The Characterisation of CDP-Diacylglycerol Synthases In

The Characterisation of CDP-Diacylglycerol

Synthases in Lipid Droplet Dynamics and

Adipocyte Development

A thesis presented for the degree of Doctor of Philosophy

Tamar Sonia Kapterian

School of Biotechnology and Biomolecular Sciences

University of New South Wales

August, 2013

Originality Statement

ORIGINALITY STATEMENT

AUTHENTICITY STATEMENT

Acknowledgements

ACKNOWLEDGEMENTS

I would like to thank my supervisor Prof Rob Yang for giving me the opportunity to undertake my PhD in his lab. Thanks for being a really great supervisor and for giving me such an interesting project! The last 3.5 years have flown by and that’s largely due to having such a great work environment and a project that I really enjoyed. With that being said, I also have to thank all Yang lab members, past and present (especially

Robin, Stephen, Jack and Truong). You truly made the lab such a wonderful place to work. Thanks for all the discussions, answering my questions and basically just the endless help! I would also like to thank Prof Ian Dawes and the Dawes lab for providing me with helpful discussions and resources.

Lastly, I would like to thank my mum, dad and sister. There are too many things to thank you for, so I’m going to keep this short and sweet. Your support throughout this entire PhD has been nothing but positive. I probably wasn’t the easiest person to put up with so thank you for always being there and being so understanding.

III

Abstract

ABSTRACT

Changes in the cellular dynamics of lipid droplets (LDs) are associated with human metabolic disorders such as obesity. Adipocyte differentiation is the process where preadipocytes differentiate into mature adipocytes. Understanding the molecular mechanisms involved in LD dynamics and adipocyte development will provide insights into how therapeutic treatments can be developed against human metabolic diseases.

In mammals, there are two cytidine-diphosphate-diacylglycerol (CDP-DAG) synthases,

CDS1 and CDS2. CDP-DAG synthases catalyse the formation of CDP-DAG by utilising phosphatidic acid (PA), the precursor for all phospholipids and triacylglycerols (TAGs).

This study has characterised CDS1 and CDS2 as enzymes involved in lipid storage, at both the cellular and systemic level. The down-regulation of CDS1 or CDS2 resulted in the formation of “Supersized” LDs (SLDs) in cultured cells. To identify the impact of knocking down CDS1 and CDS2 on lipid profiles, lipid analysis was carried out on extracted neutral lipids. Mass spectrometry indicated a significant increase in total cellular PA of siCDS1 cells. Fluorescence microscopy revealed that PA strongly accumulates in the endoplasmic reticulum (ER) of HeLa cells down-regulated with CDP-

DAG synthases. Importantly, depleting CDS1 almost completely blocked the differentiation of 3T3-L1 preadipocytes, whereas depleting CDS2 had a moderate inhibitory effect on adipocyte differentiation. Furthermore, transient siRNA transfections in mature adipocytes showed that CDP-DAG synthases are involved in the fusion of large LDs. A role for CDS1 and CDS2 in isoproterenol stimulated lipolysis was also revealed, with a decrease in phosphorylated-perilipin when CDS1 or CDS2 was depleted. This study also established a functional interaction between CDS1/2 and

Abstract

Fld1p/Seipin. These findings indicate that CDP-DAG synthases are more than just enzymes involved in phospholipid synthesis. They are regulators of lipid storage and play important roles in LD growth, adipogenesis and adipocyte physiology.

Abbreviations

ABBREVIATIONS

°C Degrees celcius

µM Micromolar ade Adenine

AGPAT 1-acylglycerol-3-phosphate-O-acyltransferase aP2 Adipocyte fatty acid binding protein

ATGL Adipose triglyceride lipase

BCA bicinchoninic acid

BSA Bovine Serum Albumin

BFP Blue fluorescent protein

C. elegans Caenorhabditis elegans

C/EBP CCAAT/enhancer binding protein

Casp3 Caspase 3

CDP-DAG Cytidine diphosphate-diacylglycerol

CDS Cytidine diphosphate-diacylglycerol synthase

CEPT choline/ethanolamine phosphotransferase

CIDE Cell death-inducing DNA fragmentation factor-like effector

DAG Diacylglycerol

DGAT Diacylglycerol acyltransferase

DIC Differential interference contrast

DMEM Dulbecco's Modified Eagle Medium

E. coli Escherichia coli

ER Endoplasmic Reticulum

Abbreviations

FA Fatty Acid

FBS Fetal Bovine Serum

FSP27 fat specific protein 27 g centrifugal acceleration with respect to gravity (9.0806 m/s2)

GFP Green fluorescent protein

GFP-PASS Green fluorescent protein- phosphatidic acid sensor with specific sensitivity

GPAT glycerol phosphate acyl transferase h hour his Histidine

HSL Hormone sensitive lipase

IBMX isobutylmethylxanthine

INO1 Inositol-3-phosphate synthase

LDs Lipid droplets leu Leucine

Lsd Drosophila Lipid Storage Droplet gene m Minute mito Mitochondria mM Millimolar mTOR Mammalian target or rapamycin

NCS Newborn calf serum nM Nanomolar

OP3 Phospholipid methyltransferase

PA Phosphatidic acid

PAH1 phosphatidate (PA) phosphatase

VII

Abbreviations

PC Phosphatidylcholine

PDE Phosphodiesterase

PE Phosphatidylethanolamine

PEMT Phosphatidylethanolamine N-methyltransferase

PG Phosphatidylglycerol

PI Phosphatidylinositol

PIS Phosphatidylinositol synthase

PKA Protein Kinase A

PPARγ Peroxisome proliferator activated receptor gamma

Pref1 Preadipocyte factor 1

RFP Red fluorescent protein rpm Revolutions per minute s Second

S. cerevisiae Saccharomyces cerevisiae

S6K S6 kinase

SC Synthetic complete

SDS Sodium dodecyl sulfate shRNA Short hairpin ribonucleic aicd siCDS1 CDS1 siRNA siCDS2 CDS2 siRNA siCTRL Universal negative control siRNA siLPIN1 Lipin-1 siRNA siRNA Small interfering ribonucleic acid

SLDs Supersized lipid droplets

SREBP1c Sterol regulatory element binding protein 1c

VIII

Abbreviations

TAG Triacylglycerol

TLC Thin layer chromatography trp Tryptophan ura Uracil

V Volts

VEGFA Vascular endothelial growth factor

WT Wild type

X-gal 5-bromo-4-chloro-3-indolyl-beta-D-galactopyranoside

YPD Rich glucose medium

Table of Contents

TABLE OF CONTENTS

ORIGINALITY STATEMENT ...... I

AUTHENTICITY STATEMENT ...... II

ACKNOWLEDGEMENTS ...... III

ABSTRACT ...... IV

ABBREVIATIONS ...... VI

TABLE OF CONTENTS ...... X

LIST OF TABLES ...... XIII

LIST OF FIGURES ...... XIV

PUBLICATION RECORD AND CONFERENCE PRESENTATIONS ...... XVII

PUBLICATION RECORD ...... XVII CONFERENCE PRESENTATIONS ...... XVII 1.0 INTRODUCTION ...... 1

1.1 MAMMALIAN FAT STORAGE ...... 2 1.2 LIPID DROPLETS AT THE CELLULAR LEVEL...... 2 1.2.1 The Occurrence of Lipid Droplets ...... 2 1.2.2 The Composition and Morphology of Lipid Droplets ...... 3 1.2.3 Phospholipid Biosynthesis Pathway ...... 5 1.3 LIPID DROPLET FUNCTION ...... 6 1.4 LIPID DROPLET BIOGENESIS ...... 7 1.4.1 The “budding” model ...... 8 1.4.2 The “hatching” model ...... 9 1.4.3 Alternative models ...... 10 1.5 THE REGULATION OF LIPID DROPLET GROWTH ...... 11 1.5.1 Phospholipids ...... 11 1.5.2 Proteins ...... 14 1.5.3 Other protein factors ...... 20 1.6 ADIPOGENESIS (ADIPOCYTE DIFFERENTIATION) AT THE SYSTEMIC LEVEL ...... 22 1.6.1 Peroxisome Proliferator-Activated Receptor γ (PPARγ) ...... 24 1.6.2 CCAAT/ enhancer binding protein family (C/EBP) ...... 24 1.7 LIPOLYSIS ...... 25 1.8 MAMMALIAN TARGET OF RAPAMYCIN (MTOR) PATHWAY ...... 27 1.9 CDP-DAG SYNTHASES ...... 30 1.10 LIPID DROPLETS IN PREVAILING DISEASES ...... 32 1.11 SUMMARY ...... 33 1.12 AIMS ...... 34 2. MATERIALS AND METHODS ...... 36 X

Table of Contents

2.1 GENERAL MATERIALS AND METHODS ...... 36 2.2 GENERAL BUFFERS ...... 40 2.3 QRT-PCR PRIMER DESIGN ...... 41 2.4 PLASMIDS ...... 43 2.4.1 Vector Construction ...... 44 2.5 MAMMALIAN CELL CULTURE ...... 47 2.5.1 Adipocyte Differentiation ...... 47 2.5.2 Cell Transfection ...... 48 2.5.3 Retroviral shRNA Stable Transfections ...... 48 2.5.4 Fatty Acid-Supplemented Medium ...... 49 2.6 YEAST STRAINS ...... 50 2.6.1 Yeast Growth Conditions ...... 50 2.6.2 Yeast Transformation (TRAFO method) ...... 51 2.7 MICROSCOPY ...... 52 2.7.1 Fluorescence Microscopy ...... 52 2.7.2 Live Cell Imaging ...... 53 2.7.3 Immunofluorescence ...... 53 2.7.4 Fluorescence Resonance Energy Transfer (FRET) by Acceptor Photobleaching ...... 54 2.8 LIPID ANALYSIS ...... 55 2.8.1 Oleate incorporation...... 55 2.8.2 Neutral Lipid Extraction ...... 55 2.8.3 Phospholipid Extraction ...... 56 2.8.4 Thin Layer Chromatography (TLC) ...... 56 2.8.5 Mass Spectrometry ...... 57 2.8.6 Oil Red O Stain ...... 57 2.9 RNA EXTRACTION AND CDNA SYNTHESIS ...... 58 2.10 QUANTITATIVE RT-PCR (QRT-PCR) ...... 58 2.11 PROTEIN ANALYSIS ...... 58 2.11.1 Western Blotting ...... 58 2.11.2 Co-Immunoprecipitation by Dynabeads ...... 61 2.11.3 DUALmembrane yeast two-hybrid system ...... 62 2.12 BIOCHEMICAL ASSAYS ...... 63 2.12.1 Lipolysis Assay ...... 63 2.13 STATISTICAL ANALYSIS ...... 63 3. CHAPTER 3. CDP-DAG SYNTHASES REGULATE THE GROWTH OF LIPID DROPLETS AND ADIPOCYTE DEVELOPMENT ...... 65

3.1 BACKGROUND ...... 65 3.2 REVERSE GENETIC SCREEN IN YEAST IDENTIFIED CDS1 AS A ‘SUPERSIZED’ LIPID DROPLET MUTANT ...... 67 3.3 DOWN-REGULATION OF CDP-DAG SYNTHASES FORMED “SUPERSIZED” LIPID DROPLETS ...... 68 3.3.1 mRNA Expression Levels of CDP-DAG Synthases ...... 70 3.4 SUPERSIZED LIPID DROPLETS EMERGE FROM CLUSTERED LIPID DROPLETS ...... 71 3.5 THE LOCALISATION OF CDS1 AND CDS2 ...... 73 3.6 THE SYNTHESIS OF TRIACYLGLYCEROLS (TAG) IS INCREASED IN SICDS1 & SICDS2 CELLS ...... 76 3.7 PA METABOLISM UPON KNOCKING-DOWN CDS1 OR CDS2 ...... 78 3.8 OVEREXPRESSION OF CDP-DAG SYNTHASES INHIBITS LIPID DROPLET SYNTHESIS ...... 83 3.9 OVEREXPRESSION OF CDS INHIBITS THE FORMATION OF SLDS ...... 84 3.10 DOWN REGULATION OF CDS1 INHIBITS ADIPOCYTE DIFFERENTIATION ...... 87 XI

Table of Contents

3.10.1 Down-regulation of CDS2 using shRNA (shCDS2) inhibits adipocyte differentiation ...... 92 3.11 DISCUSSION ...... 94 3.12 CONCLUSION ...... 100 4. CHAPTER 4. THE CHARACTERISATION OF CDP-DAG SYNTHASES IN MATURE ADIPOCYTES ...... 101

4.1 BACKGROUND ...... 101 4.2 DOWN-REGULATION OF CDP-DAG SYNTHASES IN ADIPOCYTES RESULTS IN ‘SUPERSIZED’ LIPID DROPLETS ...... 102 4.3 LIPID DROPLET FUSION IN SICDS1 AND SICDS2 MATURE ADIPOCYTES ...... 103 4.4 DOWN-REGULATION OF CDP-DAG SYNTHASES SUPPRESS ADIPOCYTE MAINTENANCE ...... 104 4.5 CDP-DAG SYNTHASES ARE INVOLVED IN LIPOLYSIS ...... 105 4.5.1 Down-Regulation of CDS1 Diminished Phosphorylated Protein Kinase A (PKA) and Phosphorylated Perilipin Expression ...... 107 4.5.2 CDP-DAG Synthases Decrease Lipolytic Activity ...... 110 4.6 CDP-DAG SYNTHASES ARE IMPLICATED IN INSULIN SIGNALLING ...... 111 4.6.1 Down-Regulation of CDP-DAG Synthases Increased the Expression of Phosphorylated Akt .. 113 4.6.2 CDP-DAG Synthases alter mTOR Expression during Lipolysis ...... 114 4.7 DISCUSSION ...... 117 4.8 CONCLUSION ...... 120 5. CHAPTER 5: CDP-DAG SYNTHASE FUNCTIONALLY INTERACTS WITH FLD1 .. 122

5.1 BACKGROUND ...... 122 5.2 FLD1P PHYSICALLY INTERACTS WITH CDS1P UNDER THE DUALMEMBRANE YEAST TWO HYBRID SYSTEM ...... 123 5.3 HUMAN CDP-DAG SYNTHASE1 DOES NOT PHYSICALLY INTERACT WITH SEIPIN ...... 125 5.4 OVEREXPRESSION OF CDS1 PARTIALLY RESCUES THE ‘SUPERSIZED’ PHENOTYPE IN FLD1∆ CELLS ...... 127 5.5 KNOCKOUT OF FLD1 IN TET-CDS1 CELLS FORMS “SUPERSIZED” LIPID DROPLETS ...... 129 5.5.1 Supersized Lipid Droplets Emerge from Clustering in siSeipin HeLa Cells ...... 132 5.6 CDS2 AND SEIPIN LOCALISE TO THE ENDOPLASMIC RETICULUM (ER) ...... 133 5.6.1 CDP-DAG Synthase2 and Seipin do not Interact by FRET/Acceptor Photobleaching (AB) ...... 134 5.7 DISCUSSION ...... 135 5.8 CONCLUSION ...... 139 6. CHAPTER 6: FUTURE DIRECTIONS AND FINAL CONCLUSIONS ...... 140

6.1 CDP-DAG SYNTHASES REGULATE LIPID DROPLET GROWTH AND ADIPOCYTE DEVELOPMENT ...... 140 6.2 THE CHARACTERISATION OF CDP-DAG SYNTHASES IN MATURE ADIPOCYTES ...... 144 6.3 CDP-DIACYLGLYCEROL SYNTHASE FUNCTIONALLY INTERACTS WITH FLD1 ...... 147 6.4 PUTTING IT ALL TOGETHER – CDP-DIACYLGLYCEROL SYNTHASES AND HUMAN LIPID STORAGE...... 149 REFERENCES ...... 152

APPENDIX ...... 168

FIGURE LEGENDS OF DIGITAL FILES ...... 168

XII

List of Tables

LIST OF TABLES

Table 1-1 Materials used in this study 36

Table 1-2 Nucleotide sequence of qRT-PCR primers designed in this study 41

Table 1-3 Plasmids used in this study that were obtained as gifts 43

Table 1-4 Primer sequences for designed constructs 45

Table 1-5 List of amino acids added to SC medium 51

Table 1-6 Recipe for buffers used in immunofluorescence 53

Table 1-7 Recipe for Oil Red O solution 57

Table 1-8 Buffers used in western blotting 60

XIII

List of Figures

LIST OF FIGURES

1-1 Lipid Droplet Composition 4

1-2 Phospholipid Biosynthesis Pathway 6

1-3 Proposed Models for Lipid Droplet Biogenesis 8

1-4 The Regulation of LD growth is mediated by phospholipids and proteins 13

1-5 Adipocyte Differentiation 23

1-6 Lipolysis 26

1-7 The mTOR Signalling Pathway 29

1-8 Role of CDP-DAG Synthase in lipid synthesis 31

3-1 CDS1 Expression in S. cerevisiae 68

3-2 Transient siRNA knock-down of CDS1 and CDS2 69

3-3 qRT-PCR of siCDS1 and siCDS2 Transfected Cell Lines 71

3-4 Time Course of Oleate Treated HeLa Cells 72

3-5 Co-localisation of Rab10 and CDP-DAG Synthases 74

3-5-1 Co-localisation of Rab10 with lipid droplets 75

3-6 Co-localisation of GFP-PIS and CDP-DAG synthases 76

3-7 Neutral lipid quantification in siCTRL, siCDS1 and siCDS2 HeLa cells 77

3-8 Quantification of total PC, PE, PG and PI in siCTRL, siCDS1 and siCDS2 cells 78

3-9 Quantitative analysis of PA species by mass spectrometry 80

3-9-1 Quantitative analysis of PA species in siCDS2 cells by mass spectrometry 81

3-10 Co-localisation of pmGFP-PASS and DsRed-ER in siCDS1 and siCDS2 cells 82

3-11 Co-localisation of pmGFP-PASS and Mito-RFP in siCDS1 and siCDS2 cells 83

XIV

List of Figures

3-12 Overexpression of CDP-DAG synthases 84

3-13 CDS Overexpression in siLPIN1 cells 86

3-14 PAH1 overexpression in Tet-CDS1 cells 87

3-15 Expression of CDS1 and CDS2 during adipocyte differentiation 88

3-16 Down-regulation of CDS1 during adipocyte differentiation 89

3-17 Oil Red O staining and Imaging on siCDS1 induced 3T3-L1s 90

3-18 Down-regulation of CDS2 during adipocyte differentiation 91

3-19 Oil Red O staining and Imaging on siCDS2 induced 3T3-L1s 92

3-20 Down-regulation of CDS2 using shRNA 93

4-1 Down-regulation of CDP-DAG synthases in mature adipocytes 103

4-2 qRT-PCR of siCDS1 and siCDS2 mature adipocytes 105

4-3 CDP-DAG synthases are involved in lipolysis 107

4-4 Expression of p-PKA and p-perilipin in siCDS1 and siCDS2 mature adipocytes 109

4-5 Free glycerol assay on siCDS1 and siCDS2 mature adipocytes 111

4-6 CDP-DAG synthase expression upon wortmannin treatment 112

4-7 Phosphorylated Akt expression in siCDS1 and siCDS2 mature adipocytes 114

4-8 mTOR expression during lipolysis in siCDS1 and siCDS2 mature adipocytes 116

5-1 Deletion of FLD1 and down-regulation of CDS1 results in “supersized” lipid droplets 122

5-2 Interaction of Fld1p and Cds1p using the DUALmembrane system 124

5-3-1 Co-immunoprecipitation of CDS1 and Seipin 126

5-3-2 qRT-PCR expression in siRNA transfected cells 127

5-4-1 Overexpression of CDS1-GFP in fld1Δ cells 128

List of Figures

5-4-2 Seipin-mCherry overexpression 129

5-5-1 Knockout of FLD1 from Tet-CDS1 cells 130

5-5-2 Double knock-down of CDP-DAG synthase and Seipin 131

5-5-3 Down-regulation of Seipin in oleate treated HeLa cells 132

5-6-1 Co-localisation of Seipin and CDS2 134

5-6-2 FRET analysis of Seipin and CDS2 135

6-1 Down-regulation of CDP-DAG synthase in PA metabolism 150

XVI

Publication Record and Conference Presentations

PUBLICATION RECORD AND CONFERENCE PRESENTATIONS

PUBLICATION RECORD

 Fei W, Shui G, Zhang Y, Krahmer N, Ferguson C, Kapterian TS, Lin RC, Dawes IW, Brown AJ, Li P, Huang X, Parton RG, Wenk MR, Walther TC, Yang H. A role for phosphatidic acid in the formation of supersized lipid droplets, PLoS Genetics, 2011, 7(7): e 1002201.  Fei, W., Li, H., Shui, G., Kapterian, T.S., Bielby, C., Du, X., Brown, A. J., Li, P., Wenk, M.R., Liu, P., and Yang, H. and Molecular characterization of seipin and its mutants: implications for seipin in triacylglycerol synthesis J. Lipid Res, 2011. 52(12): p. 2136- 2147.  Minh T. Ta, Tamar S. Kapterian, Weihua Fei, Ximing Du, Andrew J. Brown, Ian W. Dawes and Hongyuan Yang. Accumulation of squalene is associated with the clustering of lipid droplets. FEBS Journal, 2012. 279(22): p. 4231-4244.  Kapterian, T.S., Zhang, Y., Fei, W., Dawes, I.W and Yang H. 2013, CDP-diacylglycerol synthases regulate the growth of lipid droplets and adipocyte development. 2013, to be submitted

CONFERENCE PRESENTATIONS

 ComBio2012: 23 - 27 September 2012, Adelaide Convention Centre, South Australia Oral Presentation: The Role of CDP-DAG Synthases in Lipids Droplet Dynamics and Adipocyte Development. Tamar S Kapterian & Hongyuan Yang.  Gordon Research Conference, The Molecular Biology of Lipids, 21 – 26 July 2013, Waterville Valley, New Hampshire, USA Poster Presentation: The Characterisation of CDP-DAG Synthases in Lipids Droplet Dynamics and Adipocyte Development. Tamar S Kapterian & Hongyuan Yang.

XVII

Introduction

1.0 INTRODUCTION

Lipid droplets (LDs) are no longer only regarded as an inert reservoir of fat. They are highly dynamic organelles that are also involved in many cellular functions including protein storage and degradation, as well as membrane and lipid trafficking [1, 2]. LDs contain a highly hydrophobic neutral lipid core surrounded by a phospholipid monolayer [3]. In humans, the lipid core comprises mainly triacylglycerols (TAGs) and cholesteryl esters (CEs). The biogenesis of LDs remains poorly understood. While several models have been proposed, the prevailing theory is that LDs originate from the endoplasmic reticulum (ER) [4]. The size of LDs varies within different tissues and even within the same cell type [5]. ‘Supersized’ LDs in the budding yeast

Saccharomyces cerevisiae, often exhibit up to 50 times the volume of wild-type cells[6].

Changes in the cellular dynamics of LDs are associated with human metabolic disorders such as obesity. Over the past decade, the occurrence of obesity and its associated disorders has significantly increased in Australia and other Western countries [7]. The development of obesity is characterised by the accumulation of adipocytes and the expansion of lipids, due to at least in part, enhanced adipocyte differentiation [8].

Understanding the physiological relevance of ‘supersized’ LDs and the molecular mechanisms involved in LD dynamics and adipocyte development will provide insight into how therapeutic strategies can be developed against human metabolic diseases.

Introduction

1.1 Mammalian Fat Storage

The capacity to store TAG in LDs is an evolutionarily conserved process, and has been studied in yeast through to higher mammals. LDs can vary in size, from 400nm in yeast, to >100µm in white adipocytes. White adipocytes are known as the major energy reservoir of TAG [9]. Unlike white adipocytes, brown adipocytes have small LDs rather than large LDs and induce energy expenditure in the form of thermogenesis [10].

As a result of internal and external factors, LDs can differ in size within the same cell.

Traditionally, adipocytes have been regarded as the ultimate fat storing organelle. By forming one large unilocular LD, white adipocytes display an efficient lipid storing mechanism, particularly during lipotoxicity [11]. In contrast, during excess lipid accumulation, non-adipocytes respond to lipid storage by either increasing or decreasing the number and size of LDs [12]. Altering the number of LDs is beneficial during lipolysis as it provides a greater surface area for lipases.

1.2 Lipid Droplets at the Cellular Level

1.2.1 The Occurrence of Lipid Droplets

Lipid droplets are found in all types of eukaryotic cells such as yeast, algae, plants and mammals [13-15]. A small collection of prokaryotes are known to accrue lipids in the form of LDs. These prokaryotes belong to the Gram-positive bacterium [16]. In mammalian adipocytes, TAGs are the predominant lipid ester. In ovarian, testicular and adrenocortical cells, CE is the major lipid required for hormone synthesis [17].

Introduction

1.2.2 The Composition and Morphology of Lipid Droplets

Lipid droplets are dynamic organelles that contain a highly hydrophobic neutral lipid core surrounded by a phospholipid monolayer [3]. In Saccharomyces cerevisiae, triacylglycerols (TAG) and ergosterol esters form the predominant mass of the lipid core. In humans, the lipid core comprises mainly of TAGs and CEs (Figure 1-1). Other endogenous lipids like retinol ester and ether-linked glycerolipids can also be found in the core [18-20].

The composition of the phospholipid monolayer of the LD is similar to that of the endoplasmic reticulum (ER) membrane. This suggests that the ER is a possible site for

LD biogenesis. Leber et al. revealed that the phospholipid arrangement of the LD resembled that of the ER membrane, given that phosphatidylcholine (PC), phosphatidylethanolamine (PE) and phosphatidylinositol (PI) are enclosed in this array

[13, 17, 21]. In contrast, Tauchi-Sato et al. demonstrated by means of mass spectrometry that the fatty acid composition of PC in LDs varies from that of the ER

[18].

Introduction

Figure 1-1: Lipid Droplet Composition. Lipid droplets comprise a neutral lipid core which is primarily made up of TAGs and CEs in mammals. The lipid core is covered by a layer of phospholipids. Proteins on the LD surface are believed to function in lipid homeostasis, lipid structure and rigidity.

In mammals, a copious amount of proteins are localised to the LD surface. This suggests that they may play a significant role in LD stability, structure and function [22,

23]. The most well-known LD surface proteins belong to the PAT (Perilipin, Adipose

Differentiation Related Protein and the Tail Interacting Protein of 47kDa) family of proteins. Perilipin and adipose differentiation related protein (ADRP) are LD specific while TIP47 and S3-12 are not confined to LDs [22, 23].

Perilipin is a phosphoprotein that is expressed in adipocytes and steroidogenic cells, and is likely to have a significant role in cAMP-dependent lipolysis [24, 25]. Perilipin specifically localises to the surfaces of intracellular LDs [22, 26, 27]. ADRP is a protein that has been associated with adipocyte differentiation. Unlike perilipin, ADRP is expressed in all mammalian tissues [24, 28]. The expression of ADRP on the LD surface is greatly associated with the amount of neutral lipid in the cell [29]. Unlike Perilipin

Introduction

and ADRP, TIP47 is abundant throughout the cytosol and has broad tissue distribution

[30, 31].

1.2.3 Phospholipid Biosynthesis Pathway

Phospholipids are vital for the growth of LDs, with both structural and signalling functions [32, 33]. In mammals, the phospholipid biosynthesis pathway has been well studied and consists of enzymes which are evolutionarily conserved (Figure 1-2). The

LD surface is covered with a range of phospholipids, with PC and PE the most abundant

[34]. Glycerol-3-phosphate is converted to lysophosphatidic acid (LPA) by the enzyme,

GPAT [35]. Phosphatidic acid (PA) is the key lipid intermediate responsible for the de novo synthesis of all phospholipids and TAG storage. It is a product of lysophosphatidylacyltransferase (LPAAT), which converts LPA to PA.

In mammals, PA is utilised by CDP-DAG synthases, CDS1 or CDS2 for the synthesis of

CDP-DAG, the precursor to the synthesis of phosphatidylinositol (PI), phosphatidylglycerol (PG) and cardiolipin (CL) [36]. PA phosphatases (Lipin) convert PA to diacylglycerol (DAG), which is then converted to TAG through diacylglycerol- acyltransferase (DGAT). Unlike in yeast where PS, PE and PC can be synthesised from the CDP-DAG pathway, the synthesis of PE and PC in mammals primarily requires DAG, through the Kennedy pathway.

Introduction

Figure 1-2: Phospholipid Biosynthesis Pathway in Mammals. The precursor to the de novo synthesis of all phospholipids is phosphatidic acid (PA), which is utilised by CDP-DAG synthase (CDS) and converted to CDP-DAG, or, by PA phosphatases such as Lipin for the synthesis of DAG and TAG. Abbreviations are: Glycerol-3-phosphate acyltransferases (GPAT); 1- acylglycerol -3-phosphate acyltransferase (AGPAT); cytidine-diphosphate-diacylglycerol synthase (CDS); phosphatidylinositol synthase (PIS); phosphatidylglycerolphosphate synthase (PGPS); cardiolipin synthase (CLS); diacylglycerol kinase (DAGK); diacylglycerol acyltransferases (DGAT); CDP-Ethanolamine (CDP-Eth); ethanolaminephosphotransfease (EPT); phosphoethanolamine methyltransferase (PEMT); choline phosphotransferse (CPT); cholinetransferase (CT); CDP-phosphate (CDP-P); choline kinase (CK); phosphatidylserine synthase-1 (PSS1); phosphatidylserine synthase-2 (PSS2), ethanolaminetransferase (ET); ethanolamine-phosphate (Ethanolamine-P) and ethanolamine kinase (EK).

1.3 Lipid Droplet Function

Traditionally, LDs have been regarded as a reservoir for storing excess lipids. Emerging evidence proposes that LDs function as dynamic organelles with a key role in lipid metabolism, membrane trafficking, membrane (phospholipid) and steroid synthesis [1,

37]. Furthermore, LDs have been found to not only engage in intracellular cholesterol homeostasis, but they might also be involved in unrelated activities such as cell 6

Introduction

signalling, protein storage and protein degradation [38-40]. Emerging evidence has also revealed that LDs are storage sites for unstable proteins [41].

Pathogens utilise LDs for their survival and proliferation. For example, Hepatitis C virus

(HCV) requires LDs during its proliferation [42]. HCV targets capsid core proteins to

LDs, as well as using the lipoprotein synthesis pathway to generate its own lipoviroparticles [43]. While many diverse LD functions have been identified, advances in LD biology are certain to reveal more.

1.4 Lipid Droplet Biogenesis

The biogenesis of LDs is poorly understood, while several models have been proposed the prevailing theory is the “budding” model (Figure 1-3). This theory hypothesises that LDs are synthesised between the two leaflets of the endoplasmic reticulum (ER)

[4, 44, 45]. The “hatching” model assumes that the LD forms by “hatching” once the lipid ester globule detaches from the ER along with the two membrane leaflets [3]. The

“vesicular budding” model proposes that LDs are originally formed within the vesicle bilayers. The alternative model suggests that the LD is cupped like an “egg” between two opposing ER membranes. The apparent correlation between LDs, the PAT proteins and ER membranes suggests that they may contribute to LD biogenesis. During the initial stages of LD biogenesis, it is assumed that the LD is too small to be seen with conventional microscopy. Thus, advanced high-resolution microscopy would be needed to detect any form of contact in the early stages of biogenesis.

Introduction

A B C

Figure (Text/Chart/Diagram etc.) has been removed due to Copyright restrictions

Figure 1-3. Proposed Models for Lipid Droplet Biogenesis. (Extracted from Ohsaki et al, 2008, BBA.) (A) The “budding” model proposes that LDs grow from the ER bilayer and eventually

bud off. (B) The “hatching” model suggests that newly synthesised lipids accumulate between the leaflets of the ER before being excised. (C) The “alternative” model in bacteria predicts that the wax ester/diacylglycerol transferase (WS/DGAT) is a TAG synthesising enzyme, which cups the lipid like an “egg” between two opposing membranes.

1.4.1 The “budding” model

The “budding model” suggests that newly synthesised lipid ester is accumulated in the membrane. As the lipid ester accumulates a certain amount of TAG, it reaches a

“threshold”. The lipid ester begins to isolate between the two membrane leaflets by phase separation and steadily projects towards the cytoplasm, eventually detaching from the mother membrane as a budding LD. Although this model offers a reasonable explanation on how a LD surrounded by a phospholipid monolayer originates from the bilayer, experimental evidence remains inadequate [4]. In cultured hepatocytes,

Ohsaki et al. recognised a structure that relates to the lipid ester globule between the

Introduction

two membrane leaflets of the ER [46]. Abnormal binding of lipidated apolipoprotein B to the ER membrane generated the structure. However, a comparable structure has never been observed in normal cells, possibly because the budding LD segregates from the ER membrane before it reaches a visible size. H-NMR and atomic force microscopy suggests the budding LD may be too small to record even by electron microscopy [17]

[3].

A study by Robenek et al., represents a considerable challenge to the prevailing hypothesis of LD biogenesis [47]. Robenek and colleagues observed that caveolin-1 is found in lumenal leaflets of ER membranes but not in the cytoplasmic leaflets. These findings are incompatible with the current hypothesis of LD biogenesis because it is believed that the cytoplasmic leaflet of the ER membrane is thought to cover the lipid ester globule and become the surface phospholipid monolayer of the cytoplasmic lipid droplet (CLD) [48].

1.4.2 The “hatching” model

The “hatching” model hypothesises that LDs form by “hatching”, where the lipid ester globule detaches as a bicellar structure, carrying both the cytoplasmic and lumenal membrane leaflets. In this model, a temporary pore is assembled in the ER membrane, created by the fusion of the cytoplasmic and lumenal leaflets. A LD would then be surrounded by a phospholipid monolayer, which in part, would be surrounded by the

ER-lumenal leaflet and its inserted proteins. The pore would then remove LDs from the

ER membrane [49]. A significant feature of this hypothesis is that the correlation of LDs with ER like membranes does not indicate the site of LD origin, but somewhat a means of LD growth through the transfer of LD-associated proteins [50]. Ducharme and Bickel 9

Introduction

hypothesised that the correlation of LDs with ER like membranes does not indicate the site of LD origin but somewhat a means of LD growth through the transfer of lipid droplet associated proteins [50].

In the proposed “budding” and “hatching” models, the lipid ester is supposed to project outwards as a globule within a reasonably small area of the membrane, however, it is not clear whether such an arrangement takes place based on the lipid properties alone. It is likely that certain protein-based mechanisms confine the lateral spreading of the lipid ester between the two membrane leaflets [3].

1.4.3 Alternative models

The alternative model was devised from the study of bacteria, where wax ester synthase/diacylglycerol acyltransferase (WS/DGAT) is accountable for TAG synthesis

[51]. The alternative model suggests that the LD is cupped like an “egg” between two opposing ER membranes rather than growing between the two leaflets. Lipids are transferred from the ER to the surface of the LD by adipophilin rich clusters. In contrast to the “budding” model, the LD envelope might not result from the budding portion of the ER membrane. The bacterial WS/DGAT shows no resemblance in sequence to that of diacylglycerol acyltransferases (DGATs) in eukaryotes, but the suggestion that the cytoplasmic LD is produced on the cytosolic surface of the ADRP- positive ER domain might be based on a similar extra-membrane mechanism [3, 47].

The challenge with this proposed model is that the formation of the phospholipid monolayer has yet to be explained.

Introduction

The “vesicular budding” model proposes that LDs are originally formed within the vesicle bilayers [52]. Here, a vesicle is formed and remains within close distance to the

ER membrane where neutral lipids may be packed into the vesicle [52]. The remaining lumen of the vesicle could fuse to the outer leaflet of the LD or stay within the LD.

1.5 The Regulation of Lipid Droplet Growth

1.5.1 Phospholipids

LD growth requires the accumulation of both phospholipids at the droplet surface and neutral lipids (TAG & CE) inside the lipid core. The most abundant phospholipids on the

LD surface are PC, a cylindrical shaped lipid, and PE, a conical shaped lipid. Their biosynthesis occurs primarily in the ER. Other phospholipids such as PA, PS, PG and PI are also found in smaller volumes.

A recent genome wide screen in the budding yeast, S. cerevisiae, identified 10

‘supersized’ LD mutants [53]. A common characteristic amongst these mutants was their increase in the level of PA, as well as their involvement in the metabolism of phospholipids. PA is the precursor to the de novo synthesis of all major phospholipids and TAG (Figure 1-2). Different PA subclasses may explain the diverse functions of PA.

Further investigation found that altered levels of PE and PA affected the growth and formation of SLDs. Upon treatment of ethanolamine, yeast mutants displayed an increase in the formation of SLDs. Moreover, mutants with elevated levels of both PE and PA had a greater number of SLD producing cells. As the surface of the LD contains

Introduction

a phospholipid monolayer, PA could alter the growth of LDs due to its shape, and induce membrane curvature [54].

A genomic screen in Drosophila S2 cells revealed that an increase in PC produced small

LDs [55]. The study by Krahmer et al., showed that PC is critical for the stabilisation of growing LDs, as well as the prevention of fusion [55]. The study also indicated that the assembly of LDs for lipid storage requires a particular increase in the synthesis of PC at the ER. Unlike most eukaryotes and higher mammals, PC synthesis in nematodes such as C. elegans is mediated through the phosphoethanolamine N-methyltransferase

(PEAMT) pathway. C. elegans contains two PEAMTs, PMT-1 and PMT-2, which are both involved in the synthesis of phosphocholine (PPC), the precursor to the synthesis of PC.

Although C. elegans lack adipose tissue, their main source of lipid storage is the intestinal cavity (also referred to as fat cavity/body). Down-regulation of pmt-1 led to the accumulation of large LDs in the intestinal cavity of the worm. This caused an increase in the total level of TAG and a decrease in PC [56]. Furthermore, it has been suggested that a decrease in the ratio of total surface phospholipids-to-neutral lipids could control the morphology of LDs through fusion (Figure 1-4). This fusion would carry on until the surface-to-volume ratio of LDs mirrored the phospholipid to TAG ratio. These findings indicate that PC, a potential regulator in LD growth, is involved in controlling the synthesis of TAG, a main component in the lipid core.

Introduction

Figure 1-4: The regulation of LD growth is mediated by phospholipids and proteins.

(Extracted from Yang et al, 2012, Curr Opin Cell Biol.) (Ai) LD delivery is targeted at the LD-ER contact site, where TAG accumulates, possible through FITM1/2. (Aii) TAG and CDP-Choline are synthesised on LDs from their respective synthesising enzymes. (Bi) Fsp27 enriched LD- contact sites facilitate the exchange of neutral lipids to regulate their size. (Bii) Phospholipids can facilitate rapid LD fusion. Under low PC and increased PA conditions, fusion may occur.

In mammals, PC in synthesised via two pathways: (1) The Kennedy (CDP-choline) pathway, and (2) The PEMT pathway. In agreement with the above findings, down- regulation of PEMT in 3T3-L1 adipocytes caused a significant decrease in the level of total TAG, as well as a reduction in PC [57]. The authors hypothesised that this might be due to either a defect in the synthesis of PC from PE via methylation, or, the utilisation of residual PC by PSS1. Furthermore, 3T3-L1 adipocytes with low PEMT expression presented decreased LD stability [57].

Introduction

In the glycerolipid synthesis pathway, AGPAT2 encoding 1-acyl-glycerol-3-phosphate

O-acytransferase-2, converts lysophosphatidic acid (LPA) to phosphatidic acid (PA)

[58]. Lipin, a phosphatidate phosphatase, utilises PA and converts it to diacylglycerol

(DAG), the precursor to the synthesis of TAG [58]. These key phospholipid enzymes have been implicated in generalised lipodystrophy, a genetic disease symbolised by a complete loss of adipose tissue [59, 60].

1.5.2 Proteins

Fld1p/Seipin

Two independent studies of the non-essential gene collection in yeast identified FLD1 as a ‘supersized’ LD mutant (SLD) [6, 61]. SLDs contain a lipid volume 30 times greater than that of the wild-type. Fluorescence microscopy revealed that fld1∆ cells not only displayed SLDs, but they contained densely packed (clustered) LDs in ~60% of the population. This led Fei et al. to reason that clustering is a requirement in LD fusion [5].

Upon treatment with cerulenin, fld1∆ cells displayed a significantly slower rate of TAG mobilisation than that of the wild-type, suggesting Fld1p functions as a regulator in LD growth. Microarray data indicated that INO1 and OPI3 (enzymes in phospholipid biosynthesis pathway of yeast) were the only transcripts to display a significant upregulation in fld1∆ cells. INO1 expression was derepressed when the concentration of

PA increased within the cell. Further analysis showed a substantial increase in PA isolated from the microsomes of the ER in fld1∆ cells. This data supports the notion

Introduction

that Fld1p could regulate the metabolism of phospholipids and neutral lipids required in LD growth.

The mammalian ortholog of Fld1p is seipin (BSCL2). In Drosophila, the absence of

Drosophila seipin (dSeipin) caused less TAG in the fat body and adipose tissue, which led to the accumulation of ectopic LDs in the salivary gland [62]. Lipidomic analysis revealed an increase in the total amount of unsaturated DAG and TAG species in the salivary gland. A potential role for dSeipin in the regulation of LD growth involves its function in lipogenesis. Genetic interactions concerning dSeipin and lipogenic genes such as DGAT indicate the contribution of dSeipin in the lipogenic pathway.

Furthermore, dSeipin has been implicated as a cofactor in the metabolism of PA.

In cultured mammalian cells, the knock down of seipin resulted in the accumulation of

LDs [63]. This finding was consistent with that of the fruit fly and yeast [6, 61, 62].

Similarly, the accumulation of oleate-induced TAGs led to an increase of LDs. In contrast, the overexpression of seipin caused an inhibition in the development of LDs.

Lipidomic analysis revealed a reduction in the level of TAG, indicating the seipin is a regulator in TAG metabolism. In vitro data revealed that seipin knock-down cells failed to maintain the expression of PPARγ, hence, inhibiting adipocyte differentiation [64].

Chan and colleagues showed that seipin -/- mice also displayed the evolutionarily conserved phenotype of SLDs, along with fewer LDs [65]. In addition, seipin -/- mice displayed severe generalised lipodystrophy [66]. This data indicates that seipin is a regulator in the formation of LDs.

Introduction

In humans, mutations of the gene which encode for seipin (BSCL2) are associated with

Berardinelli-Seip congenital lipodystrophy 2. BSCL is a human genetic disorder; it is distinguished by the almost complete absence of adipose tissue, and the accretion of ectopic TAG in both the liver and muscle. Along with BSCL2, AGPAT2 has also been implicated in the disorder [67].

Pah1p/Lipin

Lipins are phosphatidate phosphatases which converts PA to DAG. The yeast ortholog of lipin is Pah1, which has been shown to regulate the formation of LDs. The deletion of PAH1 produced a lower number of LDs and an accumulation of neutral lipids within the membrane. In addition, pah1∆ mutants displayed a reduction in the assembly of

LDs [68]. PAH1 contains a catalytically active PA phosphatase, which is required in LD formation. Adeyo et al. showed that the activator of Pah1p, a protein phosphatase,

Nem1p, localised near the LDs in approximately 80-85% of the cases. While the molecular mechanism underlying the growth of LDs in not fully understood, Goodman and colleagues provide an insightful model as to how PAH1 regulates LD formation.

Here, a LD is formed through the acquisition of DAG from Pah1p. The addition of DAG can aid in membrane curvature to promote the formation of LDs [68, 69].

In Drosophila, deletion of Drosophila lipin (dLipin) has been found to impair the development of the fat body, the key tissue in the fruit fly for TAG storage. dLipin mutants displayed smaller LDs and lower TAG levels, suggesting that the phospholipid to neutral lipid ratio had increased. In contrast, dLipin mutants displayed significantly

Introduction

larger fat body cells when compared to the control [70]. dLipin has been shown to be a gene which is essential in regulating TAG storage and controlling the size of LDs [71].

In agreement with the findings in Drosophila, lipin has also been found to be involved in TAG synthesis in C.elegans [72]. Unlike mammals, C. elegans has one lipin homolog, lipin-1. The knock-down of lipin-1 in worms resulted in a reduction in the accumulation of TAG, indicating that lipin-1 mutants developed a defect in lipid storage [72]. It is possible that lipin plays an essential role in the regulation of LD growth by controlling the total volume of TAG.

Unlike lower eukaryotes, higher organisms like mammals have 3 lipin genes. Lipin1 has been shown to be involved in the regulation of LDs through the accumulation of fat. In the mature adipocytes of transgenic lipin overexpression mice, a significant accumulation of fat was observed in the muscle specific tissues [73]. In contrast, lipin overexpression mice (with adipose tissue-specific expression) did not display an accumulation of TAG in any other tissues other than the adipose. Different levels of lipin expression in mice were found to alter genes which regulate TAG storage and fatty acid uptake [73]. In addition, the deletion of lipin-1 in mice causes lipodystrophy.

Chen et al. revealed that the loss of lipin1 did not impair the synthesis of hepatic TAG, possibly due to the high expression level of lipin2 [74]. Lipin-1γ is an isoform of lipin-1.

Recent studies have shown that upon oleate treatment, lipin-1γ localised to LDs [75].

Furthermore, lipin-1γ transfected COS-7 cells displayed small, clustered LDs with no change in total cellular TAG. This phenotype indicates that there is an increase in the surface area of LDs, which causes an increase in the phospholipid to neutral lipid ratio.

Introduction

Hence, lipin-1γ might function as a regulator of phospholipid synthesis by providing

DAG rather than TAG [75].

Perilipin1

The most widely studied protein on the surface of the LD is the phosphoprotein,

Perilipin 1. Perilipin 1 is a member of the PAT family of proteins. In Drosophila, two proteins have been identified with the same sequence homology to that of the PAT proteins, Lsd1 and Lsd2. In Drosophila, Lsd2 mutants exhibited a reduced level of TAG, as well as a deficiency in lipid storage [76]. Moreover, LSD2 has been found to be a regulator in the motor-driven movement of LDs in Drosophila embryos [77].

While Perilipin1 is known to be a regulator of lipolysis [78, 79], recent studies have emerged indicating a role in the expansion and regulation of LD growth. Perilipin A, the most abundant protein on the surface of the LD in adipocytes, has been shown to regulate the synthesis of TAG in LDs [80]. In adipocytes, as LDs expand and mature, the proteins which coat the surface of the LDs change. Upon increased TAG synthesis, perilipins 3-5 (encoded by TIP47, S3-12 & OXPAT) relocate onto the surface of the LD.

Perilipin 1 & perilipin 2 (ADRP) remain tightly packed at the LD surface [81] until perilipin2 is fully replaced by perilipin1. This indicates that perilipin1 is a potential cofactor involved in the regulation of LD growth. It is possible that during LD expansion, perilipin1 may be required to control the level of TAG. Alternatively, it may indirectly regulate LD growth by supporting LD expansion proteins/cofactors. In addition, mutations in perilipin have been found in patients with partial lipodystrophy

[82]. Although the mutated perilipin localised to the surface of the LD, the size of the

Introduction

LDs were much smaller. Adipose tissue from these patients revealed that the adipocytes were smaller in size, implying that mutations in perilipin have a defect in storing neutral lipids.

FSP27

Recent studies have begun to cast light on the importance of the regulation of LDs by identifying protein factors involved in LD expansion. The LD associated protein, Fat

Specific Protein of 27 kDa (FSP27), has emerged as a key regulator in the expansion of

LDs. FSP27 belongs to the cell death–inducing DFF45-like effector (CIDE) family, which are associated with obesity, type 2 diabetes and non-alcoholic fatty liver (NAFL) disease. FSP27, a novel regulator of LD growth, promotes the transfer and expansion of LDs at the LD contact site (LDCS) [83]. Enrichment of FSP27 at the LDCS could promote LD stability to allow for the efficient transfer of neutral lipids (TAG). Because

FSP27 localises to the surface of LDs, it may interact with other factors to distort the monolayer of phospholipids [83]. This distortion could activate the newly created LD contact site and direct lipid transfer proteins to allow lipid exchange, thereby regulating the growth of LDs through expansion/fusion. Similarly, Cidea, another LD associated protein identified in the CIDE family, has also been shown to regulate LD formation and reduce the hydrolysis of TAG [84, 85]. Cidea has previously been recognised as brown adipose tissue in mice, recent studies by Puri et al., showed that it is highly abundant in white adipose tissue and promotes the development of large LDs.

These results indicate the importance of the protein in the storage of neutral lipids.

Introduction

A common characteristic of the ob/ob mice is the development of non-alcoholic fatty liver disease. Studies have shown that the expression of Cidea and FSP27 are up- regulated in the liver, further implicating that the proteins may serve as regulators of

LD growth. The overexpression of FSP27 in the hepatocytes of the ob/ob mice showed an accumulation of TAG. In contrast, knock-down of FSP27 partially improved ob/ob liver steatosis [12]. Furthermore, mutations in FSP27 have been found in patients with partial lipodystrophy, fatty liver and many other human metabolic diseases. Mutations in FSP27 resulted in multilocular LDs, with an inability to increase LD size [86]. FSP27 has been implicated as a regulator in LD growth, required for the formation of unilocular LDs and lipid storage in white adipose tissue [86].

1.5.3 Other protein factors

LD Surface proteins- GPAT4/DGAT2

TAG synthesis enzymes such as GPAT4 have been implicated in the regulation of LD growth. Wilfling and colleagues identified two classes of LDs during oleate loading; small LDs which do not acquire TAG synthesising enzymes and remain small after their formation, and large LDs which require TAG synthesising enzymes to travel between the ER and LD [87]. Further analysis showed that GPAT4 relocalises to the ER domains during LD growth and to the phospholipid monolayer of mature LDs. The relocalisation of GPAT4 from the ER to the LD arises from ER bridges/tubules, potentially mediating

LD growth by producing TAG on LDs. Wilfling et al further identified that the function of GPAT4 in LD growth is an evolutionarily conserved process.

Introduction

FIT1 & FIT2

From lower eukaryotes to higher mammals, the fat storage-inducing transmembrane proteins (FIT) are evolutionarily conserved [88]. While FIT1 is expressed in the skeletal muscle and heart, FIT2 is ubiquitously expressed at low levels in both human and mouse tissue [89]. The FIT proteins have been shown to directly bind to DAG and TAG.

The binding activity is strongly correlated with LD size [88]. How the proteins form expanding LDs remains unknown.

SNARES

It has been reported that SNARE proteins facilitate the fusion of LDs. SNAP23, syntaxin-

5 and VAMP4 were found to localise on LDs along with NSF and α-SNAP. SNARE proteins were present during the fusion between LDs and are believed to be regulators of LD growth [90]. Furthermore, the GTPase ADP-ribosylation factor related protein 1

(ARFRP1) has been revealed as an essential regulator in LD growth [91]. Arfpr1 -/- mice displayed smaller LDs with altered levels of LD protein composition than that of the control. Arfpr1 deficient MEFS were unable to form normal LDs [91]. microRNAs

From yeast to mammals, phospholipid and protein cofactors have been associated with LD growth. A recent study on microRNA (miRNA) target analysis revealed that miRNAs control LD growth and formation in hepatocytes. Alterations in the miR-181 family were found to affect intracellular lipid content by reducing TAG levels of up to

60%. Of the miRNAs identified, 3 of the 11 were found to be altered in non-alcoholic

Introduction

fatty liver diease. The findings by McDonough and colleagues strongly suggest that miRNAs have a role in lipid metabolism [92].

1.6 Adipogenesis (adipocyte differentiation) at the Systemic Level

The development of adipocytes from mesenchymal stem cells involves a highly coordinated series of gene expression activation [93]. While the main transcription factors have been known for some time, recent studies have emerged with new cofactors and regulators that are involved in adipogenesis. After proliferation of preadipocytes, adipocyte differentiation is promoted by several families of adipogenic markers. Two members of the CCAAT-enhancer binding protein (C/EBP) family, C/EBP-

β and C/EBP-δ are first expressed [94]. This increase in expression leads to the activation of peroxisome-proliferator activated receptor-γ (PPAR-γ[95], which forms a heterodimer with retinoid X receptor (RXR)[96-98]. PPARγ in turn activates C/EBP-α expression [99, 100] (Figure 1-5). The sterol responsive element binding protein 1c

(also named adipocyte determination and differentiation factor 1; ADD1/SREBP1) is able to improve the transcriptional activity of PPAR-γ [101]. During adipogenesis,

PPARγ and C/EBP-α transactivate a subset of genes which give rise to the adipocyte phenotype. This is signified by the appearance of triglyceride accumulation [102].

Model cell lines have proven to be an invaluable tool in studying the process of adipocyte differentiation. Preadipocyte cells lines such as 3T3-L1 cells have become the most reliable system in replicating the differentiation process [103, 104]. Once preadipocytes reach growth arrest, a “cocktail” of inducing agents is added. The induction activates the cAMP response element binding protein (CREB) which becomes

Introduction

phosphorylated and eventually stimulates the expression of C/EBPβ, triggering the adipogenic signalling cascade [105, 106].

Figure 1-5. Adipocyte Differentiation. The master regulator of adipogenesis, PPARγ, is activated through the expression of C/EBPβ and C/EBPδ. PPARγ forms a heterodimer with RXR to main and induce differentiation. The expression of PPARγ activates C/EBPα which maintains a positive feedback loop to induce adipogenesis. SREBP1c helps generate a ligand to increase the transcription activity of PPARγ along with FOXC2.

Recent studies have shed light on the importance of lipid intermediates as regulators of gene expression during adipogenesis [107]. Lipin-1 has been implicated during the early stages of adipogenesis and has been shown to be a transcriptional activator of

PGC-1α, a co-activator of PPARγ. Zhang et al showed that the PAP activity of lipin-1, but not the co-activator of PPARγ, was sufficient enough to rescue the expression of

PPARγ during adipogenesis [108]. Similarly, Stapleton et al demonstrated that the

Introduction

overexpression of GPAT1 in Chinese Hamster Ovary (CHO) cells resulted in an increase in the level of LPA, which increased the activity of PPARγ [109]. Conversely, the overexpression of AGPAT2 decreased the activity of PPARγ. Through the manipulation of endogenous LPA and PA levels, these studies have indicated that PA has an inhibitory effect on the expression levels of PPARγ during adipogenesis and may act as a possible antagonist.

1.6.1 Peroxisome Proliferator-Activated Receptor γ (PPARγ)

The master regulator of adipogenesis, PPARγ, is a ligand-activated transcription factor of the nuclear hormone receptor superfamily [110-113]. Through alternative promoter usage and alternative splicing, three isoforms of PPARγ exist [114]. While PPARγ1 is found in many tissues, PPARγ2 is exclusively found in brown and white adipose tissue

[115]. The expression of PPARγ alone is sufficient enough to induce differentiation

[111]. Experimental evidence supports the notion that PPARγ activation in adipose tissue could increase insulin sensitivity. Activation of PPARγ in adipocytes results in the upregulation of genes that stimulate free fatty acid (FFA) release. PPARγ ligands such as thiazolidinediones (TZDs) are used in the treatment of type II diabetes [116].

1.6.2 CCAAT/ enhancer binding protein family (C/EBP)

C/EBPs belong to the family of basic-leucine zipper transcription factors. Three isoforms are associated in adipogenesis; C/EBPα, C/EBPβ and C/EBPδ [117]. Following the induction of preadipocytes, C/EBPβ and C/EBPδ are the first transcription factors activated, which stimulate PPARγ to maintain a positive feedback loop with C/EBPα.

Introduction

Similar to PPARγ, C/EBPα transactivates the expression of lipogenic, lipolytic and insulin sensitive genes, including; GLUT4, aP2, AGAPT2, leptin and perilipin [100].

1.7 Lipolysis

The process of lipolysis in adipocytes involves the hydrolysis of TAG to FFAs and glycerol [118]. Thus far, three enzymes have been implicated in the catabolism of TAG; adipose-triglyceride lipase (ATGL), hormone-sensitive lipase (HSL) and monoglyceride lipase (MGL) [119, 120]. Fatty acids (FAs) are vital as substrates for lipid synthesis and energy production. Regardless of their biological importance, too much FA is damaging at the cellular level. The highly regulated process of lipogenesis provides an effective system by removing FAs when they are not required and supplying FAs in times of need.

ATGL performs the rate-limiting step of hydrolysing TAGs to generate DAG (Figure 1-6).

It was first described by three separate groups as a lipolytic enzyme, which belongs to the patatin domain-containing proteins [119, 121, 122]. While ATGL can be found in many tissues, BAT and WAT display the highest mRNA expression levels. Studies have shown that in adipocytes the activity of ATGL and HSL is accountable for ~90% of the lipolytic activity [123]. During lipolysis, ATGL requires a coactivator, comparative gene identification-58 (CGI-58) for complete hydrolysis [124].

Unlike HSL, ATGL-deficient mice are unable to hydrolyse TAGs effectively, resulting in severely reduced lipolysis and an increase in overall lipid accumulation [125]. Similarly,

ATGL mutations are correlated with TAG accumulation and excessive accumulation of

Introduction

fat in the heart causing cardiac myopathy [126]. These findings emphasise the importance of ATGL as a regulator in TAG catabolism.

Figure 1-6. Lipolysis. During lipolysis, ATGL requires its coactivator, CGI-58 to hydrolyse TAG and generate DAG and free fatty acid. In DAG catabolism, HSL is phosphorylated by cAMP-

dependent PKA on six consensus serine residue. Phosphorylated HSL interacts with perilipin-1 to produce MAG and free fatty acid. MGL catalyses the final step in lipolysis, converting MAG to glycerol and free fatty acid.

HSL is an enzyme of dual-purpose capabilities; hydrolysing TAG and DAG. During TAG hydrolysis, HSL is the rate-limiting enzyme in DAG catabolism [127]. Similar to ATGL,

HSL exhibits the highest expression levels in white and brown adipose tissue. In adipocytes, HSL activity is strongly regulated by two separate processes upon β- adrenergic stimulation [128, 129]. Firstly, unlike ATGL, HSL is phosphorylated by cAMP- dependent PKA on six consensus serine residues [130, 131] and secondly, phosphorylated HSL interacts with perilipin-1 [132]. Recent studies have shown that the phosphorylation at Ser650 and Ser660 are particularly important for increased hydrolytic activity [133]. Mice deficient in perilipin-1 are unable to interact with HSL at

Introduction

the LD surface, preventing the induction of lipolysis [25]. While in non-adipose tissues, the increase in insulin levels prevent the phosphorylation of HSL and perilipin, reducing the activity HSL and ATGL [134].

Lipin1 -/- mice display reduced lipolytic activity under basal and isoproterenol-treated conditions in adipocytes. As well as a reduction in the level of ATGL, HSL and p-PKA, the level of LPA and PA increased in the WAT [135].

Recent studies of lipolysis have identified many regulators which depict a more complex process than previously thought. The role of phosphorylation on enzyme activity, along with the factors which regulate the lipolysis pathway during hormonal stimulation need to be further investigated. Also, understanding the involvement of

PA accumulation during lipolysis will provide insight into the role of phospholipids and lipolytic regulators in human metabolic diseases.

1.8 Mammalian Target of Rapamycin (mTOR) Pathway

The mTOR pathway is a complex network that regulates an array of cellular processes, from protein and lipid synthesis to energy metabolism and cell survival. mTOR signalling has been implicated in metabolic diseases, as well as LD associated processes such as adipogenesis and lipolysis. mTOR is a serine/threonine kinase which is evolutionarily conserved [136]. It belongs to the phosphoinositide3-kinase (PI3K)- related kinase family and interacts with various proteins to forms two complexes; mTORC1 and mTORC2. Both complexes share the mTOR subunit and the mammalian lethal with sec-13 protein 8 (mLST8, also referred to as GβL) [137]. The complete unit of mTORC1 also consists of the regulatory associated protein of mammalian target of

Introduction

rapamycin (raptor). In contrast, the complete subunit of mTORC2 comprises of rapamycin-insensitive companion of mTOR (rictor) [138].

Of the two complexes, mTORC1 is better characterised. Through the phosphorylation of substrate 6 kinase 1(S6K1) and the eukaryotic translation initiation factor 4E-binding protein 1 (4E-BP1), mTORC1 controls a variety of biological processes such as protein and lipid synthesis, autophagy, lysosome biogenesis and energy metabolism [139-141]

(Figure 1-7). mTORC2 regulates cell survival and cytoskeletal organization through the phosphorylation of kinases such as Akt [142, 143].

Since the mTOR pathway responds to high levels of nutrients, such as lipid uptake in the adipose tissue, its role in controlling metabolic diseases has been of interest.

Studies have shown that the down-regulation of mTORC1 inhibits adipogenesis, while the overexpression of mTORC1 promotes adipogenesis [144, 145]. Mice lacking adipose-specific mTORC1 display a lean phenotype and are impervious to obesity under high-fat-diet conditions [146]. In contrast, mTORC2 adipose-specific depleted mice exhibit the wild-type fat phenotype [147]. Furthermore, S6K1 inhibition of insulin signalling results in increased insulin resistance in the adipose tissue. Loss of mTORC1 in the muscle of mice increases the activation of Akt, which promotes glycogen accumulation, while loss of mTORC2 causes minor systemic glucose intolerance.

Recent studies by several groups have shown that the activation of mTORC1 promotes lipogenesis through the regulation of SREBP1c [148-150]. mTORC1-mediated down- regulation of ATGL is one way of lipolysis regulation [151].

Introduction

In order to maintain membrane synthesis and stimulate cell growth and proliferation, studies have shown that PA is required in mTORC1 and mTORC2 stability [152, 153].

Although mTORC1 is more sensitive to PA and the changes in PA level, it is believed that rapamycin and PA compete with one another to form a stable complex with mTOR. The most commonly derived PA involved in mTOR regulation is from the synthesis of phospholipase D (PLD), an enzyme which hydrolyses PC to PA [154, 155].

While mTOR -/- mice are embryonically lethal, PLD1 -/- mice are viable, indicating that

PA generated from other lipid-synthesising enzymes are also necessary for PA/mTOR stability.

Figure 1-7. The mTOR signalling pathway. The rapamycin-sensitive mTORC1 contains the mTOR kinase along with mLST8 and raptor, while mTORC2 comprises mainly of the mTOR

kinase, mSin, mLST8 and rictor. The activation of mTORC1 results in the phosphorylation of S6K1, which inhibits IRS and down-regulates PI3K/Akt. Conversely, mTORC2 phosphorylates and activates Akt in response to IRS. Both mTORC1 and mTORC2 are activated by growth factors, while nutrient availability also activates mTORC1. 29

Introduction

Mitra et al identified that the dephosphorylated pool of PA in Lipin1 -/- mice could also regulate mTOR activity. The deletion of Lipin1 increased the phosphorylation of mTOR at Ser2448 as well as S6K. In lipin1 knock-down 3T3L1 adipocytes, the treatment of

Torin (mTOR inhibitor) increased PDE activity (phosphodiesterase enzymes which regulate the signalling of PKA in adipocytes) [135]. This is consistent with the idea that

PDE activity is affected by the level of PA, through the mTOR machinery.

1.9 CDP-DAG synthases

To identify novel gene products regulating the cellular dynamics of LDs, genome-wide screens were carried out in the budding yeast, S. cerevisiae. A previous genome-wide screen identified Fld1p (yeast homologue of human seipin) as a major regulator of LD dynamics. Fld1p has been implicated in the metabolism of fatty acids and phospholipids [6, 53, 61], as well as in TAG synthesis. Importantly, seipin has also been shown to be involved adipocyte differentiation [63, 156]. Therefore, seipin can regulate both cellular and systemic lipid storage. Subsequent genome-wide screens showed that knocking-down CDP-diacylglycerol synthase 1 (CDS1) formed “supersized”

LDs in yeast.

CDS1 is an essential gene which is evolutionarily conserved from bacteria to humans

[157]. In mammals, there are two CDP-diacylglycerol synthases (CDS, also known as

CTP-phosphatidic acid cytidyltransferase), CDS1 and CDS2 [158, 159]. In higher eukaryotes, CDS has been associated with eye disease and angiogenesis (the development of new blood vessels from pre-existing ones). In Drosophila, CDS has been implicated in phototransduction (light excitation in photoreceptors), a process mediated by the phosphoinositide signalling cascade. Wu et al., showed that cds

Introduction

mutants displayed light dependent retinal degeneration [160, 161]. Additionally, Pan et al., revealed that CDS2 is vital for VEGFA-induced angiogenesis. The down-regulation of CDS2 resulted in the defective development of endothelial cells and arterial differentiation [162].

In humans, CDS1 is predicted to have 7 transmembrane domains, while CDS2 is anticipated to have 8 transmembrane domains. CDS1 and CDS2 catalyse the formation of CDP-DAG by utilising PA. In this reaction, CTP and phosphate are converted to CDP and diphosphate to produce CDP-DAG [163] (Figure 1-8). CDP-DAG is believed to be the rate-limiting intermediate for downstream lipid synthesis. Cds1p and Cds2p localise to two subcellular domains, the inner mitochondrial membrane, where it has been implicated in the synthesis of phosphatidylglycerol (PG) and cardiolipin (CL) and the endoplasmic reticulum, where it is involved in the synthesis of phosphatidylinositol

(PI) [164, 165]. A recent study in yeast showed that CDS1 solely resides in the ER, and that another CDP-DAG synthase, Tam41, exists in the mitochondria, required for the synthesis of cardiolipin [166]. Although the biochemical function of CDS1 and CDS2 has been characterised, little is known about their involvement during LD formation and adipocyte differentiation.

Figure 1-8. The Role of CDP-DAG Synthase in Lipid Synthesis. CDS1/2 converts CTP and phosphate to CDP-diphosphate to catalyse PA to CDP-DAG. CDP-DAG is believed to be the rate-limiting lipid intermediate for the synthesis of PG, CL and PI.

Introduction

1.10 Lipid Droplets in Prevailing Diseases

Changes in the cellular dynamics of lipid droplets (LDs) are associated with human metabolic disorders although the mechanisms remain unknown. Therefore, it is necessary to understand the molecular mechanisms governing LD biogenesis and growth. Defects in the accumulation of LDs are associated with many human metabolic disorders such as obesity, type II diabetes and atherosclerosis.

While the accumulation of cholesteryl ester in LDs has been associated with foam cell formation in atherosclerosis, the accumulation of triacylglycerol in LDs has been linked to obesity, non-alcoholic fatty liver disease and type II diabetes. The accumulation of lipids in the muscle [167] as well as the liver [168, 169] is highly related with the development of insulin resistance and type 2 diabetes (for review see [170]). The accumulation of neutral lipids, particularly cholesterol esters in the arterial wall can cause atherosclerosis. Bostrom et al. showed that when muscle cells are incubated with fatty acids, an increase in LDs and insulin resistance arises [19]. The development of insulin resistance is a key variable in developing premature atherosclerosis [171].

Lipodystrophy is the illness described by the partial or complete loss of adipose tissue.

It has been connected to many loci including, Berardinelli-Seip congenital lipodystrophy

2 (BSCL2) [172, 173]. Also, mutations in ATGL and CGI-58 have been shown to cause neutral lipid storage disease (NLSD) [174]. An autosomal recessive disease which is characterised by the extreme accumulation of neutral lipids in a variety of tissues.

Introduction

1.11 Summary

Although recent advances in LD biology have provided some insight into how LD growth is regulated, many questions remain unanswered. For example, what controls the directed transfer of neutral lipids, TAG and CE, to the LDs? Of all the factors regulating LD growth, the level of phospholipids, particularly PA and PC, appear to be evolutionarily conserved from lower eukaryotes to higher mammals. Are there protein factors which modify the phospholipid monolayer to regulate LD growth? If so, are they conserved between all species? Furthermore, PA also appears to be a major contributor in a number of cellular processes such as adipogenesis, lipolysis and mTOR signalling. Whether PA has tissue specific roles would need to be further investigated.

In addition, PA derived from the glycerolipid synthesis pathway and PA derived from

PC hydrolysis may have different roles in biological processes. Future research should help answer these questions.

Introduction

1.12 Aims

CDP-DAG synthases catalyse the formation of CDP-DAG from PA. By manipulating the expression levels of CDS1 and CDS2, the level of endogenous PA was modulated. This study hypothesised that PA promotes the formation and fusion of LDs, and that the accumulation of PA in preadipocytes would have a negative effect on adipocyte differentiation. Furthermore, the study also hypothesised that the accumulation of PA resulted from the down-regulation of CDP-DAG synthases, which would have an effect on lipolysis and the mTOR signalling pathway.

The central aim underlying this project was to characterise CDP-DAG synthases in LD dynamics and adipocyte development.

 Aim 1: To examine the role of CDP-DAG synthases in lipid droplet dynamics

Inhibiting the expression of CDS1 in yeast cells produces supersized LD

mutants. To observe whether the ‘supersized’ phenotype is evolutionarily

conserved, the down-regulation of CDS1 and CDS2 in the mammalian system

was investigated (Chapter 3). HeLa cells and 3T3-L1 preadipocytes were used to

explore the role of CDS1 and CDS2 in lipid droplet dynamics.

 Aim 2: To examine the role of CDP-DAG synthases in adipocyte development

Previous data has shown that the down-regulation of CDS1 in yeast causes

significant accumulation of PA. If PA does inhibit adipogenesis as hypothesised,

then inhibiting CDS1 and CDS2 would result in a negative effect on adipocyte

differentiation. For the purpose of the experiment, 3T3-L1 preadipocytes were

used as an in vitro model of adipogenesis (Chapter 3).

Introduction

 Aim 3: To identify if CDP-DAG synthases are involved lipolysis and mTOR

signalling

PA has been implicated in both lipolysis and mTOR signalling. By down-

regulating the expression of CDS1 and CDS2 in mature adipocytes, the

expression level of lipolytic and mTOR associated proteins was examined

(Chapter 4).

 Aim 4: To investigate whether and how the interaction between CDS1 and FLD1

occurs in the phospholipid biosynthesis pathway

Previous genome wide screens of the yeast gene collection identified CDS1 and

FLD1 as the only ‘supersized’ LD mutants in enriched (YPD) media. Protein-

protein interactions were employed to assess whether the interaction of Cds1p

and Fld1p was functional or physical (Chapter 5).

Materials and Methods

2. MATERIALS AND METHODS

2.1 General Materials and Methods

Commercial suppliers are listed in alphabetical order along with the materials purchased. Sequences of commercially purchased siRNAs are also listed in Table 1-1.

Table 1-1. Materials used in this study. Materials Supplier/Description

RFP Antibody Abcam

Actin Antibody Abcam

CDS1 Antibody Abcam

Perilipin Antibody Abcam

Acetic Acid Glacial Ajax FineChem

Ethylenediaminetetraacetic Acid (EDTA) Ajax FineChem

Methanol Ajax FineChem

N-Hexane Ajax FineChem

Potassium Chloride (Kcl) Ajax FineChem

Chloroform Ajax FineChem

Betafluor Ajax FineChem

Ethanol Absolute Ajax FineChem

D-Glucose Anhydrous Ajax FineChem

Diethyl Ether Ajax FineChem

Materials and Methods

Ethanol Absolute Ajax FineChem

Tris (Hydroxymethyl) Aminomethane (Tris) Ajax FineChem

Sodium Hydroxide (Naoh) Ajax FineChem

Sodium Chloride (Nacl) Ajax FineChem

Hydrochloric Acid (HCL) Ajax FineChem

Isopropanol Ajax FineChem

Phospholipid Internal Standard Mix Avanti Polar Lipids

Bacto Tm Yeast Extract B.D. Becton, Dickinson + company

Yeast Nitrogen Base Without Amino Acids And B.D. Becton, Dickinson + company

Ammonium Sulfate (YNB W/O A.A +

(NH4)2SO4)

Free Glycerol Assay Biovision

HSL Antibody Cell Signaling

Phospho-HSL Antibody Cell Signaling

ATGL Antibody Cell Signaling mTOR Antibody Cell Signaling

Phospho-mTOR Antibody Cell Signaling

S6k Antibody Cell Signaling

Phospho-S6k Antibody Cell Signaling

Akt Antibody Cell Signaling

Phospho-Akt Antibody Cell Signaling

DUAL membrane kit DualsystemsBiotech

Tryptone Fluka

Materials and Methods

KAPA SYBR® FAST qPCR Kits KapaBiosystems

Dynabeads® Magnetic Beads Life Technologies

CellLight® Mitochondria-RFP BacMam 2.0 Life Technologies

Dulbecco’s Modified Eagle Medium (DMEM) Life Technologies

Fetal Bovine Serum (FBS) Life Technologies

Neonatal Calf Serum (NCS) Life Technologies

Penicillin-Streptomycin (10,000 U/mL) Life Technologies

OPTI-MEM Life Technologies

TrypLE™-Express Life Technologies

D-PBS Life Technologies

ProLong® Antifade Gold Reagent Life Technologies

Alexa Fluor Antibodies Life Technologies

TRIzol® TM Reagent Life Technologies (Invitrogen)

Lipofectamine RNAiMAX® Reagent Life Technologies (Invitrogen)

Lipofectamine® LTX with PLUS™ Reagent Life Technologies (Invitrogen)

Lipofectamine® 2000 Reagent Life Technologies (Invitrogen)

DECP treated water – DNAse/ RNAse free Life Technologies (Invitrogen)

SuperScript® VILO™ cDNA Synthesis Kit Life Technologies (Invitrogen)

BODIPY 493/503 Life Technologies (Invitrogen)

PCR purelink purification kit Life Technologies (Invitrogen)

Plasmid Mini-Prep Kit Life Technologies (Invitrogen)

Puromycin Life Technologies (Invitrogen)

TLC Silica Gel Glass Plates Merck

Materials and Methods

TLC aluminium sheet Merck

Oleic acid [1-14C] MP Biomedicals

CYTIDINE 5′-TRIPHOSPHATE (CTP) [5-3H] MP Biomedicals

Restriction Enzymes New England Biolabs

KOD Hot Start DNA polymerase Novagen

Bacteriological Peptone Oxoid

All amino acids purchased in powder form Sigma Aldrich

Flag Antibody Sigma-Aldrich

Human CDS1 siRNA Sigma-Aldrich

5’- 3’ CUCACUUCCACCCUUUCUA

Human CDS2 siRNA Sigma-Aldrich

5’- 3’ CUUGUUAUCCACAACCUAU

Mouse CDS1 siRNA Sigma-Aldrich

5’- 3’ CAGGUUUGACUGUCAGUAU

Mouse CDS2 siRNA Sigma-Aldrich

5’- 3’ GAUUAAGUGUUUCCAUGAA

Human LPIN1 siRNA Sigma-Aldrich

5’- 3’ GCAUGAAUCAUCCUCCAGU

Mouse LPIN1 siRNA Sigma-Aldrich

5’- 3’ GAGACAACGGAGAAGCAU

Isoproterenol Sigma-Aldrich

Bovine Serum Albumin (BSA) Sigma-Aldrich

Materials and Methods

Oleic Acid >99% GC Sigma-Aldrich

Insulin Solution 10mg/mL Sigma-Aldrich

3-Isobutyl-1methylxanthine ≥99% (HPLC), Sigma-Aldrich

powder

Dexamethasone Sigma-Aldrich

Nile Red Sigma-Aldrich

Bicinchoninic Acid (BCA) Kit for Protein Sigma-Aldrich

Determination

Iodine Sigma-Aldrich

Triacylglycerol + Sterol Ester Standard Sigma-Aldrich

Doxycycline Sigma-Aldrich

Ampicillin Sigma-Aldrich

Kanamycin Sigma-Aldrich

Phospho-perilipin Antibody Vala Sciences

2.2 General Buffers

Unless stated otherwise, all buffers used in this study were made up with MilliQ water

(MilliQ system, Millipore NSW). All solutions were either sterilised by filtration, through a 0.22µm filter, or, by autoclaving for 15m at 121°C. pH calibration was carried out using HCl or NaOH. Prior to disposal, all biological waste was either autoclaved or bleached.

Materials and Methods

2.3 qRT-PCR Primer Design cDNA sequences were obtained from http://www.ncbi.nlm.nih.gov/gene (NCBI). qRT-

PCR primers were designed using the software, Primer 3.0. Primer sequences are listed in Table 1-2, in alphabetical order.

Table 1-2. Nucleotide sequence of qRT-PCR primers designed in this study. Primer (qRT-PCR) Sequence

36B4 5’- CCCACTTACTGAAAAGGTCA

3’- TTAGTCGAAGAGACCGAATC

aP2 (mouse) 5’- ACATGAAAGAAGTGGGAGTG

3’- GGTTATGATGCTCTTCACCT

ATGL (mouse) 5’- TGTGGCCTCATTCCTCCTAC

3’- TCGTGGATGTTGGTGGAGCT

BSCL2 (human) 5’- CTCCTGCTATTTGGCTTTGC

3’- GCTGAGGAAGGTGAAGTTGC

BSCL2 (mouse) 5’- GACCAGATCAAAGGA

3’- AAGGATGGTGCAGAAGAGC

C/EBPα (mouse) 5’- CAAGAACAGCAACGAGTACC

Materials and Methods

3’- TTGACCAAGGAGCTCTCAG

Caspase3 (mouse) 5’- ACTTCCATAAGAGCACTGCA

3’- ACCATGGCTTAGAATCACAC

CDS1 (human) 5’- CCTTGTCATCCAAAATCTGT

3’- GAATCCTTCCCAAGTCTTTT

CDS1 (mouse) 5’- CGCACGTCACTTTACTGATA

3’- GAATCCTTCCCAAGTCTTCT

CDS2 (human) 5’- GGCTACAACGTCTACCACTC

3’- GAACCGGTGGTATTTACTGA

CDS2 (mouse) 5’- TGGTTTATTGTTCCCATCTC

3’- AGCATCTGTACCCAGACATC

GAPDH 5’- AGAAGGCTGGGGCTCATTTG

3’- AGGGGCCATCCACAGTCTTC

HSL (mouse) 5’- CTGAACAGTCAGACAAGCAA

3’- CTGTGACCCACTCAGAAAGT

LPIN1 (human) 5’- CGACCTTCAACACCTAAAAG

Materials and Methods

3’- AGTGTCTGAAGATTCGCTGT

LPIN1 (mouse) 5’- CCAGAATGGCTACAAGTTTC

3’- CTTTTCTGGCTTCTTTTCAA

PPARγ (mouse) 5’- ACCACAGTTGATTTCTCCAG

3’- TAGAGCTGGGTCTTTTCAGA

Pref1 (mouse) 5’- CCATGAAAGAGCTCAACAAG

3’- TACTGCAACAGGAGGTTCTT

β-Actin (human) 5’- AGCGAGCATCCCCCAAAGTT

3’- GGGCACGAAGGCTCATCATT

2.4 Plasmids

The plasmids listed in Table 1-3 were gifts from collaborators and peers.

Table 1-3. Plasmids used in this study that were obtained as gifs. Plasmids Purchased

BFP-Rab10 Gift from Gia Voeltz

CDS1-flag (PPJ25) Gift from Suzanne Jackowski

Materials and Methods

DsRed-ER Commercially purchased

GFP-PIS Gift from Gia Voeltz

mCH-CEPT1 Gift from Gia Voeltz

mCH-PIS Gift from Gia Voeltz

mCH-Rab10 Gift from Gia Voeltz

pCI-neo-flag-hSeipin Designed by Weihua Fei

pEGFP-mSeipin Designed by Weihua Fei

yEp-FLD1-GFP Designed by Weihua Fei

pmCherry-hSeipin Designed by Weihua Fei

pmGFP-2XPASS Gift from Guangwei Du

pmGFP-PASS Gift from Guangwei Du

pYEX4T1-FLD1 Designed by Weihua Fei

RFP-PASS Gift from Guangwei Du

2.4.1 Vector Construction cDNA sequences were retrieved from NCBI and Yeast Genome Database

(http://www.yeastgenome.org/).

Materials and Methods

Primers were designed without a stop codon and were in-frame of the multiple cloning site (MCS). DNA was amplified by polymerase chain reaction (PCR) using KOD Hot Start

DNA polymerase. PCR parameters were; one cycle at 95°C for 2m, followed by 40 cycles of denaturation at 95°C for 20s, annealing at the lowest primer Tm°C for 10s

(usually 60°C) and extension at 70°C between 45-75s (depending on target size). DNA fragments were confirmed by agarose gel and purified using the PCR purification kit.

Purified PCR products and vectors were digested with the required restriction enzymes and ligated at a vector to insert ratio of 1:3. Ligated products were sub-cloned into E. coli DH5α for plasmid amplification and grown on antibiotic sensitive LB (Luria-Bertani)

(1% (w/v) Bacto tryptone, 0.5% (w/v) yeast extract, 0.5% (w/v) NaCl) plates. Positive constructs were then grown in LB broth with antibiotics and purified using the plasmid mini-prep purification kit. Primer sequences for constructs designed in this study have been listed in Table 1-4. Underlined nucleotides represent the overhang sequence, nucleotides in red characterise the restriction enzyme used and nucleotides in black represent the target gene sequence.

Table 1-4. Primer sequences of designed constructs Primers Sequence

CDS1-myc 5’- TAGATGGATCCATGTCTGACAACCCTGAGATGAA

3’- TAGATGTCGACAGAGTGATTGGTCAATGATTTC

mCherry-N1-hCDS1 5’- TAGATAGATCTATGTTGGAGCTGAGGCACCGG

Materials and Methods

3’- TAGATCCGCGGTACCTTCAAGGTGGGTTGTAGGAT mCherry-N1-hCDS2 5’- TAGATAGATCTATGACAGAGCTGAGGCAGAGG

3’- TAGATCCGCGGCTCGTCCTCTGTGGTGGATGTCAG mCherry-N1-mCDS1 5’- TAGATAGATCTATGCTGGAGCTGCGGCACCGC

3’- TAGATCCGCGGCACCTTCAAGGTGGGCTGCAGGA mCherry-N1-mCDS2 5’- TAGATAGATCTATGACCGAACTACGGCAGAGG

3’- TAGATCCGCGGCTCATCTTCCAAGGCAGATGT pPR3N-CDS1 5’-

TAGATGGCCATTACGGCCATGTCTGACAACCCTGAGATGAAACC

3’- TAGATGGCCGAGGCGGCCTCAAGAGTGATTGGTCAATGA

PYEX4T1-CDS1 5’- TAGATGAATTCATGTCTGACAACCCTGAGATGAAACC

3’- TAGATCTCGAGTCAAGAGTGATTGGTCAATGA yEp-CDS1-GFP 5’- TAGATAAGCTTGGCAATGGAGCTGTTAGATGC

3’- TAGATTCTAGAAGAGTGATTGGTCAATGA

Materials and Methods

2.5 Mammalian Cell Culture

HeLa cells were grown in Dulbecco’s Modified Eagle Medium (DMEM) with 10% Fetal

Bovine Serum (FBS) and 1% Penicillin/Streptomycin (P/S). Cells were maintained at

37°C with 5% CO2 where the media was refreshed every two days. 3T3-L1 preadipocytes were grown in DMEM supplemented with 10% Newborn Calf Serum

(NCS) and 1% P/S before adipocyte differentiated was induced. Similar to HeLa cells,

3T3-L1 preadipocytes were maintained at 37°C with 5% CO2 where the media was refreshed every two days.

2.5.1 Adipocyte Differentiation

To induce adipocyte differentiation, 3T3-L1 preadipocytes were grown in DMEM containing 10% NCS and 1% Penicillin/Streptomycin. Two days post confluence, cells were induced in DMEM/FBS/PS supplemented with insulin (5 μg/ ml), dexamethasone

(1 μM) and isobutylmethylxanthine (IBMX) (0.5 mM) (day 0). Two days later, the medium was changed to DMEM/FBS/PS supplemented with insulin (5 μg/ ml) (day 2).

An additional two days later, the medium was changed to DMEM/FBS/PS (day 4).

Media was changed every two days until the desired day [175]. Quantitative real-time

PCR (qRT-PCR) was carried out in cells 6 days after differentiation. For adipocyte experiments, 3T3-L1 preadipocytes were differentiated up to day 8 on a 15cm dish.

3T3-L1 adipocytes were split by adding 2ml of trypsin (TrypLE) to the cultured dish, removing 1.5ml of TrypLE and incubating the cells at 37°C with 5% CO2 for 3m. Pre- warmed collagenase was used to detach the adipocytes from the dish. Adipocytes

Materials and Methods

were spun down at 400x g for 5m and resuspended in DMEM/FBS/PS ready for plating

[176].

2.5.2 Cell Transfection

Prior to transient transfections, cell culture media was replaced with DMEM and 10%

FBS, without antibiotics. For transient small interfering RNA (siRNA) transfections,

Lipofectamine RNAiMAX was used according to the manufacturer’s instructions. For transient plasmid (DNA) transfections, Lipofectamine LTX was employed. Co- transfections of siRNA and plasmid (DNA) required Lipofectamine 2000, all reagents followed the manufacturer’s instructions. Transfections using CellLight mito-RFP were carried out using the manufacturer’s guidelines. For transient siRNA transfection in

3T3-L1 adipocytes, adipocytes were split into 6-well plates for one day or until re- attachment occurred. Adipocytes were then transfected with Lipofectamine RNAiMAX according to the manufacturer’s instructions. Forty eight-hours after transfection, cells were treated with 10nM isoproterenol and were harvested after 4h for western blotting [135]. Wortmannin was added to the adipocytes at a concentration of 100nM for 45m followed by harvesting and protein analysis.

2.5.3 Retroviral shRNA Stable Transfections

Virus production: Phoenix-E cells were plated at 1-2x106 cells /10cm dish and incubated overnight at 37°C with 5% CO2. Cells were transfected using Lipofectamine

LTX according to the manufacturer’s instructions. Approximately 6µg of plasmid DNA

Materials and Methods

(pBABe-puro) was required per 10cm dish. Cells were incubated with the transfection complex for 48-72h. Prior to harvesting the virus, 3T3-L1 preadipocytes were grown in a 6-well plate for cell infection.

Harvesting virus: Virus media was transferred into a 10ml syringe and polybrene (5-

10µg/ml) was added. The virus was filtered through a 0.45µm filter and collected in a new tube.

Cell Infection: Between 1-2ml of virus media and 1-2ml of DMEM/NCS was added to the 6-well plate with 3T3-L1 preadipocytes. Cells were incubated overnight with the virus media followed by trypsinisation to a 10cm dish. After cell attachment, media was changed to DMEM/NCS containing 4µg/ml of puromycin as the selective agent.

Cells were selected for 72h followed by a further selection for 48h

2.5.4 Fatty Acid-Supplemented Medium

To stimulate neutral lipid synthesis, mammalian cells were supplemented with oleate- coupled BSA. 20mM sodium oleate was added to 5% BSA at a molar ratio of ~8:1.

Oleate-coupled BSA was added to DMEM/10% FBS/P/S at a molar concentration of ~

6.6:1. The fatty acid medium was filtered through a 0.22µm filter tip. The final concentration of oleate in the medium was 400µM [177]. Cells were treated with oleate from 4h to 14h.

Materials and Methods

2.6 Yeast Strains

S. cerevisiae wild type (WT) strain BY4741 (MATa; his3Δ1; leu2Δ0; met15Δ0; ura3Δ0) and the non-essential gene-deletion collection were obtained from European

Saccharomyces cerevisiae Archives for Functional Analysis (EUROSCARF). The S. cerevisiae WT strain, R1158 (URA3::CMV-tTA MATa his3-1 leu2-0 met15-0) was purchased from Open Biosystems and used in the expression of the essential yeast Tet- promoters Hughes collection (yTHC), controlled by the regulatable TetO7 promoter.

2.6.1 Yeast Growth Conditions

Yeast cells were proliferated in YPD medium containing 1% yeast extract, 2% (w/v) bacteriological peptone and 2% (w/v) D-glucose. Essential genes were repressed in the presence of doxycycline (15µg/ml) in YPD medium [6, 178]. Synthetic complete (SC) media contained 2% (w/v) D-glucose, 0.17% yeast nitrogen base (without ammonium sulfate and amino acids) and 0.5% ammonium sulphate, as well as the appropriate amino acids (Table 1-5). SC medium was needed for the selection of yeast cells which had been transformed with plasmids, with the suitable amino acids absent from the media. For example, SC medium lacking tryptophan is symbolised as SC-trp.

Materials and Methods

Table 1-5. List of amino acids added to SC media

Amino Acids added to SC medium

L-Lysine

L-Leucine

L-Adenine

L-Histidine

L-Tryptophan

L-Methionine

Uracil

Amino acid stock (L-Arginine, L-aspartic acid,

L-Isoleucine, L-Phenylalanine, L-Proline, L-

Serine, L-Threonine and L-Valine)

2.6.2 Yeast Transformation (TRAFO method)

Yeast cells were transformed with plasmids according to the TRAFO method [179].

Cells were harvested in a 50mL tube, centrifuged at 4000rpm for 10m and resuspended in 25mL sterile H2O. Centrifugation was repeated and 1mL of 100mM

LiOAc solution was added to the cells. The cells were then resuspended in 400µl of

100mM LiOAc and centrifuged again. The following products were mixed into the tube;

Materials and Methods

240µl PEG 3350, 36µl 1M LiOAc, 10µl single stranded salmon sperm DNA, 1 µl plasmid

DNA from miniprep and 60µl of sterile H2O. The contents were vortexed for 1m, incubated at 30˚C for 30m followed by heat shock at 42˚C for 30m. Tubes were centrifuged at 8000 rpm for 15 seconds and resuspended in 200µl of sterile H2O. The transformation mix was plated onto a SC-selective plate and incubated for 3 days at

30˚C.

2.7 Microscopy

2.7.1 Fluorescence Microscopy

Mammalian cells grown on coverslips were fixed with 4% paraformaldehyde (PFA) for

20m at room temperature. Mammalian LDs were stained with freshly prepared 2µg/ml

BODIPY 493/503 in 150mM NaCl for 10m [177]. Yeast LDs were stained with Nile Red in acetone (20µg/ml) at a dilution of 1:100 [53]. Coverslips were mounted onto slides using ProLong® Antifade Gold Reagent. Cells were imaged using a Leica CTR5500 microscope (Wetzlar, Germany) equipped with an EL6000 fluorescent lamp and a

DFC300 FX digital camera. Z-stack confocal images and still images were carried out using the Leica SP5 CW STED microscope and Zeiss LSM 780 microscope. 3D rendering was carried out using Huygens Essential Software. For confocal microscopy, cells were imaged using the 100x 1.4 objective with immersion oil. The green fluorescent protein

(GFP) was imaged using a 488nm Argon-ion laser line. The red fluorescent protein

(RFP) was imaged using a 561nm diode pumped solid state laser, while blue fluorescent protein (BFP) images were taken with a 405nm laser line.

Materials and Methods

2.7.2 Live Cell Imaging

For live cell imaging, cells were grown on a 35mm glass dish with a bottom thickness of

0.17µm. Cells were imaged using the Nikon Biostation which was supplemented with

5% CO2 at 37°C. Cells were imaged every 15m for 16h by a 12 bit charged-coupled device camera using the phase filter. Images were saved in AVI format and viewed using Windows Live Movie Maker.

2.7.3 Immunofluorescence

Transfected cells grown on coverslips were fixed with 4% PFA for 15m at room temperature, followed by three washes of 1X PBS. Cells were permeabilised in saponin for 30m at room temperature and blocked for 1.5h at room temperature (Table 1-6).

Cells were incubated with the primary antibody for 1h at room temperature in blocking solution (1:500 dilution), followed by three 10m washes in 3% BSA/PBS. Cells were protected from direct light and further incubated with the secondary antibody

(Alex Fluor 1:500 dilution) in blocking solution for 1h at room temperature [180].

Subsequently, cells were washed with 3% BSA/PBS for 10m and mounted using

ProLong® Gold Antifade reagent. Immunofluorescence images were captured using the

Leica SP5 CW microscope.

Table 1-6. Recipe for buffers used in immunofluorescence. Buffer Recipe

Permeabilisation Buffer 0.1% saponin/1X PBS

Blocking Buffer 3% BSA/1X PBS/0.05% saponin

Materials and Methods

2.7.4 Fluorescence Resonance Energy Transfer (FRET) by Acceptor Photobleaching

EGFP (donor) and mCherry (acceptor) were used as the FRET pairs [181]. For FRET,

Seipin-EGFP and CDS2-mCherry were co-transfected into HeLa cells for 24h. Cells were fixed with 4% PFA and underwent FRET using the 63x 1.2 water objective on the Leica

SP5 CW microscope. Argon and DPSS 561 lasers were required, with three sequential scans taking place. The scan of the acceptor was carried out first using laser 561, with

5% laser intensity and a wavelength of 600-750nm. The donor scan was second, which required laser 488 and a wavelength of 493-550nm. Lastly, the FRET pair was scanned at a wavelength of 600-750nm using laser 488. FRET values were observed using LAS

AF software.

Acceptor Photobleaching: For acceptor photobleaching, the FRET AB wizard was opened up using LAS AF software. The donor (EGFP) and acceptor (mCherry) parameters were set. Cells were first identified using bright field and a region of interest (ROI) was selected. Bleaching was carried out with laser 561 at 100% intensity.

Automated evaluation of the efficiency before and after bleaching was observed.

Materials and Methods

2.8 Lipid Analysis

2.8.1 Oleate incorporation

HeLa cells were grown in a 6-well plate and transfected with siRNA overnight at 37°C

14 with 5% CO2. For oleate incorporation, cells were treated with 1µCi/per well of C- oleate for 30m at 37°C with 5% CO2. For steady state, cells were treated with 1µCi of

14 C-oleate for 20h at 37°C with 5% CO2 [63]. Lipid extraction was performed as mentioned in the 2.8.2. Neutral lipids were spotted onto a thin-layer silica TLC sheet.

The TLC sheet was exposed to a BAS-MS imaging plate (Fujifilm, Tokyo, Japan) for 96h.

The imaging plate was visualised using the FLA-5100 phosphorimager (Fujifilm). The relative intensities of bands relating to triacylglycerol were quantified using ImageJ

(NIH).

2.8.2 Neutral Lipid Extraction

HeLa cells grown in 10cm dishes were washed in PBS and lysed with 0.1M NaOH for

10m at room temperature, with rocking. The dishes were washed with 1X PBS and 1M

HCl. The cell lysate was pooled in a screw-capped tube and the protein concentration was determined using a BCA protein assay kit. To the cell lysate, 2ml of methanol was added and mixed by shaking, followed by 2ml of hexane [63]. The solution was vortexed for 30s and spun for 5m at 1000 x g. The top layer was transferred to a glass vial and air-dried overnight. Protein concentrations were normalised to total neutral lipids and resuspended in 60µl hexane.

Materials and Methods

2.8.3 Phospholipid Extraction

Phospholipids were extracted according to Folch’s protocol [182]. Briefly, cells were washed with 1X PBS and scraped into a falcon tube. Aliquots were used to determine the protein concentration of transfected cells using a BCA protein assay kit. Cells were spun at 4000 rpm for 5m, and the supernatant was discarded. Cell pellets were homogenised with 2ml chloroform/methanol (2:1) and shaken for 15m at room temperature. To recover the liquid phase, 250µl of water was added to the solvent mixture and vortexed for 1m. The mixture was spun down for 5m at 3000 rpm and the lower phase was transferred to a new tube. The solvent was washed with 400µl of water, briefly vortexed, and spun at 2000 rpm for 10m until two distinct phases were evident. The bottom phospholipid phase was transferred to a glass vial and air-dried overnight. Phospholipids were dissolved in chloroform: methanol at a ratio of 2:1.

2.8.4 Thin Layer Chromatography (TLC)

Extracted neutral lipids were spotted onto a Silica Gel 60 F254 TLC plate and run in an equilibrated tank of hexane/diethyl ether/acetic acid (85:15:1), with triolein and cholesteryl ester as the standard. Colourisation of lipids was observed after a 30m incubation of the TLC plate in the iodine tank. The TLC plate was scanned using EPSON

PERFECTION 4490 PHOTO and the image was analysed using Image J software.

Materials and Methods

2.8.5 Mass Spectrometry

Prior to phospholipid extraction, 50µl of internal standard was added to the scraped cells. For mass spectrometric analysis, phospholipids were dissolved in chloroform: methanol: ammonium acetate (2: 1: 300mM ammonium acetate). Phospholipid samples were acquired using the ABI QStar Pulsar mass spectrometer. PA, PG, PI and

PE samples were acquired using the automated negative electrospray ionisation mode, while PC samples were analysed in the positive electrospray ionisation mode [183].

Lipids were normalised to the internal standard and analysed using the software,

LipidView.

2.8.6 Oil Red O Stain

Adipocytes were fixed with 4% PFA for 1h at room temperature. To prevent drying, dishes were wrapped with parafilm and covered in aluminium. Following incubation,

100% isopropanol was added to PFA treated dishes. The medium was removed and the cells were washed with 60% isopropanol. After the removal of isopropanol, the dish was left to completely dry. Oil Red O working solution was added to completely cover the surface of the dish [184]. Adipocytes were stained with Oil Red O for 10m (Table 1-

7). Once Oil Red O was removed, adipocytes were washed four times with dH2O.

Dishes were scanned using a UMAX scanner.

Table 1-7. Recipe for Oil Red O solutions. Oil Red O Solution Recipe

Stock solution 175mg Oil Red O: 50mL Isopropanol

Working solution Stock solution: dH2O (3:2)

Materials and Methods

2.9 RNA Extraction and cDNA Synthesis

Total RNA was extracted using Trizol TM reagent following the manufacturer’s protocol. Two micrograms of RNA were transcribed into cDNA using the Superscript

VILO cDNA synthesis kit, with a total volume of 20µl per reaction. Reverse transcription was carried out at 25°C for 10m, 42°C for 1h and 85°C for 5m.

2.10 Quantitative RT-PCR (qRT-PCR) qRT-PCR was performed using SYBRGreen mix. Samples were held at 95˚C for 3m followed by 40 cycles of denaturation at 95˚C for 5s, annealing at 60˚C for 20s and extension at 72˚C for 30s. The threshold cycle (Ct) value was determined in each run.

The mRNA expression levels were normalised against the housekeeping gene and compared to the expression levels in the control cells. Primers for qRT-PCR were designed using the software, Primer 3.0 (Table 1-2).

2.11 Protein Analysis

2.11.1 Western Blotting

Cell lysis and protein concentration: Cells were lysed with lysis buffer containing protease and phosphatase inhibitors. After lysis, cell lysates were passed through a 22- gauge needle 10 times and vortexed for 15m. The protein concentration was determined using a protein BCA assay kit according to the manufacturer’s protocol.

The assay mixture was incubated at 37°C for 30m and cooled to room temperature.

Within 10m of cooling, the absorbance of the BSA standard and protein samples was measured on a spectrophotometer at 562nm. A standard curve was used to determine

Materials and Methods

the concentration of the protein samples, by measuring each BSA standard vs. unknown protein.

Protein loading buffer: Normalised proteins were mixed with equal volumes of protein lysate and 0.02 % (w/v) bromophenol blue and 5 % (v/v) β-mercaptoethanol

(BME), in a 1:1 ratio. Samples were heated at 90°C for 5m on a heat block and were loaded onto a 10% (w/v) SDS-PAGE gel.

Western Blotting – Samples loaded onto the SDS-PAGE gel were run at 150 V for 1.5h. The gel was transferred onto a nitrocellulose membrane by either the trans- blot SD semi-dry transfer at 17V for 1h, or, the wet transfer system at 100V for 1h. The membrane was stained with Ponceau Red C to verify protein expression and then rinsed once with PBS-T or TBS-T. The membrane was blocked with 20 mL of 5% skim milk in PBS-T or 5% BSA in TBS-T (filtered) for 1h at room temperature. The membrane was incubated overnight at 4°C with the primary antibody (1:1000 dilution) in 5% skim milk/PBS-T or BSA/TBS-T, followed by three 10m washes in PBS-T or TBS-T. The membrane was then incubated with the secondary antibody (1:5000 dilution ) in 5% skim milk/PBS-T or BSA/TBS-T for 1h at room temperature, followed by a further three washes. Protein expression was detected using the enhance chemiluminesscence (ECL) detection kit (1:1 ratio). Membranes were visualised using the BioRad ChemiDoc XRS+ imager and analysed using the software, Image Lab. Buffer recipes are listed in Table 1-

Materials and Methods

Table 1-8. Buffers used in western blotting. Buffer Recipe

Lysis Buffer 10mM Tris pH 7.4, 100mM

NaCl, 1% SDS, protease

inhibitor & phosphatase

inhibitor (1:100)

SDS-PAGE Gel Running Per 1L: 3.0g Tris-base, 14.4 g

Buffer Glycine, 10% SDS

Western Blot Transfer Buffer Per 1L: 3.0g Tris-base, 14.4g

Glycine and 200mL methanol

Ponceau Red Stain Per 1L: 0.5g Ponceau S and

25mL Acetic

Phosphate Buffered Saline- 3.2 mM Na2HPO4; 0.5 mM

Tween 20 (PBS-T) KH2PO4; 1.3 mM KCl; 135

mM NaCl; 0.1% Tween 20

pH 7.4

Tris Buffered Saline-Tween Per 1L: 2.42g Tris-base, 8.78g

20 (TBS-T) NaCl and 0.05% Tween 20 pH

7.6

Blocking Buffer (PBS-T) 5% skim milk/PBS-T

Blocking buffer (TBS-T) 5% BSA/TBS-T (filtered

through 0.22µm filter)

Materials and Methods

2.11.2 Co-Immunoprecipitation by Dynabeads

Co-Immunoprecipitation by Dynabeads was prepared according to the manufacturer’s instructions. Cells grown in a 10cm dish were washed with ice-cold PBS and scraped into a pre-chilled 50ml falcon tube. The cell suspension was spun at 1000x g for 5m at

4°C. Pellets were resuspended in 1ml of lysis buffer containing protease inhibitors (PI dilution 1:100) and passed 15 times through a 22-gauge needle. The cell lysate was spun at the maximum speed for 15m at 4°C and the supernatant was transferred into a new tube. Dynabeads were resuspended and 50µl of Dynabeads per sample were transferred to a microcentrifuge tube. The tube was attached to the magnetic block and the supernatant was removed.

To obtain antibody bound Dynabeads, ~10µg of antibody was diluted in 200µl of PBS-T and transferred to the tube containing Dynabeads. The solution was rotated for 10m at room temperature and attached to the magnetic block to discard the supernatant. Up to 1ml of the cell lysate was added to the antibody bound Dynabeads and rotated for

2h at room temperature. The tube was transferred to the magnetic block and the supernatant was discarded. The same volume of PBS was added to the beads and mixed by inverting; the mixture was then transferred to a pre-chilled microcentrifuge tube. The supernatant was once again removed using the magnetic block. The wash step was repeated three times to remove as much supernatant as possible. 60µl of 2X

BME buffer was added to the beads and heated at 70°C for 10m on a heating bock.

Tubes were transferred to the magnetic block and the eluate was subjected to SDS-

PAGE and immunoblotting.

Materials and Methods

2.11.3 DUALmembrane yeast two-hybrid system

The yeast-two hybrid system identifies protein-protein interactions between two integral membrane proteins. Following the manufacturer’s instructions, a bait construct containing FLD1 was fused to the C terminal half of ubiquitin, pB3TC-FLD1, while the prey construct containing CDS1 was fused to the N terminal half of ubiquitin, pPR3N-CDS1.

Plasmids were transformed in yeast cells, along with the positive control, pB3TC-FLD1: pAI-Alg5 and the negative controls, pB3TC:pPR3N-CDS1 and pB3TC-FLD1:pD2-Alg5.

Cells were grown on SC-leu-his-trp-ade plates at 30°C overnight. The activation of the genes was signified by the growth of yeast on defined minimal media.

To further verify the interaction between two proteins, an X-gal agarose overlay was carried out. Briefly, 0.5g of low melting agarose was added to 100ml of stock solution and microwaved at 15s intervals until fully dissolved. 2.5µl of X-gal (100mg/ml) was added per ml of solution along with 0.5µl of BME per ml. The solution (~7ml) was applied directly onto SC-leu-his-trp-ade plates and kept away from direct light for 10m at 30°C. Strong inducers turned blue within 2h of the overlay.

Materials and Methods

2.12 Biochemical Assays

2.12.1 Lipolysis Assay

Adipocytes were split and transfected according to 2.5.2. For the lipolysis assay, media was changed to OPTI-MEM with 2% fatty acid free BSA (basal). To induce lipolysis,

10µM isoproterenol and 10µM IBMX was added to the media [11]. Media was collected and frozen immediately at the following time-points: 0m, 10m, 30m, 60m,

120m and 240m. Free glycerol assay was carried out using the manufacturer’s instructions (Biovision). Briefly, dilutions of the glycerol standard and glycerol assay buffer were mixed to prepare a standard curve, while 50µl of the unknown samples were combined with the reaction mix (glycerol assay buffer, PicoProbe™, glycerol enzyme mix and glycerol developer). The reaction was protected from the light and incubated at room temperature for 1h. Fluorescence was measured at an excitation of

535nm and emission of 587nm. The following equation was used to calculate the glycerol concentration.

Sample Glycerol concentration (C) = B/V x Dilution Factor = pmol/µl = nmol/ml = µM

2.13 Statistical Analysis

Data was analysed in GraphPad Prism 5 and GraphPad Prism 6. The error from samples was measured using the standard error mean (SEM). Statistical significance was carried out using either two-way ANOVA or one-way ANOVA. In two-way ANOVA, within each row, the columns were compared and each cell mean was compared to that of the

Materials and Methods

control. In one-way ANOVA, the mean of each column was compared with the mean of the control column.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

3. CHAPTER 3. CDP-DAG SYNTHASES REGULATE THE GROWTH OF LIPID DROPLETS AND ADIPOCYTE DEVELOPMENT

3.1 Background

Lipid droplets (LDs) are highly dynamic organelles that play a central role in mammalian energy storage. LDs are also involved in many cellular functions including, protein storage and degradation, membrane trafficking and lipid trafficking [1, 2, 87,

185]. LDs contain a highly hydrophobic neutral lipid core surrounded by a phospholipid monolayer [3]. In mammals, the lipid core comprises mainly triacylglycerols (TAGs) and cholesteryl esters (CEs). The biogenesis of LDs remains poorly understood. While several models have been proposed, the prevailing theory is that LDs originate and bud from the ER, followed by expansion and maturation [4]. The size of LDs varies within different tissues and even within the same cell type [5]. Recent studies have established the Cide family proteins, Cidec/FSP27 in particular, as key regulators of LD growth and fusion in adipocytes [11, 83]. Phospholipids, especially PC and PA, also appear to be important regulators of LD growth and proliferation. A decrease in PC or an increase in PA can both result in the formation of ‘supersized’ LDs (SLDs) [53, 55,

186]. LDs can also grow by lipid synthesis in situ, as TAG synthesis enzymes can relocalise from the ER to LD surface to mediate LD growth [87]. Despite these progresses, our understanding of LD biogenesis and growth is still at its infancy, and additional mechanisms and regulators remain to be identified.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

To identify novel gene products regulating the cellular dynamics of LDs, a genome wide screen in the budding yeast, S. cerevisiae was carried out. The initial screen identified

Fld1p (yeast homologue of human seipin) as a major regulator of LD dynamics. Fld1p has been implicated in the regulation and metabolism of fatty acids and phospholipids

[6, 53, 61]. Recently, seipin has been shown to be involved in TAG synthesis, LD formation and adipocyte differentiation [63, 156]. In yeast, the deletion of FLD1 resulted in the formation of SLDs. SLDs in the budding yeast often exhibit up to 50 times the volume of wild-type cells [6].

To identify additional “supersized” candidates, further genome-wide screens in

Saccharomyces cerevisiae were carried out. The screen found that knocking-down

CDS1, which encodes CDP- diacylglycerol synthase, can form giant LDs. In mammals, there are two CDP-diacylglycerol synthases (CDS), CDS1 and CDS2. These enzymes catalyse the formation of CDP-DAG by utilising phosphatidic acid (PA), the precursor for all phospholipids and TAG storage. Cds1p and Cds2p localise to two subcellular domains, the inner mitochondrial membrane, where it has been implicated in the synthesis of phosphatidylglycerol (PG) and cardiolipin (CL) and the endoplasmic reticulum, where it is involved in the synthesis of phosphatidylinositol (PI) [164].

Although the biochemical function of CDS1 and CDS2 has been characterised, little is known about their involvement during LD formation and adipocyte differentiation.

Understanding the physiological relevance of SLDs and the molecular mechanisms underlying the expansion of LDs and adipocyte development will provide insights into how therapeutic strategies can be developed against human metabolic diseases.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

The aim of this study was to examine the role of CDP-diacylglycerol synthases in LD dynamics and adipocyte differentiation. Here, it has been shown that CDS1 and CDS2 regulate both LD expansion and adipocyte development, most likely through modulating the level of PA.

3.2 Reverse Genetic Screen in Yeast Identified CDS1 as a ‘Supersized’ Lipid Droplet Mutant

In an attempt to identify additional regulators involved in LD dynamics, a reverse- genetic screen of the Tim Hughes collection (yTHC) of 838 essential gene strains was completed [187]. The yTHC strains were grown to mid log phase and mid-late stationary phase, stained with Nile Red and observed under the fluorescence microscope. The number, size, shape and localisation of LDs were recorded. The LD morphology of WT cells showed anywhere from 3-7 LDs per cell with an average size between 200-400nm. In the presence of doxycycline, CDS1 strains displayed SLDs, which were >1µm in diameter (Figure 3-1a) [53]. The phenotypic characteristics of

CDS1 were also observed at log-phase. When grown to log phase (OD600 = 0.8), the LDs in CDS1 showed a similar morphology to the cells grown to stationary. The overexpression of CDS1-GFP in Tet-CDS1 cells restored the LD morphology to that of the WT (Figure 3-1b). The localisation of CDS1 appeared to be the endoplasmic reticulum (Figure 3-1b).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-1: CDS1 expression in S. cerevisiae (Extracted from Tamar Kapterian’s Honours thesis 2009). (A) Wild-type and Tet-CDS1 cells were grown overnight in YPD + doxycycline. LDs were stained with Nile Red and fluorescence microscopy was carried out. (B) CDS1-GFP was overexpressed in Tet-CDS1 cells and grown in YPD + doxycycline. LDs were stained with Nile Red and imaged using fluorescence microscopy.

3.3 Down-Regulation of CDP-DAG Synthases Formed “Supersized” Lipid Droplets

To determine if CDP-DAG synthases in mammalian cells regulate the expansion/morphology of LDs, transient siRNA knock down experiments were carried out in HeLa cells and 3T3-L1 preadipocytes. Cells were treated overnight with BSA- coupled oleate and LDs were stained with BODIPY 493/503 prior to fluorescence microscopy.

Fluorescence microscopy indicated that knocking down CDS1 or CDS2 in both HeLa and

3T3-L1 cells resulted in the formation of giant/supersized LDs. In normal HeLa or 3T3-

L1 cells, the diameters of LDs were below 2 µm even after oleate treatment for 20 hours under our culture conditions. In contrast, most siCDS1 (CDS1 siRNA) and siCDS2

(CDS2 siRNA) cells displayed LDs with diameters between 3-9µm (Figure 3-2a). For the

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

purpose of SLD classification in HeLa cells and 3T3L1 preadipocytes, LDs with diameters

>2µm were arbitrarily categorised as SLDs. Cell counting revealed that 60% of siCDS1 and 50% of siCDS2 HeLa cells harboured SLDs (Figure 3-2b). 3D rendering of siCTRL, siCDS1 & siCDS2 cells confirmed the “supersized” phenotype, and further validated the difference between the volume of control LDs and SLDs (Video 1a-c).

Figure 3-2: Transient siRNA knock down of CDS1 and CDS2. A) HeLa cells and 3T3-L1 preadipocytes were transfected with siRNA negative control (siCTRL), CDS1 siRNA (siCDS1) and CDS2 siRNA (siCDS2). LDs were treated with oleate and stained with BODIPY. Cells were imaged using fluorescence microscopy. Bar= 20µm (HeLa) Bar= 45µm (3T3-L1). B) Percentage of HeLa cells with the “supersized” phenotype. n=100. P<0.001

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

3.3.1 mRNA Expression Levels of CDP-DAG Synthases

To examine the effect of gene silencing, qRT-PCR was carried out on siCDS1 and siCDS2 cells. All values were normalised to siCTRL. Compared to the control, CDS1 was down- regulated by ~75% and 50% in HeLa and 3T3-L1 cells respectively (Figure 3-3a); while

CDS2 was down-regulated by ~90% in both cell types (Figure 3-3b).

The expression levels of CDS1 and CDS2 were also investigated in their respective siRNA transfected cells. qRT-PCR showed that CDS1 was down-regulated by ~90% in siCDS1 cells, whereas in siCDS2 cells, the expression of CDS1 increased by ~3 fold.

(Figure 3-3c). CDS2 was down-regulated by ~80% in siCDS2 cells, whereas in siCDS1 cells, the expression level of CDS2 did not increase compared to the control (Figure 3-

3c). The fact the CDS1 activity was up-regulated in siCDS2 indicated that CDS1 might be the prominent CDP-DAG synthase. This notion was further supported because CDS2 mRNA levels did not increase in siCDS1 cells.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-3: qRT-PCR of siCDS1 and siCDS2 transfected Cell Lines. A) HeLa cells were transfected with human siCTRL, siCDS2 and siCDS2. qRT-PCR was carried out 24h post- transfection to observe the effect of gene silencing. B) 3T3-L1 preadipocytes were transfected with murine siCTRL, siCDS1 and siCDS2. qRT-PCR was carried out 24h post-transfection to confirm the effect of gene silencing. C) qRT-PCR of CDS1 and CDS2 expression in siRNA

transfected HeLa cells. p< 0.01.

3.4 Supersized Lipid Droplets Emerge From Clustered Lipid Droplets

To characterise the formation of SLDs in greater detail, LD formation was examined in

HeLa cells treated with oleate in a time course experiment. As shown in Figure 3-4, after 4h of oleate treatment, control HeLa cells appeared to have many and small LDs with no apparent change in distribution. After 8h of oleate treatment, HeLa cells appeared to have a similar phenotype to that of 4h, with slightly larger LDs. At 10h and

14h of oleate treatment, HeLa cells displayed many LDs, while LDs of up to 1µm in 71

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

diameter could be seen. After 20h of oleate treatment, HeLa cells displayed LDs which were 1-2µm in size. By contrast, after 4h of oleate treatment, siCDS1 and siCDS2 cells displayed many and small LDs (Figure 3-4). At 8h, siCDS1 and siCDS2 cells displayed many LDs that were densely packed and clustered (Arrow). To further confirm this clustered phenotype, 3D rendering was carried out on siCDS1 cells to understand how densely packed the LDs were after 8h of oleate treatment (Video 2). After 10h, both

CDS knock-down cells displayed large LDs that were in close proximity to one another, while at 14h (overnight) the SLD had fully emerged (Figure 3-4). The control cells however, did not form SLDs after 10h and 14h of oleate treatment. At 20h of oleate treatment, siCDS1 and siCDS2 maintained the “supersized” phenotype.

Figure 3-4: Time course of Oleate Treated HeLa Cells. HeLa cells were down-regulated with siCTRL, siCDS1 and siCDS2. Cells were treated with oleate for 4h, 8h, 10h, 14h and 20h. LDs were stained with BODIPY and imaged using fluorescence microscopy. Bar= 20µm. Arrow = Clustered LD phenotype.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

3.5 The Localisation of CDS1 and CDS2

The ER plays a major role in membrane lipid synthesis and in lipid droplet biogenesis and growth. A recent study showed that Rab10 localises to the ER and to dynamic ER- associated structures that mark the position of new ER tubule growth [188].

Interestingly, the same study found that two ER enzymes, phosphatidylinositol synthase (PIS) and choline/ethanolamine-phosphotransferase (CEPT) localised to the

Rab10 domain. This study hypothesised that CDS1 and/or CDS2 may localise to the

Rab10 domain. Also, this study was interested in the spatial relationship between LDs and the Rab10 domain. HeLa cells were transfected with BFP-Rab10, an ER specific marker, and CDS1-flag or CDS2-mCherry. Immunofluorescence was carried out on

CDS1-flag transfected cells using Alex-Fluor 594nm secondary antibody. Cells were fixed with 4% PFA and confocal microscopy was carried out. Fluorescence microscopy revealed that while CDS1 and CDS2 do localise to the ER, they do not localise with ER associated structures (Figure 3-5). Furthermore, to examine if LDs closely interact with the Rab10/PIS1 domain, HeLa cells were transiently transfected with mCherry-Rab10 and incubated with oleate for 4h, 8h, 12h and 16h. Fluorescence microscopy revealed that the ER associated structures, at best, partially co-localise with the LDs (Figure 3-5-

1).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

BFP RFP Merge

Figure 3-5: Co-localisation of Rab10 and CDP-DAG synthases. HeLa cells were co-transfected with BFP-Rab10, DsRed-ER, CDS1-flag and CDS2-mCherry. Cells were imaged using fluorescence microscopy. Arrow = Rab10 ER-associated structures.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-5-1: Co-localisation of Rab10 with lipid droplets. HeLa cells were transfected with mCherry-Rab10 and treated with oleate for 4h, 8h, 12h and 16h. LDs were stained with BODIPY and fluorescence microscopy was carried out. Arrow = Rab10 ER-associated structures.

As CDP-DAG is the precursor to the synthesis of PI, CDS1-flag and CDS2-mCherry were co-transfected with GFP-PIS1. Fluorescence microscopy showed that while both phospholipid synthesising enzymes localised to the ER, only GFP-PIS localised with ER- associated structures (Figure 3-6).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

RFP GFP Merge

Figure 3-6: Co-localisation of GFP-PIS and CDP-DAG synthases. Co-localisation of GFP -PIS with CDS1 and CDS2 showed CDP-DAG synthases do not co-localise to ER-associated

structures like PIS. Arrow = ER-associated structures.

3.6 The Synthesis of Triacylglycerols (TAG) is Increased in siCDS1 & siCDS2 Cells

As CDS is the enzyme required for converting PA to CDP-DAG, the down regulation of

CDS1/2 genes could cause an accumulation of PA and DAG, which could affect the synthesis of TAG, as well as that of PC and PE through the Kennedy pathway. In fact, the formation of SLDs in siCDS1 & siCDS2 cells could result from excessive TAG accumulation or a reduction in total phospholipids, in particular PC and PE. Steady- state levels of TAG were therefore measured, as well as the rate of oleate incorporation into TAG. While siCDS1 & siCDS2 cells both displayed a significant increase in the rate of oleate incorporation into TAG (Figure 3-7a), steady-state levels of TAG increased significantly only in siCDS1 cells (Figure 3-7b).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-7: Neutral lipid quantification in siCTRL, siCDS1 and siCDS2 HeLa cells. (A) 14C-oleate incorporation in siCTRL, siCDS1 and siCDS2 HeLa cells. After 30m of 14C-oleate treatment, neutral lipids were extracted and quantified by TLC. p < 0.05 (B) 14C-oleate steady state treatment in siCTRL, siCDS1 and siCDS2 HeLa cells. After 20h of 14C-oleate treatment, neutral

lipids were extracted and quantified by TLC. p < 0.05.

The level of PC, PE, PG and PI was also investigated in siCDS1 and siCDS2 HeLa cells.

Transient siRNA transfections of CDS1 and CDS2 were carried out in HeLa cells.

Phospholipids were extracted following Folch’s protocol and mass spectrometry was carried in the negative and positive ionisation mode. Total phospholipids were normalised to the protein concentration and analysed using the software, LipidView.

Mass spectrometry revealed that there was no significant change in the total level of

PE or PC in siCDS1 cells, while the total cellular amount of PG and PI increased significantly (Figure 3-8a). Mass spectrometry data in siCDS2 cells also showed no significant difference in the total levels of PC, PE and PI, but a significant increase in PG compared to the control (Figure 3-8b).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-8: Quantification of total PC, PE, PG and PI in siCDS1 and siCDS2 cells. HeLa cells were transfected with siCTRL, siCDS1 and siCDS2 and phospholipids were extracted. Mass spectrometry was carried out in negative and positive ionization mode. Data was analysed using the software, LipidView. A) Mass spectrometry of total PC, PE, PG and PI in siCDS1 cells. p < 0.05. B) Mass spectrometry of total PC, PE, PG and PI in siCDS2 cells. p < 0.05.

3.7 PA Metabolism upon Knocking-Down CDS1 or CDS2

CDP-diacylglycerol synthesises CDP-DAG from PA, which is the precursor to the de novo synthesis of all major phospholipids. Previous studies have shown that this cone- shaped phospholipid plays a critical role in SLD formation in yeast [53]. This study hypothesised that knocking down CDS1 or CDS2 would result in an accumulation of PA, which is an important contributing factor to the development of SLDs.

HeLa cells were transfected with CDS1/2 siRNA for 24h prior to phospholipid extraction. Phospholipids were extracted using Folch’s protocol and analysed by mass spectrometry. Mass spectrometric data was quantified using LipidView software. As expected, by two-way ANOVA, siCDS1 cells displayed a nearly 3-fold increase in total

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

PA, as well as significant increases in almost all PA species (Figure 3-9). It should also be noted that of all PA species examined, the levels of PA 34:1 and PA 40:5 were significantly reduced in siCDS1 cells (Figure 3-9). Interestingly, siCDS2 cells did not display a similar increase, with no change in the total level of PA.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-9: Quantitative analysis of PA species by mass spectrometry in siCDS1 and siCDS2 HeLa cells. HeLa cells were transfected with siCTRL, siCDS1 and siCDS2. Phospholipids were extracted for mass spectrometry and run using the negative ionization mode. Lipids were normalised to the internal standard and analysed using the software, LipidVIew. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

Mass spectrometry was repeated on siCDS2 HeLa cells, and while there was a slight increase in the total level of cellular PA, the increase was not statistically significant

(Figure 3-9-1). However, there were significant increases in PA 34:3 and 36:3 of siCDS2

HeLa cells (Figure 3-9-1).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-9-1: Quantitative analysis of PA species in siCDS2 cells by mass spectrometry. HeLa cells were transfected with siCTRL and siCDS2. Phospholipids were extracted for mass spectrometry and run using the negative ionization mode. Lipids were normalised to the internal standard and analysed using the software, LipidVIew. * p < 0.05, *** p < 0.001.

As the measurement of total cellular PA did not reveal any subcellular changes of PA, the cellular location of PA was examined. PA localisation was determined by using a PA sensor, which contains the repeat of the Spo20p51-91 domain fused to green fluorescent protein (GFP) that specifically binds PA [189]. siCTRL, siCDS1 and siCDS2

HeLa cells were co-transfected with pmGFP-PASS (from here on referred to as PASS, which expresses the PA sensor) and DsRed-ER. Fluorescence microscopy revealed that the PA sensor clearly associated with the ER in siCDS1 and siCDS2 cells but not as strongly in the control cells (Figure 3-10), indicating that PA accumulates in the ER of siCDS1 and siCDS2 cells. Together, these results suggest that PA accumulates in the ER of both siCDS1 and siCDS2 cells, although the total amount of PA did not significantly increase in siCDS2 cells.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

pmGFP-PASS DsRed-ER Merge

Figure 3-10: Co-localisation studies of pmGFP-PASS and DsRed-ER in siCDS1 and siCDS2 cells. siC TRL, siCDS1 and siCDS2 were transiently co-transfected with DsRed-ER and pmGFP-PASS. Cells were fixed with 4% PFA and imaged using fluorescence microscopy.

As CDS1 and CDS2 are believed to have possible localisation in the mitochondria, we wondered if PA accumulates in the mitochondria in the absence of CDS1 or CDS2. HeLa cells were co-transfected with siCDS1 and siCDS2 along with PASS and the mitochondrial marker, Bacmam-2.0 Mito-RFP. Interestingly, fluorescence microscopy revealed no co-localisation between PASS and Mito-RFP, indicating that PA accumulates mostly in the ER (Figure 3-11).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

pmGFP-PASS Mito-RFP Merge

Figure 3-11: Co-localisation studies of pmGFP-PASS and Mito-RFP in siCDS1 and siCDS2 cells. siC TRL, siCDS1 and siCDS2 were transiently co-transfected with Mito-RFP and pmGFP-PASS. Cells were fixed with 4% PFA and imaged using fluorescence microscopy.

3.8 Overexpression of CDP-DAG Synthases Inhibits Lipid Droplet Synthesis

To further examine the role of CDP-DAG synthases in LD formation, HeLa cells were transiently transfected with mCherry-N1 alone, CDS2-mCherry alone or mCherry-N1 & pCDNA3-CDS1-flag. CDS1-flag was used as CDS1-mCherry construct could not be generated. HeLa cells were treated with 400µM BSA-coupled oleate for 6h and were subjected to fluorescence microscopy. When CDS1 or CDS2 were overexpressed, there was a significant reduction in the formation of LDs (Figure 3-12a). In contrast, overexpression of the empty vector control, mCherry-N1, exhibited no impairment in the formation of LDs, indicating the effect of lipid droplet formation was specific to

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

CDS1 and CDS2. Immunoblotting was carried out to confirm that CDS1 was overexpressed in the transiently transfected HeLa cells (Figure 3-12b).

Figure 3-12: Overexpression of CDP-DAG synthases upon oleate treatment. A) HeLa cells were transiently transfected with the empty vector control, mCherry-N1, mCherry- N1:pCDNA3-CDS1-flag and mCherry-N1-cds2 and treated with oleate for 6h. B) Western blot of transfected HeLa cells with CDS1-flag. Arrow = Transfected HeLa cells.

3.9 Overexpression of CDS Inhibits the Formation of SLDs

Besides CDS1, the PA phosphatase lipin can also metabolise PA by converting PA to

DAG for TAG, PC and PE synthesis. Lipin-1 is encoded by LPIN1 and is a known regulator of adipogenesis [190]. To further investigate whether PA regulates the size of

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

LDs rather than TAG, HeLa cells were transfected with siLPIN1. As shown in Figure 3-

13a, knocking-down lipin-1 induced the formation of SLDs, possibly due to the accumulation of PA. When CDS1 or CDS2 were overexpressed in siLPIN1 cells, the formation of SLDs was impaired (Figure 3-13c). These results suggest that CDS can inhibit the formation of SLDs, most likely via reducing the level of PA. qRT-PCR was carried out to confirm the effect of gene silencing. In HeLa cells, the expression of

LPIN1 was down-regulated by ~ 50% (Figure 3-13b). To further support the hypothesis that PA is required in the formation of SLDs, TLC was carried out in siLPIN1 HeLa cells that were co-transfected with overexpressed CDP-DAG synthases. HeLa cells were treated overnight with 14C-oleate and neutral lipids were extracted. Lipids were spotted onto a TLC aluminium sheet and exposed for 1 day using the phosphorimager.

Image J was used to quantify the amount of TAG per cell. TAG quantification showed no change in the level of TAG in siLPIN1 cells or cells co-transfected with siLPIN1 and

CDS1/2 plasmids (Figure 3-13d).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

A) B)

Figure 3-13: CDP-DAG synthase overexpression in siLPIN1 cells. A) HeLa cells were transfected with siLPIN1 and treated with oleate overnight. LDs were stained with BODIPY and detected by fluorescence microscopy. B) qRT-PCR of siLPIN1 transfected HeLa cells. C) HeLa cells were co- transfected with siLPIN1 and pCDNA3-CDS1-flag: mCherry-N1 or CDS2-mCherry. Cells were treated with oleate overnight and imaged using fluorescence microscopy. Bar = 25 µm. D) TAG quantification of transfected HeLa cells. HeLa cells were transfected with a combination of siCTRL, siLPIN1, CDS1-flag and CDS2-mCherry. Cells were treated overnight with 14C-oleate followed by neutral lipid extraction. 86

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

To further confirm that the growth of SLDs is modulated primarily through PA, the yeast homolog of lipin, PAH1, was overexpressed under a GAL1 promoter. The overexpression of PAH1 in Tet-CDS1 cells reduced SLD formation by almost half (Figure

3-14). Fluorescence microscopy revealed that LD size had decreased, while the number of LDs per cell increased.

Figure 3-14: PAH1 overexpression in Tet-CDS1 cells. PAH1 was overexpressed under a GAL1 promoter in WT cells and Tet-CDS1 cells. Bar =10µm

3.10 Down Regulation of CDS1 Inhibits Adipocyte Differentiation

Phosphatidic acid has recently been implicated in adipocyte differentiation [191]. This study hypothesised that down regulating CDS1 or CDS2 may cause an accumulation of

PA in the ER and nuclear membrane, which could inhibit adipocyte differentiation, possibly by interfering with PPARγ function. Firstly, the expression level of CDS1 and

CDS2 were measured during the differentiation of 3T3-L1 preadipocytes. Two days post confluence, 3T3-L1 cells were induced for differentiation. qRT-PCR was carried out on the extracted RNA on days 0, 2, 4 and 6. CDS1 expression steadily increased during differentiation, reaching 7 fold after 6 days; while CDS2 had a slight increase of ~40% on day 6 after a 50% decrease on day 2 (Figure 3-15). The expression of the main adipogenic markers, PPARγ, aP2 and C/EBPα progressively increased during differentiation. Pref1 was used to determine the preadipocyte factor, while Caspase 3 measured cell death.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-15: Expression of CDS1 and CDS2 during adipocyte differentiation. 3T3-L1 preadipocytes were induced for differentiation two days post confluence. Samples were

collected at days 0, 2, 4 and 6. RNA was extracted and qRT-PCR was carried out to measure the expression levels of PPARγ, aP2, CEBPα, CDS1, CDS2, Caspase3 and Pref1.

To investigate whether the down-regulation of CDS1 and CDS2 inhibited adipocyte differentiation, transient siRNA knock-down experiments were carried out in 3T3-L1 preadipocytes which were induced for differentiation. On day 4, CDS1 was down regulated by ~40%, while the expression of PPARγ significantly decreased by ~10 fold.

On day 6, PPARγ activity further decreased by ~15 fold (Figure 3-16). Similarly, on day 4, aP2 expression was reduced by ~700 fold, and ~1200 on day 6. C/EBPα expression also significantly declined from days 2-6 by ~100 fold and ~400 fold respectively (Figure 3-

16). Compared to the control, the expression of Caspase3 in siCDS2 cells remained constant from days 2-6. Pref1 expression remained the same in siCDS1 differentiated

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

cells. Oil Red O staining further revealed the near complete inhibition of adipocyte differentiation in siCDS1 cells (Figure 3-17a). Light microscopy further confirmed the inhibition of adipogenesis in siCDS1 cells compared to the control which displayed the adipocyte phenotype (Figure 3-17b).

Figure 3-16: Down-regulation of CDS1 during adipocyte differentiation. 3T3-L1 preadipocytes were transfected with siCTRL or siCDS1. Two days post confluence, 3T3-L1

cells were induced for differentiation. qRT-PCR was used to measure the expression levels of CDS1, PPARγ, aP2, CEBPα, Caspase3 and Pref1, p < 0.0001.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-17: Oil Redo O staining of differentiated siCDS1 3T3-L1 cells. A) Oil Red O staining of

differentiated 3T3-L1 cells transfected with siCTRL and siCDS1. B) Light microscopy of differentiated 3T3-L1 cells transfected with siCTRL and siCDS1.

In cells treated with siCDS2, qRT-PCR indicated that CDS2 was initially down regulated by ~90% on day 0, followed by a ~40% reduction during differentiation. The expression of PPARγ, aP2 and C/EBPα was moderately but significantly decreased (Figure 3-18). Oil

Red O staining of siCDS2 cells showed a moderate decrease in adipocyte differentiation

(Figure 30-19a). Light microscopy also confirmed the partial inhibition in adipocyte differentiation (Figure 3-19b).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-18: Down-regulation of CDS2 during adipocyte differentiation. 3T3-L1 preadipocytes were transfected with siCTRL and siCDS2. Two days post-confluence, 3T3-L1 cells were induced for differentiation. qRT-PCR measured the expression levels of CDS2, PPARγ, aP2, CEBPα, Caspase3 and Pref1 on days 0-6. * p < 0.05, ** p < 0.01, *** p < 0.001 & **** p < 0.0001.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

Figure 3-19: Oil Red O staining of differentiated siCDS2 3T3-L1 cells. A) Oil Red O staining of differentiated 3T3-L1 cells transfected with siCTRL and siCDS2. B) Light microscopy of differentiated 3T3-L1 cells transfected with siCTRL and siCDS2.

3.10.1 Down-regulation of CDS2 using shRNA (shCDS2) inhibits adipocyte differentiation

As CDS2 moderately inhibited adipocyte differentiation, stable CDS2 shRNA 3T3-L1 cells lines were generated to validate this finding. While CDS1 shRNA did not work,

CDS2 cells were down-regulated by ~25% and also displayed the “supersized” phenotype (Figure 3-20a). Stable shCDS2-3T3-L1 and shLacZ-3T3-L1 (control) cells were induced for differentiation. qRT-PCR was carried out at days 0, 2, 4 and 6. The data confirmed the previous finding that the down-regulation of CDS2 partially inhibits adipocyte differentiation. When CDS2 was down-regulated during differentiation,

PPARγ was down regulated by ~5 fold and ~7 fold on days 4 and 6 respectively (Figure

3-20b). The expression level of aP2 was down-regulated by ~130-fold on day 6, while the activity of C/EBPα also decreased significantly by ~ 45-fold on day 6. Oil Red O staining further confirmed the moderate inhibition (Figure 3-20c).

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

C) shLacZ shCDS2

Figure 3-20: Down-regulation of CDS2 using shRNA (shCDS2). A) 3T3-L1 cells were transfected with shLacZ and shCDS2 and treated with oleate overnight. LDs were stained with BODIPY and fluorescence microscopy was carried out.. B) Stable cell lines of shLacZ and shCDS2 were induced for differentiation. qRT-PCR measured the expression CDS2, PPARγ, aP2, C/EBPα, Pref1 and Caspase3. (C) Oil Red O staining of shLacZ and shCDS2 cells induced for differentiation. * p < 0.05, ** p < 0.01, *** p < 0.001 & **** p < 0.0001. 93

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

3.11 Discussion

In this study, the role of CDP-DAG synthase, a key enzyme in phospholipid metabolism was examined in cellular lipid storage, as well as in the differentiation of adipocytes. The data produced from this study demonstrate that both CDS1 and CDS2 can regulate the expansion of LDs. Interestingly, CDS1, but not CDS2, appears to be essential for adipogenesis. This data provides an intimate link between the expansion of LDs at the cellular level and the differentiation of adipocytes at the systemic level, and suggest that phospholipids, i.e. phosphatidic acid (PA), may play an important role in the regulation of two seemingly disparate processes. By manipulating the expression levels of

CDP-DAG synthases, the level of PA was modulated. The results presented here imply that PA may regulate lipid storage at the cellular and systemic level.

LDs have now been recognised as dynamic organelles that constantly change in their size and number. However, the molecular mechanisms underlying the expansion or contraction of LDs remain to be fully elucidated. Therefore, identifying the molecular mechanisms involved in the cellular dynamics of LDs and adipocyte development is of great relevance in the fight against obesity.

A reverse genetic screen in S. cerevisiae identified CDS1 as a SLD mutant [53].

Generally, LDs in yeast WT cells range in size from 200-400nm. Under the tet- regulatable promoter, repressing the expression of CDS1 revealed LDs that were greater than ~ 1µm in diameter. The screen identified 10 SLD mutants in total and a noticeable feature of all mutants was their involvement in phospholipid metabolism

[53]. An essential role for PA in SLD formation was further established when all mutants that developed SLDs were found to accrue PA.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

To determine whether the “supersized” phenotype was evolutionarily conserved, transient siRNA transfections of CDS1 and CDS2 in HeLa cells and 3T3-L1 preadipocytes was carried out. The down-regulation of CDS1 and CDS2 revealed that SLDs were also evident in mammalian cell lines. Quantifying the mRNA level of CDP-DAG synthases in siCDS1 and siCDS2 cells revealed that CDS1 activity was significantly increased when

CDS2 was down-regulated. However, when CDS1 was down-regulated, there was no change in the expression level of CDS2. This finding indicates that CDS1 might be the main isozyme.

A recent study by English et al., found that Rab10 localises to ER-associated structures and regulates ER tubule growth [188]. The same study found that the Rab10 domain was enriched with two enzymes involved in phospholipid synthesis, PIS1 and CEPT1. As both CDS1 and CDS2 are known to localise to the ER, this study wondered whether the two enzymes localised to two different ER domains [164, 165]. Both CDS1 and CDS2 showed no co-localisation with Rab10 or PIS1. As the authors hypothesised ER tubule growth and phospholipid synthesis could be coupled, the co-localisation of Rab10 with

LDs was investigated. Fluorescence microscopy revealed at best, a partial co- localisation of Rab10 with LDs.

The growth of LDs, while not completely known, appears to be a dynamic process, regulated in part, by lipids and proteins. Recent studies have identified key proteins such as FSP27 in the growth of LDs in adipocytes, and have also revealed a role of phospholipids in determining the size of the LDs [5, 11, 83]. For instance, decreased cellular PC or increased PA can both lead to the formation of giant LDs, possibly

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

involving FSP27-independent LD fusion. A time point of oleate treated cells indicated that siCDS1 and siCDS2 cells developed clustered LDs, prior to the formation of SLDs. In yeast, enhanced fusion activities are seen in CDS1 mutants as a result of changes in phospholipid composition. In vitro studies revealed that the addition of PA led to the fusion of artificial LDs. The clustered phenotype was not observed in siCTRL cells, which appeared to have no apparent localisation during LD growth. Thin layer chromatography revealed that pulse labelling with 14C-oleate for 30m showed a significant increase in the total level of TAG in siCDS1 and siCDS2 cells. The down- regulation of CDS1/2 could cause an accumulation of PA, which might increase the overall activity of LPIN1. Enhanced LPIN1 activity may increase expression of DGAT and ultimately, increase TAG. Overnight treatment with 14C oleate showed a significant increase in siCDS1, but not siCDS2 cells. While TAG may be a contributing factor in the growth of LDs, it is most likely not the rate-limiting lipid required in the formation of

SLDs.

Phosphatidic acid is a non-bilayer forming phospholipid [54]. It contains a smaller hydrophilic head group and a larger hydrophobic domain. As the surface of the LD contains a phospholipid monolayer, PA could alter the growth of LDs due to its conical shape, and induce membrane curvature. While mass spectrometry in yeast has shown that CDS1 mutants display an increase in total cellular PA [53], little is known about their role in mammalian cells. CDP-DAG synthase sits at a key branching point of phospholipid synthesis where it converts PA to CDP-DAG for PI and PG synthesis.

Knocking down CDS1 or CDS2 could increase the amount of PA and reduce the amount of PI and PG. However, the level of PI and PG increased upon depletion of CDS1 or

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

CDS2, whereas PA did accumulate, especially on the ER. The impact of depleting CDS1 on the level of PI and PG is surprising: a previous study demonstrated that CDP-DAG synthase is probably not a determining factor in controlling the de novo rate of PI synthesis or in establishing the cellular PI content as overexpressing CDS1 does not increase the cellular levels of CDP-DAG or PI [165]. Among other possibilities, the dramatic increase in the level of PI and PG upon depleting CDS1 could be caused by enhanced expression of CDS2 in those cells. While PI and PG do not appear to be the major phospholipids involved in LD formation, the data suggests that they might be contributing factors in LD expansion. Furthermore, no dramatic changes were detected in PC and PE.

Mass spectrometry in siCDS1 cells confirmed an overall increase in the total level of PA compared to the control. Interestingly, almost all PA species in siCDS1 HeLa cells showed a significant increase in PA. This data strongly suggests that PA is a key regulator of LD growth in mammalian cells. Surprisingly; siCDS2 cells did not exhibit the same findings. While several species of PA increased in siCDS2 cells, the measurement of total cellular PA did not significantly increase. As LDs are believed to originate from the ER, it is possible that the accumulation of PA occurs at the microsomal level in siCDS2 cells, and that the localisation of PA may also influence the formation of SLDs.

Fluorescence microscopy revealed that PA accumulates in the ER of siCDS1 and siCDS2 cells. Compared to the control, siCDS1 and siCDS2 cells displayed a brighter intensity of

PA, indicating that more PA accumulates in the ER. As CDP-DAG synthases are also believed to localise to the mitochondria, a mitochondrial marker was co-transfected with the PA sensor. Fluorescence microscopy revealed no localisation of PA in siCDS1

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

and siCDS2 cells with the mitochondria, further supporting the hypothesis that PA accumulates in the ER.

This study has uncovered a role for CDP-DAG synthases in LD synthesis of cultured mammalian cells. Down-regulating CDP-DAG synthases in mammalian cells results in the formation of SLDs, whereas the overexpression of CDP-DAG synthases had an opposite effect. As this phenotype was not observed in control cells, it can be concluded that the significant reduction in LD formation was specific to CDS1 and

CDS2.

The PA phosphatase, lipin, can also metabolise PA by converting PA to DAG for TAG, PC and PE synthesis. Lipin-1 is encoded by LPIN1 and is a known regulator of adipogenesis

[191]. The critical role of PA in SLD formation was further supported when SLDs were formed upon lipin-1 knock-down. Importantly, overexpression of CDS1 or 2 restored the normal morphology in lipin-1 depleted cells, possibly through reducing the level of

PA. The total TAG levels were measured in HeLa cells transiently transfected with lipin1 siRNA. As TLC was carried out on all cells, there was no noticeable difference in the total TAG. Mass spectrometry should be carried out on cells down-regulated with lipin1 to accurately quantify the TAG amount.

Obesity can be defined as the accumulation of enlarged adipocytes loaded with giant

LDs, which is due in part, to enhanced LD expansion at the cellular level and adipocyte differentiation at the systemic level [192]. Excess adipose tissue can lead to increased lipid accumulation (hypertrophy) as well as the proliferation of white adipocytes

(hyperplasia). To examine the role of CDP-DAG synthases in adipogenesis, 3T3-L1

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

preadipocytes were induced for differentiation. During adipogenesis, CDS1 and CDS2 appear to be differentially regulated as CDS1, but not CDS2, is up-regulated. The differentiation of preadipocytes requires a transcriptional cascade that ultimately leads to the activation of the master regulator of terminal adipogenesis: peroxisome proliferator-activated receptor-γ (PPARγ). Depleting CDS1 had a strong inhibitory effect on adipocyte development, whereas depleting CDS2 had a moderate inhibitory effect. How does CDP-DAG synthase regulate adipogenesis? Given the changes in phospholipid profile upon depleting CDS1 or CDS2, this study hypothesised that PA may play a key role in this process as well.

The master driver of adipogenesis, PPARγ, has a large ligand binding pocket that can be activated by various metabolites that originate from phospholipid and fatty acids [96].

Despite the abundance of naturally occurring molecules that activate PPARγ in cell- based assays, the endogenous regulators of PPARγ that are of physiological importance remain poorly defined. The recent identification of cyclic PA (cPA) as an antagonist of PPARγ adds another layer to the regulation of PPARγ activity [193, 194].

Therefore, certain PA species could serve as direct antagonists of PPARγ. Alternatively, increased PA could affect post-translational modifications of PPARγ or could trap

PPARγ into certain membrane domains, preventing its activation. These and other possibilities need to be explored in future studies. Enzymes which regulate PA metabolism like CDS1 and CDS2 could be targeted to reduce PA production and increase adipocyte expansion for enhanced insulin sensitivity.

Chapter 3: CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

3.12 Conclusion

This study has shown that CDP-DAG synthases are more than just enzymes. They are regulators of LD growth and adipocyte development. This study has implicated that the accumulation of PA is involved in two processes; the generation of supersized LDs and the inhibition of adipogenesis. The knock-down of CDS1 and CDS2 generated SLDs, while 3D rendering confirmed the difference in the volume between the control and

CDS down-regulated cells. Overexpression of CDP-DAG synthases in HeLa cells showed that CDS1 and CDS2 inhibited LD synthesis. Mass spectrometry data revealed a significant increase in PA amongst siCDS1 cells, while fluorescence microscopy showed that PA strongly accumulates in the ER of siCDS1 and siCDS2 cells. Furthermore, at the mRNA level, the knock-down of CDS1 or CDS2 significantly decreased the expression level of PPARγ. How does PA inhibit adipogenesis? It is possible that PA is an antagonist of PPARγ, whether there are different species of PA which regulate LD development and adipogenesis are yet to be investigated.

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4. CHAPTER 4. THE CHARACTERISATION OF CDP-DAG SYNTHASES IN MATURE ADIPOCYTES

4.1 Background

Obesity can be characterised by an excess of adipose tissue (fat stores), the key factor in the development of type 2 diabetes [195]. Today, adipocytes are identified as major regulators of total body homeostasis and are known to be involved in the pathogenesis of a variety of metabolic diseases [196]. Adipocyte lipolysis involves the hydrolysis of

TAG to generate FFAs and free glycerol [197]. High insulin levels and increased FFAs produced through lipolysis are associated with insulin resistance [198]. Furthermore, increased plasma concentrations of FFAs have been implicated in obesity and type 2 diabetes [199]. Therefore, understanding adipocyte function is essential for the development of potential therapies in metabolic diseases.

Recent studies have shed light on the importance of PA in lipolysis and mTOR signalling

[135]. As mentioned in Chapter 1, lipin utilises PA and converts it to DAG for PC, PE and

TAG synthesis [58]. Lipin1 -/- mice exhibit increased levels of LPA and PA in the white adipose tissue. Also, the deletion of lipin1 in adipocytes was shown to reduce lipolytic activity under basal and isoproterenol-treated conditions, with changes in the expression of ATGL, HSL and p-PKA [135]. ATGL, the rate limiting TAG hydrolase is also a hormone-responsive enzyme. Like HSL, ATGL undergoes perilipin-dependent translocation to LDs in adipocytes treated with β-adrenergic agonists [200, 201]. ATGL initiates lipolysis by hydrolysing TAGs, and HSL mainly functions as a DAG lipase in vivo

[202].

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The aim of this study was to identify whether CDP-DAG synthases are involved lipolysis and mTOR signalling. Here, preliminary findings have shown that the down-regulation of CDP-DAG synthases in mature adipocytes affects lipolysis. Using siRNA transient knock-down experiments, this study has shown that CDP-DAG synthases are involved in a number of adipocyte functions. The preliminary findings have shown that the down-regulation of CDP-DAG synthases can affect the expression of PPARγ and

C/EBPα. Isoproterenol stimulated lipolysis revealed that the down-regulation of CDP-

DAG synthases impaired the expression of phosphorylated perilipin, while western blots shed light on the importance of CDS1 and CDS2 in the regulation of lipolysis.

Here, a range of potential functions for CDP-DAG synthases have been identified, other than their traditional role in phospholipid synthesis.

4.2 Down-Regulation of CDP-DAG Synthases in Adipocytes Results in ‘Supersized’ Lipid Droplets

As mentioned in Chapter 3, the down-regulation of CDS1 and CDS2 forms SLDs in cultured mammalian cells. Therefore, the role of CDS1 and CDS2 in LD expansion of mature adipocytes was examined. 3T3-L1 preadipocytes were differentiated until day

8 (maturation) and split. Adipocytes were transiently transfected with murine siCTRL, siCDS1 and siCDS2 for 24h. LDs were stained with BODIPY and fluorescence microscopy was carried out. Fluorescence microscopy revealed that adipocytes transfected with siCDS1 and siCDS2 bared the ‘supersized’ phenotype (Figure 4-1a).

To quantitate the effect of gene silencing, qRT-PCR was carried out on adipocytes that were transfected at day 8. In siCDS1 adipocytes, the expression of CDS1 was down- regulated by ~ 70%, while siCDS2 adipocytes were down-regulated by ~ 90% (Figure 4-

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1b). To further verify the effect of gene silencing at the protein level, immunoblotting was carried out on day 8 siCTRL, siCDS1 and siCDS2 adipocytes. While there was no working mouse CDS2 antibody at the time of this study, a CDS1 antibody was purchased from Abcam. Immunoblotting showed that the protein expression level of

CDS1 decreased in siCDS1 adipocytes (Figure 4-1c). Actin was used as the loading control.

B) C)

Figure 4-1: Down-Regulation of CDP-DAG synthases in mature adipocytes. A) 3T3 -L1 preadipocytes were induced for differentiation until day 8. Adipocytes were split ad transiently transfected with siCTRL, siCDS1 and siCDS2. LDs were stained with BODIPY 493/503 and subjected to fluorescence microscopy. Bar=20µm. B) qRT-PCR of CDS1 and CDS2 knock- down in mature adipocytes. p < 0.0001. C) Mature adipocytes were knock-down with siCTRL, siCDS1 and siCDS2. Immunoblotting using anti-CDS1 was carried out.

4.3 Lipid Droplet Fusion in siCDS1 and siCDS2 Mature Adipocytes

To investigate whether the inhibition of CDS activity caused the fusion of LDs, live cell imaging was carried out. Mature adipocytes were transiently transfected with siCTRL, siCDS1 and siCDS2. Using PHASE bright field light microscopy, live cell imaging was 103

Chapter 4: The Characterisation of CDP-DAG Synthases in Mature Adipocytes

carried out to observe LD fusion. LDs were not stained with any dye to validate that the fusion was real and not an artifact. In siCTRL adipocytes, it emerged that the larger

LDs made contact and came in close proximity of one another at a lipid-droplet- contact-site (LDCS). However, after 16h of imaging, it appeared that the large LDs in the control cells did not fuse together (Video-3a).

In contrast, adipocytes transfected with CDS1 siRNA produced large LDs that appeared to come together at the LDCS. The LD slowly fused into the large LD over several hours, eventually, two SLDs were observed in siCDS1 adipocytes (Video-3b). Another form of fusion that was observed in siCDS1 adipocytes was quick fusion. Rather than gradually fusing into a larger LD at the LDCS, quick fusion involved the fusion of LDs within minutes and sometimes seconds. In quick fusion, siCDS1 adipocytes displayed the unilocular LD phenotype (Video-3c). In Video-3c, seven LDs are within one adipocyte, through quick fusion, the smaller LDs fused together to form three SLDs. The formation of the SLDs led to the fusion of all three to generate a unilocular LD which occupies most of the surface area of the adipocyte. Similarly, siCDS2 adipocytes also displayed the phenotype of slow fusion to form large LDs. In siCDS2 adipocytes, many LDs appeared to fuse together and gradually fused to form a large LD (Video-3d). As this phenotype was not observed in siCTRL cells, these preliminary findings indicate that

CDP-DAG synthases can regulate the formation of SLDs in adipocytes.

4.4 Down-Regulation of CDP-DAG Synthases Suppress Adipocyte Maintenance

Because CDP-DAG synthases impair the differentiation of preadipocytes, the expression levels of the same adipogenic markers were examined in mature

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adipocytes. 3T3-L1 preadipocytes were induced for differentiation until day 8.

Adipocytes were split and transiently transfected with siCTRL, siCDS1 and siCDS2 prior to RNA extraction. qRT-PCR revealed that when adipocytes were down-regulated with

CDS1 by ~ 70%, the expression level of PPARγ and C/EBPα significantly decreased by ~

50% and ~ 60% respectively (Figure 4-2a). While the expression level of aP2 slightly decreased, statistical analysis showed no significance. Caspase3 showed now significant change, confirming the knock-down of CDS1 did not promote cell death.

Furthermore, when CDS2 was down-regulated by ~ 90%, the expression level of

C/EBPα significantly decreased (Figure 4-2b). Though there was a slight decrease in

PPARγ, the change was not statistically significant. Also, the expression level of aP2 and Caspase3 did not change compared to the control.

A) B)

Figure 4-2: qRT-PCR of siCDS1 and siCDS2 adipocytes. A) 3T3-L1 preadipocytes were differentiated until day 8. Mature adipocytes were split and transfected with siCTRL and siCDS1. qRT-PCR was carried out to measure the expression of CDS1, PPARγ, aP2, C/EBPα and Caspase3. p< 0.0001. B) 3T3-L1 preadipocytes were differentiated until day 8. Mature adipocytes were split and transfected with siCTRL and siCDS2. qRT-PCR was performed to measure the expression of CDS2, PPARγ, aP2, C/EBPα and Caspase3. *** p <0.001 **** p < 0.0001. 4.5 CDP-DAG Synthases are Involved in Lipolysis

Recent evidence has shown that the phosphatidic acid phosphatase (PAP) activity of lipin1 is a potential regulator of lipolysis [135]. Under basal conditions, adipose

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extracted from Adn-Lpin1-/- mice showed a reduction in the total protein level of ATGL and HSL. With this notion, it was hypothesised that the down-regulation of CDS1 or

CDS2 would cause an accumulation of PA and an increase in PA would impair lipolysis.

Day 8 adipocytes were transiently transfected with siCTRL, siCDS1 and siCDS2. Lipolysis was initiated by the addition of isoproterenol for 4h. Cells were lysed and proteins were extracted for immunoblotting against ATGL and HSL.

Under basal conditions, there was a significant decrease in the protein expression of

ATGL in both siCDS1 and siCDS2 adipocytes (Figure 4-3a). However, the protein expression of HSL and phosphorylated HSL did not change in siCDS1 and siCDS2 adipocytes. Upon isoproterenol treatment, the protein level of ATGL increased in both siCDS1 and siCDS2 when compared to the control (Figure 4-3b). While phosphorylated

HSL and total HSL appeared to have no significant change in siCDS1 adipocytes, there appeared to be a slight down-regulation in phosphorylated HSL of siCDS2 adipocytes

(Figure 4-3b). qRT-PCR revealed that the mRNA expression level of both isoforms of ATGL had decreased by ~ 60% in siCDS1 adipocytes (Figure 4-3c) and ~ 70% in siCDS2 adipocytes

(Figure 4-3d). Unlike the protein level, the mRNA level of HSL also appeared to decrease significantly in siCDS1 adipocytes, while siCDS2 appeared to have a moderate decrease. These findings implicate CDP-DAG synthases as potential regulators of lipolysis.

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A) B)

C) D)

Figure 4-3: CDP-DAG Synthases are Involved in Lipolysis. A) 3T3-L1 preadipocytes were differentiated until day 8. On day 8, adipocytes were split and transfected with siCTRL,

siCDS1 and siCDS2. Immunoblotting against ATGL, phospho-HSL and total-HSL was carried out. B) 3T3-L1 preadipocytes were differentiated until day 8. On day 8, adipocytes were split and transfected with siCTRL, siCDS1 and siCDS2. To initiate lipolysis, transfected adipocytes were treated with isoproterenol for 4h. Immunoblotting against ATGL, phospho-HSL and total-HSL was carried out. C) mRNA level of ATGL and HSL in basal siCDS1 adipocytes. p<

0.001 and p< 0.0001. D) mRNA level of ATGL and HSL in basal siCDS2 adipocytes. p< 0.01 and p< 0.0001.

4.5.1 Down-Regulation of CDS1 Diminished Phosphorylated Protein Kinase A (PKA) and Phosphorylated Perilipin Expression

Lipolytic rates are tightly regulated by PKA-mediated phosphorylation of perilipin1

[132]. Therefore, the protein expression levels of phosphorylated PKA and phosphorylated perilipin were investigated. Adipocytes were split on day 8 and

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transfected with CDS siRNA. For basal conditions, transfected adipocytes were left untreated prior to protein extraction. To induce lipolysis, transfected adipocytes were treated with 10nM isoproterenol for 4h. Western blot analysis with a phospho-PKA- substrate antibody showed that the phosphorylation of PKA at the 50 kDa mark in siCDS1 adipocytes had diminished compared to the control (arrow) (Figure 4-4a). In siCDS2 adipocytes, there appeared to be no change in the expression of p-PKA substrate. As the p-PKA substrate band displayed a decrease in protein expression at

~50 kDa, and phosphorylated perilipin is ~ 56 kDa, the reduced protein expression may coincide with perilipin. Furthermore, under basal and isoproterenol stimulated conditions, the expression of phospho-perilipin decreased significantly in siCDS1 adipocytes (Figure 4-4b). The expression of phospho-perilipin also appeared to decrease in siCDS2; however, its effect was not as dominant as that of CDS1. Total perilipin and actin showed that there were equal amounts of the respective proteins in each sample. To observe the difference in the expression of p-perilipin, a 3D view of the protein bands was carried out on the membrane. The 3D view confirmed the significant reduction of p-perilipin in siCDS1 adipocytes, as well as the moderate reduction in siCDS2 adipocytes (Figure 4-4c).

During lipolysis, through phosphorylation, perilipin1 interacts with HSL, to facilitate the translocation of HSL from the cytoplasm to the LD surface. The down-regulation of

CDP-DAG synthases could lead to an increase in PA. PA might accumulate on the surface of the LD, preventing perilipin from interacting with HSL and translocating to the LD surface. Also, the reduction in phosphorylated perilipin may suggest that CDP-

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DAG synthases are regulating adipocyte physiology i.e. perilipin is required on the surface of LDs; inhibiting CDS activity may affect perilipin function.

p-perilipin (3D)

Figure 4-4: Phospho-PKA and phospho-perilipin expression in siCDS1 and siCDS2 adipocytes during lipolysis. A) siCTRL, siCDS1 and siCDS2 adipocytes were blotted with phospho-PKA. B) siCTRL, siCDS1 and siCDS2 adipocytes were blotted with phospho-perilipin and total perilipin under basal and isoproterenol stimulated conditions. To induce for lipolysis, adipocytes were

treated with 10nM isoproterenol for 4h. C) 3D view of p-perilipin from Figure B.

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4.5.2 CDP-DAG Synthases Decrease Lipolytic Activity

Preliminary data from western blots have implicated a role for CDP-DAG synthases in lipolysis; therefore, the lipolytic rates were evaluated. The free glycerol assay measured the concentration of glycerol released from siCTRL, siCDS1 and siCDS2 adipocytes under basal and isoproterenol stimulated conditions. To induce lipolysis for the assay, transfected adipocytes were treated with 10µM isoproterenol and 10µM

IBMX for up to 4h. Aliquots of the media were assayed at 0m (untreated/basal), 10m,

30m, 60m and 240m.

Under basal conditions (0m), siCDS1 and siCDS2 adipocytes released slightly more glycerol than siCTRL adipocytes (Figure 4-5). After 10m of isoproterenol and IBMX treatment, the amount of glycerol released in siCTRL significantly increased compared to the amount released at 0m. At 30m and 60m, the amount of glycerol released decreased in siCDS1 and siCDS2 adipocytes, while the amount of glycerol released in siCTRL adipocytes increased (Figure 4-5). Two-way ANOVA showed that after 240m

(4h) of isoproterenol and IBMX treatment, the amount of glycerol released had significantly decreased in siCDS1 and siCDS2 adipocytes compared to 0m. As expected, the amount of glycerol released in siCTRL adipocytes had increased after 240m (4h) of isoproterenol and IBMX stimulation (Figure 4-5). Due to time constraints, this assay was carried out once. Additional assays would need to be performed to validate the amount of glycerol released in siCDS1 and siCDS2 adipocytes after 4h of stimulated lipolysis.

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Figure 4-5: Free glycerol assay during lipolysis in siCDS1 and siCDS2 adipocytes. Adipocytes were transfected with siRNA and treated with isoproterenol and IBMX. The media was

removed at 0m, 10m, 30m, 60m and 240m. At 0m, the amount of glycerol released in siCDS1 and siCDS2 adipocytes is higher than the siCTRL adipocytes. After 4h of induced lipolysis, the amount of glycerol released from siCDS1 and siCDS2 adipocytes had decreased compared to the glycerol released at 0m. p < 0.0001.

4.6 CDP-DAG Synthases are Implicated in Insulin Signalling

Emerging evidence has linked glycerolipid synthesising enzymes with mTOR regulation and insulin signalling [155]. During lipolysis, insulin acts as a powerful inhibitor. The effects are predominantly conveyed through the phosphorylation of insulin receptor substrates, the initiation of PI3K and the activation of Akt. To examine whether CDP-

DAG synthases might me involved in this pathway, the endogenous levels of CDS1 were observed. 3T3-L1 preadipocytes were differentiated until day 8. Mature adipocytes were examined under three conditions; adipocytes were left untreated

(basal), adipocytes were treated with 10nM isoproterenol for 4h and adipocytes were treated with 100nM wortmannin for 45m. Wortmannin is a potent inhibitor of PI3K,

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inhibiting its activity prevents the activation of Akt. Proteins were extracted and immunoblotted with anti-CDS1. Under basal and lipolytic conditions, the protein level of CDS1 did not change. However, upon wortmannin treatment, the endogenous protein levels of CDS1 increased (4-6a). During this study, no CDS2 mouse antibody had been manufactured, therefore, mRNA levels of CDS1 and CDS2 were also examined. qRT-PCR showed that the expression level of CDS1 was up-regulated by ~ 3- fold (Figure4-6b), while CDS2 activity increased by ~ 80% (~0.8 fold) (Figure 4-6c). The increase in protein and mRNA levels of CDP-DAG synthases indicated that CDS1 and

CDS2 might be involved upstream of the PI3K/insulin signalling cascade.

Figure 4-6: CDP-DAG synthase expression upon wortmannin treatment. A) Western blot of endogenous levels of CDS1 in basal, isoproterenol and wortmannin treated adipocytes.

B) qRT-PCR of CDS1 expression in basal and wortmannin treated adipocytes. C) qRT-PCR of CDS2 expression in basal and wortmannin treated adipocytes.

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4.6.1 Down-Regulation of CDP-DAG Synthases Increased the Expression of Phosphorylated Akt

Growth factors and insulin have been shown to activate Akt through PI3K [203]. The activation of Akt requires phosphorylation at threonine and serine residues. In adipocytes, the phosphorylation of Akt activates phosphodiesterase 3B, which degrades cAMP [204]. The degradation of cAMP reduces lipolysis by preventing the activation of PKA. The release of PKA blocks the phosphorylation-mediated activation of HSL and perilipin. To examine whether CDP-DAG synthases affect Akt phosphorylation during lipolysis, the protein levels of Akt were investigated in siCDS1 and siCDS2 adipocytes.

To measure the protein levels of Akt during lipolysis, 3T3-L1 preadipocytes were differentiated until day 8. Adipocytes were split and transfected with siCTRL, siCDS1 and siCDS2 for 48h. To stimulate lipolysis, transfected adipocytes were treated with

10nM isoproterenol for 4h. Proteins were extracted and immunoblotting was carried out using anti-phospho-Akt (S473/T389) and total Akt.

Upon isoproterenol treatment, there was no expression of phosphorylated Akt (S473).

However, western blotting against phosphorylated Akt (T389) showed a slight increase in siCDS1 adipocytes and a stronger increase in siCDS2 adipocytes (Figure 4-7a). 3D viewing confirmed that there was an increase. Western blotting was repeated in siCDS1 and siCDS2 adipocytes under basal and isoproterenol stimulated conditions to verify the increase in p-Akt (T389). Immunoblotting showed that in basal siCDS1 and siCDS2 adipocytes, the expression of p-Akt (T389) significantly decreased, while isoproterenol stimulated lipolysis confirmed the previous findings (Figure 4-7b). 113

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3D viewing confirmed that there was a decrease under basal conditions and an increase upon isoproterenol treatment. These findings suggested that CDP-DAG synthases may affect signalling pathways through Akt phosphorylation.

A) B)

Figure 4-7: Phosphorylated Akt Expression in siCDS1 and siCDS2 adipocytes. A) Transfected adipocytes were stimulated for lipolysis. Western blotting was carried out using anti-p-Akt (S473), anti-p-Akt (T389) and t-Akt. Transfected adipocytes were treated with 10nM isoproterenol to stimulate lipolysis. B) 3D view of image A. C) Basal and isoproterenol

stimulated lipolysis in siCTRL, siCDS1 and siCDS2 adipocytes. Western blotting was carried out using anti-p- Akt (T389).

4.6.2 CDP-DAG Synthases alter mTOR Expression during Lipolysis

Emerging evidence has mechanistically linked PA to mTOR, as well as lipolysis [155]. To understand whether the down-regulation of CDP-DAG synthases are involved in mTOR signalling during lipolysis, immunoblotting against mTOR and its downstream target,

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S6K was carried out. 3T3-L1 preadipocytes were induced for differentiation until day 8; mature adipocytes were split and transfected with siCTRL, siCDS1 and siCDS2 for 48h.

To induce lipolysis, transfected adipocytes were treated with 10nM isoproterenol for

4h. Western blotting was carried out using anti-phosphorylated-mTOR, anti-total- mTOR, anti-phosphorylated-S6K and anti-total mTOR. Under basal conditions in adipocytes, the expression of phosphorylated mTOR did not change (Figure 4-8a).

Upon isoproterenol treatment, the expression of phosphorylated mTOR slightly decreased in siCDS1 adipocytes and strongly decreased in siCDS2 adipocytes (Figure 4-

8a). This finding was confirmed by 3D viewing of the exposed membrane.

Interestingly, compared to the control, siCDS1 and siCDS2 adipocytes displayed a slight increase in the protein expression of phosphorylated S6K (Figure 4-8b). During isoproterenol stimulated lipolysis, there was a reduction of p-S6K in both siCDS1 and siCDS2 adipocytes. Western blots revealed a non-specific band > 75 kDa when immunoblotted against p-S6K. 3D viewing also confirmed the reduction of p-S6k in siCDS1 and siCDS2 adipocytes during lipolysis. The decrease of p-mTOR and p-S6K under lipolytic conditions suggests that CDP-DAG synthases might be involved downstream of the mTOR signalling pathway.

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Figure 4-8: mTOR expression during lipolysis in siCDS1 and siCDS2 adipocytes. 3T3-L1 pread ipocytes were induced for differentiation until day 8. Adipocytes were split and transfected with siCTRL, siCDS1 and siCDS2 for 48h. A) Western blots were carried out on transfected adipocytes using antibodies against p-mTOR and t-mTOR under basal and lipolytic conditions. 3D view of p-mTOR. B) Western blots were carried out on transfected adipocytes using antibodies against p-S6K and t-S6K under basal and lipolytic conditions. 3D view of p- S6K.

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4.7 Discussion

In this study, the role of CDP-DAG synthases in mature adipocytes, under basal and isoproterenol stimulated lipolysis were examined. Our preliminary findings indicate that CDP-DAG synthases may be involved in adipocyte development, particularly during lipolysis. As mentioned in Chapter 3, the down-regulation of CDS1 and CDS2 in

HeLa cells and 3T3-L1 preadipocytes resulted in the formation of SLDs. Consistent with this finding; adipocytes that were down-regulated with CDP-DAG synthase also exhibited the ‘supersized’ phenotype, with LDs up to 10µm in diameter. Current investigations have identified protein factors such as FSP27 as regulators of LD growth

[11, 83]. In adipocytes, FSP27 is localised to the LD surface (at the LD-contact site), where it facilitates the fusion and growth of LDs. Live cell imaging showed that in siCDS1 and siCDS2 adipocytes, LDs increased in size through fusion at an apparent LD- contact site. While fusion of small LDs was observed in siCTRL adipocytes, live cell imaging was unable to detect fusion of large LDs. Indicating that CDP-DAG synthases may be required in the fusion of LDs. The quick fusion observed in siCDS1 adipocytes suggests an FSP27-independent mechanism, possibly due to PA accumulation.

Evidence from Chapter 3 revealed that depleting CDS1 had a strong inhibitory effect on adipocyte differentiation, while CDS2 depletion had a moderate effect on the inhibition of adipocyte differentiation. This study hypothesised that PA may serve as a direct antagonist of PPARγ. Also, the accumulation of PA could confine PPARγ into certain membrane domains, preventing its activation. Interestingly, the down- regulation of CDS1 in mature adipocytes significantly inhibited the mRNA expression level of PPARγ and C/EBPα, while the down-regulation of CDS2 notably decreased

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C/EBPα activity. This further supports the conclusion that different species of PA may control different functions. Furthermore, it is possible that CDS1 is the main isozyme which is required for adipocyte development and maintenance.

In fatty liver dystrophic (fld) mice, constitutive loss of lipin1 results in complete inhibition of adipocyte differentiation [205]. Furthermore, recent data has associated lipin1-mediated PAP activity as a regulator in adipocyte lipolysis [135]. Therefore, this study investigated whether the down-regulation of CDP-DAG synthases would affect adipocyte lipolysis. Western blotting revealed the expression level of ATGL decreased under basal conditions in siCDS1 and siCDS2 adipocytes, while stimulated lipolysis increased the expression of ATGL in siCDS1 and siCDS2 adipocytes. Remarkably, the expression of phosphorylated-HSL showed no real change. At the mRNA level, the expression of ATGL and total HSL were down-regulated in siCDS1 and siCDS2 adipocytes. A possible explanation as to why the expression level of phosphorylated

HSL was not affected might be due to the depletion of CDP-DAG synthase. Down- regulating the activity of CDS could lead to an accumulation of PA at the LD surface; this accumulation might decrease the amount of perilipin which would prevent the phosphorylation of HSL to the LD surface, therefore inhibiting the initiation of hydrolysis. It is also possible that CDP-DAG synthases are involved in TAG hydrolysis but not DAG hydrolysis.

During lipolysis, perilipin is subject to phosphorylation by PKA. Additionally, perilipin interacts with HSL through phosphorylation, to translocate HSL to the LD surface and initiate DAG catabolism. Western blotting showed that the phosphorylated PKA substrate detected no change in phospho-Ser/Thr residues, except at ~ 50kDa in 118

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siCDS1 adipocytes, which may correlate with the expression of perilipin. The protein levels of perilipin were also examined under basal and isoproterenol treated conditions. Interestingly, under basal conditions, p-perilipin appeared to decrease in siCDS1 and siCDS2 adipocytes. Upon isoproterenol treatment, there was a significant reduction of p-perilipin in siCDS1 adipocytes, and a moderate reduction in siCDS2 adipocytes. In adipocytes, phosphodiesterase enzymes (PDE) can regulate the activity of PKA. The down-regulation of lipin1 in adipocytes has been shown to increase PDE activity upon PA accumulation [135]. As CDP-DAG synthase also utilise PA from the glycerol-3-phosphate pathway, it would be valuable to carry out a similar experiment for CDS1 and CDS2. It is possible that the down-regulation of CDS could cause an accumulation of PA, which could lead to increased PDE activity and decreased p-PKA and p-perilipin expression.

The free glycerol assay further confirmed that the down-regulation of CDP-DAG synthases impaired lipolysis to some extent. Prior to lipolytic stimulation, the amount of glycerol released in siCDS1 and siCDS2 adipocytes was greater than the control.

After 4h (240m) of isoproterenol and IBMX treatment, the amount of glycerol had decreased in siCDS1 and siCDS2 adipocytes, while the concentration of free glycerol increased in siCTRL adipocytes. Furthermore, statistical analysis confirmed the amount of glycerol released at 0m in siCDS1 and siCDS2 adipocytes had significantly decreased after 4h of stimulated lipolysis.

While PA has been implicated in lipolysis, recent studies have also shown a correlation of PA with insulin signalling and mTOR function [155]. Upon wortmannin treatment,

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the endogenous expression level of CDS1 increased at the protein level. qRT-PCR exhibited an increase of CDS1 and CDS2 at the mRNA level. These results indicate that

CDS1 and CDS2 might be involved upstream of the insulin signalling cascade. During lipolysis, the expression of phosphorylated-Akt (T308) in siCDS1 and siCDS2 adipocytes significantly increased, while basal adipocytes exhibited a significant decrease.

Phosphorylated-mTOR expression was also investigated in siCDS1 and siCDS2 adipocytes under basal and isoproterenol stimulated lipolysis. Under basal conditions, the expression of p-mTOR appeared to have no change amongst all siRNA transfected adipocytes. Upon isoproterenol treatment, the expression of p-mTOR appeared to decrease in siCDS1 and siCDS2 adipocytes. It is interesting to note that the downstream target of mTOR, S6K, when phosphorylated under basal conditions appeared to increase. In lipolysis-induced adipocytes, the expression of p-S6K slightly decreased. A negative feedback loop has been associated with mTOR and PI3K signalling. The activation of mTOR diminishes PI3K signalling by repressing the activity of IRS-1.

Therefore, future work would need to be performed to determine whether the down- regulation of CDP-DAG synthases affects mTOR expression and insulin signalling.

4.8 Conclusion

To summarise, this study has discovered novel functions of CDP-DAG synthases in adipocytes and during isoproterenol stimulated lipolysis. Down-regulation of CDP-DAG synthases in adipocytes resulted in the fusion of larger LDs. Further experiments would need to be carried out to determine whether the fusion is FSP27-mediated or FSP27- independent. While the study showed that CDP-DAG synthases in part, are involved in

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lipolysis, the most striking observation comes from the almost complete inhibition of p-perilipin in siCDS1 adipocytes. While siCDS2 adipocytes also had a strong decrease upon isoproterenol treatment, siCDS1 adipocytes exhibited a striking reduction.

Though it is unknown how CDS1 down-regulation inhibits the phosphorylation of perilipin, we hypothesise that the accumulation of PA on the membrane may prevent

HSL from translocating to the LD surface in order to initiate lipolysis. This data has also implicated a role for CDP-DAG synthases in insulin signalling. The findings from this study have shown that CDP-DAG synthases are more than enzymes in phospholipid synthesis; they are highly dynamic regulators of adipocyte function.

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5. CHAPTER 5: CDP-DAG SYNTHASE FUNCTIONALLY INTERACTS WITH FLD1

5.1 Background

Previous genome-wide screens identified Fld1p (yeast homologue of human seipin) as a major regulator of LD growth. Fld1p/Seipin has been implicated in the regulation and metabolism of fatty acids and phospholipids [6, 61, 156]. Recently, seipin has been shown to be involved in TAG synthesis, LD formation and adipocyte differentiation [63,

156]. Therefore, seipin has been implicated in the regulation of lipid storage, at both the cellular and systemic levels.

To identify additional ‘supersized’ candidates, further genome-wide screens in

Saccharomyces cerevisiae were carried out. The screen led to the identification of 10

SLD mutants, which were found to be phospholipid synthesising enzymes. We found that repressing CDS1 under the tet-regulatable promoter (here on referred to as tet-

CDS1), which encodes CDP-diacylglycerol synthase, can form giant LDs in yeast [53]. Of all the mutants identified in the screen, only FLD1 and CDS1 displayed the ‘supersized’ phenotype when grown in rich media (YPD media) (Figure 5-1). This led to the hypothesis that CDS1 and FLD1 may physically interact.

The aim of this study was to investigate whether CDS1 and FLD1 interact with one another. With the use of protein-protein interaction studies and functional studies, preliminary findings from this study have shown that Cds1p and Fld1p functionally interact.

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A) B)

Figure 5-1: Deletion of FLD1 and down-regulation of CDS1 results in ‘supersized’ lipid droplets. A) fld1∆ cells in YPD and SC media displayed SLDs. B) In the presence of doxycycline, tet-CDS1 cells displayed SLDs in YPD and SC media. Bar = 5µm

5.2 Fld1p Physically Interacts With Cds1p under the DUALmembrane Yeast Two Hybrid System

In order to identify protein-protein interactions between Cds1p and Fld1p, the

DUALmembrane system was employed [206]. This yeast-two hybrid system is used to identify protein-protein interactions between two integral membrane proteins. Firstly, the

TMHMM server was employed to predict the number of transmembrane domains in

Cds1p and Fld1p. The prediction showed that Fld1p has two transmembrane domains, while Cds1p was predicted to have 6 transmembrane domains in yeast (Figure 5-2a). A bait construct containing FLD1 was fused to the C terminal half of ubiquitin (pB3TC), while the prey construct containing CDS1 was fused to the N terminal half of ubiquitin (pPR3N).

A negative (pD2-Alg5) and positive control (pAI-alg5) were utilised from the

DUALmembrane system kit. An empty vector control (pB3TC) was also transformed with pPR3N-CDS1. The interaction between Fld1p and Cds1p resulted in transcriptional activation, which enabled the transformed yeast to grow on defined minimal medium lacking the reporter genes or the activation of lacZ encoding β-galactosidase (Figure 5-2b).

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X-gal overlay was carried out to confirm β-galactosidase was expressed. Cells expressing the control or the two interacting proteins displayed growth on the SC-leu-trp-his-ade plate, as well as the blue colourisation from the X-gal overlay. This indicated an interaction between Cds1p and Fld1p.

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Figure 5-2. Interaction of Fld1p and Cds1p using the DUALmembrane system. A) TMHMM predicted transmembrane domains of yeast Fld1p and Cds1p. B) DUALmembrane yeast two hybrid of transformed positive and negative controls along with pB3TC-FLD1: pPR3N-CDS1 and X-gal overlay. “+++” represents full growth and “-“represents no growth.

To further validate the physical interaction, a GST-pull down assay was carried out.

GST-FLD1 was co-transformed with CDS1-GFP in yeast, however, after a number of attempts, the pull down showed no noteworthy interaction. CDS1 was then used as the prey (GST-CDS1) and transformed with FLD1-flag, however, consistent with the previous finding, no real interaction was detected (data not shown).

5.3 Human CDP-DAG Synthase1 Does Not Physically Interact With Seipin

To determine whether CDS1 and Seipin interact in mammalian cells, co- immunoprecipitation was carried out on human CDS1 and human Seipin in HeLa cells.

Briefly, HeLa cells were co-transfected with the control, mCherry-N1 and pCDNA3-

CDS1-flag, or the interactors, Seipin-mCherry and pCDNA3-CDS1-flag. Western blots were carried out on the transfected cells to validate the protein expression levels. The size of mCherry-N1 was observed ~ 26kDa, while the size of CDS1-flag and Seipin- mCherry were ~ 53kDa and ~60kDa respectively (Figure 5-3-1a). Co- immunoprecipitation was carried out on the co-transfected HeLa cells, the lysate and eluates were loaded onto a SDS-PAGE gel. The input validated the expression of CDS1- flag in all samples. Interestingly, the western blot against RFP showed that the lysate of mCherry-N1: CDS1-flag displayed no RFP expression. However, its expression was detected in the eluates. Although CDS1 and seipin were detected, the IP revealed that

CDS1 was unable to pull down Seipin (Figure 5-3-1b). Instead, the eluates both had a

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significant expression level of mCherry alone. Therefore, at the mammalian level, CDS1 and Seipin do not physically interact.

Figure 5-3-1. Co-immunoprecipitation of CDS1 and Seipin. A) HeLa cells were transfected with CDS1-flag, mCherry-N1 and Seipin-mCherry . Proteins were extracted and loaded onto a SDS-PAGE gel. Blots were probed with flag-antibody and RFP-antibody. B) HeLa cells were co- transfected with mCherry-N1: CDS1-flag and Seipin-mCherry: CDS1-flag. Co- immunoprecipitation was carried out using Dynabeads and western blots were carried out using flag-antibody and RFP-antibody. The first two lanes of each gel represent the lysate, while the last two lanes on each gel are the eluates.

As mentioned in Chapter 3, this study has hypothesised that CDS1 is the major CDP-

DAG synthase. Therefore, most experiments were carried out on CDS1. However, as there are two CDP-DAG synthases, it is possible that CDS2 but not CDS1 may physically interact with Seipin. Due to time constraints, the interaction between CDS2 and Seipin remains to be investigated. To see whether the activity of CDS or Seipin was affected during gene-silencing, HeLa cells were transfected with siSeipin, siCDS1 and siCDS2. 126

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RNA was extracted and the cDNA was synthesised. qRT-PCR was carried out to determine the expression level of Seipin, CDS1 and CDS2 in the respective siRNA transfected cells. When Seipin was down-regulated by ~ 80%, the expression level of

CDS1 significantly increased by ~ 1.7-fold, while the expression of CDS2 slightly increased by ~ 20% (Figure 5-3-2a). In siCDS1 cells, the expression level of CDS1 decreased by ~ 90%, with no change in the expression level of Seipin (Figure 5-3-2b).

Interestingly, in siCDS2 cells, when CDS2 was down-regulated by ~ 80%, Seipin expression increased significantly by ~ 2-fold (Figure 5-3-2c).

B) C)

Figure 5-3-2. qRT-PCR expression in siRNA transfected cells. HeLa cells were transfected

with siRNA for 24-48h prior to RNA extraction and qRT-PCR. A) qRT-PCR of expression levels of Seipin, CDS1 and CDS2 in siSeipin HeLa cells. B) Expression levels of CDS1 and Seipin in siCDS1 HeLa cells. C) Expression levels of CDS2 and Seipin in siCDS2 HeLa cells. p < 0.001 and p < 0.0001.

5.4 Overexpression of CDS1 Partially Rescues the ‘Supersized’ Phenotype in fld1∆ Cells

Preliminary data have shown that CDS1 and FLD1 may not physically interact; therefore, the functional relationship between the two proteins in both yeast and

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mammalian cells was investigated. As the deletion of FLD1 in yeast cells forms SLDs, gene complementation studies were carried out to determine whether the overexpression of CDS1 in fld1∆ cells could rescue the ‘supersized’ phenotype. fld1∆ cells were transformed with CDS1-GFP and LDs were stained with Nile Red and observed using differential interference contrast (DIC) and GFP filter cubes. As GFP and

Nile Red have the same excitation and emission wavelengths, fld1∆ cells were overexpressed with CDS1-GFP and imaged without Nile Red. The LD phenotype was initially observed using DIC. LDs were then stained with Nile Red and counted to determine whether the size of LDs was reduced. In some fld1∆ cells, the overexpression of CDS1-GFP resulted in the restoration of the WT phenotype (Figure 5-

4-1). A cell count of 900 cells was carried out to identify whether the overexpression of

CDS1 rescued the ‘supersized’ phenotype in fld1∆ cells. Approximately 30% of the cells reverted to the WT LD morphology (Figure 5-4-1).

Figure 5-4-1. Overexpression of CDS1-GFP in fld1∆ cells. WT (BY7471) and fld1∆ cells were transformed with CDS1-GFP and grown in SC-leu with doxycycline. LDs were stained with Nile Red and observed using fluorescence microscopy. Bar = 5µm.

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To establish whether this finding was evolutionarily conserved, HeLa cells were transiently with siCDS1 and Seipin-mCherry. Transfected cells were treated with oleate and LDs were stained with BODIPY prior to fluorescence microscopy. Similar to Chapter

3, the overexpression of Seipin in HeLa cells inhibited LD synthesis (Figure 5-4-2a). The overexpression of Seipin-mCherry upon CDS1 knock-down shows a clear reduction in

LD size (Figure 5-4-2b). As CDS1 and Seipin do not physically interact, we hypothesise that the ‘rescued’ phenotype may be because Seipin is also involved in the phospholipid biosynthesis pathway, and that it may act upstream of CDS1.

Figure 5-4-2. Overexpression of Seipin upon oleate treatment A) HeLa cells were transfected with Seipin-mCherry and treated with oleate overnight. LDs were stained and fluorescence microscopy was carried out. B) HeLa cells were co-transfected with siCDS1 and Seipin-mCherry. Cells were treated with oleate overnight and LDs were stained with BODIPY. Arrow = transfected cells with mCherry signal.

5.5 Knockout of FLD1 in Tet-CDS1 Cells Forms “Supersized” Lipid Droplets

As mentioned previously, the deletion of FLD1 and down-regulation of CDS1 results in the formation of SLDs [53]. While both strains displayed few lipid droplets per cell, this study investigated whether a double knockout would cause a more dramatic LD

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phenotype. As CDS1 activity can only be repressed, FLD1 was deleted from Tet-CDS1 cells. The expression of CDS1 was then down-regulated in the presence of doxycycline.

The double knock-out mutant displayed SLDs which were greater than the ‘standard’

SLDs. In this instance, LDs were up to 4µm in diameter, which covered half the cell

(Figure 5-5-1). As CDS1 and FLD1 knock-down mutants display an increase in PA levels, it is probable that excessive PA accumulation has caused the ‘standard’ SLDs to coalesce.

Figure 5-5-1: FLD1 deletion from Tet-CDS1 cells. FLD1 was deleted from tet-CDS1 cells. Cells were grown in the presence of doxycycline and LDs were stained with Nile Red. Bar = 5µm

To further confirm the extremely ‘supersized’ phenotype, HeLa cells were down- regulated with siCDS1 or siCDS2 and siSeipin. Transfected cells were treated with BSA- coupled oleate overnight and imaged using fluorescence microscopy. The average SLD is > 4µm, while the SLDs seen in siCDS1siSeipin and siCDS2siSeipin cells were > 6µm

(Figure 5-5-2a). This supports the previous finding in yeast, and suggests an

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evolutionarily function link between the two proteins. The extremely ‘supersized’ phenotype may be due to the excessive accumulation of PA, which could promote the fusion of SLDs. To determine whether the total level of TAG increased, neutral lipids were extracted and total TAG was quantified in siCDS1siSeipin HeLa cells. Briefly, HeLa cells were co-transfected with siCDS1 and siSeipin. Transfected cells were treated with

BSA-coupled oleate overnight prior to neutral lipid extraction and TLC. After 14h of oleate treatment, siCDS1 cells displayed a ~ 60% increase in the total level of TAG, while siSeipin cells displayed an increase of ~ 25%. The double knock-down of CDS1 and Seipin revealed that the total amount of TAG increased by ~ 40% (Figure 5-5-2b).

180 B) 160

140 120 100 80 60 Relative Relative TAG 40 20 0

Figure 5-5-2: Double knock-down of CDP-DAG synthase and Seipin. A) HeLa cells were co- transfected with siCDS1siSeipin and siCDS2siSeipin. Cells were treated with oleate overnight and stained with BODIPY. Bar = 20µm. B) HeLa cells were transfected with siCDS1, siSeipin and siCDS1siSeipin. Transfected cells were treated with oleate overnight and neutral lipids were extracted for TLC. TAG bands were analysed using ImageJ software.

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5.5.1 Supersized Lipid Droplets Emerge from Clustering in siSeipin HeLa Cells

As mentioned in Chapter 3, SLDs emerged from clustered LDs in cells down-regulated with siCDS1 or siCDS2. Because fld1∆ cells displayed SLD mutants, this study hypothesised that the down-regulation of seipin (human homologue of FLD1) would also result in the formation of SLDs. HeLa cells were transfected with siSeipin and treated with oleate for from 4h-20h. Lipid droplets were stained with BODIPY and imaged using fluorescence microscopy. siSeipin cells displayed many, small and clustered LDs after 4h of oleate treatment (Figure 5-5-3). At 8h of oleate treatment, siSeipin cells were densely packed. This was consistent with the phenotype seen in siCDS1 and siCDS2 HeLa cells. By 10h of oleate treatment, siSeipin cells displayed large

LDs, while at 14h; the development of SLDs was fully established. At 20h of oleate treatment, siSeipin cells maintained the ‘supersized’ phenotype (Figure 5-5-3). These findings suggest a similar mechanism is responsible for the phenotype displayed in siCDS1/2 and siSeipin cells.

Figure 5-5-3. Down-regulation of Seipin in oleate treated HeLa cells. HeLa cells were transfected with siCTRL and siSeipin and treated with BSA-coupled oleate for 4, 8, 10, 14 and 20h. LDs were stained with BODIPY and imaged using fluorescence microscopy. Bar = 20µm

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5.6 CDS2 and Seipin Localise to the Endoplasmic Reticulum (ER)

Although co-immunoprecipitation studies between CDS1 and Seipin showed no interaction, this study also investigated the interaction between CDS2 and Seipin. For fluorescence microscopy, HeLa cells were transfected with CDS2-mCherry or EGFP-

Seipin. For co-localisation studies, HeLa cells were co-transfected with both constructs and subjected to fluorescence microscopy. Co-localisation studies revealed that both proteins localise to the ER. Interestingly, EGFP-Seipin alone displayed punctate structures (Figure 5-6-1a). This study found that when HeLa cells were co-transfected with EGFP-Seipin and CDS2-mCherry, the appearance of punctated structures was abolished, indicating a possible functional relationship between Seipin and CDS2

(Figure 5-6-1b). This was further supported by the presence of aggregates when HeLa cells were co-transfected with the ER marker, DsRed-ER, and EGFP-Seipin.

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Figure 5-6-1 Co-localisation of Seipin and CDS2. A) HeLa cells were transfected with EGFP- Seipin or CDS2-mCherry. Cells were fixed with 4% PFA and subjected to fluorescence microscopy. B) Co-transfection of EGFP-Seipin and DsRed-ER in HeLa cells, as well as EGFP- Seipin and CDS2-mCherry.

5.6.1 CDP-DAG Synthase2 and Seipin do not Interact by FRET/Acceptor Photobleaching (AB)

Acceptor photobleaching was carried out on HeLa cells transfected with EGFP-Seipin and CDS2-mCherry. Cells were scanned sequentially to confirm the localisation prior to bleaching. The region of interest was selected and bleached with the 561nm laser at

100% intensity. Co-localisation of EGFP-Seipin and CDS2-mCherry validated that both proteins localised to the ER (Figure 5-6-2a). The evaluation revealed no interaction between Seipin and CDS2 at the ER (Figure 5-6-2b). The blue coloured areas signified no interaction, while pink colouring represented a strong interaction. Although fluorescence microscopy revealed a small portion of the co-transfected cells did interact, the interaction was not observed throughout the whole cell. As FRET identifies molecules which are within close proximity to one another (10-100Å), it is

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possible that the interaction between CDS2 and Seipin interact may not be within

100Å.

Figure 5-6-2 FRET analysis of Seipin and CDS2. A) HeLa cells were co-transfected with EGFP- Seipin and CDS2-mCherry. B) Acceptor photobleaching of co-transfected HeLa cells. Evaluation of interaction after photobleaching.

5.7 Discussion

In an attempt to identify mutants with aberrant LD morphology, a genome wide screen of ~ 6000 (non-essential and essential) genes was carried out in S. cerevisiae [53]. Of all the strains screened, only FLD1 and CDS1 displayed the “supersized” phenotype under both rich and minimal growth conditions. This finding led to the hypothesis that CDS1 and FLD1 may physically interact. While the molecular function of FLD1 (yeast homologue of human Seipin) is not known, this study hypothesised that FLD1 may be

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involved in the phospholipid biosynthesis pathway, interacting with CDS1 and regulating the utilisation or production of PA.

To examine the functional interaction between CDS1 and FLD1, CDS1 was overexpressed under the native promoter in fld1Δ cells. The overexpression of CDS1 was able to rescue the “supersized” phenotype in ~ 30% of cells. The results in Chapter

3 implicated PA as a key regulator in LD expansion, hence, the restoration of the wild- type LD phenotype, may be due to the modulation of PA. While Fei et al., has shown that PA accumulates at the ER in fld1Δ cells, the data presented here suggests that the overexpression of CDS1 may reduce the level of PA. Therefore, through gene complementation, the level of endogenous PA could be normalised. This study also postulated that the interaction was conserved from yeast to humans; therefore, yeast and cell culture systems were employed. While yeast has one CDP-diacylglycerol synthase, mammals have two, CDS1 and CDS2. In HeLa cells, the overexpression of seipin inhibits LD synthesis, similar to the phenotype exhibited by CDP-diacylglycerol synthases. In HeLa cells, the down-regulation of CDS1 displayed SLDs, while the overexpression of Seipin was able to restore the LD morphology.

The physiological significance of the size of LDs remains poorly understood. However, the appearance of SLDs provides an efficient form of fat storage. The deletion of FLD1 from Tet-CDS1 cells resulted in a large unilocular LD in at least half of the cells. To further understand how this phenotype is affected at the quantitative level, future work would involve lipidomic analyses on total phospholipids and neutral lipids.

Additionally, this phenotype was further confirmed in HeLa cells when both CDS1 and

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CDS2 were knocked-down with Seipin. The double knock-down cells displayed SLDs that were ~30% larger than single knock-down cells. These preliminary findings have shown that in yeast and mammals, the down-regulation of CDS1 and Seipin form one

SLD per cell. A possible explanation for the single SLD observed is that excessive PA accumulation at the ER may enhance membrane curvature and promote quick fusion of SLDs. Mass spectrometry on siCDS1siSeipin and siCDS2siSeipin would need to be performed in the future to confirm whether an increase in PA is the key contributing factor in LD growth.

Quantification of the mRNA level of CDP-diacylglycerol synthases and Seipin revealed that there was a significant up-regulation in the activity of CDS1 but not CDS2, when

Seipin was down-regulated. During the down-regulation of CDS1, the expression of

Seipin did not change. Conversely, the down-regulation of CDS2 saw a significant increase in the expression level of Seipin. This finding indicates that Seipin may be involved in the phospholipid biosynthesis pathway. Because the down-regulation of

CDP-DAG synthases results in the formation of SLDs from clustered LDs, we wanted to see whether a similar phenotype emerged from siSeipin cells. Similar to CDS, siSeipin cells exhibited clustered LDs after 8h of oleate treatment, followed by the development of SLDs at 14h. In yeast, a genome wide screen identified SLDs, which had a common feature of an increase in PA. Therefore, SLDs in mammalian cells may develop from clustered LDs as a result of an accumulation of PA.

As mammalian cells have two CDP-DAG synthases, we hypothesised that Seipin might interact with one CDP-DAG synthase. BSCL2/Seipin has been shown to physically

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interact with transmembrane protein 19 (TMEM19), a paralog of CDS1 [207].

Therefore, it is possible that CDS2 and not CDS1 may interact with Seipin. Co- localisation studies confirmed that both CDS2 and Seipin localise to the ER.

Interestingly, the overexpression of Seipin resulted in cells displaying aggregates.

Preliminary data showed that the co-transfection of CDS2 with Seipin can prevent the formation of seipin aggregates. To identify whether there is a physical interaction between CDS2 and Seipin, FRET by acceptor photobleaching was carried out.

Fluorescence microscopy revealed that while CDS2 and Seipin do co-localise, no significant interaction was detected. FRET only detects donor and acceptor molecules which are within close proximity (10-100Å). Therefore, it is possible that CDS and

Seipin are further apart.

In order to identify protein-protein interactions between Cds1p and Fld1p, the

DUALmembrane system was employed. Growth on the selective plate signified a positive interaction, while the two negative controls were unable to grow on the quadruple dropout plate. To further validate the interaction, a GST pull-down assay was employed. The assay did not show any significant interaction between CDS1 and

FLD1. False positives are a common feature in yeast two hybrid assays; hence, co- immunoprecipitation was carried out in HeLa cells to confirm a physical interaction between the two proteins. CDS1-flag was unable to pull down Seipin-mCherry, thus, it seems that the interaction between CDP-DAG synthases and Seipin appear to be functional rather than physical. However, additional experiments employing different detergents and buffer conditions are required to conclusively determine the physical relationship between seipin and the CDS proteins.

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5.8 Conclusion

To identify novel gene products which regulate the development of LDs, a genome wide screen was carried out in S. cerevisiae. The initial screen identified Fld1p as a major regulator of LD growth. The human homologue of FLD1, Seipin, has been implicated in LD and TAG synthesis. The down-regulation of CDS1 also displayed the

“supersized” phenotype under rich and minimal growth conditions. Therefore, protein- protein interaction studies were employed to understand whether these two proteins interact. While protein-protein interaction showed no conclusive physical interaction, preliminary studies indicated that CDS1, CDS2 and Seipin functionally interact. As the physical data remain inconclusive, future experiments should employ a variety of lysis buffers and detergents to verify the interaction. While the molecular function of Seipin is not known, CDP-DAG synthase stand as a potential interactor which may help to understand the molecular function of seipin.

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6. CHAPTER 6: FUTURE DIRECTIONS AND FINAL CONCLUSIONS

This thesis has characterised CDP-DAG synthases as regulators of LD growth, adipocyte development (Chapter 3) and function (Chapter 4) for the first time in cultured mammalian cells. In addition, this study has identified a potential functional relationship between two SLD mutants, CDS1 and FLD1 (Chapter 5). In summary, this study has discovered novel functions of a classical enzyme in phospholipid metabolism,

CDP-DAG synthase. Our results implicate PA as a key regulator of LD expansion and adipogenesis, and highlight the important cellular and developmental functions of phospholipids.

6.1 CDP-DAG Synthases Regulate Lipid Droplet Growth and Adipocyte Development

This study provides an intimate link between the expansion of LDs at the cellular level and the differentiation of adipocytes at the systemic level. It suggests that phospholipids, i.e. phosphatidic acid (PA), may play an important role in the regulation of two apparently distinct processes of lipid storage. Given the emerging roles of LDs in human health and energy production, understanding the molecular mechanisms involved in LD dynamics and adipocyte development will provide insights into how therapeutic strategies can be developed against human metabolic diseases.

Additionally, identifying LD regulators may help us understand how the human body controls excess lipid intake with today’s high fat diet.

CDP-DAG synthase is an evolutionarily conserved key enzyme in phospholipid metabolism. There is one CDS gene in yeast and fly, but there are two CDS genes in mammals [165, 208]. In humans, CDS1 and CDS2 share 73% identity and 92%

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similarity. The down-regulation of CDS1 and CDS2 lead to the formation of SLDs, a phenotype which appears to be evolutionarily conserved from yeast to mammals.

While the physiological significance of SLDs remains to be investigated, it seems that

LD size is a tightly regulated process. Giant or “supersized” LDs can often be formed in diseased tissues and cells [209].

This study showed that CDS1 appears to be the main isozyme, as knocking down CDS1 led to drastic changes in the total level of PA. CDS1 and CDS2 also appear to be differentially regulated as CDS1, but not CDS2 is up-regulated during adipocyte differentiation. To determine whether CDS1 is the major enzyme, future work on the enzymatic activity of CDS1 and CDS2 should be carried out. Although PA accumulated strongly in the ER of siCDS1 and siCDS2 cells, the total level of PA only significantly increased in siCDS1 cells. Also, the down-regulation of CDS1 strongly inhibited the differentiation of preadipocytes, while the down-regulation of CDS2 moderately impaired this process. Therefore, CDS activity would also need to be measured under these conditions to validate the hypothesis that CDS1 is the major isozyme.

During this study, there was no commercially available CDS2 antibody with known species reactivity in mice. Hence, the protein expression of down-regulated CDS2 during adipocyte differentiation cannot be detected. Interestingly, the down- regulation of CDS1 and CDS2 in HeLa cells led to an increase in the total level of PG and

PI. Future work would involve performing double knock-down experiments to determine whether the increase in PG or PI was based on a compensatory effect during single siRNA transfections. The overexpression of CDS1 and CDS2 inhibited LD

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synthesis. CDS activity assays would need to be performed in the future to ensure the over-expressed protein was functional. To establish whether the enzymatic activity of

CDS1 and CDS2 is important in LD expansion, future work would involve designing a point mutation which is defective in enzymatic activity.

To determine whether PA and not TAG was the critical factor in LD expansion, the PA phosphatase, lipin-1 was knocked-down. Lipin1 converts PA to DAG, which is the precursor for PE, PC and TAG synthesis in mammals. Upon lipin1 down-regulation,

HeLa cells displayed SLDs. Overexpression of either CDS1 or CDS2 rescued the LD morphology to that of the control cells. Therefore, it was concluded that PA, but not

TAG, plays a critical role in SLD formation. Future work would involve measuring the level of PA in the lipin1 knock-down cells in the presence/absence of CDS1 and CDS2 overexpression. As the role of PA in LD formation is an important conclusion from this study, its level should be also measured. Furthermore, the PA phosphatase activity of lipin1, unlike its catalytically inactive site, has been shown to be involved in adipocyte differentiation [191]. To further demonstrate the requirement of PA in adipogenesis, the level of PA should be reduced by overexpressing LPIN1. We hypothesise that a reduction in PA should increase adipogenesis, only if it is inhibited as a result of excessive endogenous PA accumulation (i.e. knock-down of CDS1). Therefore, future work would involve knocking down CDS1 or CDS2 and overexpressing LPIN1. By manipulating the expression of these enzymes, the total level of PA could be

“normalised” to rescue the inhibition of differentiation in preadipocytes, caused by the elevated level of PA.

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This study has also provided evidence for the role of CDP-DAG synthases in lipid storage at the systemic level. We hypothesised that the down-regulation of CDS would cause an accumulation of PA, and that the increase of PA would have tissue specific functions, i.e. inhibition of adipocyte development. As PA has also been shown to be a regulator of LD growth, different species of PA may have different functions. As discussed in Chapter 3, the master driver of adipogenesis, PPARγ, has a large ligand binding pocket that can be activated by various metabolites that originate from phospholipid and fatty acids [96]. Although there are plenty of naturally occurring molecules that trigger PPARγ in cell-based assays, the endogenous regulators of PPARγ that are of physiological importance remain poorly defined.

The recent identification of cyclic PA (cPA) as an antagonist of PPARγ adds another layer to the regulation of PPARγ activity [193, 194]. However, unlike PA, cPA is a lysophosphatidic acid which does not promote membrane curvature. Furthermore, cPA synthesising enzymes have not been associated with human metabolic diseases such as lipodystrophy. Congenital generalized lypodystrophy (CGL) is characterised by the complete absence of adipose tissue, resulting in hepatic steatosis [210]. As mentioned in Chapter 1, glycerolipid synthesising enzymes such as AGPAT2 and lipin-1

(regulate PA metabolism) have been associated with this disorder. Exactly how the accumulation of PA inhibits adipocyte differentiation remains to be investigated. In addition, identification of PA species which have a high affinity for PPARγ needs to be determined.

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To summarise, the data presented in this study has given exciting new life to CDP-DAG synthase, a classical enzyme in phospholipid synthesis. The novel functions of CDP-

DAG synthase in lipid storage may provide important cellular and developmental functions of phospholipids.

6.2 The Characterisation of CDP-DAG Synthases in Mature Adipocytes

This study has provided preliminary evidence that CDP-DAG synthase may be involved in a number of adipocyte functions. Similar to Chapter 3, the preliminary data produced here has shown for the first time that CDP-DAG synthase is an enzyme of diverse functions. In mature adipocytes, the down-regulation of CDS1 and CDS2 led to the formation of fewer and larger LDs per cell. Future work would involve carrying out mass spectrometry to determine whether the knock-down of CDS increases the total amount of PA in mature adipocytes. Recent studies have shown an increasing interest in the fusion of LDs, particularly in adipocytes. In adipocytes, FSP27, a novel regulator of LD growth, promotes the transfer and expansion of LDs at the LD contact site (LDCS)

[83]. Enrichment of FSP27 at the LDCS could promote LD stability to allow for the efficient transfer of neutral lipids. The down-regulation of CDS1 and CDS2 led to the fusion of LDs in mature adipocytes. Future work would determine whether this fusion is FSP27 mediated or FSP27-independent. Also, the expression of FSP27 should be looked at in mature adipocytes which have depleted CDS1 or CDS2.

Adipocyte maintenance was briefly investigated. The down-regulation of CDS1 led to a significant decrease in PPARγ and C/EBPα. As discussed above, PA may serve as a

PPARγ antagonist. CDS2 down-regulation decreased the expression of C/EBPα but not

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PPARγ. An enzyme activity assay should be performed in the future to measure whether there expression of CDS1 and CDS2 various in adipocytes.

Preliminary findings from this study showed that CDP-DAG synthases are involved in lipolysis. While the expression of ATGL decreased when CDS1 and CDS2 were down- regulated, HSL showed no change. To establish whether ATGL and CDP-DAG synthase physically interact, protein-protein interaction studies such as co-immunoprecipitation could be carried out. Also, upon lipolytic stimulation, the expression of phosphorylated perilipin significantly decreased, particularly in siCDS1 adipocytes. Additional studies in the future would need to be performed to understand whether CDP-DAG synthase and perilipin/ the PKA signalling pathway (see below) are coordinated during lipolysis.

Furthermore, the expression of perilipin in siCDS1 and siCDS2 adipocytes under basal and lipolytic conditions should also be investigated. Moreover, the amount of glycerol released would need to be determined upon lipolytic stimulation. Preliminary data showed a reduction in the concentration of glycerol released in siCDS1 and siCDS2 adipocytes; however, a larger reduction was expected.

Mitra et al., showed that the loss of lipin1 resulted in an accumulation of PA which activated phosphodiesterase enzymes (PDE) through direct allosteric interaction. In adipocytes, cAMP-degrading PDE enzymes can regulate PKA signalling [135]. The study found that the PAP activity was responsible for the control of PA accumulation in the organisation of PDE activity in adipocytes, as well as mTOR signalling. As CDS also utilise PA, it would be of interest to measure PDE activity in mature adipocytes down- regulated with CDS1 or CDS2. While the data presented here showed that the

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expression of mTOR decreased upon isoproterenol treatment, the data is preliminary and further studies need to be performed. Similar to Mitra et al., it would be useful to treat siCDS1 and siCDS2 adipocytes with an mTOR inhibitor, Torin, and measure the activity of PDE. If the activity is increased, we would assume that the accumulation of

PA from CDS inhibition also regulates PDE activity and affects mTOR signalling.

Preliminary findings from this study have implicated CDP-DAG synthase in Akt and insulin signalling. In adipocytes treated with wortmannin, the expression of CDS1 increased. While there was no available mouse-specific CDS2 antibody to check the protein expression, the mRNA levels of both CDS1 and CDS2 increased. Further work needs to be carried out to understand whether and how CDS might be implicated in insulin signalling. It would be noteworthy to measure the total level of PI in adipocytes down-regulated with CDS1 or CDS2, and uncover whether CDS can control the insulin pathway at the level of phospholipids.

The preliminary findings in this chapter have provided novel functions for CDP-DAG synthase in adipocyte functions. Here, we have implicated CDP-DAG synthases in LD fusion and isoproterenol stimulated lipolysis. However, future work would need to be performed to comprehend how CDS are involved in lipolysis and whether the down- regulation of CDS could control the activity of lipolytic enzymes. In addition, extensive lipidomic analyses would need to be carried out to understand if PA and other phospholipids are accumulated upon CDS down-regulation.

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6.3 CDP-Diacylglycerol Synthase Functionally Interacts with FLD1

The reverse genetic screen of the entire essential and non-essential collection in S. cerevisiae was a powerful tool in identifying aberrant LD morphology. As CDS1 and

FLD1 were the only mutants to display the “supersized” phenotype under both rich and minimal growth conditions, we wondered whether the two proteins physically interact. This study identified the functional relationship between Cds1p and Fld1p.

As Seipin has established roles in TAG synthesis, adipogenesis and LD growth, it is possible that Seipin may be involved in the phospholipid biosynthesis pathway. As

CDS1 and CDS2 are enzymes involved in phospholipid synthesis, identifying potential

Seipin interactors was of interest. Interestingly, the overexpression of CDS1, CDS2 or

Seipin inhibited the development of LDs, while the down-regulation led to the formation of SLDs through clustering.

Preliminary protein-protein interaction studies were carried out; however, the data remains inconclusive on whether the two proteins physically interact. Future work would involve examining a variety of detergents and buffer conditions to optimise physical interactions between the transmembrane proteins. A recent study by Sim et al

[211], revealed seipin and lipin interact as a result of co-immunoprecipitation. Co- transfection of seipin and lipin showed that lipin1α and lipin1β were detected in the precipitates. Furthermore, the interaction required the N and C terminals of seipin. As lipin is a PA phosphatase which converts PA to DAG for TAG synthesis, it would be useful to perform the same experiment using the CDS proteins.

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Although the physical interaction between Seipin and the CDS proteins remains inconclusive, this study discovered a functional relationship between the transmembrane proteins. The overexpression of CDS1-GFP in fld1Δ yeast cells rescued the LD morphology to that of the WT in approximately one third of the cells. Similarly, in HeLa cells when CDS1 was down-regulated, the overexpression of Seipin restored the normal LD morphology. To further confirm a functional relationship between

Seipin and the CDS proteins, a CDS activity assay should be carried out in yeast and cell culture systems. In particular when CDS and Seipin are down-regulated and also when the proteins are overexpressed. Measuring the enzymatic activity would also confirm the functionality of the CDS proteins and determine whether their expression varies when transfected with Seipin.

In mammals, as there are two CDP-DAG synthases, it is possible that seipin may only interact with one of the proteins. A study by Rual et al., showed that BSCL2 (encodes for Seipin) physically interacts with transmembrane protein 19 (TMEM19), a paralog of

CDS1 [207]. Therefore, it is plausible that because BSCL2 interacts with a paralog of

CDS1, it may not interact with CDS1 but CDS2 instead. Future studies such as co- immunoprecipitation should also be performed to investigate the physical interaction between CDS2 and Seipin. Due to time constraints, only functional studies were performed using CDS2 and Seipin. Co-localisation studies confirmed the ER localisation of the transmembrane proteins. While the co-transfection of CDS2 and Seipin prevented the formation of seipin aggregates.

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This study has provided preliminary evidence for the functional relationship between the CDS proteins and Seipin/Fld1p. While the molecular function of seipin is unknown, here this study has recognised potential interactors of seipin which may help to identify its function in the future. Additional experiments utilising different lysis conditions and buffers are needed to establish whether the CDS proteins and

Seipin/Fld1p physically interact.

6.4 Putting it all Together – CDP-Diacylglycerol Synthases and Human Lipid Storage

Recent studies on phosphatidic acid have revealed the cone-shaped lipid as a key regulator in LD growth, lipolysis, mTOR and insulin signalling [53, 135, 155]. These findings have been associated with enzymes such as GPAT, AGPAT and lipin. However, no studies have been performed using CDS to modulate the endogenous levels of PA.

In yeast, the formation of SLDs is due to an increase in total cellular and microsomal PA

[53]. The PA phosphatase activity of lipin has been shown to be a regulator of adipocyte differentiation [191]. In addition, increased levels of PA from Lipin1-/- mice have been shown to impair lipolysis and mTOR through enhanced PDE activity [135].

While PA synthesised from the glycerol-3p-phosphate pathway has been shown to impair mTORC2 activity and suppress insulin signalling [155].

The key findings in this thesis have implicated CDP-DAG synthases in LD expansion and adipocyte differentiation, two major features of lipid storage. Preliminary data from this study has also implicated CDP-DAG synthases in a number of adipocyte functions; in particular, isoproterenol stimulated lipolysis. We hypothesised that the down- regulation of CDS1 or CDS2 would cause an accumulation of PA which would lead to 149

Chapter 6: Future Directions and Final Conclusions

the formation of SLDs, inhibition of adipocyte differentiation and impair lipolysis

(Figure 6-1). This thesis has shown that the down-regulation of CDS1 and CDS2 in HeLa cells leads to the accumulation of PA. The fact that PA has been shown to be involved in many processes sheds light on the importance of phospholipids as regulators of mammalian lipid storage and development.

Figure 6-1. Down-regulation of CDP-DAG Synthase in PA metabolism. The down-regulation of CDP-DAG synthase from the phospholipid synthesis pathway would cause an accumulation of PA. An increase in the level of PA may inhibit adipocyte differentiation, impair lipolysis and regulate LD growth. Glycerol-3-phosphate (G-3-P) is converted to lysophosphatidic acid (LPA), which is then converted to phosphatidic acid (PA).PA can be utilised by CDP-diacylglycerol synthase (CDS) and converted to CDP-DAG which is the precursor to the synthesis of phosphatidylinositol(PI) and phosphatidylglycerol (PG). Alternatively, PA can be metabolised by LPIN and converted to diacylglycerol (DAG), which is the precursor to the synthesis of phosphatidylethanolamine (PE), phosphatidylcholine (PC) and triacylglycerol (TAG).

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This study has uncovered important novel functions of CDS in cellular and systemic lipid storage. This project found that CDS enzymes are not only required for the normal proliferation and expansion of cellular lipid droplets, but are also required for the development of adipocytes. These results further suggest that CDS enzymes exert their effects by modulating the level and distribution of PA, which is a key regulator of the

LD expansion and adipogenesis. Preliminary findings have also implicated a role for the classical enzyme in adipocyte functions, particularly, lipolysis. These results therefore highlight the important cellular and developmental functions of phospholipids.

Understanding the molecular mechanisms that govern LD expansion and adipocyte differentiation will provide insights into the development of therapeutic strategies targeted against human metabolic diseases.

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Appendix

APPENDIX

Figure Legends of Digital Files

Video 1a (Chapter 3): 3D imaging of siCTRL HeLa cells with overnight oleate treatment.

Video 1b (Chapter 3): 3D imaging of siCDS1 HeLa cells with overnight oleate treatment.

Video 1c (Chapter 3): 3D imaging of siCDS2 HeLa cells with overnight oleate treatment.

Video 2 (Chapter 3): 3D imaging of siCDS1 HeLa cells after 8h of oleate treatment.

Video 3a (Chapter 4): Live cell imaging of siCTRL mature adipocytes.

Video 3b (Chapter 4): Live cell imaging of siCDS1 mature adipocytes (1).

Video 3c (Chapter 4): Live cell imaging of siCDS1 mature adipocytes (2).

Video 3d (Chapter 4): Live cell imaging of siCDS2 mature adipocytes.

168