The Role of the B-Type Phospholipases in S. Cerevisiae: Function, Regulation, and Physiological Relevance in Lipid Homeostasis Beth a Surlow

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Summer 2014 The Role of the B-Type Phospholipases in S. cerevisiae: Function, Regulation, and Physiological Relevance in Lipid Homeostasis Beth A Surlow

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THE ROLE OF B-TYPE PHOSPHOLIPASES IN S. CEREVISIAE: FUNCTION,

REGULATION, AND PHYSIOLOGICAL RELEVANCE IN LIPID HOMEOSTASIS

A Dissertation

Submitted to the Bayer School of Natural and Environmental Sciences

Duquesne University

In partial fulfillment of the requirements for

the degree of Doctor of Philosophy

Beth A. Surlow

August 2014

Beth A. Surlow

2014

THE ROLE OF B-TYPE PHOSPHOLIPASES IN S. CEREVISIAE: FUNCTION,

REGULATION, AND PHYSIOLOGICAL RELEVANCE IN LIPID HOMEOSTASIS

Beth A. Surlow

Approved June 5, 2014

______Dr. Jana Patton-Vogt Dr. Philip Auron Associate Professor of Biological Professor of Biological Sciences Sciences (Committee Member) (Committee Chair)

______Dr. Joseph McCormick Dr. Jeffrey Brodsky Chair and Associate Professor of Professor of Biological Sciences Biological Sciences University of Pittsburgh (Committee Member) (Committee Member)

______Dr. Philip Reeder Dean, Bayer School of Natural and Environmental Science

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ABSTRACT

THE ROLE OF B-TYPE PHOSPHOLIPASES IN S. CEREVISIAE: FUNCTION,

REGULATION, AND PHYSIOLOGICAL RELEVANCE IN LIPID HOMEOSTASIS

Beth A. Surlow

August 2014

Dissertation supervised by Dr. Jana Patton-Vogt.

Membrane phospholipid synthesis and turnover is a continual process during normal cell growth. The turnover of the glycerophospholipids by B-type phospholipases (PLBs) in

Saccharomyces cerevisiae results in the formation glycerophosphodiesters through a deacylation reaction. Here, I address several aspects of the glycerophospholipid deacylation, transport, and reutilization pathway in S. cerevisiae. First, I show the RAS GTPase-activating proteins, Ira1 and

Ira2, are required for utilization of the glycerophosphodiester – glycerophosphoinositol (GroPIns)

– as a phosphate source. Second, I demonstrate loss of the cell surface associated PLBs, Plb1-3, and/or utilization of GroPIns causes actin cytoskeleton defects and an increased cell size. For the third and major part of my dissertation, I identified a novel interaction between Ypk1 and Plb1.

Ypk1, the yeast homolog of the human serum- and glucocorticoid-induced kinase (Sgk1), affects diverse cellular activities, including sphingolipid homeostasis. Here, I report that Ypk1 also

impacts the turnover of the major phospholipid, phosphatidylcholine (PC). Pulse-chase radiolabeling reveals that a ypk1∆ mutant exhibits increased Plb1-mediated PC deacylation and glycerophosphocholine (GroPCho) production compared to wild type. Consistent with a link between Ypk1 and Plb1, the levels of both Plb1 protein and PLB1 message are elevated in a ypk1∆ strain compared to WT yeast. Furthermore, I discovered that an increase in PLB1 expression also occurs upon disruption to sphingolipid synthesis and is mediated by the Crz1 transcription factor.

Taken together, these findings suggest that sphingolipid synthesis is coordinated with PC turnover to maintain optimal lipid homeostasis.

ACKNOWLEDGEMENT

Thank you to my advisor Dr. Jana Patton-Vogt for giving me the opportunity to work on this project, and for her continual guidance, advice, and support of both my research, and teaching endeavors. She has guided my development as a scientist in basic lab skills, scientific communication, and in problem solving/reasoning. Thank you to my outside committee member,

Dr. Jeff Brodsky, for his excellent support and advice, and for allowing me to learn a few techniques in collaboration with his lab. I also want to thank my committee members, Dr. Auron and Dr. McCormick for all of their helpful suggestions and for their time spent both during and outside my committee meetings assisting in various aspects of this project.

I want to acknowledge all of the members of the Patton-Vogt laboratory, past and present, for helpful conversations, both scientific and personal, during the time we shared together in the lab, including Andrew Bishop, Kellie Rosiek, Tao Sun, Nicole Nesbitt, Sanket Anaokar, and all of the undergraduates I have worked with, especially thank you to Ben Cooley and Carole Wolfe who have processed many samples and worked independently in the lab. I am especially grateful for Andy, who helped train me initially when I joined the lab, and who went through most of the grad school experience with me. I truly appreciate the working relationship and friendship.

I am grateful for all the friendships and colleagues I have made with my fellow graduate students. Thank you especially to Tiffaney Czapski, as we worked through many of the rough patches together. I appreciate all of the scientific and social adventures with all of you, Andy, Tiff,

Kellie, Sarah, Metis, Natasha, Maria, Sumedha, Chris, Allen, Stephanie, Andre, Heather, Nick,

AnJey, Juraj, and Ruth.

A special thank you to all of my family who encouraged me in continuing this journey without giving up, especially my parents, in-laws, and siblings – Mom, Dad, Caren, John, Marie,

Vince, Sarah, and Abby. I truly appreciate all of your help especially the last two years since Heidi was born. You have each graciously volunteered to help with babysitting, laundry, cooking, etc. exactly when the extra pair of hands was most needed, and have of course provided much emotional support. A special thank you also to my grandma and late grandpa who have said countless prayers for me. I am very fortunate and thankful to have a wonderful, supportive family!

Most importantly, I want to thank my husband, Will. Your daily support over the last six plus years, and your patience, understanding, and love through all the ups and downs is greatly appreciated. Your encouragement to keep going was my motivation. Thank you for being both the dad and the “mom” for Heidi at times, especially during the past few months. And lastly, thank you to my daughter, Heidi. I can always depend on you to turn my frown upside down. Mommy is “all done” with school and I will be spending more time with you and daddy soon!

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TABLE OF CONTENTS

Page

Abstract ...... iv

Acknowledgement ...... vi

List of Tables ...... xii

List of Figures ...... xiii

List of Abbreviations ...... xv

Chapter 1 Introduction ...... 1 1.1 Saccharomyces cerevisiae as a model organism ...... 1 1.2 Major lipid classes in the S. cerevisiae plasma membrane ...... 1 1.3 Glycerophospholipid metabolism in yeast ...... 4 1.4 Role and classification of phospholipases...... 9 1.5 Glycerophosphodiester production and transport in S. cerevisiae and higher eukaryotes ...... 12 1.6 The B-type Phospholipases in S. cerevisiae ...... 15 1.7 Glycerophosphodiesterase hydrolysis of the glycerophosphodiesters ...... 17 1.8 Summary of Background and Significance...... 17

Chapter 2 Neurofibromin Homologs Ira1 and Ira2 affect glycerophosphoinositol Production and Transport in Saccharomyces cerevisiae ...... 19 2.1 Introduction ...... 20 2.2 Results and Discussion ...... 21 2.2.1 Identification of IRA genes as affecting GroPIns metabolism...... 21 2.2.2 An ira1∆ira2∆ mutant exhibits altered GroPIns transport activity...... 23 2.2.3 Deletion of IRA1 or IRA2 increases the extracellular accumulation of GroPIns...... 24 2.3 Supplement to Chapter 2 ...... 27 2.3.1 Experimental Procedures ...... 27 2.3.2 Results and Discussion...... 29 2.3.3 Conclusions ...... 33

Chapter 3 Examination of the roles and regulation of the PLBs and Git1, including examination of phenotypes associated with altered dosage of these genes ...... 35 3.1 Introduction ...... 35 3.1.1 The Git1 glycerophosphodiester transporter ...... 36

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3.1.2 The S. cerevisiae B-type phospholipases ...... 36 3.1.3 Potential ability of the PLBs to reacylate GPXs to produce lysoglycerophospholipids or glycerophospholipids ...... 37 3.1.4 Morphology and cytoskeletal structure of S. cerevisiae cells ...... 38 3.1.5 Potential functional interactions between the PLB genes and other genes implicated in the maintenance of cellular morphology ...... 39 3.2 Experimental Procedures ...... 40 3.3 Results ...... 47 3.3.1 Cell morphology is altered upon loss of the Git1 transporter and in WT cells utilizing GroPIns as a sole phosphate source ...... 47 3.3.2 Growth of the PLB mutants utilizing GroPIns as a phosphate source...... 51 3.3.3 Deletion of one or more PLB genes does not alter growth rate ...... 54 3.3.4 The plb123∆ strain has increased sensitivity to the actin destabilizer, Latrunculin A ...... 55 3.3.5 Phenotypic analysis of WT and strains bearing deletion of the PLB genes upon exposure to stressors targeting the plasma membrane and the cytoskeleton ...... 57 3.3.6 Role of phosphate and growth phase in PLB expression ...... 57 3.3.7 Lethality in the absence of PI synthase cannot be rescued by the presence of GroPIns in the medium ...... 60 3.3.8 The functional Interaction between the PLBs and other genes, specifically Ypk1 and Lge1 .... 63 3.4 Discussion ...... 64

Chapter 4 Accelerated Plb1-mediated turnover of phosphatidylcholine upon loss of Ypk1 in S. cerevisiae ...... 68 4.1 Introduction ...... 69 4.2 Experimental Procedures ...... 72 4.3 Results ...... 81 4.3.1 Plb1 mediated extracellular GroPCho production increases upon loss of YPK1...... 81 4.3.2 Turnover of another glycerophospholipid, phosphatidylinositol, is unaffected by deletion of YPK1...... 83 4.3.3 PC Synthesis via the Kennedy pathway is unaffected by loss of YPK1...... 85 4.3.4 The secreted and cell associated forms of Plb1 increase in a ypk1∆ strain...... 86 4.3.5 Compromised sphingolipid synthesis upregulates PLB1 expression and increases PC turnover...... 89 4.3.6 Crz1 mediates PLB1 upregulation either when sphingolipid biosynthesis is compromised or when Ypk1 is absent...... 91 4.3.7 Ameliorating disrupted sphingolipid synthesis by deletion of ORM1 and ORM2 in a ypk1∆ strain partially rescues PLB1 expression to WT levels...... 91 4.3.8 Overexpression of only PLB1 rescues ypk1tsykr2∆ lethality at the restrictive temperature...... 93 ix

4.3.9 Deletion of both PLB1 and YPK1 leads to synthetic interactions...... 94 4.4 Discussion ...... 96

Chapter 5 Summary and Future Directions ...... 100 5.1 Summary of results ...... 100 5.2 How does alteration in sphingolipid biosynthesis result in activation of Crz1? ...... 100 5.3 Are other genes in phospholipid synthesis affected by disrupted sphingolipid synthesis? ...... 101 5.4 Does Plb1 play a role in Slm1 relocalization from eisosomes to MCT domains leading to TORC2 and Ypk1 activation? ...... 102 5.5 Is the plasma membrane lipid composition altered in YPK1/PLB1 deficient cells? ...... 103 5.6 Is the functionality of model plasma membrane proteins, 1,3-β-glucan synthase, Git1, Pma1, and Can1, altered in YPK1/PLB1 deficient cells? ...... 103 5.7 Do the B-type phospholipases play a role in deacylation of the phosphorylated phosphoinositides in vivo? ...... 104 5.8 Final conclusions about the importance of PLBs ...... 105 References ...... 106

Appendix A ...... 121

A 1. Genes with predicted or observed phospholipase activity in S. cerevisiae...... 122 A 2. Loss of the cell surface associated PLBs results in invaginations of the cell membrane when grown in media containing GroPIns as the sole phosphate source...... 123 A 3. Exposure to LatA alters distribution of actin patches and cables upon loss of PLB1-3. .... 124 A 4. Alterations in PLB message levels in log phase compared to stationary phase...... 125 A 5. Deletion of LGE1 and PLB3 results in slowed growth and decreased colony size...... 125 A 6. Protein Architecture upon construction of the PLB1-I-3HA allele...... 126 A 7. Plb1 mediated external glycerophosphocholine production is increased upon deletion of YPK1 at multiple time points in the BY4742 background...... 127 A 8. Plb1 mediated external glycerophosphocholine production is increased upon deletion of YPK1 at multiple time points in the YPH499 background...... 128 A 9. Kennedy pathway synthesis of phosphatidylcholine is not altered upon loss of Ypk1...... 129 A 10. Deletion of the flippase activators, Fpk1 or Fpk2, or deletion of LSP1 or PIL1, involved in eisosome formation do not alter extracellular GroPCho production...... 129 A 11. Overexpression of the plasma membrane flippase, DNF1, does not result in altered extracellular GroPCho production...... 130 A 12. Secreted and cell-surface associated Plb1 protein abundance in log and stationary phase is elevated in a ypk1∆ mutant...... 131 A 13. Regulation of PLB1, PLB2, and PLB3 message levels upon loss of Ypk1 or upon loss of transcription factors, Crz1, Msn2, and Msn4...... 131 A 14. Compromised sphingolipid synthesis by the presence of myriocin results in increased extracellular GroPCho production...... 132 A 15. Overexpression of the PLB genes on a multicopy plasmid elevates corresponding PLB message levels...... 133 x

A 16. The slowed growth phenotype and sensitivity to the cell wall perturbing agent, calcofluor white, for a ypk1∆ strain is exacerbated by deletion of PLB1...... 133 A 17. 1,3-β glucan synthase activity is depressed upon deletion of PLB1 and YPK1...... 134 A 18. Altered expression of the catalytic subunit of 1,3-β glucan synthase, Fks2...... 135 A 19. Activity of the plasma membrane protein, Git1, is depressed upon loss of Ypk1 and/or Plb1...... 135 A 20. Pma1 activity is not lost by deletion of PLB1 and/or YPK1...... 136 A 21. Sensitivity to canavanine is not altered upon YPK1 and/or PLB1 deletion...... 136

LIST OF TABLES

Table 2.1 S. cerevisiae strains used in this study (Chapter 2)...... 22 Table 3.1 S. cerevisiae strains used in this study (Chapter 3)...... 42 Table 3.2 Nucleotide sequences of primers used in this study (Chapter 3) ...... 43 Table 3.3 Chemical cell stressors evaluated and their targets ...... 57 Table 4.1 S. cerevisiae strains used in this study (Chapter 4)...... 74 Table 4.2 Nucleotide sequences of primers used for gene deletions in this study (Chapter 4) .... 74 Table 4.3 Nucleotide sequences of primers used for qRT-PCR ...... 78 Table 4.4 Cell Associated Counts from Figure 4.1 ...... 85

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LIST OF FIGURES

Chapter 1

Figure 1-1. Structures of the major lipid classes in the S. cerevisiae plasma membrane...... 2 Figure 1-2. Structures of the major plasma membrane glycerophospholipids...... 5 Figure 1-3. Synthesis pathways for the major phospholipids found in yeast...... 6 Figure 1-4. Phosphorylation sites of phosphatidylinositol...... 9 Figure 1-5. Phospholipase cleavage sites...... 11 Figure 1-6. Deacylation Reaction...... 13 Figure 1-7. Metabolic pathways for GroPIns and GroPCho in S. cerevisiae...... 15

Chapter 2

Figure 2-1. Ira1 and Ira2 are required for optimal growth of cells using GroPIns as a phosphate source...... 23 Figure 2-2. Ira1 and Ira2 affect GroPIns transport activity and GroPIns production...... 24 Figure 2-3 S. PLB1 and PLB3 message levels are elevated in the ira1ira2 double mutant but not in the single deletion strains...... 31 Figure 2-4 S. Deletion of IRA1 or IRA2 decreases INO1 promoter-driven ß-galactosidase activity...... 31 Figure 2-5 S. Expression of the hyperactive RAS2V19 allele results in inositol auxotrophy...... 32 Figure 2-6 S. GIT1 expression is decreased in WT and ira1∆ in media containing inositol...... 33

Chapter 3

Figure 3-1. Activities of the PLBs and hypothetical GroPIns acyltransferase activity...... 37 Figure 3-2. WT and git1∆/GIT1 diploid strains display minor differences in cell shape...... 48 Figure 3-3. WT cells display increased aggregation in the presence of GroPIns...... 49 Figure 3-4. WT cells have slowed growth when grown in media requiring utilization of GroPIns as a phosphate source at 37ºC...... 50 Figure 3-5. GroPIns can be used as a phosphate source upon loss of the cell surface associated PLBs, but results in aberrant morphology...... 52 Figure 3-6. Cells lacking single deletions of the PLB genes display some instances of morphological abnormalities...... 53 Figure 3-7. Loss of PLB 1, 2, and 3 does not alter growth compared to WT...... 55 Figure 3-8. Loss of multiple cell surface PLBs in the BY4741 background increases sensitivity to the actin destabilizer, Latrunculin A...... 56 Figure 3-9. Glycerophosphodiester transport via Git1, not production via Plb1-3 is regulated by phosphate availability...... 59 Figure 3-10. Known PI synthesis through Pis1 activity and hypothetical PI synthesis by GroPIns acyltransferase activity...... 60

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Chapter 4

Figure 4-1. Simplified schematic of the Ypk1 signaling pathway and downstream events...... 71 Figure 4-2. Plb1-mediated extracellular GroPCho production is increased upon deletion of YPK1...... 84 Figure 4-3. Synthesis of PC is not altered upon loss of Ypk1...... 86 Figure 4-4. Plb1 protein abundance and localization...... 88 Figure 4-5. PLB1 message levels are elevated by heat stress and upon disruption of sphingolipid homeostasis...... 90 Figure 4-6. Crz1 mediates PLB1 upregulation either when sphingolipid biosynthesis is compromised or when Ypk1 is absent...... 92 Figure 4-7. Deletion of ORM1 and ORM2 in a ypk1 strain partially reduces PLB1 expression. 92 Figure 4-8. Overexpression of only PLB1 rescues the lethality of the ypk1tsykr2 strain at restrictive temperature...... 93 Figure 4-9. The slowed growth phenotype and sensitivity to the cell wall perturbing agent, calcofluor white, in a ypk1 strain is exacerbated by deletion of PLB1...... 95 Figure 4-10. Model of PLB1 transcriptional activation upon loss of Ypk1 or upon disruption to sphingolipid biosynthesis by the presence of myriocin...... 99

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LIST OF ABBREVIATIONS

CDP-DAG Cytidine diphosphate diacylglycerol CDRE calcineurin dependent responsive element CL Cardiolipin DAG Diacylglycerol F-BAR (Bin–Amphiphysin–Rvs) GDTR Glycerophosphodiester transport and reutilization pathway GPC AT Glycerophosphocholine acyltransferase GPI glycosylphosphatidylinositol GPI AT Glycerophosphoinositol acyltransferase GPX(s) Glycerophosphodiester(s) GroPCho Glycerophosphocholine GroPIns Glycerophosphoinositol IP3 Inositol triphosphate IPC inositol phosphate ceramide Lat A Latrunculin A LSC Liquid scintillation counting M[IP]2C mannosyl-diinositolphosphate-ceramide MCC Membrane compartment containing Can1 MCP Membrane compartment containing Pma1 MCT Membrane compartment containing TORC2 MIPC mannosyl-inositolphosphate-ceramide OD(U) Optical density (unit) PA Phosphatidic Acid PC Phosphatidylcholine PDME Phosphatidyldiethanolamine PE Phosphatidylethanolamine PG Phosphatidylglycerol PH Pleckstrin homology PI phosphatidylinositol PIPs Phosphorylated phosphoinositides PKA Protein Kinase A PKC Protein Kinase C PLB B-type phospholipase PM Plasma membrane PMME Phosphatidylmonoethanolamine PS Phosphatidylserine PtdIns[4,5]P2 phosphatidylinositol-4,5-bis-phosphate PtdIns[4] phosphatidylinositol-4-phosphate ROS Reactive oxygen species S. cerevisiae Saccharomyces cerevisiae SGD Saccharomyces Genome Database SPOTS SPT, Orm1/2, Tsc1, Sac1 complex SPT Serine palmitoyl transferase TAG Triacylglycerol TORC2 Target of rapamycin 2 UASINO inositol response upstream activating sequence WT Wild type xv

Chapter 1 Introduction

1.1 Saccharomyces cerevisiae as a model organism

Saccharomyces cerevisiae (S. cerevisiae) the common budding yeast, is a well-studied model eukaryote. Yeast was first used as an experimental organism in the early twentieth century and has proven to be an asset in studying basic cellular processes (1). S. cerevisiae is a unicellular fungus in the phylum Ascomycota that can exist stably in culture as a haploid or diploid. The genome of S. cerevisiae has approximately 6,000 ORFs contained on 16 chromosomes. Several characteristics of yeast have made it ideal to use in research. For example, the environmental conditions can be controlled by using defined media, and altering the composition as needed. The generation time for yeast is about 90 minutes, which allows for quick mass production.

Additionally, many yeast strains containing drug resistance, auxotrophic markers, and specific mutations are readily available for use, including the yeast deletion set (2). A variety of other tools are available for yeast researchers, including those associated with the Saccharomyces Genome

Database (SGD), where there is catalogued information on each gene (http://yeastgenome.org/).

Many cellular functions of yeast, including many aspects of lipid metabolism, are conserved among higher eukaryotes, which make yeast a valuable model organism for studies of humans, animals, and plants.

1.2 Major lipid classes in the S. cerevisiae plasma membrane

The primary structural components of the S. cerevisiae cell membrane include glycerophospholipids, sphingolipids, and sterols, which are regulated by a variety of mechanisms in order to maintain optimal lipid homeostasis (3). A structural illustration of an example from each class of lipids is shown in Figure 1-1. The plasma membrane of the cell not only acts as a boundary, but also mediates transport of molecules and acts as a platform for signaling pathways

(4,5). In response to cellular stress, the regulatory networks of these lipids must coordinately adapt to maintain cell viability. Cells need to maintain the most favorable ratios of lipids in the plasma membrane, as well as in the membranes of the different organelles (6). The mechanisms for lipid trafficking between organelles, establishing and maintaining precise lipid compositions of specific compartments, and changing of lipid environments in response to environmental signals are not completely elucidated.

Glycerophospholipid (Phosphatidylcholine, PC)

Sphingolipid (Inositol phosphoceramide, IPC)

Sterol (Ergosterol)

Figure 1-1. Structures of the major lipid classes in the S. cerevisiae plasma membrane. Representative examples showing the basic structure of glycerophospholipids, sphingolipids, and sterols. The head groups are circled in red, with choline and inositol for the glycerophospholipid and sphingolipid respectively. Both the glycerophospholipid and sphingolipid have a phosphate group attached to the head group, and the respective backbone, glycerol and ceramide. Structures were generated using the LIPID MAPS structure database (7,8).

The major glycerolipids include the glycerophospholipids, – phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI) – phosphatidylglycerol (PG), and cardiolipin (CL). Other phospholipid intermediates found in the cell include phosphatidic acid (PA), diacylglycerol (DAG), triacylglycerol (TAG), CDP-

diacylglycerol (CDP-DAG), phosphatidylmonoethanloamine (PMME), phosphatidyldiethanloamine (PDME), phosphorylated phosphoinositides (PIPs), and lysophospholipids (LysoPL) (9). Structurally the glycerophospholipids are composed of a glycerol backbone with two fatty acids attached by ester linkages, a phosphate group, and various head groups. The primary fatty acids found in the glycerophospholipids are oleic, palmitoleic, palmitic and stearic acids (5,9). The fatty acid and head groups both affect lipid packing in the membrane, therefore influencing membrane fluidity. Significant progress has been made, and extensively reviewed, in elucidating the aspects of glycerophospholipid metabolism, including gene-enzyme relationships in biosynthetic pathways, localization of these processes, and regulation of synthesis and turnover (9). The glycerophospholipids are the primary focus of this dissertation.

The classification of sphingolipids includes sphingosines, ceramides, sphingomyelin, and phosphorylated or glycosylated derivatives of these metabolites (10,11). Sphingolipids have a basic structure containing a ceramide moiety attached to a fatty acid by an amide linkage. The three complex sphingolipids in S. cerevisiae are inositol phosphate ceramide (IPC), mannosyl- inositolphosphate-ceramide (MIPC), and mannosyl-diinositolphosphate-ceramide (M[IP]2C).

Greater than 90% of the cell’s complex sphingolipids are found in the plasma membrane (12). The regulation of sphingolipid synthesis has largely been uncharacterized until very recently. The Orm family of proteins were characterized and found to negatively regulate serine palmitoyl transferase, the enzyme catalyzing the rate limiting step in sphingolipid biosynthesis (13) (Figure 4-5).

Understanding of sphingolipid metabolism will be important in Chapter 4 of this dissertation.

In S. cerevisiae, ergosterol is the most abundant sterol in the plasma membrane, which serves a similar purpose to cholesterol, the main sterol lipid in higher eukaryotes. The sterols are necessary to maintain membrane fluidity, protein function, and organelle differentiation (14). The

research described here examined a potential connection to ergosterol metabolism, but was not pursued in detail.

Together with membrane proteins, these lipids – glycerophospholipids, sphingolipids, and ergosterol – make up the membranes of all cellular organelles, and the outer plasma membrane boundary of the cell, which is greatly enriched in the complex sphingolipids and ergosterol. Lipid asymmetry contributes to the physical properties of the membrane, with PE, PI, and PS enriched in the intracellular leaflet, and with PC and sphingolipids primarily present in the extracellular leaflet (5,15). Furthermore, enrichment of certain lipids and proteins into microdomains in the membrane confers regulated functionality of the membrane as a platform for signaling cascades

(4,16). Considering the lipid ratios determine the functional characteristics of the membrane, it is likely that crossover in regulation of lipid metabolism exists. Understanding the biosynthesis and turnover of the glycerophospholipids and sphingolipids, in particular, will be essential for the research described here.

1.3 Glycerophospholipid metabolism in yeast

S. cerevisiae has proven to be a powerful eukaryotic model for phospholipid metabolism since many synthesis pathways are similar to those in mammalian cells (9,17-19). Phospholipids are not only vital components of biological membranes, but many also act as signaling molecules or precursors of signaling molecules that impinge upon a variety of cellular processes such as cell growth and metabolism. Glycerophospholipids consist of two fatty acid chains bound via acyl bonds to the SN1 and SN2 hydroxyl groups of a glycerol backbone. The primary glycerophospholipids found in S. cerevisiae include phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylinositol (PI), and phosphatidylserine (PS), all are shown in Figure 1-2 (9,17). All of these major glycerophospholipids are derived from phosphatidic acid (PA) and can be synthesized via the CDP-DAG pathway (20). Alternatively, PC and PE can 4

be synthesized via the Kennedy pathway, primarily when choline and ethanolamine are supplemented in the media, although both pathways occur regardless of choline and ethanolamine availability (9,17). The basic biochemical pathways for the synthesis of the major phospholipids are summarized in Figure 1-3.

Figure 1-2. Structures of the major plasma membrane glycerophospholipids. The structure shown at the top displays a generic glycerophospholipid with stearic and palmitic acid acyl chains. The “X” can represent any of the head groups shown in the diagram to produce the corresponding glycerophospholipid.

Figure 1-3. Synthesis pathways for the major phospholipids found in yeast. The basic pathways for phospholipid synthesis and the gene products catalyzing the reactions are shown. The major lipid species are shown in red boxes with yellow text. The glycerophosphodiesters are shown in red boxes with blue text. The phosphorylated phosphoinositdes are shown in red boxes with black text. Gene names shown in red are regulated by inositol and contain the UASINO element in their promoters. Other gene products not regulated by inositol are shown in black. Metabolic intermediates are shown in black at the end of arrows. Dotted line indicates several steps in the pathway were omitted. Gluc-6-P, glucose-6- phosphate; Ins-1-P, inositol-1-phosphate; Gro-3-P, glycerol-3-phosphate; lyso-PA, lysophosphatidic acid; PA, phosphatidic acid; CDP, cytidine diphosphate; DAG, diacylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; PE, phosphatidylethanolamine; PMME, phosphatidylmonomethylethanolamine; PMDE, phosphatidyldimethylethanolamine; PC, phosphatidylcholine; Cho, choline; Cho-P, choline phosphate; Etn, ethanolamine; Etn-P, ethanolamine phosphate; PIP, phosphorylated phosphatidylinositol phospholipids; GroPCho, glycerophosphocholine; GroPIns, glycerophosphoinositol.

The synthesis of these phospholipids is regulated by genetic and biochemical factors at the transcriptional level, by mRNA stability, and post-translationally (9,17,20,21). Enzymatic activity is additionally regulated by lipids, phospholipid precursors and products, and by covalent modifications (20). Phospholipid biosynthetic genes are regulated by factors including nutrient

availability, growth stage, pH, temperature, and by message stability (9,17). The promoters of several genes in the CDP-DAG and Kennedy pathways contain the UASINO element (inositol response upstream activating sequence), which is regulated by the transcription factors Ino2, Ino4, and Opi1(20). The UASINO element, also known as the inositol/choline response element (ICRE), contains a consensus binding site, 5’CATGTGAAT3’ (22), for the Ino2/Ino4 heterodimer complex, which activates transcription of several genes involved in glycerophospholipid synthesis

(9). Repression of transcription occurs via Opi1 binding to Ino2, hence preventing the Ino2/Ino4 complex from binding to the UASINO elements. (23,24).

These transcription factors are regulated by a series of mechanisms including the presence or absence of inositol and choline (9). Several observations have supported the hypothesis proposed by Carman and Henry that the mechanism by which the cells sense inositol, choline, and other phospholipid precursors is through their effects on phospholipid synthesis (9). Furthermore, evidence shows PA is the signaling lipid controlling the UASINO co-regulated genes. PA co- regulates these genes by a mechanism in which Opi1 binds PA and Scs2, an integral ER membrane protein, at the ER/nuclear membrane thereby preventing Opi1 translocation into the nucleus (25).

The binding affinity of Opi1 to PA is strongly affected by cellular pH, and in turn the ionization state of PA, instead of the absolute abundance of PA (26). However, when PA levels are depressed, such as when inositol is added to the medium, Opi1 can enter the nucleus and interact with Ino2 to prevent transcription (17,23).

PA content and UASINO containing genes are controlled, in part, by inositol (27) and zinc

(17). Inositol availability causes an increase in PI synthesis due to excess substrate availability, which then results in depletion of the PA pool for synthesis of other phospholipids. Lower levels of PA allow Opi1 repression of UASINO containing genes encoding enzymes in the CDP-DAG and

Kennedy pathways causing an increase in PI and a decrease in PA, PS, and PC levels (20).

Depletion of zinc activates PI synthase resulting in decreased PA levels and repression of UASINO containing genes involved in PE synthesis (21). As regulators of phospholipid synthesis, inositol and zinc both regulate through PA levels. Briefly, PA levels are also regulated by the PA phosphatase encoded by PAH1 and by DAG kinase encoded by DGK1 (28). The activity of several phospholipid synthesis genes are additionally regulated by phosphorylation by PKA, PKC, and casein kinase II (23). While PA has an important role in regulating glycerophospholipid synthesis in S. cerevisiae, it is also implicated in multiple physiological processes in plant and animal cells, with PA formation occurring in response to stress, such as pathogen infection or drought.

One UASINO containing gene commonly evaluated to assess overall misregulation of the

UASINO containing genes is INO1 (24). INO1 encodes inositol-3-phosphate synthase, which converts glucose-6-phosphate to inositol-3-phosphate (29). INO1 is regulated by inositol and choline at the transcriptional level. In the absence of inositol and choline, Ino2/Ino4 activates transcription of INO1 through the UASINO element (24). Inositol auxotrophy and excess inositol excretion can indicate misregulation of INO1 and other UASINO containing genes (23). For example, ino2∆ and ino4∆ mutants are inositol auxotrophs, while opi1∆ mutants excrete excessive inositol into the media. Testing for inositol auxotrophy and excess inositol secretion is commonly employed to check for proper regulation of the phospholipid biosynthetic genes.

Synthesis of the primary glycerophospholipids mentioned thus far is clearly regulated at various levels. These glycerophospholipids have biological importance as is, but are also available for subsequent metabolism and modification. Multiple phosphorylated PI species (PIPs), highlighted in Figure 1-4, are also present in S. cerevisiae. These PIPs are generated by the action of kinases and/or phosphatases, which covalently modify three of the hydroxyl groups of the

inositol ring on PI. There are seven major PIPs derived from PI, each with a unique function and localization: phosphatidylinositol-4-phosphate (PtdIns[4]P), phosphatidylinositol-4,5-bisphosphate (PtdIns[4,5]P2), phosphatidylinositol-5-phosphate (PtdIns[5]P), phosphatidylinositol-3- phosphate (PtdIns[3]P), phosphatidylinositol-3,4-bis-phosphate (PtdIns[3,4]P2), phosphatidylinositol-3,5-bis-phosphate (PtdIns[3,5]P2), and phosphatidylinositol-3,4,5- triphosphate (PtdIns[3,4,5]P3) (30). The PIPs found specifically in S. cerevisiae include,

PtdIns[3]P, PtdIns[4]P, PtdIns[3,5]P2, and PtdIns[4,5]P2 (31). Additionally, PtdIns[3,4,5]P3 has been detected in the fission yeast Schizosaccharomyces pombe, but not in S. cerevisiae (31). The

PIPs each have distinct functions. The catabolism of PI and PC by the B-type phospholipase will be the major focus of the study, but great interest lies in further examination of potential PIP catabolism by the PLBs.

Figure 1-4. Phosphorylation sites of phosphatidylinositol. Phosphatidylinositol can be phosphorylated at three of the hydroxyl groups on the inositol ring to produce the phosphorylated phosphoinositides (PIPs). The hydrophilic inositol head group protrudes from biological membranes making the hydroxyls in positions 3, 4, and 5 accessible to kinases and phosphatases found at the membrane or in the cytosol.

1.4 Role and classification of phospholipases

During the course of normal cell growth, phospholipids are continually synthesized and degraded in a regulated fashion. The catabolic reactions occur through the action of phospholipases. Phospholipases are the enzymes that hydrolyze phospholipids to produce a variety of metabolites, some of which have signaling functions, and some of which are shuttled back into

phospholipid synthesis or other metabolic pathways. Phospholipases, which have been identified in both prokaryotes (32,33) and eukaryotes (34), are involved in a diverse range of processes including membrane homeostasis, nutrient acquisition, interaction with host cells, and signaling molecule production (35). In some organisms, phospholipases have a key role in virulence (36,37), and are found in venoms (38,39).

Phospholipases are generally classified as types A (1 or 2), B, C, or D, depending on where they cleave the phospholipids (Figure 1-5). The phospholipases A1 and A2 hydrolyze the SN1 and

SN2 fatty acid chains, respectively, producing a free fatty acid and a lysophospholipids

(phospholipids with only a single fatty acid). The B-type phospholipases (PLBs) have both A1 and

A2 activity meaning they can deacylate both the SN1 and SN2 acyl groups of glycerophospholipids with minimal accumulation of lysophospholipids. Phospholipase C cleaves just before the phosphate group producing diacylglycerol (DAG) and phosphate monoester (eg. inositolphosphate). Phospholipase D activity produces a free head group (eg. choline) and phosphatidic acid. Lysophospholipase and transacylase enzymes also assist in catabolism and modification of glycerophospholipids. Some fungal PLBs can have all three functions: phospholipase A1/A2 activity, lysophospholipase activity, and transacylase activity (35). Lysophospholipases deacylate lysophospholipids, while transacylases act by transferring the acyl group from one lysophospholipid to another. There are currently sixteen S. cerevisiae genes listed in the NCBI

Gene database with observed or putative phospholipase activity or domains (Appendix A 1).

PLA2 enzymes were the first phospholipases discovered and the superfamily has greatly expanded the classification into six types, that each consist of one or more groups and subgroups:

1) secreted PLA2 (sPLA2) including Groups I, II, III, V, IX, X, XI, XII, XIII, and XIV each having a His/Asp catalytic dyad, 2) cytoplasmic PLA2 (cPLA2) Group IV with a conserved catalytic

Ser/Asp and critical Arg in proximity of the catalytic center, 3) calcium independent PLA2 (iPLA2)

Group VI also having a Ser/Asp catalytic site, 4) platelet activating factor acetylhydrolyase (PAF-

AH) Groups VII and VIII which have a conserved catalytic Ser/His/Asp, 5) lysosomal PLA2

(LPLA2) Group XV with a Ser/His/Asp catalytic site, and 6) adipose PLA2 (AdPLA2) Group XVI containing a His/Cys catalytic site (40). The older classification system by the “Type” designation has become more ambiguous with the continual discovery of new PLA2 enzymes, and with new information obtained about the already identified PLA2s, such as the finding that most of the types are all found in the cytoplasm, and several of the groups outside of the iPLA2 Type are also independent of calcium. The group designations alone, which are based on the organism and sequence homology, more accurately classify the differences between the PLA2s (41,42). Various

PLA2 enzymes exist in bacteria, fungi, and mammalian cells, which are all included as part of the aforementioned classification system (35).

Figure 1-5. Phospholipase cleavage sites. Generic phospholipid shown with the cleavage sites for the different phospholipases. Only phospholipase B has the ability to hydrolyze both of the acyl groups.

Many fungi have multiple PLB genes – S. cerevisiae has four known PLB genes and C. albicans has five (35). In S. cerevisiae, the four genes encoding enzymes with phospholipase B activity are PLB1, PLB2, PLB3, and NTE1 (43). PLB1, PLB2, and PLB3 have >60% identity at the DNA level (44) and share a predicted catalytic center with conserved Arg/Ser/Asp (45). In yeast, phospholipases help regulate many processes, including, but not limited to, growth, nutritional/stress response, meiosis/mitosis cell cycle, and secretion (46).

1.5 Glycerophosphodiester production and transport in S. cerevisiae and higher eukaryotes

In S. cerevisiae, the deacylation of the glycerophospholipids via phospholipases of the B- type produce glycerophosphodiesters (GPXs) and free fatty acids (43). The B-type phospholipases

(PLBs) in S. cerevisiae are most similar to the mammalian cPLA2 Group IV phospholipases (47-

49). The deacylation of phosphatidylinositol (PI) and phosphatidylcholine (PC) in yeast produces the intracellular and extracellular glycerophosphodiesters, glycerophosphoinositol (GroPIns) and glycerophosphocholine (GroPCho), respectively (Figure 1-6). In mammalian cells, PLA1/2 activity followed by lysophospholipase activity will release arachadonic acid, lysophosphatidylinositol

(LysoPI), and the glycerophosphoinositols (49-51). Bacteria (52,53), mammals including humans

(54-57), and fungi (35,36) all possess enzymes with PLB activity. A human homologue of PLB was first identified in the epidermis, suggesting a function involving hydrolysis of lipids for use as components of the epidermal barrier (49). A PLB precursor has also been identified in human neutrophils, which has the ability to deacylate PC, PI, PE, and their lysophospholipids, suggesting a role in defense against microbes or involvement in inflammation (49,58).

Glycerophosphodiesters can also be formed by catabolism of the phosphorylated phosphoinositols (PIPs) to produce the collectively termed, glycerophosphoinositol glycerophosphodiesters (GroPIPs). The GroPIPs are water-soluble metabolites initially thought to be byproducts of PIP metabolism. However, evidence suggests these metabolites have their own 12

functions and metabolism (49,51). In mammalian cells, PLA2IVα produces GroPIns and

GroPI[4]P via its dual phospholipase and lysophospholipase activity on PI and PtdIns[4]P respectively (49). PLA2IVα activity is suggested to be regulated by calcium, phosphorylation,

PtdIns[4,5]P2 binding, and hormonal stimulation (40,49). Production of GroPIns and GroPI[4]P have been reported in many cell types upon different types of stimulation, such as by calcium or hormones (50). A few examples include GroPIns production upon stimulation of platelets, macrophages, neutrophils, thyroid cells, kidney slices, and brain slices. GroPI[4]P production has been observed in cell types such as fibroblasts, thyroid cells, and hepatocytes.

Figure 1-6. Deacylation Reaction. The glycerophospholipid, phosphatidylcholine (PC), is deacylated via the action of a B-type phospholipase enzyme to produce the glycerophosphodiester, glycerophosphocholine (GroPCho) and two free fatty acids.

The deacylation of the PIPs in yeast is also a regulated process that is influenced by several conditions. In S. cerevisiae, GroPIns, GroPI[4]P and GroPI[4,5]P2 release has been observed upon stimulation by glucose (59). Importantly, no studies have addressed the role of Plb1-3 in GroPI[4]P and GroPI[4,5]P2 release, although published literature states the PLBs also act on the PIPs to

produce GroPIPs (44,45). This extracellular release of the GroPIPs occurs in both yeast and mammalian cells (49,60). Upon release in mammalian cells, the GroPIPs may be further metabolized by extracellular phosphodiesterases, act as paracrine or autocrine signals, or can be transported back into the cell (49).

The glycerophosphodiesters are hydrophilic and cannot enter the cell without the action of a transporter. The glycerophosphodiester transporter, Git1, was first discovered in yeast when

GroPIns was observed to be secreted and taken back up (61). The human protein with the highest identity to Git1 was later identified as Glut2 (62). Glut2 mediates uptake of GroPIns and is primarily expressed in cells of the liver, kidney, and small intestine (49). GroPIns is able to get into cells not expressing Glut2, indicating there are likely other unidentified GroPIns transporters in mammalian cells (49).

In S. cerevisiae, both GroPIns and GroPCho can be transported into the cell via the permease encoded by GIT1. Once inside the cell, GroPIns and GroPCho are metabolized. In fact, exogenous GroPIns and GroPCho can individually be provided as the sole phosphate source for the cells (63,64). GroPCho is broken down into choline and glycerol-3-phosphate via the glycerophosphodiester phosphodiesterase, Gde1 (65,66). The enzyme facilitating the catabolism of GroPIns and the subsequent products is currently unknown. The biochemical pathways specific to GroPIns and GroPCho in S. cerevisiae are shown in Figure 1-7. The deacylated glycerophosphoinositide (GroPIPs) derivatives exit and enter mammalian cells as well (50,67,68).

Figure 1-7. Metabolic pathways for GroPIns and GroPCho in S. cerevisiae. GroPIns and GroPCho metabolism and transport are shown with the corresponding genes that encode the enzymes required for the reactions to take place. Git1 transports GroPIns and GroPCho into the cell.

1.6 The B-type Phospholipases in S. cerevisiae

Some information about substrate preferences and factors affecting each of the PLB gene products is available through gene deletion and overexpression studies. Plb1 primarily deacylates

PC producing GroPCho, and less robustly produces GroPEtn from PE (69). Plb2 is responsible for exogenous phospholipid hydrolysis (45). Plb3, which only uses PtdIns and PtdSer as substrates, is thought to be mainly responsible for extracellular GroPIns production with Plb1 having some additional influence (45). Plb1, Plb2, and Plb3 have phospholipase B, lysophospholipase, and lysophospholipid acyltransferase (LPAT) activities (45,46,69). The other enzyme with phospholipase B activity in S. cerevisiae, Nte1, controls internal GroPCho production and has been

suggested to be localized to the endoplasmic reticulum (70). At a lower efficiency Plb1 and Plb2 can hydrolyze PA and PtdIns[4,5]P2 (44).

Biochemical studies have predicted Plb1 and Plb3 localization at the plasma membrane and periplasmic space in stationary phase S. cerevisiae cells (43,45,69). In addition to localization in the plasma membrane and periplasmic space, Plb2 is predicted to be found in the supernatant

(43,45). However, large-scale microscopic datasets employing fluorescent tags fused to the C- terminal end and expressed under control of their endogenous promoters, YPL+ (71,72) and

YeastGFP (73-75), and a study utilizing C-terminal fusions to an epitope tag with the target genes expressed under control of the GAL promoter, Organelle DB (76), contain no data or inconclusive data for Plb1, Plb2, and Plb3 localization, as reported on SGD. In addition, a localization screen of proteins involved in lipid metabolism, in which Plb1 was C-terminally tagged and expressed under the TEF promoter, showed ER and vesicle localization, but no PM localization, and the medium was not analyzed (77). Furthermore, Plb1 and Plb3 have predicted glycosylphosphatidylinositol (GPI) – anchor attachment sites (78). Interestingly, recent evidence in addition to the data presented in Chapter 4, suggests Plb1 may have an intracellular role.

Analysis of the enzyme kinetics for the B-type phospholipases, and all phospholipases, is complex given that the enzymes act on individual lipids that are aggregated together to form micelles, vesicles, and membrane bilayers at the lipid-water interface (40). Frequently, a “surface dilution kinetic model” is employed to study many enzymes in lipid metabolism in vitro, in which activity is measured in the presence of cell-free mixed micelles containing lipids and a detergent

(79). The affinity for different substrates of the B-type phospholipases in vivo mentioned above is different from the primary substrates observed in vitro (44). Furthermore, in vitro experiments have shown pH, and the presence or absence of various cations, Ca+2, Mg+2, Fe+3, and Al+3, affect

the activity of Plb1, Plb2, and Plb3 (44). Of particular interest, at pH = 2.5 the presence of any of the listed cations caused decreased activity of Plb1, but at pH = 5.5 Plb1 activity was increased by the presence of the cations. In vivo Plb1-mediated GroPCho production increased with increasing pH from 3.5 to 7.5, contrary to the in vitro observations that increasing pH resulted in decreased

Plb1 activity (44). Importantly, Plb1 activity can clearly be affected by pH and cations.

1.7 Glycerophosphodiesterase hydrolysis of the glycerophosphodiesters

A number of glycerophosphodiesterases (GDEs) that catalyze the breakdown of the glycerophosphodiesters have been characterized in bacteria, fungi, plants, and animals (51). In S. cerevisiae, the glycerophosphodiesterase Gde1 hydrolyzes GroPCho to choline and glycerol-3- phosphate, but the enzyme facilitating breakdown of GroPIns is unknown (65,66). In mammalian cells, seven different GDEs (GDE1 – 7) have been identified each with specific substrate specificity, cellular localization, and tissue specificity (51). The mammalian GDE enzymes are relatively newly characterized proteins, and unlike bacterial GDE enzymes, can hydrolyze substrates other than the glycerophosphodiesters, including GPI anchors (51). Both mammalian

Gde1 and Gde3 prefer GroPIns over GroPCho, but they hydrolyze different bonds. Gde1 activity produces inositol and glycerol phosphate, while Gde3 acts like a PLC, producing glycerol and inositol-1-P (49,51). All but one of the mammalian GDEs are predicted to be membrane bound, so the GPXs would need to reach the plasma membrane for further catabolism (49) supporting the role of PLB or PLA activity at the plasma membrane.

1.8 Summary of Background and Significance

The PLBs deacylate phospholipids, thereby changing the phospholipid composition of the cell membrane. Several studies have shown that certain signaling pathways, protein activities, and cytoskeletal elements require specific membrane phospholipid composition for proper functioning.

In addition, the PLB products, the glycerophosphodiesters, have been shown to elicit physiological

effects in mammalian cells, including those involved in cytoskeletal function and cell morphology.

A more thorough examination of PLB function and regulation in the yeast S. cerevisiae, as described here, provides novel evidence for how phospholipase B activity impacts cellular physiology.

The overall goal of my project was to further elucidate the physiological functions and regulation of the B-type phospholipases (PLBs). My central hypothesis was that the PLBs are important for membrane homeostasis and cellular functions under certain environmental conditions. The specific objectives of my work as detailed in the following three chapters were as follows: 1) Determine how alterations in the RAS/cAMP signal transduction pathway affect PLB1,

PLB2, and PLB3 gene response, 2) Examine phenotypes associated with deletion of one or more of the PLB genes and identify potential genetic interactions that could contribute to the elucidation of the PLB function, and 3) Elucidate the mechanism by which Ypk1 regulates Plb1 activity.

Objectives 1 and 2 were initiated for the intended completion of the M.S. degree, but instead led into my transition to the Ph.D. program, at which time I began the major project for my Ph.D. work, Objective 3. Each of these objectives will be discussed individually in Chapters 2, 3, and 4 respectively, with Chapter 4 serving as the majority of my Ph.D. work. Together, the data provided here furthers our understanding of the role and regulation of the PLBs.

Chapter 2 Neurofibromin Homologs Ira1 and Ira2 affect glycerophosphoinositol Production and Transport in Saccharomyces cerevisiae

Running Title: Glycerophosphoinositol metabolism

Reprinted from Eukaryotic Cell, Vol. 8 No. 12, pages 1808-1811. Bishop, A. C.*, Surlow, B.A.*, Anand, P., Hofer, K., Henkel, M., Patton-Vogt, J. "Neurofibromin homologs Ira1 and Ira2 affect glycerophosphoinositol production and transport in Saccharomyces cerevisiae." Copyright (2009) with permission from the American Society of Microbiology.

*A.C. Bishop and B.A. Surlow contributed equally to this work.

Chapter 3 Attributions: Beth Surlow, I completed the growth studies shown in Figure 1 and the real time qRT-PCR of the PLB genes in the IRA mutant strains (data not shown in publication). Additional experiments I completed pertaining to this project, but excluded from final submission of the publication are included in the Supplement to Chapter 2.

Andrew Bishop, a graduate student in the Patton Vogt lab, completed the GroPIns transport and production experiments shown in Figure 2.

Puneet Anand, a Masters student in the Patton-Vogt lab, performed the initial screen of the yeast knockout collection to identify strains bearing compromised growth when given GroPIns as the sole phosphate source.

Katherine Hofer, an undergraduate in the Patton Vogt lab performed anion exchange chromatography for separation of inositol containing compounds in Figure 2.

Abstract Saccharomyces cerevisiae produces extracellular glycerophosphoinositol through phospholipase-mediated turnover of phosphatidylinositol and transports glycerophosphoinositol into the cell upon nutrient limitation. A screening identified the RAS GTPase-activating proteins

Ira1 and Ira2 as required for utilization of glycerophosphoinositol as the sole phosphate source, but the RAS/cyclic AMP pathway does not appear to be involved in the growth phenotype. Ira1 and Ira2 affect both the production and transport of glycerophosphoinositol.

2.1 Introduction

Membrane phospholipids are continually synthesized and degraded as cells grow and respond to environmental conditions. A major pathway of phosphatidylinositol (PI) turnover in

Saccharomyces cerevisiae is its deacylation to produce extracellular glycerophosphoinositol

(GroPIns) (60). Plb3, an enzyme with phospholipase B (PLB)/lysophospholipase activity, is thought to be primarily responsible for the production of extracellular GroPIns, with Plb1 playing a lesser role (44,45,69). GroPIns is transported into the cell by the Git1 permease (61). GIT1 expression is upregulated by phosphate limitation and inositol limitation. In fact, GroPIns can act as the cell’s sole source of both inositol (61) and phosphate (63).

A screening for gene products involved in the process by which GroPIns enters the cellular metabolism identified Ira1 and Ira2, yeast homologs of the mammalian protein neurofibromin.

Alterations in NF1, the gene encoding neurofibromin, are associated with the pathogenesis of neurofibromatosis type 1, an autosomal dominant genetic disease (80-82). Ira1 and Ira2 and neurofibromin function as RAS GTPase-activating proteins (RAS GAPs). S. cerevisiae Ras1 and

Ras2 activate adenylate cyclase to modulate cyclic AMP (cAMP) levels. The binding of cAMP to the regulatory subunits of protein kinase A (Bcy1) results in dissociation and activation of the catalytic subunits (Tpk1 to Tpk3). Ira1 and Ira2 inactivate RAS and thereby downregulate the pathway (83,84). Hydrolysis of cAMP by the phosphodiesterases encoded by PDE1 and PDE2 also downregulates the pathway (85-87). The RAS/cAMP pathway responds to nutrient signals to modulate fundamental cellular processes, including stress resistance, metabolism, and cell proliferation (85,87,88).

2.2 Results and Discussion

2.2.1 Identification of IRA genes as affecting GroPIns metabolism.

To identify genes involved in the metabolic process by which GroPIns is used as a phosphate source, we screened the MATα viable yeast knockout collection (Research Genetics) for strains displaying compromised growth when GroPIns rather than low Pi was supplied as the phosphate source. Synthetic complete medium, high-Pi (10 mM KH2PO4) medium, low-Pi (0.2 mM KH2PO4) medium, and GroPIns+ (70 µM) medium were made as described previously (64).

Mutants were transferred by hand pinner from a master plate to a 96-well plate containing low-Pi medium and allowed to grow at 30°C for 3 days. From low-Pi medium, cell inocula were transferred to plates containing GroPIns+ medium. Growth was monitored at 595 nm after 4 days of incubation at 30°C. The ratio of absorbance in low-Pi medium to absorbance in GroPIns+ medium was determined. This screening was performed twice. Mutants with a value of 3 or greater for the ratio of absorbance in low-Pi medium to absorbance in GroPIns+ medium were subjected to a second screening by the monitoring of their growth in test tubes. As expected, the GIT1 gene was required for growth, and no other mutant displayed a complete growth abatement phenotype

(Figure 2-1 A). In particular, we did not identify a gene likely to encode a glycerophosphodiesterase responsible for GroPIns catabolism. A likely explanation for this result is that multiple gene products are involved in the mechanism(s) by which GroPIns enters cellular metabolism. However, a number of strains exhibited a slow-growth phenotype, one of which was the ira2∆ mutant (Figure 2-1 A). To confirm the validity of this result, we analyzed growth on

GroPIns in strains of the unrelated ∑1278b background (gifts from M. Cardenas, Duke

University), and found that not only the ira2∆ mutant but also the ira1∆ mutant and the ira1∆ira2∆ double mutant (Table 1) exhibited greatly reduced growth on GroPIns (Figure 2-1 B). The ira1∆ mutant was not present in the deletion set screened initially. 21

Table 2.1 S. cerevisiae strains used in this study (Chapter 2)

To probe the role of the RAS/cAMP in GroPIns metabolism, we analyzed other strains bearing alterations in the pathway. Interestingly, a pde2∆ mutant that, like the ira2∆ mutant, exhibits increased cAMP levels (89) was not defective for growth on GroPIns. Similarly, a strain bearing a plasmid-borne hyperactive allele of RAS2, RAS2-V19 (a gift from P. Herman, Ohio State

University) (90-92), was not defective for growth on GroPIns (Figure 2-1 A). Thus, upregulation of the RAS/cAMP pathway may not be the primary reason for the growth defect seen in the absence of Ira1 or Ira2. Not surprisingly, ras1∆ and ras2∆ mutants, expected to downregulate the

RAS/cAMP pathway, were not defective for growth on GroPIns (data not shown).

A trivial explanation for the inability of strains bearing mutations in the IRA genes to utilize

GroPIns as a phosphate source is that the cells are less robust than WT cells and die before they are able to induce the metabolism required to utilize GroPIns. To test this, we compared the survival of the WT and an ira1∆ira2∆ mutant upon phosphate starvation and found no difference between the two strains (data not shown).

Figure 2-1. Ira1 and Ira2 are required for optimal growth of cells using GroPIns as a phosphate source. Strains in the BY4742 (A) or ∑1278b (B) background were grown overnight in low-Pi medium. Cells were transferred to low-Pi or GroPIns+ medium (A600 ≈ 0.005). Absorbance was recorded after 48 h of growth at 30°C. WT cells transformed with empty vector pRS415 (vector) or plasmid pPHY453, containing the hyperactive RAS2 allele, RAS2-Val19 (RAS2V19), were also assayed (A). Readings are means ± standard errors of the means of results of at least two independent experiments.

2.2.2An ira1∆ira2∆ mutant exhibits altered GroPIns transport activity.

To determine the cause of the defect in GroPIns utilization that occurs upon deletion of

IRA1 or IRA2, we measured GroPIns transport activity (63) under low-Pi conditions. In cells harboring single-deletion mutations, GroPIns transport was similar to that of the WT strain or only slightly reduced (Figure 2-2 A). However, the simultaneous deletion of IRA1 and IRA2 resulted in a clear reduction in activity. These results do not explain the growth defects seen in the single- deletion mutants, but they do indicate a role for Ira1 or Ira2 in regulating Git1. A complete understanding of the nature of the regulation of GroPIns transport by Ira1 or Ira2 will be the focus of future studies.

Figure 2-2. Ira1 and Ira2 affect GroPIns transport activity and GroPIns production. (A) Cells grown to log phase in low-Pi medium were assayed for GroPIns transport activity. Data represent means ± standard errors of the means of results of two independent experiments performed in duplicate. (B) Cells pre-labeled with [3H]-inositol were re-inoculated into fresh high-Pi medium, and the amount of extracellular GroPIns was determined following 22 h of growth. Data are means ± standard errors of the means of results of at least three independent experiments and are reported as the percentage of total [3H]- inositol label found as extracellular GroPIns normalized for culture density.

2.2.3 Deletion of IRA1 or IRA2 increases the extracellular accumulation of GroPIns.

We next monitored the effect of Ira1 or Ira2 upon extracellular production of GroPIns via

PI deacylation. Cells were grown overnight in medium containing 3 µCi/ml of [2-3H]-inositol.

Cells were harvested, washed, and reinoculated to an A600 of 0.2 in high-Pi medium, the condition under which extracellular GroPIns accumulates (61). At various times, aliquots of the cultures were centrifuged, and the inositol compounds in the supernatant were separated (93). The ira1∆, ira2∆, and ira1∆ira2∆ mutants accumulated roughly twice as much extracellular GroPIns as the

WT strain (Figure 2-2 B) upon entrance to stationary phase. The lack of an additive effect upon

GroPIns formation in the double mutant compared to formation in the single mutants may be explained by substrate availability. In other words, there may be a limited amount of plasma

membrane-associated PI available for PLB-mediated turnover. Whereas an ira1∆ira2∆ double mutant exhibits an increase in PLB1 and PLB3 transcript levels as measured by real-time reverse transcriptase PCR (data not shown, Supplemental Figure 2-3 S), no such increase was detectable in the ira1∆ or ira2∆ mutant. Thus, Ira1 or Ira2 regulation of the PLB activity leading to GroPIns formation likely occurs at multiple levels and will be the focus of future studies. Another explanation is that yet another, uncharacterized, phospholipase exists for the formation of GroPIns in the absence of Ira1 or Ira2. Importantly, GroPIns transport does not occur under the growth conditions of the experiment (high Pi), so diminished transport cannot explain the increased accumulation of GroPIns in the absence of Ira1 or Ira2.

In summary, this study indicates a role for Ira1 and Ira2 in regulation of both the production of extracellular GroPIns and its transport into the cell. In addition, we show that Ira1 and Ira2 are required for the utilization of GroPIns as a phosphate source under conditions in which transport of GroPIns into the cell is normal. However, we were unable to show a linkage between hyperactivation of the RAS/cAMP pathway, which occurs upon deletion of IRA1 or IRA2, and the growth phenotype. Future studies will focus on determining the mechanism(s) for the growth disturbance, including whether it involves the RAS/cAMP pathway or other, less defined functions of Ira and Ira2. Growth is a multifactorial process, and a combination of disturbances, some involving heightened signaling through the RAS/cAMP pathway and some not, may contribute to alterations in the cell’s ability to utilize GroPIns as a phosphate source. Ira1 and Ira2 are large proteins, each over 3,000 amino acids. The RAS GAP-related domain of each protein is contained within a few hundred amino acids, and C-terminal regions of approximately 200 amino acids bind to Gpb1 or Gpb2 and are important for protein stability (94). In addition, yeast Ira1 and Ira2 proteins and the human neurofibromin type 1 protein all contain bipartite phospholipid binding

modules consisting of a Sec14 homologous segment and a pleckstrin homology-like domain (95).

Both in vitro phospholipid binding and lipid exchange activity have been documented for neurofibromin (96), suggesting that Ira1 and Ira2 may have similar activities. Indeed, Ira1 appears to play a membrane-anchoring role for adenylate cyclase in addition to its role as a RAS GAP (97).

It is tempting to speculate that Ira1 or Ira2 might also play a role in the membrane association or activation of an enzyme or enzymes responsible for the metabolism of internalized GroPIns. A test of that hypothesis awaits the identification of the gene products involved in the process by which

GroPIns enters cellular metabolism.

This work was supported by National Institutes of Health grant GM59817 to J.P.-V. We thank M. Cardenas for the gift of strains and P. Herman for the gift of plasmids. We thank Lindsay

Chromiak and David Ekpin for technical assistance.

2.3 Supplement to Chapter 2

The experiments described here were completed as part of the project described in the

Chapter 2 manuscript, but were excluded from the publication. The purpose of these supplemental studies was to examine PLB1-3 and INO1 transcriptional expression in response to alterations in the Ras/cAMP pathway

2.3.1 Experimental Procedures

Strains, plasmids, and growth conditions. The S. cerevisiae strains used in this study are shown in Table 2.1. Strains were grown aerobically with shaking at 30ºC. Turbidity was monitored by measuring the optical density at 600nm (A600) on a BioMate 3 Thermo Spectronic spectrophotometer. Media used for this study included rich yeast extract peptone dextrose (YEPD) media, or synthetic complete yeast nitrogen base (YNB) media made as described previously, but containing 2% glucose (93). To control media phosphate concentration, KH2PO4 (1 g/L) was excluded from the synthetic mix and replaced with KCl (1 g/L). KH2PO4 was added back at a concentration of 10 mM (high) or 200 µM (low) as indicated. YNB containing 75 µM inositol (I+) or lacking inositol (I-) was used as noted.

The plasmids pRS416 (JVE288) and pJR1040-RASVal19 (JVE290) were kindly provided by

Paul Herman (90) (The Ohio State University). Plasmids pBH315-GIT1-lacZ (JVE294) and pJH359-INO1/CYC1/lacZ (98) were available or made previously in the lab. Transformation of plasmids into yeast strains was performed using standard lithium acetate chemical transformation.

β-galactosidase assays. Gene promoter activity was measured as a function of β-galactosidase

(β-gal) activity using either the Pierce β-galactosidase kit per the manufacturer’s instructions or using the lab β-gal protocol. Cells were grown to late log phase in YNB medium lacking the specified amino acid for selection, harvested, and an aliquot was assayed for β-gal activity. Briefly,

using the Pierce kit, 350 µL of working solution (50:50, 2X β-gal assay buffer: Y-PER) was added to 350 µL of culture to start the assay and incubated at room temperature for 6.5 minutes. β-gal stop solution (300 µL) was added, solution agitated with a Vortex mixer, and centrifuged for 30s at 13,000 x g. Supernatant absorbance was measured at 420 nm and used to calculate β-gal activity

([1000 x A420nm]/[time in minutes x Volume in mL x OD600 at harvest]). For the lab procedure, 10

ODUs (optical density units) of cells were harvested and resuspended by Vortex agitation in 150

µL of Z buffer, 50 µL chloroform, and 20 µL of 0.1% SDS. 0.7 mL of ONPG (1.2 mg/mL) was added to start the reaction and incubated at room temperature for 10 minutes. The reaction was stopped with 0.5 mL of 1M Na2CO3 and centrifuged for 2 minutes. The absorbance of the supernatant was measured at 420 nm and used to calculate β-gal activity (A420nm x 1000 x (1/time in minutes)).

RNA Extraction and real time qRT-PCR gene expression analysis. Cultures were grown in

YNB medium containing 75 µM inositol under the conditions specified in the text. RNA was extracted using a hot phenol-chloroform extraction (99). A 7 µg sample of RNA was DNase treated with 2 units of DNase at 37ºC for 30 minutes using the TURBO DNA-free™ Kit (Applied

Biosystems) per the manufacturer. Message abundance was quantified by quantitative reverse transcriptase PCR (qRT-PCR) using the Verso™ SYBR® Green 1-Step QRT-PCR ROX Kit

(Thermo Scientific) on the Applied Biosystems StepOnePlus Real Time PCR System. The primers used are shown in Table 3.2. Standard curves for each primer pair were optimized to determine acceptable primer concentration (70-100 nM) and template concentrations (15-25 ng/25

µL reaction). Reverse transcription was carried out at 50C for 15 min followed by 95C for 15 min for RT inactivation and polymerase activation. Amplification parameters were 40 cycles at

95C 15s, 54.5C 30s, and 72C 40s. Primer specificity was verified by melt curve analysis and

visualization of amplicons by gel electrophoresis. Primer sets did not produce amplicons when

RNA from the respective deletion strains was used as template. No template control (NTC) and minus reverse transcriptase (-RT) reactions were performed to verify there was no genomic DNA contamination in the RNA samples and/or the reagents. RNA from at least three independent cultures of all strains was analyzed in triplicate reactions per primer set. Data was normalized to the endogenous control gene, SNR17, and analyzed using the comparative ∆CT approach

(100,101).

2.3.2 Results and Discussion

An ira1∆ira2∆ mutant exhibits altered PLB1-3 transcript levels.

The B-type phospholipase encoded by PLB3 is primarily responsible for the production of

GroPIns via PI deacylation, with Plb1 having some contribution (44). To determine if transcriptional regulation of the PLB genes is responsible for the increase in GroPIns production observed upon deletion of IRA1 and IRA2, real-time qRT-PCR was performed. The transcript level of PLB2 was also monitored. Deletion of both IRA1 and IRA2 resulted in an increase in

PLB1 and PLB3 message (Figure 2-3 S). However, deletion of IRA1 or IRA2 individually had little or no effect on PLB1 transcript levels and actually decreased PLB3 transcript levels. Hence, these results do not support a model for increased PLB gene expression causing the increased GroPIns production occurring upon deletion of IRA2 (Figure 2-2 B), but do indicate that deletion of both

IRA1 and IRA2 affects PLB1 and PLB3 message levels. With regard to PLB2, there was a clear decrease in transcript levels in the ira1∆, ira2∆, and ira1∆ ira2∆ mutants (Figure 2-3 S).

INO1 expression is altered in the IRA1 and IRA2 mutants.

During the course of the transport experiments, we discovered that ira1∆ira2∆ mutants are inositol auxotrophs. Inositol auxotrophy is typically the result of misregulation of INO1, the gene

encoding inositol-3-phosphate synthase (Figure 1-3), which is the rate-limiting step in inositol biosynthesis (29). In a WT strain, INO1 is derepressed in cells grown in inositol-free medium, and repressed in cells grown in inositol containing medium. We hypothesized the inositol auxotrophy in the IRA mutants is due to misregulation of the INO1 gene. INO1 expression was monitored in response to phosphate and inositol concentrations by transforming strains with a reporter construct, in which the INO1 promoter is fused to lacZ (98). Because the ira1∆ira2∆ mutant cannot grow in the absence of inositol, INO1 expression was monitored in the presence of 10M inositol, a concentration that supports growth and only partially represses INO1 expression in a WT strain.

Deletion of either IRA1 or IRA2 reduces INO1 expression, and the deletion of both simultaneously increases the effect (Figure 2-4 S). Also noteworthy is that INO1 expression, in general, is reduced under low phosphate conditions. Importantly, the reduced INO1 expression of the ira1∆, ira2∆ and ira1∆ira2∆ double mutants cannot explain the growth defect observed when GroPIns was supplied as a phosphate source, as the growth experiments shown in Figure 2-1 were performed in the presence of inositol.

Upregulation of the Ras/cAMP pathway with the RasV19 allele results in inositol auxotrophy

Because prior experiments in the lab showed ira1,2∆ is an inositol auxotroph, and ira1∆, ira2∆, and ira1,2∆ have defective growth on GroPIns we hypothesized that upregulation of the

Ras/cAMP pathway by introduction of a constitutively active form of RAS, RasVal19, may have a similar affect (90). A strain bearing a hyperactive allele of RAS2, RAS2V19 (91) exhibited moderate inositol auxotrophy (Figure 2-5 S), suggesting that upregulation of the RAS/cAMP pathway is involved in mediating the inositol auxotrophy phenotype.

Figure 2-3 S. PLB1 and PLB3 message levels are elevated in the ira1ira2 double mutant but not in the single deletion strains. Relative message abundance of PLB1, PLB2, and PLB3 in WT (JPV 569), ira1∆ (JPV 573), ira2∆ (JPV 574), and ira1∆ira2∆ (JPV 575) (∑1278b background) grown in YNB highPi medium to log phase (OD600 ≈ 0.7) at 30ºC. Total RNA was extracted from cells and transcript levels were analyzed by real-time qRT- PCR using the comparative ∆CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT cells. Values represent the mean ± S.E. of at least 3 individual cultures each assayed in triplicate.

Figure 2-4 S. Deletion of IRA1 or IRA2 decreases INO1 promoter-driven ß-galactosidase activity. WT (JPV 467), ira1 (JPV 573), ira2 (JPV 574), and ira1,2 (JPV 575) were transformed with the plasmid pJH359-INO1/CYC1/lacZ (JVE3) and grown to late log phase (OD600 ≈ 0.8-1.0) in YNB –URA media with the indicated phosphate and inositol concentrations. Cells were harvested and assayed using the Pierce β-gal assay kit. Data represent the mean ± S.E. of three individual transformants in each strain assayed in duplicate.

Figure 2-5 S. Expression of the hyperactive RAS2V19 allele results in inositol auxotrophy. WT (JPV 569) transformed with empty vector (pRS416 empty, JVE289) or plasmid pJR1040 containing a hyperactive RAS2 allele (RAS2V19) (JVE291) were spotted onto YNB – URA synthetic medium containing (I+) or lacking (I-) inositol. 5 l of cells normalized to A600=0.5 were spotted in a series of four 10-fold dilutions.

Lack of inositol increases GIT1 expression in the WT and ira1 strains

Clearly Ira1 and Ira2 affect inositol biosynthesis and GroPIns transport. Since GIT1 expression is upregulated by phosphate starvation (102) and by inositol limitation in a WT strain

(Figure 2-6 S), GIT1 expression upon inositol limitation in the IRA mutants was monitored using a plasmid with β–gal activity driven by the GIT1 promoter. Lack of inositol increases GIT1 expression similar to WT in the ira1 strain, while GIT1 expression in the ira2 strain is unchanged by presence or absence of inositol (Figure 2-6 S). Although there is increased GroPIns accumulation in the IRA mutants as compared to the WT (Figure 2-2), this cannot be explained by the potential lack of GroPIns transport due to a decrease in GIT1 expression because in I+ conditions the mutants and WT have similar GIT1 expression. Furthermore, the transport experiments were performed in abundant phosphate conditions in which Git1 activity is turned off

(102).

Figure 2-6 S. GIT1 expression is decreased in WT and ira1∆ in media containing inositol. WT (JPV 569), ira1∆ (JPV 573), and ira2∆ (JPV 574) were transformed with GIT1-lacZ containing plasmid (JVE 294). At least three individual transformants were grown to late log phase at 30°C in -URA 200µM phosphate +/- inositol media and assayed for β-gal activity using the lab protocol. Data are means ± S.E.

2.3.3 Conclusions

A relationship between Ira1/2 and inositol biosynthesis was uncovered by the finding that the ira1∆ira2∆ mutant is an inositol auxotroph. Misregulation of INO1, which is normally derepressed in the absence of inositol, is typically the cause of inositol auxotrophy. Indeed, INO1- driven -galactosidase activity was decreased upon deletion of IRA1 or IRA2. The fact that a hyperactive RAS2 allele exhibited moderate inositol auxotrophy suggests that this phenotype, unlike the growth phenotype, is attributable to hyperactivation of the RAS/cAMP pathway.

In addition to affecting GroPIns transport and inositol biosynthesis, our results indicate that

Ira1/2 affect GroPIns formation, which occurs via PLB-mediated turnover of PI. Although an increase in GroPIns formation is seen in the ira1 and ira2 mutants, an increase in PLB1 or PLB3 transcript only occurs in the double mutant (Figure 2-3 S). Thus, Ira1/2 regulation of the PLB activity leading to GroPIns formation likely occurs at multiple levels. Potential post-transcriptional mechanisms of regulation include localization or phosphorylation status. An alternate explanation

is that yet another, uncharacterized, phospholipase exists for the formation of GroPIns in the absence of Ira1/2. Importantly, the increased extracellular accumulation of GroPIns seen in the absence of Ira1/2 cannot be explained by a decrease in its transport into the cell, as under the growth conditions of the experiment (I+, high Pi), GIT1 is not expressed (102) (data not shown).

In total, this study indicates a role for IRA1/2 in the ability of cells to use GroPIns as a phosphate source, in both GroPIns formation and transport, and in inositol biosynthesis. Future studies could focus on determining the mechanisms for those disturbances, including whether they involve the RAS/cAMP pathway or other, less defined, functions of Ira1/2. Further study of PLB expression and activity in response to alterations in the Ras/cAMP pathway could help elucidate a link between GroPIns metabolism and this pathway. However, the Ras/cAMP pathway was not addressed further in this dissertation.

Chapter 3 Examination of the roles and regulation of the PLBs and Git1, including examination of phenotypes associated with altered dosage of these genes

Note: The work described in this Chapter was pursued as exploratory work to initially finish the intended M.S. degree. In completing this probing work, and finding an interesting connection between Ypk1 and Plb1, the project was extended into the basis for my Ph.D. dissertation work.

Therefore, many of the described experiments were not investigated in detail. None of the findings in Chapter 3 are published and upon transition from the M.S. to Ph.D. requirements my project was further defined. However, some of the cursory findings presented here helped lead us to the

Ypk1-Plb1 interaction thoroughly examined in Chapter 4 as the major project for my dissertation.

3.1 Introduction

The phospholipids of the plasma membrane are continually being synthesized and degraded in response to environmental conditions. One aspect of this metabolism is carried out through a recycling process consisting of i) deacylation of glycerophospholipids via the B-type phospholipases (45,69,103), ii) the resultant production of intracellular and extracellular glycerophosphodiesters (GPXs), iii) transport of the GPXs back into the cell via the transporter,

Git1 (64,93,102), and iv) the catabolism of and reutilization of the GPXs in cellular metabolism

(43,66). This recycling pathway will be referred to as the glycerophospholipid deacylation, transport, and reutilization (GDTR) pathway. In this chapter, I report on preliminary studies on the GDTR pathway that were initiated, but were not pursued to the level of publication. The topics are as follows: i) morphological assessment of cells upon loss of Git1, loss of PLB1-3, and in cells

utilizing of GroPIns as a sole phosphate source, ii) potential reacylation of GroPIns, and iii) genetic interactions with the PLBs, one of which was pursued for completion of my Ph.D.

3.1.1 The Git1 glycerophosphodiester transporter

The Git1 permease of S. cerevisiae is a H+ symporter and a member of the major facilitator superfamily (MFS) of proteins responsible for transporting GroPIns, and at a lower affinity

GroPCho, into the cell (104,105). Four homologues to S. cerevisiae GIT1 have been characterized in the pathogenic yeast, C. albicans, named CaGIT1–4, each with varied affinity for particular

GPXs (104,106). In mammalian cells, Glut2 is the only GPX transporter characterized to date, but others likely exist (67). GPX transporters are not essential proteins, but in phosphate limiting conditions or in circumstances when GPXs are available, either in the growth medium or in the human body, the ability to transport and utilize the GPXs can be advantageous, and even contribute to virulence of an organism (106).

3.1.2 The S. cerevisiae B-type phospholipases

The three cell surface PLBs, Plb1, Plb2, and Plb3 deacylate the glycerophospholipids at the plasma membrane to produce the GPXs that can then be transported into the cell by Git1

(45,61,69). The other PLB, Nte1, produces intracellular GroPCho at the ER membrane (103).

Published information about these enzymes in S. cerevisiae is reviewed in Chapter 1. Limited studies have focused on characterizing the PLBs so there are many gaps in our knowledge of their roles, specifically in how they affect cellular physiology. Moreover, the purpose of the production and recycling of the glycerophosphodiesters is largely unknown.

O PI CO CH 2 OH O OH CO CH O OH

H2C O P O Fatty acid source OH OH PLB O- PLB LysoPI HO (Transacylase) CH2 O OH CO CH O OH OH

H2C O P O Fatty acid source OH OH PLB O- GPI AT or PLB ?? (lysophospholipase)

HO CH2 OH OH HO CH O OH GroPIns + Free Fatty Acids H2C O P O OH OH O-

Figure 3-1. Activities of the PLBs and hypothetical GroPIns acyltransferase activity. PLB enzymes are known to hydrolyze PI to lysophosphatidylinositol (LysoPI), which can be hydrolyzed again by the lysophospholipase activity of the PLB to produce GroPIns. PLBs are also able to add a fatty acid to LysoPI in the presence of a fatty acid source, typically another phospholipid, via transacylase activity. The existence of a PLB activity reacylating GroPIns has not been characterized. GPI AT, GroPIns acyltransferase.

3.1.3 Potential ability of the PLBs to reacylate GPXs to produce lysoglycerophospholipids or glycerophospholipids

The fate of the glycerophosphodiesters produced by PLB activity is only partially understood. While Gde1 hydrolyzes GroPCho to produce glycerol-3-phosphate and choline (66), no enzyme catalyzing the catabolism of GroPIns has been identified in yeast, although human glycerophosphodiesterases have activity on GroPIns (51). One hypothesis is that the glycerophosphodiesters, GroPCho and GroPIns, can be reacylated to produce LysoPC or LysoPI, followed by additional acylation to produce the full lipids, PC and PI. This GroPCho acyltransferase (GPC AT) activity has been observed in yeast and mammalian cells, but has

remained controversial since enzymes catalyzing this activity remain elusive (107). Many putative acyltransferase enzymes in yeast have been examined, but none were found to possess GPC AT activity. The PLBs possess phospholipase, transacylase, and lysophospholipase activity, but acyltransferase activity with a GPX as the substrate has not been attributed to the PLBs (Figure

3-1).

Importantly, PI is required for cell viability and is synthesized by the action of PI synthase, encoded by PIS1 (Figure 1-3) (108). In the absence of Pis1, cells cannot make PI and will die.

However, if supplied with GroPIns and reacylation of GroPIns occurs, cells with non-functional

Pis1 hypothetically would survive. I explored this possibility in Section 3.3.7.

3.1.4 Morphology and cytoskeletal structure of S. cerevisiae cells

All cells must maintain an optimal architecture for survival. This includes a diverse range of properties such as aggregation, gross morphology, and cytoskeleton structure. Yeast cells can vary in shape and size depending on genetic background, but generally diploids are larger and ellipsoidal, and haploids are smaller and more spherical (109). Screens of the yeast deletion set have yielded morphological classifications of mutants into categories including WT, elongate, round, football, clumpy, and other (110) and classified mutants as large (uge) or small (whi)

(111,112). These morphological classifications are based on cell size, shape, and aggregation compared to WT cells. Cell size regulation through a balance of growth and proliferation is a tightly controlled process in all eukaryotic cells.

Eukaryotic cells have the ability to aggregate together, resulting in formation of tissues and organs in higher eukaryotes, or forming flocs, biofilms, and psuedohyphae in the single celled yeast (113). These non-sexual aggregates in yeast can serve various purposes from protection of the innermost cells, response to nutrient starvation, or a change in membrane properties causing

sticking (114,115). This clumping, or flocculation, occurs due to interactions between the surface glycoproteins encoded by the flocculation (FLO) genes, but recent data indicates involvement of additional genes (113).

The yeast cytoskeleton, composed of actin filaments, microtubules, and septins, functions in multiple processes in the cell including regulation of cell polarity, endocytosis, cell division, maintenance of cell morphology, and intracellular vesicular transport (116,117). Contrary to mammalian cells, yeast actin only forms three primary structures in the cell, actin patches, actin cables, and the actinomyosin ring. Actin patches are dense networks of actin filaments localized to the polarized growth zone and at sites of clathrin-mediated endocytosis, while actin cables are bundles of actin filaments extending the long axis of the cell. The actinomyosin ring forms during cytokinesis at the division site (reviewed in (116)).

Here, I will examine these structural properties – aggregation, morphology, and actin cytoskeleton structure – upon disruption to the GDTR pathway.

3.1.5 Potential functional interactions between the PLB genes and other genes implicated in the maintenance of cellular morphology

To better understand the roles of the PLBs in physiology, I proceeded to identify potential functional interactions between the PLBs and other genes. The resources on the Saccharomyces

Genome Database, including the BioGRID interaction database, were used to narrow down a list of interesting interactions from the total genetic and physical interactions identified for each of the

PLB genes (yeastgenome.org). Based on results that will be described in this chapter, interactions that were potentially involved with cell morphology or actin cytoskeleton maintenance were of primary interest, namely the genetic interactions between Ypk1 – Plb1 and Lge1 – Plb3. The

interaction between Plb1 and Ypk1 proved to be the most interesting and was therefore pursued in great detail in Chapter 4.

Because the glycerophospholipids are a significant structural component of the cell, I hypothesized that alterations in various aspects of the GDTR pathway, might contribute to gross morphological features of the cell. This hypothesis led me to ask, is gross morphology altered in yeast cells upon loss of various genes in the GDTR pathway, specifically GIT1 and PLB1-3, and furthermore, what particular cellular processes are impacted by PLB activity? Here, I investigated several aspects of the GDTR pathway and how defects in the genes mediating this pathway can affect cell physiology. This chapter will focus on four major areas: i) cellular morphology upon loss of Git1 and upon utilization of GroPIns as a phosphate source in WT cells, ii) cellular morphology and utilization of GroPIns upon loss of the PLBs and factors regulating the PLBs, iii) ability of the cell to reacylate GroPIns to produce PI through activity of the PLBs, and iv) functional interactions of the PLBs with other proteins affecting cell morphology and the actin cytoskeleton.

3.2 Experimental Procedures

Strains and Media. The S. cerevisiae strains used in this study are shown in Table 3.1.

Strains were grown aerobically with shaking at 30ºC or 37ºC, as noted. Turbidity was monitored by measuring the optical density at 600nm (A600) on a BioMate 3 Thermo Spectronic spectrophotometer. The cells were grown in various media as indicated. Yeast peptone dextrose

(YPD) was purchased from Fisher Scientific. Synthetic defined medium of yeast nitrogen base

(YNB) was made as described previously, but containing 2% glucose (93). YNB medium was supplemented with either 200 µM KH2PO4 defined as low phosphate (LoPi) or 10 mM KH2PO4 defined as high phosphate (HiPi), or contained 6.8 mM phosphate if not specified. When indicated,

YNB medium was supplemented with 75 µM inositol (I+) or did not contain inositol (I-). Plates were prepared by adding 2% agar.

Yeast Transformation. Plasmid DNA and PCR products were transformed into yeast by high efficiency lithium acetate chemical transformation procedure (118-120). Yeast cells were grown to log phase, made competent with lithium acetate treatment, transformed with the desired

DNA, and transformants were selected on medium appropriate for the specific selectable markers.

When transforming cells with a drug resistance marker, as opposed to a nutritional marker, cells were grown in non-selective rich medium for several hours prior to plating on selective medium.

PCR-mediated gene deletion. Gene deletions were performed by replacing all or part of the targeted gene with a selectable marker (121). Primers were designed with approximately 60 base pairs of homology to the target genomic sequence to be replaced and 20 base pairs homology to the sequences of the selectable marker. The PCR products were purified, verified for correct size by agarose gel electrophoresis, and extracted from the gel. The purified DNA fragments were transformed into the desired yeast strains. Transformants were identified on selective medium.

Genomic DNA was isolated from potential deletion strains using a phenol-choloroform extraction, and the gene deletion was verified by PCR. Verification of deletions was performed using two primer sets, with both primers outside of the targeted deletion area, and the other set containing one primer outside of the targeted deletion and one primer binding inside the inserted selectable marker. Primers are listed in Table 3.2.

Calcofluor Staining. Cells were grown to an OD600 = 0.5. Calcofluor was added at a final concentration of 0.2 mg/mL to an aliquot of culture. Suspension was mixed by vortex agitation and incubated at room temperature for 5 minutes. Cells were pelleted and washed three times with water. An aliquot of resuspended cells was examined by fluorescent microscopy.

Table 3.1 S. cerevisiae strains used in this study (Chapter 3) Strain Genotype Source

JPV 13 WT EG123; leu2-3, 112 ura3-52 trp1-1 his4 can1R derived from EG123, D. Levin JPV 140 leu2-3, 112 ura3-52 trp1-1 his4 can1R plb1∆plb2∆ EG123, Susan Henry JPV 201 a/alpha his3/his3 leu2/leu2 met15/MET15 LYS2/lys2 BY4743 heterozygous diploid ura3/ura3 git1∆/GIT1 JPV 203 WT BY4741; his3∆1 leu2∆0 met15∆0 ura3∆0 Research Genetics JPV 212 his3 leu2 lys2 ura3 git1::KanMX Research Genetics JPV 381 a/alpha his3/his3 leu2/leu2 ura3/ura3 MET15/met15 LYS2/lys2 Research Genetics JPV 399 WT BY4742; his3∆1 leu2∆0 lys2∆0 ura3∆0 Research Genetics JPV 467 E1278b; ura3-52 Heitman lab at Duke University JPV 469 E1278b; ura3-52 git1∆::KanMX our lab, Claudia Almaguer JPV 477 leu2-3, 112 ura3-52 trp1-1 his4 can1R plb1∆ EG123, SH1082, Susan Henry JPV 478 leu2-3, 112 ura3-52 trp1-1 his4 can1R, plb2∆ EG123, SH1083, Susan Henry JPV 479 leu2-3, 112 ura3-52 trp1-1 his4 can1R plb3∆ EG123, SH1084, Susan Henry JPV 480 leu2-3, 112 ura3-52 trp1-1 his4 can1R plb1∆, plb2∆, plb3∆ EG123, SH1085, Susan Henry JPV 569 WT ∑1278b; ura3-52 M. Cardenas JPV 573 ∑1278b; ura3-52 ira1::LoxP-nat M. Cardenas JPV 574 ∑1278b; ura3-52 ira2::LoxP-nat M. Cardenas JPV 575 ∑1278b; ura3-52 ira1::LoxP-nat ira2::LoxP-nat M. Cardenas JPV 617 BY4741; plb1::hph our lab, Beth Surlow JPV 618 BY4741; plb2::KanMX our lab, Beth Surlow JPV 619 BY4741; plb3::nat1 our lab, Beth Surlow JPV 620 BY4741; plb1::hph, plb2::KanMX our lab, Beth Surlow JPV 621 BY4741; plb1::hph, plb3::nat1 our lab, Beth Surlow JPV 622 BY4741; plb2::KanMX, plb3::nat1 our lab, Beth Surlow JPV 623 BY4741; plb1::hph, plb2::KanMX, plb3::nat1 our lab, Beth Surlow JPV 624 ∑1278b; ura3-52, plb1::hph our lab, Beth Surlow JPV 625 ∑1278b; ura3-52, plb2::KanMX our lab, Beth Surlow JPV 626 ∑1278b; ura3-52, plb3::nat1 our lab, Beth Surlow JPV 627 ∑1278b; ura3-52, plb1::hph, plb2::KanMX our lab, Beth Surlow JPV 628 ∑1278b; ura3-52, plb1::hph, plb3::nat our lab, Beth Surlow JPV 629 ∑1278b; ura3-52, plb2::KanMX, plb3::nat our lab, Beth Surlow JPV 630 ∑1278b; ura3-52, plb1::hph, plb2::KanMX, plb3::nat our lab, Beth Surlow JPV 636 BY4742; ypk1::KanMX Research Genetics JPV 645 BY4742; lge1::KanMX, plb3::nat our lab, Beth Surlow JPV 668 BY4742; plb1::hph our lab, Beth Surlow JPV 669 BY4742; ypk1::KanMX plb1::hph our lab, Beth Surlow JPV 679 BY4742; nte1::ble1 our lab, Beth Surlow JPV 680 BY4742; plb1::hph, plb2::KanMX, plb3::nat, nte1::ble1 our lab, Beth Surlow JPV 691 BY4742; ypk1::KanMX, nte1::ble1 our lab, Beth Surlow

Table 3.2 Nucleotide sequences of primers used in this study (Chapter 3) Name Sequence Notes snr17 rev 5' CGG TTT CTC ACT CTG GGG TAC - 3' RT-PCR primer for SNR17 snr17 for 5' TTG ACT CTT CAA AAG AGC CAC TGA - 3' RT-PCR primer for SNR17 PLB1 For. 5' - GCA TAC ACC AAG GAG GCT TTG - 3' RT-PCR primer for PLB1 PLB1 Rev. 5' - GAG TGG ATA GCA AGG AAG TGT CAC - 3' RT-PCR primer for PLB1 PLB3 For. 5' - CCT GAC TTG GTC TTC CAT TAG GGA T - 3' RT-PCR primer for PLB3 PLB3 Rev. 5' - GAT ACC TAC CGT CCG CAA - 3' RT-PCR primer for PLB3 plb2 ForB 5' - ATC TCT GAG GCT ACT GAC GGT - 3' RT-PCR primer for PLB2 plb2 RevB 5' - CGA AGA GTT TGA GTT AGA AGT GAC A - 3' RT-PCR primer for PLB2 Plb1/Hph F 5’ CAA AAC CAT GAA GTT GCA GAG TTT GTT GGT Primer used for PLB1 deletion TTC TGC TGC AGT TTT GAC TTC TCT AAC CAG CTG AAG CTT CGT ACG C ‘3 Plb1/Hph R 5’ AGA GCC TAA ATT AGA CCG AAG ACG GCA Primer used for PLB1 deletion CTA ATG ACA CTT AAG ACA CCA ATA AAA GGC ATA GGC CAC TAG TGG ATC TG ‘3 Plb2/KanMX 5’ CTT ACG CAG ATG CAA TTA CGG AAC ATA TTA Primer used for PLB2 deletion F CAG GCT AGC TCG CTA ATT TCT GGA CTT TCG CTC ATT TAG CCC ATA CAT CC ‘3 Plb2/KanMX 5’ CAG AAG GGC TTC TAA ATT AGT CCA AAT AGT Primer used for PLB2 deletion R CCA AAT AAT GCT GTT ATG GCA CCT AAC AAA CTC ATT AGA GTG GGC AGA TGA TGT CGA GG‘3 Plb3/Nat F 5’ TAA GTA ATG ATA CGT CCA TTA TGT TCA AAA Primer used for PLB3 deletion ATT ATT ATC AGT TAC ATA TTC GCA ACA GCT GAA GCT TCG TAC GC ‘3 Plb3/Nat R 5’ GCA AGA GAT ATT ATA CTG CTC CGG TAA ACA Primer used for PLB3 deletion TCA ATA AAG TAA GCA TAA TCA TAG CGC ATA GGC CAC TAG TGG ATC TG ‘3 Nte1/ble F 5’ GAA TAG AAA GGC TTG GCT CTA GTT TTT GCA Primer used for NTE1 deletion TGC GTT CAA TGA ATT GCA CTA CGA ACA ACG ACA ACC CTT AAT ATA ACT TC ‘3 Nte1/ble R 5’ CAA AAA ACG GTC ACC AAG GCG TTG GAA Primer used for NTE1 deletion GAT AGA AAT CTA TCA AAT ACT ATT TCT TCT ATG ATC TGA TAT CAC CTA ATA AC ‘3 PLB1 XhoI 5’ CAT TAG CTC GAG GGT TTT GTT TTA TTT Primer used for PLB promoter Reverse TCT TTC CCG '3 insertion into pBH315 PLB1 BglII 5’ CAT TAG AGA TCT CTT CAG CCT ATT ATT Primer used for PLB promoter Forward TAC GCA CCG '3 insertion into pBH315 PLB2 XhoI 5’ CAT TAG CTC GAG CTG CGT AAG AAG TGT Primer used for PLB promoter Reverse TAC TTT AC '3 insertion into pBH315 PLB2 BglII 5’ CAT TAG AGA TCT CGG GCA ATG CTT TGG Primer used for PLB promoter Forward TGA ATT ATT C '3 insertion into pBH315 PLB3 XhoI 5’ CAT TAG CTC GAG TAC TTA TAT TAT TAT Primer used for PLB promoter Reverse GTG TGT AAT TTC C '3 insertion into pBH315 PLB3 BglII 5’ CAT TAG AGA TCT ACT ACC GCT TGT GCG Primer used for PLB promoter Forward GAC CTG C '3 insertion into pBH315 LGE1/Leu2 5’ CCC TAC TGC ATT AAT AAC AAT GAG TCC AGC Primer used for LGE1 deletion reverse TTT TCT TGG GTC AGT TGA ATA TTT AAT GCA GAA TCT TTT TAA GCA AGG AT 3’ Lge1/Leu2 5’ GTA TGA GCG GAT ACA CGG GAA ATA ATT Primer used for LGE1 deletion forward ATA GCA GAT ACT CAT CCA CTC CTC CGA GAC AAA CAC ATA CCT AAT ATT ATT GC 3’

Actin Staining with Phalloidin-FITC. The protocol for actin staining was adapted from

Reizman et al., Pringle et al., and the current protocols in molecular biology (122,123). Cells were grown in YPD media to log phase at an OD600 ~ 0.8. To 1 mL of culture 100 µL of 37% formaldehyde was added to fix the cells. The suspension was incubated at room temperature for 1 hour followed by centrifugation at 8,000 rpm for ~5 mins to collect the pellet. Cells were washed three times with 1mL aliquots of phosphate buffered saline (PBS). The pellet was resuspended in

1 mL of 0.2% Triton-X100/PBS solution and incubated at RT (25C) for 10 minutes. The cells were pelleted and washed three times with 1 mL aliquots of PBS followed by resuspension in 100

µL of PBS. To 50 µL of the suspension, 8 µM FITC-phalloidin methanol stock solution was added followed by incubation on ice in the dark for 90 minutes. Cells were washed five times with 1 mL aliquots of PBS. Final pellet was resuspended in PBS. An aliquot was placed onto a microscope slide and allowed to air dry in the dark. 10 µL of ProLong Gold was placed over the sample for preservation and for adherence of the cover glass (#1.5 thickness). The slides were stored overnight in the dark to allow the Prolong gold to solidify. All slides were stored at -20C until viewing by confocal microscopy.

Microscopy. Brightfield and fluorescent microscopy were performed on a Nikon

Microphot-SA microscope with attached camera. Confocal microscopy was performed on the

Zeiss Axio Observer Z1 confocal microscope, and images were obtained with HCImage software

(Zeiss). Scanning Electron Microscopy (SEM) was performed by Dr. Donna Stolz at the University of Pittsburgh.

Construction of the PLB-lacZ containing plasmids. Plasmids containing each of the PLB promoters were constructed previously in the lab (JPV 226, 228, and 230) but were non-functional so new constructs were made. The PLB promoters were amplified by PCR from the respective

plasmids using hybrid primers. The primers were engineered to contain BglII and XhoI restriction sites (Table 3.2). PCR products were purified from agarose gels using a gel extraction kit according to the manufacturer instructions (Qiagen). The PLB promoter products were cloned into the intermediate TOPO pCR2.1 vector using the TOPO TA Cloning Kit with One Shot TOP10 electrocompetent cells (Invitrogen). The potential pCR2.1-PLB promoter plasmids were isolated from transformants and checked for the correct insertions by restriction digest with EcoRI. The constructed intermediate pCR2.1-PLB promoter vectors (JPV 325, 327, and 328) and the pBH315

(JPV 161) were restriction digested with BglII and XhoI. The digested DNA was gel purified and extracted. The pBH315 backbone was treated with CIP followed by gel purification and extraction.

Ligation of the pBH315 backbone with the PLB-promoter DNA was carried out with T4 Ligase at

16ºC for 16 hours. Ligation products were transformed into E. coli competent cells. Plasmids isolated from transformants were checked for the correct PLB promoter insertion by restriction digest with BglII and XhoI as well as by BglII and SphI digestion.

RNA Extraction and real time qRT-PCR gene expression analysis. Cultures were grown in

YNB medium under the conditions specified in the text. RNA extraction and qRT-PCR was performed as described in Section 2.3.1.

β-galactosidase assays. Using cells grown in liquid media β-gal assays were performed as described in Chapter 2.3.1. β-gal activity assays from cells grown on plates were performed by plating the strains transformed with lacZ expression plasmids onto YNB – URA I+ plates containing X-GAL and incubated at desired temperature. Formation of a blue color in the cell patch indicated β-gal activity.

PIS1 promoter replacement with the GAL1 promoter. The PIS1 promoter was replaced with the GAL promoter in the genome by amplifying DNA for insertion by PCR from the plasmid

pFA6a-KanMX6-PGAL1-3HA with hybrid primers named pFA6a-KanMX6-PGAL1-3HA F1

(5'GTACCGATGAGATGAGATGAG…3') and pFA6a-KanMX6-PGAL1-3HA R1

(5'ATTGGGAATATA….3'), each containing homology to the GAL promoter and region surrounding the PIS1 promoter. The PCR product was gel purified and transformed into JPV 13 and JPV 203 and plated onto YT+Galactose+G418 plates.

Sporulation and tetrad dissection. Sporulation media used included, KAc (0.05% glucose,

1% potassium acetate, 0.1% yeast extract in DI water), and minimal media (10% potassium acetate and 0.05% zinc acetate). Sporulation media supplemented with amino acids and bases complementary to the strain auxotrophies were also used. Sporulation and tetrad dissection was carried out as published in Ausubel et. al. (99). Diploids to be sporulated were grown to an OD600

= 3 in 5 mL of YPD medium, cells were harvested for 5 minutes at 1200 x g, washed once in water and resuspended in 1 mL of sporulation medium. Cells were sporulated at 30ºC for up to two weeks or until spores were microscopically visible. Cells were washed three times in sterile water and resuspended in 5 mL of sterile water. 30 µL of the cell suspension was diluted with 270 µL of sterile water and 3 µL of diluted glusulase was added to disrupt asci by vortex agitation.

Suspension was incubated at room temperature until half of all tetrads had a disrupted ascus wall

(2 - 6 minutes). Tetrad dissection was performed by placing treated spores in two parallel lines across the surface of the spore dissection plate (medium specified in text). Using the dissection needle, groups of four spores were separated into a line adjacent to the original position of the tetrad. After dissection, plates were incubated at 30ºC.

3.3 Results

3.3.1 Cell morphology is altered upon loss of the Git1 transporter and in WT cells utilizing

GroPIns as a sole phosphate source

To begin understanding the role of Git1 in cellular morphology, in conjunction with its substrate, GroPIns, several phenotypic analyses were performed. Previous large scale morphological screens of the yeast deletion set classified the heterozygous git1∆/GIT1 diploid strain as football shaped (110). Here, microscopic examination of WT and the heterozygous git1/GIT1 diploid strains revealed minor differences in cell shape between the two strains grown in YPD, YNB I+ HiPi, and YNB I+ LoPi media (Figure 3-2). Each cell was classified as “football” shaped, defined as being pointed on both ends, or as “other”, based upon subjective visual examination and tallying. The percentage of football shaped cells in the git1∆/GIT1 strain was roughly the same in each of the media examined, suggesting phosphate concentration does not significantly affect cell shape.

Transported into the cell by Git1, GroPCho or GroPIns can be supplied as the sole phosphate source for yeast cells (61). These metabolites are also found at physiologically relevant concentrations throughout different regions of the human body (51,124). We sought to determine if the presence of GroPIns in the medium facilitated any changes in the morphology of WT cells.

Microscopic observations of WT cells grown in YNB and YPD medium, containing GroPIns in addition to KH2PO4 as an endogenous phosphate source, resulted in increased cellular aggregation

(Figure 3-3).

Figure 3-2. WT and git1∆/GIT1 diploid strains display minor differences in cell shape. WT (JPV 381) and git1∆/GIT1 (JPV 201) were grown to log phase in YPD, YNB I+ HiPi, and YNB I+ LoPi media. A, An aliquot of each culture was examined under the light microscope. Strains were grown in duplicate and pictures of at least 5 frames per culture were obtained. Cells grown in YPD at 400X magnification are shown as a representative example. B, Images were examined and the percentages of football shaped cells was determined for each strain and condition. Cells in the budding stage were only considered when they were over half the size of the parent cell. All classifications were subjective based on my discretion.

Figure 3-3. WT cells display increased aggregation in the presence of GroPIns. WT cells (JPV 13) were grown to single colonies on YPD and I+ plates at 30ºC. A portion of an individual colony was resuspended in 15 µL of water, agitated by vortex mixing for 30 seconds, and sonication for 30 seconds. An aliquot of the suspension was viewed under the light microscope at 200X magnification. At least 3 colonies under each condition were observed. Pictures are representative of the entire data set.

Over the course of my experiments, I observed that WT yeast cannot grow with GroPIns as the phosphate source when grown at an elevated temperature of 37ºC, prompting me to examine this phenomenon in more detail. WT strains in several genetic backgrounds (EG128, JPV 13;

BY4741, JPV 203; ∑1278b, JPV 569) can grow on plates with GroPIns as a phosphate source at

37ºC (Figure 3-4A), but were defective in growth in liquid media with GroPIns as the phosphate source (Figure 3-4 B and C). When WT cultures are started at an OD600 ≤ 0.02, after 60 hours of growth at 37ºC the culture in YNB I+ NoPi + 200 µM GroPIns media does not reach an OD600 >

0.08, but in YNB I+ LoPi media the OD600 > 1.5 (data not shown). When WT cells were grown in

YNB I+ NoPi + 200 µM GroPIns liquid media initially at 30ºC, and then shifted to 37ºC, growth did not stop, but the cultures reached stationary phase at a lower density (Figure 3-4B).

Furthermore, the lower the cell density at the time of temperature shift, the greater the amount of

reduction in final density (data not shown). Preliminary studies to determine if a potential 96-well plate screen could be performed revealed WT could grow in YNB I+ NoPi + 200 µM GroPIns media at 37ºC in a 96-well plate without shaking, albeit minimally compared to 30ºC (Figure

3-4C). To use as a screen in the future, tests would need to be completed in 96-well plates with shaking at 30ºC and 37ºC, to mimic traditional liquid growth in 5 mL cultures grown in a rotating wheel.

Figure 3-4. WT cells have slowed growth when grown in media requiring utilization of GroPIns as a phosphate source at 37ºC. A, WT (JPV 203 and JPV 569) were grown in YNB I+ LoPi media at 30ºC and diluted to equal densities for plating to single colonies. Aliquots were spread onto YNB I+ LoPi and YNB I+ NoPi + GroPIns plates and incubated at 30ºC and 37ºC for 2 days. B, WT (JPV 203) cultures were started at OD600 = 0.005 in 5mL cultures in listed media and grown at 30ºC in rotator wheel, and subsequently split after 12 hours (red line), for growth at 30ºC and 37ºC. C, WT (JPV 203) cultures were started at OD600 = 0.002 in 200 µL of media in a 96-well plate in the listed media and grown at 30ºC and 37ºC. Data are means ± S.E. of at least triplicate cultures.

3.3.2 Growth of the PLB mutants utilizing GroPIns as a phosphate source.

The preliminary data suggesting WT cells display altered growth and aggregation in the presence of GroPIns prompted me to examine potential physiological or morphological alterations in cells lacking the B-type phospholipases, which produce GroPIns, when grown in the presence or absence of GroPIns in the medium. The plb123∆ (JPV 630) strain can grow with GroPIns as the sole phosphate source at 30⁰C, although slower than WT (Figure 3-5A). Microscopic exam of the cultures revealed the plb123∆ strain displays larger average cell size compared to WT (Figure

3-5 B and C). Furthermore, abnormally large (XL) cells, with large vacuoles, were observed in plb123∆ cells when grown with 200 µM GroPIns as the phosphate source, and no XL cells were observed under the same condition in WT or in corresponding cultures containing 200 µM KH2PO4

(LoPi) as the phosphate source. Cells were also examined in YNB I+ LoPi medium containing additional 200 µM GroPIns, but a notable increase in XL cells was not observed in this medium for WT or plb123∆ (data not shown). This suggests that the forced utilization of GroPIns as a phosphate source in the absence of the PLBs results in the most extreme phenotype. Suspension of cells grown in corresponding media, but on agar plates, produced trends similar to those observed in the liquid media (data not shown). In media requiring utilization of 200 µM GroPIns as the phosphate source, XL cells were observed occasionally in the single and double PLB mutants examined, and subjectively appeared to be most prevalent in the plb2∆, plb12∆, and plb123∆ strains (Figure 3-6). However, since there was not an obvious change in morphology compared to WT a more complete quantitative analysis was not performed in the single or double

PLB deletion strains.

Preliminary scanning electron microscopy (SEM) images of WT and plb123∆ cells

(Courtesy of Dr. Donna Stolz at the University of Pittsburgh) revealed large invaginations in the

surfaces of these XL cells (Appendix A 2). The cause of this observation is unknown, but suggests major cellular alterations when PI deacylation is disrupted, and GroPIns is used as a phosphate source. This result may indicate PLB mediated turnover is required for normal physiology when

GroPIns is incorporated into cellular metabolism.

Figure 3-5. GroPIns can be used as a phosphate source upon loss of the cell surface associated PLBs, but results in aberrant morphology. A, WT (JPV 13) and plb123∆ (JPV 480) were grown in YNB I+ LoPi media and inoculated into listed test media at starting OD600=0.005. B, Aliquots of culture were removed and examined microscopically. Quantitation of cell area from log phase cells growing in equivalent concentrations of KH2PO4 or GroPIns was performed using ImageJ software. Several fields from at least 3 separate cultures were analyzed. C, Phase contrast microscopy of representative fields at 400X magnification after 48 hours of growth.

Figure 3-6. Cells lacking single deletions of the PLB genes display some instances of morphological abnormalities. WT (JPV 13), plb1∆ (JPV 477), plb2∆ (JPV 478), plb3∆ (JPV 479), plb12∆ (JPV 140) and plb123∆ (JPV 480) were grown in in I+ LoPi media and inoculated into test media at OD600=0.005. Cultures were examined by phase contrast microscopy at 400X magnification after 38 hours of growth. Frames are representative of each condition.

Together the morphological abnormalities identified in the plb123∆ strain and in cells grown on GroPIns as a phosphate source could indicate potential alterations in the cytoskeleton cell wall, plasma membrane, and/or vacuolar morphology. Before pursuing other tests, including growth and drug susceptibility assay, another set of PLB mutants was constructed in two different strain backgrounds detailed in the next sections.

3.3.3 Deletion of one or more PLB genes does not alter growth rate

Preliminary experiments with PLB mutants in the EG123 background produced inconsistent results, prompting me to make all of the deletion strains in our standard laboratory strains. Deletion of each of the PLBs, PLB1, PLB2, and PLB3 were made singly and in combination in the BY4741 and ∑1278b backgrounds. Additionally, NTE1 was deleted singly and in the plb123∆ strain in BY4741. Growth of all the newly constructed PLB deletion strains (JPV

617 – 630, Table 3.1) was monitored in several types of media including: YNB LoPi, YNB HiPi, and YNB NoPi + GroPIns, all with and without inositol. No major differences in growth between

WT and plb123∆ in respective media were observed (Figure 3-7). As demonstrated in previous publications, and shown in Figure 3-7, growth on GroPIns results in a long lag phase. Microscopic examination of the plb123∆ strain when grown on GroPIns as the phosphate source revealed the presence of XL cells consistent with the observations in the EG128 background. Interestingly, WT cells in both the BY4742 and in the ∑1278b backgrounds also appeared to have more XL cells when grown on a medium requiring GroPIns utilization compared to low or high phosphate media, although this was a visual observation and not quantified.

Figure 3-7. Loss of PLB 1, 2, and 3 does not alter growth compared to WT. Indicated strains were pre-grown in YNB I- LoPi medium and inoculated at a low starting density into the desired medium. Optical density (A600) of the cultures was monitored over time.

3.3.4 The plb123∆ strain has increased sensitivity to the actin destabilizer, Latrunculin A

Actin cytoskeleton disruption can cause abnormalities in cell morphology. Furthermore, the yeast fitness database identified plb3∆ as having increased sensitivity to Latrunculin A (LatA), an actin destabilizing drug that inhibits actin polymerization. LatA inhibited growth of various combinations of the double PLB mutants, and most severely inhibited growth of the triple plb123∆ mutant, whereas Lat A did not affect growth of the single PLB deletion mutants (Figure 3-8). Cell viability of WT and plb123∆ strains after growth in the presence of LatA was similar based upon

plating equal amounts of cells and counting individual colonies (data not shown). Observation of growth sensitivity to LatA prompted us to microscopically examine the actin cytoskeleton.

Preliminary observations revealed that in fixed cells stained with TRITC-phalloidin in the absence of LatA, the actin cytoskeleton of the plb123∆ cells appeared to be similarly organized to the WT cells, both strains exhibiting actin patches and cables, typical of yeast cells (Appendix A 3). In the presence of LatA, actin patches are visible in the WT, but interestingly, no defined organized actin structures are visible in plb123∆ (Appendix A 3). This microscopic observation was not pursued quantitatively and would require more replicate measures.

Figure 3-8. Loss of multiple cell surface PLBs in the BY4741 background increases sensitivity to the actin destabilizer, Latrunculin A. The cells were grown from equal starting densities at 37C in YPD media with 1µM LatA (in DMSO) or with solvent. Inhibition by LatA for each strain was determined by calculating the percentage of growth in LatA containing media compared to growth in the control for the single (A), double (B), and triple (B) mutants after 25 hours of growth. Growth curves of WT and plb123∆ with and without presence of LatA (C). Chemical structure of Latrunculin A (D).

3.3.5 Phenotypic analysis of WT and strains bearing deletion of the PLB genes upon exposure to stressors targeting the plasma membrane and the cytoskeleton

To continue our analysis of PLB function, growth was examined in the presence of several stressors shown in Table 3.3. In summary, no differences in visible growth sensitivity to the mentioned stressing agents were noted between WT and plb123∆ strains under the conditions examined (liquid and/or plate growth assays in BY4741 and ∑1278b backgrounds, data not shown). The strain bearing deletion of all four S. cerevisiae PLBs (plb123∆ nte1∆) was constructed in the BY4741 background and growth studies showed no alteration in sensitivity to calcofluor,

NaCl, or SDS compared to WT (data not shown).

Table 3.3 Chemical cell stressors evaluated and their targets Drug/Compound Targeted process/Molecular target Name SDS, NaCl, Plasma membrane disturbing agents CdCl2, CuSO4 Calcofluor Cell wall destabilizer Benomyl Microtubule depolymerizing drug Nocodazole Interferes with microtubule polymerization Amphotericin B Ergosterol metabolism; Associates with ergosterol causing ion leakage through the membrane Terbinefine HCl Ergosterol metabolism; Inhibits the enzyme squalene monoxygenase preventing the conversion of squalene to lanosterol in the ergosterol biosynthetic pathway Ketoconazole Ergosterol metabolism; Inhibits enzyme that converts lanosterol to ergosterol Nystatin Ergosterol metabolism; Binds to ergostrol causing K+ ion leakage through the membrane

3.3.6 Role of phosphate and growth phase in PLB expression

In addition to identifying physiological processes associated with PLB activity, I wanted to examine how the PLB genes are regulated at the transcriptional level, and how that regulation impacts the broader GDTR pathway. I hypothesized that the PLB genes may be regulated by phosphate, as is the GroPIns transporter GIT1. Additionally, PLB transcript levels were compared

in log versus stationary phase cells, at 30ºC versus 37ºC, and in cells exposed to various stressors

(SDS, NaCl, calcofluor, amphotericin B, LatA, choline, terbinefine, and nystatin).

Plasmids containing each of the PLB promoters controlling the lacZ gene were constructed as described in Experimental Procedures (Chapter 3.2). The lacZ vectors and β-gal assays were used to obtain preliminary data for PLB expression (data not shown), however, real time qRT-

PCR was used as a more sensitive method to examine message levels under conditions of interest.

Additionally, using the Yeast Tract database (yeastract.com), I identified potential transcription factors with binding sites in the promoters of the PLB genes that were likely involved in altered morphology and/or disrupted actin cytoskeleton. Strains bearing deletion of the genes encoding the transcription factors Msn2, Msn4, Crz1, Sfl1, Xbp1, Hac1, Sko1, and Yap1 were screened for changes in PLB gene expression. Noticeable differences in PLB promoter driven β–gal activity were observed in the msn2∆, msn4∆, and crz1∆ strains, but these were preliminary and not followed up at this stage. However, β–gal assays in WT strains transformed with the PLB-lacZ vectors suggested all three genes encoding the cell surface PLBs (PLB1-3), have greater expression at 37ºC compared to 30ºC.

Since the Git1 transport activity is regulated by phosphate levels (102), I examined PLB gene regulation by phosphate levels. In agreement with the transport data, GIT1 transcript levels measured by qRT-PCR were found to be depressed in YNB HiPi medium compared to YNB LoPi medium (Figure 3-9B). Unlike GIT1, the PLB1-3 genes are not highly regulated by phosphate availability (Figure 3-9C). No difference in message abundance was noted in PLB1 or PLB2 in

WT grown in HiPi versus LoPi medium, while PLB3 message was approximately twice as abundant in cells grown in HiPi compared to LoPi. Message abundance of PLB1-3 in WT cells gown in YNB + GroPIns media compared to YNB LoPi media was not significantly different

(Figure 3-9D). I also report that in WT cells PLB2 and PLB3 message levels decrease in stationary phase, while the PLB1 message increases in stationary phase compared to levels in log phase cells

(Appendix A 4).

A. * B.

Y N B L o P i Y N B H iP i 1 6 1 .5

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Figure 3-9. Glycerophosphodiester transport via Git1, not production via Plb1-3 is regulated by phosphate availability. A – C, Relative message abundance of GIT1, PLB1, PLB2, and PLB3 in a WT strain grown in low Pi compared to high Pi containing medium (A and B) or low Pi compared to GroPIns containing medium (C). Total RNA was extracted and transcript levels were analyzed by real-time RT-PCR using the comparative ∆∆CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT cells grown in high Pi medium. Values represent the means ± S.E. of at least 3 individual cultures each assayed in triplicate. Student’s t-test was performed to compare expression of each gene in media compared to YNB LoPi (A, p = 0.015; B, p = 0.028).

3.3.7 Lethality in the absence of PI synthase cannot be rescued by the presence of GroPIns in the medium

A series of experiments were performed in an attempt to determine if a glycerophosphodiester (GPX) reacylation pathway exists in S. cerevisiae, and if so, does it require the PLBs. GroPIns was chosen as the GPX substrate since GroPIns was shown to be the most abundant glycerophosphodiester accumulating in the media of S. cerevisiae, with GroPCho and glycerophosphoethanolamine present at much lower levels (125). We hypothesized that GroPIns enters cellular metabolism through reacylation to LysoPI and/or PI. Several approaches to determine if a reacylation synthesis pathway for PI exists in the cell will be briefly summarized here, however no evidence for reacylation of GroPIns to PI was obtained. The experimental goal in each of these approaches was to show that when the essential gene PIS1 is deleted or made nonfunctional, the cells can retain viability if the medium is supplemented with GroPIns, suggesting that GroPIns is reacylated to produce the essential PI (Figure 3-1 and Figure 3-10).

Figure 3-10. Known PI synthesis through Pis1 activity and hypothetical PI synthesis by GroPIns acyltransferase activity. PI is synthesized from CDP-DAG and inositol through a condensation reaction catalyzed by PI Synthase (Pis1) and can be catabolized through the action of Plb3 to produce GroPIns. A PI synthesis pathway by GroPIns reacylation through GroPIns acyltransferase (GPI AT) activity is currently uncharacterized.

Using published methods described for chromosomal promoter replacement and PCR based gene deletion, the endogenous PIS1 promoter was replaced with the GAL1-inducible promoter (121,126). Transformants containing the GAL1 promoter were obtained, but while the transformants grew better on medium with galactose, cells continued to grow on glucose medium.

In conclusion, the GAL1 promoter was too leaky to perform the depletion experiment. A few years later, the Lopes lab published work on a pis1∆ haploid strain containing a GAL-PIS1 plasmid

(127). They kindly sent us this strain, but preliminary tests revealed that it still grows fairly well on medium containing glucose, as well as on galactose medium, corroborating my attempt.

A diploid pis1∆/PIS1 (JPV 605) strain was transformed with the pYES2CT-HA-PIS1 plasmid and transformants were sporulated in several types of sporulation media. Tetrads were dissected to isolate individual spores and the resulting individual colonies were tested to determine which were haploid by a mating assay, tested for PIS1 deletion by G418 resistance, and checked for presence of the PIS1 containing plasmid by growth on –URA plates. Only two of the spores screened met all of the requirements. These strains was propagated on galactose containing plates.

Equal starting amounts of cells were grown in liquid and on plates in the following media: YNB

I- LoPi, YNB I- LoPi + GroPIns + 5-FOA, and YNB I- LoPi + 5-FOA (some containing 5-FOA to induce loss of the plasmid and hence produce a pis1∆ strain). Growth occurred on all of these media indicating that even though the strain was confirmed to have PIS1 deleted and contains the

PIS1 plasmid, something else was compensating for the deletion, or the cell had another copy of

PIS1. Checking for PIS1 deletion by PCR revealed bands consistent with the presence of both a

WT copy of PIS1 and a copy in which URA3 was inserted, suggesting that they are not truly haploid, that a recombination event occurred with PIS1 on the plasmid, or that mixed colonies were obtained. This question was not pursued further.

Direct deletion of PIS1 in a haploid strain in the presence of GroPIns was attempted. Since

PIS1 is an essential gene this attempt would only be successful if GroPIns can be reacylated to rescue the PIS1 deletion. Homologous recombination by insertion of KanMX and transformation onto plates containing GroPIns resulted in transformants, but the insert was never in the correct location. As a control, PIS1 was successfully deleted in a diploid strain using the same methods used for the haploid. Because selection for KanMX resulted in significant background growth,

PIS1 was also directly deleted by homologous recombination by insertion of URA3. Many successful transformants appeared in the control diploid strains, and a few transformants appeared in the haploid strain. Despite these strains being verified to be haploids through a mating assay, and PCR checking revealed several transformants contained the URA3 insert in the correct location, interestingly a band indicating the presence of a WT copy of PIS1 also appeared. The results suggest there was an extra copy of the chromosome, or this region was under pressure to duplicate.

A pis1/PIS1 diploid was constructed in a diploid strain known to have a high sporulation efficiency (JPV 381). The deletion was made by insertion of URA3 and verified by PCR. The newly constructed diploid pis1∆/PIS1 (JPV 616) was efficiently sporulated in sporulation media

(1% potassium acetate, 0.1% Bacto Yeast Extract, 0.05% dextrose). Tetrad analysis by pulling spores directly onto plates containing GroPIns produced maximum of two developing colonies per tetrad indicating that the pis1∆ haploid strain is unable to be rescued by the presence of GroPIns in the medium. Therefore, it is unlikely that GroPIns is able to be reacylated in order to rescue a pis1∆ mutant.

In summary, I was unable to rescue a pis1∆ mutant through GroPIns supplementation. This could mean either that GroPIns cannot be reacylated, cannot be reacylated efficiently enough to

rescue the lethality, that genes responsible were not expressed under the conditions used here, or that Pis1 has another essential function besides synthesis of PI. To date, no enzyme has been characterized with acyltransferase activity to reacylate GroPIns, but recent work in the Patton Vogt lab in collaboration with the Stymne lab indicates GroPCho can be reacylated by a novel gene product in both yeast and plants.

3.3.8 The functional Interaction between the PLBs and other genes, specifically Ypk1 and

Lge1

The described results concerning morphological and actin cytoskeletal abnormalities prompted me to examine the genetic interactions between Ypk1 – Plb1 and Lge1 – Plb3 because of their known involvement in actin dynamics and cell morphology, respectively. The Ypk1 – Plb1 interaction became the primary focus for this dissertation and is discussed in Chapter 4. A genome- scale interaction study identified a negative genetic interaction between LGE1 and PLB3, in which the lge1∆ plb3∆ double mutant exhibited slower growth than either single mutant alone (128).

Notably, the LGE1 mutant exhibits a large cell phenotype under optimal growth conditions (129).

LGE1 is a verified ORF, predicted to be localized to the nucleus, but the function of the protein is currently unknown. Together with my data concerning the PLBs, I hypothesized this interaction impacted processes involved with maintaining proper cell morphology. Only preliminary data was collected examining the Plb3-Lge1 interaction. The lge1∆ plb3∆ strain was constructed to begin assessing the interaction between the two genes. Consistent with the published large scale genetic screen, the lge1∆ plb3∆ strain displayed slowed growth compared to the single deletion strains

(Appendix A 5). No further studies were performed with the lge1∆ plb3∆ strain in this dissertation, but a follow-up of this interaction may elucidate a novel role for the PLBs, specifically Plb3, in maintaining optimal morphology.

3.4 Discussion

Perturbation to the GDTR pathway by deletion of GIT1 or the PLB genes, or by forcing utilization of GroPIns as a sole phosphate source results in a variety of morphological changes in the cell. These morphological changes are known to be regulated by and/or affect a diverse range of processes. Since all of the players in the pathway have functional homologs in higher eukaryotes and because GroPIns is present in the human body, this has relevance for higher eukaryotes.

Preliminary microscopic studies indicate the presence of GroPIns in the medium causes increased clumping in WT cells (Figure 3-3). Genetically, aggregation of cells generally occurs upon induction of the FLO genes, which are in part regulated by various environmental conditions including cations, pH, temperature, sugars, and ethanol (115). In some fungi aggregation can lead to formation of hyphae and biofilm structures, which contribute to pathogenesis. Possibly, GroPIns could be a signal promoting aggregation. Future studies may examine the expression of the FLO genes, or other genes implicated in flocculation. A sedimentation test would provide a quantitative assessment of differences in aggregation (113).

Morphological changes in the cell can provide clues for elucidating the function of particular proteins. The football shaped cells observed upon loss of GIT1 are consistent with a published screen classifying the git1∆ mutant as football shaped (110). Altered cell shape can be due to a variety of factors involving the cell wall, plasma membrane, and cytoskeleton. Most interestingly, loss of the three cell surface PLBs resulted in XL cells, membrane invaginations, and actin cytoskeletal defects when forced to utilize GroPIns. My results suggest that the presence of

GroPIns exacerbates disruptions to cell morphology in strains or conditions where morphology is already compromised, such as upon loss of the PLBs or upon heat stress (Figure 3-4 and Figure

3-4).

To elucidate the underlying mechanism for the observed changes in cell size and morphology upon loss of the PLBs, I examined the effect of stressor known to perturb the cell wall, plasma membrane, and cytoskeleton. Increased sensitivity to LatA in plb123∆ yeast (Figure

3-8) and actin staining (Appendix A 3) indicate loss of the PLBs results in a compromised actin cytoskeleton, providing a possible explanation for some of the observed morphological alterations.

However, growth alone is not a complete measure of defects in plasma membrane and cell wall stability so these preliminary studies do not entirely rule out plasma membrane or cell wall defects in plb123∆ yeast cells.

Membrane expansion and failure to divide can result in XL cells to develop (111). An experiment involving synchronization of cells and subsequent monitoring of division could also be employed. Since cultures did not produce uniformly XL cells, a series of measurements from mother and daughter cells would be useful to determine if there is a significant alteration in cell size in plb123∆ yeast.

There are a few descriptions in the literature describing cellular circumstances that produce large invaginations of the yeast cell membrane, however the cause of the large invaginations observed in plb123∆ cells is unknown. Published observations have demonstrated the presence of invaginations in cells including furrow like invaginations at the membrane compartment containing Can1 (MCC) domain and the sub-adjacent eisosomes (130-133), invaginations in regions termed PtdIns[4,5]P2 enriched structures (PES) (132), and invaginations in cells that are lacking two of the three genes encoding Sac1-containing phosphatases, INP51, INP52 and INP53

(133,134). A recurring theme relating to cellular invaginations are the importance of phospholipids, particularly the phosphorylated phosphatidylinositols (PIPs), also essential regulators of the actin cytoskeleton (135,136). Interestingly, addition of glucose or ergosterol will

induce major changes in the PIPs (137,138). For example, addition of glucose causes rapid hydrolysis of the PIPs to produce the GroPIPs (59), an activity that has been suggested to be mediated by the PLBs. Therefore, it is tempting to speculate that these structures could be related to PtdIns[4,5]P2 levels and Sac1 function. Evaluation of these possibilities requires further experimentation.

Here, I show that the PLBs are transcriptionally regulated by temperature. Although preliminary experiments by β-gal accumulation assays did not indicate changes in transcription upon LatA treatment, it would be interesting to examine PLB message by the more sensitive qRT-

PCR method. Furthermore, using the β-gal assays I was unable to conclusively identify transcription factors strictly regulating the PLB genes, however further examination of the TFs of interest, Msn2/4 and Crz1, under various stress conditions will shed light on PLB gene regulation

(Chapter 4.3.6). Regulation data compiled on SGD suggests several other transcription factors of interest that are shown to regulate PLB message levels under elevated temperature, various media, exposure to drug perturbations, or in different growth phases, indicating the combinatorial control regulating the PLB genes.

The PLBs have phospholipase, transacylase, and lysophospholipase activities (45,69). The ability of the PLBs to acylate GroPIns to PI has not been shown, and the studies described in this chapter suggest the cell does not have the capability to re-acylate GroPIns, or not sufficiently enough, to rescue growth in the absence of PI synthase under the conditions tested here. However, the ability of yeast and plants cells to reacylate GroPCho has been shown in vitro (107,139), but the activity remained controversial without identification of the gene product. Recently, a paper submitted for publication from our lab in collaboration with the Stymne lab, in which I am a co- author, reveals more evidence for GroPCho acyltransferase (GPC AT) activity in plants and in

yeast (submitted and in revision, Lager et al., 2014). This activity results in acylation of GroPCho to produce LysoPC, either by transferring an acyl group from an acyl-CoA or from LysoPC, followed by subsequent acylation to the full lipid, PC, by PLB or Ale1 transacylase activity. The

GPC AT activity is not from PLB activity, but instead from an uncharacterized protein. Studies on this gene are currently underway in the Patton-Vogt lab in collaboration with the Stymne lab.

Pursuing other functional relationships with the PLB genes, including the Plb3-Lge1 interaction, could provide more complete understanding of the physiological significance of the

PLBs. Lge1 plays a role in ubiquitination of histone H2B in the nucleus, but minimal new information on Lge1 has been published (140). Moreover, hundreds of physical and genetic interactions with Lge1 have since been identified making the pursuit of the Lge1-Plb3 interaction less attractive (yeastgenome.org). Although, the interaction was initially interesting because of the large cell phenotype, which I thought could correspond to the XL plb123∆ cells we observed.

However, other interactions such as a recently identified interaction between a vacuolar protein,

Mon2, and PLB1, 2, and 3 (141) are of greater interest especially given the results that will be described in Chapter 4. Briefly, strains bearing deletion of one of the PLB genes displayed altered vacuolar fragmentation, an effect not observed in mutants with defective PLC or PLD pathways

(141).

In summary, loss of the cell surface associated PLBs and/or utilization of GroPIns causes alterations in the actin cytoskeleton and an increased cell size. Furthermore, PLB1-3 are regulated at the transcriptional level by temperature and growth phase in WT yeast. A more in depth characterization of the PLBs is essential in understanding of the GDTR pathway. Confirmation of and follow up of the observations presented in this chapter would provide more insight into the function and purpose of the PLBs in the GDTR pathway and in cellular physiology.

Chapter 4 Accelerated Plb1-mediated turnover of phosphatidylcholine upon loss of Ypk1 in S. cerevisiae

Submitted to the Journal of Biological Chemistry.

Surlow, B.A.1, Cooley, B.1, Needham, P.2, Brodsky, J.2, Patton-Vogt, J.1§ (2014).

Chapter 4 Attributions: I completed all of the work in this chapter except those noted here.

Benjamin Cooley, an undergraduate student in the Patton-Vogt lab, assisted in completion of sample processing for separation of choline containing metabolites by anion exchange chromatography and aided in sample preparation and running samples by mass spectrometry.

Patrick Needham, a research assistant professor in the Brodsky lab, taught me how to prepare a sucrose gradient and together we performed the sucrose gradient experiment for Plb1 localization.

1From the Department of Biological Sciences, Duquesne University, Pittsburgh, PA

2From the Department of Biological Sciences, University of Pittsburgh, Pittsburgh, PA

*Running title: Ypk1 regulates phosphatidylcholine turnover

§ To whom correspondence should be addressed: Jana Patton Vogt, Department of Biological Sciences, Duquesne University, Pittsburgh, PA, USA, Tel.: (412) 396-1053; Fax: (412)396- 5907; E-mail: [email protected]

Keywords: Yeast, Phosphatidylcholine, Glycerophosphocholine, Sphingolipids, Phospholipid turnover, Phospholipase, Protein Kinase

Background: Ypk1 is a protein kinase known to regulate sphingolipid homeostasis. Plb1 deacylates phosphatidylcholine to produce glycerophosphocholine. Results: Loss of Ypk1 or disruption of sphingolipid synthesis by other means elevates Plb1- mediated phosphatidylcholine turnover. Conclusion: Accelerated turnover of phosphatidylcholine compensates for aberrant sphingolipid synthesis. Significance: Sphingolipid synthesis is coordinated with phosphatidylcholine metabolism to maintain membrane lipid homeostasis.

ABSTRACT

Ypk1, the yeast homolog of the human serum- and glucocorticoid-induced kinase (Sgk1), affects diverse cellular activities, including sphingolipid homeostasis. We now report that Ypk1 also impacts the turnover of the major phospholipid, phosphatidylcholine (PC). Pulse-chase radiolabeling reveals that a ypk1∆ mutant exhibits increased PC deacylation and glycerophosphocholine (GroPCho) production compared to WT. Deletion of PLB1, a gene encoding a B–type phospholipase that hydrolyzes PC, in a ypk1∆ mutant curtails the increased PC deacylation. In contrast to previous data, we find that Plb1 resides in the ER and in the medium.

Consistent with a link between Ypk1 and Plb1, the levels of both Plb1 protein and PLB1 message are elevated in a ypk1∆ strain compared to WT yeast. Furthermore, deletion of PLB1 in a ypk1∆ mutant exacerbates phenotypes associated with loss of YPK1, including slowed growth and sensitivity to cell-wall perturbation, suggesting that increased Plb1 activity buffers against the loss of Ypk1. Since Plb1 lacks a consensus phosphorylation site for Ypk1, we probed other processes under the control of Ypk1 that might be linked to PC turnover. Inhibition of sphingolipid biosynthesis by the drug myriocin or through utilization of a lcb1-100 mutant results in increased

PLB1 expression. Furthermore, we discovered that the increase in PLB1 expression observed upon inhibition of sphingolipid synthesis or loss of Ypk1 is under the control of the Crz1 transcription factor. Taken together, these results suggest a functional interaction between Ypk1 and Plb1 in which altered sphingolipid metabolism upregulates PLB1 expression via Crz1.

4.1 Introduction

Ypk1, the yeast homolog of the human serum- and glucocorticoid- induced kinase (Sgk1), is a serine/threonine protein kinase known to affect multiple downstream processes, including lipid homeostasis (142-146), actin dynamics (147,148), cell wall integrity (147,148), endocytosis

(144,149), and lipid flippase activity (142). Ypk1 has a functionally redundant homolog, Ykr2.

Either Ypk1 or Ykr2 is required for cell viability, but Ypk1 plays a more prominent role in several processes (147,150). For example, deletion of YPK1 results in a growth defect and increased sensitivity to several drugs, phenotypes that are not observed in the ykr2∆ mutant (147). Ypk1, a cytosolic protein, is recruited to the plasma membrane through its interaction with Slm1/2, proteins which contain a pleckstrin homology (PH) domain that mediates binding to PI[4,5]P2 (151,152).

At the plasma membrane Ypk1 is first activated by target of rapamycin complex 2 (TORC2)

(151,152) and then by Pkh1/2 phosphorylation (147). TORC2 has been shown to be activated by sphingolipid depletion (153) and elevated reactive oxygen species (ROS) (154). In turn, Pkh1 and

Pkh2, homologs of the mammalian phosphoinositide-dependent protein kinase-1 (PDK1), preferentially phosphorylate and activate Ypk1 and Ykr2, respectively, although some crossover is observed (147). Pkh1 and Ypk1 are localized exclusively to the cytosol, while Pkh2 and Ykr2 also enter the nucleus. Exogenous addition of the long chain base (LCB) phytosphingosine (PHS) activates Pkh1/Pkh2 in vitro as measured by an increase in PKC phosphorylation (155), while a more recent paper suggests that the ability of Pkh1/2 to activate Ypk1/2 is unaffected by LCBs, but instead requires the complex sphingolipid, mannosylinositol phosphorylceramide (MIPC)

(142).

The known targets of Ypk1 phosphorylation, Fpk1/2 (142), Orm1/2 (143), and Gpd1 (156) are all involved in some aspect of lipid metabolism. Ypk1 phosphorylates and thereby inactivates

Fpk1 and Fpk2, which are upstream activators of the lipid flippase complexes, Lem3-Dnf1 and

Lem3-Dnf2, respectively (142,157). Ypk1 phosphorylation of the endoplasmic reticulum (ER) transmembrane proteins, Orm1 and Orm2, renders them unable to bind to and inhibit serine palmitoyl transferase (SPT), the rate-limiting step in sphingolipid biosynthesis (13,143,158).

Finally, Ypk1 phosphorylates and inactivates Gpd1, glycerol-3-phosphate dehydrogenase, which reduces dihydroxyacetone phosphate (156). The product of this reaction, glycerol-3-phosphate, can be shuttled into multiple metabolic pathways, including phospholipid biosynthesis. A simplified schematic of the Ypk1 signaling pathway is shown in Figure 4-1.

POTENTIAL EFFECTS OF YPK1 DELETION ON PHOSPHOLIPID HOMEOSTASIS:

↑Flippase Activity = Less PE and PS in outer leaflet of PM

↓SPT activity = Decreased basal levels of sphingolipids

↓Ceramide Synthesis = Decreased complex sphingolipids and more sphingosine phosphates

↑Glycerol-3-P = More flux into PC synthesis

Figure 4-1. Simplified schematic of the Ypk1 signaling pathway and downstream events. Ypk1/Ypk2 are activated when phosphorylated by the kinases Pkh1 and TORC2. Ypk1/2 phosphorylate and negatively regulate the flippase activator proteins, Fpk1/2. Fpk1/2 can, in turn, phosphorylate and negatively regulate Ypk1. Ypk1 phosphorylates Orm1/2, negative regulators of the sphingolipid biosynthetic genes LCB1/2. Orm1/2 normally binds and inhibits serine palmitoyl transferase so when Orm1/2 are phosphorylated inhibition is relieved and sphingolipid synthesis can proceed. Ypk1 phosphorylates Gpd1, glycerol-3-phosphate (G3P) dehydrogenase, turning off G3P production. Glycerol 3 phosphate dehydrogenase reduces DHA to glycerol-3-phosphate. G3P can act as a precursor to phospholipids. Ypk1 phosphorylates and activates Lag1 and Lac1, components of the ceramide synthase (142,143,146,147,156,159) and unpublished results from the J. Thorner Lab).

Another finding linking Ypk1 to lipid metabolism is that overexpression of the B-type phospholipase, PLB1, rescues the lethality of the temperature sensitive ypk1tsykr2∆ mutant (1).

The B-type phospholipases (PLB) catalyze the deacylation of glycerophospholipids to produce the glycerophosphodiesters and free fatty acids (Fig. 1D). In S. cerevisiae, four genes encode proteins with PLB activity, PLB1, PLB2, PLB3, and NTE1, which function in membrane remodeling in the cell. Based on in vivo studies, Plb1 primarily deacylates phosphatidylcholine (PC) to produce external glycerophosphocholine (GroPCho) (69). Biochemical studies have suggested that Plb1 is plasma membrane-associated and secreted from the cell (43,45,69), whereas Nte1 is localized to the endoplasmic reticulum and is responsible for the production of internal GroPCho (103). The primary substrate for Plb2 appears to be exogenous phospholipids, and Plb3 acts on phosphatidylinositol (PI) to produce extracellular glycerophosphoinositol (GroPIns) (43,45).

PC synthesis and catabolism are tightly regulated in yeast (9) and other eukaryotic cells

(18), and disruptions to PC metabolism can result in aberrant physiological conditions. In mammalian cells increased choline uptake and incorporation into PC is a marker for proliferative growth (160), and conversely, inhibition of choline synthesis results in apoptosis (161). Aberrant

PC levels and/or alterations in GroPCho have also been observed in patients with various cancers, such as colorectal (162) and prostate (163) cancer, and in Alzheimer’s disease (164,165). What has remained unknown, however, is how protein kinases and members of the B-type phospholipases, which deacylate phospholipids to produce GroPCho, are coordinately regulated to ensure lipid homeostasis. Here, we describe a novel role for Ypk1, the yeast homolog of Sgk1, in regulating Plb1 mediated PC turnover.

4.2 Experimental Procedures

Strains, plasmids, media, and growth conditions. The S. cerevisiae strains used in this study are shown in Table 1. Strains were grown aerobically with shaking at 30ºC or 37ºC, as noted. 72

Turbidity was monitored by measuring the optical density at 600nm (A600) on a BioMate 3 Thermo

Spectronic spectrophotometer. Empty vectors, YEp351 and pRS426, and vectors overexpressing each of the PLB genes, YEp351-PLB1, YEp351-PLB2, YEp351-PLB3, and pRS426-NTE1, were gifts from Susan Henry (YEp351 series) (45) and Chris McMaster (pRS426-NTE1) (166). Media used for this study included rich yeast extract peptone dextrose (YPD) medium purchased from

Fisher Scientific, or synthetic complete yeast nitrogen base (YNB) medium made as described previously, but containing 2% glucose and 75 µM inositol (93).

Construction of strains. The S. cerevisiae WT strain BY4742 was purchased from Open

Biosystems (Thermo Scientific, Huntsville, AL, USA). The deletion strains were constructed using standard homologous recombination techniques (121). Drug resistant markers, hphMX, KanMX, natMX, and ble were amplified from plasmids pAG32, pUG6, pAG25, and pUG66, respectively, using the primers listed in Table 2, and the resulting DNA fragments were inserted in place of the genes targeted for deletion. The plasmids were received from Euroscarf (167,168). For deletion of

YPK1 in the crz1∆ strain, the URA3 cassette was amplified from plasmid pPMY-3xHA (169), and the resulting DNA fragment was used to disrupt YPK1.

Table 4.1 S. cerevisiae strains used in this study (Chapter 4) Strain Genotype Source

JPV 399 WT, BY4742 BY4742; MATα his3∆1 leu2∆0 lys2∆0 ura3∆0 Research Genetics JPV 636 ypk1∆ BY4742; ypk1∆::KanMX Research Genetics JPV 668 plb1∆ BY4742; plb1∆::hph This study JPV 766 ypk1∆plb1∆ BY4742; ypk1∆::KanMX plb1∆::hph This study JPV 744 WT PLB1-I-3xHA BY4742; PLB1-I-3xHA This study JPV 745 ypk1∆ PLB1-I-3xHA BY4742; ypk1::KanMX PLB1-I-3xHA This study JPV 671 crz1∆ BY4742; crz1∆::KanMX Research Genetics JPV 782 crz1∆ypk1∆ BY4742; crz1∆::KanMX ypk1∆::URA3 This study JPV 680 plb123∆nte1∆ BY4742; plb1∆::hph plb2∆::KanMX plb3∆::nat This study nte1∆::ble YPH 499 WT, YPH499 MATa ade2-101och his3∆200 leu2∆1 lys2-801am trp1∆1 (147) ura3-52 YES 3 ypk1∆ YPH499; ypk1-∆1::HIS3 (147) YES 1 ykr2∆ YPH499; ykr2-∆1::TRP1 (147) YPT 40 ypk1tsykr2∆ YPH499; ypk1-1ts::HIS3 ykr2-∆1::TRP1 (147) JPV 772 plb1∆ YPH499; plb1∆::hph This study JPV 773 ypk1∆plb1∆ YPH499; ypk1-∆1::HIS3 plb1∆::hph This study JPV 774 ypk1tsykr2∆plb1∆ YPH499; ypk1-1ts::HIS3 ykr2-∆1::TRP1 plb1∆::hph This study YFR 276 ypk1tsykr2∆orm1∆ YPH499; ypk1-1ts::HIS3 ykr2-∆1::TRP1 orm1∆::hphNT1 (143) YFR 279 ypk1tsykr2∆orm2∆ YPH499; ypk1-1ts::HIS3 ykr2-∆1::TRP1 orm2∆::URA3 (143) YFR 283 ypk1tsykr2∆orm1∆orm2∆ YPH499; ypk1-1ts::HIS3 ykr2-∆1::TRP1 orm1∆::hphNT1 (143) orm2∆::URA3 RH 406 WT MATα leu2 trp1 ura3 bar1 (170) RH 3804 lcb1-100 MATα leu2 trp1 ura3 bar1 lcb1-100ts (170)

Table 4.2 Nucleotide sequences of primers used for gene deletions in this study (Chapter 4) Gene Name Primer Sequence 5' - 3' PLB1 Forward CAAAACCATGAAGTTGCAGAGTTTGTTGGTTTCTGCTGCAGTTTTGACTTCTC TAACCAGCTGAAGCTTCGTACGC PLB1 Reverse AGAGCCTAAATTAGACCGAAGACGGCACTAATGACACTTAAGACACCA ATAAAAGGCATAGGCCACTAGTGGATCTG CRZ1 Forward CCATCACAGCAGTTTAGTACGATTATACACTCTGAACGCGAGTAAAACTAGG TTGTATTTCAGCTGAAGCTTCGTACGC CRZ1 Reverse TACAAGGGAGTGATGCATCTTCTACCACTTTCAGTTTTAAAATGCCTACCAA GAGCAGCATAGGCCACTAGTGGATCTG YPK1 Forward AGTTCCTCTCATATCAACAAACATTAATACAGTTCCTGAAAATGTATTCTTGG AAGTCAAAGCGAACAAAAGCTGG YPK1 Reverse ATGTATTCACTAAGTCTATCTAATGCTTCTACCTTGCACCATTGAGCTACCTA GCTGTTCCTGTAGGGCGAATTGGG PLB1-I- Forward CTAACAGAGAACGTTAACGCTTGGTCACCAAATAACAGTTACGTC 3xHA AGGGAACAAAAGCTGG PLB1-I- Reverse TTCTCTGACTAAGTTAATATCATCATCACAGGTTACGTTCGCAGGCTGTAGG 3xHA GCGAATTGGG Bolded sequences are homologous to the template plasmids.

[14C]-Choline labelling and preparation of external, internal, and membrane fractions.

Cells were grown to uniform labeling in YNB medium containing 20 µM choline and 1 µCi/mL

[14C]-Choline Chloride of negligible concentration (Perkin Elmer, NEC141VU250UC or

American Radiolabelled Chemical, ARC 0208). Cells were harvested in log phase, washed free of excess label, and re-inoculated at OD600 = 0.2 into YNB medium containing 10 mM nonradioactive choline. At various time points, 1 mL aliquots of the chase cultures were removed and separated into external, internal, and membrane fractions (93,171). In brief, cells were pelleted by centrifugation and the supernatant was removed and retained as the extracellular fraction. The cell pellet was resuspended in 0.5 mL of 5% trichloroacetic acid (TCA) followed by a 20 minute incubation on ice. Cells were pelleted again by centrifugation and the supernatant was removed as the intracellular (soluble) fraction. The pellet was then resuspended in 0.5 mL of 1 M Tris (pH =

8), centrifuged, and the supernatant was added to the intracellular fraction. The final cell pellet was resuspended in 0.5 mL of 1 M Tris (pH = 8) and saved as the membrane fraction. Total counts

(external + internal + membrane) recovered at each time point as well as the percentages of counts in each fraction were tracked by liquid scintillation counting (LSC).

Analysis of choline-containing metabolites. The water soluble choline containing metabolites were separated by anion exchange chromatography (172). Internal and external samples were diluted 5-fold in deionized H2O, applied to a 250 µL Dowex 50Wx8 200 – 400 anion exchange column, and eluted as described previously (172). [14C]-GroPCho was eluted with a 1

14 mL and 2 mL H2O wash. If present, [ C]-choline phosphate was eluted with a subsequent 3 mL

14 H2O wash. [ C]-choline was eluted with 5 mL of HCl. Standards were used to verify the separation procedure, and label incorporated into each metabolite was quantified by LSC. To confirm results by independent methods, [14C]-choline containing metabolites from selected samples were

separated by TLC using previously published methods (173) and/or by HPLC with an in-line radioactive detector (65,66). For TLC, samples were dried down, resuspended in 1:1 methanol:H2O, were spotted on Silica plates, and metabolites separated with a mobile phase of

CH3OH:0.6% NaCl:NH4OH (50/50/5, v/v/v).

Phospholipids from the membrane fraction were extracted as described previously (174).

The membrane pellet was resuspended in 1 mL of ESOAK (95% ethanol: diethylether: H2O: pyridine: NH4OH, 15:5:15:1:0.036) and incubated at 60ºC for one hour. The suspension was centrifuged and the lipid containing supernatant was extracted with 2.5 mL of choloroform:methanol (2:1). Low speed centrifugation separated the layers and the bottom layer containing the glycerophospholipids was dried under N2 and the residue was resuspended in choloroform:methanol (2:1). Radiolabelled PC and LysoPC were resolved by TLC on Silica Gel

60A (Whatman) plates in chloroform: ethanol: H2O: triethylamine (30/35/7/35, v/v/v/v) mobile phase (171,175).

Analysis of PC synthesis rate. Strains were grown to log phase (OD600 = 0.5) in YNB 75

µM inositol medium, harvested by centrifugation, and concentrated to equivalent densities in fresh

YNB containing 75 µM inositol and 100 µM choline. To start the assay, 50 µL of 0.005 mCi/mL

[C14]-choline chloride was added and cells were allowed to incubate for 30 minutes at 37°C. The cells were pelleted by centrifugation, washed with water, and suspended in 0.5 mL of 5% TCA for

10 minutes. After pelleting, the supernatant was discarded and the membrane pellet was washed with water. Lipids were extracted from the remaining pellet as described above. Incorporation of

[C14]-choline chloride into PC was quantified by LSC and verified by TLC.

LC-MS analysis of extracellular choline and GroPCho. Cells were grown in YNB medium containing 75 μM inositol, 20 μM choline, and 2 mM KH2PO4 instead of 7 mM KH2PO4. The cells

were pelleted by centrifugation at 1,600 x g for 3 minutes and the supernatant containing the extracellular metabolites was filtered through 0.2 µM cellulose acetate filters. The method of analysis for these extracellular metabolites by mass spectrometry was described previously

(104,176). A 250 µL aliquot of supernatant was spiked with an internal standard (choline-d9) and water soluble lipid metabolites were extracted by addition of 2.25 mL of cholorform:methanol

(2:1, v/v). The suspension was agitated with a Vortex mixer followed by centrifugation for 3 minutes at 1,600 x g. The upper aqueous phase was dried under N2 and resuspended in 125 μL of acetonitrile:methanol (75:25, v/v) with 10 mM ammonium acetate, pH 4.5. After centrifugation at

16,000 x g for 10 minutes, an aliquot of the supernatant was transferred to a LC vial and diluted

20-fold in acetonitrile:methanol (75:25, v/v). A 10 µL volume was injected into the Agilent 1200 series Rapid Resolution LC system coupled to an Agilent 6440 Triple Quadrupole Mass

Spectrometer. Separations were performed on a Waters Xbridge HILIC (150×4.6 mm, 5 μm) column with an isocratic elution at a flow rate of 0.5 mL/min at room temperature. The mobile phase was acetonitrile/water (70:30 v/v) with 10 mM ammonium acetate. Electrospray mass spectrometry (ESI-MS) was performed with the scan mode set to multiple reaction monitoring

(MRM) targeting GroPCho (258.0 104.0), choline (104.1  60.1), and choline-d9 (113.1 

69.1) in fast switch ionization mode. The MS parameters were set as previously optimized: 10eV collision energy, 3.5 kV capillary voltage, 50 V fragmentor voltage, 200 ms dwell time, 300ºC drying gas temperature, and 325ºC sheath gas temperature. Sheath gas flow and drying gas flow rates were 10 and 8 L/min respectively. Each sample was run in duplicate. Data were acquired and analyzed using MassHunter Work Station software. The peak areas of the target metabolites were normalized to the internal standard and to the density of cells (OD600) from which metabolites were extracted.

RNA Extraction and real time qRT-PCR gene expression analysis. Cultures were grown in

Biosystems) per the manufacturer’s specifications. Message abundance was quantified by quantitative reverse transcriptase PCR (qRT-PCR) using the Verso™ SYBR® Green 1-Step QRT-

PCR ROX Kit (Thermo Scientific) on the Applied Biosystems StepOnePlus Real Time PCR

System. The primers used are shown in Table 4.3. Standard curves for each primer pair were optimized to determine acceptable primer concentrations (70-100 nM) and template concentrations

(15-25 ng/25 µL reaction). Reverse transcription was carried out at 50C for 15 minutes followed by 95C for 15 minutes for RT inactivation and polymerase activation. Amplification parameters were 40 cycles at 95C 15s, 54.5C 30s, and 72C 40s. Primer specificity was verified by melt curve analysis and visualization of amplicons by gel electrophoresis. Primer sets did not produce amplicons when RNA from the respective deletion strains was used as template. No template control (NTC) and minus reverse transcriptase (-RT) reactions were performed to verify the absence of genomic DNA contamination in the RNA samples and/or the reagents. RNA from at least three independent cultures of all strains was analyzed in triplicate reactions per primer set.

Data were normalized to the endogenous control gene, SNR17, and analyzed using the comparative

∆∆CT approach (100,101).

Table 4.3 Nucleotide sequences of primers used for qRT-PCR Gene Name Primer Sequence 5' - 3' SNR17 Forward TTG ACT CTT CAA AAG AGC CAC TGA SNR17 Reverse CGG TTT CTC ACT CTG GGG TAC PLB1 Forward GCA TAC ACC AAG GAG GCT TTG PLB1 Reverse GAG TGG ATA GCA AGG AAG TGT CAC

Construction of the chromosomal PLB1-I-3xHA allele. The plasmid pPMY-3xHA (169), kindly provided by Nancy Hollingsworth (Stony Brook University), was used as the template to amplify a 3xHA-URA3 module for insertion into the genome between amino acids 30 and 31 of

Plb1. Because Plb1 is predicted to be GPI anchored to the plasma membrane, insertion at this point would avoid cleavage of the HA tag from either end since both the N-terminal localization signal and C-terminal GPI anchor attachment site are cleaved during processing and GPI anchor attachment. Insertion did not interfere with the predicted catalytic site for Plb1. The primers used to amplify the 3xHA-URA3 module are listed in Table 4.2. The PCR product was transformed into

WT and ypk1∆ strains and transformants were selected on YNB plates lacking uracil. To verify integration at the correct location DNA was extracted and used as template with two sets of primers. Verified transformants were grown in YPD liquid medium to allow for recombination and loss of the URA3 marker. Cells that had lost the URA3 marker were selected on YNB plates containing 5-FOA. DNA was extracted from the 5-FOA resistant colonies and used as a template for PCR. Strains producing the expected amplicon size, indicating insertion of the 3xHA and loss of the URA3, were named JPV 767 (WT + PLB1-I-3xHA) and JPV 745 (ypk1∆+ PLB1-I-3xHA).

These strains therefore contain DNA encoding an internal 3xHA tag between nucleotides 90 and

91 of PLB1 in their genomic DNA. The function of Plb1 containing the 3xHA tag was verified by observing the expected Plb1 activity in PC turnover/GroPCho production experiments.

Protein extraction and sucrose gradient separation. For total protein extraction from cell lysates, cultures were grown in YNB+75 µM inositol medium to log phase (OD600 ≈ 0.7). Cells were collected by centrifugation of a 125 mL culture and the supernatant was removed for evaluation of secreted protein. The supernatant was concentrated 100-fold using an Amicon Ultra

30K centrifugal filter (Millipore). The cell pellet was resuspended in 200 µL SUME protein lysis

buffer (1% SDS, 8 M urea, 10 mM MOPS, 10 mM EDTA, 1 mM AEBSF, 1X Yeast/Fungal

Protease Arrest™ G-biosciences 786-333), and lysed with glass beads by eight 1-minute alternate cycles of agitation on a Vortex mixer and incubation on ice. Centrifugation at 2000 x g for 5 minutes removed unbroken cells and debris. Protein concentration in the concentrated supernatant and the cell lysate was determined by Bradford assay kit (Thermo Fisher).

Sucrose gradient separation was performed as described (177,178). A 100 mL culture was grown to log phase and 2 mL of 0.5 M NaN3 was added at time of harvest. The cells were collected by centrifugation and broken by glass bead lysis in a 10mM Tris-HCl, pH 7.5, 1mM EDTA, 10% sucrose solution. Cell lysates were cleared of debris by low speed centrifugation and the resulting lysate was layered onto an 11 mL gradient of 30-70% sucrose in the lysis buffer. The gradient was centrifuged at 100,000 x g in a Beckman SW41 rotor for 18 hours at 4ºC. One mL fractions were collected from the top of the gradient by pipetting. Proteins of interest were analyzed by SDS-

PAGE and Western blot analysis.

Western blot analysis. Total protein extracts were analyzed by SDS-PAGE and Western blotting. Equal amount of protein was loaded onto NuPAGE Bis-Tris gels (Invitrogen, IM-8042) and transferred to a nitrocellulose membrane using the XCell Blot Module (Invitrogen).

Membranes were incubated in blocking buffer (phosphate buffered saline (PBS) containing 5% casein in 0.1M NaOH, 5% BSA, phenol red, and NaN3) for 30 minutes at room temperature.

Primary antibody against HA-Plb1 (monoclonal antibody HA-11, Covance catalog number MMS-

101P) was added to the solution at a 1:10,000 dilution and incubated overnight at 4ºC. Primary antibody was removed and the membrane was washed in PBS. The membrane was incubated in blocking buffer, without NaN3, containing a 1:125,000 dilution of secondary antibody (HRP- labeled affinity–purified antibody to mouse IgG (H+L), KPL catalog number 074-1806) for 1 hour

at room temperature. The membrane was washed in PBS and incubated with SuperSignal West

Femto extended duration substrate reagent (Peirce, #34095) before exposure on either a Kodak

440CF Image Station or exposure to Kodak Biomax MR Film for visualization.

Sucrose gradient fractions were analyzed by SDS-PAGE and Western blot analysis as described for total protein extracts, but equal volumes of each fraction were run on the SDS-PAGE gel instead of equal amounts of protein. Membranes were probed with primary antibodies against

HA-Plb1, Sec61 (1:1,000), and Pma1 (1:4,000). Anti-Pma1 was provided by Dr. Carolyn

Slayman, Yale University, and anti-Sec61 was obtained as described (179). The blots were then incubated with the appropriate secondary antibody, HRP-conjugated anti-rabbit or HRP- conjugated anti-mouse. For quantification of Western blots, densitometry was carried out using the Kodak 1D software.

Statistical Analysis. GraphPad Prism was used for statistical analysis by Student’s t – test or

ANOVA as indicated.

4.3 Results

4.3.1 Plb1 mediated extracellular GroPCho production increases upon loss of YPK1.

Plb1 deacylates phosphatidylcholine to produce free fatty acids and extracellular glycerophosphocholine (GroPCho) (69) (Figure 4-2 C). To better define the link between Ypk1 and Plb1 activity, turnover of phosphatidylcholine (PC) and the concomitant production of the

GroPCho were examined. Cells grown to uniform labeling with C14-choline chloride were washed free of label and re-inoculated into fresh medium containing 10 mM choline at 37°C. The chase was performed at elevated temperature to maximize turnover, since the Ypk1 pathway is activated by heat stress (11,142) and because PLB1 expression is upregulated by elevated temperature (see

Figure 4-5a). These experiments were performed in both the BY4742 and YPH499 backgrounds because of slight differences in phenotypes observed in ypk1∆plb1∆ double mutants constructed

in the two backgrounds, as described below. At the indicated times, cells were harvested, the percentages of the total radioactivity in the extracellular, intracellular, and membrane fractions were determined, and the metabolites produced were identified. Radioactivity in the extracellular fraction, which is used to monitor in vivo Plb1 activity, consisted of C14-GroPCho and free C14- choline. As shown in Figure 4-2, deletion of YPK1 resulted in increased production of extracellular

GroPCho that is ameliorated by deletion of PLB1. This held true for both strain backgrounds, although the magnitude of the change varied from nearly 3-fold for YPH499 to roughly 5-fold for

BY4742. The rest of the counts at each time point were in either the membrane or the intracellular fraction (Table 4.4).

Examination of the intracellular fractions revealed there is a decrease in internal GroPCho production in both the ypk1∆ and the ypk1∆plb1∆ strains (Table 4.4). Internal GroPCho production occurs primarily through Nte1 mediated turnover (103), suggesting that there may be a trade-off between Nte1 and Plb1 activity upon loss of Ypk1. Loss of Ypk1 also appears to cause increased extracellular choline. The phospholipase D (PLD) that hydrolyzes PC into free choline and phosphatidic acid (180), Spo14, could be responsible for the increase. Although quantitative differences in the levels of choline metabolites were observed as a function of strain background, the general trends were consistent. Also, deletion of YKR2, the paralog of YPK1, did not affect

GroPCho production (Figure 4-2 B).

To confirm the results obtained through radiolabeling, and to obtain a more complete picture of the extracellular choline metabolites produced, we performed tandem mass spectrometry

(LC-MS/MS). Through radiolabeling, we were able to monitor the turnover of Kennedy pathway- derived PC synthesized during the labeling period. Through MS, we could monitor all GroPCho produced during the experiment through both the PE methylation pathway and through the

Kennedy pathway (9). Importantly, the MS data confirmed that GroPCho production rose upon deletion of YPK1, and that Plb1 plays a major role in mediating the increase (Figure 4-2 C). Using

MS, we also detected a decrease in extracellular GroPCho production by plb1∆ yeast as compared to WT cells that was not apparent through pulse-chase radiolabelling. As indicated by both the MS data and the radiolabeling data, Plb1 cannot be the only phospholipase responsible for extracellular

GroPCho production, as strains lacking Plb1 still produce GroPCho. In fact, a strain in which all of the known PLBs are deleted, plb123∆ nte1∆ exhibits greatly reduced, but not completely eliminated GroPCho production (Figure 4-2 C), suggesting that one or more deacylating phospholipases have yet to be identified.

4.3.2 Turnover of another glycerophospholipid, phosphatidylinositol, is unaffected by deletion of YPK1.

In a WT strain, GroPIns produced through the Plb3-mediated deacylation of PI is the major phospholipid metabolite found in growth medium (45). To monitor this metabolite, WT and ypk1∆ cells grown to uniform labeling with [H3]-inositol were chased with non-radiolabelled inositol, similar to the conditions used in the C14choline labeling experiments above. However, we observed no significant difference between WT and ypk1∆ strains in the rate of PI turnover or in the percentage of water soluble inositol-containing counts found in extracellular (WT 31% ± 1 and ypk1 25% ± 2) or intracellular (WT 22% ± 1 and ypk1 26% ± 3) fractions following a 5-hour chase.

A. External 4 hours, BY4742 B. External 4 hours, YPH499

35 35 14 [14C] GroPCho [ C] GroPCho

) ) 14 14

e 30 e 30

l [ C] Choline [ C] Choline

* s

l * l 25 25 a

n e 20 20

a 15 15

T 10

x T 10

( ( 5 5

0 0 WT plb1 ypk1 ypk1 WT plb1 ypk1 ypk1 ykr2 plb1 plb1

Figure 4-2. Plb1-mediated extracellular GroPCho production is increased upon deletion of YPK1. A and B, Strains in the BY4742 (A) and YPH499 (B) backgrounds were pre-grown at 30⁰C in YNB medium containing 0.2 Ci/mL [14C]-Choline chloride, followed by a chase at 37⁰C in YNB medium containing 10 mM choline. Extracellular, intracellular, and membrane fractions were isolated, and [14C]-choline- containing metabolites from extracellular fractions were separated by anion exchange column chromatography. Values are the means ± S.E. of triplicate cultures and are representative of multiple experiments. One-way ANOVA was performed indicating a significant difference between GroPCho produced in the ypk1∆ strain compared to GroPCho produced by each of the other strains listed (A, p ≤ 0.011; B, p ≤ 0.015). C, Cells grown to log phase were harvested and the extracellular medium analyzed for GroPCho by LC-MS. Relative abundance is normalized to OD600 and to the internal standard. Values are the means ± S.E. of at least two independent cultures. D, Plb1 mediated deacylation of PC to GroPCho.

Table 4.4 Cell Associated Counts from Figure 4.1 A. Cell Associated Counts in BY4742 (% of total counts) Time, Internal Internal Membrane Total Strain hours Choline GroPCho PC Counts 0 2 ± 0 34 ± 1 58 ± 3 94 WT 4 2 ± 0 81 ± 1 10 ± 0 93 0 1 ± 0 38 ± 1 61 ± 3 99 plb1∆ 4 1 ± 0 76 ± 1 15 ± 1 92 0 1 ± 0 31 ± 1 66 ± 3 98 ypk1∆ 4 1 ± 0 50 ± 4 18 ± 2 69 0 1 ± 0 35 ± 3 57 ± 4 92 ypk1∆ plb1∆ 4 1 ± 0 55 ± 3 24 ± 1 80

B. Cell Associated Counts in YPH499 (% of total counts) Time, Internal Internal Membrane Total Strain hours Choline GroPCho PC Counts 0 0 ± 0.2 7 ± 1 90 ± 1 97 WT 4 2 ± 0.4 14 ± 2 62 ± 2 82 0 2 ± 1 13 ± 1 82 ± 1 97 plb1∆ 4 1 ± 0.1 9 ± 4 63 ± 1 82 0 2 ± 1 11 ± 1 84 ± 1 97 ypk1∆ 4 2 ± 0.2 10 ± 2 53 ± 7 69 0 3 ± 1 15 ± 2 80 ± 3 98 ypk1∆ plb1∆ 4 3 ± 0.5 8 ± 2 68 ± 3 70 * Values for the internal and membrane metabolites are derived from the experiments described in Figs. 1A and 1B.

4.3.3 PC Synthesis via the Kennedy pathway is unaffected by loss of YPK1.

Since an increased rate of PC deacylation and GroPCho production was observed upon loss of Ypk1, we next tested if the rate of PC synthesis was affected. The logic underlying this experiment was that Cki1 (choline kinase) and Pct1 (cholinephosphate cytidylyltransferase), both of which are involved in PC synthesis by the Kennedy pathway, contain potential consensus Ypk1 phosphorylation sites [R-x-R-x-x-S/T-ϕ where ϕ is a preference for hydrophobic amino acids

(150)]. To detect changes in synthesis rate via the Kennedy pathway, we measured the incorporation of [C14]-choline into PC following a 30 minute pulse, but no difference was found

between WT and ypk1∆ yeast (Figure 4-3). This result also suggests that choline uptake was not significantly affected by loss of Ypk1.

12000

)

/ p 8000

(

h C 4000

]

[

0 WT ypk1

Figure 4-3. Synthesis of PC is not altered upon loss of Ypk1. WT and ypk1∆ strains in the BY4742 background were grown to log phase, harvested, and resuspended to equal densities in YNB medium. [C14]-choline was added and the amount of incorporation into PC after 30 minutes was determined. Values are the means ± S.E. of three independent cultures assayed in duplicate. Similar results were observed in the YPH499 background (data not shown).

In contrast, the Plb1 sequence lacks the Ypk1 consensus phosphorylation site. Furthermore,

Plb1 was shown to have a low likelihood value for phosphorylation by Ypk1 in a large-scale peptide screen for kinase activity (181). Similarly, the Plb1 sequence does not contain the Fpk1 phosphoacceptor site, RxSLD(x)15-21RxSLxD (142). Thus, direct regulation of Plb1 through phosphorylation by Ypk1 or its downstream kinase, Fpk1, is unlikely. In addition, a strain bearing deletion in FPK1 did not significantly alter GroPCho production. The percentages of total [C14]- choline label incorporated into external GroPCho in the fpk1 and WT strains following an eight hour chase, were 8 ± 1% and 5 ± 1%, respectively. Together, these data strongly suggest that Fpk1 is not involved in the regulation of Plb1 by Ypk1.

4.3.4 The secreted and cell associated forms of Plb1 increase in a ypk1∆ strain.

Plb1 activity has been reported in plasma membrane fractions, in the periplasm, and in the culture supernatant (45,69). However, localization data were absent in large-scale analyses using fluorescent tags fused to the C-termini of yeast ORFs expressed under control of their endogenous promoters (71-75), and

in a study utilizing C-terminal fusions to an epitope tag with the target genes expressed under control of the

GAL promoter (76). In addition, a localization screen of proteins involved in lipid metabolism in which

Plb1 was C-terminally tagged and expressed under the control of the TEF promoter, demonstrated ER and vesicle localization, but plasma membrane localization was not apparent; the medium/secreted fraction was not analyzed in this study (77). Complicating the question of Plb1 localization is that it has a predicted GPI- anchor, which should lead to plasma membrane residence (78,182). During processing, GPI-anchored proteins are cleaved just after the anchor attachment site, which removes the C-terminus (22). Therefore, if

Plb1 were GPI-anchored, C-terminal tagging—as used in several of the studies cited above—would be unable to detect the final processed protein.

To establish the localization of Plb1, and to have a reliable epitope-tagged Plb1 for Western analysis, we constructed a WT and ypk1∆ strain in which HA was inserted into the genomic copy of PLB1 near the N-terminal end, after the signal sequence, but before the catalytic site. This PLB1-

I-3xHA construct was judged functional by confirming that ypk1∆ containing PLB1-I-3xHA displayed elevated GroPCho production indistinguishable from the ypk1∆ strain (see

“Experimental Procedures”; data not shown). Consistent with increased GroPCho production, ypk1∆ containing the PLB1-I-3xHA allele displayed greater Plb1 abundance in total cell lysates and in the medium as compared to the WT strain ( Figure 4-4 A and B). To examine intracellular localization, we performed sucrose gradient fractionation using lysates from WT strains that contained the integrated, functional PLB1-I-3xHA gene. Interestingly, we found Plb1 associated with the ER (as indicated by co-migration with Sec61), as expected for a protein trafficking through the secretory pathway, but the protein was absent from plasma membrane fractions, as indicated by the migration of Pma1 (Figure 4-4C). These results, which are consistent with fluorescence microscopy of C-terminally tagged Plb1 under the control of the TEF promoter (77), lead us to conclude that the protein is not plasma membrane associated, and that if it is GPI-anchored, it is efficiently released

from the plasma membranes under the conditions used here. We suspect that the cell surface-associated

Plb1 activity that releases GroPCho into the medium is due to its periplasmic localization.

Figure 4-4. Plb1 protein abundance and localization. A, WT and ypk1∆ yeast containing the PLB1-I-3xHA allele were grown in YNB medium at 30⁰C to log phase. The supernatant was concentrated 100-fold, and total protein was extracted from the cells. Western blot was performed on both fractions using an anti-HA-Plb1 primary antibody and HRP-conjugated anti- mouse secondary. GAPDH was used as a loading control for the lysate fractions. B, Amount of Plb1 protein was quantified by densitometry of western blots and normalized to the culture density (OD600) and sample volume. Values represent the fraction of total Plb1 protein in the WT strain. C, WT yeast containing PLB1- I-3xHA were grown in YNB medium, lysed, and subjected to sucrose gradient centrifugation. The gradient was fractionated and the migration of Plb1 and the indicated marker proteins, Pma1 (plasma membrane) and Sec61 (ER), were determined by SDS-PAGE and Western blot analysis. Fraction number 1 on the left represents the top of the gradient.

Although Plb1 is predicted to be 72 kDa, the species found in the lysate migrates at about

145 kDa. While the 145 kDa band is the most abundant band in the medium, a less abundant, diffuse band around 260 kDa is also present. Plb1 is known to be N-glycosylated (183,184), and multiple glycosylated forms of secreted phospholipase B have been detected previously

(69,185,186). Upon treatment with Endo Hf, the protein migrates as one band at approximately 72 kDa (data not shown).

4.3.5 Compromised sphingolipid synthesis upregulates PLB1 expression and increases PC turnover.

Consistent with the observed increases in PC deacylation and Plb1 abundance (Figure 4-2 and Figure 4-4), PLB1 transcript levels also increased upon loss of YPK1 at both 30°C and after a

37°C temperature shift (Figure 4-5 A). In both WT and ypk1∆ yeast, a shift to 37°C increased PLB1 message levels. The ypk1∆ strain in the YPH499 background displayed a similar trend (data not shown).

The major role for Ypk1 described in the literature is the regulation of sphingolipid homeostasis (143). If the mechanism by which loss of Ypk1 upregulates Plb1 expression involves compromised sphingolipid synthesis, we would expect reduction of sphingolipid synthesis by other means to have a similar effect on Plb1 expression. Indeed, PLB1 expression was elevated in a WT strain treated with either myriocin or aureobasidin (Figure 4-5 B and C), which inhibit SPT and inositolphosphoceramide (IPC) synthase, respectively (187,188) (Figure 4-5 D). We also monitored PC turnover in WT cells treated with myriocin, and saw the expected increase in external GroPCho production (Figure 4-5E). Similar trends were observed in a YPH499 strain background. Sphingolipid synthesis can also be diminished genetically through the use of the lcb1-100 strain, which harbors temperature-sensitive SPT activity (170). As anticipated, the lcb1-

100 strain shifted to the restrictive temperature also exhibited increased PLB1 expression as compared to WT (Figure 4-5F).

A. B. C.

D. E.

Figure 4-5. PLB1 message levels are elevated by heat stress and upon disruption of sphingolipid homeostasis. A, Strains in the BY4742 background were grown to early log phase in YNB medium at 30°C. Cultures were split and incubated at 30⁰C or 37⁰C, as indicated, for 3 hours. B, Cells were grown in YNB medium at 30°C to an OD600 ≈ 0.7, myriocin (dissolved in methanol) was added to a final concentration of 1 µg/mL, and cultures were incubated for 2 hours at 30°C. C, Cells were grown in YNB medium at 30°C to an OD600 ≈ 0.4, aureobasidin (dissolved in methanol) was added to a final concentration of 1 µg/mL and the cells were incubated for 3 hours at 30°C. A-C, At harvest, total RNA was extracted and PLB1 transcript levels were analyzed by real-time qRT-PCR using the comparative CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT control cells. Values represent the means ± S.E. of at least 3 independent cultures assayed in triplicate. D, Simplified schematic of sphingolipid synthesis in S. cerevisiae including targets of inhibiting drugs. E, WT cells in the BY4742 background were pre-grown at 30⁰C in YNB medium containing 0.2 Ci/mL [14C]-Choline

chloride, followed by a chase at 30⁰C in YNB medium containing 10 mM choline and 1 g/mL myriocin in methanol or an equivalent volume of methanol. Extracellular, intracellular, and membrane fractions were isolated, and [14C]-choline containing metabolites from extracellular fractions after 3 hours were separated by anion exchange chromatography. Values are the means ± S.E. of at least duplicate cultures and are representative of multiple experiments. Similar trends were observed in the YPH499 background (data not shown). F, The lcb1-100 and WT strains were grown in YNB medium at room temperature (25°C) to an OD600 ≈ 0.4, then shifted to the restrictive temperature, 37°C, for 90 minutes. RNA was extracted and analyzed as in A-C. Student’s t-test was performed for statistical analysis (B, p = 0.05; C, p = 0.008; F, p = 0.0006).

4.3.6 Crz1 mediates PLB1 upregulation either when sphingolipid biosynthesis is compromised or when Ypk1 is absent.

A recent microarray study showed that many of the up/down regulated genes — including

PLB1 — identified in a ypk1∆ykr2∆ strain bearing an analogue inactivated allele of YPK1 (YPK1as) contain binding sites for the Crz1 transcription factor (151,152). Thus, we examined the role of

Crz1 in mediating PLB1 expression in response to altered sphingolipid biosynthesis or when YPK1 is deleted. A ypk1∆ mutant exhibited increased PLB1 message levels as compared to WT yeast, while PLB1 message levels were unchanged in a ypk1∆crz1∆double mutant (Figure 4-6 A). In addition, upregulation of PLB1 expression in response to myriocin treatment (Figure 4-5 B) observed in a WT strain did not occur in the crz1 strain (Figure 4-6 B). This data indicates that

Crz1 is required for the upregulation of PLB1 expression that occurs upon loss of Ypk1.

4.3.7 Ameliorating disrupted sphingolipid synthesis by deletion of ORM1 and ORM2 in a ypk1∆ strain partially rescues PLB1 expression to WT levels.

A primary mechanism by which Ypk1 regulates sphingolipid synthesis is through phosphorylation of Orm1 and Orm2 (13,189). Orm1 and Orm2 are ER transmembrane proteins that regulate sphingolipid biosynthesis by binding to and inhibiting SPT (13,158). Upon phosphorylation by Ypk1, the interaction between Orm1/2 and SPT is lessened, and sphingolipid biosynthesis is upregulated. Thus, if Ypk1 regulates Plb1 via an Orm1/2 dependent effect on sphingolipid synthesis, we would expect PLB1 expression to mirror WT levels in a

ypk1tsykr2∆orm1∆orm2∆ strain grown at the restrictive temperature. In fact, we found that deletion of ORM1/2 only partially reduced PLB1 expression compared to the ypk1tsykr2∆ strain (Figure

4-7). These results suggest that Ypk1 regulates sphingolipid synthesis via a mechanism that is partially independent of Orm1/2 phosphorylation.

25 A. B. WT crz1 30C 2.0

d 20

l 37C

L *

L 1.5

o o 15

s 10 i

o l 1.0

5 o

N x 0.5 0 E WT ypk1 crz1 ypk1 0.0 crz1 Methanol Myriocin

Figure 4-6. Crz1 mediates PLB1 upregulation either when sphingolipid biosynthesis is compromised or when Ypk1 is absent. A, Strains (BY4742) were grown in YNB medium at 30°C to an OD600 ≈ 0.7 (log phase). An aliquot was diluted to an OD600 ≈ 0.2 and grown for 3 hours at 37°C. B, WT and crz1 strains were grown in YNB medium at 30°C to an OD600 ≈ 0.7, myriocin (dissolved in methanol) was added to a final concentration of 1ug/mL, and incubated for 3.5 hours at 30°C. A and B, RNA was extracted and PLB1 message levels were analyzed by qRT-PCR using the comparative CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT cells grown at 30ºC (A) and WT cells grown with vehicle, methanol (B). Values represent the means ± S.E. of 3 independent cultures assayed in triplicate. Student’s t-test was performed for statistical analysis, p ≤ 0.0001.

5 *

1 4

o 3

m 2

E 1

0 ts ts WT ypk1 ypk1 ypk1 ykr2 ykr2 orm1orm2

Figure 4-7. Deletion of ORM1 and ORM2 in a ypk1 strain partially reduces PLB1 expression. Cells were grown in YNB medium at 30ºC to early log phase followed by a shift to 35ºC for 1.5 hours. Total RNA was extracted and PLB1 transcript levels were analyzed by real time qRT-PCR using the comparative ∆∆CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT cells. Values represent the means ± S.E. of 3 independent cultures assayed in triplicate. Student’s t-test was performed for statistical analysis, p ≤ 0.04.

4.3.8 Overexpression of only PLB1 rescues ypk1tsykr2∆ lethality at the restrictive temperature.

Plb1 was previously identified as a multicopy suppressor of lethality in the ypk1tsykr2∆ strain at the restrictive temperature of 37°C (147). To determine if rescue was PLB1-specific or could be conferred by other PLB-encoding genes, growth was monitored in the ypk1tsykr2∆ strain overexpressing PLB1, PLB2, PLB3, or NTE1. The genes were expressed from their endogenous promoters on multicopy plasmids (see “Experimental Procedures”). Even though there was a 200-

300-fold increase in PLB1, PLB2, and PLB3 message levels and a 50-fold increase in NTE1 message levels, only overexpression of PLB1 rescued ypk1tsykr2∆ lethality at the restrictive temperature (Figure 4-8).

Figure 4-8. Overexpression of only PLB1 rescues the lethality of the ypk1tsykr2 strain at restrictive temperature. A. WT and ypk1tsykr2 strains (YPH499) harboring the indicated plasmids were pre-grown in selective medium, harvested, and resuspended to equivalent densities. Cells from 5-fold serial dilutions of each culture were spotted onto selective medium and grown for 3 days at the permissive (30⁰C) and restrictive (35⁰C) temperatures.

4.3.9 Deletion of both PLB1 and YPK1 leads to synthetic interactions.

Based on the data presented in Figure 4-8, PLB1 and YPK1 appear to genetically interact.

To confirm this hypothesis, we examined specific phenotypes in strains lacking both genes. As shown in Figure 4-9A, a negative genetic interaction was observed between YPK1 and PLB1 when growth was monitored, although the severity of the defect varied somewhat depending upon the genetic background employed. On solid YNB medium, ypk1∆plb1∆ isolates constructed in both the BY4742 or YPH499 backgrounds displayed a growth defect as compared to ypk1∆ at 30ºC and

35ºC (Figure 4-9 A). The ypk1∆plb1∆ strain constructed in the YPH499 background also exhibited a growth defect on YPD plates (Figure 4-9B).

The Ypk1 pathway influences several downstream events, including cell wall integrity

(147,148). Therefore, we examined the effect of the cell wall perturbing agent, calcofluor white, on growth. A ypk1∆plb1∆ mutant exhibited increased sensitivity to calcofluor white as compared to ypk1∆ (Figure 4-9B). This result was most striking in the YPH499 background, where no growth was observed in the ypk1∆plb1∆ strain in the presence of calcofluor white when present at a final concentration of 8.5 µg/ml. These findings, together with the fact that PLB1 is a multicopy suppressor of the lethality of the ypk1tsykr2∆ strain, indicate that increased PC turnover via Plb1 activity increases the fitness of yeast when Ypk1 is absent. More generally, these findings suggest that sphingolipid synthesis is coordinated with PC turnover to maintain optimal lipid homeostasis.

Figure 4-9. The slowed growth phenotype and sensitivity to the cell wall perturbing agent, calcofluor white, in a ypk1 strain is exacerbated by deletion of PLB1. A, Strains in the BY4742 and YPH499 backgrounds were pre-grown in YNB medium, harvested, and resuspended to equivalent densities. Cells from 5-fold serial dilutions were spotted onto YNB plates and incubated for 48 (BY4742) or 32 hours (YPH499). C and D, WT and deletion strains were pre-grown in YNB medium, harvested, and resuspended to equivalent densities. 5-fold (BY4742) or 10-fold (YPH499) dilutions were spotted onto YPD and YPD + 8.5g/mL calcofluor white plates and incubated at 30⁰C for 32 or 48 hours, respectively. Data displayed is representative of multiple experiments.

4.4 Discussion

PC is the most abundant glycerophospholipid in eukaryotic membranes and its synthesis and catabolism are highly regulated. In yeast, the known routes of PC catabolism occur through the action of the phospholipase D encoded by PLD1/SPO14 (180), the ER localized PLB encoded by NTE1 (103), and PLB1 (69). Here we show that Plb1 activity (Figure 4-2) and expression

(Figure 4-5, Figure 4-6) were modulated by the serine/threonine kinase Ypk1. Furthermore, we show that regulation occurs via a mechanism in which the cell can detect compromised sphingolipid biosynthesis (Figure 4-6). Blockage of SPT activity through use of an lcb1-100 mutant or by treatment with myriocin also results in increased PLB1 expression. Treatment of cells with aureobasidin A, which blocks downstream steps in the sphingolipid biosynthetic pathway, led to a similar effect, indicating that the formation of the complex sphingolipid, IPC, at a minimum, is required to regulate PLB1 (Figure 4-6).

Plb1 has been widely accepted as a secreted and plasma membrane-associated protein based on cellular fractionation studies and its role in the production of extracellular GroPCho (69).

Our results, utilizing a Plb1 construct with an internal HA tag to account for a potential GPI anchor, indicate that Plb1 is secreted (Figure 4-5A) and fractionates with the ER marker, Sec61, but not with the plasma membrane marker, Pma1 (Figure 4-5C). As a secreted and glycosylated protein, it is not unexpected to find Plb1 at the ER. However, our data along with that of others suggest the intriguing possibility that Plb1 may be active intracellularly as well as at the cell surface, since it has been shown to metabolize excess intracellular lysophospholipids (190) and to be involved in

PC acyl chain remodeling (191). The secreted/periplasmic form of Plb1 is most likely responsible for the production of extracellular GroPCho. Notably, extracellular GroPCho is not a dead-end metabolite, but can be recycled into PC synthesis following its uptake via glycerophosphodiester transporters that have been identified in both S. cerevisiae (61) and C. albicans (104,106). 96

Ypk1 regulates sphingolipid synthesis by phosphorylating Orm1/2 (143). Phosphorylated

Orm1/2 cannot bind to SPT, resulting in upregulated sphingolipid synthesis. However, our results show that the increase in PLB1 expression observed in a ypk1∆ strain is not entirely mediated by

Orm1/2, since PLB1 transcript levels are not depressed to WT levels in the ypk1tsykr2∆orm1∆orm2∆ strain (Figure 4-8). This result suggests that Ypk1 may affect sphingolipid synthesis via a mechanism that goes beyond regulation of Orm1/2 activity.

Alternately, deletion of Orm1/2 may have additional consequences that impinge upon Plb1 expression and phospholipid metabolism. For example, an orm1∆orm2∆ strain is an inositol auxotroph, indicative of disrupted regulation of phospholipid metabolism (158). Furthermore, auxotrophy can be reversed by growth in the presence of myriocin, indicating a connection between sphingolipid and phospholipid metabolism upon loss of Orm1/2 (158). These data suggest a complicated relationship between Orm1/2 and its role in regulating sphingolipid and phospholipid metabolism.

Phosphorylation of the lipid flippase activator, Fpk1, by Ypk1 downregulates the Dnf1 and

Dnf2 flippases (142). In addition, Fpk1 has been linked to elevated levels of ROS that are produced upon loss of Ypk1 (154). Our results do not indicate a connection between Plb1 activity and Fpk1, as a fpk1∆ strain does not display increased PC turnover and Plb1 does not have a Fpk1/2 phosphoacceptor site. We also discovered that the calcineurin dependent responsive element

(CDRE) transcription factor, Crz1, is a key regulator of the increased PLB1 expression observed upon disruption of sphingolipid biosynthesis. Interestingly, Crz1 is not responsible for increased

PLB1 transcription observed upon heat stress (Figure 4-7A), suggesting multiple levels of PLB1 regulation, depending on the applied stress. Consistent with our findings, a ypk1∆ykr2∆ strain bearing a YPK1as allele displays increased expression of several genes, many that have Crz1

binding sites in their promoters, including PLB1 (151). Based on these and other data, a model was proposed in which the TORC2 and calcineurin pathways act antagonistically to maintain the optimal cell growth and survival (151). When Ypk1 is recruited to the plasma membrane by

Slm1/2, there is a shift toward downregulation of the calcineurin pathway and upregulation of

TORC2 signaling. Therefore, in a ypk1∆ strain, activation of the calcineurin pathway would predominate, activating the downstream transcription factor, Crz1, which in turn activates various

CDRE genes, including PLB1 (Figure 4-10). Interestingly, treatment of cells with myriocin results in TORC2-dependent phosphorylation of Ypk1 in a manner that requires the Ypk1 recruiting proteins, Slm1/2, to translocate from eisosomes to membrane compartments containing TOR

(152,153). Since Slm1 relocalization was suggested to be a result of an altered sphingolipid environment, and an increase in ROS (154), it is tempting to speculate that alterations in the eisosome lipid environment due to Plb1-mediated PC turnover may also contribute to Slm1 relocalization.

Our phenotypic analysis of the ypk1∆plb1∆ double mutant suggests that upregulated Plb1 activity upon loss of Ypk1 is a functionally important compensatory mechanism, since slowed growth and sensitivity to calcofluor white are exacerbated upon deletion of PLB1 in the ypk1∆ strain (Figure 4-9). Two lines of evidence indicate Plb1 activity, as opposed to activity of the other

PLBs, mediates this phenomenon: i) only overexpression of PLB1 rescues growth of the ypk1tsykr2∆ strain at restrictive temperature (Figure 4-8), and ii) turnover of PI, the primary substrate of Plb3, is not altered in the ypk1∆ strain.

Figure 4-10. Model of PLB1 transcriptional activation upon loss of Ypk1 or upon disruption to sphingolipid biosynthesis by the presence of myriocin. The left panel displays events occurring in a WT cell where the TORC2/Ypk1 signaling pathway predominates resulting in a basal level of PLB1 trnscription and basal level of Plb1-mediated PC turnover. The right panel displays event occurring upon loss of YPK1 where the Calcinuerin signaling pathway predominates causing Crz1 dephosphorylation and resultant elevation of PLB1 trnscript levels and elevation of Plb1-mediated PC turnover. The right panel (gray) also displays elevation of Crz1 activated PLB1 transcription in the presence of myriocin. The dotted line indicates movement TORC2, Target or rapamyacin complex 2; SPT, serine palmitoyl transferase.

In summary, our results indicate that Ypk1, in addition to regulating sphingolipid synthesis and plasma membrane flippase activity, also regulates the turnover of the major phospholipid, PC.

Future studies will focus on the mechanism by which altered sphingolipid biosynthesis, as engendered by loss of Ypk1, activates the myriad of genes under the control of Crz1, such as Plb1.

Acknowledgements – We thank Jeremy Thorner and Yusef Hannun for the gifts of yeast strains and Nancy Hollingsworth, Susan Henry, and Chris McMaster for the gifts of plasmids.

*This work was supported by the National Institute of Health Grants R15 GM104876 to JPV, and

R01 GM75061 and P30 DK79307 to JLB.

Note: Appendices A 6 - A 16 contain additional data from Chapter 4 that were excluded from the submitted manuscript.

Chapter 5 Summary and Future Directions

5.1 Summary of results

My dissertation work has resulted in novel contributions to our understanding of the B- type phospholipases in S. cerevisiae. First, Ira1 and Ira2 were implicated in regulation of both the production of extracellular GroPIns and its transport into the cell. However, I was unable to implicate the Ras/cAMP pathway in this regulation suggesting that other undefined functions of

Ira1/2 might be involved. Second, examination of various aspects of the GDTR pathway revealed loss of the cell surface associated PLBs, Plb1-3, and/or utilization of GroPIns as a phosphate source results in abnormal actin cytoskeleton and an increased cell size. Regarding the reutilization aspect of the GDTR pathway, I was unable to determine if reacylation of GroPIns can occur in the cell.

Third, I identified a novel interaction between Ypk1 and Plb1 demonstrating that in addition to regulating sphingolipid synthesis and plasma membrane flippase activity, Ypk1 also regulates the

Plb1-mediated turnover of the major phospholipid, PC.

Additionally, my work has suggested many new avenues that could be pursued in order to better understand the relationship between phospholipid turnover via phospholipases of the B- type, sphingolipid metabolism, and the Ypk1 kinase. Here, I discuss what I feel are the most pressing open questions and future directions that could be pursued.

5.2 How does alteration in sphingolipid biosynthesis result in activation of Crz1?

I have documented a novel role for Ypk1 in regulation of Plb1-mediated PC turnover. This data adds to the published findings indicating that Ypk1 contributes to the maintenance of both sphingolipid and phospholipid homeostasis. Ypk1 does not appear to regulate Plb1 by direct kinase

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activity, but instead Plb1 activity is elevated in ypk1∆ yeast due to the disruption of sphingolipid metabolism (Figure 4-5, Figure 4-6). This disruption causes transcriptional upregulation of PLB1 in a Crz1-dependent manner.

To identify genes involved in Crz1-mediated gene regulation, a screen of selected deletion mutants bearing a plasmid (CDRE plasmid) containing a promoter region with binding sites for the transcription factor Crz1 linked to the lacZ gene could be utilized. This plasmid has been constructed by other researchers and is available (11). Transcriptional induction could be monitored by measuring β-galactosidase activity upon treatment of the cells with myriocin or another drug disrupting sphingolipid metabolism. Mutants to be tested would include those that have a link to calcium metabolism, since Crz1 is a calcineurin/calcium-activated transcription factor. For example, strains lacking the plasma membrane calcium channel complex, Mid1/Cch1, and the vacuole cation channel Yvc1, both of which allow calcium into the cytoplasm, are prime candidates to be tested (192).

5.3 Are other genes in phospholipid synthesis affected by disrupted sphingolipid synthesis?

Since Ypk1 appears to be a major regulator of lipid homeostasis, additional genes transcriptionally regulated by loss of Ypk1 and/or disruptions to sphingolipid biosynthesis could be identified. Genes of interest may include genes containing potential Crz1 binding sites in their promoters or genes involved in some aspect of lipid metabolism that I believe are linked to Ypk1 function. Several genes of interest involved in lipid metabolism that also have Crz1 binding sites in their promoter regions include: ORM2, SCS2, CHO2, NTE1, INO1, OPI3, and LAC1

(YEASTRACT), while other genes of interest involved in lipid metabolism include SPO14 and

GDE1. Specifically, measuring message levels by qRT-PCR of the selected genes could be performed in two sets of samples: i) in a wild type strain as compared to a ypk1∆ mutant and ii) in

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wild type cells either treated or not treated with the sphingolipid synthesis inhibiting drug, myriocin.

5.4 Does Plb1 play a role in Slm1 relocalization from eisosomes to MCT domains leading to TORC2 and Ypk1 activation?

As discussed in Chapter 4, the antagonism between the TORC2 (promotes cell growth) and the calcineurin (promotes cell survival) pathways suggests a potential explanation for the increase in Plb1-mediated turnover upon loss of Ypk1 (151,152,193), with cell survival and Crz1 activation predominating in the absence of Ypk1. The Slm proteins must be translocated from eisosomes to the plasma membrane MCT in order for Ypk1 to be recruited to the plasma membrane for activation by TORC2 (153). Recent literature suggests that elevated levels of ROS are the essential signal for Slm1 relocalization to the MCT (154), and others have demonstrated that depletion of sphingolipids, hypo-osmotic stress, and mechanical membrane stress all cause Slm1 relocalization from eisosomes to MCT patches (152). While it has been suggested that sphingolipid levels may be important for Slm1 movement, it would be interesting to see if Plb1 and PC levels also play a role. For these experiments localization of fluorescently tagged Slm proteins could be compared in a WT and plb1∆ strains with and without myriocin treatment. Ypk1 phosphorylation by TORC2 could also be monitored in these strains. If Slm protein localization is altered in plb1∆ cells, ROS levels could be compared in WT, ypk1∆, plb1∆, and ypk1∆plb1∆ strains since ROS is the proposed essential mediator of Slm protein relocalization. Fluorescently tagged Slm constructs are available from other labs (153), and published procedures exist for measuring Ypk1 phosphorylation (153) and ROS abundance (154). Further understanding of how lipids regulate the Slm proteins and the resultant Ypk1 activation by TORC2 is of interest because both TORC2 and ROS are elevated in cancer cells (154).

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5.5 Is the plasma membrane lipid composition altered in YPK1/PLB1 deficient cells?

Evaluation of the plasma membrane phospholipid composition in the YPK1/PLB1 mutants should be completed to better understand if phospholipid composition itself, in addition to sphingolipids, may influence Ypk1 signaling and/or cellular processes. Lipid composition specifically in the plasma membrane, ER, and vacuole fractions of WT, ypk1∆, plb1∆, and ypk1∆plb1∆ strains could be compared. Each membrane fraction can be isolated by published techniques and lipids – phospholipids, sphingolipids, neutral lipids, and sterols – can be extracted and their abundances quantified. Particular lipid composition has been found to be essential for some plasma membrane enzymes (194,195). Turnover events, through the action of the B-type phospholipases and other enzymes continually causes alterations to the plasma membrane lipid composition and is equally important. While my data indicates that PC turnover is altered in the ypk1∆and ypk1∆plb1∆ strains, turnover of the other major glycerophospholipids, phosphatidylinositol (PI), phosphatidylserine (PS), and phosphatidylethanolamine (PE) and the concomitant production of their metabolites, GroPIns, GroPSer, and GroPEtn could be thoroughly analyzed. Furthermore, more rigorous evaluation of methylation pathway synthesized PC turnover is also warranted.

5.6 Is the functionality of model plasma membrane proteins, 1,3-β-glucan synthase, Git1, Pma1, and Can1, altered in YPK1/PLB1 deficient cells?

Disrupted lipid regulation occurring upon YPK1 deletion may in turn abolish the proper lipid environment required for activity of essential plasma membrane enzymes. Phospholipase activity has been shown to alter plasma membrane enzyme function including lysophospholipase overexpression dependent inhibition of amiloride-sensitive epithelial sodium channel (ENaC) activity in mammalian cells (196), or PLA2 activity and lysophospholipid inhibition of the 1,3-β- glucan synthase in yeast (197). My preliminary experiments have identified several potential

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enzymes whose activities are altered upon deletion of PLB1 and YPK1, including 1,3-β-glucan synthase (Appendix A 17), Git1 (plasma membrane GPI transporter protein) (Appendix A 19), and

Pma1 (plasma membrane ATPase) (Appendix A 20), and an enzyme whose activity is not altered,

Can1 (arginine transporter) (Appendix A 21).

Additional enzymes of interest, particularly spanning localization in different membrane compartments could be identified and analyzed for altered function. Upon verification that enzyme activity is indeed altered, the level of control causing the alterations in enzyme activity could be determined: transcription (mRNA levels), post-transcriptional (protein levels), activity, and/or localization. These protein activities could be expected to be involved in phenotypes reported in this dissertation. Furthermore, if these proteins or others are verified to have altered activity upon loss of YPK1 and/or PLB1, future studies could evaluate the requirement for specific lipid environments for their optimal functionality by employing synthetic liposomes.

5.7 Do the B-type phospholipases play a role in deacylation of the phosphorylated phosphoinositides in vivo?

The roles of the PIPs have been extensively studied in yeast and higher eukaryotes demonstrating their key regulatory role in fundamental eukaryotic cell processes such as endo/exocytois, membrane trafficking, cell division, actin organization, and in the localization of proteins that bind to lipids, such as those with PH domains (31). PIP catabolism mostly occurs through the action of PLCs, but it can also occur by the action of PLA2 and/or PLB enzymes with concomitant production of the GroPIPs, lysophospholipids, and free fatty acids, with the GroPIPs being the least studied metabolite (51). Glucose addition to stationary phase S. cerevisiae cultures was shown to result in release of GroPIns, GroPI[4]P, and GroPI[4,5]P2 into the extracellular medium (59). It is known that deacylation reactions are one method of PIP turnover in yeast (45).

In addition, the literature suggests that both PIPs and their deacylation products, the GroPIPs, play

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a role in actin cytoskeleton function. Thus, the potential role of the PLBs in this process is obvious considering my work showing that strains with PLB1-3 deleted have an increased sensitivity to the actin destabilizer, Lat A, as well as have an abnormal actin organization in the presence of

Latrunculin A. Examination of PLB activity on the PIPs is also warranted for their potential role in the Ypk1-Plb1 signaling interaction as the Slm proteins have PH domains that are required for

Ypk1 recruitment to the plasma membrane. Thus, I propose that determining if the PLBs are active against the PIPs in vivo is warranted.

Although reproducible detection of the GroPIPs has proven difficult since they are physiologically present at low concentrations, new techniques and dyes specific to PIPs may aid in future PIP/GroPIP analyses. HPLC or LC/MS analysis of GroPIns and the GroPIPs would need to be employed. Fusion of known PIP binding domains to specific flares tagged with GFP have been constructed to use in detection of PtdIns[4,5]P2 and PtdIns[4]P (198,199) and Annexin V conjugated to Alexa Fluor 488 is available for specific binding to PS (200). Complete analysis of the PIPs and GroPIPs in the YPK1 and PLB1-3 mutant strains would require development and optimization of additional techniques to monitor these metabolites at their physiologically relevant concentrations in vivo.

5.8 Final conclusions about the importance of PLBs

The experiments completed thus far provide novel insights into the roles and regulation of the B-type phospholipases, and how regulation of lipid metabolism and turnover can affect many cellular processes. Long-term goals could include further elucidating the mechanism by which

Ypk1 regulates Plb1, and developing an understanding of how this regulation impacts global cellular processes such as, cell wall integrity, actin dynamics, eisosome assembly, endocytosis, and general cell health. The outcomes of the studies would further our understanding of how lipid metabolism is integrated with other important aspects of cellular physiology in a eukaryotic cell. 105

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Appendix A

Preliminary data from exploratory experiments pertaining to or referenced in Chapter 3 are found in Appendices A 2 - A 5. Appendices A 6 - A 16 contain data collected as part of the project described in Chapter 4, but were not included with the submitted manuscript. No narrative will be included with those figures. Additional preliminary experiments performed by me that were not pursued further, or that will be continued by other members of the lab are included in Appendices

A 17 - A 21 and are referred to in Chapter 5 for discussion purposes.

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A 1. Genes with predicted or observed phospholipase activity in S. cerevisiae. (Gene descriptions from SGD, yeastgenome.org) GeneID GENE Aliases Description from Saccharomyces Genome Database Mitochondrial cardiolipin-specific phospholipase; functions upstream of Taz1p to generate monolyso-cardiolipin; transcription increases upon genotoxic stress; involved in restricting Ty1 853007 CLD1 YGR110W transposition; has homology to mammalian CGI-58

Protein with a role in maintaining mitochondrial morphology; also involved in maintaining normal cardiolipin levels; mitochondrial inner membrane protein; proposed to be involved in N- 856001 FMP30 YPL103C acylethanolamine metabolism; related to mammalian N-acylPE-specific phospholipase D Inositol phosphosphingolipid phospholipase C; mitochondrial membrane localized; hydrolyzes complex sphingolipids to produce ceramide; activates genes required for non-fermentable carbon source metabolism during the diauxic shift; activated by phosphatidylserine, cardiolipin, and 856739 ISC1 YER019W phosphatidylglycerol; mediates Na+ and Li+ halotolerance Peroxisomal matrix-localized lipase; required for normal peroxisome morphology; contains a peroxisomal targeting signal type 1 (PTS1) and a lipase motif; peroxisomal import requires the PTS1 receptor, Pex5p and self-interaction; transcriptionally activated by Yrm1p along with genes 854251 LPX1 YOR084W involved in multidrug resistance; oleic acid inducible Serine esterase, phospholipase B; homolog of human neuropathy target esterase (NTE); Nte1p- mediated phosphatidylcholine turnover influences transcription factor Opi1p localization, 854943 NTE1 YML059C affecting transcriptional regulation of phospholipid biosynthesis genes Protein of the endoplasmic reticulum; required for GPI-phospholipase A2 activity that remodels YCR044C, the GPI anchor as a prerequisite for association of GPI-anchored proteins with lipid rafts; 850411 PER1 COS16 functionally complemented by human ortholog PERLD1 Phosphatidyl Glycerol phospholipase C; regulates the phosphatidylglycerol (PG) content via a phospholipase C-type degradation mechanism; contains glycerophosphodiester 855895 PGC1 YPL206C phosphodiesterase motifs

Phospholipase B (lysophospholipase) involved in lipid metabolism; required for efficient acyl chain remodeling of newly synthesized phosphatidylethanolamine-derived phosphatidylcholine; required for deacylation of phosphatidylcholine and phosphatidylethanolamine but not 855020 PLB1 YMR008C phosphatidylinositol; PLB1 has a paralog, PLB3, that arose from the whole genome duplication Phospholipase B (lysophospholipase) involved in lipid metabolism; displays transacylase activity 855018 PLB2 YMR006C in vitro; overproduction confers resistance to lysophosphatidylcholine

Phospholipase B (lysophospholipase) involved in lipid metabolism; hydrolyzes phosphatidylinositol and phosphatidylserine and displays transacylase activity in vitro; PLB3 854151 PLB3 YOL011W has a paralog, PLB1, that arose from the whole genome duplication Phospholipase C; hydrolyzes phosphatidylinositol 4,5-biphosphate (PIP2) to generate the signaling molecules inositol 1,4,5-triphosphate (IP3) and 1,2-diacylglycerol (DAG); involved in regulating many cellular processes; Plc1p and inositol polyphosphates are required for acetyl- 855860 PLC1 YPL268W CoA homeostasis which regulates global histone acetylation Meiosis-specific prospore protein; required for meiotic spindle pole body duplication and separation; required to produce bending force necessary for proper prospore membrane assembly 855720 SPO1 YNL012W during sporulation; has similarity to phospholipase B

Phospholipase D; catalyzes the hydrolysis of phosphatidylcholine, producing choline and phosphatidic acid; involved in Sec14p-independent secretion; required for meiosis and spore formation; differently regulated in secretion and meiosis; participates in transcription initiation and/or early elongation of specific genes; interacts with "foot domain" of RNA polymerase II; YKR031C, deletion results in abnormal CTD-Ser5 phosphorylation of RNA polymerase II at specific 853902 SPO14 PLD1 promoter regions Multifunctional lipase/hydrolase/phospholipase; triacylglycerol lipase, steryl ester hydrolase, and Ca2+-independent phospholipase A2; catalyzes acyl-CoA dependent acylation of LPA to YKR089C, PA; required with Tgl3p for timely bud formation; phosphorylated and activated by Cdc28p; 853964 TGL4 STC1 TGL4 has a paralog, TGL5, that arose from the whole genome duplication Putative protein of unknown function; localizes to the membrane fraction; possible Zap1p- regulated target gene induced by zinc deficiency; YJL132W is a non-essential gene; contains 853309 YJL132W YJL132W domain for Zinc dependent phospholipase C Acyl-protein thioesterase responsible for depalmitoylation of Gpa1p; green fluorescent protein (GFP)-fusion protein localizes to both the cytoplasm and nucleus and is induced in response to YLR118C, the DNA-damaging agent MMS ; contains conserved domain for Abhydrolase 2, 850809 YLR118C APT1 Phospholipase/Carboxylesterase Putative carboxylic ester hydrolase; similar to bovine phospholipase A1; the authentic, non- 854187 YOR022C YOR022C tagged protein is detected in highly purified mitochondria in high-throughput studies

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A 2. Loss of the cell surface associated PLBs results in invaginations of the cell membrane when grown in media containing GroPIns as the sole phosphate source. A, WT (JPV 13) and plb123∆ (JPV 480) were grown on YNB I+ LoPi and YNB I+ NoPi +200 µM GroPIns agar plates. A sample of the cells from each condition were analyzed by SEM. B, Magnified image of plb123∆ in GroPIns media Arrows point to cells displaying large invaginations. Photos courtesy of Dr. Donna Stolz, University of Pittsburgh.

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A 3. Exposure to LatA alters distribution of actin patches and cables upon loss of PLB1-3. Cells grown as described in Fig. 3-14. TRITC-phalloidin staining of the actin cytoskeleton in the absence (A) and presence (B) of Latrunculin A (LatA). Visualized by the Zeiss Axio Observer Z1 confocal microscope, 100x 1.4NA objective. Images obtained with HCImage software.

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A 4. Alterations in PLB message levels in log phase compared to stationary phase. Total RNA was extracted from harvested cells followed by DNase treatment. Transcript levels were analyzed by real-time RT-PCR using the comparative ∆∆CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT cells in log phase. Values represent the mean ± S.E. of at least 3 individual cultures each assayed in triplicate.

A 5. Deletion of LGE1 and PLB3 results in slowed growth and decreased colony size. WT (JPV 399), plb3∆, lge1∆ (JPV 637), and lge1∆plb3∆ patched to single colonies on YPD plates at 30ºC (A) and growth curve in YPD media at 30ºC (B). In the lower panels (A), two independent isolates are streaked from single colonies.

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A 6. Protein Architecture upon construction of the PLB1-I-3HA allele. (Supplement to Experimental Procedures, HA-allele). The HA tag was inserted into the PLB1 gene without interference with the localization signal, the GPI signal, and the catalytic site.

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A 7. Plb1 mediated external glycerophosphocholine production is increased upon deletion of YPK1 at multiple time points in the BY4742 background. (Supplement to Fig. 4.1A and Table 4.1A). Separation of metabolites from external, internal, and membrane fractions is shown for multiple time points, after 0, 2, 4, and 13 hours of chase. Values are the means ± S.E. of at least triplicate cultures.

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A 8. Plb1 mediated external glycerophosphocholine production is increased upon deletion of YPK1 at multiple time points in the YPH499 background. (Supplement to Fig. 4.1B and Table 4.1B). Separation of metabolites from external, internal, and membrane fractions is shown for multiple time points, after 0, 4, and 8.5 hours of chase. Values are the means ± S.E. of at least triplicate cultures.

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A 9. Kennedy pathway synthesis of phosphatidylcholine is not altered upon loss of Ypk1. (Supplement to Fig. 4.3). Strains in the YPH499 background were grown to log phase in YNB 75 µM inositol media. [C14]-choline was added to equivalent amounts of cells in YNB media and the amount of incorporation into PC after 30 minutes was determined. Lipids were extracted and [C14]-choline incorporation was measured by LSC (A) and verified to be incorporated into PC by TLC (B). A non- radioactively labeled standard for PC was visualized by staining the TLC plate with Thioflavin S solution. Values are the means ± S.E. of three independent cultures assayed in duplicate.

A 10. Deletion of the flippase activators, Fpk1 or Fpk2, or deletion of LSP1 or PIL1, involved in eisosome formation do not alter extracellular GroPCho production. (Supplement to Chapter 4). Strains in the BY4742 background bearing the indicated gene deletions were pre-grown at 30⁰C in YNB media containing 0.2µCi/mL [14C]-Choline chloride, followed by a chase in YNB media containing 10 mM choline at 37⁰C. Extracellular, intracellular, and membrane fractions were separated from harvested cells at the listed time points. Separation of [14C]-choline containing metabolites from extracellular and intracellular fractions was performed using anion exchange column chromatography, but no differences from WT were observed, except in the ypk1∆ strain as previously discussed. Data shown is the mean ± S.E. of at least duplicate cultures. 129

A 11. Overexpression of the plasma membrane flippase, DNF1, does not result in altered extracellular GroPCho production. Supplement to Chapter 4. WT (BY4742) was transformed with pRS423 (JVE 338) and pRS423-DNF1 (JVE 339) and pre-grown at 30⁰C in YNB media lacking uracil containing 0.2µCi/mL [14C]-Choline chloride, followed by a chase in YNB media containing 10mM choline at 37⁰C. Extracellular, intracellular, and membrane fractions were separated from harvested cells at the listed time points. Separation of [14C]- choline containing metabolites from extracellular and intracellular fractions was performed using anion exchange column chromatography. Data shown is the mean ± S.E. of triplicate transformants.

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A 12. Secreted and cell-surface associated Plb1 protein abundance in log and stationary phase is elevated in a ypk1∆ mutant. (Supplement to Figure 4-4). A, WT + PLB1-I-3xHA and ypk1∆ + PLB1-I-3xHA were grown in YNB media at 30⁰C to log and stationary phase. Supernatant was concentrated 100-fold, while total protein was extracted from the cells. Western blot was performed on all fractions using an anti-HA-Plb1 primary antibody and HRP-conjugated anti-mouse secondary. Note there is less protein in the stationary phase samples than the log phase samples.

A 13. Regulation of PLB1, PLB2, and PLB3 message levels upon loss of Ypk1 or upon loss of transcription factors, Crz1, Msn2, and Msn4. Strains in the BY4742 background were grown to early log phase in YNB media at 30°C. Total RNA was extracted and PLB transcript levels were analyzed by qRT-PCR using the comparative ∆∆CT method. Data were normalized to the amount of the endogenous control mRNA, SNR17, and expressed relative to the value in WT cells. Values represent the mean ± S.E. of three cultures assayed in triplicate. Results of interest from this experiment include upregulation of PLB3 in ypk1 cells compared to WT, however a concurrent increase in Plb3-mediated GroPIns production was not observed (Chapter 4). Msn2 and Msn4 did not appear to regulate PLB1 or PLB3 under the growth conditions tested, but may be involved in depressing PLB2 message. Crz1 appears to have regulate PLB1 and possibly PLB3 message levels.

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A 14. Compromised sphingolipid synthesis by the presence of myriocin results in increased extracellular GroPCho production. (Supplemental to Fig. 5E.) Separation of metabolites from external, internal, and membrane fractions is shown for multiple time points, after 0, 3, and 6 hours of chase. Values are the means ± S.E. of at least triplicate cultures.

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A 15. Overexpression of the PLB genes on a multicopy plasmid elevates corresponding PLB message levels. (Supplemental to Figure 4-8). A, WT cells transformed with the PLB overexpression plasmids, YEp351- PLB1, YEp351-PLB2, YEp351-PLB3, and pRS426-NTE1, were grown to log phase in selective media at 30°C. Total RNA was extracted and PLB1, PLB2, PLB3, and NTE1 transcript levels were analyzed by real- time qRT-PCR using the comparative ∆∆CT method. Message levels were normalized to the endogenous control, SNR17, and expressed relative to the message in corresponding WT + empty vector. Values represent the mean ± S.E. of at least 3 individual transformants assayed in triplicate. 2-way ANOVA showed no statistical difference between values for PLB1, PLB2, or PLB3, but expression of NTE1 was significantly different.

A 16. The slowed growth phenotype and sensitivity to the cell wall perturbing agent, calcofluor white, for a ypk1∆ strain is exacerbated by deletion of PLB1. (Supplemental to Figure 4-9). Strains in the YPH499 (A and B) and BY4742 (C and D) backgrounds were grown in YNB media containing 75 µM inositol at 30⁰C and 37⁰C. Data represent the mean and standard error of at least three replicates. E, Cells from 10-fold serial dilutions of strains in the YPH499 background were spotted onto YPD, YPD + 0.5 M NaCl, and YPD + 8.5 µg/mL calcofluor white plates and incubated at both 30⁰C and 35⁰C for 3 days. None of the strains were able to grow on YPD + 8.5 µg/mL calcofluor white plates at 35⁰C. Data displayed is representative of multiple experiments.

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A 17. 1,3-β glucan synthase activity is depressed upon deletion of PLB1 and YPK1. Plasma membrane isolated from WT (JPV 399), plb1 (JPV 668), ypk1 (JPV 636), and ypk1plb1 (JPV 669) grown in YNB I+ media to log phase (OD600 ≈ 0.7) at 30C, harvested, washed with 1 mM EDTA, and assayed for 1,3-β-glucan synthase activity according to published procedures (201-203). Acid washed glass beads (2 mL) and 8 mL of cold Breaking Solution (0.5 M NaCl, 1 mM EDTA, 1 mM PMSF) were added to cell pellet on ice. Vortex and incubations on ice were done alternately for 1-2 minutes and repeated 10 times. Cell lysate was transferred to a fresh tube and beads were washed with an aliquot of breaking buffer and added to the lysate. Centrifuging 5 minutes at 1500 x g at 4⁰C cleared unbroken cell and debris. Supernatant (membrane fraction) was transferred to ultracentrifuge tubes and breaking solution was added up to top of tube. Membranes were pelleted at 100,000 x g for 30 minutes at 4⁰C. Pellet was resuspended in 1mL of membrane buffer (50 mM Tris-HCl pH = 7.5, 1 mM EDTA, 1 mM β-mercaptoethanol, 33% glycerol) and homogenized on ice with the Dounce homogenizer. Protein concentration was determined by Bradford assay. 20µg of protein was assayed for 1 hour at 30°C in a reaction containing 2 µL of 0.1 M UDP-glucose, 1.25 µL of UDP-[14C]-glucose (0.02 µCi/mL), 3 µL of 10% BSA, 2 µL of 0.5 M KF, 3 µL of 10mM EDTA, 4µL of 1mM GTP, and water up to 40µL. The reaction was stopped by addition of 1mL 10% TCA and filtered through glass fiber (GF/C, Whatman 1822-025). Filters were washed with three 1 mL aliquots of 10% TCA followed by two 1 mL washes with 95% ethanol. Incorporation of [C14]-UDP- glucose into cells was quantified by liquid scintillation counting. Data represent mean ± S.E. of four independent cultures. The ypk1plb1 1,3-β-glucan synthase activity is from multiple colonies of only one deletion isolate.

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A 18. Altered expression of the catalytic subunit of 1,3-β glucan synthase, Fks2. WT (JPV 399), plb1 (JPV 668), ypk1 (JPV 636), and ypk1plb1 (JPV 669) transformed with pFKS2- lacZ (JVE 336) were grown to log phase (OD600 ≈ 0.8) in YNB –URA media at 30ºC and 37ºC. Harvested cells were assayed for β-gal activity as described in Experimental Procedures (Chapter 2.3.1). Data represent the mean ± S.E. of four independent transformants.

A 19. Activity of the plasma membrane protein, Git1, is depressed upon loss of Ypk1 and/or Plb1. WT (JPV 399), plb1 (JPV 668), ypk1 (JPV 636), and ypk1plb1 (JPV 669) grown in YNB I- 200µM 3 KH2PO4 media were harvested and assayed for their ability to transport [H ]-GroPIns in a 6 minute assay at 30ºC as described in Materials and Methods (Chapter 2). Data represent the mean ± S.E. of three independent cultures assayed in duplicate.

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A 20. Pma1 activity is not lost by deletion of PLB1 and/or YPK1. WT (JPV 399), plb1 (JPV 668), ypk1 (JPV 636), and ypk1plb1 (JPV 669) cells grown in YNB I+ media to log phase at 30ºC were harvested and membranes isolated as described in Galan et al.(204). Sodium azide was added to culture, cells were pelleted and washed, and resuspended in lysis buffer (0.1 M Tris-HCl pH = 7.5, 0.15 M NaCl, 5 mM EDTA, protease inhibitor mix) to a concentration of 50 ODU/0.2 mL. Chilled glass beads were added to meniscus of suspension and vigorously agitated with a vortex between incubations on ice. Lysate was removed to a fresh tube and beads were washed with aliquot of lysis buffer that was added to cell lysate. Centrifugation at 3,000 rpm (should have been slower because likely lost Pma1 at this speed) removed unbroken cells and debris. Supernatant was removed as the PM enriched fraction and centrifuged for 45 minutes at 12,000 rpm x g. Pellet containing membranes was resuspended in 10X diluted lysis buffer. Protein concentration was measure by Bradford assay. 40 µg of protein was assayed for ATPase activity as described in Koland et al.(205). Protein sample was added to 1 mL of ATPase assay buffer (50 mM MES, 5 mM MgCl2, 5 mM NaN3, pH = 6 with Tris) with and without 100 µM orthovanadate and pre-incubated for 5 minutes at 30ºC. To start assay 20 µL of 0.1 M ATP (sodium salt) was added and incubated for 10 minutes at 30ºC. The reaction was stopped with 2mL of stop solution (2% sulfuric acid, 0.5% ammonium molybdate, 0.5% SDS). The product corresponding to ATPase activity, phosphomolybdate, was reduced by addition of 20 uL of 10% ascorbic acid. The reaction sat for 5minutes at RT before absorbance was read at 750 nm. Blanks containing no enzyme and no ATP were run to subtract out background. Data trends are representative multiple experiments.

- Arginine - Arginine + Canavanine

plb1∆ WT ypk1∆plb1∆ ypk1∆ can1∆ WT

A 21. Sensitivity to canavanine is not altered upon YPK1 and/or PLB1 deletion. Indicates Can1 arginine transport is not compromised in these strains. WT (JPV 399), plb1 (JPV 668), ypk1 (JPV 636), ypk1plb1 (JPV 669), WT (JPV 523), ypk1 (JPV 768), ypk1tsykr2 (JPV 771) and can1 cells were grown in YNB media, diluted to OD600 = 1.0 in water. Four 10-fold serial dilutions were performed and 5 µL of each dilution was spotted onto YNB – arginine plates ± 7.5 µg/mL canavanine. Plates were incubated at 30ºC and 37ºC for 48 hours.

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