The Glucose Signal Transduction Pathway Mediated by the Rgt2 and Snf3 Glucose Sensors in the Budding Yeast Saccharomyces Cerevisiae

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The Glucose Signal Transduction Pathway Mediated by the Rgt2 and Snf3 Glucose Sensors in the Budding Yeast Saccharomyces cerevisiae

by Adhiraj Roy

B.Sc. in Microbiology, May 2005, University of Calcutta, India M.Sc. in Microbiology, May 2007, University of Calcutta, India

A Dissertation submitted to

The Faculty of The Columbian College of Arts and Sciences of The George Washington University in partial fulfillment of the requirements for the degree of Doctor of Philosophy

May 18, 2014

Dissertation directed by

Jeong-Ho Kim Assistant Professor of Biochemistry and Molecular Medicine

The Columbian College of Arts and Sciences of The George Washington University certifies that Adhiraj Roy has passed the Final Examination for the degree of Doctor of

Philosophy as of March 18, 2014. This is the final and approved form of the dissertation.

The Glucose Signal Transduction Pathway Mediated by the Rgt2 and Snf3 Glucose Sensors in the Budding Yeast Saccharomyces cerevisiae

Adhiraj Roy

Dissertation Research Committee:

Jeong-Ho Kim, Assistant Professor of Biochemistry and Molecular Medicine, Dissertation Director

William Weglicki, Professor of Biochemistry and Molecular Medicine, Committee Member

Wenge Zhu, Assistant Professor of Biochemistry and Molecular Medicine, Committee Member

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Dedication

I dedicate my dissertation to my family, my grandparents, (Late) Sourindra K. Roy and

Mrs. Jyotsna Roy, my parents, Mr. Soven Roy and Mrs. Mamata Roy and my beloved sister Ms. Sanjana Roy.

Acknowledgements

This dissertation would not have been possible without the help of so many people in so many ways. It is also a product of a large measure of serendipity, fortuitous encounter with people who changed the course of my academic career. First and foremost, I express my deepest gratitude to my mentor, Dr. Jeong-Ho Kim for providing me the opportunity of being his graduate student and working on this wonderful project. As my

“Academic Father”, Dr. Kim inculcated in me the flavor of yeast genetics and molecular biology. His constant guidance, support and encouragement helped me to think “out of the box”, analyze biological phenomenon critically and finally resulted in the growth of my scientific intellect.

I would also like to thank my committee members, Dr. Rakesh Kumar, Dr. William

Weglicki, Dr. Wenge Zhu and Dr. Paul Brindley for their insightful comments and constant encouragement. A special thanks goes to Dr. Linda Werling, Program Director,

IBS for her endless support throughout my graduate career.

I will be always indebted to my former colleague Dr. Yong Jae Shin who has been like a mentor to me and taught me many aspects of molecular biology and I am grateful for his constant support. Dr. Shin, Gamsahapnida! Furthermore, I thank my colleague Dr.

Yong Bae Kim and Dr. Sujit Nair, Dr. Kazufumi Oshiro from Dr. Rakesh Kumar’s lab for their help and our departmental administrative staffs including Debby, Nichole and

Laura. Thank you all.

The 5 year long journey for the quest of knowledge would not have been possible without unconditional love and support of my friends. Ananda Banerjee, you have been my best friend ever and I thank you for being with me in the ups and downs of this long

v journey. Somenath Chakraborty, how can I forget your support and encouragement which were always needed in my tough times? Kankana, Shubham, your support has always been with me, I thank you all for everything.

I must take a moment to express my heartily gratitude to Dr. Ananta K. Das, former

Chair, Microbiology department of my undergraduate college at The University of

Calcutta. Sir, you have taught me everything, from preparing bacterial media to immunohistochemistry. It was you who has been my inspiration all the time and I am indebted to you forever. I am so lucky to have you as my mentor during the foundation period of my academic career.

Most importantly, I would like to thank my family for their unconditional love and moral support. I am forever grateful to my parents for their commitment and dedication they showered upon me in every hardship of my journey. Ma, Bapi, you are the best! My little sister missed me for last few years but nevertheless, always encouraged me. I am blessed to have a sibling like Sanjana. I love you. I thank my grandmother, uncles, aunts and all my cousins for their encouragement. Without their support, I could not have gone this far.

Abstract of Dissertation

The Glucose Signal Transduction Pathway Mediated by the Rgt2 and Snf3 Glucose Sensors in the Budding YeastSaccharomyces cerevisiae

Sensing and signaling the presence of extracellular glucose is crucial for yeast

Saccharomyces cerevisiae because of its fermentative metabolism, characterized by high glucose flux through glycolysis, mediated by expression of glucose transporter genes

(HXTs). Yeast cells mediate aerobic glycolysis in part, by the crosstalk between two glucose signaling pathways: 1) the Rgt2/Snf3 glucose induction pathway and 2) the

Snf1/Mig1 glucose repression pathway. The yeast Rgt1 repressor inhibits transcription of

HXT genes in the absence of glucose by recruiting general transcription repressor

Ssn6/Tup1 and the HXT corepressor Mth1. In response to glucose, Rgt1 is phosphorylated by the cAMP-activated protein kinase A (PKA) and dissociates from the

HXT promoters and no longer interacts with Ssn6/Tup1, resulting in the HXT gene expression. Glucose regulates Rgt1 function by primarily modulating the Rgt1-

Ssn6/Tup1 interaction and Rgt1 removal from DNA occurs but not necessary for expression of HXT genes. Ssn6/Tup1 interferes with Rgt1 DNA-binding activity in the absence of Mth1, and that Rgt1 function abrogated by Ssn6 overexpression is restored by cooverexpression of Mth1. Mth1 acts like a scaffold-like protein to facilitate interaction between Rgt1 and Ssn6/Tup1 by blocking PKA-dependent Rgt1 phosphorylation. The cell surface glucose receptors Rgt2 and Snf3 and the glucose transporter (Hxt1) are stable and functional only in the presence of glucose but are removed from the plasma membrane through ubiquitination and subsequent vacuolar degradation via endocytosis in the absence of glucose.

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Table of Contents

Dedication…………………………………………...…………………………………....iv

Acknowledgements………………………………………………………………………..v

Abstract…………………………………………………………………………………..vii

Table of Contents……………………………...... ………………….………………viii

List of Figures………...... …………………………….….……………………xi

List of Tables...... xiv

List of Symbols/Abbreviations…………………………………………………...……...xv

Chapter I:

Introduction………………………………………………………………….....……….…1

1.1.Background…………………………………………………………………....1

1.2. Aerobic Glycolysis in Yeast...... 4

1.3.Key Components of the HXT Gene repressor Pathway...... 5

1.4. Key Components of the HXT Gene Induction Pathway……………………...8

1.5. Crosstalk Between Glucose Induction and Repression Pathways…………..12

1.6. Research Significance…………………………………………………....….15

Chapter II: Understanding the Mechanism of Glucose-induced Relief of Rgt1- mediated Repression of HXT Gene Expression in Yeast...... …....20

2.1 Introduction………………………………………………….…...... …….….21

2.2. Materials and Methods………………………………………………...... …23

2.3. Results……………………………………………………………....……….26

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2.4. Discussion…………………………………………………………………...32

Chapter III: Mth1 Regulates the Interaction between the Rgt1 Repressor and

the Ssn6-Tup1 Corepressor Complex by Modulating PKA-dependent

Phosphorylation of Rgt1...... ….40

3.1 Introduction………………………………………….....……………...….….41

3.2. Materials and Methods…………………………………....……...….……...44

3.3. Results………………………………………………………...... ….…...... 48

3.4. Discussion………………………………………………………………...... 59

Chapter IV: Endocytosis and Vacuolar Degradation of the Yeast Cell Surface

Glucose Sensors Rgt2 and Snf3………………………………………………...... …68

4.1 Introduction…………………………...…………….…………...….……….69

4.2. Materials and Methods………………………………………...………....…71

4.3. Results………………………………………………………………..….….74

4.4. Discussion……………………………………………………………...…...85

Chapter V: Glucose Starvation-induced Turnover of the Yeast Glucose

Transporter Hxt1…………………………………………………………...……...….....93

5.1 Introduction………………………………………………….…………...... 94

5.2. Materials and Methods……………………………………………...……...96

5.3. Results…………………………………………………………………...…99

5.4.Discussion…………………………………………………...... …….106

Conclusions and Future Direction……………………………………………...... …...110

References……………………………………………..………………………...... …112

Appendix I: Construction of Yeast Strains Useful for Screening Drugs that

Inhibits Glucose Uptake and Glycolysis…………………………………..…...... …..132

A1.1 Introduction…………….……………………………………..……..………..132

A1.2 Construction and Properties of the HXT-NAT Reporter Strains…………...... 133

A1.3 Conclusion…………………………………………………....……....………137

List of Figures

Figure 1. The Aerobic Glycolysis, a Hallmark of Cancer...... 3

Figure 2. Aerobic Glycolysis in Yeast…………………………….…………...... ……5

Figure 3. Structures of Glucose and Related Hexoses………….……………………...... 9

Figure 4. Rgt2/Snf3 Mediated Glucose Induction Pathway of HXT

Gene Expression….…………………………………...... …………...... ………….13

Figure 5. Schematic Diagram of the Crosstalk between Glucose Signaling

Pathways in Yeast...... …...15

Figure 6. Dissociation of the Rgt1 Repressor From HXT Promoters is Not Required

for Glucose-induction of HXT1 Gene Expression…………..……………………...... …28

Figure 7. Mth1 Does Not Directly Regulate the DNA-binding Ability of Rgt1…...... 30

Figure 8. Rgt1 Phosphorylation at the PKA Sites is Required For Glucose- induction of HXT Gene Expression…………...... …………...... …...35

Figure 9. The Interaction of Rgt1 With Ssn6-Tup1 is Critically Regulated by its

Phosphorylation State……………………………………...... ….…36

Figure 10. Glucose Regulates the Function of the Two Major Glucose Responsive

Repressors Rgt1 and Mig1 in a Similar Manner………………….……...... …37

Figure 11. qRT-PCR Analysis of HXT1 Gene Expression…………….………...... …37

Figure 12. Rgt1-HA and LexA-Rgt1 Have Different Affinities for Ssn6-Tup1….....…...38

Figure 13. Ssn6-Tup1 Negatively Regulates the Ability of Rgt1 to Bind to its

Target Promoters……………………………………………...... …49

Figure 14. Ssn6 Overexpression Induces Derepression of HXT Expression in

Repressing Conditions…………………………...... …...... ……..……53

Figure 15. Mth1 is Required For the Interaction of Rgt1 With Ssn6-Tup1...... ….…57

Figure 16. Rgt1 Function is Regulated Positively and Negatively by Mth1 and

Ssn6, Respectively……………………...... ……………...... …58

Figure 17. Rgt1 Phosphorylation by PKA Leads to the Disruption of its Interaction

With Ssn6-Tup1…………...……………...... …63

Figure 18. Ssn6-Tup1 is Recruited to the HXT1 Promoter in an Rgt1-dependent

Manner...... 64

Figure 19. Repression of the MTH1 Gene by the Mig1-Ssn6-Tup1 Complex…...... …65

Figure 20. Rgt1-S5A-HA Constitutively Represses Expression of the HXT1 Gene….....65

Figure 21. Ssn6 Overexpression Abrogates the DNA-binding Ability of Rgt1, But

Not of Mig1…………………………………...... ……...66

Figure 22. Rgt2 is Degraded in the Absence of Glucose…………………………...... 75

Figure 23. Glucose Starvation-induced Degradation of Rgt2 is not Dependent on

Proteasome……………………………………………………...... ……….76

Figure 24.Rgt2 Undergoes Endocytosis and Subsequent Vacuolar Degradation in

Glucose Starved Ccells……………………………………...... ………78

Figure 25. Snf3 Levels are Regulated by both Transcriptional and

Translational Mechanisms………………………...... …………...……...... ……....80

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Figure 26. Ubiquitination of the Cytoplasmic Tail Domain of Rgt2 is Required

for its Endocytosis………………………………………...... …...... ………..82

Figure 27. Constitutively Active Rgt2-1 and Snf3-1 Glucose Sensors do not

Undergo Endocytosis...... 86

Figure 28. Signaling Defective Rgt2 Glucose Sensor is Constitutively

Endocytosed……………………………………………………………………....……...87

Figure 29. Glucose Regulation of the Yeast Glucose Sensors………………...... …..91

Figure 30. Hxt1 Protein Levels are Posttranslationally Downregulated in

Response to Glucose Starvation……………………………………...... 100

Figure 31. Glucose Starvation Induces Endosytosis and Subsequent Vacuolar

Degradation of Hxt1…………………………...... …....102

Figure 32. The N-terminal Cytoplasmic Domain of Hxt1 is Required for

Turnover…...... 107

Figure 33. Hxt1 is Ubiquitinated by the Ubiquitin Ligase Rsp5…………………....….108

Figure 34. K12 and K39 Serve As Putative Ubiquitin Acceptor Lysine Residues….....109

Figure 35. The HXT-NAT Reporter Strains Exhibit Strict Growth Dependence on Glucose…………………………………………...... …....136

Figure 36. 2-DG Inhibits Growth of the HXT-NAT Reporter Strains…………...... 137

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List of Tables

Table 1.1. S. cerevisiae strains used in the study of Chapter 2…………...... 39

Table 1.2. Plasmids used in the study of Chapter 2...... 39

Table 2. S. cerevisiae strains used in the study of Chapter 3...... 67

Table 3.1. S. cerevisiae strains used in the study of Chapter 4...... 92

Table 3.2. Plasmids used in the study of Chapter 4...... 92

TABLE 4.1: S. cerevisiae strains used in the study of Chapter 5...... 97

TABLE 4.2: Plasmids used in the study of Chapter 5...... 97

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List of Symbols/Abbreviations

AMPK AMP-activated Kinase

ChIP Chromatin Immunoprecipitation

CHX Cycloheximide

CK1γ Casein Kinase 1-gamma

DAPI 4´,6-diamidino-2-phenylindole

DMSO Dimethyl Sulfoxide

ECL Enhanced Chemiluminiscence

EDTA Ethylene Diamine Tetracaetic Acid

GFP Green Fluorescence Protein

GLUT Glucose Transporter

HA Hemeagglutinin

HXT Hexose Transporter

IB Immunoblot

IP Immunoprecipitation

NADH Nicotinamide Adenine Dinucleotide (Reduced)

NAT Nourseothricin Acetyl Transferase

ORF Open reading Frame

PCR Polymerase Chain Reaction

PET Positron Emission Tomography

PKA Protein Kinase A

PVDF Polyvinylidene Fluoride

TAP Tandem Affinity Purification

TCA Trichloro Acetic Acid

TPR Tetratrico Peptide Repeat

TZD Thiazolidinediones qRT-PCR Quantitative Real Time PCR

WB Western Blot

2DG 2-deoxy-Glucose

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CHAPTER I

INTRODUCTION

1.1. Background

Unicellular organisms such as microbes reproduce as quickly as possible when nutrients are available and their metabolic control systems have evolved to sense an adequate supply of nutrient and consume the requisite carbon, nitrogen and free energy for generating building blocks to produce new cells (Vander Heiden et al., 2009). In multicellular organisms, most cells are exposed to constant supply of nutrient and survival of the organism requires control systems that prevent aberrant individual cell proliferation when nutrient availability exceeds the level needed to support cell division.

In order to proliferate, cells must obtain necessary energy required for many vital processes such as macromolecular biogenesis, DNA replication and cell structure maintenance. Glucose serves as a metabolic substrate as well as a signaling molecule that governs a wide variety of physiological and pathological processes (Busti et al., 2010;

Ozcan and Johnston, 1999; Rolland et al., 2001b; Santangelo, 2006; Towle, 2005).

Mammals maintain a constant level of glucose in the bloodstream despite intermittent supplies from the gut, and impaired regulation of blood glucose levels results in severe disorders including diabetes (Busti et al., 2010; Kolb, 2012; Towle, 2005). In the presence of oxygen, normal cells primarily metabolize glucose to carbon dioxide by oxidation of glycolytic pyruvate in the mitochondrial tricarboxilic acid (TCA) cycle, which produces NADH [nicotinamide adenine dinucleotide (NAD+), reduced] and fuels oxidative phosphorylation to maximize ATP production (Figure 1). In contrast, metastasized tumor cells metabolize large amounts of glucose through glycolysis and

1 produce copious amount of lactic acid even in the presence of oxygen (Hsu and Sabatini,

2008; Vander Heiden et al., 2009), called the Warburg effect, a hallmark of cancer

(Warburg, 1956). In spite of the decrease in energy yield as a consequence of the

“glycolytic phenotype”, this seems to allow an increase in cell proliferation rate and be applicable to other fast growing cells (Brand, 1997). The well-established elevated glucose consumption of malignant tissue forms the basis of the clinical imaging of cancer, PET (positron emission tomography) (Mandelkern and Raines, 2002).

The budding yeast Saccharomyces cerevisiae has the ability to adapt its metabolic profile to the environmental conditions. When glucose level is high, the yeast carries out

“aerobic glycolysis” as its main metabolic pathway (the Crabtree effect) (Crabtree, 1929;

Lagunas, 1979) and when glucose is scarce, it can switch to oxidative metabolism

(Thevelein, 1994). Regarding energy metabolism, yeast and tumor cells share several similarities. In both cell types, the glucose-induced downregulation of oxidative metabolism is observed along with an enhanced fermentation despite the presence of oxygen (Diaz-Ruiz et al., 2009). As tumor cells, yeast overexpress all glycolytic enzymes in response to glucose (Entian et al., 1984; Takeda, 1981) and the activity and/or expression pattern of glycolytic enzymes are also modified in yeast (Entian et al., 1984;

Muratsubaki and Katsume, 1979). Expression of human glucose transporters (GLUTs) is upregulated by Hif-1α in tumor cells (Ebert et al., 1995). The high affinity glucose transporters Glut1 and Glut3 are overexpressed in several cancer cell lines (Macheda et al., 2005). Yeast also upregulates expression of glucose transporters (HXTs) to increase glucose uptake (Boles et al., 1997). Snf1/AMPK is a low energy checkpoint, acting as the prime energy sensor in response to energy depletion, and its activation mechanism is

2 conserved in eukaryotes and actively involved in glucose uptake and metabolism. Several current diabetes therapeutics, such as metformin and thiazolidinediones (TZDs), are thought to lower blood glucose and increase insulin sensitivity by activating AMPK in peripheral tissues (Shaw et al., 2005). Further understanding glucose sensing and signaling mechanisms in yeast may reveal the molecular basis of enhanced glucose uptake in cancer cells and provide clues for developing therapeutic strategies.

Figure 1. The aerobic glycolysis, a hallmark of cancer. In the presence of oxygen, normal cells metabolize glucose to pyruvate via glycolysis and then oxidize most of the pyruvate in the mitochondria (mt) to CO2 by oxidative phosphorylation. In oxygen- limiting condition, cells convert pyruvate to lactate primarily by anaerobic glycolysis. Tumor cells convert most glucose to lactate regardless of the presence of oxygen (aerobic glycolysis), a phenomenon called “Warburg effect”. Aerobic glycolysis is less efficient than oxidative phosphorylation producing only 2 ATPs per glucose molecule (Vander Heiden et al., 2009)

1.2. Aerobic glycolysis in yeast

Glucose is by far the preferred energy source of the budding yeast S. cerevisiae,

because glucose regulation dictates the organism’s distinctive fermentative lifestyle,

aerobic ethanol fermentation (the Crabtree effect) This is mediated, in part, by the

crosstalk between two glucose signaling pathways: 1) the Rgt2/Snf3 glucose induction

pathway responsible for glucose uptake (Johnston and Kim, 2005; Kaniak et al., 2004;

Ozcan and Johnston, 1999); 2) the Snf1/Mig1 glucose repression pathway that negatively

regulates the genes involved in glucose oxidation and the use of alternative sugars

(Figure 2) (Carlson, 1999; Gancedo, 1998; Hedbacker and Carlson, 2008). The poorly

understood mechanistic basis of Warburg effect in tumor cells shows some remarkable

similarities to the mechanism responsible for this phenomenon in yeast. Since energy

generation by fermentation is insufficient, yeast pumps a large amount of glucose through

glycolysis by expressing glucose transporters (HXT) genes and therefore, enhancing the first rate-limiting step of glucose metabolism---its uptake (Boles and Hollenberg, 1997;

Ozcan and Johnston, 1999). S. cerevisiae possesses at least six members of glucose transporter family (Hxt1, 2, 3, 4, 6 and 7) each with different affinities for glucose in order to cope with environmental changes in glucose availability (Boles and Hollenberg,

1997; Reifenberger et al., 1997). Yeast possesses cell surface glucose receptors Rgt2 and

Snf3 which generate intracellular glucose signal upon binding to glucose, to upregulate expression of HXT genes. In this study, I examined the crosstalk between the glucose signaling pathways that leads to induction of HXT gene expression in yeast, as a model to investigate how cells sense glucose and adapt their gene expression program to glucose availability.

Figure 2. Aerobic glycolysis in yeast. Yeast cells uptake extracellular glucose by glucose transporters (Hxts) and convert it to pyruvate via glycolysis. Due to its fermentative lifestyle, yeast cells prefer to further convert pyruvate to carbon dioxide and ethanol rather than oxidizing it in mitochondria (mt) by electron transfer chain (ETC). Because fermentation produces only 2 ATP molecules per glucose molecule compared to ~36 ATPs in glucose oxidation, yeast cells consume glucose vigorously by upregulating glucose transporter genes (HXTs). Glucose itself induces glycolysis by glucose induction pathway and represses glucose oxidation and metabolism of other alternative carbon sources by glucose repression pathway (Ozcan and Johnston, 1996).

1.3. Key components of the HXT gene repressor pathway

In the absence of glucose, the expression of HXT genes is repressed by a multi- protein repressor complex formed on the HXT promoter element, consisting of the HXT repressor Rgt1, glucose responsive corepressor Mth1/Std1 and general transcription repressor complex Ssn6-Tup1 (Flick et al., 2003; Kim et al., 2006; Kim et al., 2003;

Lakshmanan et al., 2003; Malave and Dent, 2006; Ozcan and Johnston, 1999; Ozcan et al., 1996b; Smith and Johnson, 2000b) (Figure 4). Each of the components of the HXT

gene repression pathway are briefly discussed below.

1.3.1. The Rgt1 repressor

Rgt1 is a member of the Gal4 family of transcription factors containing the zinc

binuclear cluster (Cys6Zn2) DNA-binding domain (Ozcan et al., 1996b). Unlike most

members of the family that bind as dimer to two ‘CGG’ triplets, Rgt1 lacks the coiled-

coil dimerization domain and thus binds DNA as a monomer to a specific sequence

‘CGGANNA’ (Kim et al., 2003). This synergistic repression is probably due to efficient

recruitment of Rgt1 to multiple binding sites and hence, Rgt1 as a monomer, functions

more efficiently through its multiple sites (Kim, 2009). This is reminiscent of the

Aspergillus nidulans transcription activator AlcR, which binds DNA as a monomer but functions synergistically through multiple sites (Panozzo et al., 1997). Rgt1 is dissociated from HXT promoters after addition of glucose to glucose-depleted cultures. Rgt1 is phosphorylated at basal level in the absence of glucose and hyperphosphorylated by PKA in high levels of glucose. Hyperphosphorylated Rgt1 does not bind DNA, whereas dephosphorylation of Rgt1 in vitro restores its DNA-binding ability (Flick et al., 2003;

Kim et al., 2003; Mosley et al., 2003).

1.3.2. The glucose responsive transcription factors Mth1 and Std1

Mth1 and Std1 are paralogous proteins that play a key role in regulation of Rgt1 function (Hubbard et al., 1994; Schmidt et al., 1999). The main lines of supporting evidence are: 1) HXT gene expression is constitutive in the absence of Mth1 and Std1

(Hubbard et al., 1994; Lafuente et al., 2000; Lakshmanan et al., 2003; Schmidt et al.,

1999); 2) Rgt1 does not bind to HXT promoters in yeast cells lacking both MTH1 and

STD1 genes (Flick et al., 2003; Lakshmanan et al., 2003); 3) Mth1 and Std1 directly

interact with Rgt1 (Lakshmanan et al., 2003; Polish et al., 2005; Tomas-Cobos et al.,

2004). Since Rgt1 is hyperphosphorylated in the mth1std1 mutant, Mth1 and Std1 are thought to serve as Rgt1 regulators that modulate the phosphorylation state of Rgt1

(Mosley et al., 2003). Mth1 and Std1 are downregulated by glucose; they are ubiquitinated by the SCFGrr1 ubiquitin ligase complex and degraded via the 26S

proteasome in response to glucose (Flick et al., 2003; Moriya and Johnston, 2004; Pasula

et al., 2010; Pasula et al., 2007). There are also dominant mutations in the MTH1 gene

(HTR1-23, DGT1 or BCP1) (Gamo et al., 1994; Schulte et al., 2000) that render Mth1

resistant to glucose-induced degradation (Kim et al., 2006). Accumulating evidence

shows that Mth1 and Std1 may be not functionally redundant. Deletion of the STD1 gene

alone has little effect on the phosphorylation (Polish et al., 2005) and DNA-binding of the

Rgt1 repressor (Flick et al., 2003) and the expression of HXT genes (Kim et al., 2006).

Furthermore, transcriptome analysis shows that expression of major HXT genes is profoundly upregulated in an mth1 mutant but is not significantly changed in a std1 mutant (Sabina and Johnston, 2009). Therefore, Mth1 but not Std1 seems to be the major regulator of Rgt1.

1.3.3. The general corepressor complex Ssn6-Tup1

The Ssn6-Tup1 complex is a general transcriptional corepressor complex, composed of one molecule of Ssn6 and four molecules of Tup1 (Varanasi et al., 1996). The complex contains the tetratrico peptide repeat (TPR) and WD domains, respectively, which serve as protein-protein interaction motifs (Jabet et al., 2000; Schultz et al., 1990; Smith and

Johnson, 2000a; Smith et al., 1995; Sprague et al., 2000). Ssn6-Tup1 is recruited to its

7 target promoters by sequence-specific DNA-binding repressors (Malave and Dent, 2006;

Smith and Johnson, 2000b) and mediates transcriptional repression by recruiting global corepressors such as chromatin and nucleosome remodelers (Davie et al., 2003;

Edmondson et al., 1996) and/or by interacting with the RNA transcription machinery

(Malave and Dent, 2006; Smith and Johnson, 2000b). Ssn6-Tup1 also appears to be involved in the induction of gene expression (Mennella et al., 2003; Papamichos-

Chronakis et al., 2004; Proft and Struhl, 2002) and recruited to its target promoters in a manner independent of sequence-specific DNA-binding proteins (Buck and Lieb, 2006;

Desimone and Laney, 2010; Hanlon et al., 2011; Papamichos-Chronakis et al., 2004). In addition, Ssn6-Tup1 is shown to exert its function by masking the activation domain of a

DNA-binding repressor and thereby preventing recruitment of the coactivators necessary for transcriptional activation (Wong and Struhl, 2011).

1.4. Key components of the HXT gene induction pathway

Glucose induces expression of the HXT genes by inhibiting the function of Mth1 and

Rgt1. Two cell surface glucose receptors Rgt2 and Snf3 recognize the presence of extracellular glucose and generates an intracellular glucose signal that leads to disruption of HXT repressor complex and derepression of HXT gene expression (Figure 5). The properties of each of the key components of the HXT gene induction pathway are summarized below.

1.4.1. Cell surface glucose receptors Rgt2 and Snf3

The glucose signal that leads to inactivation of Rgt1 function appears to be generated

by Rgt2 and Snf3 (Ozcan and Johnston, 1996), that are similar to glucose transporters but

lost their ability to transport glucose (Ozcan et al., 1998). Although most of the Hxt

proteins are very similar to each other, sharing 50-100% amino acid sequence identity,

Rgt2 and Snf3 are most divergent members, being only ~25% similar to their relatives

(Bisson et al., 1993; Boles and Hollenberg, 1997; Kruckeberg, 1996). A distinguishing

characteristic of Rgt2 and Snf3 is their unusually long C-terminal tail (~200 amino acids) that is predicted to reside in the cytoplasm (Carlson, 1999) and transduce the glucose

Figure 3: Structures of glucose and related hexoses. The configurations with regard to C-3 and C-4 of the hexoses (grey circles) seem to be important for the interaction of the hexoses with the glucose sensors.

signal for expression of HXT genes (Ozcan et al., 1998). Rgt2 and Snf3 have separate but overlapping function in recognizing the presence of extracellular glucose (Ozcan et al.,

1998). Snf3 functions as a sensor of low levels of glucose as it is required for the expression of high affinity glucose transporter genes such as HXT2. Rgt2 recognizes the presence of high levels of glucose and is required for full induction of the low affinity glucose transporters such as HXT1. Hence, Rgt2 and Snf3 act as high and low affinity

9 glucose sensors, respectively. Hence, Rgt2 and Snf3 act as low and high affinity glucose sensors, respectively. Glucose and galactose are epimers; they only differ with respect to the C-4, yet glucose sensors generate signal in response to glucose, but not galactose.

These observations suggest that the glucose sensors and transporters have remarkable substrate specificity and the configuration of the hexoses with regard to C-3 and C-4 seem to be crucial factor in activating the sensors (Figure 3, Jouandot et al., 2011).

1.4.2. Yeast casein kinase I (Yck1 and Yck2) and SCFGrr1 ubiquitin ligase

A central player in the transduction of glucose signal generated by Rgt2 and Snf3 is the yeast casein kinase I (Yck1 and Yck2), tethered to the plasma membrane via palmitoylation of the C-terminal Cys-Cys sequence by the palmitoyl transferase Akr1

(Babu et al., 2004; Feng and Davis, 2000; Roth et al., 2002). Deletion of both the YCK1 and YCK2 genes results in non-viable phenotype (Robinson et al., 1992). Yeast casein kinase I is involved in various cellular functions such as morphogenesis (Robinson et al.,

1993), polarized growth and proper septin organization (Robinson et al., 1999). It is also required for turnover of many cell surface receptors and transporters such as uracil permease Fur4 (Marchal et al., 2000), alpha-factor receptor Ste2 (Hicke et al., 1998) and maltose permease Mal61 (Gadura and Michels, 2006; Gadura et al., 2006). Yck1/2 plays a major role in Rgt2/Snf3 mediated glucose induction of HXT genes. A current view is that glucose binding to Rgt2 and Snf3 induces a change in their conformation that activates Yck1/Yck2 (Moriya and Johnston, 2004), which in turn catalyzes phosphorylation of Mth1 and Std1 (Moriya and Johnston, 2004; Pasula et al., 2010).

Phosphorylated Mth1 and Std1 are ubiquitinated by SCFGrr1 ubiquitin protein ligase and

subsequently degraded by 26S proteasome (Flick et al., 2003; Kim et al., 2006; Pasula et

al., 2007) leading to PKA-dependent phosphorylation of Rgt1 (Kim and Johnston, 2006)

and derepression of HXT gene expression. A recent study showed that Cu/Zn superoxide

dismutase Sod1 is required for stabilization of Yck1/2 and glucose repression of

respiration (Reddi and Culotta, 2013) which further favors aerobic glycolysis in yeast.

1.4.3. The yeast glucose transporters (Hxt)

S. cerevisiae possesses at least six members of glucose transporter family (Hxt1, 2, 3,

4, 6 and 7), each with different affinities for glucose in order to cope with environmental

changes in glucose availability (Boles and Hollenberg, 1997; Reifenberger et al., 1997;

Reifenberger et al., 1995). The Hxt proteins belong to the major facilitator superfamily

(MFS) with predicted twelve transmembrane domains and cytoplasmic amino and caoxy-

terminal domains (Lewis and Bisson, 1991). Hxt1 is a low affinity glucose transporter

with Km value around 100 mM for glucose and expressed when glucose levels are high (>

~1%); the HXT2 and HXT4 genes encode glucose transporters with moderate affinity for

glucose (Km values around 10 mM) and their expression is induced in the presence of low

amount of glucose (~ 0.2%) (Maier et al., 2002; Reifenberger et al., 1997). Hxt3 has low

affinity for glucose and induced in both low and high amounts of glucose (Km value

around 30-60 mM) (Boles et al., 1997; Ozcan and Johnston, 1999; Reifenberger et al.,

1997). The HXT6 and HXT7 genes encode high affinity glucose transporters with Km

value around 1 mM for glucose and their expression is induced by low concentration of

glucose as well as in the presence of non-fermentable carbon sources such as ethanol, but

is repressed at high glucose concentrations (Diderich et al., 1999; Liang and Gaber, 1996;

Ye et al., 1999).

1.5. Crosstalk between glucose repression and induction pathways to ensure

stringent regulation of glucose consumption

1.5.1. Integration of glucose induction and glucose repression pathways in a

regulatory network

A gene expression profiling study showed that the Rgt2/Snf3 glucose induction and

the Snf1/Mig1 glucose repression pathways are intertwined in a regulatory network

(Kaniak et al., 2004). Glucose stimulates the proteasome-mediated degradation of Mth1 by the Rgt2/Snf3 pathway and also reduces MTH1 expression by the Snf1/Mig1 pathway

(Flick et al., 2003; Kim et al., 2006; Moriya and Johnston, 2004), which pose the inhibitory effect of glucose on Mth1 function and ensure maximal glucose induction of

Rgt1-repressed genes. Glucose inhibits Std1 function by stimulating its degradation and also induces STD1 gene expression by Rgt2/Snf3 pathway (Kaniak et al., 2004).

Therefore, the feedback control of STD1 gene expression counteracts the glucose-induced

Std1 degradation and provides for a rapid reestablishment of repression when glucose is exhausted. This may contribute to the regulation of glucose repression, because Snf1 activity is enhanced by Std1 (Kuchin et al., 2003). Mig2 is a glucose repressor and collaborates with Mig1 in repression of most glucose-repressed genes. Mig1 and Mig2 bind to the identical DNA sequences but are differently regulated (Lutfiyya et al., 1998;

Lutfiyya and Johnston, 1996). Mig2, unlike Mig1, is not regulated by Snf1, but its expression is induced by glucose via the Rgt2/Snf3 pathway (Kaniak et al., 2004).

Therefore, glucose repression is a result of outputs from two glucose signal transduction

pathways: the Mig1 component regulated by the Snf1 kinase and the Mig2 component

regulated at the level of their transcription by the Snf3/Rgt2 signaling pathway.

Expression of the SNF3 gene is repressed by the Snf1-Mig1 pathway in high levels of glucose, enabling Snf3 to function only in low glucose conditions (Kaniak et al., 2004).

Hxk2 is the most active hexokinase isoenzyme during growth on glucose in the cytoplasm, while it interacts with components repressing expression of several glucose- repressed genes in the nucleus (Busti et al., 2010). Glucose induces expression of the

HXK2 gene via the Rgt2/Snf3 pathway, providing another functional link between glucose induction and repression pathways (Palomino et al., 2006).

Figure 4. Rgt2/Snf3 mediated glucose induction pathway of HXT gene expression. In the absence of glucose, Rgt1 forms a repression complex on the HXT promoters along with Mth1/Std1 and Ssn6-Tup1 and represses HXT gene transcription. Presence of extracellular glucose is sensed by two cell surface glucose receptors Rgt2 and Snf3 and they generate an intracellular signal which activates casein kinase I (Yck1/2). Activated yck1/2 phosphorylates Mth1/Std1 which triggers its degradation via SCFGrr1 ubiquitination and 26S protasomal pathway. Rgt1 is phosphorylated by cAMP-activated protein kinase A (PKA) and released from the DNA resulting in disruption of the

repression complex and derepression of HXT genes. The glucose transporters (Hxt) now migrate to the cell surface and transport glucose inside the cell to carry out glycolysis.

1.5.2. The glucose repression signal is generated through glucose metabolism

The AMP-activated protein kinase (AMPK) is known to be activated by an elevated

AMP:ATP ratio upon glucose depletion and the AMP:ATP ratio increases more than

200-fold upon glucose removal and decreases when glucose is added. However, Snf1

activity is not directly regulated by the AMP:ATP ratio (Hardie et al., 1998; Wilson et al.,

1996). Instead, it has been speculated that glucose-6-phosphate (G-6-P) might serve as a

glucose repression signal based on the observations that: (1) Snf1 is constitutively active

in a hxk1hxk2glk1 deletion mutant; (2) addition of the glucose analog 2-deoxyglucose, which can be phosphorylated but not further metabolized, to glycerol-grown cells results in the decreased activity of Snf1; (Ma et al.) Snf1 activity is not affected by 6- deoxyglucose, which cannot be phosphorylated (Hedbacker and Carlson, 2006; Meijer et al., 1998). However, the observations that glucose repression may be related to the glucose concentration rather than glucose flux suggest that the glucose repression signal appears to be upstream of G-6-P (Meijer et al., 1998; Ye et al., 1999). In this regard, it was demonstrated that glucose repression of the Mig1 target genes is abolished in a mutant lacking all 17 hexose transporters or the Rgt2 and Snf3 glucose sensors (Ozcan,

2002). Hence, regulation of glucose-induction of HXT gene expression plays an important role in the generation of the glucose repression signal, highlighting how glucose induction and glucose repression of gene expression is coordinated to ensure tight control of glucose uptake and metabolism (Figure 5).

Figure 5. Schematic diagram of the crosstalk between glucose signaling pathways in yeast. Yck I (Yck1 and Yck2) phosphorylates Mth1 and Std1 upon activation by glucose-bound Rgt2 and Snf3 glucose sensors. Phosphorylated Mth1 and Std1 are ubiquitinated by the SCFGrr1 complex and degraded by the proteasome. The PKA phosphorylation sites in the amino terminal region of Rgt1 are exposed and available for phosphorylation when Mth1 is degraded. Phosphorylated Rgt1 is dissociated form Ssn6- Tup1 and subsequently from DNA, leading to derepression of Rgt1 target genes, such as the HXT and HXK2 genes. The Rgt2/Snf3 pathway regulates itself through glucose- induction of STD1 gene expression. Consequently, the STD1 gene is expressed at the same time that the Std1 protein is degraded in response to glucose (Kaniak et al., 2004). By contrast, glucose stimulates Mth1 degradation but also represses Mth1 expression via Mig1 and Mig2. Glucose uptake is required for the generation of the glucose repression signal that leads to inactivation of the Snf1 kinase (Ozcan, 2002). Expression of the MIG2 gene is induced by glucose via the Rgt2/Snf3 pathway. Glucose-repression of SNF3 gene expression by Mig1 reflects the probable function of Snf3 as a high affinity glucose sensor, representing another important feature of the interaction between the glucose induction and repression pathways.

1.6. Research Significance

Upon addition of glucose to the glucose-depleted medium, Rgt1 was shown to be phosphorylated by PKA and dissociated from DNA (Kim and Johnston, 2006; Kim et al.,

2003). In my preliminary result, I found that two chimeric Rgt1 proteins (LexA-Rgt1 and

GFP-Rgt1) bind DNA constitutively but cannot repress HXT gene expression in the presence of glucose. Therefore, in the chapter entitled “Understanding the mechanism of glucose-induced relief of Rgt1-mediated repression in yeast”, I investigated the mechanism of glucose regulation of Rgt1 function. I showed that Mth1 inhibits glucose induced phosphorylation of Rgt1 that leads to dissociation of Ssn6-Tup1 from the DNA- bound Rgt1 constructs, resulting in derepression of HXT gene expression. Thus, I have presented direct evidence that removal of Rgt1 from the DNA upon glucose induction is not necessary for glucose induction of HXT genes expression, suggesting that glucose regulates Rgt1 function by primarily modulating the Rgt1 interaction with Ssn6-Tup1.

Mig1 is a transcription factor that represses the expression of genes required for the use of alternative carbon sources such as SUC2 and GAL1 in the presence of glucose and the regulation of Rgt1 function is mechanistically similar to Mig1, which is phosphorylated and negatively regulated by Snf1 kinase (Hedbacker and Carlson, 2008;

Treitel et al., 1998). Unphosphorylated Mig1 interacts with Ssn6-Tup1 in the presence of high glucose and mediates repression of Mig1-target genes. Mth1 is a glucose responsive transcription factor that is required by Rgt1, not Mig1. Hence, in my next chapter entitled

“Mth1 regulates the interaction between the Rgt1 repressor and the Ssn6/Tup1 corepressor complex by modulating PKA-dependent phosphorylation of Rgt1”, I investigated the role of Mth1 in repression of HXT gene expression. I found that

Ssn6/Tup1 interferes with the DNA-binding activity of Rgt1 in the absence of Mth1, and that Rgt1 function abrogated by Ssn6 overexpression is restored by cooverexpression of

Mth1. Thus, Mth1 acts as a scaffold-like protein and mediates interaction between Rgt1 and Ssn6-Tup1 by preventing PKA-dependent phosphorylation of Rgt1. This study

16 provided a new insight into eukaryotic transcription regulation mechanism where a transcription repressor complex (Ssn6-Tup1) inhibits the DNA-binding activity of the master regulator (Rgt1) and this inhibitory effect is stabilized by another transcription factor (Mth1) for fine-tuning of transcription regulation of target genes (HXTs).

Many small molecule transporters function as sensors. Bacteria sense certain sugars by sugar transporters (Postma et al., 1993). Because signal generation is coupled to transport and metabolism, this phenomenon cannot be viewed as a receptor-mediated event. Neurospora crassa glucose transporter Rco3 is required for expression of glucose transporters and it may function as Saccharomyces cerevisiae glucose sensors Rgt2 and

Snf3 (Madi et al., 1997). Although mammals lack well-defined glucose sensor systems like Rgt2 and Snf3, it is possible that similar glucose receptors may be found in β-cells in pancreas which must sense the availability of glucose. Therefore, to gain more insight into the mechanism of glucose regulation of the glucose sensors Rgt2 and Snf3, I investigated how Rgt2 and Snf3 generate glucose signal for the expression of HXT genes and how the signal is desensitized in the absence of glucose in the chapter entitled

“Glucose regulation of the yeast cell surface glucose sensors Rgt2 and Snf3”. I found that Rgt2 and Snf3 are functional over a broad range of glucose concentrations (2-0.05%) and are ubiquitinated and degraded in vacuole via endocytosis in the absence of glucose, whereas, constitutively active sensors (Rgt2-1, Snf3-1), though to be in glucose-bound conformation, are resistant to degradation. These findings reinforce the fact that yeast cells maintain their unique fermentative lifestyle by stabilizing the glucose sensors in cell surface and sensing the presence of fluctuating levels of glucose in changing environment.

Glucose sensor-mediated intracellular signal targets inactivation of Rgt1 and

derepression of HXT genes for increased uptake of extracellular glucose. In the following

chapter entitled “Glucose starvation-induced turnover of the yeast glucose transporter

Hxt1”, I analyzed the role of glucose in maintaining the structural and functional

properties of the yeast glucose transporters using low affinity glucose transporter Hxt1 as

a model. I showed that the amino-terminal cytoplasmic domain of Hxt1 is required for

glucose starvation-induced endocytosis and vacuolar degradation of Hxt1. I showed that

the stability and cell surface level of Hxt1 may be associated with its kinetic properties

and structure and provided a mechanistic insight into how glucose regulates the stability

of glucose transporters to carry out glycolysis. I have also identified amino acid residues

that are, in part, required for glucose transport activity of Hxt1.

The studies in this dissertation identified the precise role of the transcription factor Mth1

in glucose regulation of HXT gene expression and provided a novel mechanism of

eukaryotic gene regulation. Many yeast nutrient and metal transporters are produced in

the absence of their substrates, but undergo endocytosis and subsequent vacuolar

degradation when their substrates become available in excess. In my study, I introduced

novel concept of cell surface receptor biology. Ligand-induced internalization and lysosomal degradation constitutes a primary mechanism responsible for receptor downregulation. The endocytosis of many signaling receptors is stimulated by ligand-induced activation (Doherty and McMahon, 2009; Polo and Di Fiore, 2006). My findings show that the glucose sensors are subjected to endocytosis and subsequent vacuolar degradation in the absence of their ligand, glucose, but are stable in the presence of glucose. This enables yeast cells to consume glucose vigorously. This discovery sheds the

18 light on robust glucose consumption associated with aerobic glycolysis, the prime characteristic of the Warburg/Crabtree effect. Further characterization of glucose-binding site of glucose transporters in pathogenic yeasts such as Candida albicans could provide therapeutic interventions against pathological conditions including Candidiasis.

CHAPTER II

Understanding the mechanism of glucose-induced relief of Rgt1- mediated repression of HXT gene expression in yeast

Abstract

The yeast Rgt1 repressor inhibits transcription of the glucose transporter (HXT) genes in the absence of glucose; it does so by recruiting the general corepressor complex Ssn6-

Tup1 and the HXT corepressor Mth1. In response to glucose, Rgt1 is phosphorylated by the cAMP-activated protein kinase A (PKA) and dissociates from the HXT promoters, resulting in expression of HXT genes. In this study, using Rgt1 chimeras that bind DNA constitutively, I investigate how glucose regulates Rgt1 function. My results show that the DNA-bound Rgt1 constructs repress expression of the HXT1 gene in conjunction with

Ssn6-Tup1 and Mth1, and this repression is lifted when they dissociate from Ssn6-Tup1 in response to glucose. Mth1 prevents PKA phosphorylation of the Rgt1 constructs, and glucose downregulation of Mth1 enables PKA to phosphorylate the Rgt1 constructs. This phosphorylation induces dissociation of Ssn6-Tup1 from the DNA-bound Rgt1 constructs, resulting in derepression of HXT gene expression. Therefore, Rgt1 removal from DNA occurs in response to glucose but is not necessary for glucose induction of

HXT gene expression, suggesting that glucose regulates Rgt1 function by primarily modulating the Rgt1 interaction with Ssn6-Tup1.

2.1. Introduction

The yeast Rgt1 repressor is a DNA-binding transcription factor that regulates

expression of glucose responsive genes, including genes encoding a family of glucose

transporters (Hxts) (Ozcan and Johnston, 1996, 1999). Rgt1 brings about repression of

HXT gene expression by recruiting the general corepressor Ssn6-Tup1 to the HXT

promoters in the absence of glucose (Kim et al., 2003; Ozcan et al., 1996b). Ssn6-Tup1

mediates transcriptional repression by recruiting global corepressors, such as chromatin

and nucleosome remodelers, or by directly interacting with the RNA transcription

machinery (Malave and Dent, 2006; Smith and Johnson, 2000b). Ssn6-Tup1 also

functions by masking the activation domain of a DNA-binding repressor and thereby preventing recruitment of the coactivators necessary for transcriptional activation in the absence of glucose(Wong and Struhl, 2011). Thus, Ssn6-Tup1 may act differently on

different repressors, but an efficient recruitment of Ssn6-Tup1 by gene specific repressors

may be critical for establishing repression.

Rgt1-dependent, Ssn6-Tup1-mediated repression occurs in conjunction with the

paralogous proteins Mth1 and Std1. Early observations showed that HXT genes are

constitutively expressed in cells lacking Mth1 and Std1 (Schmidt et al., 1999), and that

Rgt1 directly interact with Mth1 and Std1 and binds to the HXT promoters in an Mth1

and Std1-depedent manner (Flick et al., 2003; Kim et al., 2006; Lafuente et al., 2000;

Lakshmanan et al., 2003; Polish et al., 2005; Tomas-Cobos and Sanz, 2002). Mth1 and

Std1 were also shown to be degraded in the 26S proteasome in cells grown in high glucose medium, implicating Mth1 and Std1 as Rgt1 regulators (Kaniak et al., 2004).

However, recent studies have shown that deletion of the STD1 gene alone has little effect

21 on the regulation of HXT gene expression (Flick et al., 2003; Moriya and Johnston,

2004). Furthermore, glucose appears to inactivate Rgt1 function by primarily down- regulating Mth1 levels: it not only induces proteasome degradation of Mth1 (Kaniak et al., 2004; Lakshmanan et al., 2003; Moriya and Johnston, 2004; Pasula et al., 2007;

Spielewoy et al., 2004) but also represses expression of the MTH1 gene (Flick et al.,

2003; Moriya and Johnston, 2004). Therefore, it is likely that Mth1 acts as a central player in establishing Rgt1-mediated repression (Kim et al., 2013; Spielewoy et al.,

2004).

Rgt1 is phosphorylated and dissociated from the HXT promoters in response to glucose (Kim et al., 2003; Mosley et al., 2003). Rgt1 is a phosphoprotein; it is phosphorylated at a basal level in the absence of glucose, but is hyperphosphorylated by

PKA in high levels of glucose (Kim and Johnston, 2006; Palomino et al., 2006; Toda et al., 1987a; Toda et al., 1987b). Rgt1 is phosphorylated at four serine residues within its amino-terminal region, but this does not occur until Mth1 is degraded (Jouandot et al.,

2011). More importantly, phosphorylation of Rgt1 inhibits its interaction with Ssn6-Tup1

(Roy et al., 2013a). Thus, Mth1 plays a key role in mediating the interaction between

Rgt1 and Ssn6-Tup1 by inhibiting PKA phosphorylation of Rgt1. Moreover, the finding that Rgt1 bound to the HXT1 promoter does not inhibit glucose induction of HXT1 gene expression in cells lacking Ssn6 or Tup1 raises a possibility that and glucose-induced

Rgt1 removal from DNA is not the primary cause of glucose induction of HXT gene expression (Roy et al., 2013a). The relief of Ssn6-Tup1-mediated repression comes about through the destruction or inactivation of the individual repressors, leading to dissociation of the repressors from Ssn6-Tup1 and/or DNA (Smith and Johnson, 2000b). Based on

22 previous observations, I have suspected that dissociation of Rgt1 from DNA occurs in response to glucose, but is not required for glucose induction of HXT gene expression, and that rather dissociation of Ssn6-Tup1 from Rgt1 is sufficient to lift Rgt1-mediated repression.

Here, I investigate how glucose regulates Rgt1 function using the LexA-Rgt1 and

GFP-Rgt1 fusions that bind DNA constitutively. I found that the Rgt1 constructs repress

HXT1 gene expression in conjunction with Mth1 and Ssn6-Tup1 in the absence of glucose, and that this repression is lifted when the Rgt1 constructs are phosphorylated and dissociated from Ssn6-Tup1 in response to glucose. In contrast, the Rgt1 constructs lacking PKA phosphorylation sites did not dissociate from Ssn6-Tup1, resulting in constitutive repression of the HXT1 gene. My results provide direct evidence that glucose induction of HXT gene expression results primarily from the disruption of the

Rgt1-Ssn6-Tup1 interaction, rather than from Rgt1 removal from DNA.

2.2. Materials and Methods

2.2.1. Yeast strains and Plasmids

Yeast strains used in this study are listed in Table 1.1. Except where indicated, yeast strains were grown in YP (2% bacto-peptone, 1% yeast extract) and SC (synthetic yeast nitrogen base media containing 0.17% yeast nitrogen base and 0.5% ammonium sulfate) supplemented with the appropriate amino acids and carbon sources. Plasmids used in this study are listed in Table 1.2.

2.2.2. Chromatin Immunoprecipitation (ChIP)

ChIP was performed as described previously (Kim et al., 2003). Briefly, Yeast strains

were grown till mid-log phase (O.D600nm = 1.2-1.5) and incubated with formaldehyde

(1% final concentration) at room temperature for 15 to 20 min. The cross-linking reaction

was quenched by adding glycine to a final concentration of 125 mM for 5 min. The cells

were disrupted by vortexing with acid-washed glass beads in ice cold ChIP lysis buffer

(50 mM HEPES-KOH, pH 7.5, 150 mM NaCl, 1% Triton X-100, 0.1% Na-deoxycholate)

containing protease and phosphatase inhibitors. The lysate was sonicated (ultrasonic cell

disruptor with a microtip) five times with 10 sec pulse. The genomic DNA fragments

were immunoprecipitated with anti-HA, LexA, GFP or Ssn6 antibody (Santa Cruz)

conjugated with agarose beads. After washing the immunoprecipitated beads, DNA was

eluted from both immunoprecipitated and 1/100 input samples. The immunoprecipitated

DNA was PCR-amplified using primer pairs directed against the HXT1 promoter. As a

negative control, primer sets were designed to amplify the actin gene promoter region.

DNA-binding of Rgt1 was determined by running the PCR products of linear range in

1.5% agarose gel and visualizing by ethidium bromide staining.

2.2.3. Western blot and Immunoprecipitation (IP) analysis

For Western blot analysis, yeast cells (O.D600 = 1.5) were collected by centrifugation

at 3,000 rpm in a table-top centrifuge for 5 min. The cell pellets were resuspended in 100

µl of SDS-buffer (50 mM Tris-HCl, pH 6.8, 10% glycerol, 2% SDS, 5% β-

mercaptoethanol) and boiled for 5 min. After the lysates were cleared by centrifugation at 12,000 rpm for 10 min., soluble proteins were resolved by SDS-PAGE and transferred

to PVDF membrane (Millipore). The membranes were incubated with appropriate

antibodies (anti-HA, anti-LexA, anti -GFP and anti-TAP antibodies, Santa Cruz) in TBST buffer (10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.1% Tween-20) and proteins were detected by the enhanced chemiluminescence (ECL) system. For IP, yeast cells were disrupted by vortexing with acid-washed glass beads in ice cold NP40 buffer (1% NP40,

150 mM NaCl, 50 mM Tris-HCl, pH 8.0) containing protease and phosphatase inhibitors.

The cell lysates were incubated with appropriate antibodies at 4 oC for 3 hr and further

incubated with protein A/G-conjugated agarose beads at 4 oC for 1 hr. The precipitated

agarose beads were washed three times with ice cold NP40 buffer containing protease

and phosphatase inhibitor cocktails (Sigma P8215 and Sigma P0044, respectively) and

boiled in 50 µl of SDS-PAGE buffer. The resulting proteins were analyzed by Western

blot using appropriate antibodies.

2.2.4. β-galactosidase assay

To assay β-galactosidase activity with yeast cells expressing the HXT1-LacZ reporter,

the yeast cells were grown to mid-log phase and the assay was performed as described

previously (Kaniak et al., 2004). Results were given in Miller Units [(1,000 x

O.D420nm)/(T x V x O.D600nm), where T was the incubation time in minutes, and V is the volume of cells in milliliters]. The reported enzyme activities were averages of results from triplicates of three different Transformants with error bars representing standard deviation (S.D).

2.2.5. Quantitative Real Time PCR (qRT-PCR)

Total RNA was extracted by RNeasy mini kit (Qiagen) following manufacturer’s

protocol and 2µg of total RNA was converted to cDNA by qScript cDNA supermix

(Quanta Biosciences). cDNA was analyzed by qRT-PCR using SsoFast Evagreen reagent

(Bio-Rad) in CFX96 Real-time thermal cycler (Bio-Rad). ACT1 was used as an internal control to normalize expression of HXT1 gene. All of the shown quantification data were

the averages of three independent experiments with error bars representing standard

deviations (S.D).

2.3. Results

2.3.1. Glucose induction of HXT gene expression does not require the

dissociation of Rgt1 from the HXT promoters

The prevailing view is that expression of HXT genes is induced when Rgt1 is

dissociated from DNA in response to glucose (Kim et al., 2003). However, my recent

study shows that Rgt1 can bind DNA constitutively without inhibiting glucose induction

of HXT gene expression in cells lacking Ssn6 or Tup1 (Roy et al., 2013a). This finding

has raised the question of whether derepression of HXT genes does not require Rgt1 removal from the HXT promoters. To answer this question, I tested three Rgt1 fusion proteins—Rgt1-HA (3x HA at its C-terminus, 27aa) (Mosley et al., 2003), LexA-Rgt1

(LexA at its N-terminus, 87 aa) (Ozcan et al., 1996b) and GFP-Rgt1 (GFP at its N- terminus, 239 aa) (Kim et al., 2003) —for their ability to bind to the HXT1 promoter

(Figures 6A and 6B). My ChIP assays showed that Rgt1-HA binds to the HXT1 promoter

26 in the absence of glucose, but is dissociated from DNA in response to glucose (Figures

6C and 6D). However, LexA-Rgt1 and GFP-Rgt1 were shown to bind DNA constitutively. Thus, Rgt1-HA, like the native, untagged Rgt1, binds DNA in a glucose- regulated manner as reported previously (Mosley et al., 2003; Roy et al., 2013a).

Whereas DNA binding by LexA-Rgt1 and GFP-Rgt1 is constitutive. These results suggest that the addition of the LexA or GFP epitope to the N-terminus of Rgt1 affect its

DNA-binding property.

I next assessed the ability of the Rgt1 fusions to repress HXT1 gene expression. The

HXT1-NAT reporter strain expresses the NAT resistance gene under the control of the

HXT1 promoter. Therefore, the strain is susceptible to nourseothricin in the absence of glucose (2% Gal+ NAT), but exhibits resistance to the antibiotic when glucose is present

(Roy et al., 2013b). The reporter strains expressing the Rgt1 fusions were shown to grow only on glucose-containing medium (Figure 6E), suggesting that the Rgt1 constructs are fully functional in the absence of glucose and negatively regulated by glucose. I also found that expression of the HXT1-lacZ reporter is repressed by all the Rgt1 fusions in the absence of glucose (vector vs Rgt1 fusions) and that glucose induces expression of the reporter by ~52-fold (Gal vs Glu) in cells expressing Rgt1-HA and by ~11- and ~15- fold (Gal vs. Glu) in cells expressing LexA-Rgt1 and GFP-Rgt1, respectively (Figures 6F and 6A). This indicates that expression of the HXT1-lacZ reporter is largely induced by glucose in cells expressing the Rgt1 fusions, but is still ~4-5-fold repressed by LexA-

Rgt1 and GFP-Rgt1 (vector vs LexA- or GFP-Rgt1 fusions) in the presence of glucose.

Figure 6. Dissociation of the Rgt1 repressor from HXT promoters is not required for glucose-induction of HXT1 gene expression. (A) Schematic diagram of the structures of Rgt1 and its constructs. The map shows the DNA-binding domain, the Ssn6- Tup1 interaction site and the PKA phosphorylation sites. (B) Yeast cells (rgt1∆) expressing Rgt1-HA (KFP60), LexA-Rgt1 (pBM3580) or GFP-Rgt1 (pBM3911) were grown in SC-2% galactose medium (-) till mid-log phase and shifted to SC-4% glucose medium (+) for 1 hr. Rgt1 was subjected to Western blot analysis using anti-HA, anti- LexA or anti-GFP antibody. (C) ChIP analysis of Rgt1 binding to the HXT1 promoter. Yeast cells (rgt1∆) expressing Rgt1-HA, LexA-Rgt1 or GFP-Rgt1 were grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr, and cross-linked chromatin was precipitated using anti-HA, anti-LexA or anti-GFP antibody. Representative PCRs are shown for amplification of HXT1 promoter. As a negative control of Rgt1 DNA binding, primer sets were designed to amplify the actin gene promoter region (pACT1), which does not contain the Rgt1-binding sequence (5’- CGGANNA-3’). (D) qPCR analysis of Rgt1-binding to the HXT1 promoter. The amount of immunoprecipitated (IP) DNA was quantified by qPCR with primer pairs directed against the HXT1 promoter (pHXT1). IP/Input ratio was determined by the ratio of IP/pHXT1 relative to the IP/pACT1 divided by the ratio of input/pHXT1 relative to the INPUT/pACT1. The data shown are averages of three independent experiments with error bars showing mean ± S.D. (E) The HXT1 ORF was replaced by the NAT ORF by homologous recombination. Empty vector, Rgt1-HA, LexA-Rgt1 and GFP-Rgt1 were expressed in the reporter strain. The reporter cells were spotted on SC-Leu or SC-Ura plate containing either 2% galactose or 4% glucose supplemented with 100µg/ml NAT sulfate. The first spot of each row represented a count of 5×107 cells/ml, which is diluted

1:10 for each spot thereafter. The plates were incubated for 3 days and photographed. (F) Yeast cells (rgt1∆) coexpressing pHXT1-LacZ reporter plasmid and Rgt1-HA, LexA- Rgt1 or GFP-Rgt1 were grown as described in (B) and assayed for β-galactosidase activity. An empty vector served as a control.

2.3.2. Mth1 does not directly regulate the DNA-binding ability of LexA-Rgt1

The DNA-binding activity of Rgt1 is regulated by Mth1 (Flick et al., 2003; Moriya

and Johnston, 2004). Given that LexA-Rgt1 and GFP-Rgt1 binds DNA constitutively

(Figure 6), I investigated whether the DNA-binding activity of these Rgt1 constructs is regulated by Mth1. The glucose signal that leads to HXT gene expression is generated by the Rgt2 and Snf3 glucose sensors at the plasma membrane (Ozcan et al., 1996a) (Figure

7A). Dominant mutations in the glucose sensor genes (SNF3-1 and RGT2-1) have been

identified that cause Mth1 degradation and thereby HXT gene expression in a glucose-

independent manner (Ozcan et al., 1998; Pasula et al., 2010; Pasula et al., 2007).

Consistent with these observations, Rgt1 does not bind DNA regardless of the presence of glucose in mth1Δ or RGT2-1 strain (Flick et al., 2003). My ChIP analysis showed that the Rgt1-HA fusion does not bind DNA and that, by contrast, the DNA-binding of the

LexA-Rgt1 fusion is constitutive, in mth1Δ or RGT2-1 strain (Figure 7B). Thus, Mth1 is not required for the DNA-binding of LexA-Rgt1. Despite of this discrepancy, neither

Rgt1-HA nor LexA-Rgt1 was able to repress expression of HXT1-lacZ reporter in the strain (mth1Δ or RGT2-1) (Figures 7C and 11B). These results suggest that Mth1 may regulate the function of LexA-Rgt1 without directly affecting its DNA-binding ability.

These observations are in line with the previous observations that the role of Mth1 in

Rgt1-mediated repression is to mediate the interaction between Rgt1 and Ssn6-Tup1

(Roy et al., 2013a).

Figure 7. Mth1 does not directly regulate the DNA-binding ability of Rgt1. (A) The current model of glucose-induction of HXT gene expression. The Rgt2 and Snf3 glucose sensors undergo a conformational change upon glucose binding and generate a signal that leads to proteasomal degradation of Mth1 and Std1. PKA phosphorylation of Rgt1, which occurs when Mth1 and Std1 are degraded, induces its dissociation from both Ssn6-Tup1 and its target promoters, leading to the expression of the HXT genes. RGT2-1 and SNF3- 1 are dominant mutations that are thought to convert the proteins into the glucose-bound forms and cause glucose-independent expression of the HXT genes. (B) ChIP analysis of Rgt1-binding to the HXT1 promoter. Top: Yeast cells of indicated genotypes expressing Rgt1-HA or LexA-Rgt1 were grown in SC-2% galactose medium (-) and shifted to SC- 4% glucose medium (+) for 1 hr and cross-linked chromatin was precipitated using anti- HA or anti-LexA antibody, and representative PCRs were shown for amplification of HXT1 promoter. As a negative control of Rgt1 DNA-binding, primer sets were designed to amplify the actin gene promoter region (pACT1). Bottom: qPCR analysis of Rgt1- binding to the HXT1 promoter, as described in Figure 6D. (C) Yeast cells (mth1∆ and RGT2-1) coexpressing pHXT1-LacZ reporter plasmid and Rgt1-HA or LexA-Rgt1 were grown as described in (B) and assayed for β-galactosidase activity. Empty vector served as a control.

2.3.3. LexA-Rgt1 function is regulated by its phosphorylation state

My findings that LexA-Rgt1 binds DNA constitutively without significant inhibition of glucose-induction of HXT gene expression support the view that Rgt1 dissociation

from DNA may not be required for glucose-induction of HXT gene expression (Figure 6).

Rgt1 function is critically regulated by its phosphorylation by PKA (Jouandot et al.,

2011; Kim and Johnston, 2006; Palomino et al., 2006). Thus, I examined whether the

DNA-bound LexA-Rgt1 is regulated by its phosphorylation state by exploring the ability

of the wild type LexA-Rgt1 and the mutant LexA-Rgt1 (5SA) lacking the PKA phosphorylation sites (S96, S146, S202, S283 and S284) (Kim and Johnston, 2006) to

regulate the HXT1 promoter. Both LexA-Rgt1 and LexA-Rgt1 (5SA) were shown to bind to the HXT1 promoter constitutively, suggesting that the phosphorylation state of

LexA-Rgt1 does not regulate its DNA-binding ability (Figure 8A, top). However, the colony assay, performed as described above (Figure 6E), demonstrated that LexA-Rgt1

(5SA), but not by LexA-Rgt1, constitutively inhibits the expression of the NAT resistant gene and thus cell growth in the glucose medium (Figure 8B). Consistently, the HXT1- lacZ reporter assay showed that LexA-Rgt1 (5SA) inhibits glucose-induced expression of the reporter gene (Figures 8C and 11C). These results suggest that LexA-Rgt1, bound to the HXT promoters constitutively, is critically regulated by its phosphorylation state.

2.3.4. The phosphorylation state of LexA-Rgt1 regulates its affinity for Ssn6-

Tup1

Glucose-induced expression of HXT genes requires the dissociation of Rgt1 from

Ssn6-Tup1 (Roy et al., 2013a). I examined the ability of LexA-Rgt1 and LexA-Rgt1

(5SA) to recruit Ssn6-Tup1 to the HXT1 promoter by ChIP analysis using the anti-Ssn6 antibody. LexA-Rgt1 appeared to recruit Ssn6-Tup1 to the HXT1 promoter in a glucose- dependent manner; Ssn6-Tup1 is associated with the HXT1 promoter in the absence of

glucose but is largely dissociated from the promoter when glucose is present (Figure 9A).

However, LexA-Rgt1 (5SA) recruits Ssn6-Tup1 constitutively to the HXT1 promoter,

suggesting that blocking glucose-induced PKA phosphorylation of Rgt1 enables it to

recruit Ssn6-Tup1.

Finally, I explored the effect of the phosphorylation defective mutation of Rgt1 (5SA)

on the interaction between LexA-Rgt1 and Ssn6-Tup1. To do so, I coexpressed LexA-

Rgt1 or LexA-Rgt1 (5SA) and Ssn6-TAP, and performed co-immunoprecipitation

experiments with the anti-LexA antibody. The interaction of LexA-Rgt1 with Ssn6-TAP was strongly detected in galactose-grown cells, but significantly reduced in glucose-

grown cells. Notably, the interaction between LexA-Rgt1 (5SA) and Ssn6-TAP occurs

constitutively, reinforcing the view that the ability of LexA-Rgt1 to recruit Ssn6-Tup1 is

regulated by its phosphorylation state (Figure 9B).

2.4. Discussion

In this study, I provide evidence that glucose induction of HXT gene expression

results primarily from the disruption of the Rgt1-Ssn6-Tup1 interaction, rather than from

Rgt1 removal from the HXT promoters. It has been well established that Rgt1 binds DNA

in a glucose-dependent manner. ChIP analyses demonstrate that Rgt1 binds to its target

promoters in the absence of glucose and dissociates from DNA in response to glucose

(Flick et al., 2003; Kim, 2004, 2009; Kim et al., 2003; Lakshmanan et al., 2003; Mosley

et al., 2003). An in vitro experiment showed that nuclear extracts from cells grown in

glucose-depleted medium, but not in glucose-containing medium, can make a DNA-

protein complex with a synthetic DNA sequence containing an Rgt1 recognition site

(Rgt1HXK2 probe) (Palomino et al., 2006). Here, I used three Rgt1 constructs—Rgt1-HA,

LexA-Rgt1 and GFP-Rgt1 fusions—to study the glucose regulation of Rgt1 function.

Rgt1-HA behaves like the native, untagged Rgt1, as reported previously (Lakshmanan et

al., 2003; Mosley et al., 2003; Roy et al., 2013a). However, LexA-Rgt1 and GFP-Rgt1 bind to the HXT1 promoter constitutively (Figure 6), suggesting that the N-terminal LexA or GFP moiety of the Rgt1 fusion modulates its DNA-binding property by affecting the zinc cluster DNA-binding domain at its N-terminus (aa 46-76). LexA-Rgt1 is shown to be released from DNA in high glucose-grown cells in ChIP experiments using an antibody that specifically recognizes the C-terminus of Rgt1 (Kim et al., 2003; Polish et al., 2005). Glucose induces an intramolecular interaction between the central region of

Rgt1 and its N-terminal DNA-binding domain (Polish et al., 2005), suggesting the view that the C-terminus of LexA-Rgt1 may be hidden and unavailable for antibody recognition in high glucose conditions.

Glucose induces the expression of the HXT1 gene in cells expressing Rgt1-HA,

LexA-Rgt1 and GFP-Rgt1 by ~50, ~10, and ~15 folds, respectively. Thus, taken at face value, DNA-binding alone (by LexA-Rgt1 and GFP-Rgt1) accounts for ~ 4-5-fold repression (Figure 6F). However, I argue that this repression may be associated with the ability of the Rgt1 constructs to interact with Ssn6-Tup1, rather than their ability to bind

DNA. The supporting evidence is that, in response to glucose, Rgt1-HA largely dissociates from Ssn6-Tup1 and that, by contrast, LexA-Rgt1 substantially associate with

Ssn6-Tup 1 (Figures 11 and 12). The Rgt1 interaction with Ssn6-Tup1 is regulated by its phosphorylation state (Roy et al., 2013a). Hence, LexA-Rgt1 may be less efficiently phosphorylated by PKA than Rgt1-HA, enabling it to recruit Ssn6-Tup1 even in the

presence of glucose. This may explain the repression of HXT1 gene expression mediated

by LexA-Rgt1 and GFP-Rgt1 in high glucose conditions. my observations provide

significant insights into the mechanism of glucose regulation of Rgt1 function: 1) Mth1

does not directly regulate the DNA-binding ability of Rgt1; rather, it mediates the Rgt1 interaction with Ssn6-Tup1 by modulating PKA phosphorylation of Rgt1 (Roy et al.,

2013a); 2) dissociation of Rgt1 from DNA occurs in a glucose-dependent manner, but is not absolutely required for the derepression of its target genes; 3) disruption of the Rgt1-

Ssn6-Tup1 interaction is necessary and sufficient to lift Rgt1-mediated repression; 4) the interaction of Rgt1 with Ssn6-Tup1 may be regulated by its phosphorylation state.

In Kluyveromyces lactis, expression of the glucose transporter gene RAG1 is repressed by the Rgt1 ortholog klRgt1 in the absence of glucose. Of note, glucose induction of RAG1 gene expression does not require dissociation of klRgt1from the

RAG1 promoter; klRgt1 remains bound to the RAG1 promoter even in high glucose conditions (Rolland et al., 2006). These results reinforce the view that the primary mechanism of glucose induction of HXT gene expression is not Rgt1 release from HXT

promoters but its dissociation from Ssn6-Tup1. Glucose regulates Rgt1 in a similar

manner, as it does to the glucose repressor Mig1 (Figure 10). Mig1 recruits Ssn6-Tup1

for repression in high glucose conditions (Rolland et al., 2006); however, it dissociates

from Ssn6-Tup1 upon phosphorylation by the Snf1 kinase in glucose-depleted conditions,

resulting in derepression of its target genes (Hedbacker and Carlson, 2008; Treitel et al.,

1998). Thus, Mig1 binds to its target promoters under either repressing or inducing

condition, supporting the view that Snf1 controls glucose repression by modulating the

Mig1-Ssn6-Tup1 interaction (Papamichos-Chronakis et al., 2004). Likewise, PKA

regulates glucose induction by controlling the Rgt1 interaction with Ssn6-Tup1.

Figure 8. Rgt1 phosphorylation at the PKA sites is required for glucose-induction of HXT gene expression. (A) ChIP analysis of LexA-Rgt1-binding to the HXT1 promoter was carried out as described in Figure 6C (left). The anti-LexA antibody was used to precipitate chromatin. Right: qPCR analysis of Rgt1-binding to the HXT1 promoter, as described in Figure 6D. (B) The pHXT1-NAT reporter strain (JKY98) expressing empty vector, LexA-Rgt1 or LexA-Rgt1 (5SA) was spotted on SC-Leu plate containing either 2% galactose or 4% glucose supplemented with 100µg/ml NAT sulfate, as described in Figure 6D. The plates were incubated for 3 days and photographed. (C) Yeast cells (rgt1∆) coexpressing the pHXT1-LacZ reporter plasmid and LexA-Rgt1 or LexA-Rgt1 (5SA) were grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr and assayed for β-galactosidase activity.

Figure 9. The interaction of Rgt1 with Ssn6-Tup1 is critically regulated by its phosphorylation state. (A) ChIP analysis of the interaction of Ssn6 with Rgt1. Left: Yeast cells (rgt1∆) expressing LexA-Rgt1 or LexA-Rgt1 (5SA) were grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr, and cross-linked chromatin was precipitated using anti-Ssn6 antibody. Representative PCRs are shown for the amplification of HXT1 promoter. As a negative control, primer sets were designed to amplify the actin gene promoter region (pACT1). Right: qPCR analysis of Rgt1-binding to the HXT1 promoter, as described in Figure 6D. (B) Co-IP analysis of the interaction of Rgt1 with Ssn6. Yeast cells coexpressing Ssn6-TAP and LexA-Rgt1 (left) or LexA- Rgt1(S5A) (right) were grown in SC-2% galactose medium (-) till mid-log phase and shifted to SC-4% glucose medium (+) for 1 hr. Cell extracts were immunoprecipitated with anti-LexA antibody (IP) and immunoblotted with either anti-LexA or anti-TAP antibody. Expression of Ssn6-TAP was analyzed by Western blot (Input). Quantification of Ssn6-TAP immunoprecipitated with LexA-Rgt1 protein is shown (bottom).

Figure 10. Glucose regulates the function of the two major glucose responsive repressors Rgt1 and Mig1 in a similar manner. (A) A proposed model for glucose induction of HXT gene expression. High glucose disrupts the HXT gene repressor complex formed by Rgt1, Ssn6/Tup1 and Mth1/Std1 by proteasomal degradation of Mth1 via the Rgt2/Snf3 pathway. The PKA-dependent phosphorylation of Rgt1 leads to its dissociation from DNA and HXT gene is expressed. (B) (B) In glucose-limited conditions, the Snf1 kinase phosphorylates and negatively regulates Mig1 by preventing the interaction between Mig1 and Ssn6-Tup1. In high glucose condition, however, Snf1 is inactive, and thereby Mig1 is dephosphorylated and recruit Ssn6-Tup1 to mediate the repression of its target genes. Therefore, the role of phosphorylation of Mig1 and Rgt1 repressors in inducing conditions is to prevent their interaction with Ssn6-Tup1.

Figure 11. qRT-PCR analysis of HXT1 gene expression. (A) Yeast cells (rgt1∆) expressing either empty vector, Rgt1-HA, LexA-Rgt1 or GFP-Rgt1 were grown in SC- 2% galactose medium (-) till mid-log phase (O.D600nm = 1.2-1.5) and shifted to SC-4%

glucose medium (+) for 1 hr. (B) Yeast cells (mth1∆ and RGT2-1) expressing either empty vector, Rgt1-HA or LexA-Rgt1 were grown as described in A. (C) Yeast cells (rgt1∆) expressing either LexA-Rgt1 or LexA-Rgt1 (5SA) were grown as described in A. The HXT1 mRNA levels in A, B and C were quantified by qRT-PCR. The data shown are mean ± SD.

Figure 12. Rgt1-HA and LexA-Rgt1 have different affinities for Ssn6-Tup1. (A) This figure is adapted from the Fig. 4B. Co-IP analysis of the interaction between LexA- Rgt1 and Ssn6-TAP. Yeast cells (WT) coexpressing LexA-Rgt1 and Ssn6-TAP were first grown in SC-2% galactose medium (Gal) till mid log phase and shifted to SC-4% glucose medium for 1 hr. Cell extracts were immunoprecipitated with anti-LexA (IP) and immunoblotted with either anti-LexA or anti-TAP antibody. Expression of Ssn6-TAP was analyzed by Western blot using anti-TAP antibody (Input). Densitometry analysis to quantify Ssn6-TAP immunoprecipitated with LexA-Rgt1 protein was shown (bottom). (B) This figure is adapted from the reference (Roy et al., 2013a). Co-IP analysis of the interaction between Rgt1-HA and LexA-Ssn6. Yeast cells (WT) coexpressing Rgt1-HA and LexA-Ssn6 were grown as described in A. Cell extracts were immunoprecipitated with anti-HA (IP) and immunoblotted with either anti-HA or anti-LexA antibody. Expression of LexA-Ssn6 was analyzed by Western blot using anti-LexA antibody (Input). Densitometry analysis to quantify LexA-Ssn6 immunoprecipitated with Rgt1-HA protein was shown (bottom). (C) This figure is adapted from the reference (Roy et al., 2013a). Co-IP analysis of the interaction between LexA-Ssn6 and Rgt1-HA. Yeast cells (WT) coexpressing Rgt1-HA and LexA-Ssn6 were grown as described in A. Cell extracts were immunoprecipitated with anti-LexA (IP) and immunoblotted with either anti-LexA or anti-HA antibody. Expression of Rgt1-HA was analyzed by Western blot using anti- HA antibody (Input). Densitometry analysis to quantify Rgt1-HA immunoprecipitated with LexA-Ssn6 protein was shown (bottom).

Table 1.1. S. cerevisiae strains used in this study

Strain Genotype Source BY4741 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ (Giaever et al., 2002) FM557 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 (Giaever et al., rgt1::kanMX 2002) YM6545 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 RGT2- (Kaniak et al., 1 2004) JKY98 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study rgt1::kanMX pHXT1::NAT KFY35 Matα his3Δ1 leu2Δ0 ura3Δ0 met15Δ (Giaever et al., mth1::kanMX 2002) KFY56 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ SSN6-TAP- (Ghaemmaghami HIS3MX6 et al., 2003)

Table 1.2. Plasmids used in this study

Plasmid Description Source KFP69 pAD80, 3x-HA-CYC1 terminator, Leu2, 2µ (Kim, 2009) KFP60 pAD80-PRGT1-Rgt1-3x-HA (Mosley et al., 2003) pBM3580 PADH1-LexA-Rgt1, Leu2, 2µ (Ozcan et al., 1996b) pBM3911 PMET25-GFP-Rgt1, Ura3, CEN (Kim et al., 2003)

CHAPTER III

Mth1 regulates the interaction between the Rgt1 repressor and the Ssn6-Tup1 corepressor complex by modulating PKA-dependent phosphorylation of Rgt1

Abstract

Glucose uptake, the first, rate-limiting step of its utilization, is facilitated by glucose transporters. Expression of several glucose transporter (HXT) genes in yeast is repressed

by the Rgt1 repressor, the glucose responsive transcription factor Mth1 and the Ssn6-

Tup1 general corepressor complex when glucose is absent; however, it is derepressed

when Mth1 is inactivated by glucose. Here, I show evidence that Ssn6-Tup1 interferes

with the DNA-binding ability of Rgt1 in the absence of Mth1, and that Rgt1 function

abrogated by Ssn6 overexpression is restored by cooverexpression of Mth1. Thus, Mth1

likely regulates Rgt1 function not by modulating its DNA-binding activity directly but by

functionally antagonizing Ssn6-Tup1. Mth1 does so by acting as a scaffold-like protein to recruit Ssn6-Tup1 to Rgt1. Supporting evidence shows that Mth1 blocks the PKA- dependent phosphorylation of Rgt1 that impairs the ability of Rgt1 to interact with Ssn6-

Tup1. Notably, Rgt1 can bind DNA in the absence of Ssn6-Tup1 but does not inhibit transcription, suggesting that dissociation of Rgt1 from Ssn6-Tup1, but not from DNA, is necessary and sufficient for the expression of its target genes, such as HXT genes.

Together, these findings demonstrate Mth1 as a transcriptional corepressor that facilitates the recruitment of Ssn6-Tup1 by Rgt1.

3.1. Introduction

High aerobic glycolysis—high propensity to ferment rather than oxidize glucose even

when oxygen is abundant—is a hallmark of glucose metabolism in many types of cancer

cells and the budding yeast Saccharomyces cerevisiae (Johnston and Kim, 2005). A key

characteristic of this phenomenon is the increased glucose uptake as a result of elevated

expression of the glucose transporter genes. The budding yeast has at least 6 glucose

transporter genes (HXT1-4, HXT6 and HXT7) whose expressions are induced by glucose

but repressed when glucose is depleted (Diderich et al., 1999; Ko et al., 1993; Ozcan and

Johnston, 1999). Repression of the HXT genes is largely controlled by the HXT repressor

Rgt1, a member of the Gal4 family of transcription factors that contains the zinc

binuclear cluster (Cys6Zn2) DNA-binding domain (Ozcan et al., 1996b). Rgt1 recognizes

a specific DNA sequence 5’-CGGANNA-3’ via the DNA-binding motif in its amino- terminus in vitro (Kim, 2004, 2009; Kim et al., 2003) and synergistically binds to

multiple copies of the sequence in the upstream regions of HXT genes in vivo (Kim et al.,

2003).

Ssn6-Tup1 is a general transcription corepressor complex composed of one molecule

of Ssn6 and four molecules of Tup1 (Varanasi et al., 1996). The complex lacks DNA-

binding ability but is instead recruited to its target promoters by sequence-specific DNA-

binding repressors (Malave and Dent, 2006; Smith and Johnson, 2000b). Ssn6 and Tup1

contain the tetratrico peptide repeat (TPR) and WD domains, respectively, that serve as

protein-protein interaction motifs through which they interact with different binding

partners (Jabet et al., 2000; Schultz et al., 1990; Smith and Johnson, 2000a; Smith et al.,

1995; Sprague et al., 2000). The mechanism of Ssn6-Tup1-mediated transcriptional

repression involves the recruitment of global corepressors such as chromatin and

nucleosome remodelers and the interaction with the RNA transcription machinery

(Malave and Dent, 2006; Smith and Johnson, 2000b). For example, the corepressor

promotes gene repression by associating with histone deacetylases (HDACs) including

Rpd3, Hos1 and Hos2 (Davie et al., 2003). Tup1 interacts with histone H3 and H4, and

its binding to hypoacetylated histones flanking repressor binding sites leads to

nucleosome positioning (Davie et al., 2003; Edmondson et al., 1996). Ssn6-Tup1 is

involved in repression of the genes regulated by diverse signaling pathways (Malave and

Dent, 2006). Among them are HXT genes (Ozcan and Johnston, 1999), glucose-

repressible genes (Nehlin and Ronne, 1990), hypoxia-induced genes (Balasubramanian et al., 1993), DNA damage-response genes (Huang et al., 1998) and haploid-specific genes

(Johnson and Herskowitz, 1985; Komachi et al., 1994). The relief of Ssn6-Tup1- mediated repression comes about through the destruction or inactivation of the individual repressors that leads to dissociation of the repressors from Ssn6-Tup1 and DNA (Smith and Johnson, 2000b).

Rgt1-mediated repression of the HXT genes occurs by a mechanism that requires the paralogous proteins Mth1 and Std1 (Hubbard et al., 1994; Schmidt et al., 1999). Key lines of supporting evidence are: 1) HXT expression is constitutive in yeast cells lacking

MTH1 and STD1 genes (Lafuente et al., 2000; Lakshmanan et al., 2003; Schmidt et al.,

1999); 2) Mth1 and Std1 directly interact with Rgt1 (Polish et al., 2005; Tomas-Cobos et al., 2004); 3) Mth1 and Std1 might form a repression complex with Rgt1 and Ssn6-Tup1 in the absence of glucose (Kim et al., 2003; Lakshmanan et al., 2003). Mth1 and Std1 are

42 degraded by proteasome in the presence of high levels of glucose, resulting in disruption of the represssor complex and thereby derepression of HXT expression (Flick et al., 2003;

Moriya and Johnston, 2004; Pasula et al., 2007). The glucose signal transduction pathway that leads to degradation of Mth1 and Std1 begins at the plasma membrane with the two glucose transporter-related sensor proteins Rgt2 and Snf3 (Ozcan et al., 1998). There are also dominant mutations in the MTH1 gene (HTR1-23, DGT1 or BCP1) that render Mth1 resistant to glucose-induced degradation (Kim et al., 2006), resulting in the constitutive repression of HXT expression (Gamo et al., 1994; Schulte et al., 2000). Expression of the

MTH1 gene is also repressed by Mig1 in high glucose conditions, reinforcing the inhibitory effect of glucose on Mth1 function (Kaniak et al., 2004; Kim et al., 2006). The ability of Rgt1 to bind to HXT promoters is correlated with its phosphorylation state:

Rgt1 is phosphorylated at a basal level and binds to the promoters in the absence of glucose; it is hyperphosphorylated by PKA and dissociated from the promoters when glucose levels are high (Kim and Johnston, 2006; Palomino et al., 2006). Rgt1 is also hyperphosphorylated and does not bind DNA in cells lacking Mth1 (Flick et al., 2003;

Mosley et al., 2003), leading to the hypothesis that Mth1 and Std1 prevent the PKA- dependent phosphorylation of Rgt1 that impairs the DNA-binding ability of Rgt1.

The aim of this study is to investigate the role of Mth1 in the mechanism of Rgt1- mediated repression. I show that glucose-induced, PKA-dependent phosphorylation is a crucial step leading to dissociation of Rgt1 from Ssn6-Tup1 and consequently to derepression of HXT gene expression. Mth1 blocks such phosphorylation by mediating the interaction of Rgt1 with Ssn6-Tup1, thereby facilitating the formation of a functional repressor complex that inhibits transcription of HXT genes. I further show that Mth1 acts

to antagonize Ssn6-Tup1 inhibition of Rgt1 function; however, its expression is repressed

by Ssn6-Tup1 via Mig1 in high glucose conditions. Together, these results identify a

functional interaction between Mth1 and Ssn6-Tup1 and provide a novel insight into the

mechanism of Rgt1-Ssn6-Tup1-mediated repression.

3.2. Materials and Methods

3.2.1. Yeast strains, Gene deletion, and Plasmids

Yeast strains used in this study were listed in Table 2. Except where indicated, yeast

strains were grown in YP (2% bacto-peptone, 1% yeast extract) and SC (synthetic yeast

nitrogen base medium containing 0.17% yeast nitrogen base and 0.5% ammonium

sulfate) supplemented with the appropriate amino acids and carbon sources. Genes were

disrupted with NatMX or KanMX cassette by homologous recombination method

(Goldstein et al., 1999; Wach et al., 1994). The HXT-NAT reporter strains were constructed by replacing HXT1, HXT2, and HXT3 ORFs with the NatMX coding region by homologous recombination. JKP173 (LexA-Ssn6) and JKP231 (Ssn6-HA) were constructed by cloning the SSN6 gene into the LexA and HA plasmids, respectively.

JKP232 (Rgt1 (210-250Δ)-HA) and JKP233 (Rgt1 (310-360Δ)-HA) were constructed by

‘gap repair’.

3.2.2. Chromatin immunoprecipitation (ChIP)

ChIP was performed as described previously (Kim et al., 2003). Briefly, Yeast strains were grown till mid-log phase (O.D600nm = 1.2-1.5) and incubated with formaldehyde

(1% final concentration) at room temperature for 15 to 20 minutes. The cross-linking

reaction was quenched by adding glycine to a final concentration of 125 mM for 5

minutes. The cells were disrupted by vortexing with acid-washed glass beads in ice cold

ChIP lysis buffer (50 mM HEPES-KOH, pH 7.5, 150 mM NaCl, 1% Triton X-100, 0.1%

Na-deoxycholate) containing protease and phosphatase inhibitors. The lysate was

sonicated (ultrasonic cell disruptor with a microtip) five times with 10 sec pulse. The

genomic DNA fragments, with averaged 200-500 bp size, were immunoprecipitated with

HA or Myc antibody (Santa Cruz) conjugated with agarose bead. After washing the immunoprecipitated beads with ChIP high salt buffer (ChIP lysis buffer containing 500

mM NaCl instead of 150 mM NaCl) and then ChIP wash buffer (10 mM Tris-HCl, pH

8.0, 250mM LiCl, 0.5% NP40, 0.5% Na-deoxycholate, 1 mM EDTA), DNA was eluted from both immunoprecipitated and 1/100 input samples by incubating the samples in

ChIP elution buffer (50 mM Tris-HCl, pH 8.0, 1% SDS, 10 mM EDTA) at 65 oC for 6-8

hr. Finally, the DNAs were purified by QIAquick PCR purification kit (Qiagen). The

amount of immunoprecipitated DNA was quantified by real-time PCR using SsoFast

Evagreen reagent (Bio-Rad) in CFX96 Real-time thermal cycler (Bio-Rad) using primer

pairs directed against HXT1, HXT2, HXT3, SUC2 or GAL1 promoters. As a negative

control, primer sets were designed to amplify the actin gene promoter region. DNA- binding of Rgt1 or Mig1 was determined by the ratio of immunoprecipitated (IP)/target promoters relative to the IP/ACT1 promoter divided by the ratio of input (INPUT)/target promoters relative to the INPUT/ACT1 promoter. All of the shown data were the averages of three independent ChIP experiments with error bars representing standard deviations (S.D). The sequences of the primers used for ChIP were: HXT1, 5’-

ATATAATTCCCCCCTCCTGAAG-3’ and 5’-TGATTCTACGTTTTTGCAAGC-3’;

HXT3, 5’-CTTCT-CGAGATAACACCTGG-3’ and 5’-CCACGAAGCTTTCTCTGTG-

3’; SUC2, 5’-GTAGTTCTCGCTC-CCCCAG-3’ and 5’-

TGGGGTCGATTAACGCTACG-3’; GAL1,5’-CGAATCAAATTAACAACCATA-

GGATGATA-3’ and 5’-TATAGTTTTTTCTCCTTGACGTTAAAG-3’; ACT1, 5’-

CCTGAACGAAAC-CACTCAGAAGAA-3’ and 5’-

TTAAGGGTTTTGAGGATCCGATAAGG-3’.

3.2.3. Western blot and Immunoprecipitation (IP) assays

For Western blot analysis, yeast cells (O.D600 = 1.5) were collected by centrifugation

at 3,000 rpm in a table-top centrifuge for 5 min. Cell pellets were resuspended in 100 µl

of SDS-buffer (50 mM Tris-HCl, pH, 6.8, 10% glycerol, 2% SDS, 5% β- mercaptoethanol) and boiled for 5 min. After the lysates were cleared by centrifugation at 12,000 rpm for 10 min., soluble proteins were resolved by SDS-PAGE and transferred to PVDF membrane (Millipore). The membranes were incubated with appropriate antibodies (anti-HA, anti-LexA, anti-Myc, anti-Ssn6, and anti-GFP antibodies, Santa

Cruz) in TBST buffer (10 mM Tris-HCl, pH, 7.5, 150 mM NaCl, 0.1% Tween-20) and proteins were detected by the enhanced chemiluminescence (ECL) system (Pierce). For

IP, yeast cells were disrupted by vortexing with acid-washed glass beads in ice cold

NP40 buffer (1% NP40, 150 mM NaCl, 50 mM Tris-HCl, pH 8.0) containing protease inhibitors. The cell lysates were incubated with appropriate antibodies at 4 oC for 3 hr and

further incubated with protein A/G-conjugated agarose beads (GE Healthcare) at 4 oC for

1 hr. The precipitated agarose beads were washed three times with ice cold NP40 buffer

containing protease inhibitors and boiled in 50 µl of SDS-PAGE buffer. The resulting proteins were subjected to Western blot analysis.

3.2.4. Quantitative RT-PCR (qRT-PCR)

Yeast cells were grown in YP medium containing 2% galactose till mid-log phase

(O.D600nm = 1.2 ~ 1.5) and shifted to YP medium containing 4% glucose for 1hr. Total

RNA was extracted by RNeasy mini kit (Qiagen) following manufacturer’s protocol and

2µg of total RNA was converted to cDNA by qScript cDNA supermix (Quanta

Biosciences). cDNA was analyzed by qRT-PCR using SsoFast Evagreen reagent (Bio-

Rad) in CFX96 Real-time thermal cycler (Bio-Rad). ACT1 was used as an internal control to normalize expression of HXT1, HXT2 or HXT3 gene. All of the shown quantification data were the averages of three independent experiments with error bars representing standard deviations (S.D).

3.2.5. β-galactosidase assay

To assay β-galactosidase activity with yeast cells expressing the HXT1-LacZ reporter, the yeast cells were grown to mid-log phase and the assay was performed as described previously (Kaniak et al., 2004). Results were given in Miller Units ((1,000 x

O.D420nm)/(T x V x O.D600nm), where T was the incubation time in minutes, and V was the

volume of cells in milliliters). The reported enzyme activities were averages of results

from triplicates of three different transformants.

3.3. Results

3.3.1. The DNA-binding activity of Rgt1 is antagonistically regulated by Ssn6-

Tup1 and Mth1.

Rgt1 forms a repressor complex with Ssn6-Tup1 on and mediates repression of the HXT

promoters in the absence of glucose (Kim et al., 2003) and, this occurs in an Mth1/Std1-

dependent manner (Flick et al., 2003; Mosley et al., 2003; Schmidt et al., 1999).

However, the underlying mechanism of this process is not yet understood. To understand

the roles of Ssn6-Tup1, Mth1, and Std1 in the formation of the complex, I first

determined whether these components regulate the ability of Rgt1 to bind to HXT

promoters using ChIP analysis. The RGT2-1 mutation causes constitutive (glucose- independent) expression of HXT genes (Ozcan and Johnston, 1996), probably by inducing degradation of Mth1 and Std1 in the absence of glucose (Pasula et al., 2007).

ChIP analysis showed that Rgt1-binding to the HXT1 promoter is significantly abolished in mth1Δstd1Δ and RGT2-1 mutants, as reported previously (Flick et al., 2003; Pasula et al., 2007), but is constitutive in ssn6Δ and tup1Δ mutants (Figure 13A), without significant changes in Rgt1 protein levels (Figure 13A, top right panel). More importantly, the DNA-binding defect of Rgt1 in mth1Δstd1Δ and RGT2-1 cells was restored by the removal of the TUP1 (mth1Δstd1Δtup1Δ) or SSN6 (RGT2-1 ssn6Δ) gene from the mutants (Figure 13B). An mth1Δstd1Δssn6Δ mutant displays an extremely slow growth phenotype, compared with that of the mth1Δstd1Δtup1Δ mutant, so that it could not be used in this study. These results suggest that Rgt1 by itself is able to bind to its

DNA target sites but this ability is positively and negatively regulated by Mth1/Std1 and

Ssn6-Tup1, respectively.

Figure 13. Ssn6-Tup1 negatively regulates the ability of Rgt1 to bind to its target promoters. (A) ChIP analysis of Rgt1-binding to the HXT1 promoter in yeast cells expressing Rgt1-HA. Yeast cells of the indicated genotypes were grown in SC-2% galactose (-) and shifted to SC-4% glucose (+) for 1 hr. Cross-linked chromatin was precipitated for ChIP analysis using anti-HA antibody, and representative PCRs were shown (left panel). As a negative control of Rgt1 DNA-binding, primer sets were designed to amplify the Actin gene promoter region (pACT1), which does not contain the Rgt1-binding sequence (5’CGGANNA3’) (middle panel). Western blot analysis of Rgt1- HA expression using anti-HA antibody (right panel). (B) qPCR analysis of Rgt1-binding to the HXT1 promoter in the designated yeast strains. The amount of immunoprecipitated (IP) DNA was quantified by qPCR with primer pairs directed against the HXT1 promoter (pHXT1). IP/Input ratio was determined by the ratio of IP/pHXT1 relative to the IP/pACT1 divided by the ratio of INPUT/pHXT1 relative to the INPUT/pACT1. The data shown were averages of three independent experiments with error bars showing mean ± S.D. (C) Western blot analysis of expression of Mth1-myc using anti-Myc antibody. (D) Western blot analysis of expression of Ssn6 using anti-Ssn6 antibody. Actin serves as a loading control in panels C and D. (E) qRT-PCR analysis of HXT1 mRNA expression. The quantification data of HXT1 mRNA expressions were averages of three independent experiments with error bars showing mean ± S.D

These findings prompted us to examine whether Ssn6-Tup1 and Mth1 downregulate

each other. Western blot analysis showed that Mth1 levels are elevated in glucose-grown

ssn6Δ and tup1Δ mutants, compared with those of wild type cells (Figure 13C). Elevated levels of Mth1 in ssn6Δ and tup1Δ mutants is perhaps due to derepression of MTH1

expression (Figure 19), consistent with our previous reports that MTH1 expression is

repressed by Mig1-Ssn6-Tup1 complex at high concentration of glucose (Kaniak et al.,

2004; Kim and Johnston, 2006). Mth1 was barely detectable in RGT2-1 and RGT2-1

ssn6Δ mutants, suggesting that Ssn6-Tup1 is not involved in degradation of Mth1. I also

confirmed that there are no appreciable changes in the levels of Ssn6 in mth1Δstd1Δ and

RGT2-1 mutants (Figure 13D). Taken together, these results suggest that Ssn6-Tup1

negatively regulates the DNA-binding ability of Rgt1 by repressing expression of the

MTH1 gene.

3.3.2. Rgt1 binds to its target promoters in the absence of Ssn6-Tup1 but does

not repress transcription

My finding that Rgt1 binds to the HXT1 promoter in the absence of Ssn6-Tup1

(Figure 13A) raised the possibility that glucose might not regulate the DNA-binding

ability of Rgt1. To test this possibility, I examined the expression of HXT1 mRNA in the

mutants tested above using qRT-PCR analysis (Figure 13E). The HXT1 mRNA is

constitutively expressed in mth1Δstd1Δ and RGT2-1 mutants, perhaps due to inability of

Rgt1 to bind to the HXT1 promoter in the mutants (Figure 13A), as reported previously

(Mosley et al., 2003; Pasula et al., 2007). Most notably, however, Rgt1 was shown to be

able to bind to the promoter constitutively in cells lacking SSN6 (ssn6Δ and RGT2-1

ssn6Δ mutants) or TUP1 (tup1Δ and mth1Δ std1Δ tup1Δ mutants) (Figure 13A), but did

not significantly inhibit glucose-induction of HXT1 expression (Figure 13E). Thus,

myresults suggest that dissociation of Ssn6-Tup1 from Rgt1 is sufficient for derepression

of HXT gene expression regardless of the presence of both glucose and Mth1/Std1.

3.3.3. Mth1 acts to functionally antagonize the ssn6-Tup1 complex

Because Rgt1 DNA-binding is oppositely regulated by Mth1/Std1 and Ssn6-Tup1

(Figure 13A), I assessed the functional interaction between Mth1 and Ssn6-Tup1 for

regulating Rgt1 function in yeast cells overexpressing Ssn6 or cooverexpressing Ssn6 and

Mth1. Towards this aim, I constructed HXT reporter strains that express the

nourseothricin (NAT) resistance gene under the control of the HXT1, HXT2 or HXT3

promoter (Figure 14A). Hence, the growth of the HXT reporter strains in a NAT-

containing medium is dependent on the activity of the HXT promoters. I observed that

HXT1-NAT and HXT3-NAT reporter strains grow only in high-glucose medium, whereas

cells carrying the HXT2-NAT reporter grow in raffinose (low glucose) medium; however,

none of them grow in glycerol/ethanol medium. These results are consistent with the

previous report that various HXT promoters are expressed differently in the various

conditions (Ozcan and Johnston, 1995). Interestingly, however, the HXT-NAT reporter strains, the HXT1-NAT reporter strain in particular, were able to grow in a galactose or glycerol/ethanol medium when Ssn6 is overexpressed from a high copy plasmid (2µ)

(Figure 14B). The expression patterns of HXT mRNAs in cells overexpressing Ssn6 were similar to the growth patterns of the cells (Figure 14C). ChIP analysis also showed that

Rgt1-binding to the HXT1 promoter is significantly reduced when Ssn6 is overexpresssed

(Figure 14D, ChIP-Rgt1, Ssn6). Next, I asked whether Ssn6 overexpression inhibits the

DNA-binding ability of other Ssn6-Tup1 recruiters. The glucose repressor Mig1 is activated and mediates repression of its target genes such as SUC2 and GAL1 by

recruiting Ssn6-Tup1 in high glucose conditions (Treitel and Carlson, 1995). I found that

Mig1 binds to its target promoters constitutively (Figure 14D, right panel), as reported

previously (Papamichos-Chronakis et al., 2004); however, the DNA-binding ability of

Mig1 is not significantly affected by Ssn6 overexpression (Figure 14D, ChIP-Mig1,

Ssn6).

Expression of HXT genes is regulated by not only Rgt1 but also other transcriptional

factors, such as Sko1 (Tomas-Cobos et al., 2004) and Mig1 (Ozcan and Johnston, 1996),

raising a possibility that association of these proteins with HXT promoters may influence

Rgt1 function. To eliminate this possibility, I used a plasmid reporter that contains 6

copies of the Rgt1 DNA-binding sequence (Rgt1-BS) without intervening sequences

between Rgt1-BSs followed by the lacZ gene. Because expression of this reporter gene

is solely regulated by Rgt1, this reporter system has been successfully used to measure

the transcriptional repression activity of Rgt1 (Kim, 2009). I confirmed that expression of

this reporter gene is negatively regulated by Mth1/Std1 and Ssn6 (Figure 14E, left panel).

More significantly, Ssn6 overexpression (Ssn6-HA) resulted in derepression of the lacZ gene in yeast cells grown under repressing conditions (galactose or glycerol + ethanol); however, this derepression was significantly suppressed by cooverexpression of Mth1

(Ssn6-HA + LexA-Mth1) (Figure 14E, right panel). These results and those in Figure 13 suggest that Ssn6-Tup1 inhibits the ability of Rgt1 to bind to its target promoters and this

inhibition is overcome by Mth1.

Figure 14. Ssn6 overexpression induces derepression of HXT expression in repressing conditions. (A) HXT-NAT reporter strains were streaked in YP plate containing 4% glucose (Glu), 5% glycerol + 2% ethanol (Gly/EtOH) or 2% raffinose (Raf) supplemented with 100µg/ml NAT-sulfate. (B) Ssn6-HA (JKP231) was overexpressed from a high-copy 2µ plasmid in all three NAT reporter strains. Yeast cells were spotted on YP plate containing 5% glycerol + 2% ethanol (Gly/EtOH) or 2% galactose (Gal) supplemented with 100 µg/ml NAT-sulfate. The first spot of each row represents a count of 5 x 107 cell/ml, which is diluted 1:10 for each spot thereafter. Cells were incubated for 2 days in Gal + NAT plate and 3 days in Gly/EtOH plates, respectively. (C) qRT-PCR analysis of mRNA expression of HXT1, HXT2 and HXT3 genes. mRNA was isolated from yeast cells (BY4741) expressing either empty HA vector (V, 2µ vector only) or Ssn6-HA (Ssn6, JKP231) were grown in SC medium containing 5% glycerol + 2% ethanol (Gly/EtOH) or 2% galactose (Galan et al.) till mid- log phase (OD600nm = 1.2-1.5). The data were averages of three independent experiments with error bars showing mean ± S.D. (D) Yeast cells (BY4741, WT) cooverexpressing Ssn6-HA with either Rgt1-HA or Mig1-myc (pBM3076) were grown in SC-2% galactose

(-) and shifted to SC-4% glucose (+) for 1 hr. Cross-linked chromatin was precipitated for ChIP analysis of the indicated Rgt1 (pHXT1, pHXT2, and pHXT3) and Mig1 (pSUC2 and pGAL1) DNA target sites and representative PCRs were shown (upper panels). The results of qPCR analysis of the binding of Rgt1 and Mig1 to their respective target promoters in yeast cells (lower panels) were expressed as IP/Input ratio as described above (Figure 13B). The data were averages of three independent experiments with error bars showing mean ± S.D. (E) Yeast cells of indicated genotypes were transformed with a plasmid containing 6 copies of Rgt1-binding DNA sequence fused to the lacZ gene (6x Rgt1-BS-lacZ (JHB93)) (left). BY4741 (WT) was transformed with JHB93 along with an empty HA plasmid, a plasmid expressing Ssn6-HA, or plasmids expressing Ssn6-HA and LexA-Mth1 (pBM4150) (right). Transformants were grown in SC-5% glycerol + 2% ethanol medium (white bars), shifted to SC-2% galactose (grey bars) or SC-4% glucose (black bars) media for 1 hr, and assayed for β-galactosidase activity.

3.3.4. Mth1 mediates the interaction of Rgt1 with ssn6-Tup1, enabling Rgt1 to recruit Ssn6-Tup1 to HXT promoters

Given that Ssn6-Tup1 negatively regulates Rgt1-mediated repression (Figures 13 and

14), I examined the role of Mth1 in regulating Rgt1 function by assessing the interaction between Rgt1 and Ssn6 by co-IP and Western blot analyses. My results showed that Rgt1 interacts with both Mth1 and Ssn6 in cells grown in glucose-depleted medium, as reported previously (Polish et al., 2005), and that Rgt1-Mth1 interaction is not affected by removal of the SSN6 gene (Figure 15A, left panel). Surprisingly, however, Rgt1-Ssn6 interaction was abolished in the mth1Δ mutant (Figure 15A, right panel) and in the RGT2-

1 mutant (Figure 15B, RGT2-1), where Mth1 is constitutively degraded by proteasome

(Pasula et al., 2007). However, this interaction occurs constitutively in the HTR1-23 mutant expressing a degradation-resistant Mth1 (Figure 15B, HTR1-23).

The domains of Rgt1 responsible for the interaction with Ssn6 (aa 210-250) and Mth1

(aa 310-360), respectively,were previously identified (Polish et al., 2005). To obtain compelling evidence whether Rgt1 interaction with Ssn6 or Mth1 affects Rgt1 function, I tested mutant Rgt1 proteins that lack the Ssn6- and Mth1-binding sites for their ability to

54 interact with Ssn6 and Mth1, respectively. Rgt1 (∆210-250) was able to interact with

Mth1, but not with Ssn6 (Figure 16C). Furthermore, this mutant Rgt1 was able to bind to the HXT1 promoter (Figure 16B), but did not repress transcription (Figure 16A). In contrast, Rgt1 (∆310-360) did not strongly interact with both Mth1 (Figure 16D) and the

HXT1 promoter (Figure 16B), leading to derepression of the HXT1 gene (Figure 16A).

More importantly, Rgt1 (∆310-360) was not able to interact with Ssn6 either (Figure

16D), consistent with the results that Mth1 is required for the interaction between Rgt1 and Ssn6 (Figure 15A). Taken together, these results suggest that Mth1-dependent interaction of Rgt1 with Ssn6-Tup1 enables Rgt1 to bind HXT promoters and thus leads to the formation of a functional Rgt1-Ssn6-Tup1 repressor complex on the promoters.

3.3.5. Mth1 prevents the PKA-dependent phosphorylation of Rgt1 that impairs the ability of Rgt1 to interact with Ssn6-Tup1

The ability of Rgt1 to bind to HXT promoters is largely correlated with the phosphorylation state of Rgt1 regulated by Mth1 (Flick et al., 2003; Kim et al., 2003;

Mosley et al., 2003). Because our findings show that the DNA-binding ability of Rgt1 is inhibited by Ssn6-Tup1 (Figure 13), I determined whether Ssn6-Tup1 modulates Rgt1 function by regulating its phosphorylation state using Western blot analysis. Rgt1 was shown to be hyperphosphorylated constitutively in the mth1Δ mutant (Figure 17A), as reported previously (Flick et al., 2003; Mosley et al., 2003). However, the phosphorylation state of Rgt1 was not significantly changed in ssn6Δ and tup1∆ mutants, as compared with that of wild-type cells, regardless the presence of glucose. Rgt1 was

55 also shown to be constitutively hyperphosphorylated in mth1Δstd1Δ, RGT2-1, RGT2-1 ssn6Δ, and mth1Δstd1Δtup1Δ mutants (Figure 17A).

Rgt1 is hyperphosphorylated in the presence of high levels of glucose (Flick et al.,

2003; Kim et al., 2003; Mosley et al., 2003), and this phosphorylation is catalyzed by

PKA (Kim and Johnston, 2006; Palomino et al., 2006). To obtain direct evidence that

Mth1 regulates Rgt1 phosphorylation by PKA, I determined the phosphorylation state of

Rgt1 in yeast cells lacking functional Mth1, PKA or both. My results showed that hyperphosphorylation of Rgt1 in the mth1∆ mutant is attenuated when the mutant PKA with retarded activity (tpkw1 (Toda et al., 1987b)) is expressed in the mutant (mth1Δ tpkw1) (Figure 17B). I also found that Rgt1 binds to and represses the HXT1 promoter in mth1 ∆tpkw1 mutant constitutively (Figures 17C and 17D). These observations suggest that Mth1, but not Ssn6-Tup1, inhibits Rgt1 phosphorylation by PKA.

Given that Mth1 is required for the interaction between Rgt1 and Ssn6-Tup1 (Figure

15) and that Mth1 regulates Rgt1 phosphorylation (Figure 17B), I investigated whether the phosphorylation state of Rgt1 affects its interaction with Ssn6-Tup1 using co-IP and

Western blot analysis. Rgt1 interaction with Ssn6 was not observed in the mth1Δ mutant but was strongly detected in the mth1∆ tpkw1 mutant (Figure 17E). Furthermore, a mutant

Rgt1 that lacks PKA phosphorylation sites (Rgt1-S5A; Ser96, Ser146, Ser202, Ser283, and Ser284 (Kim and Johnston, 2006) was shown to interact with Ssn6 constitutively

(Figure 17F), thereby leading to constitutive repression of HXT1 expression ( Figure 21).

Thus, these results suggest that hyperphosphorylated Rgt1 does not interact with Ssn6-

Tup1 and that the role of PKA-catalyzed Rgt1 phosphorylation is to dissociate Rgt1 from

Ssn6-Tup1 and consequently from HXT promoters.

Figure 15. Mth1 is required for the interaction of Rgt1 with Ssn6-Tup1. (A) Co-IP analysis of the interaction of Rgt1 with Mth1 or Ssn6. Yeast cells of indicated genotypes coexpressing Rgt1-HA and either GFP-Mth1 (pBM4748) (left) or LexA-Ssn6 (JKP173) (right) were grown in SC-2% galactose medium. Cell extracts were immunoprecipitated with anti-HA (IP) and immunoblotted with either anti-GFP or anti-LexA antibodies. Expression of GFP-Mth1 (left) or LexA-Ssn6 (right) was analyzed by Western blot using either anti-GFP or anti-LexA antibody (Input). (B) Yeast cells of indicated genotypes were co-transformed with plasmids expressing Rgt1-HA and LexA-Ssn6. Cells were first grown in SC-2% galactose medium (Gal) and shifted to SC-4% glucose medium (Glu) for 1 hr. Cell extracts were subjected to IP with anti-HA or anti-LexA antibody and followed by immunoblotting with anti-HA or anti-LexA antibody. Expression of LexA- Ssn6 or Rgt1-HA was analyzed by Western blot using either anti-LexA or anti-HA antibody (Input).

Figure 16. Rgt1 function is regulated positively and negatively by Mth1 and Ssn6, respectively. (A) Yeast cells (rgt1Δ) were co-transformed with each of the three plasmids expressing Rgt1-HA (KFP60), Rgt1 (Δ210-250)-HA (JKP232), and Rgt1 (Δ310-360)- HA (JKP233) and the pHXT1-lacZ reporter plasmid (pBM2636). Transformants were first grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr and assayed for β-galactosidase activity. (B) ChIP analysis of Rgt1-binding to the HXT1 promoter in yeast cells (rgt1Δ) expressing Rgt1-HA, Rgt1 (Δ210-250)-HA and Rgt1 (Δ310-360)-HA using anti-HA antibody. The results of qPCR analysis of Rgt1- binding to the HXT1 promoter in yeast cells expressed as IP/Input as described in Figure 12B using the pACT1 promoter as a negative control of Rgt1 DNA-binding (right panel). (C) Co-IP analysis of the interaction of Rgt1 with Ssn6. Yeast cells (rgt1Δ) coexpressing Rgt1-HA (1-1170 (full-length) or Δ210-250) with LexA-Ssn6 were grown in 2% galactose till mid-log phase (OD600nm = 1.2-1.5). Cell extracts were immunoprecipitated with anti-HA antibody and followed by immunoblotting with anti-LexA antibody. (D) Co-IP analysis of the interaction of Rgt1 with Mth1. Yeast cells (rgt1Δ) coexpressing Rgt1-HA (1-1170 or Δ310-360) with Mth1-myc were grown as described in panel C, and cell extracts were immunoprecipitated with anti-HA antibody and followed by immunoblotting with anti-Myc antibody. Expression of LexA-Ssn6 or Mth1-myc was analyzed by Western blot using either anti-LexA or anti-Myc antibody in panels C and D (input).

3.3.6. The Ssn6-Tup1 complex is recruited to HXT promoters in an Rgt1- dependent manner

Although my results suggest that Rgt1 interaction with Ssn6-Tup1 is regulated by glucose (Figure 15), it is not clear whether Ssn6-Tup1 is recruited to HXT promoters through this interaction. To assess the recruitment of Ssn6-Tup1 to HXT promoters, I performed ChIP analysis of Ssn6-binding to the HXT1 promoter using anti-Ssn6 antibody. I found that Ssn6 binds to the promoter in the absence of glucose, but is removed from the promoter in the presence of high levels of glucose (Figure 18).

However, this binding was not observed in the rgt1∆ mutant, suggesting that recruitment of Ssn6-Tup1 to HXT promoters occurs in an Rgt1-dependent manner. Similar observations were made in the mth1∆ mutant, perhaps due to the inability of Rgt1 to bind to the promoter in the mutant (Figure 13A), highlighting the role of Mth1 as a mediator for the interaction of Rgt1 with Ssn6-Tup1 (Figure 15).

3.4. Discussion

This study investigates the role of Mth1 in the mechanism of Rgt1-mediated repression, with the aim of understanding the glucose regulation of HXT expression.

New findings include: (1) Mth1 mediates the interaction of Rgt1 with Ssn6-Tup1, leading to the formation of a functional repressor complex on Rgt1 target genes; (2) mutational removal or glucose inactivation of Mth1 leads to the PKA-dependent phosphorylation of

Rgt1, which keeps Rgt1 from associating with Ssn6-Tup1; (Ma et al.) dissociation of

Rgt1 from Ssn6-Tup1 is the most critical event for glucose-induction of HXT genes.

These findings support our previous observations that downregulation of Mth1 level by

glucose is a critical event for derepression of Rgt1 target genes (Kaniak et al., 2004; Kim

and Johnston, 2006). Mth1 is a common target of two glucose signaling pathways: it is

degraded in proteasome via the Rgt2/Snf3 pathway; its expression is repressed by the

Snf1-Mig1 pathway. Mth1 inactivation facilitates Rgt1 phosphorylation by the cAMP-

PKA pathway, leading to dissociation of Rgt1 from Ssn6-Tup1 and consequently from its

DNA target sites (Jouandot et al., 2011). Therefore, these three different glucose

signaling pathways converge on Rgt1 to regulate expression of HXT genes (Figure 18C).

Notably, I show that Ssn6-Tup1, although required for Rgt1-mediated repression, acts

to inhibit, rather than stimulate, Rgt1 function. Ssn6-Tup1 does so by inhibiting the

DNA-binding ability of Rgt1 in the absence of glucose (which is antagonized by Mth1)

and by repressing MTH1 expression via the Snf1-Mig1 pathway in high glucose conditions. This suggests that Ssn6-Tup1 can negatively regulate its recruiting DNA-

binding transcription factor. As evidenced in Figure 14, Mig1, an Ssn6-Tup1 recruiter, is

not negatively regulated by Ssn6-Tup1, supporting the view that Ssn6-Tup1 acts

differently on different repressors in yeast cells under identical growth conditions. The

biological significance of this phenomenon is not fully understood, but might be related

to the differential regulation of Ssn6-Tup1 target genes in response to the same stimulus.

For instance, Mig1 and Rgt1 are positively and negatively regulated by glucose. Ssn6-

Tup1 in high glucose condition is recruited to Mig1 but should not be associated with

Rgt1. In addition, Mig1 occupies GAL1 continuously under either repressing or inducing

conditions (Papamichos-Chronakis et al., 2004). Therefore, the corepressor complex in those conditions may actively inhibit its interaction with Rgt1 while associating with

Mig1, thereby avoiding dysregulation of genes regulated by the two glucose-responsive

transcription repressors.

It is not known how Ssn6-Tup1 interferes with the DNA-binding activity of Rgt1.

The purified N-terminal fragment of Rgt1 containing a DNA-binding motif is able to

bind DNA in the absence of Mth1 in vitro, suggesting that the Rgt1 DNA-binding

domain, by itself, can bind DNA (Kim et al., 2003). Ssn6 and Tup1 contain TPR and

WD40 domains, respectively (Jabet et al., 2000; Schultz et al., 1990; Sprague et al.,

2000), and appear to interact with different repressors via the domains in different

manners (Smith et al., 1995; Tzamarias and Struhl, 1995). The Ssn6-binding region in

Rgt1 (aa 210-250) is located close to the Zn cluster DNA-binding motif (Polish et al.,

2005). These observations suggest that Rgt1-Ssn6-Tup1 interaction is transient, but sufficient to induce a conformational change of Rgt1 and lead to dissociation of Rgt1 from HXT promoters. A physical interaction of Mth1 with Rgt1 prevents this from happening, enabling Rgt1 to form a functional repressor complex with Ssn6-Tup1 on

HXT promoters.

Previous evidence shows that Ssn6-Tup1 is also actively involved in induction of gene expression (Mennella et al., 2003; Papamichos-Chronakis et al., 2004; Proft and

Struhl, 2002) and can be recruited to its target promoters in a manner independent of sequence-specific DNA-binding proteins (Buck and Lieb, 2006; Desimone and Laney,

2010; Hanlon et al., 2011; Papamichos-Chronakis et al., 2004). A recent work also shows that Ssn6-Tup1 exerts its function by masking the activation domain of a DNA-binding repressor and thereby preventing recruitment of the coactivators necessary for transcriptional activation (Wong and Struhl, 2011). Glucose-induction of HXT expression

61 is not inhibited by deletion of SSN6 or TUP1 gene, suggesting that Ssn6-Tup1 does not act as an activator of the HXT genes (Ozcan and Johnston, 1996). My findings in this study also indicate that Ssn6-Tup1 is recruited to HXT promoters by Rgt1 and this recruitment occurs in an Mth1-dependent manner in the absence of glucose. However,

Ssn6-Tup1 is dissociated from Rgt1 and consequently from the HXT promoters upon glucose-induced downregulation of Mth1, reinforcing the view that Mth1 plays a key role in recruitment of Ssn6-Tup1 to Rgt1.

Regulation of Rgt1 function is mechanistically similar to that of Mig1, which is phosphorylated and negatively regulated by Snf1 kinase (Hedbacker and Carlson, 2008;

Treitel et al., 1998). Ssn6-Tup1 is recruited to only unphosphorylated Mig1 in the presence of high glucose and mediates repression of Mig1-target genes including SUC2.

Mig1-Ssn6-Tup1 interaction is disrupted when Mig1 is phosphorylated by Snf1 in low glucose conditions, leading to derepression of these genes (Papamichos-Chronakis et al.,

2004). Similarly, Rgt1-Ssn6-Tup1 interaction is disrupted when Rgt1 is phosphorylated by PKA in high levels of glucose, leading to derepression of HXT gene expression.

Therefore, it is likely that the role of phosphorylation of Mig1 and Rgt1 repressors in inducing conditions is to prevent them from associating with Ssn6-Tup1. Furthermore,

Rgt1 binds to the HXT1 promoter in the absence of Ssn6 or Tup1 in high glucose-grown cells, but does not repress the promoter (Figure 13), reinforcing the view that glucose- induction of HXT expression is primarily due to disruption of the interaction of Rgt1 with

Ssn6-Tup1, rather than due to dissociation of Rgt1 from HXT promoters.

Figure 17. Rgt1 phosphorylation by PKA leads to the disruption of its interaction with Ssn6-Tup1. (A) Glucose-induced phosphorylation of Rgt1. Yeast cells expressing Rgt1-HA were grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr. Rgt1-HA was subjected to Western blot analysis using anti-HA antibody. (B) PKA phosphorylation of Rgt1. Western blot analysis of Rgt1, as described in panel A. (C) ChIP analysis of Rgt1 binding to the HXT1 promoter. Representative PCRs were shown (left panel). The results of qPCR analysis of Rgt1-binding to the HXT1 promoter expressed as IP/Input ratio as described in Figure 13B using the pACT1 promoter as a negative control of Rgt1 DNA-binding (right panel). (D) Induction of HXT1 expression in yeast cells carrying the pHXT1-lacZ reporter plasmid (pBM2636). Yeast cells were grown as described in panel A and assayed for β-galactosidase activity. (E) Co-IP analysis of Rgt1-HA and LexA-Ssn6 in mth1Δ and mth1Δ tpk1w1. Yeast cell extracts were immunoprecipitated with anti-LexA antibody and followed by immunoblotting with anti-HA antibody. Expression of Rgt1-HA was analyzed by Western blot using anti-HA antibody (Input). (F) Co-IP analysis of Rgt1-S5A-HA and LexA-Ssn6. Yeast cells (WT and mth1Δ) expressing Rgt1-S5A-HA (JKP234, Rgt1-S5A) and LexA-Ssn6 (JKP173) were grown as described in panel A. Yeast cell extracts were immunoprecipitated with anti-LexA antibody and followed by immunoblotting with anti- HA antibody. Expression of Rgt1-S5A-HA was analyzed by Western blot using anti-HA antibody (input). a) Results of the Ssn6 interaction with wild-type Rgt1 protein depicted in Figure 15B were shown for comparison.

Figure 18. Ssn6-Tup1 is recruited to the HXT1 promoter in an Rgt1-dependent manner. (A) Yeast cells (WT, rgt1Δ and mth1Δ) were grown in YP-2% Galactose medium (-) and shifted to YP-4% glucose medium (+) for 1 hr. Cross-linked chromatin was immunoprecipitated with anti-Ssn6 antibody, and representative PCRs are shown (left and middle panels). Input or immunoprecipitated DNA was PCR amplified with primers specific to pHXT1 or pACT1 as a negative control. Western blot analysis of Ssn6 expression using anti-Ssn6 antibody (right panel). (B) qPCR analysis of Ssn6-binding to the HXT1 promoter in yeast cells. The results were expressed as IP/Input ratio as described in Figure 13B using the pACT1 promoter as a negative control. (C) A proposed model of the role of Mth1 in Rgt1-mediated repression. In the absence of glucose, Ssn6- Tup1 interferes with Rgt1 DNA-binding, but is antagonized by Mth1. Mth1 mediates the interaction between Rgt1 and Ssn6-Tup1 by blocking the PKA-dependent phosphorylation of Rgt1 that impairs the ability of Rgt1 to associate with Ssn6-Tup1 and with its target DNA sites. Therefore, Mth1 acts as a scaffold-like protein to recruit Ssn6- Tup1 to Rgt1. This complex is disrupted upon glucose-induced proteasomal degradation of Mth1 via the Rgt2/Snf3 pathway and consequent phosphorylation of Rgt1 by PKA. Expression of MTH1 gene is also repressed by the Snf1-Mig1 pathway. Phosphorylated Rgt1 is dissociated from Ssn6-Tup1 and eventually from DNA, leading to derepression of Rgt1 target genes.

Figure 19. Repression of the MTH1 gene by the Mig1-Ssn6-Tup1 complex. Yeast cells carrying the MTH1-LacZ reporter plasmid were first grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr and assayed for β- galactosidase activity.

Figure 20. Rgt1-S5A-HA constitutively represses expression of the HXT1 gene. Yeast cell (rgt1Δ) expressing Rgt1-S5A-HA was grown in SC-2% galactose medium (-) and shifted to SC-4% glucose medium (+) for 1 hr and assayed for β-galactosidase activity.

Figure 21. Ssn6 overexpression abrogates the DNA-binding ability of Rgt1, but not of Mig1. Yeast cells (BY4741, WT) cooverexpressing Ssn6-HA with Rgt1-HA were grown in SC-2% galactose (-) and shifted to SC-4% glucose (+) for 1 hr. Cross-linked chromatin was precipitated for ChIP analysis of the indicated Rgt1 (pHXT1, pHXT2, and pHXT3) (A) and Mig1 (pSUC2 and pGAL1) (B) DNA target sites. Representative PCRs of ChIP analysis were shown (upper panels). The amount of immunoprecipitated DNA was quantified by qRT-PCR with primer pairs directed against the HXT1, HXT2, HXT3, SUC2, and GAL1 promoters. As a control, primer sets were also used for Actin promoter (pACT1). qPCR analysis of the DNA-binding of Rgt1 and Mig1 was expressed as IP/Input ratio which was determined by the ratio of immunoprecipitated (IP) pHXT relative to the IP pACT1 divided by the ratio of INPUT pHXT relative to the INPUT pACT1 (bottom panel). The data shown were averages of three independent experiments with error bars showing mean ± S.D.

Table 2. S. cerevisiae strains used in this study

Strain Genotype Source BY4741 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ (Kaniak et al., 2004) BY4743 Mata/α his3Δ1/ his3Δ1 leu2Δ0/ leu2Δ0 ura3Δ0/ (Kaniak et al., ura3Δ0 met15Δ0/MET15 lys2Δ0/LYS2 2004) FM557 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 (Kaniak et al., rgt1::kanMX 2004) FM645 Mata his3Δ1 leu2Δ0 ura3Δ0 trp1 ade8 tpk1w1 (Toda et al., tpk2::HIS3 tpk3::TRP1 bcy1::URA3 1987) YM6266 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 (Kim et al., mth1::kanMX 2006) YM6545 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 (Kaniak et al., RGT2-1 2004) YM6684 Mata/α his3Δ1/his3Δ1 leu2Δ0/leu2Δ0 (Kaniak et al., ura3Δ0/ura3Δ0 met15Δ0/MET15 lys2Δ0/LYS2 2004) mig1::kanMX/mig1::kanMX mig2::kanMX/mig2::kanMX mig3::kanMX/mig3::kanMX JKY 32 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study mth1::kanMX2 std1::NAT JKY 66 Mata his3Δ1 leu2Δ0 ura3Δ0 trp1 ade8 tpk1w1 This study tpk2::HIS3 tpk3::TRP1 bcy1::URA3 mth1::NAT JKY 83 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study RGT2-1 ssn6::NAT JKY 87 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study mth1::kanMX std1::kanMX ssn6::NAT JKY 88 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study pHXT1-NAT JKY 89 Mata his3Δ1 leu2Δ0 ura3Δ0 m et15Δ LYS2 This study pHXT2-NAT JKY 90 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study pHXT3-NAT JKY 91 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 This study mth1::kanMX std1::kanMX tup1::NAT JKY 93 Mata ura3-52 his3-11 leu2::kanMX6 15MAL2 This study SUC2 GAL MET HTR1-23

CHAPTER IV

Endocytosis and vacuolar degradation of the yeast cell surface glucose sensors Rgt2 and Snf3

Abstract

Sensing and signaling the presence of extracellular glucose is crucial for the yeast

Saccharomyces cerevisiae because of its fermentative metabolism, characterized by high

glucose flux through glycolysis. The yeast senses glucose through the cell surface

glucose sensors Rgt2 and Snf3, which serve as glucose receptors that generate the signal

for induction of genes involved in glucose uptake and metabolism. Rgt2 and Snf3 detect

high and low glucose concentrations, respectively, perhaps due to their different affinities for glucose. Here, we provide evidence that cell surface levels of glucose sensors are regulated by ubiquitination and degradation in the vacuole. The glucose sensors are removed from the plasma membrane through endocytosis and targeted to the vacuole for

degradation upon glucose depletion. The turnover of the glucose sensor is inhibited in

endocytosis defective mutants, and the sensor proteins with a mutation at their putative

ubiquitin-acceptor lysine residues are resistant to degradation. Of note, the low affinity

glucose sensor Rgt2 remains stable only in high-glucose grown cells, and the high

affinity glucose sensor Snf3, only in cells grown in low glucose. Besides, constitutively

active, signaling forms of glucose sensors do not undergo endocytosis, whereas signaling

defective sensors are constitutively targeted for degradation, suggesting that the stability

of the glucose sensors may be associated with their ability to sense glucose and that the

amount of glucose available dictates the cell surface levels of the glucose sensors. In this

manner, yeast cells may maintain glucose sensing activity at the cell surface over a wide

68 range of glucose concentrations, enabling them to respond rapidly to changing glucose levels.

4.1. Introduction

Most organisms have evolved numerous mechanisms for sensing and signaling the availability of glucose, the universal fuel for life, ensuring its optimal utilization (Rolland et al., 2001a; Towle, 2005). Glucose is by far the preferred energy source of the budding yeast S. cerevisiae, because regulation of cellular function by glucose dictates the organism’s fermentative lifestyle (Holsbeeks et al., 2004; Ozcan and Johnston, 1999).

The propensity of the yeast to ferment rather than oxidize glucose demands high glycolytic flux, and, therefore, yeast cells consume the available glucose vigorously by increasing glucose uptake through glucose transporters (HXTs) (Johnston and Kim, 2005;

Ozcan and Johnston, 1999).

Expression of the HXT genes is repressed in the absence of glucose by a multi-protein repressor complex, composed of the HXT gene repressor Rgt1, the general corepressor

Ssn6-Tup1 and the glucose responsive transcription factor Mth1 (Kim, 2004, 2009; Kim et al., 2003; Ozcan and Johnston, 1996; Schmidt et al., 1999). Mth1 blocks PKA (cAMP- activated protein kinase A) phosphorylation of the Rgt1 repressor, enabling it to recruit

Ssn6-Tup1 to the HXT promoters (Kim and Johnston, 2006; Mosley et al., 2003;

Palomino et al., 2006). Addition of glucose to glucose-depleted cells induces degradation of Mth1 (Flick et al., 2003; Kim et al., 2006; Moriya and Johnston, 2004; Pasula et al.,

2010; Spielewoy et al., 2004) and consequent phosphorylation of Rgt1 by PKA, leading to Rgt1 dissociation from DNA and thus to HXT gene expression (Kim et al., 2006;

Palomino et al., 2006). Hence, multiple mechanisms are involved for fine-tuned regulation of HXT gene expression (Kim et al., 2013)

The signal that leads to proteasomal degradation of Mth1 is generated by the two cell

surface glucose sensors Rgt2 and Snf3 (Johnston and Kim, 2005). The glucose sensors

are evolutionarily derived from glucose transporters but appear to have lost the ability to

transport glucose into the cell; instead, they function as glucose receptors (Ozcan et al.,

1998; Ozcan et al., 1996a). This view is strongly supported by the identification of a dominant mutation in the glucose sensor genes (RGT2-1 and SNF3-1), which is thought to convert the sensors into the glucose-bound and therefore glucose signaling forms

(Ozcan et al., 1996a). Indeed, Mth1 degradation and subsequent HXT gene expression occur constitutively in Rgt2-1 and Snf3-1 mutant cells (Pasula et al., 2007). These observations have led to the view that glucose acts like a hormone to initiate receptor- mediated signaling, and glucose sensors function in a similar way as mammalian cell surface receptors (Busti et al., 2010; Johnston and Kim, 2005).

The yeast cells possess multiple glucose transporters with different affinities for glucose, enabling them to grow well over a wide range of glucose concentrations, from a few micromolar to a few molar (Ozcan and Johnston, 1999). They sense extracellular glucose levels through the two glucose sensors, which have different affinities for glucose. Rgt2 has a low affinity and Snf3, a high affinity, for glucose (Ozcan et al.,

1998). This difference is presumably due to differences in the amino acid residues of the sensors that form the glucose-binding site. Thus, it has been proposed that Rgt2 functions as a low affinity glucose receptor that senses high concentrations of glucose, whereas

Snf3 serves as a high affinity glucose receptor that senses low levels of glucose (Ozcan et

70 al., 1998; Ozcan et al., 1996a). However, it remains unknown whether the abundance and function of cell surface levels of the glucose sensors are associated with their affinity for glucose and thus affect glucose signaling.

Here, I provide evidence that cell surface levels of glucose sensors are regulated by ubiquitination and degradation in the vacuole. Our results indicate that the stability of glucose sensors are correlated with their affinity for glucose and that constitutively active, signaling form of glucose sensor mutants are stable against degradation. These observations suggest that conformation of the glucose sensors is critical for their stability.

I discuss the biological significance of this observation in the perspective of the fermentative metabolism of yeast, characterized by high glucose uptake, increased glycolytic activity.

4.2. Materials and Methods

4.2.1. Yeast Strains

The Saccharomyces cerevisiae strains used in this study are listed in Table 3.1. Cells were grown in YP (2% bacto-peptone, 1% yeast extract) and SC (synthetic yeast nitrogen base medium containing 0.17% yeast nitrogen base and 0.5% ammonium sulfate) media supplemented with the appropriate amino acids and carbon sources.

4.2.2. Plasmid Construction

The plasmids used in this study are listed in Table 3.2. The plasmids were constructed using standard molecular biology techniques as described below. Plasmids containing

Rgt2-HA, Rgt2 (1-545)-HA, Rgt2 (1-620)-HA, Rgt2 (1-720)-HA and Rgt2-1-HA under

its endogenous promoter (1000 base pairs) were constructed in two steps. First, the

promoter element was PCR amplified from genomic DNA isolated from wild type yeast

strain as EcoRI-BamHI fragment and cloned in the empty HA vector (KFP 69, C- terminal 3xHA fusion vector). Next, the RGT2 ORFs were fused as BamHI-XbaI fragments after the promoter region. Plasmids containing Rgt2K637A-HA, Rgt2K657A-HA,

Rgt2K637,657A-HA, Rgt2W529F-HA and Rgt2W529Y-HA were mutagenized by QuikChange

Site-Directed Mutagenesis Kit (Stratagene) according to manufacturer’s protocol.

Plasmids containing GFP-Rgt2 and GFP-Rgt2-1 were constructed by fusing RGT2 and

RGT2-1 ORFs as BamHI-XhoI fragment in pUG34 vector. Plasmids containing Snf3-HA

and Snf3-1-HA under its endogenous promoter (1000 base pairs) were constructed in two

steps. First, the promoter element was PCR amplified from genomic DNA isolated from

wild type yeast strain as SacI-XbaI fragment cloned in the empty HA vector (KFP69).

Then, SNF3 and SNF3-1 ORFs were fused as XbaI-SphI fragments. Plasmids containing

GFP-Snf3 and GFP-Snf3-1 were constructed by ‘gap repair’ of BamHI-EcoRI linearized

pUG34 vector.

4.2.3. Yeast Membrane Preparation and Western Blotting

Membrane enriched fractions were essentially prepared as described previously

(Galan et al., 1996). Briefly, after washing with phosphate buffer, pH 7.4 containing 10 mM sodium azide, the cell pellet was resuspended in ice cold membrane isolation buffer

(100 mM Tris-Cl, pH 8, 150 mM NaCl, 5 mM EDTA) containing 10 mM sodium azide, protease and phosphatase inhibitors and vortexed with acid-washed glass beads. After

diluting the samples with the same buffer, unbroken cells and debris were removed by

centrifugation and membrane enriched fraction was collected by centrifuging the samples

at 12,000 rpm for 40 min at 4ºC. The pellets were resuspended in the aforementioned

buffer containing 5M urea and incubated for 30 min on ice and further centrifuged at

12,000 rpm for 40 min at 4ºC. The proteins were precipitated with 10% TCA, neutralized

with 20 µl of 1M Tris base and finally dissolved in 80 µl of SDS buffer (50 mM Tris-

HCl, pH, 6.8, 10% glycerol, 2% SDS, 5% β-mercaptoethanol).

For Western blotting, proteins were resolved in 10% SDS-PAGE, transferred to PVDF

membrane (Millipore) and the membranes were incubated with appropriate antibodies

(anti-HA, anti-Myc, anti-GFP or anti-actin antibody, Santa Cruz) in TBST buffer (10 mM

Tris-HCl, pH, 7.5, 150 mM NaCl, 0.1% Tween-20) and proteins were detected by the

enhanced chemiluminescence (ECL) system (Pierce).

4.2.4. Quantitative Real Time PCR (qRT-PCR)

Total RNA was extracted by RNeasy mini kit (Qiagen) following manufacturer’s

protocol and 2µg of total RNA was converted to cDNA by qScript cDNA supermix

(Quanta Biosciences). cDNA was analyzed by qRT-PCR using SsoFast Evagreen reagent

(Bio-Rad) in CFX96 Real-time thermal cycler (Bio-Rad). ACT1 was used as an internal control to normalize expression of HXT1, RGT2 and SNF3 genes. All of the shown quantification data were the averages of three independent experiments with error bars representing standard deviations (S.D). Statistical significance was defined by P-values:

*P<0.05, **P<0.001 as compared with control.

4.2.5. Microscopy and Image Analysis

To visualize yeast cells expressing various GFP fusion proteins, cells were stained with

FM4-64 (lipophilic styryl dye for selectively staining vacuolar membrane, 5mg/ml stock in DMSO) and examined with Olympus FluoView confocal microscope under 63X oil immersion objective lens using GFP or Texas Red filters. Images from confocal microscope were captured by FluoView software (Olympus) and NIH ImageJ v1.4r software was used to quantify fluorescence intensities from un-manipulated raw images.

Regions of interest on plasma or vacuolar membrane and in an area outside the cell

(background) were traced using the free-hand tool and mean fluorescence intensities

(both GFP and FM4-64) were measured. After background subtraction, the GFP signals in the plasma membranes were normalized to the FM4-64 signal of vacuolar membrane.

At least 200 cells were counted and the data represented were the averages with error bars representing standard deviation (S.D).

4.3. Results

4.3.1. Glucose starvation induces endocytosis and vacuolar degradation of Rgt2

To test the hypothesis that the cell surface levels of Rgt2 glucose sensor may be regulated by glucose concentration, I determined its expression levels in yeast cells grown in different glucose concentrations. Western blot analysis showed that the cell surface levels of Rgt2-HA are greater in high glucose-grown cells (2%) than in cells grown in low glucose medium (~ 0.1%) and are very low in cells grown in the absence of

glucose (Galan et al.) (Figure 22A). However, RGT2 mRNA levels were not significantly different between yeast cells incubated with different concentrations of glucose (Figure 22B), and the treatment of the protein synthesis inhibitor cycloheximide did not greatly affect Rgt2 turnover (Figure 22C).

Figure 22. Rgt2 is degraded in the absence of glucose. (A) Western blot analysis of Rgt2-HA levels at the plasma membrane. Yeast cells (WT) expressing Rgt2-HA were grown in SC-2% glucose medium till mid log phase (O.D600nm=1.2- 1.5) and equal amounts of cells were shifted to SC medium containing different concentrations of glucose (0-2%) for 30 min. Membrane fractions were analyzed using anti-HA antibody. (B) qRT-PCR analysis of mRNA expression of RGT2 (mRNA) in yeast cells grown as described in Figure A (mRNA) and densitometric quantification of the intensity of each band on the blot in A (Protein). C. Yeast cells (WT) expressing Rgt2-HA were grown in SC-2% glucose (+) medium till mid log phase and shifted to 2% galactose (-) medium with or without cycloheximide (CHX, 50 µg/ml) for times as indicated. Membrane fractions were immunoblotted with anti-HA antibody (top panel), and the intensity of each band on the blot was quantified by densitometric scanning (bottom panel).

There are many proteins that are degraded via proteasome, such as Mth1, Std1 (Kim

et al., 2006), Rpn4 (Wang et al., 2010). I first asked if glucose starvation-induced downregulation of Rgt2 is proteasome-dependent. Rgt2-HA was expressed in yeast cells

(pdr5Δ) and the cells were treated with proteasome inhibitor drug MG132, in the absence or presence of glucose (Figure 23A). I also expressed Rgt2-HA in temperature sensitive strain pre2-2 where function of proteasomal activity is diminished (Figure 23B). Rgt2 was shown to be degraded in the absence of glucose in either cases implying the fact that downregulation of Rgt2 is proteasome-independent phenomenon.

Figure 23. Glucose starvation-induced degradation of Rgt2 is not dependent on proteasome. (A) Yeast cells (pdr5∆) coexpressing Rgt2-HA and Mth1-Myc were first grown in 2% glucose (+) medium till mid log phase and shifted to 2% galactose (-) or 2% glucose medium with or without proteasome inhibitor drug MG132 (50 µg/ml) for 30 min. Membrane and soluble fractions were analyzed by immunoblotting using anti-HA or anti-Myc antibody. (B) Yeast cells (WT and pre2-2ts) expressing Rgt2-HA were first grown in 2% glucose (+) medium till mid log phase and shifted to 2% galactose (-) medium for 30 min. membrane fractions were analyzed by immunoblotting using anti- HA antibody. Actin was served as loading control in A and B.

Because a number of yeast plasma membrane receptors and transporters are

downregulated by endocytosis and degradation in the vacuole (Kriel et al., 2011;

Rutherford and Bird, 2004), I examined expression levels of Rgt2-HA in the end3Δ

mutant defective in the internalization step of endocytosis and the pep4Δ mutant

defective in vacuolar protease processing. Rgt2-HA levels in glucose-grown wild type

cells were reduced by ~ 50% within 20 min after the cells were shifted to glucose-

depleted (galactose) medium, but this reduction was not observed in the end3Δ and pep4Δ

strains (Figure 24A). Consistently, the amounts of immunodetected Rgt2-HA was

markedly increased within 30 min after addition of glucose to glucose-starved medium

(Figure 24B).

Confocal microscopy demonstrated that GFP-Rgt2 is present at the cell surface in glucose-grown cells and that ~80% of GFP-Rgt2 is removed from there when the yeast cells are shifted from glucose to galactose medium (Figure 24D, WT). However, GFP-

Rgt2 was constitutively detected at the cell surface of the end3Δ mutant (Figure 24D) and

the pep4Δ mutant (Data not shown). It was also shown that substantial amounts of GFP-

Rgt2 were localized to the vacuole in a glucose-independent manner, suggesting

constitutive internalization and degradation of Rgt2 (Figure 24D, FM4-64). Glucose and

galactose only differ with respect to C-4, yet galactose does not activate the glucose

sensors, suggesting that the glucose sensors display remarkable substrate specificity (27).

Consistently, we found that Rgt2-HA levels are downregulated in the cells grown on

galactose, raffinose, or ethanol (Figure 24C). These data indicate that Rgt2 is stable

against degradation in the presence of high concentrations of glucose but endocytosed

and degraded in the vacuole when glucose is absent or present only in small quantities.

4.3.2. Snf3 expression is regulated at both transcriptional and posttranslational

levels

The pattern of the glucose regulation of Snf3 expression appeared to be opposite to

that of Rgt2: the amounts of Snf3-HA at the plasma membrane were greater in low glucose-grown cells than in cells grown in high glucose medium (Figure 25A).

Expression of the SNF3 gene was shown to be repressed by high concentrations of glucose (Figure 25B), as we observed previously (Kaniak et al., 2004). To examine whether Snf3 levels are

Figure 24: Rgt2 undergoes endocytosis and subsequent vacuolar degradation in glucose starved cells. (A) Yeast cells (WT, end3∆ and pep4∆) expressing Rgt2-HA were grown without cycloheximide as described in Figure 22C. Yeast cells were harvested at

different time points as indicated, and membrane fractions were immunoblotted with anti- HA antibody (top panel) and the intensity of each band on the blot was quantified by densitometric scanning (bottom panel). (B) Yeast cells (WT, end3∆ and pep4∆) expressing Rgt2-HA were grown in SC-2% glucose medium (+) till mid log phase and shifted to SC-2% galactose medium (-) for 30 min and again shifted to SC-2% glucose medium for 30 min. Membrane fractions were immunoblotted with anti-HA antibody. (C) Yeast cells (WT) expressing Rgt2-HA were grown in SC-2% glucose (Glu) medium till mid log phase and shifted to SC medium containing either 2% galactose (Gal), 2% raffinose (Raf) or 2% ethanol (EtOH) and incubated for 30 min. Membrane fractions were immunoblotted with anti-HA antibody (top panel). qRT-PCR analysis of mRNA expression of RGT2 (mRNA) and densitometric quantification of the intensity of each band on the blot (Protein) (bottom panel). (D) GFP-Rgt2 was expressed from the MET25 promoter in wild type and end3∆ strains. Yeast cells expressing GFP-Rgt2 were grown as described in Figure 23B. Confocal microscope images (top panel) and quantification of relative GFP fluorescence in the plasma membrane (bottom panel, **P < 0.001) were shown. Relative GFP fluorescence intensities were plotted with the fluorescence of WT cells (2% glucose condition) set to 100%. The data represented were averages of at least 50 cell counts with error bars representing standard deviations (S.D). Actin was served as loading controls in A, B and C.

regulated at a posttranslational level, we expressed GFP-Snf3 under the control of the

MET25 promoter, which is not regulated by glucose (Kim et al., 2006; Pasula et al.,

2007). The cell surface levels of GFP-Snf3 were greater in low glucose grown cells (~

0.1%) than in cells grown in the absence of glucose (Galan et al.) or in high glucose-

grown cells; however, the glucose regulation of Snf3 expression levels was not observed

in the end3Δ mutant (Figure 25C).

The transcriptional and posttranslational regulation of Snf3 expression was

recapitulated in cells expressing Snf3-HA (expressed from the SNF3 promoter) and GFP-

Snf3 (expressed from the MET25 promoter). Both Snf3-HA and GFP-Snf3 levels were

very low in glucose-grown cells and high in cells grown on raffinose (equivalent to low

glucose). But, GFP-Snf3, unlike Snf3-HA, was detected at low levels in galactose or

ethanol-grown cells (Figure 25D). Consistent with these observations, the plasma

membrane localization of GFP-Snf3 was observed only in raffinose-grown cells but was

FIGURE 25. Snf3 levels are regulated by both transcriptional and translational mechanisms. (A). Western blot analysis of the plasma membrane levels of Snf3-HA. Yeast cells (WT) expressing Snf3-HA were grown as described in Figure 22A and membrane fractions were immunoblotted with anti-HA antibody. (B) qRT-PCR analysis of mRNA expression of SNF3 (mRNA) in yeast cells grown as described in Figure 22A (mRNA) and densitometric quantification of the intensity of each band on the blot in A (Protein). (C) GFP-Snf3 was expressed from the MET25 promoter in wild type and end3∆ strains. Yeast cells expressing GFP-Snf3 were grown as described in Figure 22A, and membrane fractions were immunoblotted with anti-HA antibody (top panel). The intensity of each band on the blot was quantified by densitometric scanning (bottom panel, *P < 0.05, **P < 0.001). D. Western blot analysis of Snf3-HA and GFP-Snf3 levels at the plasma membrane. Yeast cells (WT) expressing Snf3-HA or GFP-Snf3 were grown as described in Figure 24C. GFP-Snf3 was expressed from the MET25 promoter. Membrane fractions were immunoblotted with anti-HA antibody (top panel), and the intensity of each band on the blot was quantified by densitometric scanning (bottom panel, *P < 0.05, **P < 0.001). (E) GFP-Snf3 was expressed from the MET25 promoter

80 in wild type and end3∆ strains in glucose (High), raffinose (Low) or galactose (No) medium. Confocal microscope images (top panel) and quantification of relative GFP fluorescence in the plasma membrane (bottom panel, **P < 0.001) were shown. Actin was served as a loading control in A, C and D.

constitutively detected, regardless of carbon sources, in the mutant cells lacking the

END3 gene (Figure 25E). These results suggested that Snf3 is internalized and degraded in the vacuole in the absence of glucose or when glucose levels are high. Hence, Snf3 levels are regulated by both feedforward and feedback mechanisms: Snf3 degradation is reinforced by glucose repression of SNF3 gene expression but is dampened by derepression of the gene in the absence of glucose. Taken together, these results indicate that the low affinity glucose sensor Rgt2 is stable in the presence of high glucose concentrations and the high affinity glucose sensor Snf3, in low glucose conditions, and thus suggest that stability of the glucose sensors is associated with their ability to sense glucose.

4.3.3. Rgt2 degradation is ubiquitin-dependent.

Ubiquitination is a signal for endocytosis of plasma membrane proteins (Hicke and

Dunn, 2003; Medintz et al., 1998). The Doa4 ubiquitin isopeptidase and the Rsp5 ubiquitin ligase are known to be involved in the ubiquitination of many plasma membrane receptors and transporters in yeast (Amerik et al., 2000; Gadura and Michels,

2006; Rotin et al., 2000; Springael and Andre, 1998). To determine whether Rgt2 downregulation is mediated by ubiquitination, I investigated glucose regulation of Rgt2 in the strain carrying the doa4Δ or rsp5-1ts mutation (Liu et al., 2007). Rgt2-HA levels were constitutively high in both the doa4Δ mutant (Figure 26A, left) and in the rsp5-1ts

FIGURE 26. Ubiquitination of the cytoplasmic tail domain of Rgt2 is required for its endocytosis. (A) Western blot analysis of Rgt2-HA levels at the plasma membrane. Yeast cells (WT, doa4∆ and rsp5-1) expressing Rgt2-HA were grown as described in Figure 23B. Membrane fractions were immunoblotted with anti-HA antibody. (B) GFP- Rgt2 was expressed from the MET25 promoter in wild type, doa4∆ and rsp5-1 strains). Yeast cells expressing GFP-Rgt2 were grown in glucose (+) or galactose (-) medium, as described in Figure 23B. Confocal microscope images (left panel) and quantification of relative GFP fluorescence in the plasma membrane (right panel, *P < 0.05, **P < 0.001) were shown. (C) Schematic maps of Rgt2 constructs (WT, 1-545 aa, 1-620 aa or 1-720 aa) showing lysine residues at N and C-terminal domains. The K637 and K657 residues are shown as putative ubiquitin acceptor sites. (D) and (E). Yeast cells (WT) expressing indicated Rgt2-HA constructs were grown as described in Figure 23B, and membrane fractions were immunoblotted with anti-HA antibody.

mutant incubated at 37º C (Figure 26A, right), compared with those in wild type cells.

Consistently, GFP-Rgt2 was shown to remain stable at the plasma membrane in those

mutants (Figure 26B). To identify the ubiquitination sites in Rgt2, I constructed a series

of deletion mutants of Rgt2 and used them to map the regions that are important for its

stability (Figure 26C). Rgt2 degradation is abolished by the deletion of the entire C-

terminal cytoplasmic domain (1-545) or significantly inhibited by the deletion of the last

143 amino acids (1-620) (Figure 26D). However, the deletion of the last 13 amino acids

of Rgt2 (1-720) did not affect its stability, implicating that the 100 amino acids between residues 620 and 720 that contain the two lysine residues, K637 and K657, may be necessary for Rgt2 ubiquitination. Indeed, substitution of the two lysine residues by alanine (K637A and K657A) markedly increased Rgt2 stability in glucose-starved cells, suggesting that the two lysine residues may serve as major ubiquitination sites (Figure

26E).

Endocytosis-mediated degradation of Snf3 is dampened by glucose regulation of the expression of the SNF3 gene, suggesting that Snf3 levels are mainly regulated by

transcriptional control. For this reason, ubiquitination of Snf3 was not thoroughly

examined in this study.

4.3.4. Constitutively active glucose sensors are stable against degradation

There are dominant mutations in the glucose sensor genes (RGT2-1 and SNF3-1) that lock the sensor proteins into a glucose-bound conformation and cause constitutive, glucose-independent expression of HXT genes (Ozcan et al., 1998). (Figure 27A). I examined the stability of the active forms of the glucose sensors by Western blotting and found that, compared with wild type glucose sensors, both Rgt2-1 and Snf3-1 sensors are stable in both the presence and the absence of glucose (Figures 27B and 27C). It was noted again that the low levels of the Snf3-1-HA, compared with those of GFP-Snf3-1, in glucose-grown cells are probably due to the glucose repression of SNF3-1 gene

expression (Figure 27C, Glu). We also examined whether the degradation resistant

glucose sensor mutants (Rgt2-1 and Snf3-1) can generate a signal even in the absence of

glucose that leads to constitutive expression of HXT genes. To this end, Rgt2-1 was

expressed in the HXT1-NAT reporter strain in which the NAT (nourseothricin) resistance

gene is expressed under the control of the HXT1 promoter (Roy et al., 2013b). Colony

assays showed that expression of Rgt2-1 sensor enables the yeast cells to grow in

medium lacking glucose (Figure 27D). Similar results were obtained in the expression of

Snf3-1 sensor in the HXT2-NAT reporter strain in which expression of the HXT2 gene is

induced by low glucose (Figure 27E, raffinose). I found that glucose repression of SNF3-

1-HA expression may be responsible for the poor growth of the HXT2-NAT reporter strain

in glucose medium (Figure 27E, Snf3-1-HA vs. GFP-Snf3-1). It was also noted that the

HXT2-NAT reporter stain expressing Snf3-HA is able to grow in glucose-depleted medium (Figure 27E, Gal + NAT), mostly due to glucose depletion-induced derepression of Snf3-HA (Figure 27C, Gal). These results are consistent with Figure 25 and support the view that glucose depletion-induced Snf3 degradation is attenuated by the derepression of

SNF3 gene expression. Consistently, confocal microscopy demonstrated that Rgt2-1 and

Snf3-1 glucose sensors, compared with wild type Rgt2 and Snf3 sensors, remain stable at the plasma membrane, regardless of glucose concentration (Figure 27F). These results suggested that conformation of the glucose sensors may be critical for their stability.

4.3.5. Signaling defective Rgt2 mutant is constitutively targeted for vacuolar

degradation

To corroborate our hypothesis that glucose sensors may be stable in their glucose-bound,

signaling state, we examined the stability of signaling defective glucose sensors against

degradation. The yeast galactose transporter Gal2 can recognize both galactose and

glucose, and F504 of Gal2, which corresponds to W529 of Rgt2, is critical for substrate

recognition (Kasahara and Kasahara, 2000). I replaced Trp at position 529 with aromatic

amino acids Phe and Tyr using site-directed mutagenesis and determined the stability of

the resulting Rgt2 mutants Rgt2W529F and Rgt2W529Y in high glucose-grown cells. The

results showed that, in contrast to wild type Rgt2, the mutant Rgt2 sensors, Rgt2W529Y in

particular, was endocytosed and degraded even in the presence of glucose (Figure 28A),

leading to inhibition of the glucose induction of HXT1 gene expression (Figure 28B).

Thus, Rgt2W529Y was not able to complement the growth defect of the rgt2snf3 double mutant in glucose medium (Figure 28C). High glucose-induced proteasomal degradation of Mth1 is triggered by glucose activation of the Rgt2 sensor (Flick et al., 2003; Moriya

and Johnston, 2004). Western blot analysis showed that glucose-dependent Mth1 degradation occurs in cells expressing the wild type Rgt2 sensor but not the Rgt2W529Y

sensor (Figure 28D). These observations support the view that the stability of the glucose

sensors may be determined by their ability to sense glucose.

4.4. Discussion

Many yeast nutrient receptors and transporters, such as Zrt1 (Gitan et al., 1998), Ctr1

(Liu et al., 2007), Fth1 (Stearman et al., 1996), Smf1 (Reddi et al., 2009), Fur4 (Galan et

al., 1996) and Gap1 (Springael and Andre, 1998), are regulated by a homeostatic fashion.

They are induced in the absence of their ligands but internalized and targeted for

degradation in the vacuole when their ligands become available in excess (Kriel et al.,

2011; Rutherford and Bird, 2004). Hence, endocytic degradation of these plasma

FIGURE 27. Constitutively active Rgt2-1 and Snf3-1 glucose sensors do not undergo endocytosis. (A) A schematic diagram of the predicted secondary structure of the Rgt2 glucose sensor showing 12 transmembrane domains, cytoplasmic N- and C-terminal tails, two constitutive mutations (RGT2-1 (R231K) and SNF3-1 (R229K)) and two putative ubiquitin-acceptor lysine residues (K637 and K657). (B) and (C) Yeast cells (WT) expressing indicated Rgt2 proteins (B) or Snf3 proteins (C) were grown in glucose (High), raffinose (Low) or galactose (No) medium, and membrane fractions were immunoblotted with anti-HA or anti-GFP antibody. Actin was served as a loading control. GFP-fusions of glucose sensors (GFP-Rgt2, GFP-Rgt2-1, GFP-Snf3, and GFP- Snf3-1) were expressed from the MET25 promoter. (D) The PHXT1-NAT reporter strain

(JKY88) expressing indicated Rgt2 proteins was spotted on SC-2% glucose (Glu), SC- 2% raffinose (Raf) or SC-2% galactose (Gal) plates supplemented with 100 µg/ml NAT- sulfate. The first spot of each row represents a count of 5 x 107 cell/ml, which is diluted 1:10 for each spot thereafter. The glucose plates and the galactose and raffinose plates were incubated for 2 and 3 days, respectively. (E) The PHXT2-NAT reporter strain (JKY89) expressing indicated Snf3 proteins was spotted and photographed as described in D. (F) Yeast cells (WT) expressing GFP-Rgt2, GFP-Rgt2-1, GFP-Snf3 and GFP-Snf3-1 were grown as described above (B and C), were analyzed by confocal microscopy.

FIGURE 28. Signaling defective Rgt2 glucose sensor is constitutively endocytosed. (A) Yeast cells (WT and end3∆) expressing indicated Rgt2-HA proteins were grown as described in Figure 23B and membrane fractions were immunoblotted with anti-HA antibody (top panel). The intensity of each band on the blot was quantified by densitometric scanning (bottom panel, *P < 0.05, **P < 0.001). (B) Yeast cells (rgt2∆snf3∆) expressing indicated Rgt2-HA proteins were grown as described in Figure 28B and the mRNA levels of HXT1 were quantified by qRT-PCR. The values shown are mean ± SD (*P < 0.05, **P < 0.001). (C) Yeast cells (rgt2∆snf3∆) expressing indicated Rgt2-HA proteins were spotted on 2% glucose plate supplemented with Antimycin-A (1µg/ml) (2% Glu + AA) or SC-2% galactose plate (2% Gal) and photographed as described in Figure 27D. (D) Yeast cells (rgt2∆snf3∆) coexpressing Mth1-Myc and indicated Rgt2-HA proteins were grown as described in Figure 23B and cell lysates were immunoblotted with anti-Myc antibody (left panel, Mth1-Myc). Actin was served as a loading control (right panel, Actin). Quantification data of Mth1-Myc protein by densitometry are shown (bottom panel, *P < 0.05, **P < 0.001).

membrane proteins functions as a homeostatic regulatory loop to prevent excessive

ligand-induced activation of downstream effectors (Hanyaloglu and von Zastrow, 2008;

Sorkin and Goh, 2009). By contrast, we show here that the glucose sensors are subjected

to endocytosis and vacuolar degradation in the absence of their ligand glucose. Of note,

the stability of the Rgt2 and Snf3 glucose sensors at the plasma membrane is correlated

with their ability to sense glucose, leading to the view that the actively signaling state of

glucose sensors is protected from degradation.

This view is supported by the findings that the conformation of the glucose receptors

determines their stability (Figure 27). The RGT2-1 or SNF3-1 mutation has been postulated to lock the glucose sensor in a glucose-bound, signaling form; leading to the hypothesis that glucose-binding to the glucose sensors suffices to initiate signaling(Ozcan et al., 1998; Ozcan et al., 1996a; Wu et al., 2006). These constitutively active glucose sensor Rgt2-1 (Rgt2R231K) and Snf3-1 (Snf3R229K) do not undergo endocytosis and remain

stable on the cell surface regardless of the presence of glucose. I also identify an RGT2

mutation that converts Rgt2 sensor to a constitutively inactive form and show that this

signaling defective Rgt2 mutant (Rgt2W529Y) is constitutively targeted to the vacuole for

degradation (Figure 28). Glucose binding likely induces a series of structural changes in

glucose sensors and transporters. Glucose transporters may undergo a conformational change upon glucose binding from the outward-facing, signaling conformation to the inward-facing, nonsignaling conformation that allows glucose to be released inside the cell; in contrast, the glucose sensors cannot switch to the inward-facing conformation

(Van Zeebroeck et al., 2009). The nature of RGT2 W529Y mutation is not well understood, but we surmise that the Rgt2 mutant (Rgt2W529Y) may not be able to sense

glucose or to be converted into the outward-facing, signaling conformation after binding

to glucose.

The yeast cells cope with environmental changes in glucose availability by

expressing at least six members of the glucose transporter family with different affinities

for glucose (Boles and Hollenberg, 1997; Ko et al., 1993; Kruckeberg, 1996;

Reifenberger et al., 1995). They express only those glucose transporters most appropriate

for the amounts of glucose available in the environment (Ozcan and Johnston, 1995). The

glucose sensors have different roles in glucose signaling: the low affinity glucose sensor

Rgt2 is responsible for expression of the low affinity glucose transporter Hxt1; the high

affinity glucose sensor Snf3 regulates the expression of the high affinity glucose

transporters Hxt2, Hxt3 and Hxt4 (Ozcan et al., 1998). This is consistent with my

findings that Rgt2 remains stable only in high-glucose grown cells, and Snf3, only in

cells grown in low glucose, reinforcing the view that the stability of the glucose sensors is

correlated with their affinity for glucose. Moreover, the glucose sensors are localized to

the vacuole in both presence and absence of glucose (Figures 24D and 25E). These

observations suggest that the glucose sensors may be inherently unstable but stabilized by

glucose.

My findings provide a conceptual framework to explain the regulation of glucose sensing

activity at the yeast cell surface that directly affects the organism’s ability to adapt to

fluctuating glucose levels. Snf3 expression is regulated by both feedforward and feedback

mechanisms: Snf3 protein is internalized and degraded in glucose-depleted cells and cells grown on high glucose; expression of the SNF3 gene is repressed by Mig1 and Mig2

repressors in the presence of high glucose concentrations but is derepressed in in the absence of glucose. Thus, high glucose-induced Snf3 degradation is reinforced by glucose repression of SNF3 gene expression, whereas glucose depletion-induced Snf3 degradation is dampened by derepression of SNF3 gene expression. As a result, substantial amounts of Snf3 are present at the cell surface even in the absence of glucose

(Figure 25A). This should serve to provide for a rapid reestablishment of induction of

HXT gene expression when glucose is available in the medium. Consequently, one or the other, or both of the glucose sensors may be present at the plasma membrane at a given glucose concentration. Snf3 may be the predominant sensor in low levels of glucose and

Rgt2, in high glucose conditions. Both Rgt2 and Snf3 may coexist in an intermediate between high and low levels of glucose (Figure 29). In this manner, yeast cells can keep glucose sensing activity constant at the plasma membrane over a wide range of glucose concentrations, enabling them to respond rapidly and appropriately to changing glucose levels and thereby to enhance glucose uptake and utilization.

FIGURE 29. Glucose regulation of the yeast glucose sensors. (A) Cell surface levels of Rgt2-HA and Snf3-HA are compared (adapted from Figures 22A and 25A). Lanes 1-7 denote different concentrations of glucose (%w/v): 2, 1, 0.5, 0.25, 0.125, 0.05 and 0, respectively. (B) Regulation of yeast glucose sensors by glucose concentration. The turnover of the glucose sensors plays an important role in the adaptation to changing glucose levels. The low affinity glucose sensor Rgt2 is stable and functional in the presence of high levels of glucose and the high affinity glucose sensor Snf3, in low glucose conditions. In this manner, yeast cells may produce the glucose transporters most appropriate for the amount of glucose available in the environment.

Table 3.1. S. cerevisiae strains used in this study

Strain Genotype Source BY4741 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ (Kaniak et al., 2004) YM6870 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ rgt2::KanMX (Kaniak et al., snf3::KanMX 2004) KFY122 Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 doa4::KanMX This study KFY123 Mata his3-1 leu2-0 ura3-0 RSP5 (Liu et al., 2007) KFY124 Mata his3-1 leu2-0 ura3-0 rsp5-1/smm1 (Liu et al., 2007) KFY127 Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 end3::KanMX This study KFY128 Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 pep4::KanMX This study JKY88 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 pHXT1-NAT (Roy et al., 2013) JKY89 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ LYS2 pHXT2-NAT (Roy et al., 2013)

Table 3.2. Plasmids used in this study

Plasmid Description Source KFP69 pAD80, 3x-HA-CYC1 terminator, Leu2 (Kim, 2009) JKP253 pAD80-PRGT2-Rgt2-3xHA This study JKP252 pAD80-PRGT2-Rgt2 (1-545)-3xHA This study JKP299 pAD80-PRGT2-Rgt2 (1-620)-3xHA This study JKP300 pAD80-PRGT2-Rgt2 (1-720)-3xHA This study JKP301 pAD80-PRGT2-Rgt2 (K637A)-3xHA This study JKP302 pAD80-PRGT2-Rgt2 (K657A)-3xHA This study JKP308 pAD80-PRGT2-Rgt2 (K637A, K657A)-3xHA This study JKP303 pAD80-PRGT2-Rgt2 (W529Y)-3xHA This study JKP304 pAD80-PRGT2-Rgt2 (W529F)-3xHA This study JKP295 pAD80-PRGT2-Rgt2-1-3xHA This study JKP298 pAD80-PSNF3-Snf3-3xHA This study JKP311 pAD80-PSNF3-Snf3-1-3xHA This study pBM3842 pUG34, PMET25-GFP-CYC1 terminator, His3 (Kim et al., 2006) JKP293 pUG34-PMET25-GFP-Rgt2 This study JKP314 pUG34-PMET25-GFP-Rgt2-1 This study JKP309 pUG34-PMET25-GFP-Snf3 This study JKP310 pUG34-PMET25-GFP-Snf3-1 This study

CHAPTER V

Glucose starvation-induced turnover of the yeast glucose transporter Hxt1

Abstract

The budding yeast S. cerevisiae possesses multiple glucose transporters with different

affinities for glucose enabling it to deal with changes in glucose availability. The

stability of glucose transporters is regulated by glucose concentration. High affinity

glucose transporters such as Hxt2, Hxt6 and Hxt7 are endocytosed and degraded in the

vacuole in response to high glucose concentrations. Western blotting and confocal

microscopy were performed to evaluate glucose regulation of the low affinity glucose

transporter Hxt1. I found that Hxt1 is removed from the plasma membrane and targeted

to the vacuole for degradation when cells are shifted to the low-glucose or glucose-free

medium. However, the steady-state levels of Hxt1 are constitutively high in end3 and

doa4 mutants. My data also show that the two lysine residues of Hxt1, K12 and K39,

serve as the putative ubiquitin-acceptor sites by the Rsp5 ubiquitin ligase. The low affinity glucose transporter Hxt1 undergoes endocytosis and vacuolar degradation in response to glucose starvation. These findings, together with previous results of glucose-

induced degradation of high affinity glucose transporters, suggest that the stability of

glucose transporters may be closely correlated with their affinity for glucose.

5.1. Introduction

Glucose serves as the primary carbon and energy source of a multitude of cells,

varying in complexity from unicellular microorganisms to higher eukaryotes, and most organisms have developed mechanisms for sensing glucose and utilizing it efficiently

(Rolland et al., 2001a; Towle, 2005). Glucose is by far the preferred energy source of the budding yeast S. cerevisiae, because glucose regulation of cellular function dictates the organism’s distinctive fermentative lifestyle (Carlson, 1999; Gancedo, 1998). The yeast prefers to ferment rather than oxidize glucose even when oxygen is abundant (Crabtree,

1929; Lagunas, 1979). Because ATP production by fermentation is largely inefficient compared with respiration, the yeast cells consume the available glucose vigorously to meet cellular ATP demands and they do so by increasing glucose uptake through glucose transporters (HXTs) (Johnston and Kim, 2005; Lagunas, 1986; Ozcan and Johnston,

1999; Rolland et al., 2001b).

S. cerevisiae copes with changes in glucose availability by expressing at least six members of the glucose transporter family with different affinities for glucose (Hxt1, 2,

3, 4, 6, and 7) (Boles and Hollenberg, 1997; Reifenberger et al., 1997; Reifenberger et al.,

1995). The yeast cells detect extracellular glucose over a broad concentration range and

express only those glucose transporters best suited for the amount of glucose available in

the medium (Gancedo, 2008; Horak, 2013; Ozcan and Johnston, 1995; Rolland et al.,

2002): (1) Hxt1 is a low affinity glucose transporter with a Km value around 100 mM for

glucose and expressed when glucose levels are high (> ~1%), (2) The HXT2 and HXT4

genes encode glucose transporters with moderate affinity for glucose (Km values around

10 mM) and their expression is induced in the presence of low levels of glucose (~0.2%)

(Maier et al., 2002; Reifenberger et al., 1997), (Ma et al.) Hxt3 has a low affinity for

glucose (Km values around 30 - 60 mM) and induced by both low and high levels of

glucose (Boles and Hollenberg, 1997; Ozcan and Johnston, 1999; Reifenberger et al.,

1997) and (4) The HXT6 and HXT7 genes encode high affinity glucose transporters with

a Km value around 1 mM for glucose and their expression is induced by low

concentrations of glucose or by non-fermentable carbon sources such as glycerol or

ethanol (Diderich et al., 1999; Liang and Gaber, 1996; Ye et al., 2001).

Expression of the HXT genes is repressed by the Rgt1 repressor in the absence of glucose (Kim, 2009; Ozcan et al., 1996a). Rgt1 does so by recruiting the general

corepressor complex Ssn6-Tup1 to the HXT promoters in a manner that requires the HXT

corepressor Mth1 (Kim et al., 2003; Lakshmanan et al., 2003; Roy et al., 2013a; Schmidt

et al., 1999). High glucose induces expression of the HXT genes by inhibiting the

function of Mth1 and Rgt1. Mth1 mRNA and protein levels are downregulated by high

glucose via the Snf1 (AMPK)-Mig1 and Rgt2/Snf3 pathways, respectively (Flick et al.,

2003; Kaniak et al., 2004; Kim et al., 2006; Moriya and Johnston, 2004; Spielewoy et al.,

2004). Glucose-induced downregulation of Mth1 enables the phosphorylation of Rgt1 by

PKA (cAMP-PKA pathway), leading to dissociation of Rgt1 from DNA and thereby

expression of HXT genes (Jouandot et al., 2011; Kim and Johnston, 2006; Palomino et

al., 2006). Thus, three glucose signaling pathways converge at multiple points for fine-

tuned regulation of HXT gene expression (Kim et al., 2013).

The steady-state levels of the yeast glucose transporters are also regulated

posttranslationally. The high affinity glucose transporters such as Hxt2, Hxt6 and Hxt7

are internalized and targeted to the vacuole for degradation in cells grown in high glucose

medium by a process, known as catabolite degradation (Horak, 2003; Krampe et al.,

1998; Kruckeberg et al., 1999). Recently, it has been shown that the low affinity glucose

transporter Hxt3 is endocytosed and degraded in the vacuole when glucose-fed cells are

exposed to glucose-free medium (Snowdon and van der Merwe, 2012). These

observations lead to the view that the stability of glucose transporters may be regulated

by glucose concentration. In this study, we examined glucose regulation of the major low

affinity glucose transporter Hxt1 and provide evidence that Hxt1 undergoes ubiquitin-

linked endocytosis followed by degradation in the vacuole in response to glucose

starvation. My results, together with previous observations, show that, at high glucose

concentrations, the low affinity glucose transporters Hxt1 and Hxt3 localize to the plasma

membrane to function, whereas the high affinity glucose transporters Hxt2, Hxt6 and

Hxt7 are targeted to the vacuole for degradation. Therefore, glucose may not be directly

involved in the ubiquitination of glucose transporters, which in turn signals their

endocytosis and subsequent degradation in the vacuole, as suggested previously (Horak,

2003). Rather, the stability of the glucose transporters may be directly correlated with

their affinity for glucose.

5.2. Materials and Methods

5.2.1. Yeast strains and growth condition

The Saccharomyces cerevisiae strains used in this study are listed in Table 5.1. Cells were grown in YP (2% bacto-peptone, 1% yeast extract) and SC (synthetic yeast nitrogen base medium containing 0.17% yeast nitrogen base and 0.5% ammonium sulfate) media supplemented with the appropriate amino acids and carbon sources.

Table 5.1. S. cerevisiae strains used in this study

Strain Genotype Source BY4741 Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ (Spielewoy et al., 2004) KFY63 Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 bcy1::KanMX This study KFY101 Matα hxt1-17∆ gal2∆ agt1∆ stl1∆ his3∆1 ura3∆0 (Boles and leu2∆0 Hollenberg, 1997) KFY122 Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 doa4::KanMX This study KFY123 Mata his3-1 leu2-0 ura3-0 RSP5 (Liu et al., 2007) KFY124 Mata his3-1 leu2-0 ura3-0 rsp5-1/smm1 (Liu et al., 2007) KFY127 Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 end3::KanMX This study

5.2.2. Plasmid construction

The plasmids used in this study are listed in Table 5.2. Plasmids containing Hxt1-

GFP, Hxt1 (Q209A)-GFP, Hxt1 (Q335A)-GFP, Hxt1 (Q336A)-GFP, Hxt1 (S363A)-

GFP, Hxt1 (N370A)-GFP and Hxt1 (W473A)-GFP were constructed by gap repair, as described previously (Roy et al., 2013a).

Table 5.2. Plasmids used in this study

Name Description Source JKP315 Hxt1-GFP, Ura3, CEN This study JKP323 Hxt1 (60-570)-GFP, Ura3, CEN This study JKP324 Hxt1 (1-512)-GFP, Ura3, CEN This study JKP325 Hxt1 (60-512)-GFP, Ura3, CEN This study JKP332 Hxt1 (K12A)-GFP, Ura3, CEN This study JKP333 Hxt1 (K27A)- GFP, Ura3, CEN This study JKP334 Hxt1 (K35A)- GFP, Ura3, CEN This study JKP335 Hxt1 (K39A)- GFP, Ura3, CEN This study

5.2.3. Yeast membrane preparation, Western blotting and protein half-life measurement.

Membrane enriched fractions were essentially prepared as described previously

(Galan et al., 1996), with minor modifications. Briefly, after washing with 10 mM phosphate buffer at pH 7.4, the cell pellet was resuspended in membrane isolation buffer

(100 mM Tris-Cl, pH 8, 150 mM NaCl, 5 mM EDTA) containing 10 mM sodium azide, protease and phosphatase inhibitors and vortexed with acid-washed glass beads.

Membrane enriched fraction was collected by centrifugation at 12,000 rpm for 40 min at

4º C and suspended in the aforementioned buffer containing 5 M urea. The proteins were precipitated with 10% TCA, neutralized with 20 µl of 1 M Tris base and finally dissolved in 80 µl of SDS buffer (50 mM Tris-HCl, pH, 6.8, 10% glycerol, 2% SDS, 5% β- mercaptoethanol).

For Western blotting, proteins were resolved by SDS-PAGE (10%) and transferred to

PVDF (Polyvinylidene fluoride) membrane (Millipore). The membranes were incubated with appropriate antibodies (anti-HA, anti-GFP or anti-Actin antibody, Santa Cruz) in

TBST buffer (10 mM Tris-HCl, pH, 7.5, 150 mM NaCl, 0.1% Tween-20) and proteins were detected by the enhanced chemiluminescence (ECL) system (Pierce). The half-life of Hxt1-GFP was measured as described previously. The band intensities were measured by densitometry using ImageJ v1.4r software (NIH) and normalized with the intensities of actin, and the values were plotted on a semi-logarithmic graph against time and further fitted to an exponential line.

5.2.4. Microscopy and image analysis

Yeast cells expressing Hxt1-GFP were stained with FM4-64 (lipophilic styryl dye to

stain the vacuolar membrane, 1 µg/ml) and analyzed with Olympus FluoView confocal

microscope under 63X oil immersion objective lens using GFP or Texas Red filter.

Images from confocal microscope were captured by FluoView software (Olympus). At

least 200 cells showing the respective makers (e.g., FM6-64) were analyzed per each

condition. Standard deviations were calculated from three or more independent

experiments and are shown as error bars.

5.3. Results

5.3.1. Hxt1 protein levels are posttranslationally downregulated in response to

glucose starvation.

The expression levels of yeast nutrient transporters are regulated at both the

transcriptional and posttranslational levels (Kriel et al., 2011). To understand glucose

regulation of Hxt1 at a posttranslational level, we examined the abundance of Hxt1 at the

plasma membrane by Western blotting. To this end, the HXT1 gene was fused to the

GFP (green fluorescent protein) coding sequence on a centromeric plasmid (PMET25-

HXT1-GFP). Because HXT1 gene expression is repressed by Rgt1 under glucose

starvation conditions (Kim et al., 2003; Ozcan and Johnston, 1996), we interrupted

glucose regulation of HXT1 by replacing its promoter with the promoter of MET25,

which is not regulated by glucose (Kim et al., 2006; Pasula et al., 2007). We found that

the cell surface levels of Hxt1-GFP are greater in cells grown on high glucose (2%) than

in cells grown in the low glucose (~0.05%) or glucose-free medium (Gal, as 0% glucose)

(Figure 30A). These results, together with the previous observations of glucose induction

of HXT1 gene expression (Johnston and Kim, 2005; Kim et al., 2013), suggest that Hxt1

expression levels are regulated at both transcriptional and translational levels. We also

found that the amounts of immunodetected Hxt1-GFP in glucose-grown cells were

reduced by ~ 50% within 5 hr after the cells were shifted to galactose medium (Figures

30B and 30C).

Figure 30. Hxt1 protein levels are posttranslationally downregulated in response to glucose starvation. (A) Western blot analysis of the expression levels of Hxt1-GFP at the plasma membrane. Yeast cells (WT) expressing Hxt1-GFP were grown in SC-2% glucose medium to mid log phase (O.D600nm = 1.2-1.5) and equal amounts of cells were shifted to SC medium containing different glucose concentrations. Membrane fractions were immunoblotted with anti-GFP antibody (top panel), and the intensity of each band on the blot was quantified by densitometric scanning (bottom panel). (B) Yeast cells (WT) expressing Hxt1-GFP were grown in SC-2% glucose (Glucose) medium to mid log phase and equal amounts of cells were shifted to SC-2% galactose medium and incubated for indicated times. Membrane fractions were immunoblotted with anti-GFP antibody. (C) Determination of the half-life of Hxt1-GFP protein. The Western blot images in (B) were scanned and the half-life was determined, as described in the materials and methods.

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5.3.2. Glucose starvation induces endocytosis and subsequent vacuolar degradation of Hxt1.

Given that Hxt1-GFP protein levels are diminished in glucose-starved cells, we

examined whether this downregulation occurs via endocytosis by examining the stability

of Hxt1-GFP in the end3Δ mutant, which is defective in the internalization step of

endocytosis. We found that the cell surface levels of Hxt1-GFP in the end3Δ mutant are

high in both glucose- and galactose-grown cells (Figure 31A). Confocal microscopy also

demonstrated that Hxt1-GFP accumulates at the cell surface in cells grown in high

glucose medium and that ~90% of plasma membrane-localized Hxt1-GFP disappears

when cells are exposed to glucose-free medium (2% gal) (Figure 31B). However, Hxt1-

GFP was shown to be accumulated at the plasma membrane constitutively in the end3Δ

mutant, suggesting that Hxt1 may be endocytosed in response to glucose starvation

(Figure 31B). Of note, Hxt1-GFP seems to be constitutively localized to the vacuole,

and this occurs even in the end3Δ mutant (Figure 31B), raising the possibility that Hxt1 may be directed to the vacuole from the Golgi directly, as suggested previously for the vacuolar accumulation of the uracil permease Fur4 (Volland et al., 1994) and the maltose permease Mal61 (Gadura and Michels, 2006).

We next determined whether Hxt1-GFP degradation is stimulated by glucose starvation or by specific carbon sources. Raffinose is a trisaccharide, consisting of fructose-glucose-galactose that is equivalent to low glucose, because yeast cells cleave the fructose-glucose bond by invertase inefficiently and thus eventually obtain low levels of glucose from it (Ozcan and Johnston, 1999). Glucose and galactose only differ with respect to C-4, yet galactose does not enter through glucose transporters, suggesting that

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the glucose transporters display remarkable substrate specificity. Both Western blot and

confocal microscopy analyses revealed that Hxt1-GFP levels are high in cells grown on

glucose or raffinose and are very low in cells grown on galactose or ethanol (Figures 31C and 31D). These results support the view that Hxt1 is subjected to endocytosis and degradation in the vacuole under glucose starvation conditions. Hxt1-GFP levels are high

in cells grown on glucose or raffinose and are very low in cells grown on galactose or

ethanol (Figures 31C and 31D). These results support the view that Hxt1 is subjected to

endocytosis and degradation in the vacuole under glucose starvation conditions.

Figure 31. Glucose starvation induces endocytosis and subsequent degradation of Hxt1. (A) Yeast cells (WT and end3∆) expressing Hxt1-GFP were grown in SC-2% glucose (Glu) medium to mid-log phase and shifted to SC medium containing 2% galactose (Gal) for 6 hr. Membrane fractions were immunoblotted with anti-GFP antibody. (B) Yeast cells (WT and end3∆) expressing Hxt1-GFP was grown as described in (A). Confocal microscope image (top panel) and quantification of relative GFP

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fluorescence in the plasma membrane (bottom panel, *P < 0.05, **P < 0.001) were shown. (C) Yeast cells (WT) expressing Hxt1-GFP were grown in SC-2% glucose medium to mid-log phase and shifted to SC medium containing either 2% raffinose (Raf), 2% galactose (Galan et al.) or 2% ethanol (EtOH) for 6 hr. Membrane fractions were immunoblotted with anti-GFP antibody. (D) Yeast cells (WT) expressing Hxt1-GFP were grown as described in (C). Confocal microscope image (top panel) and quantification of relative GFP fluorescence in the plasma membrane (bottom panel, **P < 0.001) were shown. The FM6-64 dye was used to stain the vacuolar membrane (red), and actin was served as loading control in (A) and (C). Pma1 is frequently used as a loading control for membrane fractions; it, however, is not appropriate for this study because its expression is critically regulated by glucose.

5.3.3. The amino-terminal cytoplasmic domain of Hxt1 regulates its glucose

starvation-induced turnover.

To identify the regions of Hxt1 that regulate its turnover, we constructed deletion

mutants of Hxt1 that lack its amino (N)- or carboxy (C)-terminal cytoplasmic domain and

examined their stability in glucose-free medium (Figures 32A and 32B). We found that

Hxt1 degradation in galactose-grown cells is abolished by the deletion of its entire N-

terminal cytoplasmic domain (residues 1-59) (Figure 32C, ΔN) and that, by contrast, the

deletion of the C-terminal domain of Hxt1 (residues 513-570, Hxt1-GFP-ΔC) renders it

unstable in both glucose and galactose-grown cells. These results suggest that glucose

starvation-induced Hxt1 turnover may be regulated by its N-terminal domain and raise

the hypothesis that the C-terminal domain might play a role in the turnover of Hxt1 by

regulating the N-terminal domain (Figure 32C, ΔC). We tested this hypothesis by

examining the stability of Hxt1-GFP-ΔNΔC that lacks both the N- and C-terminal

domains and found that this protein, like Hxt1-GFP-ΔC, is inherently unstable (Figure

32C, ΔNΔC). These results suggest that the C-terminal domain of Hxt1 may not be directly involved in its glucose starvation-induced degradation but may contribute to its structural stability.

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To understand the effects of the N- or C-terminal deletion of Hxt1 on its function, we

analyzed glucose transport activity of the Hxt1 deletion constructs. A yeast mutant

lacking all 17 glucose transporters are unable to grow on glucose and this growth defect

was complemented by expression of any one of HXT genes (Ozcan and Johnston, 1999).

The hxt null strain was transformed with plasmids encoding wild type or truncated Hxt-

GFP proteins and scored for growth in glucose medium containing the respiratory

inhibitor Antimycin A (Figure 32D). We found that the growth defect of the hxt null

strain on glucose was restored by the expression of wild type Hxt1-GFP or Hxt1-GFP-

ΔN, but not of GFP-Hxt1ΔC or GFP-Hxt1ΔNΔC (Figure 32D) and that this may be

correlated with their stability. Of note, the growth rate of cells expressing Hxt1-GFP-ΔN

was faster than that of cells expressing wild type Hxt1-GFP. Confocal microscopy

showed that Hxt1-GFP-ΔN accumulates constitutively at the plasma membrane and that

the vacuolar localization of Hxt1-GFP-ΔN is markedly reduced, compared with that of wild type Hxt1-GFP1 (Figure 32E). As a result, the plasma membrane levels of Hxt1-

GFP-ΔN were 2-3 folds higher than those of wild type Hxt1-GFP, suggesting that Hxt1

localization to the vacuole is regulated by its N-terminal domain (Figure 32F).

Consequently, deletion of the N-terminal domain of Hxt1-GFP likely leads to its accumulation at the plasma membrane, enabling cells expressing Hxt1-GFP-ΔN to grow faster than cells expressing Hxt1-GFP (Figure 32D).

5.3.4. Rsp5 ubiquitin ligase mediates ubiquitination of Hxt1. Ubiquitination is a common signal for endocytosis and subsequent degradation (Hicke

and Dunn, 2003). Because a number of yeast nutrient transporters are ubiquitinated by the ubiquitin ligase Rsp5 and this process is affected by the ubiquitin isopeptidase Doa4

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which is required for recycling ubiquitin from ubiquitinated substrates (Amerik et al.,

2000; Rotin et al., 2000; Springael and Andre, 1998), we examined the stability of the

Hxt1 transporter in the strain carrying the doa4Δ or rsp5-1ts mutation. Western blot analysis indicated that Hxt1-GFP levels are constitutively high in the doa4Δ (Figure 33A, top) and rsp5-1ts mutants (Figure 33B, top). Consistently, Hxt1-GFP accumulates

constitutively at the plasma membrane in those mutant strains (the bottom panels of

Figures 33A and 33B). We also observed the absence of intracellular GFP signal in the

doa4Δ and rsp5-1 mutants, suggesting that ubiquitination plays a key role in the

movement of Hxt1 transporter into intracellular compartments. Similar observations were

made in cells expressing Hxt1-GFP-ΔN, in which the truncated Hxt1 transporter accumulates at the plasma membrane but its vacuolar localization is abolished (Figures

32E and 32F). These results are supported by previous findings that ubiquitin is involved in intracellular trafficking of plasma membrane proteins and mediates vacuolar targeting of the maltose permease Mal61.

5.3.4. K12 and K39 at the N-terminal domain of Hxt1 serve as putative ubiquitination sites.

Given that the N-terminal cytoplasmic domain of Hxt1 regulates its turnover, we examined whether the domain is responsible for its ubiquitination. We changed the four lysine residues present in the N- terminal domain of Hxt1—K12, K27, K35 and K39—to alanines and examined the stability of the resulting Hxt1 mutants. Substitutions of two of these residues (K12A and K39A) resulted in a marked increase of Hxt1 levels in galactose-grown cells, suggesting that the two lysine residues may serve as putative

105

ubiquitination sites (Figure 34A). Consistently, Hxt1K12A-GFP and Hxt1K39A-GFP were

accumulated at the plasma membrane in both glucose and galactose-grown cells (Figure

34B). Notably, these two Hxt1 mutants were resistant to degradation but were able to localize to the vacuole (Figure 34B). Therefore, it is likely that ubiquitination of Hxt1 at

K12 and K39 may be essential for its glucose starvation-induced vacuolar degradation but may not be involved in its vacuolar targeting. While the N-terminal domain of Hxt1 is important for its vacuolar accumulation (Figures 32E and 32F), any of the four lysine residues in the domain are not involved in its localization to the vacuole (Hxt1K27A-GFP and Hxt1K35A-GFP are also localized to the vacuole, data not shown). These observations

suggest that glucose starvation–induced downregulation of Hxt1 and its accumulation in

the vacuole may occur by separate mechanisms.

5.4. Discussion

This study demonstrates that the low affinity glucose transporter Hxt1 accumulates at

the plasma membrane in glucose-grown cells but is subjected to endocytosis and

subsequent vacuolar degradation in glucose-starved cells. As discussed above, high

affinity glucose transporters such as Hxt2 (Kruckeberg et al., 1999) and Hxt6/Hxt7

(Krampe et al., 1998) are downregulated by endocytosis in response to high glucose

concentrations (catabolite degradation) (Horak, 2003, 2013). The molecular mechanism

underlying this phenomenon is currently unknown but is proposed to be a signal-induced process in which glucose triggers the ubiquitination of glucose transporters that signals their endocytosis and subsequent degradation in the vacuole. However, our data showing that glucose starvation-induced Hxt1 degradation is also ubiquitin-dependent suggest that

106

glucose may not play a role in the ubiquitination of glucose transporters. Rather, we

support the view that the stability of the glucose transporters may be correlated with their

affinity for glucose and/or with their kinetic properties.

Figure 32. The N-terminal cytoplasmic domain of Hxt1 is required for its turnover. (A) Schematic diagram of predicted secondary structure of Hxt1. Twelve transmembrane domains and cytosolic N- and C- terminal tails are shown. (B) Schematic maps of Hxt1 constructs (Wt, 60-570 aa (∆N); 1-512 aa (∆C); 60-512 aa (∆N∆C)) showing lysine residues at its N-terminal domain. (C) Yeast cells (WT) expressing indicated Hxt1-GFP proteins were grown as described in Figure 31A, and plasma membrane fractions were immunoblotted with anti-GFP antibody (left panel), and the intensity of each band on the blot was quantified by densitometric scanning (right panel, **P < 0.001). Actin was served as loading control. (D) Yeast cells (hxt∆) expressing indicated Hxt1-GFP proteins were spotted on 2% glucose plate supplemented with Antimycin A (1µg/ml). The first

107 spot of each row represents a count of 5 x 107 cell/ml, which is diluted 1:10 for each spot thereafter. The plate was incubated for indicated times and photographed. (E) Yeast cells (WT) expressing indicated Hxt1-GFP proteins were grown as described in Figure 31A and analyzed by confocal microscopy. (F) Yeast cells (WT) expressing indicated Hxt1- GFP proteins were grown in SC-2% glucose medium to mid log phase and stained with FM6-64 (red). Confocal microscope images (top panel) and quantification of relative fluorescent intensity of Hxt1-GFP at the plasma membrane (bottom panel, * P <0.05) were shown.

Fig. 33. Hxt1 is ubiquitinated by the ubiquitin ligase Rsp5. (A) and (B) Yeast cells of indicated genotypes expressing Hxt1-GFP were grown as described in Figure 31A. Plasma membrane fractions were immunoblotted with anti-GFP antibody (A), and subcellular localization of Hxt1-GFP was analyzed by confocal microscopy (B). The FM6-64 dye was used to stain the vacuolar membrane (red), and actin was served as loading control.

108

Figure 34. K12 and K39 serve as putative ubiquitin-acceptor lysine residues. Yeast cells (WT) expressing indicated Hxt1-GFP proteins were grown as described in Figure 31A. Plasma membrane fractions were immunoblotted with anti-GFP antibody (A), and subcellular localization of Hxt1-GFP (Wt, K12A and K39A) was analyzed by confocal microscopy (B).

109

Conclusion and Future Direction

The ability to recognize and metabolize extracellular glucose is a fundamental

process crucial for survival of eukaryotic cells. The budding yeast Saccharomyces

cerevisiae has evolved a complex interconnected signaling pathway to utilize glucose optimally. In my dissertation research, I investigated the molecular basis of Rgt1- mediated repression of HXT genes and elucidated the role of HXT corepressor Mth1 and general transcription repressor complex Ssn6/Tup1 in bringing about repression of HXT gene expression in the absence of glucose. I showed that although Rgt1 is released from the HXT promoters upon being hyperphosphorylated by PKA in the presence of high

glucose conditions, this dissociation is not required for HXT gene repression, rather,

phosphorylation state of Rgt1 (which is controlled by Mth1) and its interaction with

Ssn6/Tup1 is necessary and sufficient to turn off HXT gene transcription. Yeast cells cope

with fluctuating concentrations of glucose in the environment and to carry out “aerobic

glycolysis”, they must efficiently recognize and utilize available glucose. I found that one

of the glucose sensors (Rgt2 and Snf3) or both, may be present in the plasma membrane

at a given glucose concentration. Snf3 may be major glucose sensor in low levels of

glucose and Rgt2, in high glucose conditions. The glucose sensors are degraded in the

vacuole by ubiquitination and endocytosis in the absence of glucose. Hence, in the

presence of a broad range of glucose concentrations, yeast cells maintain a constant

glucose sensor activity at the plasma membrane and rapidly respond to the changing

glucose levels. I have also studied the molecular mechanism of glucose regulation of

yeast glucose transporter Hxt1 and showed that like the glucose sensors, Hxt1 is also

downregulated post-translationally by ubiquitination and endocytosis-mediated vacuolar

110 degradation in the absence of glucose. Besides, I have identified several amino acid residues in Hxt1 which are important for glucose binding and may form the glucose transport channel.

Yeast cells are able to sense changes in nutrient status or binding of ligand by rapidly removing or degrading plasma membrane proteins. The mammalian casein kinase 1- gamma (CK1γ) homologs Yck1 and Yck2 are essential for nutrient sensing (Liu et al.,

2008; Moriya and Johnston, 2004). They are also involved in downregulation of various permeases and receptors such as uracil permease Fur4 (Marchal et al., 2000), the α-factor receptor Ste2 (Hicke et al., 1998) and the a-factor receptor Ste3 (Feng and Davis, 2000).

Yck1 and Yck2 are essential components of the Rgt2/Snf3 glucose signaling pathway where deletion of these two gene pairs completely abolishes the expression of HXT genes

(Moriya and Johnston, 2004). A recent study has demonstrated that Cu/Zn superoxide dismutase SOD1 binds at the C-terminus of Yck1/Yck2 and represses respiration (Reddi and Culotta, 2013). Although, it has been proposed that upon recognizing glucose, the glucose sensors activate Yck1/Yck2 to generate intracellular signal that induces HXT gene expression (Moriya and Johnston, 2004), the mechanism remains elusive. Further understanding the interplay between the glucose sensors and Yck1/Yck2 kinase will help us to elucidate the role of kinases in signal transduction pathways. A comprehensive mutational analysis of the glucose sensors Rgt2 and Snf3 may also be useful to identify the amino acid residues responsible for dictating the affinity of the sensors for glucose and understand the unique fermentative lifestyle of the budding yeast.

111

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APPENDIX I

Construction of yeast strains useful for screening drugs that inhibit glucose uptake and glycolysis

Abstract

The budding yeast Saccharomyces cerevisiae expresses different isoforms of glucose transporters (Hxts) in response to different levels of glucose. Here, I constructed reporter strains in which the nourseothricin (NAT) resistance gene is expressed under the control

of the HXT1, HXT2 or HXT3 promoter. The resulting HXT-NAT reporter strains exhibited

a strict growth dependence on glucose, and their growth could be easily controlled and

optimized by adjusting glucose concentration, demonstrating the value of the reporter

strains for studying the molecular basis of differential expression of HXT genes, as well

as for screening drugs that inhibit glucose uptake and glycolysis.

A1.1. Introduction

Cancer cells consume glucose avidly to maintain their high rate of aerobic glycolysis,

a phenomenon referred to as the Warburg effect (Warburg, 1956). A characteristic of this

phenomenon is increased glucose uptake as a result of overexpression of glucose

transporters. The budding yeast Saccharomyces cerevisiae also displays aerobic

glycolysis and thus consumes glucose vigorously even in the presence of oxygen by

upregulating expression of glucose transporter genes (Johnston and Kim, 2005). The

yeast has at least six glucose transporters (Hxt 1-4, Hxt6 and Hxt7) with various affinities

for glucose. The expression of HXT genes encoding glucose transporters is induced by

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glucose, but the yeast expresses only the glucose transporters best suited to the amount of

glucose available in the environment, as discussed below, implicating a correlation

between transporter function and regulation (Ozcan and Johnston, 1995, 1999). For

example, the low affinity glucose transporter gene HXT1 is expressed only in the

presence of high glucose concentration; in contrast, expression of high affinity glucose

transporter genes such as HXT2, HXT4 and HXT6 is induced by low levels of glucose. It

is widely known that expression of the Hxt1 transporter in the presence of high levels of

glucose leads to a reduction in the concentration of glucose in the culture medium due to

glucose uptake by Hxt1, which in turn leads not only to induction of the high affinity

glucose transporters but also to repression of Hxt1 expression. This regulatory property

of glucose transporters hampers the development of effective methods to determine

differential expression of different HXT genes in response to different levels of glucose.

Here, I present a novel method to overcome this limitation by constructing yeast reporter

strains that express only the transporters most appropriate for the given glucose

concentration in the presence of the antibiotic nourseothricin (NAT).

A1.2. Construction and properties of the HXT-NAT reporter genes.

I constructed HXT-NAT reporter strains that express NAT resistance genes under the control of the HXT1, HXT2 and HXT3 promoters by using PCR-based gene deletion strategy (Baudin et al., 1993; Wach et al., 1994). To do this, the NAT open reading frame

(ORF) was PCR-amplified from the NATMX cassette (Goldstein et al., 1999), and the

BY4741 yeast was transformed with the PCR product. To identify transformants in which the NAT gene was integrated with the HXT gene sequences, genomic DNA was isolated

133 and used as template in PCRs using two primer sets: one primer that anneals within the

NAT gene and another primer that anneals to the HXT gene locus outside the region altered. PCR products of the expected size confirmed the heterologous integration of the

NAT gene into the HXT gene sequence (Figure 35A).

HXT-NAT reporter strains were spotted on YP (2% bacto-peptone, 1% yeast extract) plates supplemented with various concentrations of glucose (0-2%) supplemented with

100 µg/ml NAT-sulfate. The first spot of each row represented a cell count of 5 × 107/ml, which was diluted 1:10 for each spot thereafter and incubated for 3 days. HXT-NAT reporter strains were shown to be susceptible to nourseothricin in the absence of glucose

(glycerol/ethanol medium), and exhibited resistance to the antibiotic only when glucose was present in the growth medium (Figure 35B). I found that the growth of the HXT1-

NAT reporter strain was more easily detected in high (2%) rather than low glucose concentration and that the HXT2-NAT reporter strain grows better in low glucose medium such as raffinose containing medium (raffinose is equivalent to low levels of glucose) than in high glucose medium. The HXT3-NAT reporter strain was shown to grow in both low and high glucose media, as reported previously (Ozcan and Johnston, 1995). These results indicate that the growth of three reporter strains in the presence of nourseothricin is regulated by glucose concentration. Glucose induction of HXT gene expression requires inactivation of Rgt1 repressor, a master repressor of HXT genes (Kim et al.,

2003). Rgt1 appears to mediate repression of all HXT genes by binding and recruiting the general transcription repressor complex Ssn6-Tup1 to the upstream regions of the genes

(Kim, 2004, 2009). It remains elusive how Rgt1 is inactivated by low and high glucose signals. In this regard, the system developed in this study would provide a novel, facile

134 and highly controllable method to study regulatory mechanisms of expression of specific

HXT genes in vivo. As Hxt2 is high affinity glucose transporter, similar to Hxt4, Hxt6 and

Hxt7, in that they all are expressed in low concentrations of glucose, we constructed only the HXT2-NAT reporter strain as a representative of the whole group.

Next I investigated whether our system is useful for screening drugs that inhibit glucose uptake and glycolysis by examining the effect of the glucose analog 2-deoxy-D- glucose (2-DG) on the growth of the HXT-NAT reporter strains. 2-DG, a competitive inhibitor of glucose, is transported into the cells by glucose transporters but cannot be metabolized. To this end, the HXT-NAT reporter strains were tested for growth in glucose plus NAT-sulfate plates containing various concentrations of 2-DG and incubated for 3 days. The results showed that growth of all three reporter strains tested was significantly inhibited by 2-DG and that expression of the HXT1 was more sensitive to 2-DG than that of the HXT2 and HXT3 gene (Figure 36). These observations are consistent with our previous finding that 2-DG inhibits the induction of expression of HXT genes (Jouandot et al., 2011) and also support the view that the expression of different HXT genes may be regulated by different mechanisms (Johnston and Kim, 2005). The critical components for the glucose induction of HXT expression are the glucose sensors Rgt2 and Snf3, plasma membrane proteins that sense the presence of extracellular glucose and generate a signal that leads to inactivation of Rgt1 (Johnston and Kim, 2005; Ozcan and Johnston,

1999). The glucose sensors are very similar to glucose transporters in their structures but appear to be unable to transport glucose (Ozcan and Johnston, 1999). Therefore, my results suggest that the HXT-NAT reporter strains may be useful for screening drugs that inhibit glucose sensors and/or glucose transporters, as well as components of the glucose

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signaling pathways involved in regulation of HXT gene expression (Johnston and Kim,

2005).

Figure 35. The HXT-NAT reporter strains exhibit strict growth dependence on glucose. (A) Schematic representation of the construction of the HXT-NAT reporter strains. For example, the HXT1 ORF was replaced with the NAT ORF, which was PCR- amplified from the NAT cassette, using the PCR-based gene deletion method. Expression of the NAT gene is under the control of the respective HXT promoter. (B) Yeast cells (BY4741, Matα his3∆1 leu2∆0 ura3∆0 met15∆1) were spotted on YP (2% bacto- peptone, 1% yeast extract) plates containing 5% glycerol (Gly) + 2% ethanol (EtOH), 2% glucose (Hanyaloglu and von Zastrow) or 2% raffinose (Raf) supplemented with 100 µg/ml NAT-Sulfate. The first spot of each row represents a count of 5 × 107 cells/ml, which was diluted 1:10 for each spot thereafter. The plates were incubated for 3 days.

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Figure 36. 2-DG inhibits growth of the HXT-NAT reporter strains. Yeast cells (BY4741, Matα his3∆1 leu2∆0 ura3∆0 met15∆1) were spotted on YP (2% bacto- peptone, 1% yeast extract) plates containing 2% glucose, 100 µg/ml NAT-Sulfate and various concentrations of 2-DG. The first spot of each row represents a count of 5 × 107 cells/ml, which was diluted 1:10 for each spot thereafter. The plates were incubated for 3 days.

A1.3. Conclusion

In this study, I constructed several yeast HXT-NAT reporter strains that are resistant to the antibiotic nourseothricin only in the presence of glucose. My results demonstrating that the growth of the reporter strains can be easily controlled by glucose concentration suggest that the reporter strains may serve as useful tools for studying the regulatory mechanism of differential expression of HXT genes in response to various concentrations of glucose. Increased glucose uptake associated with aerobic glycolysis is a hallmark of cancer and its inhibition is a cancer therapeutic strategy that is being intensively studied.

This phenomenon is also observed in budding yeast, allowing us to consider the reporter strains useful for screening drugs that inhibit glucose uptake and glycolysis in yeast cells as well as cancer cells.

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