proved to be not as beneficial as

FunSpec, as the top clusters were usually repetitive and unspecific. However, FunSpec is limited in its visual outputs, as its only output is in a table format.

Due to the differences and redundancy in the databases, as well as mass amount of information for each gene, relationships between genes and terms proved to be difficult to understand and visualize (Bindea et al., 2009). Therefore, the ClueGO plugin for

Cytoscape (Shannon et al., 2003) was used, as it obtains complementary data from both

GO and KEGG databases to extract non-redundant biological information for large lists of genes (Bindea et al., 2009). ClueGO provided a variety of outputs, such as histograms, pie charts, and maps, displaying the relationships of GO terms based on the similarity of their associated genes and their significance (Bindea et al., 2009). An additional plugin,

CluePedia, created maps linking the terms with their related genes (Bindea et al., 2013). In the maps, nodes represent the enriched terms, while the edges that connect the nodes are

72 calculated using kappa statistics (Huang et al., 2007). In brief, the kappa score is a chance- corrected value of co-occurrence between two genes, taking into account both the observed and chance co-occurrence (Huang et al., 2007). P-values were calculated with the Fisher

Exact Test and corrected using the Bonferroni step-down method (Bindea et al. 2009).

2.12 Statistical analysis

Statistical analyses were done using Graphpad Prism and Excel software and T-test was used for statistical significance analysis.

2.13 Comparative Genomic Survey

As described in (Smart et al., 2014).

Chapter 3 –Phylogenetic study of fungal acyltransferases

Results obtained from these studies were published in 2014 (Smart et al. 2014).

3.1 Introduction

GPATs are key metabolic players in lipid metabolism that understanding their evolutionary history allows us to discover the cellular evolution of membrane biogenesis, trafficking, signalling and to linking organellar and metabolic evolution in this set of eukaryotic taxa.

Despite of the discovery of GPATs in bacteria and mitochondria (Lightner et al.,

1983; Shin et al., 1991), eukaryotic microsomal GPATs remained elusive for several

73 decades and just recently the new genes encoding these enzymes have been identified

(Zheng et al., 2001; Cao et al., 2006; Lewin et al., 2010).

Studies in model organisms like mice and yeast resulted in the identification of the genes involved in the de novo glycerolipid biosynthetic pathways. There are four identified mammalian GPAT isoforms encoded by independent genes, two isoforms mainly associate with mitochondria (GPAT1 and GPAT2) and the other two (GPAT3 and GPAT4) have an

ER localization (Gimeno et al., 2008; Lewin et al., 2010). There is an additional acyltransferase enzyme in animals with a peroxisomal localization which catalyzes a similar reaction, but instead of G-3-P, dihydroxy-acetone phosphate (DHAP) is used as a substrate, therefore called dihydroxy-acetone phosphate acyltransferase (DHAPAT)

(Ofman et al., 1994).

Two unique genes, SCT1/GAT2 and GPT2/GAT1, code for two GPATs were identified in Saccharomyces cerevisiae, both of which are integral membrane proteins located in the ER (Zheng et al., 2001; Bratschi et al., 2009).

Redundancy in this first acylation step catalyzed by GPATs is observed in a broad range of organisms spanning form yeast through a complex organism like human, but intriguingly fungal enzymes share a low sequence identity (~11%) with their known animal counterparts. We were intrigued by this lack of sequence homology and by the fact that no mitochondrial GPATs were identified in yeast, although GPAT activity has been detected in mitochondrial fractions.

3.2. Goals

The aim of this phylogenetic study was to understand the evolutionary history of

GPAT proteins in opisthokonts, which includes fungi and metazoans. This would help us

74 to determine how representative the Saccharomyces cerevisiae GPAT complement is of other fungal taxa; and how representative the mammalian complement is of animal taxa; and critically assess how the sets of enzymes correspond to one another.

By organizing all available genomic databases, we created a systematic structure to investigate the possible evolutionary link among GPATs in animal and fungal species and also to cluster the related sequences for identifying unique protein features within each category.

The additional goal was to ensure Gpt2p and Sct1p were the only GPATs present in yeast.

3.3 Results

3.3.1. Searching for Sct1p/Gpt2p identifies two undescribed deep clades of fungal GPATs

To probe human-like and yeast-like GPATs, valid protein sequence of a broad range of organisms from both Metazoa and Fungi were used (Table 3.1). Yeast GPAT homologues in these organisms were first identified and validated using proper sequence data base programs.

The preliminary step was done by comparing the sequence similarity between Sct1p and Gpt1p that represent the entire GPAT complement in Saccharomyces cerevisiae

(Zheng et al., 2001; Zaremberg et al., 2002). By analyzing a broad variety of both fungal and metazoan lineages including 43 databases form available genome sequencing projects, major branching points in the opisthokont clade which includes fungi and metazoans, were recovered (Figure 1) (Smart et al. 2014). This set of analysis revealed high-scoring hits for

Sct1p/Gpt2p in all fungal species, except in E. cuniculi, a parasite with a reduced and compact genome (Hardie and Carling 1997; Katinka et al. 2001). Surprisingly, nonfungal unicellular opisthokonts from the Filasterea, Ichthyosporea, and Choanozoa also acquired

Sct1p/Gpt2p homologues (Figure 3.1) (Smart et al. 2014).

Figure 3.1- Opisthokont relationships and GPAT evolution. A) Tree illustrating the relative evolutionary history for the eukaryotic emergence, loss and diversification/duplication of GPATs in opisthokonts, in lineages with genome sequences available in the fungi and metazoans with additional taxa from the base of the Opisthokonta supergroup. The apusozoan, T. trahens, was used as the outgroup. The group labeled “SACK” belongs to Saccharomycotina and involves S. cerevisiae, A. gossypii, C. glabrata, and K. lactis. To note the Pezizomycotina and Taphrinomycotina form a paraphyletic group of ‘non- Saccharomycotina ascomycota’ which we found to be a useful grouping when considering the evolution of GPATs and are thus colour coded identically. Similarly, all non-tetrapoda metazoans are coded in the same colour. In order to avoid false negative or false positives in our evolutionary deductions, we do not infer any events on

76 lineages where only a single genome was examined. B) Results from the comparative genomic survey and phylogenetic analysis. Individual species from the survey are color coded as in A) and grouped according to established taxonomic classification. The dendrogram is schematic of relationships only and the branch length is not representative of evolutionary distance. Symbols indicate the presence of at least one isoform of a given protein as verified by BLAST, Reciprocal BLAST, hidden Markov model searches, and phylogenetic data as follow: orthologs of Sct1p/Gat2p( ), Gpt2p/Gat1p( ), as well as new fungal fGPAT-A ( ) or fGPAT-B (□) and orthologs of human mitochondrial GPATs (mitoGPATs) GPAT1(▴), GPAT2(▾), putative mitoGPATs (▵), GDPAT ( ), DHAPATs (⧫) and putative DHAPATs (pDHAPAT ◊), and orthologs of human microsomal GPATs (eGPATs) GPAT3(▸) and GPAT4(◂) and putative eGPATs (→). Putative orthologs of Sct1p and Gpt2p are shown in grey while strongly supported orthologs are shown in solid black. Blank cells indicate no hits. (p) denotes GDPATs with PTS1 prediction. Designation in a particular orthologous group denotes a prediction of substrate specificity, based on assumed retention between orthologues, and provides hypotheses for future functional testing. (*) The Rhizopus acyltransferase gene groups with moderate support with the DHAPAT genes, but this placement may well be due to its highly divergent sequence. Protein sequences used to initiate the searches: yeast Sct1p (NP_009542.1), yeast Gpt2p (NP_012993.1), human GPAT1 (NP_065969.3), human GPAT2 (NP_997211.2), human DHAPAT (NP_055051.1), human GPAT3 (NP_116106.2), human GPAT4 (NP_848934.1). Note: Putative fungal GDPATs display a peroxisomal target sequence (PTS1) and were hits in a search initiated with mitochondrial human GPAT1. Figure obtained from (Smart et al. 2014).

These analyses allow us to have a better insight into the evolutionary relationships of the yeast-like GPATs identified by our homology investigations. Phylogenetic data revealed two major deep-branching clades which are strongly supported by all three applied phylogenetic analysis methods (Figure 3.2).

Figure 3.2. Phylogeny of fungal GPAT homologues found in opisthokonts. Phylogenetic tree of the fungal GPATs. In this and all other phylogenies, node support values are shown in order of Bayesian posterior probabilities, PhyML bootstrap percentages and RaxML bootstrap percentages, or are symbolized as dots, colorized as per the inset to indicate statistical strength. Clades for orthologues of Sct1p and Gpt2p are shaded. Figure made by H. Smart (Smart et al. 2014).

These two clades are only detected in members of Basidiomycota and fungal basal opisthokonts. Each clade includes a putative fungal GPAT (fGPAT) from the outgroup T trahens taxon and clearly separated the closest yeast homologues (fGPAT-A proteins) from a second fGPAT related group (fGPAT-B proteins). Categorized branching order of all fGPAT-A and Sct1/Gpt2 homologues form the largest clade. Division of branches

78 surprisingly continued to the Sct1p/Gpt2p paralogues and was restricted to the

Saccharomycetaceae lineages that diverged after the split with Yarrowia (Smart et al.

2014). These findings indicate that yeast-like GPATs exist in foundation organisms of opisthokonta lineages, which were conserved and diversified in the descendant fungal lineages but sculpted by gene loss in the metazoan ones.

Interestingly, this study identified a well-defined fungal cluster that preceded the emergence of the duplicated metazoan branches. These fungal genes, together with the homologues in the basal outgroups were defined as GDPATs (G-3-P or DHAP acyltransferases) (Figure 3.3) (Smart et al. 2014). GDPAT designation, represents the pre- duplicate status to the DHAPAT and mitoGPAT clades and explains our lack of prediction of substrate preference. Our findings strongly propose the presence of a novel pathway for glycerolipid synthesis in fungal lineages, ancestral to those currently linked to peroxisomes and mitochondria in metazoans.

Figure 3.3- Phylogenetic tree of GDPAT-related genes in opisthokonts. The clades of mitochondrial GPATs (called mitoGPAT, then enumerated), putative DHAPATs (called pDHAPAT, then enumerated) and fungal GDPATs are shaded. Darker shading in each cluster highlights biochemically characterized enzymes from vertebrates, which includes mammalian GPAT1 and DHAPAT homologues. Figure made by H. Smart (Smart et al. 2014).

3.3.2. Comparison of signature motifs between characterized and novel acyltransferases identified in this study

Discovery of new putative GPATs present in either yeast or mammalian-like enzymes allows us to investigate unique signature features of each GPAT subclasses using different protein sequences from various organisms and process comparative analysis. The

GPAT/DHAPAT proteins of both yeast and humans belong to the lysophospholipid acyltransferase (LPLAT) superfamily which contain a signature acyltransferase domain

(pfam01553) consisting of four distinct motifs (motif I, II, III and IV) arranged sequentially

(Lewin et al., 1999; Takeuchi et al., 2009). This relevant feature made us to focus on four diagnostic motifs essential for GPATs enzymatic activity (Lewin et al., 1999; Santiago et al., 2004) as these regions are potentially required for substrate specificity and catalysis within each acyltransferase subclass. Alignment analysis for each set of acyltransferase sequence was done, sequences of 4 motifs were identified and, conserved residues compared across all subclasses (Table 3.2, Figure 3.4)

Figure 3.4- Sequence alignments of the catalytic motifs. Alignment of catalytic motif sequences in strongly supported orthologs of biochemically characterized acyltransferases in each of the eGPATs, fungal GPATs (fGPATs-SACK), mitoGPATs and DHAPATs groups were compare to those of fungal GDPATs. Figure obtained from (Smart et al. 2014).

By analyzing the consensus sequences, some differences were detected in subclasses (Table 3.2). Notably, motif I of fungal GPATs (fGPAT-A and B) consist of uniform histidine and aspartic acid residues which are consistently separated by five residues (HX5D) while in all other analyzed acyltransferases, are separated by four residues

(HX4D) (Smart et al. 2014).

The largest analyzed group belongs to the fungal GPAT-As (41 sequences) with the highest degree of conservation among motifs, despite of diversity in their taxonomy.

Motif I consist of unique conversion of QF residues within the HX5D sequence. Motif III is identified as the most conserved motif including a uniform FPEGGSHD sequence in all proteins from this cluster (Figure 3.5) (Smart et al. 2014).

Figure 3.5. Sequence alignments of the catalytic motifs in strongly supported orthologs of fungal GPATs. Proteins were aligned using Uniprot Align software and acyltransferase motifs recognized as proposed in (Lewin, Wang, and Coleman 1999). Positive, negative and aliphatic residues are highlighted in green, red and yellow respectively. Abbreviations: agos, Ashbya gossypii; ama, Allomyces macrogynus; anid, Aspergillus nidulans; bdet, Batrachochytrium dendrobatidis; bfuc, Botryotinia fuckeliana; cal, Candida albicans; cgla, Candida glabrata; cimm, Coccidioidies immitis; ccin, Coprinopsis cinérea; cowc, Capsaspora owczarzaki; dha, Debaryomyces hansenii; enzn, Encephalitozoon cuniculi; gza, Gibberella zeae; klac, Kluyveromyces lactis; lbi, Laccaria bicolor; mbre, Monosiga brevicolis; ncra, Neurospora crassa; pipa, Pichia pastoris; pis, Pichia stipitis; puc or pgra, Puccinia graminis; rhiz, Rhizopus oryzae; sarc, Sphaeroforma arctica; scer, Saccharomyces cerevisiae; sja, Schizosaccharomyces

84 japonicus; spom, Schizosaccharomyces pombe; spun, Spizellomyces punctatus; sros, Salpingoeca rosetta; tadh, Trichoplax adhaerens; ttra, Thecamonas trahens; umay, Ustilago maydis; ylip, Yarrowia lipolytica. Figure obtained from (Smart et al. 2014).

3.4 Discussion and conclusions

This study identified four ancient groups of acyltransferases present in lineages named (1) fungal GPAT-A, (2) fungal-GPAT-B (3) mammalian ER-like GPATs

(eGPATs), and (4) GDPATs, which then duplicated sometime in the holozoa into (5-a) mitochondrial GPATs (mitoGPATs) and (5-b) DHAPATs (Figure 3.1). Based on our findings, it is assumed that GPATs in eukaryotic ancestors of opisthokonts were probably more complex than some of their descendants. Putative GPATs were sculpted via gene loss in fungi and metazoan lineages that can be explained as a model of reductive evolution.

The evolutionary process conducting the conservation of a pre-existing type of microsomal acyltransferase and loss of the other type highlights critical roles of GPATs in cellular mechanism specifically designed to each of these lineages. Microsomal GPATs have shown limited distribution, however the larger family of mitoGPATs, DHAPATs and novel GDPATs are identified among all metazoan and fungi lineages. This represents a novel finding of our study, as no such proteins were previously described in fungi, potentially indicative of an undescribed metabolic capacity in this lineage. According to our comparative genomic studies and phylogenetic findings, major evolutionary questions about GPATs in opisthokons have been addressed. Remarkably, we have shown that GPAT complements in S. cerevisiae and human are not entirely representative of the fungal and holozoan complements, respectively. Discovery of a GDPAT-related clade of fungal genes with putative peroxisomal targeting might reveal an undescribed metabolic capacity of this

85 lineage. The overall pattern of GPAT distribution homologues with high complexity in the ancestors of opisthokont clade and loss and sculpting of the complement in the descendent lineages highlights the existence of a reductive evolution.

Table 3.1. Names of species and specific strains used in the proteomic analysis of opisthokont GPATs. Whole reference sequence protein databases were downloaded from the NCBI (Ejsing et al. 2009) or The Broad Institute (Takeuchi et al., 2009) in FASTA format. From (Smart et al. 2014).

Species Strain Taxid Source Allomyces macrogynus Allomyces macrogynus ATCC 38327 578462 Broad Institute Amphimedon queenslandica Amphimedon queenslandica 400682 NCBI Ashbya gossypii Ashbya gossypii FDAG1 1034331 NCBI Aspergillus nidulans Aspergillus nidulans FGSC A4 227321 NCBI Batrachochytrium dendrobatidis Batrachochytrium dendrobatidis JAM81 684364 NCBI Botryotinia fuckeliana Botryotinia fuckeliana B05.10 332648 NCBI Branchiostoma floridae Branchiostoma floridae 7739 NCBI Caenorhabditis elegans Caenorhabditis elegans 6239 NCBI Candida albicans Candida albicans SC5314 237561 NCBI Candida glabrata Candida glabrata CBS 138 284593 NCBI Capsaspora owczarzaki Capsaspora owczarzaki ATCC 30864 595528 NCBI Coccidioidies immitis Coccidioides immitis RS 246410 NCBI Coprinopsis cinerea Coprinopsis cinerea okayama7#130 240176 NCBI Debaryomyces hansenii Debaryomyces hansenii CBS767 284592 NCBI Drosophila melanogaster Drosophila melanogaster 7227 NCBI Encephalitozoon cuniculi Encephalitozoon cuniculi GB-M1 284813 NCBI Gallus gallus Gallus gallus 9031 NCBI Gibberella zeae Gibberella zeae PH-1 229533 NCBI Homo sapiens Homo sapiens 9606 NCBI Kluyveromyces lactis Kluyveromyces lactis NRRL Y-1140 284590 NCBI Laccaria bicolor Laccaria bicolor S238N-H82 486041 NCBI Monosiga brevicolis Monosiga brevicolis 431895 NCBI Mus musculus Mus musculus 10090 NCBI Nematostella vectensis Nematostella vectensis 45351 NCBI Neurospora crassa Neurospora crassa OR74A 367110 NCBI Pichia pastoris Komagataella pastoris CBS 7435 981350 NCBI Pichia stipitis Scheffersomyces stipitis CBS 6054 322104 NCBI Puccinia graminis Puccinia graminis f. sp. tritici CRL 75-36-700-3 418459 NCBI Rhizopus oryzae Rhizopus oryzae RA 99-880 246409 NCBI Salpingoeca rosetta Salpingoeca sp. ATCC 50818 946362 NCBI Schizosaccharomyces japonicus Schizosaccharomyces japonicus yFS275 402676 NCBI Schizosaccharomyces pombe Schizosaccharomyces pombe 972h- 284812 NCBI Sphaeroforma arctica Sphaeroforma arctica JP610 667725 Broad Institute Spizellomyces punctatus Spizellomyces punctatus DAOM BR117 645134 Broad Institute Strongylocentrotus purpuratus Strongylocentrotus purpuratus 7668 NCBI Thecamonas trahens Thecamonas trahens ATCC 50062 461836 Broad Institute Trichoplax adhaerens Trichoplax adhaerens 10228 NCBI Ustilago maydis Ustilago maydis 521 237631 NCBI Xenopus tropicalis Xenopus (Silurana) tropicalis 8364 NCBI Yarrowia lipolytica Yarrowia lipolytica CLIB122 284591 NCBI

Table 3.2. Acyltransferase motifs and distance between motifs (DBM) in microsomal yeast-like (fGPAT-A), metazoan ER-like (erGPAT), mitoGPATs, DHAPAT and fungal GDPAT (fGDPAT) sets. From (Smart et al. 2014).

All proteins identified in this work for each category were aligned using Uniprot Align software and acyltransferase motifs recognized as proposed in (Lewin, Wang, and Coleman 1999). Number of proteins containing all four motifs used for each alignment is shown in brackets. aas proposed in (Lewin, Wang, and Coleman 1999). Nomenclature used as defined in (Aasland et al. 2002): π = P, G, A and S (short chain). φ = Hydrophobic amino acids (V, I, L, F, W, Y and M). Residues highlighted in red are the hallmark of each motif. * N.vectensis mGPAT [XP_001640722.1] and C. owczarzaki mGPAT gi|320164443| have 65 residues between Motifs 2&3.

Chapter 4 – Identification of phosphorylation sites in Gpt2p

4.1 Introduction

Signaling circuits provide a sensitive, specific and potent response to external stimuli through fine tuning of protein stability, enzyme activity, subcellular localization, and protein-protein interactions. This dynamic regulation is mediated through fast kinetics and reversibility of post-translational modifications (PTMs). Phosphorylation is one of the key PTMs occurring in eukaryotic cells where a phosphate group can be quickly added and removed at serine, threonine, and tyrosine residues by kinases and phosphatases, respectively. Indeed, adding or removing a negatively-charged phosphate group on a protein might change its physico-chemical properties, kinetics, stability and dynamics.

Phosphorylation is a reversible PTM conserved in all eukaryotic signaling pathways, and the most frequent PTM occurring in Saccharomyces cerevisiae (Vlastaridis et al. 2017).

Phospho-proteomics studies deposited in PhosphoPep (Bodenmiller et al. 2007) show that more than 2300 yeast proteins are present as phosphoproteins, covering nearly all cellular processes. Therefore, an estimated one third of the proteome is phosphorylated (Cohen

2002; Stark et al. 2010), indicating that kinase and phosphatase- mediated signaling orchestrates one of the most important regulatory mechanisms in cellular processes. A complex network of about 160 protein kinases and phosphatases controls phosphorylation in yeast (Colman-Lerner et al. 2011). There are about 127 yeast kinases (Rubenstein et al.,

2007), 97 belong to 51 subfamilies that are shared with humans (Manning et al. 2002) and there are about 33 identified phosphatases (Breitkreutz et al., 2008). Frequently, proteins contain multiple phosphorylation sites which modulate distinct aspects of the target protein, providing an expanded range of regulation in response to environmental cues

(Nishi et al., 2014). Phosphorylation at different sites on the same protein might have an opposite impact on protein behavior causing activation or inhibition (Salazar and Höfer

2009; Hoffmann et al. 1993). Phosphoproteins can be positively or negatively regulated by one or more kinases affecting their activity (in the case of enzymes), localization, stability, conformation and/or their interaction with other proteins (Swaney et al. 2013; Oliveira et al. 2012; Z. Chen et al. 2016). Yeast GPATs Gpt2p and Sct1p are phosphoproteins

(Bratschi et al., 2009; Marr et al., 2012) but is not known which aspects of these enzymes are regulated through phosphorylation and which kinases and phosphatases are directly involved in their modification.

4.2 Goals

The serine-rich C-terminal tail of Gpt2p has been identified as the region of the protein mostly targeted for phosphorylation (Pagac et al. 2012). The predicted Gpt2p topology (Pagac et al. 2012) proposes a cytosolic orientation of this serine-rich C-terminal tail (Figure 4.1) allowing it to be exposed to cytosolic kinases and phosphatases.

Figure 4.1- Predicted topology of Gpt2p. Gpt2p has six predicted transmembrane segments, with the current predicted topology positioning the N- and C- termini facing the cytosol. The serine-rich C-terminus is represented by the orange diamonds. Three of the four catalytic motifs (green boxes) are predicted to face inwards towards the ER lumen, while motif IV is predicted to be on the cytosolic side. Figure made by B. Shabits based on data from (Pagac et al. 2012).

In this thesis the role of the serine-rich region in the regulation of Gpt2p was investigated. The first step in these studies involved the identification of phosphorylation sites in Gpt2p by mass spectrometry analysis. Identified phosphorylation sites were then targeted for site directed mutagenesis to convert serine and threonine residues to alanine or aspartic acid to mimic dephosphorylated or phosphorylated states, respectively. A large toolbox of plasmids and strains expressing phospho-mutant variants were developed in collaboration with Dr K. Athenstaedt (Graz University). In this chapter the identification of phospho-residues by mass spectrometry, the modification of these residues by site directed mutagenesis and the characterization of all phospho-mutant proteins is presented.

4.3 Results & Discussion

4.3.1 Identification of phosphorylation sites in Gpt2p

Our previous studies revealed that distinct GPATs conformations can be resolved by SDS-PAGE as bands migrating at different rates, and we have previously shown that phosphatase treatment collapses all of them to one faster migrating band that closer matches the expected molecular weight of 83.7 kDa (plus tag) in the case of Gpt2p (Figure

4.2) (Bratschi et al., 2009; Marr et al., 2012).

Figure 4.2- Characteristic band pattern distinguishing phosphorylation species of Sct1p and Gpt2p. Red and blue asterisks identify P2 and P1 states, respectively. Green asterisk identifies the P0 state, the resulting band after phosphatase treatment. Sct1p-V5 and Gpt2p-V5 were overexpressed from a GAL inducible plasmid (pYES) and were visualized by western blot using an anti-V5 primary antibody. Modified from (Bratschi et al., 2009).

This suggests that a phosphorylated Gpt2p binds SDS differently, inducing anomalous membrane protein PAGE migration behavior, which has been proposed to be reflective of differences in protein-protein or protein-lipid interactions (Rath et al. 2009).

This effect of phosphorylation on Gpt2p provides a practical advantage, as SDS-PAGE analysis combined with mutagenesis can be used to identify residues with critical regulatory roles. In cells overexpressing Gpt2p the protein displays at least two major bands that we have denominated P1 and P2 (Figure 4.2).

The P2-state dominates when Gpt2p is in excess (over-expressed), while P1 has been detected when analyzing endogenous levels of the protein either in the absence of

Sct1p or in the presence of oleic acid (C18:1 Δ9) as the only carbon source (Bratschi et al.,

2009; Marr et al., 2012; Pagac et al., 2012).

In agreement with these antecedents, a differential enrichment of P2 vs P1 conformations were observed in conditions of low vs high demand for Gpt2p activity respectively. In cells carrying GPT2 in a high copy vector (pYES) under the GAL1 promoter, gene expression is induced in the presence of galactose and is repressed upon shift of cultures to glucose. This plasmid was introduced in wild type cells or in the conditional gpt2Δ sct1Δ [pGAL-GPT2] double knock out, where the shift to glucose induces cell death (Zaremberg and McMaster 2002). In this background, where Gpt2p activity is in high demand due to lack of endogenous GPATs, the dominant Gpt2p species during repression corresponded to the P1-state (Figure 4.3). In great contrast, when the plasmid was introduced in wild type cells (which have endogenous GPATs) the P2-state dominated at all times after the shift to glucose (Figure 4.3). Altogether these results and previous antecedents point to a correlation between the P2-state of Gpt2p with low cellular demand of GPAT activity. Therefore, we predicted that lack of phosphorylation in the residues related to this P2-conformation should result in increased activity of Gpt2p. The goal was then to first identify these phosphoresidues

Figure 4.3- Differential phosphorylation pattern in conditions of high vs low GPAT demand- A) Cartoon depicting the conditional lethal gpt2Δ sct1Δ double knock out strain carrying the GPT2 gene under the GAL promoter. Due to the synthetic lethality of GPT2 and SCT1 genes this strain is alive in galactose (inducer) but loses viability upon shift to medium containing glucose (repressor). B-C) Galactose to glucose shift followed by western blot detecting Gpt2p tagged with the V5 epitope. B) gpt2Δ sct1Δ double knock out cells (Bratschi et al., 2009). C) wild type transformant (blot made by J. Kearley).

To identify phosphoresidues responsible for the mobility of the P2-state in Gpt2p, three independent phospho-site mapping analyses of Gpt2p were performed by mass spectrometry using a Gpt2 with a V5-6xHis tag fused to its C-terminus. It is worth noting that C-end fusions of Gpt2 with V5-6xHis, GFP or TAP tags have been previously shown

94 to yield functional proteins (Bratschi et al., 2009; Marr et al., 2012). The source of protein for these investigations were lysates obtained from yeast lacking endogenous Gpt2p (gpt2Δ mutant) over-expressing Gpt2p-V5-6xHis from the high copy vector (pYES) under the

GAL1 promoter. In the first two rounds of MS phospho-analysis, Gpt2p was enriched by immunoaffinity purification using anti-V5 antibodies crosslinked to agarose beads while

Ni-NTA isolation was used in the last round. The protein was then separated by SDS-

PAGE, the band corresponding to Gpt2p was cut from the gel followed by trypsin digestion and LC-MS/MS analysis (see Materials & Methods for details). Several putative phosphoresidues located in the serine-rich C-terminal portion of Gpt2p were identified, with serine 668 being the residue consistently phosphorylated in all three independent rounds (Table 4.1).

Table 4.1- Results from MS analysis.

First round (anti-V5): (m/z) mass (calc) Sequence 661.47 1245.51 SNALpSRVNpSRG S664/S668

Second round (anti-V5): (m/z) mass (calc) Sequence 722.59 1442.669 GpSLTDIPIFSDAK S671 834.55 1666.636 DGYDVSpSDAESSISR S632 884.08 1765.835 SSSIHSIGSLASNALSR (either ser 1,2 or 3) S649/650/651 912.63 1822.737 RDGYDVSpSDAESSISR S632 924.21 1845.802 SSSIHSIGSLASNALSR (+ 2 Phospho S) S649/650/651 950.61 1898.913 VNpSRGSLTDIPIFSDAK S668 990.60 1978.880 VNpSRGpSLTDIPIFSDAK S668/ 671

Third round (Ni-NTA): (m/z) mass (calc) Sequence 660.64 1978.890 VNpSRGpSLTDIPIFSDAK S668/ 671 834.32 1666.630 DGYDVSpSDAESSISR S632 926.05 2775.130 SEGETSpEDEDEFDEK S693

Additional phosphoresidues at positions S664 or S671 were also present in some of the peptides containing phospho-S668 (Table 4.1). These three residues are part of a conserved motif localized to the carboxy-end of the protein, which is not only present in

Gpt2p, but also in its counterpart GPAT Sct1p, as well as in other evolutionary related

GPATs from organisms of the SACK group, including S. cerevisiae, Ashbya. gossypii,

Candida glabrata, and Kluyveromyces. lactis (Smart et al. 2014)( Figure 4.4).

Figure 4.4- Sequence alignments of serine-rich C-terminal of Gpt2p related GPATs S. cerevisiae (Scer), Ashbya gossypii (Agos), Candida glabrata (Cgla), Kluyveromyces lactis (Klac). Highlighted residues in red show conserved Ser residues among these species.

4.3.2 Site directed mutagenesis of identified phosphorylation sites in Gpt2p

Site directed mutagenesis of S664, S668 and S671 individual residues to alanine to mimic dephosphorylation were introduced in the GPT2 gene cloned in the GAL inducible pYES vector. Overexpression of the single mutants was induced by growing transformants in medium containing 2% galactose and cell lysates were obtained. Separation of the proteins in SDS-PAGE unveiled alterations in Gpt2p mobility, with S668A registering the most dramatic changes. Yet, in all cases phosphatase treatment collapsed all proteins to a faster migrating band (P0-state) (Figure 4.5).

Figure 4.5- Characteristic band pattern of overexpressed Gpt2p phosphorylation mutants. Lysates from cells overexpressing wild type or the indicated Gpt2p-V5 mutants (from a pYES vector) grown to mid-exponential phase in selective medium containing 2% galactose were visualized by western blot using an anti-V5 primary antibody. Phosphorylation pattern of proteins before phosphatase treatment (-) display bands associated with P2 or P1 states, but shift to the lowest band position (P0) after phosphatase treatment (+).

With the goal of mimicking a fully dephosphorylated P0-state or a fully phosphorylated P2-state of Gpt2p, combined triple mutations changing S664, S668 and

S671 to alanine (Gpt2-3A) or aspartate (Gpt2-3D) were generated. Interestingly, while no major changes in protein mobility were observed in the protein carrying the S to D mutations, Gpt2-3A displayed only one band that migrated near the P1-state position

(Figure 4.6). Phosphatase treatment of Gpt2-3A further changed its mobility to P0, indicating Gpt2-3A is still phosphorylated at another residue/s.

Figure 4.6- Characteristic band pattern of overexpressed Gpt2p triple phosphomutants. Lysates from cells overexpressing wild type or the indicated Gpt2p-V5 mutants (from a pYES vector) grown to mid-exponential phase in selective medium containing 2% galactose were visualized by western blot using an anti-V5 primary antibody. Phosphorylation pattern of proteins before phosphatase treatment (-) display bands associated with P2 or P1 states, but shift to the lowest band position (P0) after phosphatase treatment (+).

With the goal of producing a mutant mimicking a fully dephosphorylated P0 conformation, we aimed to introduce more Ser to Ala mutations in other phosphorylation sites identified by our MS analysis, including S632, S651, and S693 (Table 4.1). Ser to Ala mutations in each of these residues were introduced in plasmids already carrying Gpt2-3A, generating quadruple, quintuple and sextuple mutants (Figure 4.7). While subtle shifts were observed in quadruple and quintuple mutants, the sextuple mutant showed the strongest change to a faster migrating conformation, suggesting all these residues contribute to the conformation displayed by Gpt2-3A. Interestingly, the S651A mutation alone exhibited a band shift from P2 to the P1 conformation as seen for Gpt2-3A, in contrast to the introduction of a single S693A mutation which run like a wild type.

Figure 4.7- Characteristic band pattern of overexpressed Gpt2p carrying multiple mutations. Lysates from cells overexpressing wild type or the indicated Gpt2p-V5 mutants (from a pYES vector) grown to mid-exponential phase in selective medium containing 2% galactose were used to visualize Gpt2p by western blot using an anti-V5 primary antibody. Each lane was loaded with 30 µg of total protein. Loading control was stained with TCE.

During the preparation of these mutants new global studies identifying phosphoresidues in Gpt2p in response to different conditions were published as well as new entries in PhosphoGRID were registered. A complete list of all identified phosphoresidues to date, including our own MS analysis, is presented in Table 4.2.

Consistent with our findings S668 is the phosphoresidue that has been most frequently reported, followed by S671, S693 and S632 (Table 4.2). To further investigate the function and importance of the serine-rich C-terminal tail in the regulation of Gpt2p, a truncated

Gpt2p (Gpt2-Trunc) was also designed (K. Athenstaedt) missing the serine-rich C-terminal end (last 133 residues) containing 18 of the 24 residues identified as phosphorylated sites

(Figures 4.7 and 4.8).

Figure 4.8- Map of phosphorylation sites in the amino acid sequence of Gpt2p. Highlighted in red are the phosphorylation sites identified in this study and others (Table 4.2). Bold S indicates serine residues most frequently found to be phosphorylated. The four catalytic motifs are highlighted in cyan. V610 (purple) represents the last amino acid in the truncated version of Gpt2p and the 133 amino acids missing from the C-terminal end are highlighted in yellow.

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Table 4.2: Phosphorylated amino acids identified in Gpt2p. The number of times the residue has been identified in our MS analysis as well as in the literature or SGD is indicated in the # reports column. Kinase predictions are based on those detected in PhosphoGRID or global phosphoproteome studies. The phospho-sites characterized by mutagenesis in this thesis are indicated (Mutagenesis).

Gpt2 Amino Acid # reports Refere Predicted Kinase PhosphoGRID This study Mutagenesis S2 1 nces3 S27 1 9 S211 2 10 CDK1 ✓ Y315 1 3 T352 1 3 S623 1 3 S631 4 10–12 CK2 ✓ 3,10,12– S632 11 17 GSK3, PKA, CDK1, CK2 ✓ 3 ✓ S636 2 12 CDK1, GSK3 ✓ S637 4 12,13,15 CDK1 ✓ S647 2 13 CDK1, GSK3 ✓ S649 1 CDK1, GSK3 ✓

S651 6 9,12,13,16 YPK1, GSK3, CDK1, PKA ✓ 1 ✓

S654 6 12,13,16,17 PKA, CDK1 ✓ 1 S657 4 12,13,16 PKA, CDK1 ✓ S660 1 GSK3, CDK1 ✓ S664 3 13 GSK3, PKA, CDK1 ✓ 1 ✓ 3,10– S668 15 14,16–20 PAK, GSK3, PKA, PKC, CDK1, PTK1, HAL5, MKK2 ✓ 3 ✓ 3,10–14,16– S671 14 20 PAK, CDK1, PKA, PKC, cAPK, PTK1, HAL5, MKK2 ✓ 2 ✓ 3,10,12,13,16,2 T673 7 0 PKA, CDK1 ✓

S688 6 10,11,13,15,16 CK2, PKA , CDK1 ✓ 10– T692 9 14,16,18,20 CK2, PKA, CDK1 , YPK1, SNF1 ✓ 3,10,12– S693 13 16,18,20,21 CDK1, SNF1, CK2 , NKK1, SKS1 ✓ 2 ✓ S712 1 20 (3) (Pagac et al. 2012) Topology (9) (Muir et al. 2014) Ypk1 (10) (Albuquerque et al. 2008) DNA damage (11) (Helbig et al. 2010) S2 is acetylated by NatB complex, in nat3Δ background: PAK, CK2, PKA target p- sites (12) (Swaney et al. 2013) Phosphorylation/non- Ubiquitination (13) (Holt et al. 2010) Cdk1 (14) (Kanshin et al. 2015) Heat and cold stress (PKA/Cdc28) (15) (Smolka et al. 2007) DNA damage (16) (Soulard, A.; Oppliger 2010) TORC1->PKA (17) (Li et al. 2007) alpha factor arrested: PKA, PKC, CK2 (18) (Bodenmiller et al. 2010) Snf1, Ptk1, Hal5, Mkk2, Nkk1, Sks1, Ypk1 (19) (Huber et al. 2009) (20) (Saleem et al. 2010) Oleate, peroxisome (21) (Gruhler et al. 2005) Pheromone signaling

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4.3.3 Phenotypic characterization of Gpt2-phosphomutants

During characterization of the SDS-PAGE mobility of Gpt2-phosphomutants, we noticed that growth of transformants in galactose-containing medium to induce the overexpression of these proteins was unusually slow. Indeed, independent gpt2Δ transformants carrying wild type GPT2, GPT2-3A or GPT2-trunc under the GAL promoter in a pYES plasmid showed slower growth on medium containing galactose (inducer) compared to cells carrying the empty vector (Figure 4.9). While cells overexpressing wild type Gpt2p eventually reached the same maximum growth as control cells carrying the empty vector, those overproducing Gpt2-3A or Gpt2-trunc were constantly behind. These results indicate that overexpression of unregulated Gpt2p reduces fitness of yeast.

Figure 4.9- Overexpression of Gpt2-phosphomutants reduces yeast fitness- (A) Growth curves of gpt2Δ transformants overexpressing the indicated phosphorylation mutants from a pYES plasmid grown in defined medium containing 2% galactose (B) serial dilutions on plates containing 2% glucose (no induction) vs 2% galactose (inducer) grown for 3 days at 30℃.

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Inspection of cells from these cultures under the microscope suggested transformants carrying GPT2-3A or GPT2-trunc accumulated larger lipid droplets (LDs) during exponential phase. In order to look at this closer, cells were stained with BODIPY

493/503 to visualize neutral lipids stored in LDs. As shown in Figure 5.10 overexpression of unregulated Gpt2-3A or Gpt2-trunc indeed induced the accumulation of large LDs in exponential growth, while levels of proteins were in the same range. In addition these cells looked more elongated, as previously seen in the conditional lethal gpt2Δ sct1Δ [pGAL-

GPT2] double knock out (Bratschi et al., 2009).

Figure 4.10- Overexpression of unregulated Gpt2p variants increases LD size in wt-V5 trunc-V5 exponential phase. A) Empty vector or plasmids carrying GPT2 , GPT2 and 3xA-V5 GPT2 under the GAL1 inducible promoter were introduced into gpt2Δ cells. Log phase cells grown on selective defined medium containing 2% galactose were stained with Bodipy-493/503 to visualize lipid droplets (LDs). B) Microsomal fractions from cells in A were prepared. Proteins (30 ug per lane) were separated in a 10% SDS-PAGE and Gpt2p- V5 variants were visualized by Western blot using anti-V5 primary antibody.

To study the role of Gpt2p phosphorylation at these serine positions under physiological conditions we introduced the mutations in the GPT2 locus of BY4741 (wt) cells. New strains carrying each single and triple phosphorylation deficient variant in which

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S664, S668 and S671 were replaced by alanine or aspartate were obtained. Endogenously expressed Gpt2p phosphomutants (V5-taged) showed similar migration patterns in SDS-

PAGE as those observed in conditions of overexpression, with S668A displaying the strongest change among single mutants and Gpt2-3A resulting in a complete shift from P2 to the P1 conformation (Figure 4.11).

Figure 4.11- Migration of endogenously expressed Gpt2p phospho-deficient mutants. Cells expressing endogenous levels of wild type Gpt2p-V5 or indicated phospho-mutants were grown to mid-exponential phase in defined medium containing 2% glucose. Microsomal fractions were prepared and 20 µg of total protein were loaded into each lane of an SDS-PAGE gel. Gpt2 was visualized by western blot using an anti-V5 primary antibody. LC, loading control detected with TCE staining.

Altogether these results suggest that phosphorylation of S668 may regulate phosphorylation of the other serine residues, in a cascade that plays a role in switching between P2/P1 states.

As seen previously for the overexpressed protein (Figure 4.6) phosphatase treatment of Gpt2-3A expressed at endogenous levels also caused a shift to the P0 conformation indicating the presence of other phosphorylated sites even when expressed

104 at physiological levels (Figure 4.12). Unfortunately, the attempt to mimic a triple phosphorylation state in Gpt2-3D was not effective as the protein was not locked in the P1- state upon phosphatase treatment and ended behaving like wild type Gpt2p (Figure 4.12)

Figure 4.12- Characteristic band pattern of Gpt2p carrying multiple mutations expressed at endogenous levels. Endogenous Gpt2p-V5 in prepared microsomal fractionation samples of cells grown to mid-exponential phase in defined medium (SD) were visualized by western blot using anti-V5 antibody. Phosphorylation pattern of all proteins showed a shift to the P0 position after phosphatase treatment. 26 µg of proteins were loaded. Loading control is SDS-PAGE gel visualized with TCE and UV light before transfer to membrane.

Introduction of the S651A mutation at the endogenous GPT2 locus produced a protein that displayed a migration pattern comparable to that shown by Gpt2-3A and responded similarly to phosphatase treatment (Figure 4.12). Therefore, lack of phosphorylation at this single S651 residue changes the protein conformation and might play a role in regulating Gpt2p. In fact, S651 has been identified as part of a phospho- acceptor motif (646RSRSSSI652) for Ypk1p, a kinase involved in sphingolipid regulation

(Muir et al. 2015), although the involvement of this kinase in the regulation of Gpt2p has not been conclusively demonstrated.

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Given the phenotypes displayed by overexpression Gpt2-3A and Gpt2-trunc pointing to changes in glycerolipid metabolism, these unregulated forms of Gpt2p became the focus of our studies (Chapters 5 & 6). Importantly, cells bearing Gpt2-3A or Gpt2-

Trunc at physiological levels were able to support life of yeast in the absence of Sct1p, indicating these Gpt2-variants result in functional GPATs.

4.4 Concluding remarks

The yeast GPAT Gpt2p is a heavily phosphorylated protein, with at least 24 putative phosphosites reported in global phosphoproteomic studies and the PhosphoGRID database (Table 4.2). Our own MS analysis combined with site directed mutagenesis of identified phosphoresidues indicates that S664, S668, S671 and S651 are indeed phosphorylated. Mutations of these residues to Ala, to prevent phosphorylation, induce conformational changes that result in mobility shifts when the proteins are separated in

SDS-PAGE. The most drastic changes occur when S664A, S668A, S671A are combined

(3A mutant) or when S651A is introduced in Gpt2p. Unfortunately, introducing S to D mutations did not seem to mimic phosphorylation, so they were put on hold and were not incorporated in further studies.

Interestingly, overproduction of Gpt2-3A or a mutant lacking the entire serine-rich c-end tail of the protein result in reduced growth, accumulation of large lipid droplets in exponential phase and elongated cells. When these Gpt2-variants are expressed at physiological levels, migration of the proteins were similar to those shown by the overexpression system. Importantly, they yield functional GPATs as they are capable of

106 supporting life of yeast in the absence of the other GPAT, Sct1p. Characterization of all these strains is presented in Chapter 5.

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Chapter 5 – Metabolic impact of unregulated Gpt2p mutants

5.1 Introduction

Biochemical studies conducted in the 1990’s indicated that GPAT activity in yeast is distributed between microsomal fractions and lipid droplets (Athenstaedt et al., 1997;

Athenstaedt et al., 1999). The genes coding for the yeast GPATs, SCT1/GAT2 and

GPT2/GAT1, were identified in early 2000’s (Zheng et al., 2001; Zaremberg et al., 2002), which allowed to initiate investigations aimed at understanding their differential contributions to lipid metabolism. DAG consuming pathways, including TAG synthesis and the CDP-choline pathway for synthesis of phosphatidylcholine were differentially impacted by lack of Gpt2p or Sct1p (Zaremberg et al., 2002). In exponentially growing cells, perinuclear and cortical ER localizations were observed for both Gpt2p and Sct1p

(Bratschi et al., 2009). Although their sub-cellular localization mostly overlapped, further analysis of their distribution using equilibrium density gradients demonstrated a differential enrichment of Gpt2p and Sct1p in distinct ER fractions. These results suggest that distinct sub-compartments may exist in the ER, which are differentially enriched in

Gpt2p or Sct1p. Moreover, overexpression of Gpt2p and Sct1p in the absence of endogenous GPATs (i.e in a gpt2Δ sct1Δ double knock out) induced proliferation of distinct ER arrays, differentially affecting cortical ER morphology (Bratschi et al. 2009b).

Enhanced cortical localization of Sct1p was detected in the absence of Gpt2p, while no notable changes were observed in Gpt2p distribution in the absence of Sct1p. It seems that the ER distribution of Sct1p is sensitive to the presence of Gpt2p, suggesting that cellular

108 mechanisms act to adapt to a GPAT imbalance. Lack of Sct1p or Gpt2p induces a change in the phosphorylation pattern of the remaining GPAT in the cell (Bratschi et al. 2009).

Proteomic studies conducted in the last decade have consistently found Gpt2p (but not Sct1p) as part of the LD proteome (Grillitsch et al. 2011). Interestingly, association of

Gpt2p with purified LDs depends on the carbon source as it was detected in glucose but not in oleate growing cells (Grillitsch et al. 2011). Gpt2p is a transmembrane protein with hydrophilic luminal domains (Pagac et al. 2012) and therefore cannot be accommodated on the monolayer surface of an LD (Kory, Thiam, and Walther 2015) but could interact with LDs from its ER location. Indeed, we have previously observed ER-derived structures enriched in Gpt2p that were tightly associated with LDs (Figure 5.1). These ER-extensions were prominent when cells were incubated with oleate, which also induced a shift from P2 to P1-conformation in Gpt2p (Marr et al. 2012).

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Figure 5.1- Gpt2p enriched ER structures interact with lipid droplets. A) diploid cells expressing endogenous levels of Gpt2p-GFP (green) and Erg6-RFP (red) were grown in medium containing 0.1% oleic acid for 24 h and analyzed by fluorescence microscopy. DIC, differential interference contrast. B) deconvolved xyz projections of the cell depicted in A showing details of the area of interaction between the Gpt2p crescent and LD. Arrows point to the crescent structure in juxtaposition with LDs. C) three-dimensional rendering from the cell shown in A (left) and Gpt2p crescent enlargement captured from different angles to appreciate the juxtaposition of the Gpt2p crescent with LDs. For better orientation, an arrow and an asterisk were integrated into the picture. Scale bar is 1 μm. Figure obtained from Marr et al.

Therefore, changes in phosphorylation of the GPATs have been associated with a concomitant shift in their ER distributions and interactions with organelles like LDs, opening the possibility that phosphorylation/ dephosphorylation of these enzymes plays a role in their localization. Changes in cellular distribution of these proteins could be the result of specific protein-protein interactions and/or due to altered activity changing the lipid composition downstream of the GPAT. Both, changes in protein-protein interactions and activity could be regulated by the phosphorylation status of the protein.

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5.2 Goals

The main objective of this thesis was to understand how phosphorylation regulates

Gpt2p. Given the antecedents connecting changes in Gpt2p phosphorylation to LD association and TAG metabolism, the following specific goals guided the investigations described in this chapter:

to determine if constitutive dephosphorylation of Gpt2-3A

• has an impact on Gpt2p cellular distribution and association with LDs

• induces changes in neutral lipid metabolism and LD metabolic cycles

• results in changes in GPAT activity

5.3 Results & Discussion

5.3.1. Characterization of Gpt2 phospho-mutants expressed at endogenous levels

In order to evaluate the role of phosphorylation as closer to physiological conditions as possible we decided to introduce the S664A, S668A, S671A (as single mutations and all-combined in 3A) as well as 3D and the truncation (Δ133) in the GPT2 locus. As part of the toolbox of strains produced, wild type and mutant variants were fused to either a V5- x6His or GFP tags to aid in live microscopy visualization of Gpt2p (see Materials &

Methods). All mutant strains were confirmed to carry the desired mutations by sequencing.

In contrast to the overexpression system, no significant differences in growth among these strains were observed in defined medium containing 2% glucose (not shown).

Since detection of Gpt2p and its variants proved to be challenging when dealing with whole cell lysates we decided to prepare microsomal fractions to enrich in membranes containing Gpt2p, which was then detected by western blot avoiding problems related to protein degradation (Figure 5.2). We next decided to visualize Gpt2p in the new strains in

111 order to confirm they were indeed expressed and that the differences in phosphorylation status were still maintained. Gpt2p was detected by western blot (Figure 5.2). The Gpt2p phosphorylation patterns displayed by the proteins expressed at endogenous levels were in agreement with those previously observed with the overexpression system (Figure 4.7).

Furthermore, we noticed that the levels of 3A were elevated compared to the wild type sample.

Figure 5.2- Endogenous levels of Gpt2p phosphomutants. Cells expressing the indicated V5-fused proteins at physiological levels were grown to mid-exponential phase (6 hours) in defined media containing 2% glucose. Microsomes were prepared and an aliquot was treated with λ-phosphatase as described in Materials and Methods. Twenty-six (26) ug of total protein from control (-) or treated (+) samples were separated by SDS-PAGE and processed for western blot analysis using an anti-V5 primary antibody. Gpt2-V5 expected MW is ~87.4 kDa. Densitometry of Gpt2p bands treated with phosphatase was normalized according to the loading controls for each sample and expressed relative to that of wild type Gpt2p. Representative experiment out of 3 independent ones performed throughout these studies.

5.3.2 Unregulated Gpt2p- localization and impact on protein levels

Using fluorescence live microscopy, we next examined the subcellular localization of Gpt2-GFP, Gpt2-3A-GFP and Gpt2-Trunc-GFP expressed at endogenous levels. Cells grown in defined medium containing 2% glucose in both exponential (6 hrs) and stationary

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(24 hrs) phases of growth were imaged. Sct1-GFP was also included in these experiments for comparison. As shown in Figure 5.3, all actively growing (log phase) strains displayed peri-nuclear and cortical-ER localization of Gpt2p. We noticed that cells carrying the Gpt2-

3A variant displayed small LDs in log phase usually associated with Gpt2-ER enriched areas and extensions. No such structures were detected in the truncated or wild-type forms of Gpt2p or Sct1p. In addition, a hazy GFP signal localized to vacuoles was seen in cells expressing wild type Gpt2p in log phase. This signal increased in stationary phase and it was then also observed for Gpt2-3A. Remarkably, truncated Gpt2p signal was equally strong in both phases of growth and no vacuolar signal was detected. Accumulation of soluble green fluorescence in the vacuolar lumen reflects protein degradation due to membrane internalization (Zutphen et al., 2014). As GFP is more resistant to the acidic pH of the vacuole than the majority of proteins, free GFP in the vacuole can be used as a reporter of protein degradation. This implies that wild type Gpt2p was more prone to vacuolar degradation than the phosphomutant variants and its counterpart GPAT Sct1p.

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Figure 5.3. Live imaging of Gpt2p phospho-mutants during exponential and stationary phases of growth. Cells expressing the indicated GFP-fused proteins at physiological levels were grown to mid-exponential (6 hours) and stationary phases (24 hours) in defined media containing 2% glucose. Cells were analyzed by epifluorescence microscopy using a GFP filter and differential interference contrast (DIC). Images were captured using the same exposure time settings for all strains. Peri-nuclear ER (red arrow), cortical ER (blue arrow), ER extension (pink arrow), vacuole (depressed area labeled with V in DIC), lipid droplets (protruded dots pointed by yellow arrow in DIC). Scale bar is 5 µm. Representative images from one independent experiment out of 5 performed throughout these studies.

We further confirmed these observations by visualizing the proteins by western blot

(Figure 5.4). It is worth noting that in order to detect free GFP, which is a soluble protein, we had to process whole cell lysates. With the intention of minimizing post-lysate degradation of the GPATs, a new lysis buffer containing sucrose was used to stabilize the

114 protein (A. Lopez-Villalobos), (Lee et al., 1981). This allowed immediate boiling of samples after lysis preventing degradation as well as smearing of the protein during migration in SDS-PAGE (see Materials and Methods for details). In agreement with the microscopy images captured during log phase, wild type Gpt2p was the only sample displaying free GFP signal. This opened the possibility that lack of phosphorylation in

Gpt2-3A and truncated variants prevents protein degradation during exponential growth.

Figure 5.4. Protein analysis of GFP-tagged proteins in exponential phase. Cells from figure 5.3 expressing the indicated GPATs at endogenous levels and grown to log phase were lysed. Forty ug of total protein from whole cell homogenates were separated by SDS- PAGE and processed for western blot analysis using an anti-GFP primary antibody. Expected molecular weights: Gpt2-GFP ⁓110 kDa, Gpt2-trunc-GFP, ⁓90 kDa, Sct1-GFP ⁓112 kDa. and free GFP ⁓27 kDa. Representative image from one independent experiment out of 4 performed throughout these studies.

Previous work has shown that Gpt2p shifts to a P1-state when Sct1p is absent

(Bratschi et al., 2009). We therefore reasoned that since Gpt2-3A runs closer to P1 then a wild type Gpt2p in a sct1Δ background may show less degradation. In order to investigate

115 if decline of wild type Gpt2-GFP as well as phosphomutants was dependent on the presence of Sct1p, strains deleted for SCT1 but expressing all these Gpt2p variants at endogenous levels were produced. Protein analysis of these strains is presented in Figure 5.5. No significant differences in the appearance of free GFP was observed among all strains when

Sct1p was missing in exponential phase while the levels of degradation were alleviated in stationary phase. Consistent with the microscopy images (Figure 5.3) Gpt2-3A-GFP degradation increased during stationary phase, while no free GFP was detected for Gpt2- trunc. According to these results, it seems that Gpt2p lacking phosphorylation on S664,

S668, and S671 residues exhibits a different behavior than wild type during log phase but follows the same path to the vacuole during stationary phase, highlighting the importance of these three phosphorylation sites in the regulation of Gpt2p depending on the phase of growth. Constitutive dephosphorylated state in Gpt2-3A partially prevents vacuolar degradation of the protein during log phase. Moreover, our results showed that missing the serine-rich c-terminal end in Gpt2-trunc seems to have an effect on both localization and stability of Gpt2p independent of the phase of growth. Altogether it can be concluded that while S664, S668, and S671 residues regulate certain aspects of Gpt2p, lack of the entire serine-rich region has additional consequences, and presumably prevents Gpt2p from degradation in the vacuole.

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Figure 5.5- Lack of Sct1p alleviates degradation of Gpt2 in stationary phase- Cells expressing the indicated GFP-fused proteins at physiological levels were grown to mid- exponential (6 hours) and stationary phases (24 hours) in defined media containing 2% glucose. A) 25 ug of total protein from whole cell homogenates were separated by SDS- PAGE and processed for western blot analysis using an anti-GFP primary antibody. Expected molecular weights: Gpt2-GFP ⁓110 kDa, Gpt2-trunc-GFP, ⁓90 kDa, and free GFP ⁓ 27 kDa. B) Densitometry of the free GFP band was normalized to loading control

117 for each sample. The graph bar shows abundance of free GFP relative to that of wild type Gpt2p in exponential phase. Representative experiment out of 3 independent ones performed throughout these studies.

5.3.3. Unregulated Gpt2p- Impact on the timing of lipid droplet accumulation and morphology

During the in vivo localization studies with cells grown on 2% glucose we noticed that compared to wild type, cells expressing Gpt2-3A at endogenous levels exhibited large size LDs already during log-phase, which is not the typical phase where yeast accumulates neutral lipids (yellow arrow in DIC image Figure 5.3). In addition, microscopy inspection of wild type, single and 3A mutants grown to late log phase displayed enrichment of Gpt2p in specific ER areas, forming bright dots/tubules. These structures were more frequently seen in the mutants (Figure 5.6), resembling the behaviour of Gpt2p in response to oleate, when TAG synthesis and LD biogenesis is induced (Marr et al. 2012). In the early response to oleate as carbon source, Gpt2p-enriched domains were observed associated with incipient LDs (Marr et al. 2012).

Figure 5.6- Gpt2p is enriched in ER-domains in late exponential growth. Cells expressing the indicated GFP-fused proteins at physiological levels were grown to late- exponential phase (16 hours) in rich medium containing 2% glucose. Samples were imaged using epifluorescence microscopy. Overlay images obtained with the GFP filter and differential interference contrast were processed using ImageJ. Red arrows point to areas where Gpt2p accumulates. Scale bar is 5 µm.

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Since TAG synthesis increases when glucose levels decline, and reaches its maximum when cells enter stationary phase, these initial results suggested that the timing of LD accumulation was altered in these Gpt2p phosphomutants. To try to understand how alterations in Gpt2p phosphorylation may affect the formation of Gpt2p rich domains, TAG synthesis and LD dynamics, several approaches combining cell biology, biochemistry and genetic techniques were used. As a first step, cells expressing wild type Gpt2p or phosphomutants fused to GFP as well as the LD marker protein Erg6p tagged with RFP were generated. Co-localization studies revealed that GPAT-enriched domains in the ER were frequently found in close proximity to LDs (Figure 5.7, 5.8). In fact, subcellular fractionation experiments showed an enrichment of Gpt2-3A in the LD fraction compared to wild type (Figure 5.9).

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Figure 5-7. Co-localization analysis of Gpt2p and lipid droplets in log and stationary phases of growth. Cells expressing endogenous levels of the indicated GPATs fused to GFP, and Erg6-RFP (LD marker) were grown to mid-exponential phase (6 hours) (shown in panel A) and stationary phase (24 hours) (shown in panel B) in defined media containing 2% glucose. Cells were analyzed by epifluorescence microscopy using GFP and RFP filters and differential interference contrast (DIC). Images were captured using the same exposure time settings for all strains. Overlay image shows a strong association of ER extension in green with and large LDs in red in GPT23xA strain (pointed with arrows). Representative experiment out of 4 independent experiments. Scale bar is 5 µm.

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Figure 5.8- Gpt2-3A domains are intimately associated with lipid droplets. Cells expressing endogenous levels of Gpt2p-GFP (green) and Erg6-RFP (red) were grown to mid-exponential phase in SD medium and analyzed by spinning disk confocal microscopy. DIC, differential interference contrast. Three-dimensional rendering from the cell showing details of the area of interaction between the Gpt2p crescent and LD. Arrows point to the crescent structure in juxtaposition with LDs.

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Figure 5.9- Lack of phosphorylation induces Gpt2p association with lipid droplets. Cells expressing wild type Gpt2p or Gpt2-3A (endogenous levels) were grown to mid- exponential phase in defined media containing 2% glucose. Cells were collected, and lysates were subjected to subcellular fractionation (see Materials and Methods). A) Western blot analysis for microsomal fractionation (MIC) and lipid droplet isolation (LD) using anti-V5 primary antibody. B) Distribution (% of total) of Gpt2-wt and Gpt2- 3xA in microsomes and LDs after normalization of V5 bands to their respective loading controls [figure in (A) contributed by B Nagler and K Athenstaedt].

Noteworthy, cells expressing Gpt2-3A exhibited larger LDs in log phase, while differences mostly disappeared during stationary phase. These results were also observed when cells were stained with Nile Red or BODIPY 493/503 (Figure 5.10) which stain the neutral lipids stored in LDs. Quantification of LDs from these experiments indicated that during exponential phase of growth cells expressing Gpt2-3A contain larger LDs but the

123 total number of LDs does not change significantly (Figure 5.11). LDs from cells growing in exponential phase were sorted in three categories according to their diameters: small (<

0.5 µm), large (> 0.5 µm) and oversized (> 0.7 µm). While close to 75% and 70% of wild type and Gpt2-trunc cells, respectively, displayed small LDs, around 55% of Gpt2-3A cells contained LDs larger than 0.5 μm in diameter. Furthermore, 15% of Gpt2-3A cells had oversized LDs (>0.7 μm) compared to 6% and just 4% for Gpt2-trunc and wild type, respectively (Figure 5.12). These results emphasize the differences between having a

Gpt2p with constitutive dephosphorylation of S664, S668, and S671 residues, and complete lack of the serine-rich region, which displays a milder phenotype than that observed for Gpt2-3A, suggesting that other parts of the C-end combined with the 3A mutation are necessary for the enlargement of LDs.

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Figure 5.10- Gpt2-3A cells have larger LDs in exponential phase of growth. Cells with no fluorescent tag were grown to mid-exponential phase in SD media and lipid droplets (LDs) were strained with A) 2 µg/µL Nile red or B) 1 µg/µL BODIPY 493/503. Cells were visualized with epifluorescence microscopy. Representative experiment out of 3 independent experiments. Scale bar is 5 µm.

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Figure 5.11- Analysis of lipid droplet population in Gpt2p phosphomutants. Quantification analysis of LDs stained with BODIPY 493/503 in cells grown to mid- exponential phase in defined media containing 2% glucose. Box plot showing the distribution of number of large LDs (>0.5 µm diameter) per cell (A) or total number of LDs per cell (B). The box plot shows the medians (central horizontal line) with the 25th and 75th percentiles (box). The bottom line coming out of the box ends at the 5th percentile, whereas the top one ends at the 95th percentile. Asterisks *** denote significantly (P<0.001) different distribution than that of the wild type (wt).

Figure 5.12- Size distribution of lipid droplets in Gpt2p phosphomutants. Distribution of lipid droplets (LDs) based on their size from cells stained with Nile Red or BODIPY 493/503 (as shown in figure 5.10). LDs were classified in three categories according to the size of their diameter: oversized >0.7 µm, large >0.5 µm, and small <0.5 µm. Bars represent average ± SE from at least 180 cells counted for each strain. Asterisks denote significant differences compared to the wild type strain with *** P<0.001.

The contribution of Sct1p to these phenotypes was also assessed by removing SCT1 from the genome in all Gpt2-GFP strains containing Erg6-RFP. Log-phase cells expressing

Gpt2-3A grown in the presence of 2% glucose displayed a similar large population of LDs in the absence of Sct1p (Figure 5.13). No changes were observed in wild type and Gpt2-

126 trunc either. On the contrary, a marginally significative increase in LD size was registered in all strains in the absence of Sct1p. These results suggest that Sct1p does not directly contribute to the increase size of LDs when Gpt2p is in the constitutively dephosphorylated state represented by Gpt2-3A. We also noticed that in the absence of Sct1p, localization of the Gpt2-GFP signal in the cortical ER was enhanced for all strains studied, but this was not quantified.

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Figure 5.13- Analysis of lipid droplet morphology in the absence of Sct1p. Cells expressing endogenous levels of the indicated GPATs fused to GFP, and Erg6-RFP (LD marker) in a wild type or sct1Δ backgrounds were grown to mid-exponential phase (6 hours) in defined media containing 2% glucose. Cells were analyzed by spinning disk confocal microscopy. Images were captured using the same exposure time settings for all strains. Scale bar is 2 µm. (Bottom) Quantification analysis using ImageJ. LDs were classified in two groups: large LDs (>0.5 µm diameter size) and small LDs (<0.5 µm diameter size). Bars represent average ± SE, for at least 100 cells counted for each strain. Asterisks denote significant differences compared to the wild type (wt) strain * P<0.1, ** P<0.05, *** P<0.001.

5.3.4. Unregulated Gpt2p- Impact on enzyme activity and TAG metabolism

The increase in Nile Red staining signal associated with strains expressing Gpt2-

3A in log-phase was also quantified by an independent assay using a fluorescence scanner

(see Materials and Methods for details). Results from five independent experiments indicated an average 1.57 ± 0.27-fold increase in Nile Red staining for Gpt2-3A compared to the wild type strain grown to mid-log phase (Figure 5.14).

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Figure 5.14. Neutral lipid content in 3A-phosphomutant and wild type cells. Cells expressing wild type and 3A Gpt2p were grown to mid-exponential phase in defined media containing 2% glucose. Cells were stained with Nile Red and serially diluted in a 96-well plate. Signal was measured using a fluorescence scanner (532 nm excitation wavelength). Data were processed using ImageJ software. Data were normalized to the number of cells used and expressed as fold increase relative to wild type signal.

We reasoned that since Nile Red stains neutral lipids, and TAG is one of the products formed downstream of the GPAT-catalyzed reaction in the de novo glycerolipid biosynthetic pathway, the increase in Nile Red staining most probably reflects an increase in TAG rather than other neutral lipids like sterol-esters.

Elevated TAG levels may be the result of an increase in GPAT activity, so next we investigated how constitutive dephosphorylation of S664, 668 and 671 impacted GPAT activity. For these experiments oleoyl-CoA was used as it can be utilized by Gpt2p and is not a preferred substrate of Sct1p (Zheng et al., 2001). Cells expressing wild type and 3A

Gpt2p were grown for 8 hours to mid-exponential phase. Cell lysates were prepared in sucrose containing buffer and immediately used for activity assays (see Material and

Methods for details). An aliquot of the whole cell lysate was immediately boiled in

Laemmli buffer and used for protein determination and visualization of Gpt2p by western blot (Figure 5.15). Results from these experiments showed that Gpt2-3A samples displayed around a 2-fold increase in GPAT activity. Although this rise in activity consistently

130 coincided with higher levels of the mutant protein. Protein abundance assessed by densitometry in western blot images of cell lysates from 15 independent experiments showed an average of 1.43 ± 0.12 fold increase in Gpt2-3A abundance compared to wild type. These results are in agreement with those obtained with live imaging of Gpt2-GFP showing higher vacuolar degradation of the wild-type protein, while this was delayed in the case of Gpt2-3A during log-phase of growth (Figures 5.4, 5.5, 5.6). Altogether we interpreted this increase in GPAT activity as not only a partial reflection of protein levels induced by the dephosphorylated state but as an effect on the catalytic efficiency of the enzyme as well.

Figure 5.15- Constitutive dephosphorylation in Gpt2-3A impacts GPAT activity. Cells expressing wild type or 3A Gpt2p (V5 tagged) at physiological levels were grown to mid- exponential (8 hours) in defined media containing 2% glucose. Total GPAT activity was measured in cell lysates by determining the incorporation of radiolabelled [14C]-glycerol- 3-phosphate into phosphatidic acid as described in Material and Methods. Data obtained from three independent experiments were combined and expressed as average ± standard deviation. Protein samples used in activity assays were analyzed by western blot using anti- V5 antibody (a representative image from one experiment is shown). Abundance levels of proteins were measured by ImageJ software. Data were normalized to loading control for

131 each sample and expressed relative to the wildtype. A. Lopez Villalobos contribution to this figure is acknowledged.

Since in these experiments Sct1p was also present, we decided to uncouple the activity of Sct1p from Gpt2p by using strains where SCT1 was deleted. Lysates of the same strains used for live microscopy expressing native Gpt2p or Gpt2-3A fused to GFP were tested (Figure 5.16). Log-phase samples of Gpt2-3A-GFP displayed higher GPAT activity, consistent with the rise in activity previously registered with the V5-tagged version of

Gpt2-3A (Figure 5.15). Therefore, it can be concluded that the increase in GPAT activity is independent of the tag used. As previously published (Bratschi et al., 2009), lack of

Sct1p induced a mobility shift in wild type Gpt2p which correlated with higher GPAT activity (Figure 5.16). Interestingly, although no significant differences in GPAT activity between native Gpt2p and Gpt2-3A were registered in the absence of Sct1p, Gpt2-3A in sct1Δ background still displayed large LDs while LDs in sct1Δ cells seemed to slightly increase in number and size (Figure 5.13 and 5.16). To our surprise, lipid extractions and

TAG quantification from TLC plates from these cells showed no significant differences between sct1Δ expressing Gpt2-3A or the native protein (Figure 5.17). We interpreted this increase in TAG as a consequence of the shift in wild type Gpt2p phosphorylation in the absence of Sct1p, resulting also in higher GPAT activity. These results also implicated that the abnormal LD size in cells expressing Gpt2-3A is a unique trait associated with the phosphomutant. Furthermore a significant increase in DAG also accompanied cells expressing Gpt2-3A regardless of the presence of Sct1p (Figure 5.17). Remarkably, total

GPAT activity dropped in this GPT2-3A sct1Δ strain. Therefore, altogether these results indicate that Gpt2-3A is the main GPAT responsible for the rise in TAG levels observed

132 during exponential growth. However, the presence of large LDs cannot be entirely attributed to enhanced TAG synthesis due to higher GPAT activity in log phase.

Figure 5.16- Phosphorylation deficiency impacts TAG levels in the absence of Sct1p. Strains expressing native or Gpt2-3xA fused to GFP (endogenous levels) were grown to mid-exponential phase (8 hours). Cells were collected and lysed and same lysates were used to analyze Gpt2 by western blots (A) or to measure GPAT activity (C) using [14C]- glycerol 3-phosphate and oleoyl-CoA. (B) Cells from the same cultures were stained with Nile Red and large LDs quantified. Bars represent average ± SD of three independent experiments. A. Lopez Villalobos contribution to this figure is acknowledged.

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Figure 5.17. TAG and DAG levels in strains expressing endogenous levels of wild type or 3A Gpt2p in the presence or absence of Sct1p. Cells were grown to log (8.5 h) and stationary phase (24 h) in defined medium (2% glucose) and their total lipids were extracted and subjected to TLC analysis to measure A) TAG (8.5 and 24 hours samples) and B) DAG levels (24 hours samples) normalized by OD600 of cells. Bars represent average of fold change ± SD of DAG and TAG in 24 hours from two independent experiments. 8.5 hours data represent one replicate. Values are normalized relative to those of the wild type (8.5 hours in the case of TAG). A. Cardenas contribution to this figure is acknowledged.

We next evaluated if accumulation of TAG in Gpt2-3A was indeed dependent on

GPAT activity. For this purpose, we generated mutants expressing inactive Gpt2p. The mutation G261L was previously shown to yield a catalytically dead protein (Marr et al.

2012) and was therefore introduced into GPT2 and GPT2-3A strains expressing the proteins fused to V5 at endogenous levels. These catalytically dead versions were also cloned in the high copy plasmid pYES, yielding GPT2 under the control of the GAL promoter. Interestingly, no significant changes in the phosphorylation pattern of active or catalytically-dead proteins were observed (Figure 5.18) regardless of the levels of expression of the active vs catalytically dead Gpt2 native and 3A proteins. These results suggest that the phosphorylation status of Gpt2p is not directly regulated by its GPAT activity.

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Figure 5.18- Phosphorylation pattern of wild type and 3A Gpt2 catalytically-dead versions. A) Cells over-expressing wild type or the indicated Gpt2p mutants (V5 tagged) from a pYES plasmid were grown to mid-exponential (6 hours) in defined medium (2% galactose). Lysates were analyzed by Western blot. B) Cells expressing endogenous levels of indicated Gpt2p (V5 tagged) were grown to mid-exponential phase in defined medium (2% glucose). Lysates were subjected to subcellular fractionation and the microsomal fraction was analyzed by Western blot. In all cases a primary anti-V5 antibody antibody was used. LC, loading control was stained with TCE.

Since overexpression of Gpt2p phosphomutants impairs growth when induced in the presence of galactose we tested if this phenotype was also dependent on Gpt2p catalytic activity. Plasmids were transformed into yeast cells lacking endogenous Gpt2p (gpt2Δ) and were grown on selective medium containing 2% galactose to induce expression. Serial dilutions of each transformant were spotted on plates containing 2% galactose (induction) or 2% glucose (repression) and incubated for 3 days at 30℃. As expected, cells overexpressing Gpt2-3A and Gpt2-trunc displayed reduced growth, but this phenotype was prevented when Gpt2-3A was inactive, despite no difference in protein expression levels

(Figure 5.19). Therefore, excess unregulated GPAT activity results in growth defects. In an unexpected turn, TAG levels were 1.6 times higher in cells overexpressing a

135 catalytically dead Gpt2p, similar to the rise seen for overexpression of Gpt2-trunc. In agreement with these observations, no differences were observed between Gpt2-3A and its catalytically dead version as they both displayed higher TAG levels than control transformants (Figure 5.19). Interestingly, an important increase in DAG levels were observed in cells overexpressing phosphomutant Gpt2-3A or Gpt2-trunc, but this phenotype was indeed dependent on GPAT activity. Therefore, the rise in DAG levels correlate with the toxic effect seen when these phosphomutants are overproduced, independently of TAG levels. This also implicated an effect of excess Gpt2p protein (but not activity) in the induction of TAG accumulation.

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Figure 5.19- Accumulation of DAG (but not TAG) correlates with the toxic effect of unregulated Gpt2p overproduction. A) Cells overexpressing Gpt2 in a pYES vector were grown to mid-exponential phase in selective defined medium and were serial diluted onto plates containing 2% glucose (repressor) or 2% galactose (inducer). Plates were incubated for 4 days at 30 ºC. Same set of cells were grown to stationary phase (24h) and their total lipids were extracted and subjected to TLC analysis to measure B) TAG and C) DAG levels. Bars represent average fold change ± SD relative to wt of two independent experiments. Values were normalized by OD600 of cells. * P<0.1, ** P<0.05. A. Cardenas contribution to this figure is acknowledged.

These results indicate that the reduced fitness phenotype displayed by unregulated

Gpt2p mutants is dependent on GPAT activity and correlates with accumulation of DAG.

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As discussed before TAG accumulation does not depend on increased activity and can be induced by the mere excess of a catalytically dead Gpt2p.

We have previously shown that Gpt2p is required to channel oleate into TAG synthesis and dephosphorylation of Gpt2p is rapidly induced by oleate (Marr et al. 2012).

If Gpt2-3A increases oleate incorporation into TAG, then we expected that cells expressing

Gpt2-3A will be more efficient at using oleate as carbon source. To challenge this, cells were then serially diluted onto plates containing increasing concentrations of oleate and growth was assessed. No differences were observed between cells expressing wild type or

Gpt2-3A at physiological levels (Figure 5.20). We also included in these experiments cells lacking Gpt2p (gpt2Δ) which are unable to grow on oleate as the only carbon source (Marr et al. 2012).

Figure 5.20. Lack of phosphorylation on endogenously GPT2 does not change cell response to oleic acid. Cells expressing endogenous Gpt2 were grown to mid-exponential phase in defined medium and were plated on synthetic medium (STY) containing 3.5, 6, 8, and 10 mM oleic acid (OA) dissolved in NP40 as described in Material and Methods. Images show cells grown for 4 days at 30 ºC.

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In order to exacerbate the effect of Gpt2-3A, we then decided to overexpress the protein in cells lacking endogenous Gpt2p (gpt2Δ). Surprisingly, cells over-producing

Gpt2-3A or the truncated version displayed high sensitivity to oleate than those expressing native Gpt2p (Figure 5.21). Microscopy inspection of these cells stained with BODIPY

493/503 showed accumulation of oversized LDs in the Gpt2-3A phosphomutant (Figure

5.22). Noteworthy, overexpression of wild type Gpt2p induced an abnormal increase in both number and size of LDs, while Gpt2-3A displayed significantly fewer but oversized

LDs (Figure 5.22).

Altogether these results suggest that the increase LD size observed in Gpt2-3A cells was not due to just an increase in TAG biosynthetic capacity and may indeed be related to the fact that an increase in DAG is detected.

Figure 5.21- Overexpression of unregulated Gpt2p induces sensitivity to oleic acid. Cells overexpressing Gpt2p from a pYES vector were grown to mid-exponential phase in selective defined medium and were serially diluted onto synthetic media (STY) containing 3.5, 6, 8, and 10 mM oleic acid (OA) dissolved in NP40 as described in Material and Methods. Images show cells grown for 4 days at 30 ºC.

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Figure 5.22- Unregulated Gpt2p induces accumulation of oversized but few LDs. Cells overexpressing Gpt2p from a pYES vector were grown to mid-exponential phase (A) and stationary phase (B) in defined selective medium containing 3.5 mM oleic acid. Lipid droplets were stained with BODIPY 493/503 and analyzed by fluorescence microscopy. Stained LDs in cells at mid-exponential phase were quantified with (C) bar graphs representing percentage of large LDs (>0.5 µm) per cell in each strain (D) box plot showing the medians (central horizontal line) of number of LDs per cell with the 25th and 75th percentiles (box).

5.3.5 Concluding Remarks

Results from these investigations indicate that unregulated Gpt2p induces accumulation of TAG but this is probably related to increased proteins levels as catalytic activity is not needed to display this phenotype. On the other hand, only active Gpt2p phosphomutants result in DAG accumulation and abnormally oversized LDs. These phenotypes become toxic only when the mutant protein is overproduced and is catalytically active.

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Chapter 6 – Role of Gpt2p phosphorylation during growth resumption from stationary phase

6.1 Introduction

Glycerolipid homeostasis is maintained by the coordinated input from multiple biosynthetic and catabolic pathways that are regulated in response to different environmental and intracellular cues. TAG metabolism is particularly sensitive to the availability, flow and demand of fatty acids in the cell. GPATs represent the rate limiting and committed step in the synthesis of PA for the de novo synthesis of all glycerolipids, including TAG. Of relevance to this Chapter we need to consider:

1- the sources of acyl-CoA consumed by GPATs and

2- the downstream steps that regulate the flux of fatty acids (FAs) into either

membrane phospholipids or TAG for storage.

With respect to the sources of acyl-CoA used by the GPATs, biochemical characterization of yeast GPATs demonstrated that Sct1p prefers saturated fatty acids while

Gpt2p had no preference between saturated or unsaturated FAs (Z Zheng and Zou 2001).

In addition, in vivo studies determined that Gpt2p was required for the channeling of oleate into TAG and lack of Gpt2p prevented growth of yeast on oleate as the only carbon source.

To our surprise overexpression of unregulated Gpt2-3A impaired growth on oleate, close to the gpt2Δ phenotype, despite displaying higher activity. Puzzlingly, lack of Gpt2p or constitutively dephosphorylated and active Gpt2p resulted in oleate sensitivity. While in the case of gpt2Δ cells, LDs cannot be detected with Nile Red. However, cells overexpressing the unregulated Gpt2-3A enzyme accumulate few but oversized LDs when grown on oleate, resembling the phenotype displayed by cells lacking Sei1, the yeast ortholog of human BSCL2, implicated in congenital lipodystrophy (Heimo Wolinski et al.

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2015). Therefore, in one case, channeling of oleate into TAG is impaired, while in the other one, oleate is incorporated into TAG but is stored in abnormal LDs. In both cases, oleate cannot support growth of these yeast mutants when is offered as the only carbon source.

Critical downstream GPAT steps that regulate the flux of FAs into either membrane phospholipids or TAG for storage involve the interconversions between PA and DAG. PA can be dephosphorylated by PA phosphatases to become DAG, and DAG in turn can be converted back to PA by the activity of DAG kinases (only one in yeast, named Dgk1p)

(Han et al., 2008; Fakas et al., 2011; Pascual et al., 2013). Stress and nutritional conditions that induce DAG synthesis also promote TAG production and biogenesis of LDs in yeast

(Karanasios et al. 2013; Hariri et al. 2018; Pascual and Carman 2013). Channeling of DAG into LDs takes place in the endoplasmic reticulum (ER), more specifically at the nuclear envelope (Barbosa et al. 2015; Adeyo et al. 2011) and at the nuclear ER-vacuole junction

(Hariri et al. 2018). Therefore, LDs originate in specific ER-subcompartments where DAG concentrates in proximity to incipient LDs (Choudhary et al. 2018b; Adeyo et al. 2011), in a process regulated by Sei1p. LDs emerge from the ER and remain associated with the ER membrane even after maturation (N. Jacquier et al. 2011; H. Wolinski et al. 2011;

Szymanski et al. 2007; Mishra et al. 2016b; Kohlwein 2010a), which has functional consequences for lipid and protein exchange between these organelles. Gpt2p (but not

Sct1p) has been found associated with purified LDs most probably from these ER- associated compartments. TAG synthesis takes place in the ER and increases when cells enter stationary phase (Casanovas et al. 2015), inducing LD growth in number and size.

During stationary phase, vacuolar degradation of LDs through lipophagy supports viability in the absence of carbon sources allowing long-term survival (Zutphen et al., 2014).

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Compared to our knowledge on the spatiotemporal details leading to DAG channeling into

TAG synthesis and LD emergence, we know significantly less on how DAG produced from

TAG lipolysis is directed towards synthesis of phospholipids for membrane proliferation.

Upon re-entry of growth from stationary phase, cytosolic TAG lipolysis is mainly supported by LD-resident lipases, Tgl3p, Tgl4p and Tgl5p, providing fatty acids and DAG precursors for membrane lipid synthesis to boost rapid growth (Athenstaedt et al., 2003;

Kurat et al., 2006; Rajakumari et al., 2010). How the metabolic flux of DAG and FAs is channeled into phospholipid synthesis while its conversion back to TAG is prevented is not well understood. It has been suggested that turnover of LDs requires a functional interaction with the ER (Markgraf et al., 2014), positioning Gpt2p near LDs also during lipolysis. A scheme highlighting the main processes in TAG metabolism governing different phases of growth is presented in Figure 6.1.

Figure 6.1- Cycles of TAG metabolism during yeast growth. Typical growth phases of yeast grown in defined medium in the presence of glucose as carbon source and the accompanying changes observed in TAG anabolism/ catabolism.

Yeast cells can switch from TAG-synthesis to consumption responding to different cues depending on the growth phase. During late exponential-early stationary phase, the

PA phosphatase Pah1p generates DAG, and the transacylase Lro1p moves to incipient LDs, where it converts DAG into TAG generating lysolipids like LysoPC as a byproduct. Lro1p appears to have its major role in TAG synthesis during exponential growth (P. Oelkers et

144 al. 2002). Thus, it may generate the first TAG molecules when LDs are formed de novo, whereas the DAG acyltransferase Dga1p would move into the LDs and take over TAG synthesis only when the LDs have reached a sufficient size (P. Oelkers et al. 2002).

When cells exit stationary phase and resume growth, TAG is hydrolyzed to DAG and FAs by LD-resident lipases such as Tgl3p. DAG can be channeled back into the ER and utilized by Dgk1p for biosynthesis of phospholipids through the CDP‐DAG or the

Kennedy pathways (Kuchler et al., 1986; Han et al., 2008; Fakas et al., 2011; Henry et al.,

2012). DAG as a precursor of phospholipids biosynthesis needs to efficiently shuttle from

LDs to the ER membrane. Ice2p facilitates the channeling of DAG from LDs to the ER, a process that mediates the metabolism to efficient phospholipid synthesis and prevents the toxicity from accumulation of DAG (D. Markgraf et al. 2014). When TAG levels are adequately reduced, Dga1p is moved back to the ER from LDs, terminating TAG synthesis on LDs.

Recent results in mammalian cells indicate that 1,3-DAG resulted from TAG lipolysis cannot directly be utilized for phospholipid synthesis (Eichmann et al. 2012).

However, 1,3-DAG is the preferred substrate for DGAT-2, the homolog of yeast Dga1p, suggesting that a futile cycle between TAG and DAG may occur in mammals (Eichmann et al. 2012). In both mammals and yeast, isomerases probably exist, which interconvert the

DAG species. Three main factors have been proposed to regulate the switch between net

TAG-synthesis to hydrolysis (Markgraf et al., 2014): i) formation or utilization of DAG in the ER; ii). Ice2p-mediated metabolic channeling of DAG from LDs to the ER; and iii) migration of Dga1p between the LDs and ER.

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Since Gpt2p contributes to a pool of PA that could influence this switch, we next set to explore the role that phosphorylation of Gpt2p could have in preventing a futile cycle where TAG being hydrolyzed would not be re-converted back to TAG.

6.2 Goals

The fact that the catalytically dead version Gpt2-G261L-3A accumulated similar levels of TAG than Gpt2-3A pointed to mechanisms other than an increase of TAG biosynthesis due to high GPAT activity as the main reason behind it (Chapter 5). Therefore, other explanations were considered.

A standard protocol for experiments performed so far, involved an initial saturated overnight culture that was diluted in the morning and was then allowed to grow for 6 hours to reach log-phase. Considering that cells usually initiate growth resumption from stationary phase by mobilizing TAG from LDs (Figure 6.1), we reasoned that a delay in

TAG lipolysis during this period may be translated in TAG accumulating during exponential growth. Therefore, in addition to favouring TAG synthesis, unregulated Gpt2-

3A may interfere with LD consumption, leading to its accumulation during a growth period where FAs and DAG produced from TAG mobilization should be utilized for membrane proliferation.

At this point, two mutually non-exclusive possibilities were considered:

1- Unregulated Gpt2-3A increases TAG synthesis

2- Unregulated Gpt2-3A decreases TAG lipolysis

The results from investigations on the effect of unregulated Gpt2p on TAG lipolysis during growth resumption from stationary phase are reported in this Chapter.

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6.3 Results and Discussion

6.3.1 Phenotypic characterization of unregulated Gpt2p phospho-mutants during calorie restriction regimes and growth resumption from stationary phase

Metabolism of TAG during a gradual glucose consumption regime follows the biosynthetic and turnover phases depicted in Figure 6.1. It has been proposed that autophagic degradation of LDs (lipophagy) during stationary phase may be a mechanism to support viability of yeast in the absence of carbon sources (Zutphen et al., 2014). In contrast, LD consumption during re-entry of growth from stationary phase involves cytosolic lipolysis by LD-resident lipases, Tgl3p, Tgl4p and Tgl5p (Athenstaedt et al.,

2003; Kurat et al., 2006; Rajakumari et al., 2010). Therefore, cytosolic lipolysis provides

FAs and DAG as precursors to support the synthesis of phospholipids for membrane proliferation during growth resumption, while lipophagy provides FAs as an alternative source of energy during starvation. Cells that reach starvation through gradual consumption of glucose usually do not survive long-term (viability is compromised after a week). By contrast, a calorie restriction regime that permits long-term survival involves sudden glucose depletion from 2% glucose to 0.4% glucose. This acute glucose restriction triggers both bulk autophagy and µ-lipophagy which involves LD turnover (Seo et al. 2017). In this regime yeast can survive for months.

To gain insight into the role of Gpt2p phosphorylation in the regulation of TAG metabolism and LD dynamics, we explored the response of our toolbox of Gpt2-mutant strains to gradual and acute glucose restriction regimes (Figures 6.2 and 6.3). We also included in these experiments a triple tgl3Δ tgl4Δ tgl5Δ strain (3Δ), missing the three LD- associated lipases and a quadruple strain dga1Δ lro1Δ are1Δ are2Δ (4Δ) that cannot make

147 neutral lipids and LDs, accumulating DAG instead in extended ER membranes (Sorger et al. 2004). As shown in Figure 6.2 survival of Gpt2-phosphomutants was compromised in a gradual calorie restriction regimen.

Figure 6.2- Survival of Gpt2p phospho-mutants expressed at endogenous levels during gradual restriction regimes. Indicated strains were grown to mid-exponential phase in defined media containing 2% glucose as carbon source. Cultures were then split into tubes containing fresh medium with 2% glucose (gradual). Every day after, aliquots of cultures were serially diluted (1:10) and plated onto medium containing 2% glucose to monitor their cell survival. Images show survival after 5 days at 30°C. Representative images of one experiment run in duplicate out of 2 independent ones performed in total.

Figure 6.3- Survival of Gpt2p phospho-mutants expressed at endogenous levels during acute calorie restriction regimes. Indicated strains were grown to mid- exponential phase in defined media containing 2% glucose as carbon source. Cultures were then split into tubes containing fresh medium with 0.4% glucose (acute). Every day after, aliquots of cultures were serially diluted (1:10) and plated onto medium containing 0.4% glucose to monitor their cell survival. Images show survival after 50 days at 30°C. Representative images of one experiment run in duplicate out of 2 independent ones performed in total.

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Interestingly, expression of Gpt2-3A in a sct1Δ background displayed a slight improvement compared to Gpt2-3A, suggesting Sct1p contributes to the phenotype displayed by strains carrying unregulated Gpt2p. Minor differences were observed in 3Δ versus 4Δ strains, where lack of neutral lipids and in consequence also LDs in the 4Δ strain seemed to favour survival. To our surprise, all strains tested were viable at least 50 days after initiating the acute calorie restriction regime. These results indicate that Gpt2p phosphorylation is important to allow cells to effectively respond to the metabolic switches that operate during gradual consumption of glucose but does not impact longevity when these transitions are omitted in acute calorie restriction regimes.

In order to obtain direct evidence of a possible delay in TAG lipolysis in cells expressing Gpt2-3A, LD morphology was monitored upon re-entry of cells from stationary phase. As discussed above, typical growth of yeast in medium containing glucose as carbon source, is initiated with a temporary lag period where cells coming from a quiescent stationary phase, reprogram their metabolism to start proliferation. Several adjustments and modulation of lipid metabolic pathways occur during this physiological adaptation, where mobilization of TAG dominates. Fatty acids and DAG produced during TAG breakdown support the synthesis of membrane lipids, required for secretory traffic and plasma membrane expansion during the initial proliferative phase of growth (Rajakumari et al.,

2008; Zanghellini et al., 2008; Kohlwein, 2010).

Wild type and Gpt2-3A strains were grown to stationary phase for 48 hours in defined medium containing glucose, to allow for accumulation of lipid droplets and were then stained with BODIPY 493/503 before resuspending them in an agarose pad containing fresh medium containing 2% glucose. LDs were monitored in single live cells during

149 growth resumption using fluorescence microscopy. Interestingly, BODIPY intensity started to decrease after 90 min in the wild-type strain, while remained high in Gpt2-3A cells for the period analyzed (Figure 6.4). Similar results were obtained when cells were incubated with cerulenin, an antifungal antibiotic that inhibits de novo biosynthesis of fatty acids. Under this condition cells rely almost exclusively on TAG lipolysis as the cellular source of fatty acids to resume growth. Interestingly, an approximately 30 minutes delay in budding appearance was noted in cells expressing unregulated Gpt2-3A compared to wild type cells in the presence of cerulenin. Altogether these results support a delay in TAG lipolysis, which would also explain why TAG also accumulates in cells expressing catalytically-dead Gpt2p variants.

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Figure 6.4- Lack of phosphorylation on Gpt2p shows delayed TAG lipolysis detected by tracking stained lipid droplets morphology with BODIPY 493/503 over time. A) GPT2wt and GPT23xA cells were grown to late stationary phase (48 hours) in SD media to form large LDs and then diluted to fresh media to initiate growth resumption. LDs were stained with BODIPY 493/503 emitting green fluorescence and specific cells were tracked over 130 minutes. In GPT2wt large LDs started breaking down to small LDs through TAG lipolysis after 90 minutes while it happened after 110 minutes in GPT23xA. B. LDs morphology were tracked over time in the presence of 5 µg/mL Cerulenin to enhance the phenotype by inhibiting FAS (de novo fatty acids biosynthesis). LDs turnover occurred after 60 minutes in GPT2wt cells and after 90 minutes in GPT23xA. Scale bar is 5 µm.

6.3.2 Unregulated Gpt2p alters diacylglycerol metabolism and distribution

TAG metabolism is directly related to DAG either as precursor in biosynthesis or as product in turnover pathways. Thus, altered TAG metabolism is directly associated with

DAG imbalance. For example, a block in the conversion of DAG to PA by deletion of the sole yeast DAG kinase Dgk1p results in both DAG and TAG accumulation when de novo

151 fatty acid synthesis is inhibited (Fakas et al., 2011). In this case accumulation of DAG had a toxic effect on growth. Furthermore, simultaneous deletion of DGK1 and the DAG acyltransferase coding gene DGA1 causes a synthetic loss-of-growth phenotype (Garbarino et al. 2009).

Neutral lipid analysis of gpt2Δ cells overexpressing Gpt2-3A or Gpt2-trunc from the pYES plasmid showed between 3 to 4.5 times more DAG after 24 hours of growth in medium containing galactose to induce their expression (Figure 5.19, Chapter V). During this same period these transformants also showed between 1.2 to 1.6-fold more TAG compared to the control. Endogenously expressed Gpt2-3A also displayed a 1.6-fold increase in DAG after 24 hours of growth in medium containing glucose as carbon source

(Figure 5.19, Chapter V). It is possible that DAG already accumulated at earlier time points as well, but our detection method was not sensitive enough to conclusively define it.

Since DAG is produced during growth resumption, primarily from TAG lipolysis, we decided to monitor cytosolic pools of DAG in live wild type and Gpt2-3A mutant cells.

Genetically encoded chimeric constructs coupled with fluorescence microscopy have enabled the analysis of the DAG distribution in mammalian cells, with both spatial and temporal resolution. DAG-binding tandem C1 domains of several proteins including protein kinases C and D (PKC and PKD) fused to fluorescent proteins have been widely used to monitor DAG distribution in cells, and there are extensive and compelling biochemical evidences that these C1 domains specifically bind DAG with high affinity

(Codazzi et al., 2001; Baron et al., 2002; Wang et al., 2003; Stahelin et al., 2004, 2005;

Ohashi et al., 2009; Sánchez-Bautista et al., 2009). We have previously reported the use of the C1 domain of mammalian PKCδ fused to GFP to monitor DAG in live yeast cells

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(Ganesan et al., 2015). This domain used as a genetically encoded DAG sensor has unveiled the localization of cytosolic-facing pools of this lipid, clearly delineating sites of polarized growth in actively growing cells and vacuolar morphology (Ganesan et al.,

2015). The presence of DAG at the vacuolar membrane is consistent with the fact that isolated docked vacuoles display small microdomains that are enriched in DAG, which may facilitate membrane fusion by favoring nonbilayer intermediates and increased membrane curvature (Jun et al., 2004; Wickner et al., 2017). An additional pool of DAG localized to the ER was detected with the DAG sensor (Ganesan et al., 2015) in the dga1Δ lro1Δ are1Δ are2Δ quadruple mutant, known to accumulate DAG due to lack of the four enzymes responsible for the final step in the synthesis of TAG in yeast (Sorger et al. 2004;

Petschnigg et al. 2009; P. Oelkers et al. 2002; Garbarino et al. 2009; Schmidt et al. 2013;

Karin Athenstaedt 2011; Barbosa et al. 2015).

As a first step we monitored both DAG and LDs during growth resumption from stationary phase in wild type and Gpt2-3A cells. Yeast expressing the C1δ-GFP from a plasmid as well as endogenous levels of the LD marker Erg6 fused to RFP, were grown to stationary phase for 48 hours to allow accumulation of LDs. Cells grown to stationary phase for 48 hours were mounted on a solid agar sheet containing fresh medium and cultivated directly on the microscope stage to follow the subcellular distribution of DAG.

Upon growth resumption, DAG-labelled vacuolar membranes began to undergo constant morphological changes with clusters of LDs closely attached (Figure 6.5). LDs were often seen trapped into DAG-containing vacuolar membrane invaginations or sequestered between apposed vacuolar membranes and rings rich in DAG (Figure 6.5). Bright C1δ-

GFP puncta appeared adjacent to LDs positioned at vertex rings formed around apposed

153 vacuolar membranes. A three-dimensional analysis of C1δ-GFP particles revealed that these structures were juxtaposed to lipid droplets but did not co-localize with Erg6-RFP

(Figure 6.5 A-B). When this analysis was performed in cells expressing endogenous Gpt2-

3A, floral vacuolar green patterns like the one shown in Figure 6.5-C were frequently seen.

A closer look at these configurations showed that the DAG signal was coming from apposed membranes from fragmented vacuoles (Figure 6.6). Quantification analysis indicated that 20% of Gpt2-3A cells exhibited this floral phenotype while it was only detected in 4% of wild-type cells. Furthermore, bright DAG puncta were devoid of the lipid droplet staining AutoDOT dye (Figure 6.7) in both wild type and Gpt2-3A images, indicating these structures are not conventional LDs although they are intimately associated with them.

Altogether these results indicate that DAG enriched puncta adjacent to vacuolar membranes are not conventional LDs but are tightly connected to them during LD consumption. Notoriously in Gpt2-3A, the morphology of the DAG enriched areas of the vacuoles is different, and may be the result of DAG accumulation during lipolysis of TAG if we consider our previous results.

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Figure 6.5- DAG rich vacuoles associate with lipid droplets during growth resumption Cells expressing the DAG sensor C1δ-GFP from a centromeric plasmid and endogenous levels of RFP-tagged Erg6 to monitor lipid droplets were grown to stationary phase for 48 hours in selective medium at 30°C. Cells were then pelleted and resuspended in fresh media to allow growth resumption. (A) Time-lapse imaging using confocal microscopy. DAG (green) and LDs (red) were monitored during the first hour after growth resumption, with images captured every 10 min intervals. The first panel shows the overlay with the bright field image. BF, bright field. Scale bar is 2µm. (B) High-resolution images of C1δ-GFP- labelled DAG pools (green) and Erg6–RFP-labelled LDs (red) after one-hour growth re- entry. (Top) A slice (0.3 μm) cropped from a 3D dataset is shown. Arrow point to DAG rich puncta. Scale bar is 2μm. (Bottom) Deconvolved xyz projections of the cell boxed in top image, showing details of the area of interaction between the DAG rich puncta and LDs. DIC, differential interference contrast. Scale bar is 1μm. (C) Time-lapse imaging of wild type and Gpt2-3A cells as in A). Scale bar is 5 µm.

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wild type

Gpt2-3A

Figure 6.6- DAG rich vacuoles from wild type and Gpt2-3A cells. High-resolution images of cells grown as in Figure 6.5 and imaged after 1 hour of growth resumption. 3D projections were rendered from deconvolved z-stacks (0.3 μm) images acquired with a Spinning Disk Confocal Microscope (see Materials and Methods section).

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Figure 6.7- DAG enriched puncta are not lipid droplets. High-resolution images of C1δ- GFP-labelled DAG pools (green) and Erg6–RFP-labelled LDs (red) after one hour growth re-entry. Cells were then incubated with 200 µM AutoDot dye (blue) for staining neutral

158 lipids (see M&M section for details) and were subjected to spinning disk confocal microscopy. A) wild type, B) Gpt2-3A. Scale bar is 2 µm.

It was also noticed that in many cases a DAG enriched dot would break off the vicinity of the vacuole towards the plasma membrane, signposting the emergence of a budding site (Figure 6.8). We then set out to apply time-lapsed live-cell imaging to investigate this further. Using time-lapsed live-cell imaging we confirmed that around 80% of budding cells in the wild-type background exhibited a bright DAG punctum localized to the bud emergence site 15-30 minutes before the bud became noticeable in the bright field

(Figure 6.8-A). Therefore, the positioning of this DAG-rich structure seemed to correlate with polarization of growth. Notoriously, a significant delay (⁓130 min) in the budding time was registered for the mutant Gpt2-3A strain in the conditions of the experiment, resembling the delay noticed when cells were incubated in the presence of cerulenin

(Figure 6.4). This was not directly translated into a much longer lag phase when growth was monitored in liquid cultures.

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Figure 6.8- DAG polarization upon growth resumption is delayed in Gpt2-3A cells. Cells expressing C1δ-GFP (DAG) were grown to stationary phase for 48 hours in synthetic defined selective medium at 30°C, pelleted and resuspended in fresh media to allow growth resumption. Images were captured every 15 min time intervals for a total of 135 min. Arrows point to the green bright puncta that signposts the emergence of the bud. Scale bar is 5 µm. A) wild type; B) wild type and mutant Gpt2-3A. B) Quantification of budding cells over time expressed as % of total budded cells at the end of the period monitored, from fields obtained in A). At least 150 cells were counted for each strain.

6.3.3 Proteomic analysis of Gpt2p enriched membranes and lipid droplets

6.3.3.1 Assessing the importance of DAG/PA ratio and LD formation in Gpt2p stability

Up to here our investigations pointed to a role of phosphorylation in mitigating

Gpt2p activity and protein abundance during a phase of growth when LDs should be consumed and acyl-CoA pools should be utilized for membrane lipid biosynthesis. Since

Gpt2p is part of ER structures tightly associated with LDs, we wondered if Gpt2p degradation visualized by the appearance of free GFP in vacuoles was dependent on this association and the levels of DAG and PA. To test this wild-type Gpt2p fused to GFP was expressed at endogenous levels in wild type cells or mutants lacking the PA phosphatase

Pah1 (pah1Δ). These strains were transformed with plasmids carrying PAH1 or DGK1

161 under the galactose inducible promoter. Therefore, a broad range of transformants with different DAG/PA ratios was generated following the order (from high to low): wt[PAH1]> wt[E]⁓ pah1[PAH1]> wt[DGK1] >pah1[E] > pah1[DGK1]

Live imaging was performed using fluorescence microscopy and the level of Gpt2 degradation observed closely correlated with the ability of cells to form lipid droplets and the DAG/PA ratio. Under conditions where PA accumulates, in pah1[E] or pah1[DGK1],

Gpt2p was associated with membranes and no vacuolar GFP signal was detected (Figure

6.9). These conditions where PA is high and cannot be converted to DAG prevent TAG synthesis and LD biogenesis. Therefore, it seems that Gpt2p degradation in the vacuole may be associated with high levels of DAG and LD interactions with Gpt2p ER-structures.

Interestingly, we observed higher abundance of Gpt2p and Gpt2-3A in a deletion mutant of the phosphatase Nem1, which is in charge of dephosphorylating and activating Pah1, also preventing LD formation (Figure 6.10).

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Figure 6.9- Decreased vacuolar degradation levels of Gpt2 in elevated levels of PA in exponential phase. Cells expressing endogenous GFP-fused Gpt2p in a pah1Δ background carrying a plasmid overexpressing PAH1 (pPAH1) or DGK1 (pDGK1) or empty plasmid as control were grown to mid-exponential phase in Sgal-Ura (2% glucose). Cells were subjected of fluorescence microscopy.

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Figure 6.10- Lack of Nem1p phosphatase induces changes in Gpt2p abundance and a shift in the phosphorylation pattern of the unregulated Gpt2p3xA. Cells expressing physiological levels Gpt2pwt-V5-6xHis and Gpt2p3xA-V5-6xHis in wild type and nem1Δ backgrounds were grown to mid-exponential and stationary phase. Microsomal fractions were prepared and analyzed by western blot using anti-V5 antibody. Loading control displays TCE staining of total protein loaded.

Altogether these results pointed to Gpt2p-LD protein interactions resulting in Gpt2 vacuolar degradation in conditions of high DAG/PA ratios. This ratio would be high during growth resumption when TAG is metabolized to DAG and FAs. Interestingly this pool of

DAG ends accumulating in the vacuolar membrane (Figure 6.6) and Gpt2 degradation occurs in the vacuole. We speculate Gpt2-association with LDs mediates its vacuolar degradation.

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6.3.3.2 LD purification from cells re-entering growth from stationary phase

Given the tight association of Gpt2p with LDs and the differential accumulation of TAG and DAG in Gpt2-3A cells, we decided to analyze the LD proteome to identify putative interactors that could provide mechanistic insight as to why this was happening.

Many studies have analyzed the proteome of LDs isolated from yeast after 24 hours of growth (early stationary phase) but our knowledge about the changes in this proteome occurring during growth resumption from the quiescent phase have not been investigated.

The objective of this part of our study was to identify unique proteins associated with LDs in cells expressing endogenous levels of either wild type Gpt2p or Gpt2-3A. We decided to conduct these investigations in cells that in addition to V5-6xHis tagged Gpt2-proteins were expressing the LD marker Erg6p fused to RFP and the C1δ-GFP DAG-probe to be able to monitor DAG enriched fractions during purification of LDs.

To capture changes occurring during re-entry of growth from stationary phase, cells were grown for 24 hours in defined medium containing 2% of glucose and were then pelleted and resuspended in fresh medium for 45 minutes before initiating the LD purification protocol. This protocol is based on three consecutive rounds of centrifugation where a floating white layer containing LDs and a pellet containing LD associated membranes is obtained at the end. To assess the quality of the LD purification, aliquots of the fractions obtained were analyzed by western blot using antibodies against markers of distinct organelles like ER (Dpm1p); vacuole (Vma2p) and plasma membrane-Golgi

(Pma1) (Figure 6.11). We were curious as to whether the DAG enriched structures observed in live cells (Figures 6.5, 6.6) would be also found in the floating layer containing

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LDs. Consistent with the live imaging results, few distinct DAG enriched structures co- purified with LDs but were not LDs (Figure 6.11-B). The majority of the C1δ-GFP signal was found in the fraction containing vacuolar and ER membranes. Interestingly, a faint signal for a band corresponding to C1δ-GFP was detected in LD samples obtained from mutant Gpt2-3A but not from wild type cells (Figure 6.11). Both wild type and Gpt2-3A were detected in the pellet fraction, although the mutant GPAT was found at much higher levels. This probably reflects the higher stability of Gpt2-3A versus the native protein.

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Figure 6.11- LD purification from wild type and Gpt2-3A cells during growth resumption from stationary phase. A) Western blot analysis of isolated LDs and pellet fractions from cells expressing endogenous Gpt2p or Gpt2-3A (V5-tagged), the LD marker Erg6-RFP and the C1δ-GFP probe. Cells were grown for 24 hours and resuspended in fresh media to allow growth resumption for 45 min before the LD isolation protocol was initiated. Expected MWs: C1δ-GFP (DAG sensor) - 43 kDa; Erg6-RFP (LD marker) - 68 kDa; Vma2 (vacuole marker) - 58 kDa; Dpm1 (ER marker) - 30 kDa; and Pma1 (PM marker) - 100 kDa. B) Representative image of purified LD fraction obtained from wild type cells in A) expressing the DAG sensor C1δ-GFP (green) and endogenous Erg6-RFP (red) after 45 min of growth re-entry (see M&M section for details). Scale bar is 1 µm.

6.3.3.3 Proteomic analysis of LDs obtained from stationary versus growth re-entry phases

An earlier mass spectrometry analysis performed in our group looking at the proteome of LDs isolated from wild-type cells from stationary phase (Shabits, 2017) was contrasted against datasets found in several LD studies previously published (Currie et al.

2014; Grillitsch et al. 2011; Binns et al. 2006) By comparing our results to the literature, a

167 core of 15 proteins were consistently identified in all four studies investigating the yeast

LD proteome, including ours; Ayr1p, Eht1p, Erg1p, Erg6p, Erg7p, Erg27p, Faa1p, Faa4p,

Fat1p, Gtt1p, Hfd1p, Pet10p, Slc1p, Tgl1p, and Tgl3p (Figure 6.12). These proteins are most likely true LD-residents as they were found multiple times in independent experiments from four different groups.

In an independent analysis, cells expressing the V5-6xHis tagged versions of wild type or Gpt2-3A were grown for 24 hours followed by LD purification and Gpt2- enrichment by introducing an extra step where LDs were bound to a Ni-NTA column and then eluted with imidazole (Shabits, 2017). MS analysis from these purified LDs obtained from cells collected during stationary phase found a total of 58 and 113 proteins associated with wild type and Gpt2-3A LDs respectively (Figure 6.13).

Figure 6.12- Proteome of LDs from stationary phase in wild type cells identified by LC-MS/MS compared to the literature. Numbers in brackets are the total amount of proteins from each study. Adapted from (Shabits, 2017).

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Figure 6.13- Proteome analysis of wild type and Gpt2-3A LDs obtained from stationary phase. LDs were enriched in V5 proteins. Peptide threshold ≥ 80%, and a minimum of two peptides. Updated from (Shabits, 2017).

The complete core of 15 LD-resident proteins was detected in wild type LDs while the ER-associated glutathione S-transferase Gtt1p was the only protein of this group missing in LDs from the mutant Gpt2-3A. The other unique protein associated with wild type LDs not found in Gpt2-3A was Cab5, which is a subunit of the CoA-Synthesizing

Protein Complex (CoA-SPC). This complex catalyzes the last step in coenzyme A biosynthesis and has been previously reported to be associated with LDs (Currie et al.

2014) and mitochondria (Reinders et al., 2006). An additional analysis comparing our datasets with those from the literature identified a total of 14 proteins found in both populations of LDs that were never reported to be part of the LD’s proteome before (Table

6.1). Interestingly, a larger number of proteins (41) never reported before in association with LDs was found in the Gpt2-3A dataset (Table 6.2)

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Table 6.1- Proteins found in both wt and Gpt2-3A enriched LDs from stationary phase never reported to be associated with LDs before.

Genes Description ALG11 Alpha-1,2-mannosyltransferase, catalyzes addition of alpha 1,2-mannose residues to the Man5GlcNAc2- PP-dolichol intermediate during asparagine-linked glycosylation in the ER. CHS5 Component of the exomer complex, is involved in the export of select proteins, such as chitin synthase Chs3p, from the Golgi to the plasma membrane. ERG9 Farnesyl-diphosphate farnesyl transferase, joins two farnesyl pyrophosphate moieties to form squalene in the sterol biosynthesis pathway. ERO1 Thiol oxidase required for oxidative protein folding in the ER. GSF2 Endoplasmic reticulum (ER) localized integral membrane protein; may promote secretion of certain hexose transporters. IML2 Protein required for clearance of inclusion bodies; localizes to the inclusion bodies formed under protein misfolding stress. NUS1 Nuclear Undecaprenyl pyrophosphate Synthase, Forms dehydrodolichyl diphosphate syntase complex. RVB1 ATP-dependent DNA helicase, also known as pontin, involved in multiple processes such as chromatin remodeling. RVB2 ATP-dependent DNA helicase, also known as pontin, involved in multiple processes such as chromatin remodeling. SAC1 Phosphatidylinositol phosphate (PtdInsP) phosphatase, involved in protein trafficking and processing, secretion, and cell wall maintenance; regulates sphingolipid biosynthesis through the modulation of PtdIns(4)P metabolism. SEC4 Rab family GTPase; essential for vesicle-mediated exocytic secretion and autophagy. SSH1 Subunit of the Ssh1 translocon complex, involved in co-translational pathway of protein translocation. VPH1 Subunit a of vacuolar-ATPase V0 domain, is located in V-ATPase complexes of the Golgi and endosomes. YKL100C Intramembrane aspartyl protease of the perinuclear ER membrane; acts in a branch of ER-associated degradation (ERAD).

Table 6.2- Proteins found in Gpt2-3A enriched LDs from stationary phase never reported to be associated with LDs before. Highlighted in red are proteins also found in LDs from Gpt2-3A cells from growth resumption experiments.

Genes Description ALE1 Broad-specificity lysophospholipid acyltransferase, may have role in fatty acid exchange at sn-2 position of mature glycerophospholipids ALG1 Mannosyltransferase; involved in asparagine-linked glycosylation in the endoplasmic reticulum (ER) ALG12 Alpha-1,6-mannosyltransferase localized to the ER; responsible for addition of alpha-1,6 mannose to dolichol-linked Man7GlcNAc2 ALG2 Mannosyltransferase in the N-linked glycosylation pathway; catalyzes two consecutive steps in the N- linked glycosylation pathway AXL1 Haploid specific endoprotease of a-factor mating pheromone CDC19 Pyruvate kinase; functions as a homotetramer in glycolysis to convert phosphoenolpyruvate to pyruvate, the input for aerobic (TCA cycle) or anaerobic (glucose fermentation) respiration CDS1 Phosphatidate cytidylyltransferase (CDP-diglyceride synthetase), catalyzes the critical step in the synthesis of all major yeast phospholipids

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CHO2 Phosphatidylethanolamine methyltransferase (PEMT); catalyzes the first step in the conversion of phosphatidylethanolamine to phosphatidylcholine EFT2 Elongation factor 2 (EF-2), catalyzes ribosomal translocation during protein synthesis EMC1 Member of conserved endoplasmic reticulum membrane complex; involved in efficient folding of proteins in the ER ERG11 Catalyzes the demethylation of lanosterol during the biosynthesis of ergosterol ERG24 Acts in ergosterol biosynthesis ERG26 C-3 sterol dehydrogenase involved in ergosterol biosynthesis GCN1 Positive regulator of the Gcn2p kinase activity, is involved in the regulation of translational elogation; IFA38 Microsomal beta-keto-reductase; accumulate high levels of dihydrosphingosine, phytosphingosine and medium-chain ceramides LCB1 Component of serine palmitoyltransferase; responsible along with Lcb2p for the first committed step in sphingolipid synthesis LCB2 Component of serine palmitoyltransferase; responsible along with Lcb1p for the first committed step in sphingolipid synthesis MCD4 Protein involved in GPI anchor synthesis, GPI stands for glycosylphosphatidylinositol MSC7 Meiotic Sister-Chromatid recombination, Protein of unknown function OLA1 P-loop ATPase with similarity to human OLA1, identified as specifically interacting with the proteasome OST1 Alpha subunit of the oligosaccharyltransferase complex of the ER lumen, complex catalyzes asparagine- linked glycosylation of newly synthesized proteins PEX30 ER-resident protein involved in peroxisomal biogenesis PMT4 Protein O-mannosyltransferase; transfers mannose residues from dolichyl phosphate-D-mannose to protein serine/threonine residues SAC6 Fimbrin, actin-bundling protein, is involved in organization and maintenance of the actin cytoskeleton SAM2 S-adenosylmethionine synthetase; catalyzes transfer of the adenosyl group of ATP to the sulfur atom of methionine SCS2 Integral ER membrane protein regulates phospholipid metabolism SCT1 Glycerol 3-phosphate/dihydroxyacetone phosphate sn-1 acyltransferase; dual substrate-specific acyltransferase of the glycerolipid biosynthesis pathway SEC18 AAA ATPase and SNARE disassembly chaperone; required for vesicular transport between ER and Golgi SEC66 Non-essential subunit of Sec63 complex SEY1 Dynamin-like GTPase that mediates homotypic ER fusion SSB2 Cytoplasmic ATPase that is a ribosome-associated molecular chaperone, may be involved in the folding of newly-synthesized polypeptide chains STE24 Highly conserved zinc metalloprotease involved in the CAAX-box dependent processing and maturation of a-factor mating pheromone STT3 Subunit of the oligosaccharyltransferase complex of the ER lumen; complex catalyzes asparagine-linked glycosylation of newly synthesized proteins SUR4 Elongase; involved in fatty acid and sphingolipid biosynthesis; synthesizes very long chain 20-26-carbon fatty acids from C18-CoA primers; involved in regulation of sphingolipid biosynthesis TSC13 Enoyl reductase; catalyzes last step in each cycle of very long chain fatty acid elongation; localizes to ER, highly enriched in a structure marking nuclear-vacuolar junctions UBI4 Ubiquitin; becomes conjugated to proteins, marking them for selective degradation via the ubiquitin-26S proteasome system UBX3 Clathrin-coated vesicle component, regulator of endocytosis; copurifies with the DSC ubiquitin ligase complex URA2 Bifunctional carbamoylphosphate synthetase/aspartate transcarbamylase; catalyzes the first two enzymatic steps in the de novo biosynthesis of pyrimidines

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VAC8 Vacuolar membrane protein; required for homotypic vacuole-vacuole fusion and for nucleus-vacuole junction formation VMA5 ATP-binding protein that is essential for protein sorting, vesicle docking, and fusion at the vacuole; binds to SNARE domains VPS1 Dynamin-like GTPase required for vacuolar sorting; also involved in actin cytoskeleton organization, endocytosis, late Golgi-retention of some proteins, regulation of peroxisome biogenesis YNR021W Putative protein of unknown function. Has genetic interactions with CHO2, OPI3 and INO2

Enrichment analysis of these 57 unique proteins found in LDs from Gpt2-3A using the

KEGG database (ClueGO plugin, Cytoscape) showed significant categories associated with lipid metabolism including glycerophospholipid metabolism, fatty acid elongation and unsaturation, sphingolipid metabolism and sterol synthesis (Figure 6.14). In addition, phagosome (vacuole in yeast), longevity regulation, and N-glycan biosynthesis were also significantly enriched. Interestingly, Sct1p was identified in the Gpt2p-3A enriched LDs but not in those enriched in native Gpt2p. To our knowledge this is the first time Sct1p has been detected in a sample of purified LDs, suggesting that de-phosphorylation of Gpt2p may impact localization of the other GPAT.

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Figure 6.14- Enrichment categories of unique proteins (57) found in Gpt2-3A LDs obtained from stationary phase. A) Histogram with enrichment of identified genes in significant terms. Bars represent the percentage of found genes compared to all the genes associated with the term (# of genes identified/total number of genes in term multiplied by 100). Numbers at the end of the bars indicate the number of proteins identified in each term. Colour identifies groups of similar terms. Single and double red asterisks indicate terms with p-values below 0.05 and 0.005, respectively. B) Map of terms (nodes) with identified genes. Edges link genes to respective terms. Genes can belong to more than one term. Node size represents significance and colour identifies groups as in A.

Two independent LD purifications from cells resuming growth from stationary were performed. Proteins that were found associated with both LD-samples and that at least presented 2 peptides resulted in a total of 201 proteins selected for further analyses. A comparison of this dataset with those proteins identified for stationary-phase LDs identified

34 proteins in common and 167 unique to LDs from cells resuming growth (Figure 6.15).

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Figure 6.15- Proteome analysis of wild type LDs obtained from stationary phase versus growth re-entry. Only proteins detected in two independent LD isolations and presenting at least 2 peptides were included in each dataset.

Enrichment analysis of this unique group of proteins identified several categories related to glycolysis and TCA cycle, amino acid and its associated coenzyme A metabolism, RNA metabolism, and glycerophospholipid metabolism (Figure 6.16).

Figure 6.16- Enrichment categories of unique proteins (167) found in wild type LDs obtained during growth resumption.

Only 67 proteins matched the cut-off criteria in the case of LDs from Gpt2-3A. A comparison with the wild type dataset obtained for wild type LDs indicated that only 9 proteins were unique to Gpt2-3A samples (Figure 6.17). This group was enriched in proteins listed in Table 6.2 (in red) as never reported to be associated with LDs before.

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Enrichment analysis of these 9 proteins identified only the category of glycerophospholipid metabolism with high significance. A comparison of proteins found enriched in wild type samples for the same category is presented in (Figure 6.18).

Figure 6.17- Comparison between wild type and Gpt2-3A proteins identified in LDs from growth resumption.

Figure 6.18- Glycerophospholipid metabolism identified as the only lipid metabolic category in unique protein dataset from Gpt2-3A compared to wt.

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Interestingly, Sct1 was again found associated with LDs in samples obtained from cells expressing the phosphomutant Gpt2-3A. Also intriguing was the fact that the acyltransferase Ale1 capable of acylating the product of the GPAT reaction LysoPA, was consistently found with LDs in Gpt2-3A samples.

We continued our analysis as we were curious to investigate the 58 overlapped proteins resulted in figure 6.18. Clustering analysis showed enrichment of proteins in different metabolic pathways shown in Figure 6.19. We compared this group of proteins found in both wt- and 3A-LDs from growth resumption with the proteins associated with wt-LDs during stationary phase (Figure 6.20). Interestingly, clustering analysis showed association of unique proteins with LDs involved in phagocytosis during growth resumption (Figure 6.21).

Figure 6.19- Enrichment categories of proteins (58) found in both Gpt2-wt and Gpt2- 3A LDs obtained from growth resumption.

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Figure 6.20- Comparison between LD-enriched proteins found in both Gpt2-wt and Gpt2-3A cells during growth resumption (from figure 6.17) versus proteins found in Gpt2-wt LD during stationary phase.

Figure 6.21- Enrichment categories of unique proteins to Gpt2-wt LD found in stationary phase and unique proteins enriched with LDs during growth resumption in Gpt2-wt and Gpt2-3A cells.

Taking advantage of the toxic phenotype displayed by the overexpression of Gpt2-

3A, we decided to investigate the impact that deleting the LPAAT Slc1 (part of the core 15 group of LD-resident proteins) or Ale1 (never reported to be part of LD proteome) would have on the severity of the phenotype. To our surprise, cells lacking Ale1 were unable to

177 grow on galactose not only when the phosphomutant Gpt2-3A was overproduced, but also when wild type GPT2 was overexpressed (Figure 6.22). No differences in the severity of the phenotype were observed for cells depleted of Slc1p. One possible interpretation for these results is that cells lacking Ale1p may accumulate intermediate lysolipids that arise from increase GPAT activity becoming toxic to the cell.

Figure 6.22- Overexpression of wild type and phosphomutant Gpt2-3A in cells lacking LPAAT enzymes. Indicated strains overexpressing wild type or GPT2-3A from a pYES vector were grown to mid-exponential phase in Sgal-Ura medium. Serial dilutions of the cultures were then plated on medium containing glucose (repressor) or galactose (inducer). Plates were incubated for 4 days at 30 ºC.

Future investigations on understanding the role of Sct1p and Ale1p in LD metabolism is warranted. One hypothesis to test is that recruitment of a different set of acyltransferases in addition to unregulated Gpt2p and Slc1p already associated with LDs may put a constraint in the utilization of acyl-CoA pools and may change the acyl-chain profile of cellular lipids. In addition, higher flux of acyl-CoAs into an hyperactive glycerophospholipid metabolic pathway would impact other pathways that also consume acyl-CoA pools, like sphingolipids, fatty acid modifying pathways and remodelling

178 pathways. Interestingly, enzymes from each of these pathways were enriched in the protein dataset obtained for Gpt2-3A enriched LDs in stationary phase (Figure 6.15 and Table 6.2).

Since the synthesis of complex sphingolipid consumes very long chain fatty acids the enrichment seen for sphingolipids and fatty acid elongation categories could be pointing in the same direction, de novo synthesis of ceramide and derived complex sphingolipids. It is interesting to note that beyond synthesis of ceramide in the ER, the biosynthetic pathway for complex sphingolipids in the Golgi consumes phosphatidylinositol and generates an important amount of DAG. We then decided to investigate the crosstalk between glycerolipids and sphingolipid pathways.

Myriocin is an amino fatty acid antibiotic derived from certain thermophilic fungi that inhibits serine palmitoyltransferases (SPT), Lcb1p, Lcb2p, Tsc3p, and suppresses de novo biosynthesis of sphingolipids resulting in decreased levels of ceramide (Chen et al.,

1999). Aureobasidin A is a cyclic depsipeptide antibiotic which inhibits inositol phosphorylceramide (IPC) synthase, Aur1p, resulting in increased ceramide and decreased complex sphingolipids levels (Cerantola et al. 2009).

Treatment of GPAT mutants with 2 µM Myriocin resulted in growth deficiency in the wild type strain while cells carrying Gpt2-3A were resistant. Interestingly, sct1Δ in a

GPT2wt background displayed hypersensitivity to the drug, but this was reverted when the unregulated Gpt2-3A was expressed instead (Figure 6.23). Notoriously, the same set of strains treated with Auriobasidin A exhibited opposite phenotypes (Figure 6.24).

Considering that Myriocin blocks a step upstream of ceramide and Aureobasidin A targets a step downstream of ceramide (Figure 6.25), resistance to myriocin and sensitivity to

Aureobasidin A indicates the cell accumulates ceramide. This could be in fact the case of

179 cells with constitutive de-phosphorylation of Gpt2-3A. Conversely, ceramide must be lower in cells lacking Sct1p. Levels of ceramide in these strains should be determined to conclusively interpret not only the effects of these drugs but the relationship that this may have with increased DAG and TAG observed in cells with unregulated Gpt2p.

Figure 6.23. Effect of myriocin on cells expressing different sets of GPATs. Indicated strains expressing endogenous levels of GPT2-3A phospho-mutant or native GPT2 were grown to mid-exponential phase and were serially diluted (1:10) onto plates containing defined medium ± 2 µM Myriocin dissolved in methanol (control plates contained methanol vehicle only). Plates were incubated for 4 days at 30 °C.

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Figure 6.24. Effect of Aureobasidin A on cells expressing different sets of GPATs. Indicated strains expressing endogenous levels of GPT2-3A phospho-mutant or native GPT2 were grown to mid-exponential phase and were serially diluted (1:10) onto plates containing defined medium ± the indicated concentrations of Aureobasidin A dissolved in ethanol (control plates contained ethanol vehicle only). Plates were incubated for 4 days at 30 °C.

Figure 6.25. Model explaining the combined phenotypes of resistance and sensitivity to Myriocin and Aureobasidin A in Gpt2-3A cells.

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Chapter 7 - Discussions and future directions

Evidence presented in this work shows that lack of phosphorylation at Ser 664, 668 and 671 or complete removal of the last 133 amino acids containing the majority of phosphosites on Gpt2p disturbs lipid homeostasis. Most specifically seems to alter neutral lipid metabolism with a notorious effect on TAG levels. Constitutive phosphorylation can preserve the protein from vacuolar degradation during exponential phase resulting in higher abundance of the protein with highest stability displayed by the truncated Gpt2p mutant

(Figures 5.3 and 5.4).

A shift from a P2 to P1- phosphorylation state in Gpt2p can be induced by the absence of Sct1p (Figures 5.5 and 5.16) suggesting that the two GPATs have the ability to

“sense” the presence of the other one. Gpt2p seems to be more phosphorylated, and probably less abundant, in the period of fermentative growth when levels of saturated fatty acids are high. Gpt2p phosphorylation status and abundance switches as cells enter the diauxic and post-diauxic shift. Interestingly, during this period unsaturated FA cellular content goes up (Casanovas et al. 2015). Opposite protein abundance patterns are seen for

Sct1p, dropping in stationary phase. Knowing that 16:0 is the preferred substrate of Sct1p

(Zheng et al., 2001; Oelkers et al., 2016), and co-overexpression of Sct1p and Ole1p compete for the 16:0 pool of FA (De Smet et al., 2012) brings up the hypothesis that Sct1p might be more committed in exponential phase by channeling saturated FA into the sn-1 position of G-3-P and producing more phospholipids as required for cell proliferation during this phase of growth (Figure 7.1).

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Figure 7.1- Proposed model for contribution of Gpt2p and Sct1p during three different growth phases in regulated and unregulated Gpt2 backgrounds. A) exponential phase, B) stationary phase, C) Growth resumption.

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Figure 7.2- Proposed model suggesting the presence of a futile TAG cycle during growth resumption in Gpt2p-3A. During growth resumption in Gpt2-3A background, acyl-CoA instead of being used into CDP-DAG and Kennedy pathways, are channeled into an active futile cycle of TAG formation and consumption. LPAATs, lyso-PA acyltransferase; DGATs, Diacylglycerol acyltransferase; PAP, PA phosphatase.

Deletion of Sct1p decreases the cellular C16:0 content by almost 50%, whereas overexpression of Sct1p increases C16:0 levels at the expense of C16:1 and C18:1 (De

Smet et al., 2012). Overexpression of Sct1p dramatically decreases the desaturation of fatty acids and affects phospholipid composition especially on PC (De Smet et al., 2012).

Overexpression of Sct1p results in increased synthesis of PA and a consequent overall increase in cellular lipid content forming supersized LDs (Pagac et al. 2016). In contrast to deletion of SCT1, overexpression does not affect the relative rates of synthesis of TAG versus sterol esters in the neutral lipid fraction (Pagac et al. 2016).

Analysis of mRNA levels revealed that expression of SCT1 is reduced in stationary phase while expression of GPT2 is elevated (Alvarez-Vasquez et al. 2007). In agreement with this finding for Gpt2, our RT-qPCR analysis shows 1.5 ± 0.35 and 1.24 ±

0.11-fold increased for levels of GPT2 expression during stationary phase using housekeeping reference genes TFC1 and TAF10 respectively (Figure 7.3).

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Figure 7.3- Increased levels of GPT2 gene expression in stationary phase. RT-qPCR analysis showing ratio of ΔCt stationary phase / ΔCt exponential phase of GPT2 using TFC1 and TAF10 reference genes. Experiment was done in triplicates (see Material and Methods). Representative experiment out of 3 independent experiments.

We can speculate that Gpt2p is more abundant during stationary phase while Sct1p is probably more abundant during exponential phase.

Based on our results, Gpt2p-3A in exponential phase is less prone to be degraded in vacuoles, forming large LDs and is more active, probably overlapping with Sct1p therefore, total GPAT activity is at an abnormal maximum level (Gpt2-3A+Sct1p), almost

2-fold higher than wt, leading to formation of large LDs. (Figure 5.16 – Chapter V). Even higher levels as those achieved by overexpression of Gpt2p-3A triggers Sct1p degradation in exponential phase, suggesting excess level of GPAT can be sensed by the cell (Figure

7.4).

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Figure 7.4- Overexpression of Gpt2p lacking phosphorylation increases degradation of Sct1 in exponential phase. Cells expressing endogenous GFP-fused Sct1 in a gpt2Δ background carrying a pYES plasmid overexpressing Gpt2-wt or Gpt2-3A or empty plasmid as control were grown to mid-exponential phase in Sgal-Ura media (2% galactose). Cells were subjected to fluorescence microscopy. Representative experiment out of 3 independent experiments. Scale bar is 5 µm.

Overexpression of Gpt2p-3A also shows weaker growth pattern than overexpression of wt (Figure 5.19 - Chapter V) while overexpression of Gpt2-3A in a sct1Δ background grows better. On the other hand, overexpression of catalytically dead Gpt2-wt and 3A do not show growth deficiency, indicating GPAT activity mediates this effect. The growth deficiency might be caused by toxicity from elevated levels of DAG (Figure 5.19 -

Chapter V) which can be alleviated in the absence of Sct1p.

Overexpression of Gpt2p-3A increases sensitivity to oleate (18:1) (Figure 5.22 -

Chapter V) and mimics the behaviour of Sct1p in a gpt2Δ background which is not able to

186 use oleate as the only carbon source. This opens the possibility that lack of phosphorylation in Gpt2p alters acyl-CoA preference.

We have seen that lack of phosphorylation on Gpt2p results in larger LDs and delayed in TAG lipolysis upon growth resumption (Figures 5.10 and 6.4 - Chapters V and

VI). We can speculate that during stationary phase, increased levels of unsaturated FAs are channeled to TAG (Connerth et al., 2009; Grillitsch et al., 2011; Eisenberg and Buttner,

2014; Casanovas et al., 2015), it is then possible that when TAG lipolysis is triggered upon re-entry to growth from stationary phase, unsaturated FAs which are not preferred substrate for Sct1p and probably not for Gpt2-3A (more investigation is required) may cause a delay in TAG lipolysis, channeling produced acyl chains into phospholipids (Figure 7.1). This highlights the role of Gpt2p-wt in channeling of unsaturated acyl chains from TAG lipolysis. Another possibility dealing with delayed TAG lipolysis during growth resumption in unregulated Gpt2p could be the induction of a futile cycle of TAG metabolism (Figure 7.2). Produced acyl-chains seem to be recycled to form TAG instead of being channeled to form phospholipids.

It seems that phosphorylation has a key role in regulation of both Gpt2p and Sct1p responding to the growth status of the cell and metabolites fluctuations. Saturated FA can indirectly induce phosphorylation of Sct1p. Phosphorylation inactivates Sct1p and that

Cst26p, acyltransferase using saturated very long chain fatty acids, indirectly regulates

(de)phosphorylation of Sct1 (De Smet et al., 2012). In our proteomic analysis, Cst26p has been shown to be associated with LDs after growth resumption in Gpt2p-wt but not in

Gpt2p-3A. Cst26p maybe it can indirectly activate Gpt2p by inducing dephosphorylation

187 to consume saturated acyl chains produced form TAG lipolysis while it cannot regulate

Gpt2p-3A leading to a delay in TAG lipolysis.

7.1 Future directions

Further investigations are required to validate whether Gpt2 and Sct1p function are growth phase-dependent responding to the fluctuation levels of unsaturated acyl-chains.

On the other hand, as they catalyze the rate limiting step of glycerolipid biosynthesis by addition of acyl-chain with certain degree of saturation on sn-1 position of G-3-P rule out addition of acyl-chain to the next positions responding to the growth conditions which highlights their role in regulating glycerolipids composition during different growth status.

Tracking glycerolipids composition and acyl chain profiles can be analyzed in specific

GPAT backgrounds (knockout or overexpression) at different timing of growth.

It would be interesting to check abundance level of GPATs or gene expression levels in specific time point in cells grown in the presence of different fatty acid sources such as 16:0, 18:1 to investigate which GPAT is predominant for channeling specific FA at distinct phases of cell growth followed by lipid profile analysis for both neutral lipids and phospholipids.

Furthermore, phosphorylation levels of Sct1p can be manipulated to form a dephosphorylated enzyme (more activated) and check its impact on Gpt2p activity and also lipid profiles and timing of TAG mobilization. Mass spectrometry analysis can be done to study protein-protein interactions.

Since our mass spectrometry analysis has detected Sct1p associated with LDs in

Gpt2p-3A cells, and also Sct1p indirectly affects the phosphorylation status of Gpt2p, it

188 would be interesting to compare protein-protein interaction of Gpt2p-3A in the wild type and sct1Δ background to check whether association of Sct1p affects Gpt2p interactions with specific proteins depending on its phosphorylation status.

As the abundance levels of FA activator enzymes changes during growth phases

(Casanovas et al. 2015) and they might have preference in usage of acyl-chain with distinct length and unsaturation level, experiments aimed at identifying genetic interactions between unregulated Gpt2p and these set of enzymes could provide mechanistic insight to understand the phenotypes unveiled by our investigations.

189

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Appendix A Investigation of possible kinases phosphorylating Gpt2p

A.1 Introduction

Studies on mammalian GPAT show that the mitochondrial GPAT (mtGPAT1) is regulated post-translationally via phosphorylation by casein kinase-2 (CK2) causing enhanced GPAT1 activity 50–200% in liver (Onorato et al., 2005). Ser 632 and Ser 639 in its C-terminal end are identified as phosphorylation sites corresponding to CK2 consensus sequences (Bronnikov et al., 2008), but the physiological relevance of these phosphorylation events remains unknown.

AMP-activated protein kinase (AMPK), mammalian ortholog of SNF1 in yeast, a sensor of cellular energy availability can phosphorylate and inhibit GPAT1 activity

(Takeuchi et al., 2009). Recent studies have shown that GPAT3 and GPAT4 may play different roles in glycerolipid metabolism: GPAT3 is a critical regulator for lipid accumulation during adipocyte differentiation (Shan et al., 2010), whereas GPAT4 may be more responsible for constitutive glycerolipid metabolism. They have shown that function of GPAT3 and GPAT4 is controlled by insulin-mediated phosphorylation, implying their importance in lipogenesis. Three putative phosphorylation sites, S68, S77, and Y423 have been predicted in human GPAT3 by PhosphoSitePlus (www.phosphosite.org) which are supported by experimental evidence (Shan et al., 2010), but no phosphorylation site in hGPAT4 is recorded on this database.

Recent evidence indicates possible involvement of GPAT1 in the mTOR

(mammalian target of rapamycin) signaling pathway. Overexpression of GPAT1 leads to elevated PA levels causing impaired insulin signaling, reduced insulin-induced suppression

225 of gluconeogenesis, and substantially prevented mTOR complex2 (mTORC2) activity, which induced peripheral and hepatic insulin resistance (Zhang, 2011; Yu et al., 2018).

A.2 Goals

Identifying possible kinases and phosphatases regulating Gpt2p through phosphorylation enables us to understand the role of Gpt2p in the cellular response to availability of nutrients and growth status. As a preliminary approach to investigate the impact of specific signalling pathways on Gpt2p phosphorylation status we examined the effect that lack of different kinases had on the phosphorylation pattern or abundance level of Gpt2p.

A.3 Results and Discussion

Mass spectrometry analysis in our lab identified Psk1p kinase, an emerging regulator of glucose and lipid metabolism in mammals (Zhang et al., 2015), as one of the proteins interacting with Gpt2p-wt in two independent immuno-affinity purification experiments

(Shabits, 2017). Therefore, Psk1p was the first kinase of interest which is the mammalian

PAS kinase homologue. PAS kinase responds to the glucose levels and plays a critical role in glucose homeostasis in mammalian cells (Da silva et al., 2004; Wilson et al., 2005; An et al., 2006). Yeast contains two well-conserved homologs of mammalian PAS kinase,

PSK1 and PSK2 (Rutter et al., 2002; Grose et al., 2007; DeMille et al., 2014). PAS kinase activation in response to non-fermentative carbon sources is dependent on the Snf1 kinase

(DeMille et al., 2015). Snf1p, a key mediator of the regulatory switch between fermentative growth and respiratory growth, can phosphorylate and activate Psk1p by non-glucose

226 carbon sources such as raffinose (Grose et al., 2007). This signaling cascade occurs through the SNF1-dependent phosphorylation and activation of Psk1p, which in turn phosphorylates and activates poly(A)- binding protein 1 (Pbp1p), which then inhibits

TORC1 through sequestration at stress granules.

Our previous results indicated that different carbon sources affected membrane association of Psk1p (Shabits, 2017). In semi- (galactose) or poorly fermentable (raffinose) carbon sources that stimulate respiration process (Randez-Gil et al., 1998), Psk1p associated with microsomes, especially at stationary phase. On the other hand, when the cells were grown in the fermentable sugar, glucose, all Psk1p was found in the soluble fractions and none was detected in the microsomal fractions (Figure A.1).

We questioned whether Psk1p is a Gpt2p kinase which its membrane association is induced by galactose or raffinose. To answer this question, an experiment was designed to study the phosphorylation pattern and abundance levels of endogenous Gpt2p-V5-6xHis and Gpt2p-3A-V5-6xHis in cells lacking Psk1p (psk1Δ) in the presence of glucose or galactose in both exponential and stationary phases. We hypothesised that Psk1p might phosphorylate Gpt2p when its membrane association is induced in galactose. Therefore, lack of Psk1p in the presence of galactose could induce a P1 shift (dephosphorylated

Gpt2p). No notable change was observed in phosphorylation pattern of Gpt2p (Figure A.2).

Densitometry analysis of three independent experiments did not show consistent changes in the abundance levels of proteins. This result suggests that Psk1p is probably not a possible kinase of Gpt2p.

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Figure A.1- Psk1p-TAP associates with microsomes when grown to stationary phase in the presence of galactose or raffinose, but not glucose. Psk1p-TAP cells were grown in rich media containing either glucose (YPD), galactose (YPGal), or raffinose (YPRaf). Soluble (S) and microsomal (M) fractions were collected at exponential (Exp., 6 hours after growth resumption) and stationary (Stat., 24 hours after growth resumption) phases. Approximately 15% of the soluble and microsomal samples were loaded. Loading control is SDS-PAGE gel visualized with TCE and UV light before transfer to membrane. Obtained from (Shabits, 2017).

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Figure A.2- Lack of Psk1 kinase did not change phosphorylation pattern in Gpt2p-wt and Gpt2p-3A. Endogenous Gpt2-wt-V5 and Gpt2-3A-V5 in the wild type and psk1Δ backgrounds were grown to mid-exponential phase (A) and stationary phase (B) in the presence of glucose or galactose. Cell lysate samples of proteins were analyzed by western blot using anti-V5 antibody. Loading control was SDS-PAGE gel visualized with TCE and UV light before transfer to membrane. (C) Densitometry analysis for average abundance levels of proteins ± SE normalized to wt strain for three independent experiments in glucose and two independent experiments for galactose.

In the next step, Snf1p which is the regulatory enzyme upstream of Psk1p was studied as one of possible kinases of Gpt2p. In Gpt2p, Thr692 and Ser693 have been identified as targets of Snf1p (Chapter IV, Table 4.2). We wondered whether lack of Snf1p affects on phosphorylation pattern of Gpt2p in the presence of glucose and galactose in both exponential and stationary phases were examined. Western blot analysis did not show remarkable changes in the phosphorylation pattern and levels of Gpt2p-wt indicating it is presumably not regulated by Snf1p (Figure A.3). This experiment was done only once and needs to be repeated.

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Figure A.3- Phosphorylation pattern and abundance level of Gpt2 in the absence of Snf1 kinase. Cells expressing endogenous Gpt2-wt-V5 in the wild type and snf1Δ backgrounds were grown to mid-exponential and stationary phase in defined medium (2% glucose). Cells were lysed and analyzed by western blot using anti-V5 antibody.

TORC1 was the next kinase to be examined. Other than its regulatory role in response to the carbon sources, it has been also shown that TORC1 is a key regulatory enzyme controlling cellular PA:DAG ratio (Ouahoud, 2018). On the other hand, inhibition of TORC1 by Rapamycin induces LD accumulation in yeast (Madeira et al., 2014) and also can stimulate phosphorylation Orm1p and Orm2p which inhibit serine palmitoyltransferase (SPT) enzymes catalyzing the rate-limiting step of de novo synthesis of sphingolipids (Shimobayashi et al., 2013). This results in increased levels of complex sphingolipids and DAG followed by decreased levels of ceramide. So TORC1 can be a possible kinase involved in regulation of Gpt2p responding to the PA:DAG ratio and probably a crosstalk with sphingolipids biosynthesis pathway. TORC1 and PKA are in

231 distinct but parallel pathways that converge on common target proteins or processes

(Santhanam et al., 2004; Stephan et al., 2009). TORC1 and PKA which are nutrient- sensing kinases can phosphorylate downstream protein targets in the presence of nutrients leading to protein synthesis and cellular proliferation. Their response to the glucose levels is opposite of Snf1p→ Psk1p pathway (Figure A.4). Glucose starvation supresses TORC1 and PKA while Snf1p→Psk1p pathway is activated (Zhang et al., 2017).

Figure A.4- Signaling pathways responding to the glucose levels.

It has been also shown that TORC1 by acting upstream of PKA can activate PKA toward some substrates (Soulard et al., 2010). In Gpt2p, nine residues are identified that can be phosphorylated through TORC1/PKA signaling pathway (Soulard et al., 2010)

(Chapter IV, Table 4.2).

As a preliminary step of characterization, growth behaviour of Gpt2p phosphomutants were analyzed in the presence of rapamycin, an inhibitor of Torc1 activity (Sehgal,

2003). Gpt2p phosphomutants were grown to mid-exponential phase in SD and then plated

232 on SD plates containing 0.01 µg/mL Rapamycin (Figure A.5). No remarkable changes in the growth pattern of strains was observed. Our results did not suggest the presence of a regulatory effect of TORC1 on Gpt2p and need further analysis.

Figure A.5- Growth pattern of Gpt2p phosphomutants in the presence of Rapamycin. Cells were grown to the mid-exponential phase in YPD, then were serially diluted (1:10) in fresh media and plated on YPD plate + 1% DMSO as control and YPD + 0.01 µM Rapamycin. Image shows growth pattern after 4 days at 30 °C. TORC2/torc1Δ and TORC1/TORC2 strains expecting growth and no growth, respectively were used as controls.

Another kinase involved in regulation of sphingolipids biosynthesis was target of

Rapamycin complex 2 (TORC2) (Figure A.6). Orm1p and Orm2p can be stimulated by

TORC2-dependent phosphorylation of Ypk1p leading to induced activity of Lcb1p and

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Lcb2p, enzymes catalyzing the rate-limiting step of sphingolipids biosynthesis, followed by increased levels of ceramide (Roelants et al., 2011).

Figure A.6- Regulation of sphingolipids biosynthesis pathway through TORC2/Ypk1p signaling. When Sphingolipids level is compromised Torc2p phosphorylates and activates Ypk1p that can phosphorylate and supress the inhibitory effect of Orm1p/Orm2p complex on Lcb1p, Lcb2p, and Tsc3p. Red cross shows the inhibitory effect of activated Ypk1p by Torc2p.

Our earlier results indicated lack of phosphorylation in Gpt2p shows sensitivity to

Aureobasidin A which is an inhibitor of IPC synthase step. We hypothesized that TORC2 might regulate Gpt2p through phosphorylation by Ypk1p as responding to the changes in the level of ceramide. Since level of ceramide acts as a feedback loop controlling phosphorylation of Ypk1p through TORC2, YPK1 was deleted to study the effect of impaired loop and changes in the levels of sphingolipids in phosphorylation of Gpt2p

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Changes in phosphorylation pattern would suggest regulation of Gpt2p by the

TORC2/YPK1 axis.

Phosphorylation pattern of Gpt2p-wt and Gpt2p-3A were analyzed in a ypk1Δ background in the presence of glucose and galactose, since we had detected the effect of galactose on overexpression of Gpt2p phosphomutants Our results did not show relevant changes in the phosphorylation pattern or abundance of proteins suggesting Gpt2p is not the substrate of TORC2/YPK1 signaling pathway (Figure A.7). This experiment was done only once and needs to be repeated.

Figure A.7- Phosphorylation pattern of Gpt2p-wt and Gpt2p-3A in the absence of YPK1. Cells expressing endogenous Gpt2-wt-V5 and Gpt2-3A-V5 in the wild type and ypk1Δ backgrounds were grown to mid-exponential and stationary phase in defined media (2% glucose) or Sgal (2% galactose). Cell were lysed and analyzed by western blot using anti-V5 antibody.

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A.4 Concluding remarks

The western blot analysis did not show significant changes in phosphorylation pattern of Gpt2p-wt and 3A in the absence of studied kinases. Psk1p mutant showed differences in the abundance levels of Gpt2p but they were not consistent in three independent experiments. Growth behaviour of phosphomutant strains in the presence of

Rapamycin (inhibitor of TORC1) did not exhibit growth deficiency suggesting that

TORC1 probably is not a possible kinase of Gpt2p. According to the list of phosphorylation sites targeted by different kinases (chapter IV, table 4.2), CDK1 which is reported the most can be a good candidate to be examined. Our results show that there is a delay in TAG lipolysis which may lead to a delay in the budding rate after growth resumption in Gpt2p lacking phosphorylation. CDK1 which is the central regulator of cell cycle progression

(Diril et al., 2012) might regulate Gpt2p through phosphorylation responding to the growth phases. Future directions are to optimize the expression and purification protocols in order to continue investigating protein-protein interactions mediated by the C-terminus. Protein- protein interactions can be further studied by performing affinity chromatography or co- immunoprecipitations followed by mass spectrometry for protein identification. To identify kinases responsible for phosphorylation of the C-terminus, a scaled-up quantity of purified peptide can be prepared for use in kinase assays.

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Appendix B - Copy right permissions

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