The Effects of Ceramide Synthases and Protein Kinase C Epsilon on Glucose Homeostasis and Lipid Metabolism

THE EFFECTS OF CERAMIDE SYNTHASES AND PROTEIN KINASE C EPSILON ON GLUCOSE HOMEOSTASIS AND LIPID METABOLISM

Barbara Diakanastasis

Supervisor: Dr Carsten Schmitz-Peiffer

A thesis in fulfilment of the requirements for the degree of Master of Science (Research)

St Vincent’s Clinical School Faculty of Medicine University of New South Wales, Australia

2015

PLEASE TYPE THE UNIVERSITY OF NEW SOUTH WALES , Thesis/Dissertation Sheet

Surname or Family name: Diakanastasis

First name: Barbara Other name/s:

Abbreviation for degree as given in the University calendar: MSc

School: St. Vincent's Clinical School Faculty: Medicine

Title: The Effects of Ceramide Synthases and Protein Kinase C epsilon on Glucose Homeostasis and Lipid Metabolism

Abstract 350 words maximum: (PLEASE TYPE)

Insulin resistance contributes strongly to Type 2 Diabetes, which is a global epidemic. A strong link exists between dietary lipid excess and the development of insulin resistance. Two key bioactive lipid metabolites implicated in the development of insulin resistance are the sphingolipid ceramide and diacylglycerol, an activator of protein kinase c epsilon (PKCe). The mechanisms linking the actions of these metabolites to insulin resistance are unclear. The aims of this thesis were to (i) assess the effect of ceramide synthase (CerS) overexpression on key elements of skeletal muscle sphingolipid metabolism and insulin action and (ii) examine the effect of PKCe deletion solely in adipose tissue on glucose and lipid homeostasis.

CerS isoforms were overexpressed in lipid-treated LS skeletal myotubes. This did not cause compensatory changes in the mRNA expression of sphingolipid metabolism proteins. The effect of CerS overexpression on flux through ceramide synthesis pathways was assessed. Increased flux via the salvage pathway was seen after CerS overexpression. In addition, GLUT4 translocation to the plasma membrane, a key aspect of skeletal muscle insulin action, was increased or unaltered after CerS overexpression.

Glucose tolerance was tested in fat-fed mice with PKCe deletion only in adipose tissue. Improved glucose tolerance was seen in these mice during short and long term fat feeding. This was linked to a greater decrease in plasma fatty acids and without compensatory rises in insulin secretion, suggesting this deletion enhanced whole body insulin sensitivity. This was confirmed by euglycaemic-hyperinsulinaemic clamp. Improved hepatic insulin sensitivity, potentially via decreased plasma fatty acids, was shown to drive this improvement. PKCe deletion in adipose tissue was also linked to smaller adipocyte size, an indicator of improved insulin sensitivity. Gene expression analysis showed that these protective effects were not via altered mRNA expression of lipid metabolism or inflammatory genes. Increased protein expression of lipid esterification enzymes however, was observed.

This research has highlighted a novel protective role for ceramide during skeletal muscle insulin resistance upon specific CerS isoform overexpression. We have also shown for the first time that (i) exclusive PKCe deletion in adipose tissue improves glucose and lipid homeostasis during insulin resistance (ii) hepatic insulin sensitivity is modulated indirectly by PKCe.

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‘I hereby grant the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all proprietary rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation. I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstract International (this is applicable to doctoral theses only). I have either used no substantial portions of copyright material in my thesis or I have obtained permission to use copyright material; where permission has not been granted I have applied/will apply for a partial restriction of the digital copy of my thesis or dissertation.'

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2 ACKNOWLEDGEMENTS I would firstly like to thank my supervisor Dr Carsten Schmitz-Peiffer for his never- ending guidance, encouragement and training throughout my years at the Garvan Institute. This has truly enabled me to be a more perceptive, resilient and well-rounded researcher. I take this opportunity to wish you all the very best with your future endeavors. I would also like to thank my co-supervisor Professor Trevor Biden for his unwavering support and valuable input during my studies and for his strong leadership of the Cell Signalling Laboratory, which has helped it become the cohesive and dynamic team it is today. My gratitude extends to our collaborators Dr Amanda Brandon and Eurwin Suryana for performing the euglycaemic-hyperinsulinaemic clamp experiments and furthermore, for Dr Brandon’s expertise in the interpretation of associated data. Without this invaluable contribution, some of the key findings in this study would not have been attained.

I am immensely grateful to those who have assisted me by kindly donating their time, expertise and reagents toward various aspects of this project: Dr Amira Klip for her gift of the L6 rat skeletal muscle cell line, Professor David James for his gift of the AdipoQ- cre mice line, Professor Robert Brink for his gift of the Flp deleter mice line, Professor Anthony Futerman for his donation of ceramide synthase-containing mammalian plasmid constructs, Mana Liao for cloning the ceramide synthase adenoviral constructs prior to the commencement of this study, Dr Nolan Hoffman for his expert advice on the GLUT4 translocation assay, associated data analysis and donation of specialized L6 cells and other reagents for this technique, Dr Daniel Fazakerley for his expert advice on the GLUT4 translocation assay and associated data analysis, Dr Dale Hancock for performing key immunoblots for the protein kinase C epsilon study and for her theoretical input toward the ceramide-synthase and protein kinase C epsilon studies (and the Messina-related work!), Liam O’Reilly for his advice on the RT-PCR technique and associated data interpretation in addition to helping execute the plasma cytokine measurements and perform interpretation of associated data, Dr Michael Swarbrick and Rebecca Stuart for technical advice regarding the fun-filled extraction of RNA from adipose tissue and finally Dr Nancy Sue and Dr Jeng Yie Chan for their advice on the RT-PCR technique and associated data interpretation.

3 To all my fellow colleagues at the Cell Signalling Laboratory, both past and present - you have made my experience at the Garvan Institute very rewarding and memorable indeed. Your love of science has inspired me to push further to find the answers and to develop a deep respect for the research itself. I will miss our lunchtime hang-outs, post- talk drinks in de novo, Darlo Bar sessions, and all the memories shared during our “Christmas” parties. In particular, I would like to thank Ebru Boslem, Georgia Frangioudakis, Mana Liao, Gemma Pearson and Liam O’Reilly for their friendship and guidance over the years.

My final thanks are reserved for my family and friends especially my mother, my father, my parents-in law and my best friend Anthea. I would like to thank you for all your support, patience and understanding during this journey. You have all been there for me unconditionally and I look forward to spending more time with you. To my husband Theo, I love you. Though you have never stepped foot in a lab you have instilled in me many essential qualities that are required to be a good researcher and more importantly, a good human being. This thesis is for you.

4 ABSTRACT Insulin resistance contributes strongly to Type 2 Diabetes, which has become a global epidemic. A strong link exists between dietary lipid excess and the development of insulin resistance. Two key bioactive lipid metabolites implicated in the development of insulin resistance are the sphingolipid ceramide and diacylglycerol, an activator of protein kinase c epsilon (PKCε). The mechanisms linking the actions of these metabolites to insulin resistance remain unclear. The aims of this thesis were to (i) assess the effect of ceramide synthase (CerS) overexpression upon key elements of skeletal muscle sphingolipid metabolism and insulin action and (ii) examine the effect of PKCε deletion specifically in adipose tissue on glucose and lipid homeostasis. Firstly, CerS isoforms were overexpressed in lipid-treated L6 skeletal myotubes with adenovirus. This did not cause compensatory changes in the mRNA expression of sphingolipid metabolism proteins. The effect of CerS overexpression on flux through ceramide synthesis pathways was also assessed. Increased flux via the salvage pathway was seen after CerS overexpression. In addition, glucose transporter 4 translocation to the plasma membrane, a key aspect of skeletal muscle insulin action, was increased or unaltered following CerS overexpression. Secondly, glucose tolerance was tested in fat-fed mice containing PKCε deletion only in adipose tissue. Improved glucose tolerance was observed in these mice during short and long term fat feeding. This was linked to greater suppression of plasma fatty acids and without compensatory rises in insulin secretion, suggesting this deletion enhanced whole body insulin sensitivity. Investigation via euglycaemic-hyperinsulinaemic clamp confirmed this. Improved hepatic insulin sensitivity, potentially from observed decreases in plasma fatty acids, was shown to drive this improvement. PKCε deletion in adipose tissue was also linked to smaller adipocyte size – an indicator of improved insulin sensitivity. Gene expression analysis showed that these protective effects were not caused by altered mRNA expression of lipid metabolism or inflammatory genes. Increased protein expression of lipid esterification enzymes however, was observed. This research has highlighted a novel protective role for ceramide during skeletal muscle insulin resistance upon specific CerS isoform overexpression. We have also shown for the first time that (i) PKCε deletion exclusively in adipose tissue improves glucose and lipid homeostasis during insulin resistance (ii) hepatic insulin sensitivity is modulated indirectly by PKCε.

5 TABLE OF CONTENTS

CERTIFICATE OF ORIGINALITY ...... 1

ACKNOWLEDGEMENTS ...... 3

ABSTRACT ...... 5

TABLE OF CONTENTS ...... 6

LIST OF FIGURES AND TABLES ...... 10

PUBLICATIONS ...... 12

CONFERENCE PRESENTATIONS ...... 12

CHAPTER 1 GENERAL INTRODUCTION ...... 13 1.1 Glucose Homeostasis, Lipid Homeostasis and Type 2 Diabetes ...... 14 1.2 The Insulin Signalling Pathway and its Metabolic Actions ...... 16 1.3 Obesity, Diabetes and the Metabolic Syndrome ...... 20 1.4 Insulin Resistance ...... 21 1.5 The Role of Lipids in the Pathophysiology of Insulin Resistance ...... 23 1.6 Lipid-Induced Skeletal Muscle Insulin Resistance ...... 24 1.7 Mechanisms of Ceramide-Induced Insulin Resistance ...... 25 1.7.1 Inhibition of insulin signalling ...... 25 1.7.2 Inflammation ...... 26 1.7.3 Mitochondrial dysfunction ...... 26 1.7.4 Apoptosis ...... 27 1.7.5 Lipid raft alterations ...... 27 1.8 Sphingolipid Metabolism and Ceramide Synthase ...... 28 1.8.1 Ceramide metabolism ...... 28 1.8.2 Ceramide synthases ...... 31 1.8.3 Regulation of sphingolipid metabolism enzymes ...... 33 1.9 The Concept of “Many Ceramides” ...... 34 1.10 The Protective Role of Ceramide in the Pathogenesis of Insulin Resistance 35 1.11 Lipid-Induced Hepatic Insulin Resistance ...... 36 1.12 The Functional Roles of Adipose Tissue ...... 39 6 1.12.1 Lipid metabolism in adipose tissue ...... 40 1.12.1.1 Fatty acid esterification ...... 40 1.12.1.2 Lipolysis ...... 40 1.12.1.3 De novo lipogenesis ...... 43 1.12.2 Adipokine secretion ...... 43 1.12.3 Functional differences amongst regionally distinct adipose tissue depots 44 1.13 Adipose Tissue Dysfunction and Insulin Resistance ...... 45 1.14 Protein Kinase C ...... 46 1.15 Protein Kinase C epsilon ...... 49 1.16 Protein Kinase C epsilon, Lipid Metabolism and Insulin Resistance ...... 51 1.16.1 A potential role for PKCε in adipose tissue ...... 51 1.17 Summary ...... 53 1.18 Thesis Aims ...... 54

CHAPTER 2 MATERIALS AND METHODS ...... 55 2.1 Materials ...... 56 2.2 Cell Studies ...... 65 2.2.1 L6 cell culture, fatty-acid treatment and insulin stimulation ...... 65 2.2.1.1 Cell passaging ...... 65 2.2.1.2 Cell seeding, differentiation and general treatment procedure ...... 66 2.2.1.3 Generation and use of recombinant adenovirus ...... 67 2.2.1.4 Preparation of palmitate:BSA conjugate ...... 68 2.2.1.5 Preparation of insulin ...... 68 2.2.2 mRNA expression analysis ...... 68 2.2.3 Protein harvest and quantification ...... 69 2.2.4 Immunoblotting ...... 70 2.2.5 Sphingolipid flux assays ...... 70 2.2.6 GLUT4 translocation assay ...... 72 2.2.7 Sphingomyelinase inhibitor dose-response experiments ...... 73 2.3 Animal Studies ...... 74 2.3.1 Maintenance of mice ...... 74 2.3.2 Dietary treatment ...... 74 2.3.3 Generation of adipose tissue specific protein kinase c epsilon (PKCε) knockout (KO) and control mouse lines ...... 75

7 2.3.4 Genotyping ...... 76 2.3.5 Confirmation of PKCε deletion in adipocytes ...... 76 2.3.6 Intraperitoneal glucose tolerance test ...... 77 2.3.7 Tissue harvest and blood collection from 16 week fat-fed mice ...... 78 2.3.8 Adipocyte isolation ...... 78 2.3.9 Catheter insertion surgery for euglycaemic-hyperinsulinaemic clamp ...... 79 2.3.10 Euglycaemic-hyperinsulinaemic Clamp ...... 79 2.4 Metabolic and Morphological Studies ...... 81 2.4.1 Measurement of plasma insulin concentration ...... 81 2.4.2 Measurement of plasma non-esterified fatty acid concentration ...... 81 2.4.3 Measurement of plasma cytokine concentrations ...... 81 2.4.4 Measurement of adipocyte size ...... 82 2.4.5 2[14C]deoxyglucose uptake assay (Rg’ assay) ...... 82 2.5 In Vivo Expression Analysis of Gonadal Adipose Tissue ...... 84 2.5.1 mRNA expression analysis ...... 84 2.5.2 Protein expression analysis ...... 85 2.6 Statistical Analysis ...... 86

CHAPTER 3 MECHANISMS OF CERAMIDE SYNTHASE-INDUCED ALTERATIONS IN INSULIN ACTION ...... 87 3.1 Introduction ...... 88 3.2 Results ...... 90 3.2.1 CerS isoform overexpression has minimal effects on the mRNA levels of CerS isoforms and other proteins involved in sphingolipid metabolism ...... 90 3.2.2 Effects of CerS overexpression on ceramide synthesis pathway flux ...... 93 3.2.2.1 CerS isoform overexpression does not alter de novo pathway flux ...... 93 3.2.2.2 CerS1, 4 and 6 overexpression increases salvage pathway flux ...... 94 3.2.3 CerS1 overexpression increases GLUT4 transporter translocation to the plasma membrane ...... 96 3.3 Discussion ...... 98

CHAPTER 4 PHENOTYPIC CHARACTERISATION OF ADIPOSE TISSUE- SPECIFIC PROTEIN KINASE C EPSILON KNOCKOUT MICE ...... 105 4.1 Introduction ...... 106 4.2 Results ...... 108

8 4.2.1 PKCε is deleted in primary adipocytes only ...... 108 4.2.2 Minimal body-weight difference between WT and AdPKCεKO mice throughout the study ...... 109 4.2.3 Minimal differences in fasting metabolic parameters between genotypes 111 4.2.4 PKCε ablation in adipose tissue is associated with improved glucose tolerance and further suppression of plasma NEFA after a one-week HFD ...... 113 4.2.5 PKCε ablation in adipose tissue is linked with increased glucose tolerance and further suppression of plasma NEFA after 8 and 16-week HFD ...... 117 4.2.6 PKCε ablation in adipose tissue is associated with increased hepatic and whole-body insulin sensitivity in female mice following a one-week HFD ...... 122 4.2.7 PKCε ablation in adipose tissue is associated with smaller adipocyte size 127 4.3 Discussion ...... 129

CHAPTER 5 EFFECTS OF PROTEIN KINASE C EPSILON DELETION IN ADIPOSE TISSUE UPON LIPID METABOLISM AND CYTOKINE RELEASE 140 5.1 Introduction ...... 141 5.2 Results ...... 143 5.2.1 PKCε ablation in adipose tissue does not alter the expression of several genes involved in lipid metabolism, inflammation and adipocyte differentiation 143 5.2.2 PKCε ablation in adipose tissue is associated with increased protein expression of lipid esterification enzymes ...... 146 5.2.3 PKCε ablation in adipose tissue is associated with increased IL-6 and decreased TNFa plasma concentrations ...... 151 5.3 Discussion ...... 153

CHAPTER 6 SUMMARY AND FUTURE DIRECTIONS ...... 160

CHAPTER 7 APPENDICES ...... 168 Section A – Supplementary Data ...... 169 Section B – Buffer and Diet Compositions ...... 172

CHAPTER 8 REFERENCES ...... 175

9 LIST OF FIGURES AND TABLES

Figure 1-1 The canonical insulin signalling pathway and its actions on glucose metabolism and mitogenesis...... 18 Figure 1-2 The role of insulin in lipid homeostasis...... 19 Figure 1-3 Tissue-specific effects of insulin resistance...... 22 Figure 1-4 General ceramide structure...... 28 Figure 1-5 Pathways of ceramide synthesis...... 30 Figure 1-6 Sequence comparison of five CerS isoforms……………………………….32 Figure 1-7 A simplified view of lipolysis and fatty acyl-CoA esterification pathways. . 41 Figure 1-8 Main signalling pathways of lipolysis...... 42 Figure 1-9 The protein kinase C subfamilies...... 47 Figure 3-1: Effect of CerS overexpression on mRNA levels of endogenous CerS isoforms...... 91 Figure 3-2 Effect of CerS overexpression on mRNA levels of Sptlc and Ormdl isoforms 92 Figure 3-3 Radioactivity measurements of [3H]serine-labelled sphingomyelin...... 93 Figure 3-4 Representative autoradiograph of [3H]sphingosine incorporation into sphingolipids...... 94 Figure 3-5: Radioactivity measurements of [3H]sphingosine-labelled sphingolipids. .... 95 Figure 3-6: Surface and total GLUT4 transporter levels following CerS isoform overexpression...... 97 Figure 4-1 Tissue-specific protein expression of total PKCε in WT and AdPKCεKO mice...... 108 Figure 4-2 Weekly body weights of mice employed in the AdPKCεKO in vivo characterisation study...... 110 Figure 4-3 Blood glucose, plasma insulin and plasma NEFA excursions during ipGTT in male mice after 1 week high-fat or chow feeding...... 115 Figure 4-4 Blood glucose, plasma insulin and plasma NEFA excursions during ipGTT in female mice after 1-week high-fat feeding...... 116 Figure 4-5 Blood glucose, plasma NEFA and plasma insulin excursions during ipGTT in male mice after 8-week high-fat feeding...... 118 Figure 4-6 Blood glucose, plasma NEFA and plasma insulin excursions during ipGTT in female mice following 8-week high-fat feeding...... 119

10 Figure 4-7 Blood glucose, plasma NEFA and plasma insulin excursions during ipGTT in male mice after 16-week high-fat feeding...... 120 Figure 4-8 Blood glucose, plasma insulin and plasma NEFA excursions during ipGTT in female mice after 16-week high-fat feeding...... 121 Figure 4-9 Parameters of whole-body and tissue-specific insulin sensitivity measured in male mice before and during euglycaemic-hyperinsulinaemic clamp...... 125 Figure 4-10 Parameters of whole-body and tissue-specific insulin sensitivity measured in female mice before and during euglycaemic-hyperinsulinaemic clamp...... 126 Figure 4-11 Adipocytes from WT-Floxed and AdPKCεKO mice fed 16 week HFD. . 127 Figure 4-12 Adipocyte diameters of mice after 16 week HFD...... 128 Figure 5-1 mRNA levels in female murine adipose tissue...... 144 Figure 5-2 mRNA levels in male murine adipose tissue...... 145 Figure 5-3 Lipolysis protein levels in female murine adipose tissue...... 147 Figure 5-4 Lipid esterification protein levels in female murine adipose tissue...... 148 Figure 5-5 Lipolysis protein levels in male murine adipose tissue...... 149 Figure 5-6 Lipid esterification protein levels in male murine adipose tissue...... 150 Figure 5-7 Plasma cytokine concentrations in AdPKCεKO and WT-floxed mice...... 152 Figure A-1 Expression of endogenous and recombinant CerS isoforms in L6 myotubes..169 Figure A-2 The effect of acidic and neutral sphingomyelinase inhibitors on insulin signalling in L6 myotubes…………………………………………………………… 170 Figure A-3 Protein expression of PKCε in WT-Floxed and AdPKCεKO mice fed 16 week HFD. …………………………………………………………………………...171 Figure A-4 Gonadal adipose tissue weights from mice fed a 16-week HFD………...171

Table 1 Fatty acid substrate preferences and tissue-specific expression of each CerS isoform………………………………………………………………………………….33 Table 2 Materials used in this study……………………………………………………56 Table 3 Real-time PCR primers and Universal Probe Library probes used in this study ………………………………………………………………………………………….63 Table 4 Immunoblot primary antibodies used in this study……………………………64 Table 5 PCR primer sequences used for PKCεf and Adipoq-Cre genotyping…………76 Table 6 Fasting metabolic parameters for male mice before ipGTT…………………111 Table 7 Fasting metabolic parameters for female mice before ipGTT……………….112 Table 8 Parameters before and during euglycaemic-hyperinsulinaemic clamp………124 11 PUBLICATIONS In conjunction with this thesis Frangioudakis G, Diakanastasis B*, Liao BQ, Saville JT, Hoffman NJ, Mitchell TW, Schmitz-Peiffer C, 2013. Ceramide accumulation in L6 skeletal muscle cells due to increased activity of ceramide synthase isoforms has opposing effects on insulin action to those caused by palmitate treatment. Diabetologia 56(12): 2697-701. *Denotes co-first authorship

CONFERENCE PRESENTATIONS

Diakanastasis B, Brandon A, Suryana E, Hancock D, Schmitz-Peiffer C. A Role for Adipose Tissue Protein Kinase C Epsilon in Whole Body Glucose and Lipid Metabolism. Poster presentation: The Australian Society for Medical Research New South Wales Annual Scientific Meeting, Sydney, Australia, June 2014.

Diakanastasis B, Brandon A, Suryana E, Hancock D, Schmitz-Peiffer C. A Role for Adipose Tissue Protein Kinase C Epsilon in Whole Body Glucose and Lipid Metabolism.Poster presentation: The 22nd St. Vincent's Campus Research Symposium, Sydney, Australia, September 2014.

12 CHAPTER 1 GENERAL INTRODUCTION

13 1.1 Glucose Homeostasis, Lipid Homeostasis and Type 2 Diabetes Diabetes mellitus, or diabetes, is a metabolic disease defined by elevated fasting blood glucose concentrations ≥ 7.0 mmol/L [2]. This has emerged as a disease of epidemic proportions, which is projected to affect 592 million people worldwide by the year 2035 [3]. This staggering statistic highlights the increased need for more effective treatment strategies.

The two main forms of diabetes are Type 1 and Type 2 Diabetes (T2D). Type 1 Diabetes is an autoimmune disease leading to a loss of pancreatic beta cells and hence an inability to secrete sufficient levels of the hormone insulin, and is usually diagnosed early in life. T2D is defined as a failure of beta cells to secrete enough insulin to overcome a resistance to the hormone for the maintenance of normal blood glucose concentrations. This form of diabetes develops later in life, is most prevalent in obese and overweight individuals and is strongly associated with escalated morbidity and mortality relating to cardiovascular disease [4]. Current figures show that T2D accounts for up to 95% of total diabetes cases globally [3].

Impaired glucose and lipid homeostasis play a direct role in the pathogenesis of T2D. In order to completely appreciate this link, it is important to understand the fundamentals of normal glucose and lipid homeostasis. Glucose primarily acts as a source of fuel for energy production in the body. Glucose from the diet is released in the gastrointestinal tract via the degradation of complex dietary carbohydrates. This is followed by the direct absorption of glucose into the blood via facilitated entry across cell membranes by the glucose and sodium-glucose linked transporter proteins [5, 6]. Glucose is then either stored as glycogen in liver and skeletal muscle or converted to CO2 for the release of energy via aerobic or anaerobic glycolysis in skeletal muscle. Blood glucose may also originate from the breakdown of glycogen (glycogenolysis) in the liver and through its de novo synthesis in the liver and kidney (gluconeogenesis). Hyperglycaemia instigates insulin secretion from the beta cells of the pancreas, resulting in a net decrease in blood glucose concentration. Conversely, hypoglycaemia stimulates the secretion of glucagon from the alpha cells of the pancreas to increase blood glucose concentrations through the suppression of insulin secretion and the enhancement of

14 glycogenolysis. The control of glucose homeostasis hence, is highly dependent on a tightly maintained balance between the reciprocal actions of insulin and glucagon [7].

Lipids are required for a range of biological functions including energy production, cell signalling, cellular membrane formation and post-translational modifications [5]. Dietary lipids mainly occur as triacylglycerol (TAGs - three fatty acids covalently attached to a glycerol backbone) that are initially digested through partial hydrolysis in the stomach to form large fat globules, which are then further hydrolysed in the intestine to form monoacylglycerols (MAGs – one fatty acid covalently attached to a glycerol backbone) and fatty acids (FAs). These lipids are taken up into the intestinal epithelial cells, where they undergo esterification (covalent attachment of FAs) to form diacylglycerol (DAG – two fatty acids covalently attached to a glycerol backbone) and TAG, which are then coupled with apolipoprotein B to form chylomicrons and very low density lipoproteins (VLDLs) to enable DAG and TAG entry into the bloodstream via the lymphatic system [8, 9]. The chylomicrons and VLDLs interact with lipoprotein lipase (LPL) located on the inner capillary surface to release FAs through the hydrolysis of DAG and TAG. Liberated FAs either enter the cell (via the CD36 and FA transporter proteins, and to a lesser extent, membrane diffusion) or bind to albumin for re-release into the bloodstream. FAs that enter the cell are almost immediately bound to a co- enzyme A molecule (CoA) via acyl-CoA synthetase. This attachment enables FAs to remain within the cell for further metabolism [10]. In the fed state, FAs taken up by adipose tissue are esterified and stored as metabolically inert TAGs. The liver aids in this sequestration of lipids during the fed state by taking up FAs from the circulation, esterifying them into TAGs and packaging them into VLDLs which are then released into the circulation to ultimately supply lipid to adipose tissue. The fasted state causes a systemic reduction in chylomicron, VLDL and insulin concentrations and a systemic increase in catecholamine levels. This results in the hydrolysis-mediated breakdown of TAGs in adipose tissue to form FAs and glycerol that are then secreted into the circulation. Exposure to increased FA levels during fasting causes skeletal muscle and liver to switch from glucose to FA as the preferred energy substrate through altering the amounts of key intermediates in the tricarboxylic acid cycle [5, 9].

15 1.2 The Insulin Signalling Pathway and its Metabolic Actions Insulin is widely recognised as the main regulator of blood glucose and fatty acid concentrations on account of its multitude of associated actions across many tissues. Insulin maintains glucose and lipid homeostasis by i) increasing glucose uptake and storage in skeletal muscle and white adipose tissue ii) decreasing hepatic glucose production (glycogenolysis and gluconeogenesis) and output iii) increasing FA uptake and decreasing TAG breakdown (lipolysis) in white adipose tissue and iv) increasing FA production (de novo lipogenesis) in white adipose tissue and liver [11]. This further illustrates the complexity involved in the maintenance of healthy whole-body glucose and lipid homeostasis.

The canonical insulin signalling pathway is initiated when insulin binds to the alpha subunits of its heterotetrameric receptor, causing conformational changes that promote its beta subunit protein tyrosine kinase activity, initially causing autophosphorylation. The insulin signalling pathway then diverges into two main branches: the phosphatidylinositol 3-kinase/protein kinase B (P13K/Akt) pathway and the mitogen- activated protein kinase kinase/extracellular-signal-regulated kinase (MEK/ERK) pathway (Figure 1-1). In the P13K/Akt pathway, activated receptor tyrosine kinase phosphorylates insulin receptor substrate (IRS) proteins, which recruit and activate PI3K. This generates phosphatidylinositol 4,5-triphosphate (PIP3) from phosphatidylinositol 4,5-diphosphate (PIP2). PIP3 recruits phosphoinositide-dependant kinase-1 (PDK-1) and Akt to the cell membrane, positioning Akt for full activation through phosphorylation by both PDK-1 and mammalian target of rapamycin complexed with Rictor (mTORC2) [12]. Atypical protein kinase C zeta (aPKCζ) is also activated by PDK-1 and mTORC2 [13]. The activation of Akt and aPKCζ triggers a signalling cascade resulting in the translocation of the glucose transporter 4 (GLUT4) receptor from the cytosol to the cell membrane via exocytosis and subsequent uptake of glucose into the cell [14] (Figure 1-1). Activated Akt also facilitates the storage of glucose through the stimulation of glycogen synthesis via inhibitory phosphorylation of glycogen synthase kinase-3 beta (GSK-3β). This results in net dephosphorylation and activation of glycogen synthase [14] [12] (Figure 1-1). The PI3K/Akt pathway also promotes glycogen synthesis through directly inhibiting enzymes of glycogenolysis [15] (not shown) and inhibits gluconeogenesis by downregulating the transcription of

16 associated enzymes via cytosolic sequestration of the forkhead box protein O1 (FOXO1) transcription factor from the nucleus following phosphorylation by Akt [16] (Figure 1-1).

In the MEK/ERK pathway, activated insulin receptor tyrosine kinase phosphorylates the Shc adaptor protein (Shc), which then combines to the growth factor receptor-bound adaptor protein 2 (Grb2) to recruit the Son-of-sevenless (SOS) to the cell membrane. This activates the rat sarcoma (Ras) protein, triggering the sequential activation of rapidly accelerated fibrosarcoma (Raf) kinase, MEK1/2 and ERK1/2. Activated ERK1/2 then translocates to the nucleus to mediate a range of mitogenic effects including cell growth, proliferation and differentiation [11, 17] (Figure 1-1).

Figure 1-1 The canonical insulin signalling pathway and its actions on glucose metabolism and mitogenesis. Insulin binds to its receptor, activating its protein tyrosine kinases that recruit various proteins to form the P13K/Akt and MEK/ERK pathways, which mediate glucose metabolism and facilitate various mitogenic effects. For a detailed description of these pathways, refer to Section 1.2.

Insulin regulates lipid homeostasis through inhibiting lipolysis, increasing FA cellular uptake and increasing de novo lipogenesis. These actions are mainly facilitated via the PI3K/Akt pathway (Figure 1-2). Insulin stimulation of the PI3K/Akt pathway decreases lipolysis through (i) activating adipose-specific phospholipase A2 and phosphodiesterase enzymes which inhibit key lipolysis enzymes adenylate cyclase and protein kinase A (ii) decreasing the transcription of adipose triglyceride lipase, another important lipolysis enzyme and (iii) upregulating the transcription of fat-specific protein 27, an inhibitor of adipose triglyceride lipase. Insulin enhances FA uptake through upregulating the transcription, translation and activity of LPL via the PI3K/Akt pathway. FA uptake is also increased through PI3K/Akt and MEK/ERK pathway- mediated stimulation of FA transporter translocation to the plasma membrane [18].

18 Increased de novo lipogenesis elevates the amount of readily-available FAs for esterification. Insulin-stimulated protein phosphatase 1 (PP1) and the PI3K/Akt pathway facilitate the upregulation of de novo lipogenesis through elevating the transcription, post-transcriptional processing and activity of sterol regulatory element binding protein 1 (SREBP1), a transcription factor that stimulates the expression of many lipogenic genes. Insulin-stimulated glucose uptake, via the PI3K/Akt pathway, also stimulates the expression of lipogenic genes through the activation of carbohydrate response element binding protein (ChREBP). This is one example of the extensive interconnectivity between glucose and lipid metabolism [19-21].

Figure 1-2 The role of insulin in lipid homeostasis. Insulin increases de novo lipogenesis through its actions via the PI3K/Akt pathway and on PP1. FA-uptake is enhanced through the insulin-mediated initiation of the PI3K/Akt and MEK/ERK pathways. Lipolysis is decreased via the PI3K/Akt pathway.

19 1.3 Obesity, Diabetes and the Metabolic Syndrome Obesity-related disruptions to whole body glucose and lipid homeostasis can be caused by chronic exposure of major associated organs to elevated lipid concentrations. This is closely linked to the pathogenesis of T2D and forms the basis of the metabolic syndrome.

The generation of T2D is preceded by the development of insulin resistance, which is defined as the diminished response of insulin sensitive tissues to insulin [4]. During insulin resistance, the beta cells of the pancreas secrete excessive amounts of insulin to maintain healthy blood glucose concentrations. The pancreatic beta cells in genetically susceptible individuals eventually fail to keep up with the elevated insulin demand, resulting in the chronic hyperglycaemia and hypertriglyceridaemia that is characteristic of T2D [22]. Metabolic syndrome is defined as a cluster of metabolic abnormalities that culminate to accelerate the development of T2D and/or cardiovascular disease. Insulin resistance is a key element of the metabolic syndrome, which is also comprised of obesity, high blood pressure, hyperglycaemia and hypertriglyceridaemia [4].

While a strong association between lipid oversupply and insulin resistance has been long acknowledged [23], there remains an incomplete understanding of the mechanisms involved. Elucidating some of these mechanisms is one of the main objectives of this thesis.

20 1.4 Insulin Resistance Insulin resistance negatively affects the overall capability of an organ to metabolise lipid as well as glucose. This can alter the net release of lipid from a particular organ, which can induce and/or further propagate insulin resistance in other tissues. Based on this, it is widely accepted that the progression of insulin resistance to T2D involves cross talk-between insulin resistant tissues, rather than isolated contributions from each [24].

Studies in rodents have shown that insulin-sensitive tissues develop insulin resistance during different stages of high fat feeding. Furthermore, it has been recently demonstrated in mice that lipid-induced insulin resistance in a particular tissue peaks soon after its development and is not further intensified with longer-term high fat feeding. Adipose tissue and liver develop insulin resistance and glucose intolerance within one week of a high fat diet (HFD), whilst insulin resistance in skeletal muscle is not detected until three weeks of HFD [25, 26]; similar temporal trends have been observed in humans [27, 28]. These findings coincide with the idea that adipose tissue and liver are organs that initiate the generation of whole-body insulin resistance, whilst skeletal muscle develops insulin resistance during the later stages of high-fat feeding in response [5]. A major component of this thesis will investigate the role of adipose tissue in the development of the whole body insulin resistant state.

Insulin resistance severely dysregulates the normal physiological processes induced by insulin. Consequently, skeletal muscle exhibits lower glucose uptake and storage, adipose tissue displays higher FA release (resulting from increased lipolysis) and decreased FA/glucose uptake and the liver exhibits elevated glucose production (glycogenolysis and gluconeogenesis) and output (Figure 1-3). The combined effect of these factors is hyperglycaemia and hypertriglyceridaemia, which contribute toward the development of T2D in genetically susceptible individuals if not prevented by compensatory insulin secretion [29, 30] [31].

Glucose production and output

Figure 1-3 Tissue-specific effects of insulin resistance. Insulin resistant skeletal muscle presents decreased glucose uptake and storage. Adipose tissue insulin resistance causes higher amounts of FA release and decreased FA and glucose uptake. Insulin-resistant liver exhibits elevated glucose production and output.

The link between chronic lipid accumulation and insulin resistance was initially postulated by Randle and colleagues in 1963 to arise through competition between glucose and FA as the preferred substrate for oxidative energy production within muscle and adipose tissue [32]. The proposed mechanism involved elevated lipid levels that caused increased cellular uptake and oxidation of lipids resulting in an accumulation of metabolites that impaired insulin-stimulated glucose oxidation. This impairment was proposed to produce increased concentrations of glycolytic metabolites that inhibited glucose hexokinase, which at the time was considered to facilitate glucose uptake [32]. The Randle hypothesis is not however, supported by later studies showing that insulin- stimulated glycogen synthesis (which does not involve glucose oxidation) was also impaired [33] and that the predicted build up of glucose-6-phosphate in muscle does not occur [34]. It was concluded that reduced insulin-stimulated GLUT4 translocation to the cell membrane was the primary mechanism responsible for reduced glucose uptake in adipose tissue and skeletal muscle [35, 36].

22 It is generally acknowledged that mechanisms which lead to the generation and propagation of lipid-induced insulin resistance include the direct inhibition of insulin signalling, chronic low-grade inflammation, dysregulated adipose tissue protein (adipokine) secretion, mitochondrial stress and endoplasmic reticulum (ER) stress [37- 41]. The relative contribution of each pathway to the development and exacerbation of insulin resistance at the local and whole-body level is an area of research currently under extensive investigation.

1.5 The Role of Lipids in the Pathophysiology of Insulin Resistance The strong association between increased skeletal muscle TAG content and insulin resistance had led to the idea that TAG was the lipid moiety responsible for the generation of insulin resistance in insulin-sensitive organs [25, 42]. This concept was first overturned by studies demonstrating that skeletal muscle insulin resistance induced in rats via lipid infusion was associated with increased levels of DAG, but independent of changes in skeletal muscle TAG concentrations [34, 43]. The literature has also shown that increased levels of ceramide, another lipid intermediate, are present in insulin resistant skeletal muscle and adipose tissue [26, 44-46]. Hence, it is currently accepted that insulin resistance is induced by active lipid intermediates such as DAG and ceramide, rather than by TAG per se. This reconciles the Athlete’s Paradox, which highlights the fact that that endurance athletes have increased skeletal muscle insulin sensitivity despite high intramuscular TAG concentrations [47].

TAG is considered metabolically inert on account of its sequestration in the lipid droplet, and is not known to directly interfere with the pathways leading to insulin resistance. However, an increase in intracellular TAG concentrations may possibly lead to insulin resistance through providing increased amounts of substrate required to generate elevated levels of pathological lipid intermediates via lipolysis [40]. Hence, blood TAG concentrations may still serve as a useful marker of insulin resistance despite not being directly involved its development [48]. Bioactive lipid intermediates may also accumulate in insulin target tissues as a consequence of increased FA concentrations. This may occur through increased FA uptake in non-adipose tissues, increased lipolysis, decreased FA esterification and reduced FA oxidation. The relative

23 contribution of each pathway to FA accumulation differs between insulin-sensitive tissues [49].

The inhibitory effects of lipids on insulin action can occur in more than one manner. Chronic lipotoxicity is mediated through obesity-associated FA oversupply exceeding the capacity of adipose tissue to take up lipid. This causes chronic ectopic fat deposition into liver and skeletal muscle, thought to result in the generation of insulin resistance [50], most likely via the generation of inhibitory intermediates [51]. Acute exposure to FA also contributes to a reduction in insulin sensitivity of target organs. Multiple studies suggest that this occurs through transient increases in FA flux from adipose tissue to liver and skeletal muscle [52-54] [55]. Both forms of lipid-induced insulin resistance are explored in this thesis.

1.6 Lipid-Induced Skeletal Muscle Insulin Resistance Skeletal muscle accounts for up to 75% of insulin-dependent glucose uptake [56]. The appearance of insulin resistance in this tissue therefore, has a profoundly negative impact on glucose homeostasis. The most well-documented lipid mediators of skeletal muscle insulin resistance are DAG and ceramide (composed of a FA covalently attached to a sphingoid backbone - see figure 1.4).

Elevated DAG concentrations in skeletal muscle are associated with acute lipid-induced insulin resistance [57] and insulin resistance in obese rodents [58]. DAG is believed to generate skeletal muscle insulin resistance in part by activating the protein kinase c- theta (PKCθ) isoform, which may impair insulin signalling though inhibitory phosphorylation of the IRS1 protein [59, 60]. However, many studies have demonstrated a disconnect between skeletal muscle DAG concentrations and insulin resistance [61-63]. DAG species differ according to the FAs that are attached to the glycerol backbone. Based on this, it is possible that this conflict in the literature may arise from some DAG species being more involved in insulin resistance than others. Further investigation to elucidate the individual roles of particular DAG moieties however, is beyond the scope of this study.

24 Several studies report that elevated ceramide levels diminish insulin action in skeletal muscle cell models, as well as in human and rodent muscle [44-46, 64-67]. Correspondingly, there exists a positive association between skeletal muscle ceramide concentration and the severity of insulin resistance [68-70]. Furthermore, pharmacological inhibition of ceramide synthesis has been shown on multiple occasions to prevent skeletal muscle insulin resistance in cell models and rodents [45, 71-73]. Despite this mounting support for the role of ceramide as an instigator of skeletal muscle insulin resistance, Skovbro and colleagues have demonstrated a positive association between skeletal muscle ceramide concentration and insulin sensitivity in middle-aged, human males matched for lean body mass [74]. A wide range of insulin sensitivities were encompassed in this study as subjects were either Type 2 Diabetic, had impaired glucose tolerance, healthy but untrained, or healthy and endurance trained. Insulin sensitivity was determined via the euglycaemic-hyperinsulinaemic clamp, the gold standard in ascertaining this measurement [75, 76].

Overall, these conflicting studies suggest that the role of ceramide in the pathogenesis of skeletal muscle insulin resistance may not be as simple as suggested by the majority of associated literature. Ceramides, like DAGs, appear as a range of species that differ on the basis of the FA moiety attached to the sphingoid backbone. Based on this, it is plausible that these individual species may protect against, as well as promote the generation and propagation of skeletal muscle insulin resistance [77]. This idea has been explored in recent work by our laboratory and will be further investigated in this thesis.

1.7 Mechanisms of Ceramide-Induced Insulin Resistance Ceramide induces insulin resistance through many routes. Ceramide per se is most recognised to diminish insulin sensitivity through its direct actions on the insulin signalling pathway. However, it has been shown that ceramide is associated to varying extents with mitochondrial dysfunction, inflammation, apoptosis and lipid raft alterations. These links have been known to contribute to the aetiology of insulin resistance and are discussed below.

1.7.1 Inhibition of insulin signalling Ceramides in skeletal muscle inhibit insulin signalling at least in part through the deactivation of Akt, a central signal transduction node in the insulin signalling cascade. 25 This is achieved though two separate mechanisms. The first mechanism involves the dephosphorylation of Akt via the ceramide-induced activation of protein phosphatase 2A [78]. Alternatively, ceramide may induce aPKCζ to elicit phosphorylation of Akt at certain residues, preventing its translocation to the plasma membrane and subsequent activation [79, 80]. It has been demonstrated that the predominant inhibitory mechanism can differ across different cell types (as well as between different skeletal muscle cell lines), depending on variations in plasma membrane composition [81]. Ceramide-based inactivation of IRS1 and PI3K has also been documented, but findings are controversial [82].

1.7.2 Inflammation The innate immune response can be initiated via activation of toll-like receptors (TLRs), which activate the nuclear factor kappa B (NF-κB) and jun amino-terminal kinase (JNK) signalling pathways to induce the transcription of inflammatory cytokines, which are linked to insulin resistance [83]. Palmitate, a saturated FA, is known to act as an agonist for the TLR4 receptor and has been shown to induce insulin resistance through this activation [84, 85]. Furthermore, a study by Holland and colleagues in 2011 demonstrated that palmitate-induced TLR4 activation increased the transcription of various ceramide synthesis enzymes (including most ceramide synthase (CerS) isoforms) and that the resulting increase in ceramide was essential for TLR4-dependant insulin resistance [86]. In addition, palmitate-induced TLR4-dependant increases in the inflammatory cytokine tumour necrosis factor alpha (TNFα) has been proposed to increase ceramide synthesis through sphingomyelinase (SMase) activation [87, 88].

1.7.3 Mitochondrial dysfunction Mitochondria have been shown to contain a variety of sphingolipids including ceramide [89, 90]. CerS isoforms have been associated with the mitochondria as previously mentioned. Abnormalities in mitochondrial oxidative function have been proposed to cause lipid accumulation, leading to enhanced ceramide production and subsequent insulin resistance [37, 91]. Alternatively it has been shown that ceramide may induce mitochondrial dysfunction through disruptions to the electron transport chain [92], resulting in an increase in reactive oxygen species [93], which has been linked to skeletal muscle insulin resistance [40].

26 1.7.4 Apoptosis Ceramide synthesis can be induced in response to a variety of apoptotic stimuli including UV/ionizing radiation, apoptotic receptor activation and chemotherapeutic agents [94]. Furthermore, this sphingolipid has been reported to display pro-apoptotic effects [95-97]. It is proposed that ceramide mediates apoptosis by increasing mitochondrial membrane permeability through recruitment of the pro-apoptotic protein bcl-2-like protein 4, resulting in the release of cytochrome c which then activates the central facilitator of apoptosis, caspase-3 [98]. Promising evidence of a connection between ceramide-induced apoptosis and insulin resistance is presented in studies which have shown an association between palmitate-induced apoptosis and skeletal muscle insulin resistance [70, 99, 100]. Conversely, a study by Senkal and colleagues in 2010 found that distinct ceramide species and CerS isoforms demonstrated anti-apoptotic effects [101]. Another study also supports an anti-apoptotic role for distinct ceramide species and CerS isoforms. However, it is unclear whether this effect is associated with increased insulin sensitivity as this investigation was conducted in human head and neck carcinoma samples [102].

1.7.5 Lipid raft alterations Lipid rafts are microdomains comprised of clustered molecules of sphingolipids (including ceramide) and cholesterol bound to integral membrane proteins known as caveolins. Lipid rafts are distributed across the plasma membrane and function as scaffolding for the assembly of signalling complexes, including the insulin signalling pathway [103, 104]. It is therefore presumed that alterations in the ceramide content of lipid rafts could modulate initiation of the insulin signalling pathway. In fact, CerS2 has been shown to affect lipid raft composition and insulin receptor trafficking in liver, as deletion of this enzyme and resulting changes in long chain ceramide content altered hepatic insulin sensitivity [105]. More investigation is required with respect to the biological effects of ceramide-induced lipid raft alterations. This however, is beyond the scope of the current study.

27 1.8 Sphingolipid Metabolism and Ceramide Synthase The sphingolipid class of lipids comprises of a diverse range of related lipid moieties including ceramides, sphingoid bases (the canonical “backbone” of sphingolipids), sphingomyelins, glucosylceramides and glycosphingolipids. Sphingolipid metabolism is comprised of multiple pathways which converge at ceramide, making this sphingolipid the central pivot for sphingolipid metabolism [106]. Though ceramide is known to be widely implicated in skeletal muscle insulin resistance, it is also involved in a number of important physiological functions including cell cycle arrest, cell proliferation, apoptosis and most notably, membrane formation [82]. The general structure of ceramide is comprised of a sphingoid backbone acylated on its free primary amine with a fatty acyl CoA group. Ceramide species differ from one another as based on the carbon chain length and saturation of the FA attached to the sphingoid backbone (Figure 1-4).

Figure 1-4 General ceramide structure. The generic structure of ceramide consists of a sphingoid backbone acylated at its amine group to a FA of variable chain length and saturation. Figure adapted from [107].

1.8.1 Ceramide metabolism There are three known routes of ceramide generation (i) the de novo pathway (ii) the salvage (recycling) pathway and (iii) sphingomyelin (SM) hydrolysis. The de novo pathway is initiated in the ER. It begins with the condensation of serine and palmitate by serine palmitoyltransferase (SPT) to form 3-ketosphinganine. SPT can be inhibited by ORM proteins (refer to Section 1.8.3 for more details). Palmitate is selectively metabolised by SPT and therefore is the predominant substrate utilised for the initation

28 of de novo ceramide synthesis [108]. This is followed by the reduction of 3- ketosphinganine to form the sphingoid backbone (sphinganine), which is then acylated with a FA by CerS to create dihydroceramide. Desaturation of dihydroceramide forms ceramide [109] (Figure 1-5).

The salvage pathway occurs within the endo-lysosomal compartment and initially involves the uptake of glycosphingolipids and SM from the plasma membrane. These sphingolipids are then respectively degraded by acid β−glucosidase and acid SMase to form ceramide, which is further broken down to the sphingoid backbone (sphingosine) via acid ceramidase. Sphingosine leaves the lysosome and is re-converted into ceramide at other intracellular locations via the action of CerS. Ceramide produced via the de novo and salvage pathways is shuttled via facilitated transport to the Golgi for the production of the more complex sphingolipids, SM and glucosylceramide. The ceramide transport (CERT) protein delivers ceramide designated for SM synthesis, whilst vesicular trafficking delivers ceramide designated for glucosylceramides synthesis. These complex sphingolipids are transferred to the plasma membrane via vesicular trafficking (Figure 1-5).

SM hydrolysis predominantly occurs at the plasma membrane and involves the degradation of SM into ceramide through the action of neutral SMase. Due to their hydrophobicity, ceramides produced here tend to reside in the plasma membrane and can be reconverted back to SM at this site [94, 109] (Figure 1-5).

During normal physiological conditions, ceramide synthesis pathways are utilised to different extents within distinct cell types [110, 111]. Under conditions of palmitate excess however, the de novo pathway appears to predominate on account of palmitate oversupply exceeding the capacity of the mitochondria to oxidise palmitate, resulting in the shuttling of this FA towards de novo ceramide synthesis [112].

Figure 1-5 Pathways of ceramide synthesis. Ceramide synthesis can occur via three routes: the de novo pathway, the salvage (recycling) pathway and sphingomyelin hydrolysis. Refer to Section 1.8.1 for a more detailed description of these pathways. CDase – ceramidase, CerS – ceramide synthase, CERT – ceramide transfer protein, ER – endoplasmic reticulum, GCS – glucosylceramide synthase, GluCer – glucosylceramide, GSL – glycosphingolipid, PM – plasma membrane, Sph – sphingosine, SM – sphingomyelinase, SMS – sphingomyelin synthase, SPT – serine palmitoyltransferase. ORM is the actual name of the protein, not its abbreviated form. Figure adapted from [109].

30 1.8.2 Ceramide synthases The importance of CerS enzymes in sphingolipid metabolism is reflected by the fact that they are central components of both the de novo and salvage pathways of ceramide synthesis. Six CerS isoforms have been identified in mammals (CerS1-6). The molecular mass of each CerS isoform is similar, ranging between 40-46 kDa. All CerS isoforms are predicted to have six transmembrane helices except CerS6, which has been proven to possess five. Common to all CerS isoforms is the TRAM-LAG1-CLN8 domain, a stretch of approx. 200 amino acids that contains five of these predicted transmembrane helices. All CerS isoforms, except CerS1, also possess a homeobox (HOX) domain. The function of this domain however, is currently unknown. The N- terminus of CerS is predicted to reside within the lumen of the ER whilst the C- terminus is predicted to reside in the cytosolic leaflet of this organelle [1, 113]. Currently, it is unclear which domains are involved in the regulation of CerS activity. However, common to all CerS isoforms is a highly conserved 52kDa amino acid sequence known as the longevity-assurance gene 1 (Lag1p) motif located within the TRAM-LAG1-CLN8 domain [114, 115] (Figure 1-6). As such, this sequence is presumed to encode for a functional domain. This was confirmed for CerS1 and CerS5 in a study which showed that site-directed mutagenesis of Lag1p amino acids decreased the activity of these isoforms [116]. Despite high sequence homology of the Lag1p functional domain between CerS isoforms, each isoform displays an individual preference toward a subset of FAs with particular carbon chain lengths and degrees of saturation. Furthermore, each CerS isoform presents its own distinctive pattern of tissue distribution [117, 118]. The tissue-specific expression and preferred FA substrates for each CerS isoform are listed in Table 1. These differences determine the collection and abundancies of individual ceramide species within a specific tissue. The exclusive expression of CerS3 to tissues that are not implicated in insulin resistance excludes this isoform from our current study and as such, this isoform will not be further mentioned.

[1]

Figure adapted from adaptedfrom Figure Sequence comparison of five CerSisoforms. fiveSequence of comparison

6 - 1

2 1 4 5 4 5 1 5 4 4 2 1 5 2 2 1 4 5 2 1

Figure Shown is an amino acid sequence Homologous regions encoding transmembrane domains the (TM), homeobox domain (HOX) and Lag1 motif alignment between murine CerS1, CerS2, are CerS4, specified. CerS5 and CerS6. CerS CerS CerS CerS CerS6 CerS CerS CerS CerS6 CerS CerS CerS6 CerS CerS CerS CerS CerS CerS CerS6 CerS CerS CerS CerS6 CerS CerS

32 Table 1 FA substrate preferences and tissue-specific expression of each CerS isoform Data obtained from [117, 118]. Cn corresponds to number of carbons on the fatty acid molecule attached to the sphingoid backbone of ceramide.

1.8.3 Regulation of sphingolipid metabolism enzymes There is limited information within the literature concerning the regulation of enzymes involved in sphingolipid metabolism. Current knowledge with regard to this is discussed below.

SPT, an enzyme involved in the de novo ceramide synthesis pathway, is known to be negatively regulated by ORM proteins. Yeast studies indicate that ORM proteins achieve this by forming an inhibitory complex with SPT [119]. An in vitro study has also demonstrated that phosphorylation of CERT impairs its ability to deliver ceramides from the ER to the Golgi [120].

It remains uncertain whether CerS is predominantly regulated by transcription or translation. A study by Mullen and colleagues in 2011 however, clearly demonstrates the existence of counter-regulatory actions of various CerS isoforms following siRNA- mediated knockdown of specific isoforms [121]. This has also been reported in other studies [122, 123]. It has been shown that heterodimerization of CerS isoforms may enhance the activity of individual isoforms [124]. Various CerS isoforms are predicted to be phosphorylated [125]. Though the consequences of this post-translational modification remain unclear, it has been reported that phosphorylation of CerS1 by p38 mitogen-activated protein kinase and PKC increases ubiquitination and proteasomal 33 turnover of CerS1, respectively [126, 127]. Glycosylation of various CerS isoforms have been reported. Though this was a key piece of evidence to suggest that the N- terminus of CerS isoforms faces the luminal side of the ER, the functional purpose of this glycosylation is unclear [1].

The regulation of sphingolipid metabolism in skeletal muscle cells during conditions of lipid oversupply is currently not known, yet requires investigation. An improved understanding of sphingolipid metabolism over the past two decades has formed a basis for examining this particular regulation, which will be conducted in this thesis.

1.9 The Concept of “Many Ceramides” The high degree of variation stemming from the FA component of ceramide has resulted in the identification of at least 200 structurally distinct ceramide species. These species were previously conceptualised to operate as a single functional entity. The development of lipidomics analysis techniques in recent years however, has caused this paradigm to shift towards the “many ceramides” model, which postulates that individual ceramide species contain distinct functions that are dependent on both their structure and subcellular localisation [77]. Studies in agreement with this model demonstrate that ceramide entities with different fatty acyl chain lengths possess different physiological and pathophysiological functions [92, 101, 102, 128, 129] [130]. Independent roles for each CerS isoform during certain disease states are suggested by the fact that relative expression levels of CerS isoforms dictate the ceramide fingerprint for each tissue. Early evidence of this concept is provided through a study in fat fed mice, which linked alterations in CerS1 expression to changes in specific ceramide levels and glucose tolerance [45]. This concept was reinforced by a more recent study by Park and colleagues in 2013, which showed that global CerS2 knockout mice displayed hepatic insulin resistance alongside decreases in ceramides produced specifically via CerS2 [105]. Furthermore, this study specifically raises the idea that distinct ceramide species may elicit protective effects against the pathogenesis of insulin resistance.

34 1.10 The Protective Role of Ceramide in the Pathogenesis of Insulin Resistance Though there is early evidence to suggest that each CerS isoform manifests distinct biological actions, the contributions of each CerS isoform toward the development of insulin resistance in key organs remains unclear. This crucial knowledge gap motivated our laboratory to elucidate the role of individual CerS isoforms and ceramides during the inhibition of skeletal muscle insulin action by lipid treatment. We used palmitate- treated L6 rat skeletal myotubes as a cell model of skeletal muscle insulin resistance.

As expected, our data showed that each overexpressed CerS isoform altered L6 myotube ceramide profile in a highly specific manner. Surprisingly, no CerS isoforms inhibited insulin signalling or glycogen synthesis in the absence of palmitate and no CerS isoform contributed to the inhibitory effects of palmitate on insulin action. Furthermore, improvements in insulin signalling and glycogen synthesis were found to be associated with CerS1 and CerS6 overexpression in the absence of palmitate. Conversely, CerS6 siRNA knockdown impaired these physiological events [131]. The beneficial effects observed in the absence of palmitate suggest that ceramides generated from overexpressed CerS isoforms are imposing effects on insulin action independent from ceramides generated from palmitate treatment alone. Our findings indicated that specific CerS isoforms in skeletal muscle, particularly CerS6, may positively regulate glucose metabolism either by increasing potentially beneficial ceramide pools, causing compensatory decreases in potentially inhibitory ceramide pools, or both. We proposed that inhibitory ceramides could be produced via enhanced SM hydrolysis attained by palmitate-induced activation of SMase. This activation was proposed to occur through palmitate activation of the TLR4 inflammatory pathway [88, 132]. Alternatively, overexpression or knockdown of specific CerS isoforms may have lead to counter- regulation of sphingolipid metabolism making our results difficult to interpret. It remains unclear how CerS overexpression did not potentiate the effects of palmitate as expected.

A protective role for certain ceramide species and CerS isoforms have been previously reported by the aforementioned study by Park and colleagues, which showed that CerS2 and its consequent ceramides were associated with elevated hepatic insulin sensitivity [105]. In contrast, findings from two recent independent studies released after

35 publication of our results have specifically implicated CerS6 and C16 ceramide within liver, white adipose tissue and brown adipose in the pathogenesis of insulin resistance [92, 130]. These studies further highlight the need to examine CerS counter-regulation in the L6 muscle cell model.

1.11 Lipid-Induced Hepatic Insulin Resistance Hepatic insulin resistance has immensely adverse effects on whole-body glucose homeostasis. This is predominantly mediated by both acute and chronic elevations in hepatic lipid content [41]. Ceramides and CerS isoforms are implicated in the aetiology of hepatic insulin resistance. This has been demonstrated by Raichur et al., 2014 and Turpin et al., 2014 as mentioned earlier. It has been also shown that activation of hepatic adiponectin receptors lowers hepatic ceramide content. This was found to cause an improvement in whole-body insulin sensitivity [133]. The specific role of ceramide in the pathogenesis of hepatic insulin resistance however, is beyond the scope of this study. Instead, a substantial proportion of investigations in this thesis are concerned with how acute increases in hepatic FA supply could affect hepatic insulin action and whole-body glucose homeostasis.

Many studies in dogs and humans in the 1990s suggested that it was the indirect actions of insulin on peripheral tissues, rather than the direct effect of insulin on the liver, which reduced hepatic glucose production (HGP) [52-54]. These studies implemented systemic insulin infusions during which plasma FA concentrations were either allowed to decrease or were prevented from doing so via simultaneous systemic lipid infusion - a model of acute FA oversupply. Subjects administered with this lipid infusion displayed increased HGP in comparison with non-lipid infused subjects. Results from these studies therefore indicated that the anti-lipolytic effect of insulin on adipose tissue (the sole site of systemic FA release) was at least partially responsible for the regulation of HGP. It was proposed that decreased hepatic FA metabolism reduced HGP independently from insulin action through associated reductions in FA-derived energy substrates and co-factors (NADH, ATP and acetyl CoA) required for gluconeogenesis [52-54].

36 However, a seminal publication by Fisher and Kahn in 2003 demonstrated that direct insulin signalling at the liver was also required to regulate HGP. In this study, liver- insulin receptor knockout mice subjected to chronic systemic insulin infusion did not display a reduction in HGP despite the ability of wild-type (WT) controls to do so [134]. These novel findings caused a shift in focus towards determining how modulations in hepatic insulin signalling caused increases in HGP during insulin resistance.

Many subsequent studies converged toward the identification of novel PKC isoforms as mediators of hepatic insulin resistance [135-140]. Activation of these enzymes is mediated by DAG, placing these enzymes as ideal candidates for eliciting the pathological consequences of dietary lipid oversupply. As such, chronic activation of novel PKC isoforms has been reported in response to an excess of dietary lipid. The generally accepted dogma of novel PKC-induced insulin resistance stipulates that the chronic DAG-induced activation of novel PKC isoforms instigates their translocation from the cytosol to the plasma membrane. From this location, novel PKC enzymes are able to blunt insulin signalling through conducting direct or indirect (via JNK or inhibitor of nuclear factor kappa-B kinase subunit beta (ΙΚΚβ) activation) inhibitory phosphorylation of insulin receptor substrate proteins and/or physical interactions with the insulin receptor to diminish its ability to phosphorylate its substrates [141]. This inhibition of canonical insulin signalling has been linked to various physiological perturbations including decreased skeletal muscle glucose uptake and increased hepatic glucose production [41, 142]. Studies have shown that protein kinase c epsilon (PKCε) is the novel PKC isoform most commonly implicated in the generation of hepatic insulin resistance. Furthermore, the majority of this literature suggests that PKCε achieves this through direct actions in the liver [135, 136, 140]. One such publication was a study from our group [140] which demonstrated that global PKCε knockout (KO) mice displayed protection against glucose intolerance induced after a 1 week HFD, which is long enough to elicit hepatic but not skeletal muscle insulin resistance [25, 26].

To examine whether PKCε achieved this effect through direct action at the liver, our laboratory generated liver-specific PKCεKO mice and assessed them for glucose tolerance after one week of HFD. Contrary to expectation, these mice did not display an improvement in glucose tolerance (manuscript in preparation). Furthermore, a separate 37 cohort of liver-specific PKCεKO mice fed HFD for one week failed to display an improvement in whole-body insulin sensitivity during the euglycaemic- hyperinsulinaemic clamp, the gold standard technique for determining insulin sensitivity (manuscript in preparation). Though hepatic insulin resistance and reduced insulin signaling appear alongside DAG accumulation and PKCε translocation, the mechanistic links between these processes are weak [137, 140, 143, 144]. The absence of a correlation between increased hepatic DAG levels and insulin resistance has also been reported [145]. These studies, in conjunction with our own findings, suggest that PKCε may be indirectly decreasing hepatic insulin sensitivity via actions from an alternative location to the liver.

Our laboratory observed that global PKCεKO mice displayed lower plasma FA concentrations than WT controls during a glucose tolerance test (manuscript in preparation). This suggests that global PKCεKO mice may be further suppressing the acute release of FA from adipose tissue, the only organ capable of releasing FAs into the circulation [146]. Furthermore, parallel knockdown of PKCε expression in adipose tissue and liver was linked to improved insulin sensitivity in both tissues, in addition to increased suppression of FA release from adipose tissue [137]. All these findings have prompted our laboratory to revise the current dogma of lipid-induced hepatic insulin resistance by further investigating the role of adipose tissue in the regulation of HGP, with a specific focus on elucidating the potential contribution made by PKCε in adipose tissue.

38 1.12 The Functional Roles of Adipose Tissue There are two types of adipose tissue: white adipose tissue and brown adipose tissue. White adipose tissue, referred to herein as adipose tissue, constitutes for 20-30% of total body weight in the normal, healthy state and is the primary energy storage site in the body [9]. Brown adipose tissue conversely, expends energy through its thermogenic properties and is mostly present in small amounts within the neck region [147]. Research into brown adipose tissue is a thriving field but will not be discussed further.

Adipose tissue is a heterogeneous tissue composed of adipocytes, preadipocytes, fibroblasts, stromal vascular cells and immune cells such as macrophages [148]. The adipocyte is the parenchymal cell type of adipose tissue and is comprised of a large lipid droplet surrounded by a thin aqueous rim containing the cytoplasm and essential subcellular compartments [149]. Historically, adipose tissue was considered solely as a metabolically inert store of excess energy. Over the past 20 years, this interpretation has shifted toward the recognition of adipose tissue as a dynamic organ that plays a critical role in the regulation of energy balance through its capacity to buffer FA fluxes in response to altering nutritional states and to secrete adipokines [150, 151]. The tightly regulated buffering capacity of adipose tissue has two main functions (i) to prevent the excess release of FA into the circulation and to subsequently reduce exposure of tissues to elevated FA concentrations when nutrient intake is high and (ii) to facilitate the release of FA to peripheral organs when energy demand surpasses nutritional intake [150]. Insulin, in part, mediates this buffering capacity in a variety of ways as described in Section 1.2. Dysregulation of this buffering capacity is central to the development of insulin resistance in peripheral organs. Therefore, it is important that an understanding of adipose tissue function is gained.

39 1.12.1 Lipid metabolism in adipose tissue The processes of lipid metabolism that occur in adipocytes to maintain lipid homeostasis include FA uptake, FA esterification, lipolysis and, to a lesser extent, de novo lipogenesis. The process of FA uptake is summarized in Section 1.1. The remaining processes are described below.

1.12.1.1 Fatty acid esterification FA esterification is the process of FA storage through the synthesis of TAG. The main route of FA esterification is the Kennedy pathway [152], which occurs mainly at the ER and facilitates the net storage of three FAs. This process begins with the addition of a FA-CoA molecule to glycerol-3-phosphate to form lysophosphatidic acid. This step is facilitated by glycerol-3-phosphate acyltransferase (GPAT) in the ER and mitochondrial glycerol-3-phosphate acyltransferase (GPAM). Gene expression of GPAT is constitutive whilst GPAM gene expression is induced by hormonal and nutritional signals [153]. Another FA-CoA is added to lysophosphatidic acid by 1-acylglycerol-3- phosphate acyltransferase (AGPAT) to generate phosphatidic acid, which is dephosphorylated by phosphatidic acid phosphatase (PAP or LIPIN-1) to form DAG. The final step of this pathway involves the addition of a FA-CoA to DAG by diacylglycerol acyltransferase-1 and 2 (DGAT-1, DGAT-2) to form TAG [154] (Figure 1-7). Accumulation of TAG in the ER leads to the formation of a lipid droplet body, which is proposed to bud off from the ER once it exceeds a certain size [155]. Little is known about the biological signals or post-translational modifications that regulate the enzymes of the FA esterification pathway [156, 157].

1.12.1.2 Lipolysis Lipolysis is the hydrolytic breakdown of TAG into three FAs and a glycerol backbone. This process occurs in the lipid droplet and is normally induced in adipocytes under conditions of negative energy balance to increase the availability of FA as a substrate for energy production in peripheral tissues. Lipolysis begins with the hydrolysis of TAG by adipose triglyceride lipase (ATGL) to form DAG, which is hydrolysed by hormone sensitive lipase (HSL) to form MAG (Figure 1-7). The process ends with the hydrolysis of MAG by MAG lipase (MGL) to form glycerol and FA, and the subsequent release of these products into the circulation (Figure 1-7, Figure 1-8) [158-160]. At least 50% of FA molecules released by lipolysis return to adipose tissue for re-esterification. The glycerol backbone however, is unable to re-enter the adipocyte. This inability enables 40 / GPAM

direct measures of lipolysis to be made in light of the capacity for released FAs to be re- esterified [161].

/ GPAM

/ LIPIN-1

1 / DGAT2

Figure 1-7 A simplified view of lipolysis and fatty acyl-CoA esterification pathways. Shown are simplified representations of the fatty acyl-CoA esterification (TAG synthesis) and lipolysis pathways. Refer to Sections 1.12.1.1 and 1.12.1.2 for more detailed descriptions of these pathways. Figure adapted from [154].

Under basal conditions, ATGL and HSL are mostly rendered inactive by lipid droplet surface proteins known as perilipins, which act as gatekeepers to the lipid droplet. Perilipins (mainly perilipin-1) achieve this by indirectly inhibiting ATGL activity at the lipid droplet and preventing HSL entry to the lipid droplet. MGL activity however, is known to be constitutive under basal conditions. Catecholamines and natriuretic peptides control the signalling cascades which facilitate lipolysis through protein kinase A (PKA) and protein kinase G (PKG)-mediated inhibitory phosphorylation of perilipin- 1 as well as stimulatory phosphorylation of HSL to promote its translocation to the lipid droplet. These pathways are further described in Figure 1-8 [18, 162].

Figure 1-8 Main signalling pathways of lipolysis. Shown are the main signalling pathways of lipolysis which are described as follows: Gs-protein-coupled β1/β2 adrenergic receptor (β1/2-AR) induces adenylyl cyclase (AC), to generate the second messenger cAMP, which then activates protein kinase A (PKA). Stimulation of the guanylyl cyclase (GC)-coupled type-A natriuretic peptide receptor (NPR-A) increases levels of the second messenger cGMP, which then activates protein kinase G (PKG). PKA or PKG then facilitate lipolysis by (i) phosphorylating HSL, inducing its translocation to the lipid droplet (ii) phosphorylation of perilipin-1 (PLIN1) to confer conformational changes which a) enable HSL to enter the lipid droplet b) release its hold on the comparative gene identification-58 (CG1-58) protein which is then able to activate adipose TAG lipase (ATGL). Conversely, stimulation of the Gi- protein coupled α2-adrenergic receptor (α2-AR) deactivates AC, causing decreases in cAMP and subsequent suppression of the cAMP-mediated lipolysis cascade. Figure adapted from [162].

42 1.12.1.3 De novo lipogenesis De novo lipogenesis is the conversion of excess carbohydrate to FAs for their eventual storage as TAG via FA esterification. Upregulation of this process occurs in response to glucose metabolism (which provides the substrate for de novo lipogenesis) as well as through insulin and glucose-mediated activation of the SREBP1 and ChREBP transcription factors, which induce the expression of genes encoding multiple lipogenic enzymes [18, 163]. During lipogenesis, intracellular glucose undergoes glycolysis in the cytosol to form pyruvate, which is then converted to citrate via the TCA cycle in the mitochondria. Newly-formed citrate escapes the mitochondria and is converted into acetyl CoA, then carboxylated to malonyl CoA by acetyl CoA carboxylase. Fatty acid synthase then converts malonyl CoA into FA, initially palmitate, which can be further converted into other species [164].

1.12.2 Adipokine secretion Adipose tissue is known to secrete over 600 proteins, termed adipokines [165]. These proteins are actively involved in many biological processes including insulin sensitivity, glucose metabolism, inflammation, energy expenditure and appetite [166, 167].

Adiponectin and leptin are two of the most studied adipokines and are generally considered to be metabolically protective. Adiponectin predominantly increases insulin sensitivity in skeletal muscle and liver through the activation of AMP-activated protein kinase (AMPK), which enhances the catabolism of fatty acids via stimulation of beta- oxidation and/or promotes glucose uptake via increasing GLUT4 translocation to the plasma membrane (reviewed in [168]). Leptin acts in response to positive energy balance and increases insulin sensitivity by (i) suppressing food intake through altering the function of hypothalamic neurotransmitters that control appetite (ii) increasing energy expenditure by suppressing de novo lipogenesis and increasing beta-oxidation and glucose uptake in the same manner as adiponectin [169, 170].

Adipose tissue also secretes a wide range of inflammatory cytokines which include TNFα, interleukin-6 (IL-6) and monocyte chemoattractant protein 1 (MCP-1) [171]. Inflammatory cytokines are proposed to mediate insulin resistance in peripheral organs through activation of inflammatory signalling pathways that acutely interrupt insulin

43 action through inhibitory phosphorylation of the insulin receptor and IRS proteins [172], activating ceramide synthesis enzymes [86, 173] and increasing local inflammatory cytokine production which subsequently amplifies this inhibition in insulin signalling in a feed-forward manner. Furthermore, the release of MCP-1 from adipocytes causes the recruitment of pro-inflammatory macrophages. These immune cells account for a high-proportion of inflammatory cytokine release from adipose tissue and therefore also serve to propagate inflammatory capacity of adipose tissue [83].

1.12.3 Functional differences amongst regionally distinct adipose tissue depots The main adipose tissue depots are divided into two main categories: visceral (intra- abdominal) adipose tissue (VAT) and subcutaneous (SubQ) adipose tissue. VAT encompasses all adipose depots that lie within the abdominal cavity. These include gonadal fat, retroperitoneal fat and mesenteric fat1. SubQ adipose tissue is located underneath the epidermis and is more prominent in the gluteal-femoral (lower-body) region in metabolically healthy individuals [174, 175].

In recent years, it has been increasingly recognised that these regionally distinct depots differ with respect to lipid metabolism and adipokine secretion. These differences have lead to the paradigm that it is the distribution pattern of body fat which is more closely linked to the pathophysiology of insulin resistance than obesity per se. As such, individuals with higher upper body adiposity, which often occurs with increased visceral fat, are found to have a higher risk of developing insulin resistance [176-180]. Mammalian sex-based differences in fat distribution are such that males preferentially store fat in the upper-body whilst females tend to store fat in lower-body SubQ regions. Consequentially, females are found to be more insulin sensitive and less susceptible to insulin resistance than males [181-183]. This trend is further discussed in the context of findings obtained in this study.

Adipose tissue expansion during overfeeding is manifested as hyperplasia (an increase in cell number through increased adipogenesis) or hypertrophy (an increase in cell size).

1 Only the nomenclature for murine adipose tissue depots has been listed to minimise confusion with regard to later sections of this thesis.

44 Hyperplasia is viewed as a more stable and effective way of storing lipid in adipose tissue as it spreads the distribution of FA, which in turn decreases its propensity for FA release [31, 184]. Under conditions of overfeeding, lower-body SubQ adipose tissue tends to expand via hyperplasia whilst VAT is inclined to expand via hypertrophy [185, 186]. This characteristic enables SubQ to act as a sink for sequestering FAs [174, 187]. As such, VAT is associated with greater lipolysis and FA release, lower responsiveness to anti-lipolytic stimulation and higher responsiveness to lipolytic stimulation in comparison with lower-body SubQ adipose tissue [188-190]. Additionally, VAT has been reported to secrete higher amounts of pro-inflammatory adipokines than lower- body SubQ adipose tissue, which conversely is observed to release higher amounts of adiponectin and leptin than VAT [191-196]. These differences above are proposed to account for the disparate effects of VAT and lower-body SubQ adipose tissue with respect to the generation and propagation of whole body insulin resistance. However, studies have demonstrated that SubQ adipose tissue located on top of the abdominal region expands via hypertrophy and is positively linked to insulin resistance [197-199]. Though these findings add complexity to the general metabolic definition of SubQ, it further highlights the impact of anatomical location upon the metabolic characteristics of an adipose tissue depot.

1.13 Adipose Tissue Dysfunction and Insulin Resistance Normal regulation of the lipid metabolic and secretory features of adipose tissue is lost during overfeeding and/or obesity. This manifests itself as enhanced FA production/release (via decreased esterification and increased lipolysis), increased pro- inflammatory cytokine production/release, decreased adiponectin release, increased conversion of adipose tissue resident macrophages to their pro-inflammatory phenotype and elevated pro-inflammatory macrophage infiltration [31, 183, 200, 201]. Though leptin is considered a metabolically beneficial adipokine, its concentrations increase with obesity as a consequence of the central development of leptin resistance in leptin- sensitive tissues [202, 203]. It is widely proposed that this dysregulation occurs in hypertrophic adipocytes which, in comparison with smaller adipocytes (which occur in hyperplasic adipose tissue), are more susceptible to developing insulin resistance, hypoxia, mitochondrial and ER dysfunction and chronic inflammation. It is believed that the link between the adipocyte hypertrophy and the development of these

45 pathogenic processes may stem from the increased FA exposure encountered by the enzymes and intracellular organelles that facilitate these processes. Altogether, these pathogenic susceptibilities further propagate adipose tissue dysfunction in a feed- forward manner and in turn, increase the ability of adipose tissue to incite pathogenic actions that cause insulin resistance in peripheral tissues. Therefore, it is generally accepted that larger adipocytes are more insulin resistant and have a greater potential to cause insulin resistance in peripheral tissues than smaller adipocytes [31, 183, 200, 201].

Various studies in rodents and humans have suggested that the predominant mechanism that underpins HFD-induced insulin resistance changes in accordance to the duration of high-fat feeding. During short-term high-fat feeding, it is thought that acute and/or chronic lipid flux from adipose to distal tissues is the main instigator in the development of insulin resistance. Prolonged high-fat feeding on the other hand, mediates the appearance of chronic, low-grade inflammation, which is postulated as the major factor in the maintenance and propagation of insulin resistance [26, 194, 204, 205]. Further investigation is required to confirm if these effects occur temporally and to determine the mechanisms that cause the switch from one predominant system to another.

1.14 Protein Kinase C The protein kinase C (PKC) family of enzymes is a group of ten serine/threonine kinases. PKC enzymes play important roles in several signal transduction cascades which mediate a multitude of physiological processes including glucose and lipid metabolism, inflammation, cardiac ischemia, synaptic function, cell proliferation, mitosis, differentiation, cell survival and apoptosis [141, 206-210].

PKC isoforms are expressed ubiquitously, with some isoforms more prevalent in specific tissues than others [211]. All PKC isoforms can be divided into three subfamilies as based on sequence homology and modes of activation. As such, PKC isoforms are classed as either being classical, novel or atypical. All three groups are comprised of an N-terminal regulatory domain and a C-terminal catalytic domain linked together by a hinge domain. The regulatory and catalytic domains are comprised of four mainly conserved regions (C1-C4) interspersed by five main variable regions (V1-V5),

46 including the hinge domain (Figure 1-9). The regulatory domain is the binding site for PKC co-factors, which serve to regulate PKC function. The features of this domain differ between each PKC subfamily, subsequently dictating the co-factors to which they respond. Classical PKCs (cPKCs) include the PKCα, β, and γ isoforms, which are activated through binding of Ca2+ and DAG at the C2 and C1 domain, respectively. Novel PKCs (nPKCs) include the PKCδ, ε, η and θ isoforms, which contain C2-like and C1 domains in their regulatory region. As the C2-like domain is unable to bind Ca2+, only DAG is required for the activation of nPKCs [141]. Activation of the cPKC and nPKC subfamilies under normal physiological conditions occurs temporarily as a result of acute catecholamine/growth factor-induced phospholipase C–mediated hydrolysis of membrane phospholipids, which releases Ca2+ and DAG. Furthermore, the activation of cPKCs and nPKCs specifically requires DAG molecules which contain unsaturated FAs [212]. The atypical PKC (aPKC) subfamily is comprised of the PKCζ and ι/λ isoforms, which only contain a C1-like domain in the regulatory region. This domain cannot accommodate DAG binding and so neither Ca2+ and/or DAG are able to activate aPKCs. Alternatively, aPKCs are activated by PDK1 and mTORC2 phosphorylation in the catalytic domain. Contained in the regulatory domain of all PKCs is a pseudosubstrate (PS) region, which binds to the substrate-binding site of PKC in an autoinhibitory fashion. The catalytic domain is conserved between all three PKC subfamilies and facilitates PKC kinase activity by providing sites for ATP and substrate binding at the C3 and C4 domains, respectively [213] (Figure 1-9).

C1 C2

C1C1 -likelike

Figure 1-9 The protein kinase C subfamilies. Protein kinase C (PKC) isoforms are divided into conventional, novel and atypical subfamilies. For a more detailed description of the structure and regulation of these isoforms refer to Section 1.14. Figure adapted from [214].

47 The activation mechanism for PKC is as follows: newly synthesized PKC isoforms are tethered to a membrane compartment by anionic membrane lipids where they are stabilized and rendered catalytically active through a number of phosphorylation events. This process, known as maturation, is mediated at conserved sites within the catalytic domain of PKC by the enzymes PDK-1, and mTORC2 [215-217]. PKC then forms an inactive, but catalytically competent conformation (the “mature” form), characterised by binding of the PS domain to the substrate-binding domain. This is followed by detachment from the membrane and translocation to the cytosol. In response to cell stimulation by hormones or neurotransmitters, receptor-mediated phospholipase C activation results in hydrolysis of membrane phospholipids and elevation of second messengers (Ca2+ and DAG). Binding of these to the regulatory domain of cPKCs and nPKCs enables them to translocate to the membrane and undergo conformational change which then releases the PS from the substrate binding domain, allowing PKC to perform kinase reactions. In the case of cPKCs, Ca2+ binding induces translocation to the membrane followed by interaction with membrane-based DAG to elicit full activation. Translocation and full activation of nPKCs is mediated via membrane-based DAG alone. Alternatively, the kinase activity of mature aPKCs is associated with PDK1 phosphorylation. Whether this action directly causes membrane translocation and/or release of the PS domain remains unclear [213, 218].

The site of action for individual PKC isoforms is determined by their affinity for particular scaffold proteins, such as receptors for activated C kinase (RACK). These are located at various cellular membrane compartments and function to anchor PKC isoforms to these regions after the initiation of mature PKC translocation [219, 220]. The specificity of individual PKC isoforms to different scaffold proteins is defined by sequence differences between the variable regions of each PKC isoform. This results in differential patterns of membrane localisation, exposing each isoform to a unique set of substrates; this subsequently dictates the signalling pathways mediated by an individual PKC isoform. Consequentially, the possibility of functional redundancy between PKC isoforms is precluded despite the high sequence homology of the kinase domain between these isoforms [208].

The DAG-mediated activation of novel PKCs places these enzymes as ideal candidates for facilitating the pathophysiological effects associated with chronic lipid oversupply, 48 including insulin resistance. In this case, DAG is assumed to be chronically elevated during increased FA esterification, rather than through phospholipid hydrolysis. Although cPKCs are partially activated by DAG, they are not generally seen as directly stimulated in response to chronic lipid excess because parallel increases in Ca2+ (a co- factor which is also required for full activation of cPKCs) during these conditions do not necessarily occur [141]. As mentioned in earlier sections, the nPKC subfamily is most implicated in the pathogenesis of insulin resistance. PKCθ is most often implicated during skeletal muscle insulin resistance, most likely on account of its relatively high levels of expression in this tissue in comparison to other PKC isoforms [221-224]. Of particular interest to our study however, is the close association between PKCε and hepatic insulin resistance [135-137, 140, 225]. PKCε will be further discussed in the remaining sections of this literature review.

1.15 Protein Kinase C epsilon The PKCε protein is most abundant in brain tissue and readily detectable in skeletal muscle, liver and adipose tissue [226]. The general mechanism of activation for PKCε is outlined in section 1.14. Further to this, the maturation of PKCε specifically requires phosphorylation at threonine 566, serine 729 and threonine-710 [206]. PKCε has been reported to localise in a number of subcellular locations including the plasma membrane [227], cytoskeleton [228], Golgi apparatus [229], mitochondria [230] and the nucleus [206]. This has implicated PKCε in a variety of signalling events which facilitate cytoskeletal remodeling [231], nociception, alcohol tolerance [232], carcinogenesis [233], cardiac preconditioning [214] and inflammation [234], in addition to insulin resistance [141]. Consequently, PKCε has increasingly been recognised as a key mediator of many pathologies including T2D, cardiovascular disease and various cancers.

The subcellular localisation of PKCε is dictated in part by its co-factor DAG, which directs this enzyme to the plasma membrane and presumably the ER, which is the intracellular site for DAG intermediates generated during FA esterification [154, 213]. Furthermore, the localisation of PKCε is influenced by protein interactions between PKCε and various scaffolding proteins. For instance, the binding of PKCε to actin

49 mediates its proximity to the cytoskeleton, whilst its interaction with the adaptor protein myeloid differentiation primary response gene 88 (Myd88) docks PKCε to the inflammatory signalling receptor TLR4 at the plasma membrane [228, 235]. However, it is PKCε binding to RACK scaffolding proteins that has garnered extensive research. The V1 domain of PKCε is known to bind to RACK2 (also known as beta’-COP) and, to a much lesser extent, RACK1. These interactions anchor PKCε to the Golgi apparatus and plasma membrane, respectively [229, 236]. The exclusive specificity of RACK2 to PKCε has led the Mochly-Rosen group to develop peptides that exclusively inhibit and activate PKCε translocation by mimicking the binding properties of RACK2 at the V1 domain [237-239]. The generation of these peptides has essentially accelerated the elucidation of PKCε−specific functions, which was previously occluded to an extent due to the high sequence homology between the regulatory and catalytic domains of PKC isoforms. In one such study, our laboratory utilised the εV1-2 inhibitory peptide to inhibit PKCε in mouse models of T2D. Through the use of this peptide, we were able to further demonstrate the ability of PKCε to impair insulin secretion [240].

Despite these advances, the protein targets of PKCε that mediate its physiological and pathophysiological effects are poorly defined, particularly those involved in glucose metabolism [208]. In parallel, there is conflicting evidence in the literature regarding a mechanistic link between DAG accumulation, chronic PKCε translocation and the inhibition of canonical insulin signalling [137, 143-145] [140]. This suggests that PKCε may not be impairing glucose homeostasis through the simple inhibition of insulin signalling. Alternatively, a multitude of studies from our laboratory have shown that PKCε itself modulates lipid metabolism in a number of tissues, which may reconcile observed improvements in glucose and lipid metabolism following the global knockout of PKCε or inhibition of its activity. These investigations are discussed below.

50 1.16 Protein Kinase C epsilon, Lipid Metabolism and Insulin Resistance A direct role for PKCε in lipid metabolism was initially highlighted through a study published by our laboratory in 2007. This study demonstrated that pancreatic islets from global PKCεKO mice displayed enhanced glucose-stimulated insulin secretion in comparison with WT islets only after FA pretreatment ex vivo. This was suggested to be driven by a further increase in the FA esterification to FA oxidation ratio displayed by islets from global PKCεKO mice. A role for PKCε in FA esterification is plausible considering it is likely to localise at the ER. Overall, the findings from this study suggested that the global deletion of PKCε mediated an augmentation of the amplification pathway of insulin secretion via altered lipid partitioning during conditions of FA oversupply [240].

This was further investigated to elucidate the effects of PKCε with respect to lipid metabolism in endogenous lipid pools within islets. It was found that pre-incubation of islets from global PKCεKO mice with radiolabelled palmitate was associated with a significant increase in radiolabelled TAG. Further studies suggested that the absence of PKCε enhanced the cycling of lipids between esterification and lipolysis, rather than esterification per se. This was proposed to account for the observed enhancement of the amplification pathway and furthermore, was suggested to act an adaptive mechanism to curtail the toxic effects of lipid oversupply [241].

1.16.1 A potential role for PKCε in adipose tissue Suzuki et al. showed that PKCε and PKCη, very closely related PKC isoforms, are associated with lipid droplets [242]. Consistent with this, Kumashiro et al. determined that PKCε was localized to the lipid droplet and activated by DAG molecules resident in this compartment [243]. Although these studies were conducted using hepatocytes or liver extracts, it is possible that this distribution also occurs in the adipocyte, which is predominantly composed of a large lipid droplet and has been shown to express readily detectable levels of PKCε protein [149, 226].

51 Evidence of a role for PKCε in adipose tissue was provided by a 48-hour fasting study on global PKCεKO mice [244]. During states of negative energy balance, such as prolonged fasting, insulin levels are low and lipolysis is stimulated. Under these conditions, global PKCεKO mice displayed larger fat pad mass, decreased plasma FA concentrations, decreased liver TAG and reduced ketone bodies (which are mostly synthesised at the liver using FA as a substrate) than WT mice. This suggested global ablation of PKCε mediated a reduced mobilization of FA from adipose tissue to the liver. These findings also indicate that PKCε may be eliciting direct effects upon adipose tissue lipid metabolism independent from insulin [244]. Furthermore, as described in Section 1.11, a number of investigations from our laboratory have suggested the possibility that PKCε modulates lipid metabolism within the adipocyte, which in turn may account for associated improvements in glucose tolerance and liver insulin sensitivity observed in mouse models of global PKCε deletion. Despite these promising findings, the direct in vivo effects stemming specifically from PKCε action in adipose tissue in the context of lipid and glucose metabolism are yet to be defined.

52 1.17 Summary The association between dietary lipid oversupply and insulin resistance has long been acknowledged. The two main bioactive lipid metabolites implicated in the generation and propagation of insulin resistance upon dietary lipid excess are ceramide and DAG. A better understanding of the underlying mechanisms linking the action of these metabolites to insulin resistance is still required.

Surprisingly, our laboratory has recently demonstrated that individual ceramide species in vitro were associated with unchanged and/or improved skeletal muscle glycogen synthesis and insulin signalling after modulating the expression of CerS enzymes. These novel observations however, require further examination.

The general consensus of the current literature implies that DAG mediates hepatic insulin resistance through the activation of PKCε in the liver. We have shown that global ablation of PKCε in mice improves hepatic insulin sensitivity, but not via direct actions at the liver. In fact, the nature of our findings suggest that PKCε deletion in adipose tissue may improve hepatic insulin sensitivity through decreasing FA flux from adipose tissue to the liver. Preliminary and published studies employing various models of global PKCε deletion provide early evidence to suggest that PKCε directly affects lipid metabolism. Direct exploration of this possibility however, has not been conducted.

53 1.18 Thesis Aims The specific aims of this thesis are:

- To assess the effect of CerS overexpression in L6 myotubes upon the mRNA expression of sphingolipid metabolism proteins, flux through sphingolipid metabolism pathways and on additional components of insulin action.

- To characterise the effects of PKCε deletion specifically in adipose tissue on glucose tolerance, FA release and insulin sensitivity in mice.

- To examine the mechanisms of lipid metabolism which may underpin the link between PKCε ablation in adipose tissue and potential associated alterations in glucose tolerance, FA release and insulin sensitivity in mice.

54 CHAPTER 2 MATERIALS AND METHODS

55 2.1 Materials The table below lists materials that were used in this study, including their manufacturer and country of origin. Reagents mentioned in this chapter, but not listed in this table were prepared in-house by the Garvan Institute of Medical Research Media Preparation Facility (Garvan Institute of Medical Research, NSW, Australia).

Table 2 Materials used in this study

Material Manufacturer Location [3,3H]-glucose Perkin Elmer Waltham, MA, USA 0.5ml insulin syringe BD Biosciences San Jose, CA, USA Life Technologies 10% glycogen (DEPC-treated) (Ambion) Carlsbad, CA, USA 10% Neutral Buffered Formalin Traralgon, VIC, solution (4% formaldehyde) Australian Biostain AUS 2[14C]deoxyglucose Perkin Elmer Waltham, MA, USA [3H]-Serine Perkin Elmer Waltham, MA, USA [3H]-Sphingosine Perkin Elmer Waltham, MA, USA Foster City, CA, 7900-HT Real Time PCR System Applied Biosystems USA 96-well assay plates Corning Corning, NY, USA 96-well conical bottom plates Grenier Bio-One Frickenhausen, GER Absolve Perkin Elmer Waltham, MA, USA Accu-check II glucometer (euglycaemic-hyperinsulinaemic Roche Diagnostics Castle Hill, NSW, clamp studies only) Australia AUS AccuCheck Performa glucose Roche Diagnostics Castle Hill, NSW, monitor Australia AUS Roche Diagnostics Castle Hill, NSW, AccuChek Performa Test Strips Australia AUS Thermo Fisher Scientific Taren Point, NSW, Acetone (analytical grade) (AjaxFinechem) AUS Novo Nordisk - gift from Garvan Institute Clinical Actrarapid human insulin Facility Bagsvaerd, DEN Mountain View, CA, Adeno-X Rapid Titre Kit Clontech USA AG 1-X8 Resin (acetate form) Bio-Rad Laboratories Hercules, CA, USA MP Biomedicals Seven Hills, NSW, AIN Vitamin Mix Australia AUS Carlsbad, CA, USA / Charles Perkins Alexa Fluor 488-conjugated Centre - University fluorescent secondary antibody Life Technologies - gift of Sydney, NSW, (goat anti-mouse) from Dr. Nolan Hoffman AUS 56 Alexa Fluor 680-conjugated fluorescent secondary antibody (goat anti-rabbit) Thermo Scientific Rockford, IL, USA Amitriptyline hydrochloride Sigma-Aldrich St-Louis, MO, USA Anion exchange columns Bio-Rad Laboratories Hercules, CA, USA Anti-haemagglutinin (HA) antibody Covance Princeton, NJ, Antibiotic/antimycotic cocktail (100X stock) Life Technologies Carlsbad, CA, USA Pierce - Thermo BCA Protein Assay Kit Scientific Rockford, IL, USA Beckman LS6000SC Beta Counter Beckman Coulter Brea, CA, USA Beckman tubes Beckman Coulter Brea, CA, USA Bio-Rad Protein Standards Bio-Rad Laboratories Hercules, CA, USA Bio-Rad Protein Transfer System Bio-Rad Laboratories Hercules, CA, USA Bovine Serum Albumin (BSA) Sigma-Aldrich St-Louis, MO, USA C-terminal HA-containing primers (attached to CerS2 and CerS6 Integrated DNA Baulkham Hills, cDNA constructs) Technologies NSW, AUS CBA Enhanced Sensitivity Capture Beads and Mouse Detection Reagent Standards (all plasma cytokines measured) BD Biosciences San Jose CA, USA CerS1, Cers4, CerS5 cDNA Weizman Institute of constructs tagged at C-terminal Gift from Prof. Anthony Science, Rehovot, with HA Futerman Israel CerS2 and CerS6 cDNA American Type Culture constructs Collection Manassas, VA, USA Thermo Fisher Scientific Taren Point, NSW, Chloroform (AjaxFinechem) AUS Chloroform (molecular biology grade) Sigma-Aldrich St-Louis, MO, USA Clamp pumps (11-plus) Harvard Apparatus Holliston, MA, USA Clear bottom, black wall 96 well plates Grenier Bio-One Frickenhausen, GER Collagenase D Roche Diagnostics Mannheim, GER cOmplete, EDTA-free Protease Castle Hill, NSW, Inhibitor Cocktail Tablets Roche Diagnostics AUS Cytometric Bead Array (CBA) Mouse/Rat Soluble Protein Master Kit BD Biosciences San Jose CA, USA D-(+)-Glucose powder Sigma-Aldrich St-Louis, MO, USA DEPC-treated water Life Technologies Carlsbad, CA, USA Dimethyl Sulfoxide (DMSO) Sigma-Aldrich St-Louis, MO, USA Dispase Roche Diagnostics Mannheim, GER 3 En Hance Spray Perkin Elmer Castle Hill, NSW, AUS epMotion dispenser Eppendorf Hamburg, GER. 57 Eppendorf tubes (1.5ml, 2mL) Eppendorf Hamburg, GER. Ethanol (molecular biology grade) Sigma-Aldrich St-Louis, MO, USA FACSCantoII platform flow cytometer BD Biosciences San Jose CA, USA FCAP Array software BD Biosciences San Jose CA, USA FLUOstar Omega Plate Reader BMG Labtech Offenburg, GER Thermo Fisher Scientific Foetal Calf Serum (FCS) Australia Scoresby, VIC, AUS Gateway cloning system including pAd/CMV/V5-DEST cloning vector Life Technologies Carlsbad, CA, USA Trajan (SGE Analytical Melbourne, VIC, Glass syringe Science) AUS Lane Cove West, Glucose Solution (50%) Phebra NSW, AUS Glycerol Sigma-Aldrich St-Louis, MO, USA GraphPad Prism software (version 6) GraphPad Software Inc La Jolla, CA, USA GW4869 Sigma-Aldrich St-Louis, MO, USA Heparinised saline Astra-Zeneca London, UK HEPES Life Technologies Carlsbad, CA, USA Igepal CA-640 Sigma-Aldrich St-Louis, MO, USA National Institute of Image J 1.47v software Health Bethesda, MD, USA Immobilon PVDF Membrane-F Millipore Billerica, MA, USA Immobilon PVDF Membrane-P Millipore Billerica, MA, USA Iodine (solid) Sigma-Aldrich St-Louis, MO, USA IR Dye 800-conjugated fluorescent secondary antibody (goat anti-mouse) LI-COR Lincoln, NE, USA Isopropanol (molecular biology grade) Sigma-Aldrich St-Louis, MO, USA Melbourne, VIC, Kodak MR film Kodak AUS Gift from Dr Amira Klip, The Hospital for Sick Toronto, Ontario, L6 skeletal muscle cells Kids CAN Charles Perkins L6 skeletal muscle cells stably Centre - University expressing HA-tagged GLUT4 Gift from Dr. Nolan of Sydney, NSW, receptor Hoffman AUS Mount Waverley, Lard (Allowrie brand) Fonterra Brands VIC, AUS Leica DM4000 Leica Microscopes Wetzler, GER LI-COR Odyssey Scanner LI-COR Lincoln, NE, USA LI-COR Odyssey Software LI-COR Lincoln, NE, USA LightCycler 480 Analysis Software Roche Diagnostics Mannheim, GER

58 LightCycler 480 ProbesMaster real time-PCR master mix Roche Diagnostics Mannheim, GER LightCycler 480 System I real time-PCR machine Roche Diagnostics Mannheim, GER Lipid standard – ceramide Sigma-Aldrich St-Louis, MO, USA Lipid standard - glycosylceramide Sigma-Aldrich St-Louis, MO, USA Lipid standard – sphingomyelin Sigma-Aldrich St-Louis, MO, USA Lipid standard – sphingosine Sigma-Aldrich St-Louis, MO, USA Sigma Aldrich (Vetec Castle Hill, NSW, Methanol Fine Chemicals) AUS Minimum Essential Media-alpha Life Technologies medium plus nucleosides (GIBCO) Carlsbad, CA, USA MiniSart Syringe Filter Sartorius Stedim Göttingen, GER MOPC antibody (for GLUT-4 assay) Sigma-Aldrich St-Louis, MO, USA MOPS Sigma-Aldrich St-Louis, MO, USA NanoDrop ND-1000 Wilmington, DE, Spectrophotometer Nanodrop Technologies USA Wako Pure Chemical NEFA C kit Industries Osaka, Japan Glostrup, DEN / Charles Perkins Centre - University Dako (gift from Dr. of Sydney, NSW, Normal Swine Serum Nolan Hoffman) AUS NuPAGE 10X Sample Reducing Agent Life Technologies Carlsbad, CA, USA NuPAGE 4-12% Bis-Tris Gel 1.5mm 15 well Life Technologies Carlsbad, CA, USA NuPAGE 4X LDS Sample Buffer Life Technologies Carlsbad, CA, USA NuPAGE SDS Gel System Life Technologies Carlsbad, CA, USA NuPAGE SDS Gel System Life Technologies Carlsbad, CA, USA pAd/CMV/V5-DEST cloning vector containing b-galactosidase cDNA Life Technologies Carlsbad, CA, USA Palmitic Acid (>99% purity) Nu-Chek Prep Inc Elysian, MN, USA Electron Microscopy Paraformaldehyde 16% Services Hatfield, PA, USA Aldrich (now Sigma Phenol red poweder Aldrich) St-Louis, MO, USA Phenylmethylsulphonyl fluoride (PMSF) Sigma-Aldrich St-Louis, MO, USA PhotoshopCS5 (Version12.5 x64) Adobe Systems San Jose CA, USA Polyethylenesorbitan (Tween-20) Sigma-Aldrich St-Louis, MO, USA Polytron homogeniser Kinematica Lucerne, SUI Potassium phosphate Sigma-Aldrich St-Louis, MO, USA Integrated DNA Baulkham Hills, Primers: Adipoq-Cre genotyping Technologies NSW, AUS 59 Primers for genotyping the PKCε gene with exon 1 flanked by loxP sites Sigma-Aldrich St-Louis, MO, USA Puromyocin Sigma-Aldrich St-Louis, MO, USA Life Technologies RNase-free sodium chloride (Ambion) Carlsbad, CA, USA Doncaster, VIC, RNeasy Mini Kit Qiagen AUS Kingaroy, QLD, Safflower oil Proteco Gold AUS Sample acquisition tubes for flow cytometry BD Biosciences San Jose CA, USA Saponin Sigma-Aldrich St-Louis, MO, USA Scintillation vials (glass) - 20mL Perkin Elmer Waltham, MA, USA Sheep anti-mouse secondary antibody conjugated to Buckinghamshire, horseradish peroxidase GE Healthcare UK Hawthorn East, VIC, Skim milk powder (Western blot) Coles Supermarkets AUS Sodium azide Sigma-Aldrich St-Louis, MO, USA Sodium chloride Sigma-Aldrich St-Louis, MO, USA Sodium deoxycholate Sigma-Aldrich St-Louis, MO, USA Sodium fluoride Sigma-Aldrich St-Louis, MO, USA Sodium orthovanadate Sigma-Aldrich St-Louis, MO, USA Sodium pyrophosphate Sigma-Aldrich St-Louis, MO, USA Sonicator Ultrasonic Processor Famingdale, NY, XL Heat Systems USA Gordon's Specialty Yanderra, NSW, Standard chow diet Stockfeeds AUS College Station, TX, StataSE v9.2 software Stata Corporation USA Toongabbie, NSW, Sterile irrigation water Baxter AUS Sterile water for injection (10mL) ampule Astra-Zeneca London, UK Stripettes (plastic - 5mL, 10mL, 25mL) Corning Corning, NY, USA Syringe needles (21G, 23G) BD Biosciences San Jose CA, USA TaqMan Reverse Transcription Foster City, CA, Kit Applied Biosystems USA Test tubes (plastic) - 15mL, 50mL Corning Corning, NY, USA Thin layer chromatography silica plates Merck Millipore Billerica, MA, USA St. Helens, TAS, Thin Layer Chromatography tanks Analtech AUS Tissue culture dishes (10cm) Corning Corning, NY, USA Tissue culture flasks (T150) Corning Corning, NY, USA

60 Tissue culture plates (12, 24 well) BD Biosciences San Jose, CA, USA TRI-Reagent Sigma-Aldrich St-Louis, MO, USA Trizma base Sigma-Aldrich St-Louis, MO, USA Trypsin-EDTA (1X stock) Sigma-Aldrich St-Louis, MO, USA Ultima Gold XR Scintillation Fluid Perkin Elmer Waltham, MA, USA Ultra Sensitive Mouse Insulin ELISA Kit Crystal Chem IL, USA Western-Lightning Plus ECL Waltha reagents Perkin Elmer m, MA, USA Brookvale, NSW, X-Ray film Fujifilm AUS

61 The table on the following page lists the forward primer, reverse primer, and the identification number of the Universal Probe Library (UPL) probe required for the mRNA expression analysis of each gene in this study via real-time PCR. All primers were designed via the UPL Assay Design Centre website (Roche) listed below: http://lifescience.roche.com/webapp/wcs/stores/servlet/CategoryDisplay?catalogId=100 01&tab=Assay+Design+Center&identifier=Universal+Probe+Library&langId=-1

Primers for all genes, except srebp-1c, were manufactured by Integrated DNA Technologies (Baulkham Hills, NSW, Australia). Primers for the srebp-1c gene were manufactured by Sigma-Aldrich (St-Louis, MO, USA). All UPL probes were manufactured by Roche (Mannheim, Germany).

62 Table 3 Real-time PCR primers and Universal Probe Library probes used in this study

Probe Gene Forward primer Reverse Primer # Adiponectin GGAGAGAAAGGAGATGCAGGT CTTTCCTGCCAGGGGTTC 17 Agpat-2 AAGACGAAGCTCTTCACCTCA TCTGTCAGACCATTGGTAGGG 42 Atgl TGACCATCTGCCTTCCAGA TGTAGGTGGCGCAAGACA 104 Cd68 GACCTACATCAGAGCCCGAGT CGCCATGAATGTCCACTG 96 CerS1 GCCTCTTCCTATGCGTTCC CAGCTGCACATCGCTGAC 4 CerS2 ACCGGTCAGCTTTGCACT CGTTCCCACCAGAAGTAGTCA 50 CerS4 TGAAGCAGAGACCAGTGGAG AATCTGCCGCAACGTGAG 79 CerS5 GACAGTCCCATCCTCTGCAT TGTTCGTGTGTGTGGTCTCA 5 CerS6 TGGTTTCGACAAAGGCGTA AGAGGTAAAAGGAAAATCTCCACA 2 Cyclophillin B TTCTTCATAACCACAGTCAAGACC ACCTTCCGTACCACATCCAT 20 Dgat-1 ACCTGGCCACAATCATCTG TGGAGTATGATGCCAGAGCA 3 Dgat-2 GGCGCTACTTCCGAGACTAC TGGTCAGCAGGTTGTGTGTC 42 F4/80 CCTGGACGAATCCTGTGAAG GGTGGGACCACAGAGAGTTG 1 Gpam GGAAGGTGCTGCTATTCCTG TGGGATACTGGGGTTGAAAA 33 Hsl AGCGCTGGAGGAGTGTTTT CCGCTCTCCAGTTGAACC 3 Il-1β TGTAATGAAAGACGGCACACC TCTTCTTTGGGTATTGCTTGG 78 Inos GGAGCCTTTAGACCTCAACAGA AAGGTGAGCTGAACGAGGAG 3 Leptin CAGGATCAATGACATTTCACACA GCTGGTGAGGACCTGTTGAT 93 Lipin-1 CGCAACATCTTGCCAAACT CCACGTAATTCATTGTCTGAACC 3 Mcp-1 CATCCACGTGTTGGCTCA GATCATCTTGCTGGTGAATGAGT 62 Ormdl2 GGACTACAGTTTACCTCCTCACG AGAAGCTGGCCAGTAGGTAGAG 73 Ormdl3 ACCCTCACCAACCTTATCCA ATAGTCCATCTGCTCCCAGTG 66 Perilipin-1 AACGTGGTAGACACTGTGGTACA TCTCGGAATTCGCTCTCG 64 Perilipin-2 TGAGTCCCACTGTGTTGAGC CAGGACAGTCTGGCATGTAGTC 3 Phlda1 CGCACCAGCCTCTTCACT CCGAAGTCCTCAAAACCTTG 13 Pparα CCGAGGGCTCTGTCATCA GGGCAGCTGACTGAGGAA 78 Pparγ GAAAGACAACGGACAAATCACC GGGGGTGATATGTTTGAACTTG 7 Saa ATGCTCGGGGGAACTATGAT ACAGCCTCTCTGGCATCG 26 Sptlc1 GGTGCTGGTGGAGATGGT GGATTCCTTCCAGAATTAGATGG 67 Sptlc2 GGATTTGCGACAAATTCAATG TCACTCAGAATCAGGCAACCT 15 Srebp-1c CATGGATTGCACATTTGAAGA TCAGGAGAGTTGGCACCTG 3 Tbp CCCACCAGCAGTTCAGTAGC CAATTCTGGGTTTGATCATTCTG 129 Tgf-β TGGAGCAACATGTGGAACTC GTCAGCAGCCGGTTACCA 72 Tnfα TCTTCTCATTCCTGCTTGTGG GGTCTGGGCCATAGAACTGA 49

63 The table below lists all primary antibodies used for immunoblotting in this study including target protein size, manufacturer, species the antibody was raised in and working dilution.

Table 4 Immunoblot primary antibodies used in this study

Species Antibody Size (kDa) Manufacturer Raised Dilution Sigma-Aldrich β−ΑCΤΙΝ 42 St-Louis, MO, USA #Α5441 Mouse 1:2000 Abnova CerS6 33 Taipei City, Taiwan #H00253782-M01 Mouse 1:1000 Abcam DGAT-1 55 Cambridge, England, UK #ab54037 Rabbit 1:500 Santa Cruz Biotechnology DGAT-2 44 Dallas, TX, USA #sc-66859 Rabbit 1:200 Abcam GPAM 94 Cambridge, England, UK #ab69990 Rabbit 1:500 Covance Tagged HA-tag Princeton, NJ, USA protein size #MMS-101R Mouse 1:1000 Cell Signaling Technology LIPIN-1 130 Danvers, MA, USA #5195 Rabbit 1:500 Biomedical Technologies MYOSIN IIA 227 Stoughton, MA, USA (non muscle) #BT-567 Rabbit 1:5000 Cell Signaling Technology PERILIPIN-1 62 Danvers, MA, USA #9349 Rabbit 1:1000 Cell Signaling Technology Phospho-AKT 60 Danvers, MA, USA (Ser473) #9271 Rabbit 1:1000 BD Biosciences PKCε 90 San Jose CA, USA #610086 Mouse 1:500 Cell Signaling Technology Total-AKT 60 Danvers, MA, USA (pan) #4685 Rabbit 1:1000 Cell Signaling Technology Total-HSL 81-83 Danvers, MA, USA #4107 Rabbit 1:1000

64 2.2 Cell Studies

2.2.1 L6 cell culture, fatty-acid treatment and insulin stimulation The rat skeletal muscle cell line, L6, was used for the CerS experiments outlined in this thesis. The L6 cell line was developed by Yaffe in 1968 [245] and is one of the most widely used skeletal muscle cell lines for investigating the effects of insulin resistance on skeletal muscle [45, 131, 246, 247]. Under low serum concentrations in vitro, L6 cells are able to spontaneously differentiate from embryonic progenitor stem cells (myoblasts) into mature, multinucleated skeletal muscle cells (myotubes). This process involves myoblast fusion and the expression of muscle-specific proteins such as muscle actin and muscle myosin [248]. The following cell culture and treatment procedures were carried out as outlined in [45, 246]. L6 cells were used between passages 10-20 and were maintained at 37°C in T150 flasks with Minimum Essential Medium (MEM)- alpha media containing 5.6 mM glucose, 10% (v/v) heat-inactivated foetal calf serum (FCS - high serum conditions) and 1% (v/v) antibiotic/antimycotic cocktail (100 units/mL penicillin, 100µg/mL streptomycin, 0.25µg/mL amphotericin B) – referred to herein as MEM-alpha media, unless otherwise specified. Cell growth was monitored daily and media replaced every two days.

2.2.1.1 Cell passaging At 50-60% confluency (approx. every two to three days) cells were passaged as follows: MEM-alpha media was aspirated and cells were washed two times with phosphate buffered saline (PBS). Cells were then coated with 1mL of trypsin-EDTA solution (1X) then incubated at 37°C for two minutes to detach cells from the flask wall. The flask was then firmly tapped to ensure cell dislodgement. Cells were both washed off the flask wall and re-suspended with 9mL of fresh MEM-alpha media. 1mL of re- suspended cells was then transferred to a fresh T150 flask and diluted with 19mL of fresh MEM-alpha media.

65 2.2.1.2 Cell seeding, differentiation and general treatment procedure Trypsinised cells re-suspended in 9mL of fresh MEM-alpha media were counted using a haemocytometer and seeded for experiments between passages 10-20 at the following concentrations:

12-well plates: 5 x 104 cells/mL, 1mL/well MEM-alpha media 24-well plates: 5 x 104 cells/mL, 0.6mL/well MEM-alpha media 96-well plates: 100µl/well of cells harvested at 90% confluency in 10cm dishes. Cells not counted prior to seeding in 96-well plates.

At approx. 90% confluency (two days after seeding), media was replaced with an equivalent volume of differentiation media comprised of MEM-alpha media containing 5.6 mM glucose, 2% heat inactivated FCS (low serum conditions) and 1% (v/v) antibiotic/antimycotic cocktail (100 units/mL penicillin, 100µg/mL streptomycin, 0.25µg/mL amphotericin B) to induce myoblast differentiation into myotubes. Two days later, media was replaced with an equivalent volume of fresh differentiation media and infected for 24 hours with adenovirus containing cDNA encoding β-galactosidase (LacZ) or CerS isoforms to facilitate LacZ or CerS protein overexpression (see Section 2.2.1.3 for details on adenovirus generation and adenoviral doses employed in this study). Cells were then treated with lipid for 16 hours by replacing differentiation media with an equivalent volume of differentiation media containing 5% (w/v) bovine serum albumin (BSA) coupled to 0.375 mM of palmitate (see section 2.2.1.4 for details on preparation of this media). This was followed by incubating cells for a further four hours with serum-free MEM-alpha media containing 5% (w/v) BSA coupled to 0.375 mM of palmitate. Cells were stimulated with 100nM insulin for the last 10 and 20 minutes of serum starvation prior to analysis of insulin signalling (during SMase inhibition) and GLUT4 translocation, respectively. See section 2.2.1.5 for details on preparation of insulin.

66 2.2.1.3 Generation and use of recombinant adenovirus Adenovirus containing either CerS isoforms or LacZ cDNA was generated, amplified, concentrated and quantified prior to the commencement of this study. In brief, cDNA constructs encoding CerS6 as well as CerS1, CerS2 CerS4 and CerS5 tagged with C- terminal haemagglutinin (HA) were cloned into the pAd/CMV/v5-DEST vector using the Gateway cloning system kit which also contained the pAd/CMV/v5-DEST vector housing LacZ cDNA. The vectors were linearised via restriction enzyme digestion and transfected into human embryonic kidney (HEK) 293 cells to facilitate adenoviral formation and subsequent adenoviral propagation. All steps performed by Mana Liao (Garvan Institute of Medical Research, NSW, Australia). Adenoviral stocks were amplified by infecting multiple flasks of fresh HEK 293 cells with adenovirus- containing media from the original adenoviral stock flask. Infected cells were pooled and lysed. Virus was isolated and concentrated from cell lysates via caesium chloride density gradient centrifugation and viral titre estimated using the Adeno-X Rapid Titre Kit (Clontech) (both steps performed as outlined in the PhD thesis of David Pederson, University of New South Wales, NSW, Australia).

Adenoviral-mediated protein overexpression of CerS isoforms was confirmed and optimal adenoviral dose for L6 myotube infection was also determined prior to the commencement of this study. In brief, L6 myotubes were infected with control adenovirus containing LacZ cDNA or increasing doses of adenovirus containing CerS1, CerS2, CerS4, CerS5 or CerS6 cDNA. Protein from these cells was harvested, quantified and immunoblotted for HA (tagged to CerS1, CerS2, CerS4, and CerS5 cDNA) or CerS6 as per sections 2.2.3 and 2.2.4. From the immunoblot data, (shown in Figure A-1 - Appendix), we chose to infect L6 myotubes for previous L6 cell studies with 0.5 µl (CerS1), 0.2 µl (CerS2), 0.25 µl (CerS4), 0.25 µl (CerS5) and 0.5 µl (CerS6) of concentrated adenoviral stock per mL of differentiation media. The same doses were used for this study.

67 2.2.1.4 Preparation of palmitate:BSA conjugate Palmitate-containing media stocks of 20x palmitate:BSA were prepared as follows: 9.7mg of palmitic acid powder was dissolved in 990µL of absolute ethanol to yield a palmitate concentration of 37.5mM. 20g of BSA was dissolved in 100mL of differentiation or serum-free media to make 4X BSA (20% w/v) media stocks which were stored at -20°C until required. Dissolved palmitate was then diluted 1:25 with 4X BSA-media stock pre-warmed at 37°C, then incubated at 37°C for 15 minutes to facilitate palmitate coupling to BSA. The conjugate was then diluted with pre-warmed differentiation or serum free media to make a final concentration of 1X BSA (5% w/v) and 0.375mM palmitate. This was sterile-filtered through a 0.2µm filter and applied immediately to cells as described earlier. Non-palmitate 1X BSA (5% w/v) control media (for non palmitate-treated control cells) was prepared as above, except no palmitic acid was applied to absolute ethanol prior to incubation with 4X (20% w/v), BSA media.

2.2.1.5 Preparation of insulin 0.6mM insulin stock was transferred to a glass vial and diluted 1:60 with sterile saline. 10µL of diluted insulin per mL of cell media was applied to stimulate cells at a final insulin concentration of 100nM.

2.2.2 mRNA expression analysis L6 cells were seeded in 24-well plates, differentiated, infected with adenovirus to overexpress LacZ and CerS isoforms and then treated in the presence or absence of palmitate as described above. Total RNA was harvested from the L6 myotubes using the Qiagen RNeasy Mini Kit as per manufacturer’s instructions. RNA concentrations were measured using the Nanodrop ND-1000 Spectrophotometer to read the absorbance at 260nm. 450-1150ng of RNA was reversed transcribed to form cDNA using the TaqMan Reverse Transcription Kit as per manufacturer’s instructions. cDNA was stored at -20°C and RNA was stored at until -80°C until further required.

Reagents required for the real-time (RT) PCR reaction were prepared as follows. cDNA for each sample was manually dispensed in a 96-well plate. In a separate tube, the forward primer, reverse primer and UPL probe for each gene were mixed together and 68 heated to 95°C for one minute, then combined with LightCycler480 Probes Master real time-PCR master mix. A list of all primers and probes used in this study is provided in Table 3. Reagents were automatically transferred into a 384-well plate using an epMotion dispenser robot to assemble the 10µL RT-PCR reaction mixture. This was comprised of cDNA (5ng), forward primer (0.5µM), reverse primer (0.5µM), probe (0.2µM) and master mix (1X). The RT-PCR reaction was conducted in a LightCycler 480 System I RT-PCR machine. The amplification (Ct) value for each gene was automatically determined by the LightCycler 480 software accompanying the aforementioned RT-PCR machine and was corrected by a housekeeping gene (TATA box binding protein) to obtain deltaCt values which were then used in the delta-deltaCt calculation method to express mRNA expression of each gene as fold change compared to control-treated samples. Standard curves were constructed for each gene of interest using cDNA combined from all samples to determine amplification efficiency.

2.2.3 Protein harvest and quantification Following treatment procedures, L6 myotube protein lysates were prepared for protein quantification and/or immunoblotting. Cells were washed two to three times with ice- cold PBS, which was aspirated after the final wash. Protein was harvested for the sphingolipid flux assays by applying 250µL of 0.1% (w/v) SDS (in PBS) per well of a 12-well plate, scraping and transferring cells to Eppendorf tubes and then sonicating cells. For all other experiments, protein was harvested by applying 100µL (per well of a 12-well plate) of ice-cold radioimmunoprecipitation assay (RIPA) buffer (see Appendix Section B for composition), scraping cells on ice and transferring to Eppendorf tubes. Cells were homogenised via sonication and resultant lysates were separated from cell debris via centrifugation at 13000 rpm for 20 minutes at 4°C.

To quantify protein concentrations for the sphingolipid flux assays, 5µL of cell lysate was diluted in 25µL of water and quantified using the BCA protein assay kit, as per manufacturer’s instructions. For the SMase inhibitor studies, 5µL of cell lysate was diluted in 5µL of ice-cold RIPA buffer and quantified for protein with the BCA protein assay kit, as per manufacturer’s instructions in combination with BSA standards dissolved in RIPA buffer prepared in-house.

69 2.2.4 Immunoblotting Equal amounts of protein (7-30µg) were combined with relevant amounts of 4X LDS sample buffer and 10X reducing agent, then denatured at 70°C for 10 minutes. Proteins were then resolved by size on pre-cast 4-12% Bis-Tris NuPAGE gels at 150V for 60-90 minutes in 1X MOPS buffer (see Appendix Section B for composition). Samples were then transferred onto a methanol-activated Immobilon-F PVDF membrane in 1X transfer buffer (see Appendix Section B for composition) using the Bio-Rad transfer system at 0.4A for two hours, or 0.2A overnight (approximately 16 hours). Transferred membranes were then blocked with 5% (w/v) skim milk powder in 1X tris-buffered saline (TBS - see Appendix Section B for composition) at room temperature for one hour and rinsed briefly in 1X TBS with tween (TTBS - see Appendix Section B for composition). Membranes were then incubated in primary antibody diluted in antibody solution (see Appendix Section B for composition) for two hours at room temperature or overnight at 4°C. A list of all immunoblot primary antibodies (and their dilutions) used in this study is provided in Table 4. Membranes were washed four times (15 minutes each time) in 1X TTBS, then incubated with IR Dye 800-conjugated fluorescent secondary antibody (goat anti-mouse) or Alexa Fluor 680-conjugated fluorescent secondary antibody (goat anti-rabbit) diluted 1:5000 in 5% (w/v) skim milk powder in 1X TTBS (containing 0.01% SDS (w/v)) for one hour, away from light. Membranes were washed as above, but away from light. Protein bands were visualised through detecting the fluorescent signal emitted from the secondary antibody conjugates using the LICOR Odyssey imaging system. The electronic images produced by the LICOR Odyssey imaging system software were saved as TIF files for subsequent densitometry analysis using Image J software.

2.2.5 Sphingolipid flux assays L6 cells were seeded in 12-well plates, differentiated, infected with adenovirus to overexpress LacZ and CerS isoforms and only treated with palmitate as described above. During the four hour serum-free period of palmitate treatment, myotubes were radiolabeled with [3H]serine (20µCi/mL) or [3H]sphingosine (0.8µCi/mL) made up in serum-free BSA-palmitate media. Immediately after, 10µL of labeling media was transferred from each well to 5mL of scintillation fluid in a plastic vial, for subsequent beta-counting. Myotubes were washed two times with ice-cold PBS and lysed as

70 described above. 5µL of sample was transferred to separate Eppendorf tubes and stored at -20°C for subsequent protein quantification, conducted as described above. Total lipid was extracted from the remaining lysate by the Folch method of lipid extraction. Briefly, 1mL of chloroform to methanol (2:1 v/v) solution was added to remaining lysate and vortexed twice with a 10-minute gap between vortexes. Samples were then centrifuged at 10 minutes at 13000 rpm, room temperature. Upper aqueous phase and middle protein phases were discarded and remaining organic phase was washed with 125µL MilliQ water for lipid re-re-extraction. Samples were vortexed and centrifuged as before. The upper aqueous phase was removed and the tube containing the remaining organic phase was placed on a 40°C heat block and completely dried down under nitrogen gas. The dried organic phase was reconstituted in 20µL methanol to chloroform (2:1 (v/v)) solution. Silica-coated thin layer chromatography (TLC) plates were activated by washing once with acetone and left to dry in a fume hood. 10µL of each reconstituted sample and 5µL of each authentic sphingolipid standard (sphingomyelin, sphingosine, glucosylceramide and ceramide) was collected with a glass syringe and spotted onto the TLC plate under hot air. The TLC plate was placed in a pre-equilibrated TLC tank containing filter paper soaked in TLC solvent system comprised of chloroform, methanol and water (65:25:4 (v/v/v)) and filled no higher than 1cm. The lipids on the TLC plate were subsequently separated via the solvent system. Once the solvent front was 2cm from the top of the plate, the TLC plate was removed from the tank, dried under a fume hood and iodinated by closely exposing the plate to the sublimed fumes generated from solid iodine which stained resolved lipids yellow. Once the standards could be visualized (present in much higher concentrations than samples), the TLC plate was then sprayed with En3Hance autoradiography enhancer spray and dried under a fume hood for 10 minutes. The spray and dry process was repeated a further three times and the plate rotated 90° after each interval. The TLC plate was then exposed to Kodak MR film in a cassette in a dark room to detect radiolabelled lipids. The cassette was stored horizontally for 6-12 days at -80°C to continue the exposure process. Radiolabelled sphingolipids from samples were then scraped from the TLC plate (thinly covered with H2O) using the standards as a guide, then transferred to plastic scintillation vials containing 5mL of scintillation fluid, vortexed and measured for radioactivity by beta scintillation counting. Radiolabelled

71 serine and sphingosine incorporation was corrected to total protein content determined as above.

2.2.6 GLUT4 translocation assay L6 myotubes endogenously express GLUT4 transporters [249]. However for the purposes of measuring GLUT4 translocation, we required pre-engineered L6 myoblasts stably expressing a HA-tagged GLUT4 receptor. These cells were prepared by the laboratory of Professor David James (Charles Perkins Centre, University of Sydney, NSW, Australia) via retroviral infection of L6 myoblasts with cDNA encoding HA- tagged GLUT4 as described in [250]. Advice regarding all technical and analytical aspects of this assay was provided by Dr Nolan Hoffman and Dr Daniel Fazakerley (Charles Perkins Centre - University of Sydney, NSW, Australia).

These pre-engineered L6 myoblasts were maintained and passaged in 10cm dishes containing MEM-alpha media comprised of 5.6 mM glucose, 10% (v/v) heat- inactivated FCS (high serum conditions) and 2mg/mL puromyocin (to select for cells expressing the HA-tagged GLUT4 transporter). Cells were seeded between passages 10- 20 in black walled-clear bottom 96-well plates (pre-coated with gelatin) and differentiated as described above (except differentiation media contained 2mg/mL puromyocin instead of antibiotic/antimycotic). Cells were infected with adenovirus containing cDNA encoding CerS isoforms and LacZ as above, but one day later than outlined in the general L6 cell treatment procedure. This was followed by treatment in the presence or absence of palmitate and stimulation with or without insulin as described above, except differentiation media supplemented with puromyocin was employed instead of usual differentiation media for L6 cells. Subsequent steps were as detailed in [250]. Simply, cells were washed three times with ice cold PBS (100µL/well), fixed on ice for 15 minutes with 3% paraformaldehyde (100µL/well, 16% stock diluted in PBS), incubated at room temperature for 30 mins and washed once with PBS. Glycine (100µL/well, 0.05M in PBS) was applied to cells for five minutes at room temperature after last wash to quench fixation and washed once as before. Cells were blocked for 20 minutes at room temperature with either 50µL/well of 5% normal swine serum (NSS) in PBS or 5% NSS with 0.1% (v/v) saponin (to permeabilise cells) in PBS. This was followed by incubation with 30µL/well of primary HA-tag or mouse immunoglobulin G1-κ (MOPC) antibody (both are anti-mouse antibodies diluted 72 1:1000 in 2% NSS in PBS) for one hour at room temperature. After incubation, cells were washed twice with PBS (200µL/well, five minutes each time), blocked with 5% NSS in PBS (50µL/well) for 20 minutes at room temperature, incubated for one hour with Alexa Fluor 488-conjugated fluorescent goat anti-mouse secondary antibody diluted 1:100 in 2% NSS in PBS (30µL/well, away from light), washed three times (five minutes each time) with PBS (100µL/well) and replaced with fresh PBS (100µL/well). Measurement of cell surface GLUT4 (GLUT4 transporter that has translocated to the cell membrane) and total GLUT4 (amount of GLUT4 transporter both inside the cell and on the cell membrane) from saponin-treated wells was conducted by measuring fluorescence from the bottom of the plate using a FLUOstar Omega plate reader under the following settings: 485nm excitation wavelength; 520 nm emission wavelength; gain set at 2000; highest intensity sample set to 90% of the gain. Background fluorescence was corrected for using measurements from saponin and non-saponin treated wells incubated with the MOPC primary antibody.

2.2.7 Sphingomyelinase inhibitor dose-response experiments The purpose of these experiments was to determine the dose of GW4869 and amitriptyline (pharmacological SMase inhibitors) that would effectively decrease insulin signal transduction in L6 myotubes without compromising cell viability. As such, we did not overexpress LacZ or CerS isoforms in these cells. L6 myoblasts were seeded in 12-well plates, differentiated, treated in the presence or absence of palmitate and stimulated with or without insulin as described above. All cells were treated with GW4869 in DMSO applied in 8.26µL volumes to 5mL of palmitate or non-palmitate (control) coupled 1X BSA (5% w/v) media to yield final concentrations 0, 5, 10 or

20µΜ. On separate plates, all cells were treated with amitriptyline in H2O applied in 12.5µL volumes to 5mL of palmitate or non-palmitate (control) coupled 1X BSA (5% w/v) media to yield final concentrations of 0, 10, 20 or 50µΜ . Vehicle dose for GW4869 and amitriptyline experiments was the 0 µM dose. GW4869 and amitriptyline treatment commenced at the beginning of the palmitate/control incubation period and finished after insulin-stimulation. Freshly prepared GW4869 and amitriptyline was applied when changing to serum-free media during this period. Cells were lysed, harvested for protein, measured for protein concentration and immunoblotted as described in sections 2.2.3 and 2.2.4. Protein samples were immunoblotted for 73 phosphorylated Akt (Ser473), all isoforms of total-Akt and β-actin (housekeeping protein). Phosphorylated-Akt and total Akt densitometry measurements were each normalised to β-actin prior to normalisation of phosphorylated-Akt with total-Akt.

2.3 Animal Studies Animal studies in this thesis were carried out with ethics approval from the Garvan/St Vincent’s Hospital Animal Ethics Committee.

2.3.1 Maintenance of mice Mice in this study were housed at the Garvan Institute Biological Testing Facility, (Garvan Institute of Medical Research, NSW, Australia) in accordance with facility procedures. Briefly, mice were housed under conventional conditions of 12-hour light/12-hour dark cycle with either littermates (a maximum of five mice per cage) or an ovaectomised companion mouse. Male and female C57Bl/6 mice were obtained from the Australian BioResources facility (Moss Vale, NSW, Australia), studied from 8-12 weeks of age and acclimatised to the Garvan Institute Biological Testing Facility for one week prior to all animal studies, during which they were given ad libitium access to water and standard chow diet (10.88 kJ/g; 8% fat, 21% protein and 71% carbohydrate). The weight and physical well-being of each mouse was checked and recorded once a week.

2.3.2 Dietary treatment To induce insulin resistance, mice were given ad libitium access to high-fat diet (HFD) prepared in-house (19.67 kJ/g; 45% fat, 20% protein and 35% carbohydrate - see Appendix Section B for composition) for up to 16 weeks. Chow-fed mice in this study were fed the above standard chow diet for one week. The HFD used in this study was designed as based on rodent diet D12451 (Research Diets - New Brunswick, NJ, USA).

74 2.3.3 Generation of adipose tissue specific protein kinase c epsilon (PKCε) knockout (KO) and control mouse lines The mice lines used for our adipose tissue PKCε characterisation study were generated as follows: a construct comprising of both exon 1 of the PKCε gene flanked by loxP sites i.e. “floxed” PKCε (PKCεf) and the neomycin resistance gene (Neor) flanked by FRT sites was injected into stem cells. Cells that incorporated this construct into their genome were selected for via neomycin treatment. Selected stem cells were injected into C57Bl/6 pseudopregnant mice to produce PKCεf/+, Neor mice on a C57Bl/6 genetic background. All procedures were performed by Ozgene (Bentley DC, Western Australia, Australia). Staff at the Garvan Biological Testing Facility performed the following steps in-house. The Neor gene was excised from these mice by breeding them with Flp deleter mice expressing the flippase recombinase on a C57Bl/6 background (a gift from Professor Robert Brink, Garvan Institute of Medical Research, NSW, Australia), an enzyme which cleaves at the FRT. The resulting PKCεf/+mice were bred by the Australian BioResources facility (Moss Vale, NSW) with heterozygous Adipoq- Cre transgenic mice on a C57Bl/6 background (a gift from Professor David James, now at the Charles Perkins Centre, University of Sydney, Sydney, Australia), which were first developed as described in [251]. Adipoq-Cre transgenic mice express cre- recombinase only in brown and white adipocytes as a result of cre-recombinase gene expression being placed under the control of a mouse adiponectin promoter locus (Adipoq), which is only activated in brown and white adipocytes. Resulting Adipoq- Cre/+,PKCεf/+ mice hence contain PKCε deleted at exon 1 exclusively in white and brown adipocytes through the action of cre-recombinase generated specifically in these cell types. Cleavage of this exon is presumed to prevent the protein expression of the remaining PKCε exons, as based on the absence of protein bands corresponding to PKCε in immunoblots of protein from adipose tissue and isolated adipocytes of these mice (Figure 4-1; Figure A-3 – see section 2.3.5 for methodology). Adipoq- Cre/+,PKCεf/+ mice were then bred to produce Adipoq-Cre/+,PKCef/PKCef (AdPKCεKO); WT,PKCεf/PKCεf (WT-Floxed); Adipoq-Cre/+,WT (WT-Cre) and WT,WT (WT) mice. The latter two genotypes were initially employed as additional controls in this study.

75 2.3.4 Genotyping Staff at the Australian BioResources facility collected tissue from two to three week old mice via tail or ear clip and isolated genomic DNA from these tissues. DNA samples were then sent to the Garvan Molecular Genetics facility (Garvan Institute of Medical Research, NSW, Australia) for genotyping by PCR. The PCR primer sequences used for PKCεf and Adipoq-Cre genotyping are listed below.

Table 5 PCR primer sequences used for PKCεf and Adipoq-Cre genotyping Forward Sequence Reverse Sequence PKCef GTCTTTCACGACGCTCCTATCG CCTATCACCACAAGCCCTTTTGA genotyping Adipoq-Cre CCGGTCGATGCAACGAGTGAT ACCAGAGTCATCCTTAGCGCC genotyping

2.3.5 Confirmation of PKCε deletion in adipocytes Adipocyte-specific PKCε deletion was confirmed in AdPKCεKO mice by immunoblotting protein from adipose tissue, skeletal muscle, liver and isolated adipocytes. Overall procedure is described as follows. WT, WT-Cre, and AdPKCεKO mice were sacrificed via cervical dislocation. Gonadal adipose tissue, quadriceps and liver were excised and tissues were homogenised in 2mL ice cold RIPA buffer and centrifuged at 13000 rpm for 20 minutes at 4°C. The resulting supernatant (liver and skeletal muscle) or infranatant (adipose tissue) was quantified for protein with the BCA protein assay kit, as per manufacturer’s instructions in combination with BSA standards dissolved in RIPA buffer prepared in-house. Adipocytes were isolated as per section 2.3.8, combined with three times the total adipocyte volume of ice-cold RIPA and homogenised by quick vortex. Due to addition of RIPA in proportion to total adipocyte volume of each sample, protein quantification of adipocytes was not performed. All tissues were then immunoblotted with antibody specific to PKCε. Immunoblot, including sample preparation, was performed as per section 2.2.4 except protein harvested from skeletal muscle, liver and adipose tissue was immunoblotted onto an Immobilon-P membrane incubated with sheep anti-mouse secondary antibody conjugated to horseradish peroxidase (HRP - 1:5000 dilution) and detected for protein using conversion of HRP with enhanced chemiluminescent substrates and subsequent 76 exposure with X-ray film. Tissue harvest, protein isolation and immunoblot of adipose tissue, skeletal muscle and liver were performed in-house by visiting scientist Dr Dale Hancock (University of Sydney, Sydney, NSW, Australia).

2.3.6 Intraperitoneal glucose tolerance test Glucose tolerance was measured after one week of chow feeding and 1, 8 and 16 weeks of high-fat feeding using the intraperitoneal glucose tolerance test (ipGTT). Mice were fasted for six hours prior to the commencement of the test, which was instigated by intraperitoneal injection of a glucose bolus. The doses used in the ipGTT for each feeding period are listed below and are expressed as grams per kilogram of total body weight.

2g/kg (obtained directly from glucose stock): 1-week chow/HFD 1g/kg (diluted in sterile saline): 8-week HFD 0.5g/kg (diluted in sterile saline): 16-week HFD

Lower doses were used for 8 and 16-week HFD ipGTTs to avoid obtaining glucose readings above the measurement range of the glucometer (maximum 33 mM), which may have been likely to occur from further elevations in basal blood glucose levels during longer term high-fat feeding. Approximately 3mm of the tail tip was excised with a scalpel for blood collection. Blood glucose measurements were obtained using an Accu-Chek glucometer at 0, 7.5 15, 22.5, 30, 45, 60 and 90 minutes after glucose injection. Blood samples (30µl each) were collected at 0, 15, 30 and 45 minutes after glucose injection, and gently mixed with 10µl of 18mM EDTA in sterile saline at 4°C to prevent coagulation. Plasma was obtained from blood samples via centrifugation at 13000 rpm for two minutes at 4°C and subsequent transfer of the supernatant to 96-well plates. Plasma samples were stored at -20°C until required.

77 2.3.7 Tissue harvest and blood collection from 16 week fat-fed mice Blood collection and tissue harvest from 16-week fat-fed mice was conducted at least three days after the ipGTT, in collaboration with Dr Carsten Schmitz-Peiffer (Garvan Institute of Medical Research, NSW, Australia). After approximately six hours fasting, mice were weighed, measured for blood glucose (as per section 2.3.6) and anaesthetised with isoflourane (4% induction, 1.5-2% maintenance). An abdominal incision was made, followed by cutting of the rib cage to expose the heart. Up to 500µl of blood was obtained by inserting a 23 gauge needle into the heart and gradually collecting it into an attached syringe. Blood was then transferred to Eppendorf tubes containing 18mM EDTA in saline equivalent to one third of measured blood volume. Plasma was then obtained from these samples as per section 2.3.6, transferred to fresh Eppendorf tubes and stored at -20°C until required.

The following tissues were then immediately excised with surgical scissors in the following order: gonadal adipose tissue (both sides), liver (all lobes), quadriceps muscle (both legs), inguinal subcutaneous adipose tissue (one side), retroperitoneal adipose tissue (surrounding both kidneys), mesenteric adipose tissue and interscapular brown adipose tissue (both depots). Interscapular incision was required to access the interscapular fat pad, which was excised from the mouse and dissected to obtain the brown adipose tissue depots housed within. Gonadal adipose tissue, liver and brown adipose tissue were quickly weighed, then freeze-clamped in liquid nitrogen. Remaining tissues were freeze-clamped immediately after excision. All snap frozen tissues were then stored at -80°C. Prior to snap freezing the gonadal adipose tissue of selected mice, a portion of the originally excised fat pad was cleaved and placed in ice-cold 4% paraformaldehyde for subsequent measurement of adipocyte size (see section 2.4.4).

2.3.8 Adipocyte isolation Mice were sacrificed via cervical dislocation, incised above the abdomen and harvested for gonadal adipose tissue, which was rinsed in sterile saline and briefly blotted dry. The tissue was transferred to a specimen jar and digested in 1-2mL of Krebs Ringer Buffer (KRB) freshly supplemented with 6.25mg/mL of collagenase D and 1.685 mg/ml of dispase (see Appendix Part B for composition) after mincing with surgical scissors. The tissue was shaken in a 37°C waterbath at 120 cycles/min for 20 minutes.

78 The digestion mix was filtered through a handmade funnel of nylon gauze on top of a 15ml conical tube and filtrate was allowed to settle for five minutes. The bottom aqueous layer (KRB digestion buffer) was removed and remaining layer of adipocytes was gently washed three times with 1ml KRB. Adipocytes were left to accumulate at the top of the KRB for five minutes after each wash. After the final wash, KRB was removed and remaining adipocytes gently washed with 1mL of PBS and left to settle for five minutes. The bottom aqueous layer (PBS) was removed and remaining adipocytes were transferred to Eppendorf tubes, snap frozen in liquid nitrogen and stored at -80°C.

2.3.9 Catheter insertion surgery for euglycaemic-hyperinsulinaemic clamp Surgical insertion of catheters into mice was required to facilitate the glucose and insulin infusions that constitute the euglycaemic-hyperinsulinaemic (E-H) clamp. This procedure was performed by Dr Amanda Brandon (Garvan Institute of Medical Research, NSW, Australia) as follows. Mice were anesthetised with isoflurane (4% induction, 1.5-2% maintenance). Catheters were then inserted into the left carotid artery and right jugular vein, followed by subcutaneous channeling of the free catheter ends which were exteriorized at the back of the neck and heat-sealed. Following surgery, mice were housed separately and monitored daily. Heparinised saline was used to flush catheters every one to two days to prevent catheter obstruction.

2.3.10 Euglycaemic-hyperinsulinaemic Clamp The E-H clamp is the gold standard for determining insulin sensitivity. Essentially, this involves infusing insulin to create constant, excess blood insulin levels and infusing glucose at a variable rate to maintain blood glucose concentrations to normal fasting levels. The E-H clamp and calculations to determine average glucose infusion rate (GIR), hepatic glucose output (HGO) rate, average rate of whole-body glucose disappearance (Rd) and tissue-specific glucose uptake (Rg’) rate were conducted by Dr Amanda Brandon, assisted by Eurwin Suryana (Garvan Institute of Medical Research, NSW, Australia). The Rg’ assay was conducted by Eurwin Suryana and myself (see section 2.4.5). This E-H clamp procedure was performed in accordance with protocol established by [75] and [76] and is described as follows: Conscious, one-week fat-fed mice (a separate cohort to mice assessed for glucose tolerance) containing surgically inserted catheters were subjected to E-H clamp four to seven days post surgery, 79 following an approximate five-hour fast. To minimalise stress, mice were unrestrained and untouched during the E-H clamp. 90 minutes prior to insulin infusion, a primed (5 µCi), continuous infusion (0.05 µCi/min) of [3,3H]-glucose was instigated. At 30, 20, 10 and 0 minutes before insulin infusion, samples were obtained for subsequent measurement of Rd and glucose, insulin and non-esterified FA (NEFA) levels (collected 30 and 0 minutes prior to insulin infusion). At time 0, the rate of [3,3H]-glucose infusion was elevated to 0.1 µCi/min and a primed (16 mU/Kg), continuous (4 mU/kg/min) infusion of insulin was administered. Simultaneously, glucose (25% v/v) was infused at a variable rate to reach a normal fasting blood glucose concentration of approximately 8mM. Once blood glucose concentrations were consistently maintained at 8mM (termed “clamped conditions”), four consecutive blood samples were obtained for subsequent measurement of insulin, NEFA and glucose turnover. A bolus of 2[14C]deoxyglucose (2[14C]DG, 13 µCi) was then injected as a bolus through the arterial catheter, followed by collection of blood samples at 2, 5, 10, 15, 20 and 30 min after 2[14C]DG injection, for subsequent measurement of Rg’. Animals were then sacrificed via cervical dislocation and organs removed, snap-frozen and stored as per section 2.3.7. Gonadal adipose tissue, liver, quadriceps muscle, gastrocnemius muscle, heart, inguinal subcutaneous adipose tissue, retroperitoneal adipose tissue, mesenteric adipose tissue and brown adipose tissue were harvested in the above order. All tissues except liver, retroperitoneal adipose tissue and mesenteric adipose tissue were employed to determine Rg’ via 2[14C]DG uptake assay (protocol detailed in section 2.4.5).

An Accu-check II glucometer was employed to measure blood and plasma glucose levels. Plasma insulin and NEFA concentrations were determined as per sections 2.4.1 and 2.4.2, respectively. Basal and clamp Rd was calculated using steady-state equations. Clamped HGO was obtained by subtracting the GIR from Rd.

80 2.4 Metabolic and Morphological Studies

2.4.1 Measurement of plasma insulin concentration Plasma samples collected during all HFD ipGTTs and E-H clamps were defrosted on ice. 5 µL of plasma sample then was measured for insulin concentration using the Ultra Sensitive Mouse Insulin ELISA Kit, in accordance with manufacturer’s instructions. The “low-range” assay conditions of the kit were employed. Final plasma insulin concentrations obtained from ipGTT samples were corrected for dilution due to the presence of saline/EDTA.

2.4.2 Measurement of plasma non-esterified fatty acid concentration Plasma samples collected during all HFD ipGTTs and E-H clamps were defrosted on ice. 5-10 µL of plasma sample was then used to determine NEFA concentration using the NEFA-C Kit, in accordance with manufacturer’s instructions. Assay endpoints for ipGTT samples were also measured for absorbance at 450nm to calculate the contribution of haem to the dye-specific absorbance (550nm). The haem absorbance was subtracted from the NEFA-specific absorbance and the net absorbance was used to subsequently calculate NEFA concentration, which was then corrected for dilution in saline/EDTA as described above.

2.4.3 Measurement of plasma cytokine concentrations Determination of plasma cytokine concentrations was performed in collaboration with Liam O’Reilly (Garvan Institute of Medical Research, NSW, Australia) using a protocol based on the fluorescence-assisted cell sorting technique. Plasma samples (25µL) obtained from 16-week fat-fed mice as per section 2.3.7 were defrosted on ice and prepared for the measurement of cytokines (interleukin 10 (IL-10), IL-6, interleukin 1- beta (IL-1β) and TNFα) using the Cytometric Bead Array (CBA) Master Buffer Kit, and kit-prescribed detection reagents specific to each cytokine, as per manufacturer instructions. Cytokine measurement was conducted by a BD FACSCantoII platform flow cytometer in accordance with CBA Master Buffer Kit instructions. Cytokine concentrations were automatically determined using the recommended FCAP Array software.

81 2.4.4 Measurement of adipocyte size Gonadal tissue harvested from 16-week fat-fed mice was stored in ice-cold 4% paraformaldehyde at 4°C for up to 24 hours, then transferred to 70% ethanol and stored for up to three months at 4°C. Tissues were then given to the Kinghorn Cancer Centre Histopathology Facility (Kinghorn Cancer Centre, NSW, Australia) for further preparation and staining. This involved step-wise dehydration and wax perfusion of tissues, embedment into wax blocks, sectioning into 4µm slices and staining with haematoxylin and eosin for the visualization of individual adipocytes. All steps were performed in accordance with standard facility procedures established for mouse tissue. Technical advice was provided by Dr Dorit Samocha-Bonet (Garvan Institute of Medical Research, NSW, Australia) and Dr Michael Swarbrick (Westmead Millenium Institute, NSW, Australia). Images of stained tissue sections were captured using a Leica DM4000 microscope at 40x objective. Adipocytes were counted from three fields of view, 150 cells/view. Images were then converted into black and white by Photoshop CS5 (Version 12.5 x64), where white areas were representative of adipocytes. White areas were measured and expressed as pixels by Image J 1.47v using the MRI Adipocyte tools plug-in and converted into µm2 using accompanying metadata information. Images were inspected to manually exclude blood vessels and damaged cells. Imaging, counting and data analysis performed by Dr Carsten Schmitz-Peiffer.

2.4.5 2[14C]deoxyglucose uptake assay (Rg’ assay) The administration of 2[14C]DG during the E-H clamp was employed to estimate the Rg’ for specific tissues harvested at the end of the E-H clamp. 2[14C]DG enters the cell in a similar manner to glucose and is phosphorylated to form 2[14C]DG-6-phosphate (2[14C]DGP). This phosphorylated form of 2[14C]DG cannot be metabolised and therefore accumulates in the tissue, which makes this molecule a useful tool for measuring Rg’. To determine the amount of 2[14C]DGP accumulated per unit mass of tissue, approximately 20-40mg of frozen tissue was homogenised in 1.5mL of distilled

H2O using a Polytron homogeniser and centrifuged at 13,000rpm for 10 minutes at 4°C to eliminate debris. To determine total [14C] counts, 400µL of supernatant was transferred to a glass 20mL scintillation vial containing 1.6mL of dH2O and 16mL of scintillation fluid and measured for radioactivity via beta scintillation counting. To determine 2[14C]DG counts, a further 400µL of supernatant was loaded onto an anion 82 exchange column contaning hydrated AG 1-X8 resin (acetate-form). During this step 2[14C]DGP binds to the resin, and 2[14C]DG (unbound) elutes through the column. The 14 column was then washed twice with 3mL of dH2O (to completely wash 2[ C]DG from the column), eluting completely into the collected 2[14C]DG after each wash. 2mL of total eluent was transferred to a glass 20mL scintillation vial containing 16mL of scintillation fluid and measured for 2[14C]DG via beta-scintillation counting. The total [14C] radioactivity measurement was subtracted from the 2[14C]DG radioactivity measurement to determine the amount of 2[14C]DGP radioactivity.

The following equation was then used to determine Rg’ [252]

14 14 Rg’ = (2[ C]DGPtissue/ AUC 2[ C]DGplasma)* [arterial glucose]

14 14 2[ C]DGPtissue = 2[ C]DGP radioactivity in a particular tissue (in dpm/g) 14 14 AUC 2[ C]DGplasma = area under the plasma 2[ C]DG disappearance curve (dpm/min/ml) – determined from blood samples obtained during the 2[14C]DG tracer period [arterial glucose] = average blood glucose (in mmol/l) during the 2[14C]DG tracer period

83 2.5 In Vivo Expression Analysis of Gonadal Adipose Tissue

2.5.1 mRNA expression analysis Total RNA was isolated through the following procedure, based on standard phenol- chloroform extraction technique. Approximately 150mg of adipose tissue was homogenised in 1.6mL of TRI reagent under a fume hood using a Polytron homogeniser, which was cleaned in between samples via stepwise rinsing in 2% absolve, sterile irrigation water (three times) and 70% ethanol. The homogenate was transferred to an Eppendorf tube and allowed to sit for five minutes, then centrifuged for 10 minutes at 12000g, 4°C. Under a fume hood, the top organic layer was discarded and 320µL of chloroform added to the remaining layer, then inverted, left to stand for 15 minutes and centrifuged for 15 minutes at 12000g, 4°C, forming three phases. This upper clear phase (containing RNA) was removed and combined with 800µL isopropanol, 32µL of RNase-free NaCl (5M) and 2.4µL of glycogen (5mg/ml) and inverted. The mixture was either left to stand at room temperature for 10 minutes or stored at -80°C for at least one hour (to facilitate glycogen-assisted RNA precipitation), then centrifuged for 10 minutes at 12000g, 4°C. The supernatant was discarded and the remaining pellet was washed with 1.6mL of 75% ethanol, vortexed for one minute and centrifuged for five minutes at 12000g, 4°C. Supernatant was carefully removed and remaining pellet dried under a fume hood whilst on a 37°C heat block. The pellet was resuspended in 100µL of diethylpyrocarbonate (DEPC) H2O and combined with 300µL of 100% molecular biology grade ethanol and 8µL of RNase-free NaCl (5M). Mixture was inverted, stored at -20°C for approximately one hour, vortexed and centrifuged at 12000g for 15 minutes at 4°C. Supernatant was gently removed and pellet dried as before. The pellet was then reconstituted in 50µL of DEPC H2O, vortexed for one minute, incubated for 10 minutes at 55°C, vortexed for another minute and stored at - 80°C. RNA concentration was measured using a Nanodrop ND-1000 Spectrophotometer to read the absorbance at 260nm. Technical advice for this procedure was provided by Dr Michael Swarbrick and Rebecca Stuart (Westmead Millenium Institute, NSW, Australia and Garvan Institute of Medical Research, NSW, Australia). Preparation of RT-PCR reaction reagents and subsequent RT-PCR reaction was carried out as per section 2.2.2 except the housekeeping gene was cyclophillin B and the final reaction RT-PCR reaction mixture was comprised of the following: cDNA (5ng), forward primer (0.4µM), reverse primer (0.4µM), UPL probe (0.2µM) and 84 master mix (1X). Technical advice for this procedure was provided by Dr Nancy Sue, Liam O’Reilly and Dr Jeng Yie Chan (Garvan Institute of Medical Research, NSW, Australia).

2.5.2 Protein expression analysis Protein expression analysis of various lipid metabolism proteins in adipose tissue was carried out by separately immunoblotting protein harvested from the aqueous and lipid- based components of adipose tissue. Separation of adipose tissue components and protein harvest was conducted as described below. Approximately 150mg of pre-ground adipose tissue was weighed and exact weight recorded. The tissue was combined with 300µL of ice-cold RIPA buffer (see Appendix Section B for composition), vortexed and homogenised on ice with a positive displacement pipette in two separate intervals, approximately 10 seconds each. Additional RIPA buffer was added to make the total RIPA buffer volume equivalent to five times the exact recorded weight of the adipose tissue sample. This was followed by vortexing and centrifugation at 12000g, 0°C for 15 minutes to form three separate components: solid lipid, aqueous infranatant and pellet. The top layer of solid lipid was transferred to a fresh, Eppendorf tube using a metal scoop (both pre-cooled in liquid nitrogen) and weight recorded. RIPA buffer was then added to the solid lipid sample at a volume equivalent to five times the recorded weight and vortexed for approximately 10 seconds. The isolated lipid sample was centrifuged at 12000g, 0°C for 15 minutes. The buffer below the fat cake was gently removed and replaced with an equivalent volume of fresh RIPA, and the sample homogenised. The aqueous infranatant and pellet remaining from the first centrifugation step were also re- centrifuged (12000g, 0°C for 15 minutes), and 500µL of infranatant was transferred to a fresh Eppendorf tube. The lipid and infranatant samples were then immunoblotted as per section 2.2.4. except that protein quantification was not performed beforehand. Every sample densitometry measurement on a gel was normalised to a housekeeping protein (non-muscle myosin IIA (heavy chain)). Every sample densitometry measurement on a gel was then normalised by the average densitometry measurement of bands from the WT-Floxed samples (already normalised for housekeeping protein) on each gel.

85 2.6 Statistical Analysis All data presented in this thesis are expressed as ± standard error of the mean (SEM). Statistical analysis including Student’s t-test, one-way analysis of variance (ANOVA) and two-way ANOVA with post hoc testing was conducted using GraphPad Prism software (Version 6). Statistical analysis involving three-way ANOVA was conducted by Dr Carsten Schmitz-Peiffer using StataSE v9.2 software.

86 CHAPTER 3 MECHANISMS OF CERAMIDE SYNTHASE-INDUCED ALTERATIONS IN INSULIN ACTION

87 3.1 Introduction An early and crucial contributor to the development of T2D is the insulin resistance of tissues important for glucose homeostasis. One such tissue is skeletal muscle, which is a key tissue involved in blood glucose disposal [253]. In the insulin resistant state, skeletal muscle displays a diminished ability to respond to insulin stimulation, resulting in poor regulation of whole-body glucose metabolism [40].

There is a strong association between the accumulation of bioactive lipids in peripheral, non adipose tissues and the generation of insulin resistance [254]. A lipid metabolite considered to be a leading candidate in the induction of skeletal muscle insulin resistance is ceramide, the central member of the sphingolipid class of lipids [64, 255]. Ceramide is known to induce insulin resistance at multiple levels, including the inhibition of insulin signalling at the level of Akt [37, 81].

Currently, there are three known routes of ceramide generation: (i) the de novo synthesis pathway (ii) the salvage recycling pathway and (iii) SM hydrolysis (for more details on these pathways, see section 1.8). For the purposes of this chapter, it is important to reiterate that the de novo and salvage pathways both require the CerS family of enzymes for ceramide generation, whilst the SM hydrolysis pathway does not [94, 256]. The FA substrate specificity and tissue-specific expression levels unique to each CerS isoform accounts for the diverse array of ceramides present in different tissues under different conditions [1]. Studies have shown elevated levels of distinct ceramide species in the skeletal muscle of obese, insulin resistant mice and humans [26, 44-46]. In parallel, there is early evidence demonstrating an association between altered expression of individual CerS isoforms and the generation of insulin resistance [45, 105]. Collectively, this suggests that certain ceramide species are more detrimental toward insulin action than others. Identification of the ceramide species and CerS isoforms responsible for these effects however, was until very recently lacking. To address this issue, our laboratory sought to determine the role of individual CerS isoforms and ceramides during lipid-induced insulin resistance in L6 skeletal muscle cells.

Though each overexpressed CerS isoform altered L6 myotube ceramide profile in accordance with their individual lipid substrate preferences (see Table 1), no isoforms

88 inhibited insulin signalling or glycogen synthesis in the absence of palmitate. The inhibitory effects of palmitate on insulin action were also not potentiated after CerS overexpression. CerS1 overexpression was found to be associated with improvements in insulin signalling and glycogen synthesis in the absence of palmitate. Under these same conditions, CerS6 overexpression and knockdown was shown to affect these processes in a reciprocal manner [131]. Our findings suggested that ceramides generated from specific CerS isoforms, most notably CerS6, in skeletal muscle imposed beneficial actions on skeletal muscle insulin action. In parallel, we proposed that inhibitory ceramides generated by palmitate treatment were produced through the SMase pathway via palmitate-induced activation of SMase (see section 1.7.2 for more details). Currently, it is not clear how CerS overexpression improved insulin action in the absence of palmitate and furthermore, why this manipulation did not potentiate the effects of palmitate as expected. This knowledge gap is further compounded by the possibility that modulation of CerS expression may have caused compensatory changes in sphingolipid metabolism. Furthermore, assessing the effects of CerS manipulation upon other arms of insulin action beyond glycogen synthesis is required to confirm our novel findings.

In this chapter, we aimed to elucidate the potential mechanisms responsible for the improvement in insulin action after CerS overexpression by investigating (i) whether CerS overexpression caused compensatory modulations in the mRNA expression of genes encoding CerS isoforms and other sphingolipid metabolism proteins (ii) whether CerS overexpression affected the flux through the de novo and salvage pathways of ceramide synthesis. In addition, we aimed to further explore the unexpected findings from our previous work by assessing the effect of CerS overexpression upon insulin stimulated glucose uptake, another key element of skeletal muscle insulin action.

89 3.2 Results

3.2.1 CerS isoform overexpression has minimal effects on the mRNA levels of CerS isoforms and other proteins involved in sphingolipid metabolism Compensatory changes in the mRNA expression of CerS isoforms and/or other sphingolipid metabolism enzymes could explain why CerS overexpression did not potentiate the effects of palmitate in our previous work. To assess this possibility, L6 myotubes overexpressing CerS isoforms (or LacZ control) were treated with or without palmitate and harvested for mRNA. This was converted into cDNA and quantified by RT-PCR. Treatments and procedures were carried out as per sections 2.2.1.2 and 2.2.2. The following genes were quantified for mRNA expression: CerS1, CerS2, CerS4, CerS5, CerS6, Sptlc1, Sptlc2, Ormdl2 and Ormdl3. Sptlc1 and Sptlc2 encode the heterodimeric subunits of SPT which, as mentioned earlier, catalyses the first step of de novo ceramide synthesis [257] (please refer to Section 1.8.1 for more details). Ormdl2 and Ormdl3 encode different isoforms of the ORM protein, a negative regulator of de novo sphingolipid synthesis [119]. Please refer to Figure 1-5 for a diagram representing the function of all proteins investigated. Only endogenous CerS mRNA expression could be measured in these experiments because the CerS primers used were designed to target rat-based CerS sequences (L6 is a rat skeletal muscle cell line), whereas the CerS sequences in adenoviral-housed constructs (constituting overexpressed CerS) were murine based and therefore not detected by our primers. Overall, CerS overexpression caused minimal effects on the mRNA expression of endogenous CerS isoforms, Sptlc and Ormdl genes in the presence and absence of palmitate (Figure 3-1, Figure 3-2). Exceptions to this included the significant down-regulation of CerS1 and minor up- regulation of CerS5 following CerS1 overexpression under all treatment conditions. CerS1 and CerS5 mRNA expression was also significantly increased during palmitate treatment in comparison with non-palmitate treated samples (Figure 3-1).

90 # CONTROL PALMITATE * *

Figure 3-1: Effect of CerS overexpression on mRNA levels of endogenous CerS isoforms. L6 myotubes overexpressing CerS isoforms were treated with (black bars) or without (white bars) 0.375mM palmitate. mRNA was converted into cDNA and quantified by RT-PCR. Results are expressed as fold change over control (non-palmitate treated, LacZ-overexpressing cells) and are the means of three independent experiments performed in quadruplicate. ANOVA: *P<0.05 for effect of CerS1 vs. LacZ on endogenous CerS1 and CerS5 mRNA expression, #P<0.05 for palmitate effect on CerS1 expression, ##P<0.001 for palmitate effect on CerS5 expression. Figure adapted from [131].

91 CONTROL PALMITATE

Figure 3-2 Effect of CerS overexpression on mRNA levels of Sptlc and Ormdl isoforms. Details as per Figure 3-1. Figure adapted from [131].

92 3.2.2 Effects of CerS overexpression on ceramide synthesis pathway flux Flux modulation of ceramide synthesis pathways following CerS isoform overexpression could also explain why these isoforms failed to exacerbate the inhibitory effects of palmitate. Incorporation of radiolabelled [3H]serine and [3H]sphingosine into SM, sphingosine, glucosylceramide and ceramide was measured to determine if the flux through the de novo or salvage pathways of sphingolipid production were respectively altered by CerS overexpression. Treatments and procedures carried out as per sections 2.2.1.2 and 2.2.5

3.2.2.1 CerS isoform overexpression does not alter de novo pathway flux Signals corresponding to [3H]serine-labelled SM were visible on the autoradiograph of the TLC plate (not shown) whilst [3H]serine-labelled sphingosine, glucosylceramide and ceramide signals were below the level of detection. As a consequence, only SM was excised from the TLC plate and quantified for radioactivity. Beta scintillation measurements showed no changes in [3H]serine-labelled SM levels after CerS isoform overexpression compared to LacZ control (Figure 3-3). This demonstrates that flux through the de novo synthesis pathway is not modulated in response to CerS overexpression in the presence of palmitate.

PALMITATE

Figure 3-3 Radioactivity measurements of [3H]serine-labelled sphingomyelin. L6 myotubes overexpressing CerS isoforms or LacZ control were treated with 0.375mM palmitate. [3H]serine tracer was applied to cells at commencement of the four hour serum-free period of 0.375mM palmitate treatment to track flux through the de novo pathway of ceramide synthesis. Total lipid was extracted and individual lipid species were resolved by thin layer chromatography (TLC). Radiolabelled sphingolipid species were excised from the TLC plate and measured for radioactivity with beta scintillation counting. Results are expressed as percentage change compared to [3H]serine-labelled sphingomyelin in control cells and are means from three independent experiments performed in duplicate. Adapted from [131].

93 3.2.2.2 CerS1, 4 and 6 overexpression increases salvage pathway flux The representative autoradiograph in Figure 3-4 illustrates the successful detection of all sphingolipids radiolabelled with [3H]sphingosine. Measurement of [3H] sphingosine incorporation by beta-scintillation counting demonstrated that CerS1, CerS4 and CerS6 overexpression caused significant increases in [3H]sphingosine incorporation into ceramide and glycosylceramide and trending increases in [3H]sphingosine incorporation into sphingomyelin (Figure 3-5). This indicates that overexpression of these CerS isoforms up-regulates salvage pathway flux in the presence of palmitate.

Figure 3-4 Representative autoradiograph of [3H]sphingosine incorporation into sphingolipids. L6 myotubes overexpressing CerS isoforms or LacZ control were treated with 0.375mM palmitate. [3H]sphingosine tracer was applied to cells at commencement of the four hour serum-free period of 0.375mM palmitate treatment to track flux through the salvage pathway of ceramide synthesis. Total lipid was extracted and individual lipid species were resolved by TLC. Radiolabelled sphingolipid species were detected by autoradiography. Autoradiograph representative of three independent experiments performed in duplicate. Adapted from [131].

94 PALMITATE

** ** ** * * *

Figure 3-5: Radioactivity measurements of [3H]sphingosine-labelled sphingolipids. Experimental details as per Figure 3-3 except [3H]sphingosine tracer was instead applied to cells at the commencement of the four hour serum-free period of 0.375mM palmitate treatment. Results are expressed as percentage change compared to [3H]sphingosine-labelled ceramide in control cells and are means from three independent experiments performed in duplicate. ANOVA: Effect of CerS1, 4, and 6 on [3H]sphingosine incorporation into ceramide, **P<0.01; glucosylceramide, *P<0.05. Adapted from [131].

95 3.2.3 CerS1 overexpression increases GLUT4 transporter translocation to the plasma membrane Previous work from our group had demonstrated that overexpression of individual CerS isoforms either improved or did not alter insulin signalling and glycogen synthesis [131]. To further investigate these unexpected findings, we examined the effects of CerS overexpression on other important aspects of insulin action, such as glucose uptake. Insulin signalling facilitates glucose uptake by mediating the translocation of the glucose transporter, GLUT4, from intracellular vesicles to the plasma membrane [258, 259]. To assess the effects of CerS isoform overexpression on glucose uptake, GLUT4 translocation was measured in L6 cells overexpressing HA-tagged GLUT4 in addition to CerS isoforms using an immunofluorescent technique developed by the laboratory of Prof. David James (now at the Charles Perkins Centre – University of Sydney, NSW, Australia), as detailed in [250]. These cells were treated in the absence and presence of palmitate, with and without insulin stimulation. All treatments and procedures are further outlined in sections 2.2.1.2 and 2.2.6. CerS1 overexpression significantly increased surface GLUT4 levels compared to LacZ control during all treatment conditions (Figure 3-6 (a)). Overexpression of other CerS isoforms however, did not alter surface GLUT4 levels. Surprisingly, palmitate slightly increased basal surface GLUT4 levels in all samples (Figure 3-6 (a)). As expected, there was minimal variance in total GLUT4 levels across all samples (Figure 3-6 (b)). Insulin stimulation caused a 2-2.5 fold increase in surface GLUT4 levels – a typical increase for this assay (Figure 3-6 (c)). Overall, these results have provided further evidence that ceramides generated via isolated CerS isoform overexpression either improve or do not impede insulin action.

96 BASAL a INSULIN

Figure 3-6: Surface and total GLUT4 transporter levels following CerS isoform overexpression. Individual CerS isoforms were overexpressed in L6 myotubes stably expressing HA- tagged GLUT4. Cells were then treated with or without 0.375mM palmitate, stimulated with or without insulin and then fixed. To determine surface HA-tagged GLUT4 levels, cells were probed with anti-HA antibody, incubated with fluorescently labelled secondary antibody and measured for fluorescence. Levels of total HA-tagged GLUT4 were obtained as above, in permeabilised cells. Results shown in a) are expressed as fold change compared to LacZ-expressing cells not treated with 0.375mM palmitate b) are expressed as fold change relative to the mean of LacZ-expressing cells treated with and without insulin c) are expressed as fold change relative non insulin-stimulated sample. Results are means of three independent experiments executed in triplicate. ANOVA: **P<0.001 for effect of CerS1 vs. LacZ control under all treatment conditions. Figure adapted from [131].

97 3.3 Discussion Identification of the CerS enzymes most implicated in the initiation and propagation of skeletal muscle insulin resistance may enable the development of more targeted therapeutics for this condition. It was important therefore, to elucidate the roles of individual CerS isoforms and ceramide species. Findings reported by this laboratory (discussed in section 3.1) were unexpected as they showed that though overexpression CerS isoforms produced the expected alterations in individual ceramide species, it had either improved or had no effects upon insulin action in cultured L6 skeletal muscle cells [131]. It was therefore necessary to extend our findings and rule out possible compensatory mechanisms.

Here, we first investigated whether overexpression of specific CerS isoforms caused compensatory down-regulation in the mRNA expression of other CerS isoforms. mRNA expression of endogenous CerS isoforms was found to be mostly unaltered in response to CerS overexpression. An absence of expected increases in the mRNA levels of each CerS isoform upon its overexpression was most likely due to the failure of the CerS primers (designed from rat sequences) to bind to the overexpressed CerS mRNA generated from the CerS cDNA (designed from mouse sequences) packaged in the adenovirus that provides this excess CerS cDNA. This result is in contrast to reports using other models which demonstrate that the global knockout or targeted knockdown of individual CerS isoforms was accompanied by an elevation in the mRNA levels of non-targeted CerS isoforms [121-123]. Overall, our findings eliminate the possibility that compensatory changes in the mRNA levels of non-targeted CerS isoforms were responsible for beneficial effects on insulin action following CerS overexpression. An exception to this was the down-regulation of endogenous CerS1 mRNA following CerS1 overexpression in the presence and absence of palmitate. This feedback mechanism may be required to alleviate potential cellular stress caused by the observed elevation in C18:0 ceramide levels following CerS1 overexpression in our cell model [131]. The role of CerS1 and C18:0 ceramide in compromising cell viability has been well documented. In one study, C18:0 ceramide levels were decreased in human head and neck squamous cell carcinoma (HNSCC) samples. Reconstitution of C18:0 ceramide via CerS1 overexpression in the corresponding cell line however, significantly inhibited cell growth [95]. Concurrently, siRNA knockdown of CerS1 in the same

98 HNSCC cell line prevented cell death by approximately 50% in response to chemotherapeutic agents [96]. Furthermore, up-regulation of CerS1 mRNA following chemotherapy was associated with decreased proliferation of chronic myeloid leukaemia cells [97]. All studies propose apoptosis induction to be at least partially responsible for these effects. Cleaved caspase-3 is a key mediator of the lipid-induced apoptosis cascade in skeletal muscle [70]. Data from our laboratory has shown an increase in cleaved caspase-3 levels following CerS1 overexpression in palmitate- treated L6 myotubes (Barbara Diakanastasis and Carsten Schmitz-Peiffer, unpublished). However, we saw no associated change in other contributing factors towards apoptosis such ER stress or mitochondrial dysfunction (Barbara Diakanastasis and Carsten Schmitz-Peiffer, unpublished). Despite associations between skeletal muscle insulin resistance and palmitate-induced apoptosis [70, 99, 100], our results show that CerS1 overexpression increases insulin cell signaling, glycogen synthesis [131] and GLUT4 translocation. It is proposed therefore, that the accompanying increase in C18:0 ceramide may be improving insulin sensitivity and imposing cell stress simultaneously via independent, non-connected mechanisms.

CerS isoform overexpression did not significantly alter mRNA levels of the de novo synthesis pathway regulatory genes, Sptlc1, Sptlc2, Ormdl2 and Ormdl3. This further indicates that CerS isoform overexpression was not eliciting its positive effects upon insulin action in our cell model through counter-regulation of the de novo pathway, at least at the mRNA level. In contrast, it has been shown that inhibition of CerS activity decreases the mRNA expression of Sptlc1 and Sptlc2 [260]. The mRNA levels of Sptlc1, Sptlc2, Ormdl2 and Ormdl3 in our study were not altered in LacZ-infected control L6 cells after palmitate treatment. This finding contrasts with work by Verma and colleagues, which show a significant palmitate-induced increase in Sptlc1 mRNA in C2C12 skeletal muscle cells [73]. It must be noted however, that the palmitate dose used was twice the concentration of that used here.

It would beneficial to assess effects of CerS overexpression upon the protein expression of endogenous CerS isoforms, SPT1, SPT2, ORMDL2 and ORMDL3. Currently, this is not possible by immunoblotting for most CerS isoforms because of the poor performance of commercially available antibodies (not shown). Assessment of counter- regulatory changes in protein expression however, may be achieved through mass 99 spectrometry. Nonetheless, there is evidence of CerS-mediated counter-regulation of these proteins. For instance, targeted knockdown of individual CerS isoforms in MCF-7 human breast adenocarcinoma cells elevated the protein levels of non-targeted CerS isoforms [113]. In addition, the simultaneous pharmacological inhibition of all CerS isoforms in HEK293 cells and mouse liver resulted in a decrease in total ORM protein levels and an increase in SPT1 and SPT2 proteins levels, respectively [260, 261]. The activity of these proteins, as well as CerS isoforms, is also regulated via post- translational modifications [262, 263] [127]. Hence, assessment of post-translational effects on CerS isoforms, as well as on SPT1, SPT2, ORMDL2 and ORMDL3 following CerS isoform overexpression is also warranted.

It was important to analyse the relative contributions of each sphingolipid synthesis pathway when investigating the effects of CerS overexpression or knockdown, as this may help us determine which pathway/s are more implicated in generating more or less metabolically beneficial ceramides. Currently, there is limited literature that sufficiently evaluates the effect of CerS modulation on the flux of individual sphingolipid synthesis pathways in skeletal muscle [111].

In this study, we assessed whether flux through the de novo and salvage pathways was altered by CerS overexpression by tracking the fate of radiolabelled tracers specific to each pathway within palmitate-treated L6 myotubes. Overexpression of CerS1, 4 and 6 increased flux through the salvage pathway, whilst de novo pathway flux was not altered. This indicates that sphingolipids generated specifically through the salvage pathway may be eliciting beneficial effects upon muscle cell insulin action in response to CerS overexpression. This contrasts with data from [111] showing that sphingolipids were predominantly made in C212 mouse myoblasts via the de novo pathway only. It should be noted however, that the cells in that study were not lipid-treated and had no engineered alterations in sphingolipid metabolism enzyme expression. Repeating our flux assays in the absence of palmitate may confirm that increased salvage pathway flux was also responsible for the improved insulin action seen after CerS overexpression in non-palmitate treated cells (in the event that increases in salvage pathway flux occur).

Currently, it is unclear how these salvage pathway-specific ceramides may have caused the observed improvements in insulin action as shown in Figure 3-6 and in our previous 100 work [131]. Previous findings from our lab have shown that overexpressed CerS isoforms in the presence and absence of palmitate treatment did not alter mitochondrial function or the protein expression of ER stress markers (Barbara Diakanastasis and Carsten Schmitz-Peiffer, unpublished). Analysing the effects of CerS overexpression toward other elements linked to insulin resistance, such as reactive oxygen species, inflammatory markers and lipid raft composition, may provide more mechanistic insight into this phenomenon.

[3H]Serine incorporation into ceramide, glucosylceramide and sphingosine could not be quantified. This result was surprising, considering palmitate positively regulates the de novo ceramide synthesis pathway [94]. Our investigation into the effects of CerS overexpression with respect to de novo pathway flux could be improved. To attain a more accurate scope of de novo pathway flux, this experiment could be repeated using either a higher concentration of [3H]Serine tracer or an alternative radiolabelled sphingolipid tracer exclusive to the de novo pathway (such as radiolabelled sphinganine). The sole detection of [3H]serine-labelled SM, one of the final sphingolipid products of the de novo pathway, may indicate that flux through this pathway occurs more quickly than flux through the salvage pathway. As such, repeating the [3H]Serine tracer incorporation assay using shortened periods of tracer incubation may reveal alternations in de novo pathway flux in response to CerS overexpression. Repeating these experiments with non-palmitate treated cells may also uncover additional flux alternations through the de novo pathway, which may also account for the beneficial effects on insulin action previously seen in non-palmitate treated L6 cells, after CerS overexpression [131].

It is generally agreed that ceramide induces skeletal muscle insulin resistance through the deactivation of Akt. However, there exists an increasingly accepted view that examining changes in insulin signalling pathway physiological endpoints such as GLUT4 translocation (to the plasma membrane) and glycogen synthesis, rather than changes in canonical insulin signalling per se, is a more useful assessment of insulin action [40, 254]. Whilst CerS overexpression did not demonstrate any negative effects upon glycogen synthesis in our previous study [131], we could not assume that the same would apply for GLUT4 translocation. Therefore, it was essential to assess the effects of

101 CerS overexpression on GLUT4 translocation in order to evaluate the overall impact of CerS overexpression upon insulin action in muscle cells.

Significantly elevated GLUT4 levels at the cell surface following CerS1 overexpression concurred together with the beneficial effects of this isoform upon insulin signalling and glycogen synthesis [131]. The minimal variance in total GLUT4 levels across all samples confirms that this increase was not due to parallel increases in total GLUT4. An increase in surface GLUT4 levels however, was not observed in CerS6-overexpressing cells despite previously observed reciprocal effects upon insulin action following the knockdown and overexpression of this isoform [131]. The absence of noticeable decreases in insulin-stimulated GLUT4 translocation following CerS overexpression reinforces our previous findings showing that CerS overexpression did not impede insulin action nor potentiate the negative effects of palmitate. This lies in contrast with a study which demonstrated that simultaneous pharmacological inhibition of all CerS isoforms in rat brown adipocytes increased GLUT4 protein expression and elevated glucose uptake [264]. The disparity between these findings suggests that CerS isoforms may have different effects amongst various tissues with respect to insulin action.

Palmitate treatment (375 µM, 20 hours) did not produce major decreases in insulin- stimulated GLUT4 translocation, despite our previous work showing that this FA caused large reductions in insulin-stimulated glycogen synthesis [131]. Furthermore, palmitate-treated cells under basal conditions exhibited higher levels of surface GLUT4 than control cells. Similarly, [247] showed that palmitate treatment (200 µM, 24 hours) in L6 cells increased basal surface GLUT4 levels and displayed similar levels of insulin-stimulated surface GLUT4 to non-palmitate samples. These studies suggest that a higher palmitate dose is required to inhibit GLUT4 translocation. In contrast, a 50% reduction in insulin-stimulated surface GLUT4 levels was observed in L6 cells after treatment with a lower concentration of palmitate (150µM) for a similar duration of time [265]. Reasons for the absence of major decreases in GLUT4 translocation in our palmitate-treated cell model are unclear. However, our data overall further supports an association between CerS expression and beneficial skeletal muscle insulin action.

102 The opposing effects on insulin action observed between palmitate and non palmitate- treated CerS-overexpressed muscle cell samples in our previous study [131] may be due to the potential inhibitory nature of ceramides produced independently from CerS via the palmitate-activated SMase pathway. This notion is supported by previous literature demonstrating increases in Akt phosphorylation after sphingomyelinase inhibition [73, 266, 267]. This possibility was investigated in preliminary experiments which involved administering L6 myotubes with increasing doses of an acidic or neutral sphingomyelinase inhibitor (amitriptyline or GW4869) in the presence or absence of palmitate, with or without insulin stimulation (see sections 2.2.1.2 and 2.2.7 for experimental details). Neither sphingomyelinase inhibitor elicited increases in phosphorylated Akt levels in palmitate-treated cells (Figure A-2). Though these findings do not support our hypothesis, these experiments will require further optimisation before final conclusions can be made.

The findings in this chapter have provided some insight into how CerS overexpression may have improved insulin action in the absence of palmitate and how it also failed to potentiate the inhibitory actions of palmitate. Firstly we have demonstrated that the overexpression of individual CerS isoforms overall did not elicit compensatory changes in the endogenous mRNA expression of CerS isoforms and other genes encoding proteins involved in ceramide synthesis. However, the observed ability of overexpressed CerS1 to counter-regulate endogenous CerS1 expression highlights a potential anti-apoptotic role for this enzyme. It was also shown that flux through the salvage pathway of ceramide synthesis was increased following the overexpression of multiple CerS isoforms, placing this pathway as a potential source for the production of metabolically beneficial ceramides. Though the underlying mechanisms behind our findings have not been elucidated, the data does suggest that individual ceramide species may be imposing their effects upon insulin action, and possibly apoptosis, by simultaneously acting upon separate pathways that do not influence one another. Finally, we showed that GLUT4 translocation to the membrane is either enhanced or unchanged in association with CerS overexpression. Consequentially, this confirms that insulin action is either improved or not impinged when CerS isoforms are overexpressed in muscle cells. This result, in conjunction with our previous work shown in [131] has established that the effects of ceramides upon insulin action are not solely

103 inhibitory and are in fact much more complex than previously proposed by the general consensus within the literature.

104 CHAPTER 4 PHENOTYPIC CHARACTERISATION OF ADIPOSE TISSUE-SPECIFIC PROTEIN KINASE C EPSILON KNOCKOUT MICE

105 4.1 Introduction The insulin resistance of key tissues involved in maintaining glucose homeostasis and whole-body insulin sensitivity is strongly linked to the accumulation of bioactive lipid intermediates from dietary fatty acid (FA) oversupply. These intermediates operate as second messengers that modulate a variety of key signalling and metabolic pathways. The ceramide family of sphingolipids has been widely considered as examples of these pathogenic lipid moieties [37]. However, we have confirmed in the previous chapter that particular ceramide species can enhance insulin action and some aspects of insulin sensitivity within cultured skeletal muscle cells [131].

Another well-described bioactive lipid intermediate implicated in the pathogenesis of insulin resistance is DAG. Elevated intracellular DAG concentrations following dietary lipid oversupply are associated with the chronic activation of PKC isoforms. The persistent DAG-mediated activation of nPKC enzymes has been associated with the generation of insulin resistance [141]. PKC epsilon (PKCε) is a nPKC isoform that is extensively linked to the generation of lipid-induced hepatic insulin resistance. The majority of studies suggest that PKCε mediates hepatic insulin resistance through direct actions at the liver [135, 136, 140]. During one of these studies, our laboratory demonstrated that global PKCεKO mice were protected against glucose intolerance induced after a one-week HFD [140]. This suggested that an improvement in hepatic insulin sensitivity had occurred within these mice. Further investigation with liver- specific PKCεKO mice generated from our laboratory however, revealed no improvement in glucose tolerance under the same dietary conditions (manuscript in preparation). In addition, a separate cohort of liver-specific PKCεKO mice did not exhibit improved whole-body insulin sensitivity during the E-H clamp, the gold standard technique for determining insulin sensitivity (manuscript in preparation). This suggested that PKCε was indirectly decreasing hepatic insulin sensitivity from a different location.

A multitude of animal and human studies have shown that hepatic glucose production is significantly affected by FA supply from adipose tissue to the liver [52-54] [55, 268, 269]. Furthermore, our laboratory had discovered that plasma NEFA concentrations in global PKCεKO mice during the ipGTT had decreased to a greater extent than in WT

106 controls, most likely in response to insulin secretion caused by the glucose challenge (manuscript in preparation). This led us to hypothesise that PKCε caused an increase in hepatic glucose production through enhancing the FA supply from adipose tissue to the liver. In this chapter, we aimed to examine the contribution of this cross talk toward the pathogenesis and maintenance of the insulin resistant state by (i) Characterising the effects of PKCε deletion specifically in adipose tissue on glucose tolerance and NEFA release (ii) Assessing the impact of this deletion on whole-body and tissue-specific insulin sensitivity. Through this study, we expected to confirm that AdPKCε modulates the adipose tissue-liver signalling axis and emphasise this role of AdPKCε toward the pathogenesis of hepatic and subsequently, whole body insulin resistance. If achieved, this would essentially overturn the current theory that PKCε elicits hepatic insulin resistance directly at the liver and furthermore, would direct the design of therapeutics targeted toward disconnecting this cross-talk.

107 4.2 Results

4.2.1 PKCε is deleted in primary adipocytes only Confirmation of the deletion of PKCε exclusively in adipose tissue was required prior to subsequent phenotypic characterisation. AdPKCεKO and WT mice were sacrificed and tissues were harvested. Protein lysates were generated from adipose tissue and other insulin-sensitive tissues including skeletal muscle and liver. Protein lysates were also prepared from adipocytes isolated from the adipose tissue of AdPKCεKO and WT mice. All protein lysates were comprised of soluble and membrane-associated proteins (i.e. total cellular protein) and were immunoblotted for the PKCε protein. Protein harvest, adipocyte isolation and immunoblot were carried out as per section 2.3.5. WT and AdPKCεKO mice exhibited approximately the same levels of total PKCε protein in liver and skeletal muscle. Total PKCε protein level was greatly reduced in the adipose tissue of the AdPKCεKO mouse, when compared to the WT control. However, a residual total PKCε band remained in the AdPKCεKO sample. Virtual disappearance of this residual band was observed in AdPKCεKO primary adipocytes (Figure 4-1). This immunoblot data confirms that we have successfully deleted PKCε exclusively within the adipocytes of adipose tissue.

WT KO WT KO

Adipose tissue Liver

Primary Adipocytes Skeletal Muscle

Figure 4-1 Tissue-specific protein expression of total PKCε in WT and AdPKCεKO mice. Protein lysates (comprised of soluble and membrane-associated proteins) harvested from liver, skeletal muscle, adipose tissue and isolated primary adipocytes of AdPKCεKO and WT mice were immunoblotted with an antibody targeted to PKCε.

108 4.2.2 Minimal body-weight difference between WT and AdPKCεKO mice throughout the study The body weights of mice used in the AdPKCεKO in-vivo characterisation study were recorded weekly over a 16-week HFD period (males and females) and a one-week chow-feeding period (males only) in order to assess effects of PKCε deletion. Results from studies involving female one-week chow-fed mice, as well as studies involving male and female mice fed chow diet for up to 16 weeks, were not included in this thesis as they were ongoing at the time of writing. No significant differences in body weight were observed between the three genotypes within each sex, either during the high fat or chow feeding period (Figure 4-2). Average body weights across all genotypes, sexes and diets significantly increased each week in comparison to the weight from the previous week of feeding. Furthermore, the average body weight of male mice fed HFD for one week was significantly higher than the average body weight of male age- matched chow-fed controls during this period (Figure 4-2). These results indicate that the deletion of PKCε in adipose tissue does not impact upon body weight, and therefore most likely upon other related factors such as appetite and energy expenditure, although they were not directly assessed in this study.

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Body Weight (g) 20 5 0 0 1 0 1 0 1 0 1 Week Chow HFD Chow HFD WT - Floxed AdPKCεKO Figure 4-2 Weekly body weights of mice employed in the AdPKCεKO in vivo characterisation study. Body weights were recorded on a weekly basis at the commencement and during high- fat or chow feeding. a) Male (m) weekly body weights during 16-week HFD. ANOVA: p<0.001 effect of feeding over time. b) Female (f) weekly body weights during 16 week HFD. ANOVA: p<0.001 effect of feeding over time. c) Body weight comparison of one-week chow-fed and high-fat fed males. ANOVA: p<0.01 effect of feeding over time, p<0.05 HFD vs. chow. AdPKCεKO HFD (orange line/column) n(m)=14, n(f)=17; WT-Cre HFD (purple line) n(m)=10, n(f)=8; WT-Flox HFD (black line/column) n(m)=20, n(f)=20; AdPKCεKO chow (green column) n(m)=9; WT-Flox chow (blue column) n(m)=7. 110 4.2.3 Minimal differences in fasting metabolic parameters between genotypes Blood glucose concentrations were measured prior to conducting the intraperitoneal glucose tolerance test (ipGTT). Fasting plasma NEFA and fasting plasma insulin levels were measured in plasma from blood samples collected immediately prior to each ipGTT as per sections 2.4.1 and 2.4.2. All fasting metabolic parameters increased in each genotype across both sexes with increased duration of high-fat feeding (Table 6 and Table 7).

Table 6 Fasting metabolic parameters for male mice before ipGTT Body weights and blood glucose levels were measured in male mice after a six-hour fast prior to ipGTT (following one-week chow or 1, 8 or 16 week HFD). Plasma NEFA and plasma insulin were measured from plasma obtained from blood samples collected immediately prior to each ipGTT.

Metabolic Parameter WT-Cre WT-Floxed AdPKCεKO Body Weight (g) 1 week chow N/A 23.7±0.80 24.2±1.14 1 week HFD 24.2±0.46 25.8±0.63 25.3±0.56 8 week HFD 30.5±0.72 33.1±0.82 31.6±0.79 16 week HFD 36.7±1.08 39.9±1.28 38.4±1.71

Blood Glucose (mM) 1 week chow N/A 9.7±0.34 9.7±0.42 1 week HFD 11.9±0.68 11.4±0.39 10.6±0.63 8 week HFD 11.7±0.43 12.9±0.5 12±0.41 16 week HFD 11.7±0.62 12.7±0.53 11.8±0.47

Plasma NEFA (mM) 1 week HFD 0.28±0.02 0.27±0.05 0.23±0.02 8 week HFD 0.47±0.03 0.41±0.04 0.44±0.04 16 week HFD 0.60±0.04 0.64±0.04 0.54±0.06

Plasma Insulin (ng/ml) 1 week HFD 1.47±0.14 1.45±0.12 1.27±0.1 8 week HFD 1.41±0.14 2.15±0.14 2.02±0.18 16 week HFD 2.43±0.29 3.97±0.52 3.77±0.4

111 Table 7 Fasting metabolic parameters for female mice before ipGTT All fasting metabolic parameters were measured in female mice as outlined in Table 6.

Metabolic Parameter WT-Cre WT-Floxed AdPKCεKO Body Weight (g) 1 week HFD 20.3±0.98 20.3±0.48 20.1±0.38 8 week HFD 26.5±1.6 26.6±0.86 26.4±0.76 16 week HFD 31.7±2.69 33.7±1.22 31.8±1.04

Blood Glucose (mM) 1 week HFD 10.2±0.43 10.3±0.34 9.9±0.28 8 week HFD 11.2±0.46 11.4±0.49 10.6±0.29 16 week HFD 10.7±0.74 13.3±0.38 10.6±0.45

Plasma NEFA (mM) 1 week HFD 0.3±0.03 0.27±0.02 0.24±0.02 8 week HFD 0.37±0.05 0.38±0.03 0.34±0.04 16 week HFD 0.66±0.05 0.54±0.05 0.5±0.04

Plasma Insulin (ng/ml) 1 week HFD 0.92±0.2 0.82±0.06 0.73±0.07 8 week HFD 1.13±0.27 0.97±0.08 1.11±0.08 16 week HFD 1.51±0.33 1.67±0.17 1.47±0.16

112 4.2.4 PKCε ablation in adipose tissue is associated with improved glucose tolerance and further suppression of plasma NEFA after a one-week HFD To establish whether AdPKCε was associated with the development of insulin resistance via cross talk with the liver, we initially assessed glucose tolerance in male and female AdPKCεKO mice following a one-week HFD, during which only adipose tissue and liver acquire insulin resistance [25, 26]. Glucose tolerance, a measure of the body’s ability to remove glucose from the circulation, was examined in these mice via the ipGTT (see section 2.3.6), which is the most widely used assessment of glucose tolerance in research involving rodent models [270]. Glucose tolerance was also assessed in age-matched chow-fed AdPKCεKO mice to evaluate whether the beneficial effects of AdPKCεKO on glucose tolerance were solely manifested during high-fat feeding.

There was no difference in glucose excursions between male chow-fed AdPKCεKO mice and WT-Floxed controls (Figure 4-3(a)). Both fat-fed male and female AdPKCεKO mice however, displayed significantly lower glucose excursions than fat- fed WT-Floxed and WT-Cre controls during the ipGTT (Figure 4-3(a), Figure 4-4(a)). Incremental area under the curve (iAUC) measurements of glucose excursions further illustrate these decreases (Figure 4-3(b), Figure 4-4(b)). Glucose excursions in male chow-fed mice were overall much lower than those displayed by age-matched fat-fed mice. This data indicates, at least in male mice, that PKCε deletion in adipose tissue is associated with improved glucose tolerance in the context of high-fat feeding only.

NEFA concentrations decreased in all mice after commencement of the ipGTT. Both male and female AdPKCεKO fat-fed mice however, displayed further decreases in plasma NEFA concentrations in comparison to WT controls during the ipGTT (Figure 4-3(c), Figure 4-4(c)). This indicates that PKCε deletion in adipose tissue is associated with a greater suppression of NEFA release from adipose tissue upon acute glucose challenge.

All mice demonstrated slight increases in plasma insulin concentration during the ipGTT, with the exception of male WT-Cre mice (which showed a very slight decrease in plasma insulin levels). Generally however, there appeared to be no major differences 113 in plasma insulin concentrations between AdPKCεKO and WT controls during the ipGTT after one-week HFD (Figure 4-3 (d), Figure 4-4 (d)), indicating a lack of compensatory insulin release.

Overall, these results collectively suggest that PKCε deletion in adipose tissue is associated with improved glucose tolerance and enhanced suppression of plasma NEFA concentrations in the absence of compensatory insulin secretion during ipGTTs performed after a one-week HFD.

114 a 35 b 30 * 1500 ### 25 ##

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0 15 30 45 Time (min) Figure 4-3 Blood glucose, plasma insulin and plasma NEFA excursions during ipGTT in male mice after one-week high-fat or chow feeding.

Male AdPKCεKO, WT-Floxed and WT-Cre mice were fed chow or HFD for one week. ipGTT was initiated with 2g/kg glucose bolus after a six-hour fast. a) Blood glucose excursions during ipGTT. ANOVA: p<0.001 AdPKCεKO vs. all WT controls (HFD), p<0.001 HFD vs chow. b) iAUC during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls (HFD), p<0.01 AdPKCεKO (HFD) vs. AdPKCεKO (chow), p<0.001 WT- Floxed (HFD) vs. WT-Floxed (chow). c) Plasma NEFA excursions during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. WT-Cre. d) Plasma insulin excursions during ipGTT. c) and d) measured in plasma from blood samples taken after ipGTT commencement. AdPKCεKO HFD n=11 (orange line); WT-Cre HFD n=11 (purple line); WT-Flox HFD n=19 (black line); AdPKCεKO chow n=9 (green line); WT-Flox chow n=7 (blue line). 115 ** a 35 b 1500 30 1000 25

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Figure 4-4 Blood glucose, plasma insulin and plasma NEFA excursions during ipGTT in female mice after one-week high-fat feeding.

Female AdPKCεKO, WT-Floxed and WT-Cre mice were fed HFD for one week. ipGTT initiated with 2g/kg glucose bolus after a six-hour fast. a) Blood glucose excursions during ipGTT. ANOVA: p<0.001 AdPKCεKO vs. all WT controls. b) iAUC during ipGTT. ANOVA: p<0.01 AdPKCεKO vs. all WT controls. c) Plasma NEFA excursions during ipGTT. ANOVA p<0.05 AdPKCεKO vs. all WT controls. d) Plasma insulin excursions during ipGTT. c) and d) measured in plasma from blood samples taken after ipGTT commencement. AdPKCεKO n=14 (orange line); WT-Cre n=7 (purple line); WT-Flox n=18 (black line). 116 4.2.5 PKCε ablation in adipose tissue is linked with increased glucose tolerance and further suppression of plasma NEFA after 8 and 16-week HFD The improvement in glucose tolerance displayed by AdPKCεKO mice after a one-week HFD drove us to investigate if this protective effect could continue with longer term high-fat feeding, which is more representative of the obese state. The same cohorts of male and female mice employed in the one-week HFD ipGTT study were assessed for glucose tolerance following 8 and 16 weeks of high-fat feeding as per section 2.3.6.

Both male and female AdPKCεKO mice displayed significantly decreased glucose excursions during the ipGTT as compared to WT controls following 8 and 16 weeks of HFD (Figure 4-5(a), Figure 4-6(a), Figure 4-7(a), Figure 4-8(a)). These decreased glucose excursions are further illustrated by accompanying iAUC measurements (Figure 4-5(b), Figure 4-6(b), Figure 4-7(b), Figure 4-8(b)). These findings collectively suggest that PKCε ablation in adipose tissue is associated with improved glucose tolerance after 8 and 16-week HFD.

Plasma NEFA concentrations during the 8 and 16-week HFD ipGTTs were further decreased in male and female AdPKCεKO mice compared with WT-controls, suggesting that deletion of PKCε in adipose tissue was linked to further suppression of plasma NEFA during a longer-term HFD (Figure 4-5(c), Figure 4-6(c), Figure 4-7(c), Figure 4-8(c)).

Plasma insulin levels during ipGTTs after 8 and 16 week HFD were unchanged between genotypes in female mice, and between AdPKCεKO and WT-Floxed male mice. Male WT-Cre mice however, displayed noticeably lower plasma insulin levels than the other genotypes before and during both ipGTTs (Figure 4-5(d), Figure 4-6(d), Figure 4-7(d), Figure 4-8(d)). The plasma insulin profiles, with the exclusion of the male WT-Cre outlier, in turn indicated that AdPKCεKO mice displayed no additional compensatory increases in insulin release during the 8 and 16-week HFD ipGTTs.

Altogether, our findings suggest that PKCε deletion in adipose tissue is associated with improved glucose tolerance and enhanced suppression of plasma NEFA in the absence of compensatory insulin secretion during ipGTTs performed after 8 and 16-week HFD.

117 a 30 b 25 1000 20 800 15 600

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Figure 4-5 Blood glucose, plasma NEFA and plasma insulin excursions during ipGTT in male mice after eight-week high-fat feeding.

Male AdPKCεKO, WT-Floxed and WT-Cre mice were fed HFD for eight weeks. ipGTT initiated with 1g/kg glucose bolus after a six-hour fast. a) Blood glucose excursions during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls. b) iAUC during ipGTT c) Plasma NEFA excursions during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls. d) Plasma insulin excursions during ipGTT. c) and d) measured in plasma from blood samples taken after ipGTT commencement. AdPKCεKO n=12 (orange line); WT-Cre n=13 (purple line); WT-Flox n=20 (black line).

118

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0.1 (mM) PlasmaNEFA 0.0 0 15 30 45 Time (min) d 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 (ng/ml) PlasmaInsulin 0.0 0 15 30 45 Time (min) Figure 4-6 Blood glucose, plasma NEFA and plasma insulin excursions during ipGTT in female mice following eight-week high-fat feeding.

Female AdPKCεKO, WT-Floxed and WT-Cre mice were fed HFD for eight weeks. ipGTT initiated with 1g/kg glucose bolus after a six-hour fast. a) Blood glucose excursions during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls. b) iAUC of a) during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. WT-Floxed c) Plasma NEFA excursions during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls. d) Plasma insulin excursions during ipGTT. c) and d) measured in plasma from blood samples taken after ipGTT commencement. AdPKCεKO n=16 (orange line); WT-Cre n=6 (purple line); WT-Flox n=20 (black line).

119 a 30 b

20 ** 1000

15 800

10 600

Glucose (mM) Glucose 400 5 iAUC (mM.min) iAUC 200

0 0 0 15 30 45 60 75 90 WT - Cre WT - Floxed AdPKCεKO Time (min)

c 0.8 0.7 0.6

0.5 0.4 0.3 0.2

Plasma NEFA (mM) PlasmaNEFA 0.1 0.0 0 15 30 45 Time (min)

d 5.0 4.0

3.0

2.0

1.0

Plasma Insulin (ng/ml) (ng/ml) PlasmaInsulin 0.0 0 15 30 45 Time (min) Figure 4-7 Blood glucose, plasma NEFA and plasma insulin excursions during ipGTT in male mice after 16-week high-fat feeding.

Male AdPKCεKO, WT-Floxed and WT-Cre mice were fed HFD for 16 weeks. ipGTT initiated with 0.5g/kg glucose bolus after a 6-hour fast. a) Blood glucose excursions during ipGTT. ANOVA: p<0.001 AdPKCεKO vs. all WT controls. b) iAUC during ipGTT. ANOVA: p<0.01 AdPKCεKO vs. all WT controls. c) Plasma NEFA excursions during ipGTT. ANOVA: p<0.001 AdPKCεKO vs. all WT controls. d) Plasma insulin excursions during ipGTT. c) and d) measured in plasma from blood samples taken after ipGTT commencement. AdPKCεKO n=12 (orange line); WT-Cre n=11 (purple line); WT-Flox n=17 (black line). 120 a 30 b 25 1000 * 20 800

15 600

10 400 Glucose (mM) Glucose iAUC (mM.min) iAUC 200 5 0 WT - Cre WT - Floxed AdPKCεKO 0 0 15 30 45 60 75 90 Time (min)

c 0.8 0.7 0.6 0.5

0.4 0.3

0.2 Plasma NEFA (mM) PlasmaNEFA 0.1 0.0 0 15 30 45

d Time (min) 5.0

4.0

3.0

2.0

1.0 Plasma Insulin (ng/ml) (ng/ml) PlasmaInsulin 0.0 0 15 30 45 Time (min) Figure 4-8 Blood glucose, plasma insulin and plasma NEFA excursions during ipGTT in female mice after 16-week high-fat feeding.

Female AdPKCεKO, WT-Floxed and WT-Cre mice were fed HFD for 16 weeks. ipGTT initiated with 0.5g/kg glucose bolus after a six-hour fast. a) Blood glucose excursions during ipGTT. ANOVA: p<0.001 AdPKCεKO vs. all WT controls. b) iAUC during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls. c) Plasma NEFA excursions during ipGTT. ANOVA: p<0.05 AdPKCεKO vs. all WT controls. d) Plasma insulin excursions during ipGTT. c) and d) measured in plasma from blood samples taken after ipGTT commencement. AdPKCεKO n=16 (orange line); WT-Cre n=7 (purple line); WT-Flox n=16 (black line). 121 4.2.6 PKCε ablation in adipose tissue is associated with increased hepatic and whole-body insulin sensitivity in female mice following a one-week HFD The improvement in whole-body glucose tolerance and accompanying decrease in plasma NEFA exhibited by AdPKCεKO mice during the ipGTTs following short and long-term HFD suggested enhanced whole-body insulin sensitivity in these mice, especially in the context of unchanged plasma insulin concentrations. To confirm this enhancement, we performed the euglycaemic-hyperinsulinaemic (E-H) clamp technique, the gold standard for determining insulin sensitivity [75, 76]. During the E-H clamp, insulin is infused at a fixed rate to create constantly elevated blood insulin levels whilst glucose is simultaneously infused at a variable rate to maintain blood glucose concentrations to normal fasting levels (euglycaemia – “clamped” conditions) i.e. 8mM. A separate cohort of male and female AdPKCεKO and WT-Floxed mice were assessed via the E-H clamp following a one-week HFD. Since WT-Cre mice presented very similar results to WT-Floxed mice during the ipGTTs, they were no longer used as controls in this study. Details on E-H clamp technique and associated analytical methods are outlined in sections 2.3.9, 2.3.10 and 2.4.5.

No significant difference in fasting body weight, basal or clamped blood glucose concentration, plasma NEFA and plasma insulin concentrations was observed between genotypes across both sexes. Elevated insulin levels during the clamp period caused significant decreases in the blood glucose concentration of males, as well as significant decreases in plasma NEFA for both genders in comparison to basal measurements as expected (Table 8).

Male mice did not display a difference in glucose infusion rate (GIR) between genotypes (Figure 4-9(a)). As expected, the rate of whole body glucose disappearance (Rd) increased under clamped conditions for both male genotypes. This parameter however, did not change between male genotypes under both basal and clamped conditions (Figure 4-9(b)). Hepatic glucose output (HGO) decreased during clamped conditions across both male genotypes as expected. A non-significant decrease in the HGO of male AdPKCεKO mice was observed in comparison to male WT-Floxed mice (Figure 4-9(c)). Male mice presented no difference in the suppression of NEFA release between genotypes (Figure 4-9(d)). Overall, there was no difference in the rate of

122 glucose uptake (Rg’) of each tissue between male WT-Floxed and AdPKCεKO mice, with the exception of a non significant decrease in the gastrocnemius muscle (gas) Rg’ of AdPKCεKO mice as compared to WT controls (Figure 4-9(e)).

Female AdPKCεKO mice presented a significantly higher GIR than WT-Floxed mice (Figure 4-10(a)). In other words, glucose needed to be infused into female AdPKCεKO mice at a significantly higher rate than WT controls to attain euglycaemia. This indicates that female AdPKCεKO have a higher whole-body insulin sensitivity than WT-controls. As expected, Rd increased under clamped conditions for both female genotypes, but did not change between genotypes under both basal and clamped conditions (Figure 4-10(b)). HGO decreased for both female genotypes during clamped conditions, as expected. Female AdPKCεKO mice presented a significantly lower suppression of HGO during the clamp, as compared to WT-Floxed mice (Figure 4- 10(c)). This indicates that female AdPKCεKO mice have increased hepatic insulin sensitivity. It is likely that this increase mediated the observed improvement in whole body insulin sensitivity. Female AdPKCεKO mice displayed a tendency toward a greater suppression of NEFA release into plasma under clamped conditions, compared to WT controls (Figure 4-10(d)). Female AdPKCεKO mice also showed an approximate two-fold increase in Rg’ within subcutaneous (SubQ) adipose tissue and quadriceps (quad) muscle and a 30% increase in brown adipose tissue (BAT) Rg’ compared to WT controls. These differences however, were not significant. No noticeable differences in the Rg’ of gonadal fat (gonadal) and gastrocnemius (gas) muscle were observed between female genotypes (Figure 4-10(e)). This suggests that the insulin sensitivity of SubQ, BAT and quad in female mice has increased after PKCε deletion in adipose tissue.

Overall, the results from our E-H clamp experiments suggest that male AdPKCεKO mice do not have an increase in whole body insulin sensitivity that can easily be detected after a one-week HFD. However, the decrease in HGO displayed by male AdPKCεKO mice does provide promising suggestion of a subtle increase in whole- body insulin sensitivity. Our results also suggest that female AdPKCεKO mice have increased whole body insulin sensitivity after a one-week HFD. This could be due to

123 increased hepatic insulin sensitivity, which may be mediated by the observed decrease in plasma NEFA concentrations in these mice during clamped conditions.

Table 8 Parameters before and during euglycaemic-hyperinsulinaemic clamp Body weights in male and female mice were measured following an approx. five-hour fast prior to euglycaemic-hyperinsulinaemic clamp (after one-week HFD). Blood glucose was measured during the basal and clamp period. Plasma NEFA and plasma insulin were measured from plasma obtained from blood samples collected during the basal and clamp period. ANOVA: **P<0.01 for effect of clamp vs. basal (males only); ###P<0.001 for effect of clamp vs. basal (males and females). Males: AdPKCeKO n=9, WT-Floxed n=6. Females: AdPKCeKO n=6, WT-Floxed n=7.

Male Female

Metabolic Parameter WT-Floxed AdPKCεKO WT-Floxed AdPKCεKO

Fasting Body Weight (g) 24.7±0.7 23.6±0.8 21.9±0.8 20.1±0.8 Blood Glucose (mM) Basal 8.8±0.4 8.1±0.4 8.0±0.4 7.0±0.4 ** Clamp 7.8±0.2 7.6±0.2 7.7±0.2 7.6±0.3

Plasma NEFA (mM) Basal 1.03±0.08 1.00±0.08 0.97±0.12 0.98±0.08 ### Clamp 0.54±0.11 0.42±0.06 0.40±0.04 0.31±0.04

Plasma Insulin (µU/ml) Basal 29.06±4.56 29.13±2.65 23.13±5.07 18.61±4.59 ### Clamp 91.87±6.19 88.42±5.26 88.79±8.61 87.62±8.49

124 a 60 b 60 ***

40 40 WT-Flox Ad-PKCεKO

(mg/min/kg) 20 Whole Body Glucose Glucose Body Whole Glucose Infusion Rate Rate Infusion Glucose 0 Disappearance (mg/kg/min) 0 WT-Flox Ad-PKCεKO Basal Clamp Males Males c 30 *** d 80

60 20 P=0.17 40

10 (mg/kg/min) Plasma NEFA Plasma

% Suppression 20 Hepatic Glucose Output 0 0 Basal Clamp WT-Flox Ad-PKCεKO Males Males

e 500 400 300

200 30

mol/min/100g) 20 µ ( Uptake Glucose 10 0 SubQ Gonadal BAT Quad Gas Males Figure 4-9 Parameters of whole-body and tissue-specific insulin sensitivity measured in male mice before and during euglycaemic-hyperinsulinaemic clamp.

Male AdPKCεKO and WT-Floxed mice were fed HFD for one week and then fasted for approx. five-hours prior to euglycaemic-hyperinsulinaemic (E-H) clamp. a) Glucose infusion rate b) Whole-body glucose disappearance rate c) Hepatic glucose output rate d) Percentage suppression of plasma NEFA after commencement of E-H clamp e) Rate of tissue-specific glucose uptake during E-H clamp. Basal = 3[H]glucose tracer infusion period prior to commencement of E-H clamp. Clamp = 3[H] glucose tracer infusion period during E-H clamp. SubQ: subcutaneous adipose tissue; Gonadal: gonadal adipose tissue; BAT: brown adipose tissue; Quad: quadriceps muscle; Gas: gastrocnemius muscle. ANOVA: ***P<0.001 for effect of E-H clamp vs. basal. AdPKCeKO n=9, WT-Floxed n=6.

125 a b 60 ## 60 ***

WT-Flox 40 40 Ad-PKCεKO

20 20 (mg/min/kg) Whole Body Glucose Glucose Body Whole Glucose Infusion Rate Rate Infusion Glucose

0 Disappearance (mg/kg/min) 0 WT-Flox Ad-PKCεKO Basal Clamp Females Females

c *** d 30 p = 0.08 # 80 20 60 10 40 0 (mg/kg/min) Plasma NEFA Plasma

% Suppression 20 -10 Hepatic Glucose Output -20 0 Basal Clamp WT-Flox Ad-PKCεKO Females Females

e 500 400 300

200 30

mol/min/100g) 20

µ Glucose Uptake Uptake Glucose ( 10

0 SubQ Gonadal BAT Quad Gas

Females Figure 4-10 Parameters of whole-body and tissue-specific insulin sensitivity measured in female mice before and during euglycaemic-hyperinsulinaemic clamp.

Female AdPKCεKO and WT-Floxed mice were fed HFD for one week and then fasted for approx. five-hours prior to euglycaemic-hyperinsulinaemic E-H clamp. a) Glucose infusion rate b) Whole-body glucose disappearance rate c) Hepatic glucose output rate d) Percentage suppression of plasma NEFA after commencement of the (E-H) clamp e) Rate of tissue-specific glucose uptake. Basal = 3[H] glucose tracer infusion period prior to commencement of the E-H clamp. Clamp = 3[H] glucose tracer infusion period during the E-H clamp. SubQ: subcutaneous adipose tissue; Gonadal: gonadal adipose tissue; BAT: brown adipose tissue; Quad: quadriceps muscle; Gas: gastrocnemius muscle. ANOVA: *** P<0.001 for effect of E-H clamp vs. basal; #P<0.05, ##P<0.01 AdPKCεKO vs. WT-Floxed. AdPKCeKO n=6, WT-Floxed n=7.

126 4.2.7 PKCε ablation in adipose tissue is associated with smaller adipocyte size Larger adipocytes are generally considered to be more insulin resistant and have a greater potential to cause ectopic insulin resistance than smaller adipocytes [24]. We therefore compared adipocyte area between AdPKCεKO and WT mice. This was achieved through imaging cross sections of gonadal adipose tissue harvested from 16 week high-fat fed male and female AdPKCεKO and WT-Floxed mice (a subset of mice from the ipGTT cohorts in this study), and measuring the area of adipocytes using automated software (see section 2.4.4 for more details). The representative images from male AdPKCεKO and WT-Floxed mice in Figure 4-11 show that a greater proportion of AdPKCεKO adipocytes are smaller in area than WT-Floxed controls. Female mice also displayed a similar relationship between genotypes (images not shown). The adipocyte area histograms mirror these trends by showing that male and female AdPKCεKO mice have a significantly higher percentage of adipocytes within a smaller area range (3000- 7000 µm2), as well a significantly lower percentage of adipocytes in a larger area range (11000-18000 µm2) in comparison to WT-Floxed mice (Figure 4-12(a)-(b)). These results indicate that PKCε deletion in adipose tissue is associated with smaller adipocyte size (which marks higher insulin sensitivity) during long-term high fat feeding.

WT-Floxed AdPKCεKO

100 µm

Figure 4-11 Adipocytes from WT-Floxed and AdPKCεKO mice fed 16-week HFD. Gonadal adipose tissue from male (m) and female (f) mice fed HFD for 16 weeks were preserved, sectioned and stained to visualise adipocytes microscopically (40x objective). Adipocytes in this figure (from male mice) are representative of three fields of view per mouse and reflect trends in adipocyte size between WT-Floxed and AdPKCεKO mice for both genders. Males: AdPKCεKO n=7, WT-Floxed n=7. Females: AdPKCεKO n=5, WT-Floxed n=5. 127 *** *** a 14

12 WT-floxed

10 Ad-PKCεKO

6 4 Distribution (%) Distribution 2 0

Adipocyte Area - Males (µm2)

b ** * 14 12

8 6

4 Distribution (%) Distribution 2 0

Adipocyte Area - Females (µm2)

Figure 4-12 Adipocyte diameters of mice after 16-week HFD. Gonadal adipose tissue depots from male and female mice fed HFD for 16 weeks were chemically preserved, sectioned and stained to visualise adipocytes microscopically (40x objective). Areas of individual adipocytes for each sample were measured electronically from three separate fields of view (approx. 150 cells per view) and segregated into different groups based on area (groups shown on x-axis). Final number of adipocytes for area group of each sample was the average of three fields of view. a) Male adipocyte area distribution. b) Female adipocyte area distribution. Green bars = WT-Floxed, Red bars = AdPKCεKO (males n=7, females n=5 – both genotypes). ANOVA: * P<0.05, **P<0.01, ***P<0.001 AdPKCεKO vs WT-Floxed.

128 4.3 Discussion The body of work in this chapter has extensively examined the contribution of adipose tissue-specific PKCε with respect to the development, maintenance and propagation of the insulin resistant state especially via potential cross-talk to the liver. We have shown that both male and female AdPKCεKO mice display improved glucose tolerance during short and long term high-fat feeding. This was associated with further suppression of NEFA release and the absence of compensatory increases in insulin, in turn suggesting that PKCε deletion in adipose tissue mediated an elevation in whole body insulin sensitivity during various stages of high-fat feeding. Further investigation via E-H clamp studies confirmed that the increased glucose tolerance at least in female AdPKCεKO mice after a one-week HFD was due to increased whole-body insulin sensitivity driven by an improvement in hepatic insulin sensitivity. Our findings further suggest that this is mediated through further decreases in plasma NEFA levels in female AdPKCεKO mice. Adipocyte area measurements from 16-week high-fat fed mice showed that a significantly higher percentage of adipocytes from AdPKCεKO mice were in the smaller area range, whilst a significantly lower percentage were in the higher area range in comparison with WT adipocytes. This is consistent with the view that PKCε deletion in adipose tissue plays a protective role against insulin resistance.

We firstly confirmed the presence of adipocyte specific PKCε deletion in AdPKCεKO mice. Although a residual PKCε band was seen in the adipose tissue immunoblot from the AdPKCεKO mouse, it was presumed to be PKCε from the stromal vascular fraction within adipose tissue (or other non-parenchymal elements such as resident macrophages). The virtual disappearance of this band in the primary adipocyte immunoblot of AdPKCεKO mice confirmed this. Further evidence of adipose tissue- specific PKCε deletion is shown in the appendix (Figure A-3).

As expected, fasting ipGTT metabolic parameters including body weight, plasma NEFA concentration and plasma insulin concentration increased in all groups of mice, with increasing HFD duration. The minimal difference in fasting body weight between genotypes prior to the ipGTTs and E-H clamps suggests that PKCε deletion in adipose tissue is not influencing factors such as energy expenditure and appetite.

129 In this study, we aimed to investigate whether PKCε could alter lipid flux from adipose tissue to liver, as this is considered to be the dominant contributing factor toward the development of insulin resistance during short-term high-fat feeding i.e. one-week HFD [26, 205]. This was important to investigate, as it is essential to gain a better understanding of how changes in FA flux drive healthier individuals toward an insulin resistant state.

The ipGTT was employed in this study to initially ascertain whether PKCε deletion in adipose tissue was eliciting a protective response to HFD-induced glucose intolerance. The significantly lower glucose excursions displayed by chow-fed mice in comparison to age-matched mice fed HFD for one week occurred as expected, due to the fact that one-week HFD is sufficient to induce insulin resistance in the liver and adipose tissue of rodents [25, 26, 137]. The improved glucose tolerance seen in AdPKCεKO mice after one-week HFD is consistent with findings from our previous work, which demonstrated this phenotype in global PKCεKO fed HFD for the same duration of time [140]. Furthermore, the absence of improved glucose tolerance in age-matched chow-fed AdPKCεKO mice is consistent with results from our global PKCεKO study [140]. Importantly, these data collectively suggest that chronic activation of PKCε in adipose tissue by increased dietary lipid is involved in events that cause the switch from a metabolically healthy to an insulin resistant state. The extent to which global PKCεKO improved glucose tolerance however, was larger than that displayed by AdPKCεKO mice. In line with our current data, the enhanced beneficial effect displayed by global PKCεKO mice after one-week HFD was not accompanied by additional compensatory increases in plasma insulin. This suggests that global PKCεKO may mediate a greater increase in whole-body insulin sensitivity than AdPKCεKO. One explanation for this could be that global PKCε deletion may be improving insulin action in further tissues in addition to adipose tissue. Delineation of these tissues however, is beyond the scope of the current study.

Decreases in plasma NEFA across all genotypes during the ipGTT (after all high-fat feeding periods) also occurred. This is most likely due to the increase in plasma insulin (which is anti-lipolytic) that occurs during the ipGTT. There was more variation in the plasma NEFA measurements between genotypes during the ipGTT, as compared with corresponding blood glucose and plasma insulin measurements. Nevertheless, the 130 overall trend from these NEFA measurements shows that PKCε deletion in adipose tissue is associated with a greater suppression of plasma NEFA levels than with WT mice during ipGTTs following 1, 8 and 16 weeks HFD. This suggests a relationship between the ablation of PKCε in adipose tissue and decreased FA release.

The further suppression of plasma NEFA displayed by AdPKCεKO mice after one- week high fat feeding (during which only liver and adipose tissue become insulin resistant) in the absence of additional compensatory increases in insulin levels is consistent with the interpretation that the improved glucose tolerance shown by these mice was due to improved hepatic insulin sensitivity mediated via decreased supply of NEFA from adipose tissue to the liver via the actions of PKCε deletion in adipose tissue. Alteration of the inflammatory response from adipose tissue has been strongly linked to the generation of systemic insulin resistance [41, 83]. However, it has been shown that neither macrophage infiltration (a marker of inflammation) nor the mRNA expression of inflammatory markers was increased within adipose tissue of mice fed a HFD for one week [26]. Hence, any potential effects of adipose tissue-specific deletion of PKCε upon adipose tissue inflammation are unlikely to play a major role in the development of insulin resistance following a one-week HFD. It is proposed therefore, that the increased suppression of NEFA release from adipose tissue is predominantly responsible for the protection against glucose intolerance induced during short-term high fat feeding. Investigations to elucidate the pathways of lipid metabolism that are modulated by PKCε in adipose tissue will be discussed in the next chapter.

The mechanism through which NEFA increases hepatic glucose production remains incompletely understood. Insulin signalling is known to down regulate hepatic glucose production through suppressing gluconeogenesis and glycogenolysis [271]. Many studies propose that NEFA overcomes this suppression acutely through inhibiting hepatic insulin signalling via the production of diacylglycerol from NEFA, which causes PKCε activation and subsequent inhibitory actions upon insulin signalling [137, 141]. However, our laboratory did not observe any PKCε-dependent modulations in hepatic insulin signalling in global PKCεKO mice after one-week HFD [140]. Furthermore, as mentioned earlier, we have previously shown that liver-specific PKCεKO mice are not protected from glucose intolerance after one-week HFD nor display enhanced insulin sensitivity (manuscript in preparation). Increased NEFA levels 131 at the liver elevate hepatic ceramide production, causing the deactivation of Akt and subsequent increases in hepatic glucose production [71]. Alternatively, it is theorised that NEFA promotes hepatic glucose production independently of insulin signalling. This may involve the promotion of hepatic NEFA oxidation, which produces ATP, NADH and acetyl-CoA required to drive gluconeogenesis, via increased hepatic NEFA supply [269, 272]. A recent study by Perry and colleagues in 2015 has shown that male rats fed a four-week HFD displayed higher rates of lipolysis than chow-fed controls in association with higher hepatic acetyl-CoA concentrations and increased activity of pyruvate carboxylase, which catalyses the first step in gluconeogenesis and requires acetyl-CoA for activation [269]. Though we have obviated the role of hepatic PKCε in inhibiting the suppression hepatic glucose production, we are yet to elucidate the exact mechanism of NEFA-induced elevations in hepatic glucose production. Despite being of great interest, such investigation is beyond the scope of our current study.

The minimal increase in plasma insulin concentrations during the ipGTT after all high fat feeding periods, for all genotypes was typical of the C57BL/6 mouse strain used in this study [273]. Though minimal differences in plasma insulin excursions were mostly seen between all genotypes during the ipGTTs (following all high-fat feeding periods), male WT-Cre mice did present noticeably lower plasma insulin concentrations during the 8 and 16-week HFD ipGTTs. At this stage it is unclear as to why this occurred. However, the data indicate that this is not linked to improvements in glucose tolerance or suppression of NEFA release in comparison with other genotypes. Nonetheless, further investigation is required to elucidate why male WT-Cre mice presented these decreases in plasma insulin. Overall, the unchanged plasma insulin profiles between genotypes importantly indicate that AdPKCεKO mice showed no additional compensatory increases in insulin secretion during all ipGTTs in this study. This leads to the interpretation that the improved glucose tolerance and further suppression of plasma NEFA concentrations displayed by AdPKCεKO mice during these ipGTTs was due to an enhancement in whole body insulin sensitivity.

The continuation of improved glucose tolerance, greater suppression of plasma NEFA concentrations and lack of compensatory insulin release by male and female AdPKCεKO mice during the 8 and 16-week HFD ipGTTs demonstrates that the protective phenotype seen after one-week HFD was preserved with extended high fat 132 feeding during which obesity, as well as the more advanced stages of insulin resistance, have been established. It has been shown that skeletal muscle, the predominant tissue involved in insulin-stimulated glucose uptake, becomes insulin resistant after three weeks of high fat feeding [25, 26]. Hence, the improvements in glucose tolerance following 8 and 16 weeks high fat diet may be attributed to improved insulin sensitivity within skeletal muscle, as well as liver.

Global PKCεKO mice in our previous study had also displayed improved glucose tolerance after longer-term high-fat feeding (6 and 16 weeks). Once again, the extent of this improvement was larger than that displayed by AdPKCεKO mice fed HFD diet for similar periods of time. In contrast to our current findings, the improvements shown by global PKCεKO mice were associated with higher plasma insulin levels during the ipGTT [140]. This was expected, in accordance with a separate study performed by our laboratory, which showed that both global PKCε deletion in mice and the inhibition of PKCε function in islets improved glucose-stimulated insulin secretion in mice fed a HFD for 16 weeks [240]. Based on this, it was not expected that AdPKCεKO mice would elicit a protective effect against glucose intolerance during longer term high-fat feeding. The unexpected improvement in glucose tolerance seen after 8 and 16 weeks HFD, in conjunction with the protective effect from PKCε deletion in adipose tissue observed during short-term high-fat feeding strongly support the notion that PKCε within adipose tissue plays a key role in mediating the various mechanisms that initiate, maintain and propagate the insulin resistant state.

The extent of the suppression of NEFA release during the 16 week HFD ipGTT is similar to that observed during the ipGTTs after one and eight weeks HFD. However, female and male AdPKCεKO mice displayed greater protection against glucose intolerance at 16 weeks than at eight weeks. This suggests that mechanisms other than decreased lipid flux from adipose tissue to the peripheral insulin-sensitive tissues may be involved following 16 weeks high-fat feeding. Alternative mechanisms proposed to mediate insulin resistance include the production of reactive oxygen species and endoplasmic reticulum stress. Each is thought to either incite an inflammatory response and/or be amplified by inflammation [274]. Obese, insulin resistant individuals frequently display continuous, low-grade inflammation [275]. In parallel, it has been

133 shown that obese individuals can be insulin resistant without elevations in NEFA concentrations (reviewed in [276]).

Adipose tissue, in addition to its role in regulating systemic NEFA concentrations, has long been recognised to secrete a wide range of proteins, termed adipokines. These are actively involved in many biological processes including inflammation, glucose metabolism and insulin sensitivity [171]. The dysregulation of adipose tissue function during obesity results in the amplified release of inflammatory cytokines (TNFα, Il-6, MCP-1) and other adipokines that promote insulin resistance (resistin and retinol- binding protein 4 (RBP-4)) in conjunction with decreased secretion of adipokines that promote insulin sensitivity, the most well-documented being adiponectin [151, 183]. It has been generally viewed that inflammation makes a more predominant contribution toward whole-body insulin resistance than enhanced FA flux from adipose tissue to peripheral insulin sensitive tissues during long term high fat feeding [26, 205, 277]. Furthermore, plasma adiponectin is inversely correlated with increasing adiposity and insulin resistance [278, 279]. Based on this, it is suggested that the protection against glucose intolerance during long term high fat feeding displayed by AdPKCεKO mice in this study may driven by potential decreases in inflammatory pathways and increases in beneficial adipokine production/release pathways. Determination of the metabolic pathways that are modulated by PKCε in adipose tissue is the subject of further investigative studies, which are discussed in the next chapter.

A shortcoming of the GTT is its inability to reveal which aspects of glucose metabolism are implicated during the test. Furthermore, the GTT is unable to confirm which tissues are implicated in these changes. The E-H clamp technique overcomes these limitations, making it the gold standard for determining insulin sensitivity [75, 76]. To date, we have only performed E-H clamps on mice after a one-week HFD. The protection against glucose intolerance exhibited by male and female AdPKCεKO mice fed long-term high fat diets provide grounds for further investigation via the E-H clamp to both confirm an association between the ablation of PKCε in adipose tissue and improved insulin sensitivity during long term-high fat feeding and to determine the tissues and processes of glucose metabolism implicated in these improvements.

134 The higher GIR seen in female AdPKCεKO mice indicates that they have higher whole- body insulin sensitivity than WT controls after a one-week HFD. Unchanged Rd in between female genotypes under clamped conditions in our study indicates that increased whole-body insulin sensitivity in females was not attributed to increased rates of whole-body glucose uptake. However, it must be noted that unchanged whole-body Rd may not necessarily indicate unaltered glucose uptake in tissues that make smaller contributions to whole body glucose uptake, such as adipose tissue. The improved suppression of HGO in female AdPKCεKO mice indicates increased hepatic insulin sensitivity in these mice. This improvement is in agreement with the increased glucose tolerance of female AdPKCεKO mice after one-week high-fat feeding, which is predominantly due to improved hepatic insulin sensitivity.

The Rg’ of a tissue is a measure of its insulin sensitivity. The slightly higher Rg’ in quad and SubQ adipose tissue, as well as the more noticeably higher Rg’ in the BAT of female AdPKCεKO mice suggests that these tissues had increased insulin sensitivity in response to the deletion of PKCε in adipose tissue. The slight increase in Rg’ within the quad muscle and the unchanged Rg’ in gastrocnemius muscle was expected because skeletal muscle does not display insulin resistance until after three weeks of high-fat feeding [25, 26]. This is consistent with the minimal changes in skeletal muscle Rg’ seen in three-day high-fat fed male rats exhibiting PKCε knockdown in liver and white adipose tissue [137]. Importantly, the absence of major increases in skeletal muscle Rg’ of female AdPKCεKO mice agrees with the unchanged Rd between female genotypes because skeletal muscle is the primary tissue involved in insulin-stimulated glucose uptake. To assess whether PKCε deletion in adipose tissue improves skeletal muscle Rg’, and therefore Rd, it would be necessary to measure skeletal muscle Rg’ after at least three weeks of HFD.

Adipose tissue displays insulin resistance after only three days of HFD [25, 26]. Therefore it was expected that differences between the Rg’ of adipose tissues from AdPKCεKO and WT-Floxed mice would occur. Female AdPKCεKO mice displayed trending increases in SubQ adipose tissue and BAT Rg’. This may suggest that a minor improvement in insulin sensitivity is elicited by the deletion of PKCε in these tissues. Further work however, is required to confirm this. Though the general consensus is that visceral adipose tissue depots, such as gonadal fat, are implicated in the generation of 135 insulin resistance [280, 281], we did not see any improvement in female gonadal fat Rg’ following the deletion of PKCε in adipose tissue. The absence of this improvement contrasts with a study by Samuel and colleagues in 2007. In this study, three-day high- fat fed male rats subjected to antisense oligonucleotide-mediated PKCε knock down in liver and white adipose tissue presented an approximate two-fold increase in gonadal fat Rg’ following PKCε knockdown [137].

Plasma NEFA concentrations decreased as expected during clamped conditions, on account of the anti-lipolytic effect of insulin. The near-significant suppression of NEFA release displayed by female AdPKCεKO mice reinforces that insulin sensitivity within at least some adipose tissue depots has improved under stimulatory conditions following the deletion of PKCε in the adipose tissue of female mice. Furthermore, this finding supports the hypothesis that PKCε increases FA supply from adipose tissue to the liver, inhibiting the suppression of hepatic glucose production by insulin.

The unchanged GIR between male genotypes suggests that PKCε deletion in adipose tissue may not affect whole-body insulin sensitivity in male mice. Unchanged Rd and was consistent with no observed difference in GIR. However, HGO in male AdPKCεKO mice did show a tendency to decrease in comparison with WT-Floxed controls, which suggests that the E-H clamp technique was not sensitive enough to detect potential minor changes in GIR and Rd between male genotypes. The absence of improved Rd and non-significant further suppression of HGO in male AdPKCεKO mice contrasts with the aforementioned PKCε knockdown study, which showed a significant suppression of HGO and increased Rd following PKCε knockdown in adipose tissue and liver [137]. The GIR, Rd and HGO findings in male mice were unexpected in light of the increased glucose tolerance shown by male AdPKCεKO mice after a one-week HFD. However, the potential impact of PKCε deletion in adipose tissue upon net hepatic glucose uptake (NHGU) must also be considered. Hyperinsulinaemia and hyperglycaemia following a meal causes the liver to switch from a net output of glucose to a net uptake of glucose [282]. However, neither hyperinsulinaemia nor hyperglycaemia on its own can stimulate NHGU [283, 284]. It is therefore possible that NHGU occurs during a GTT (which is instigated via injection of a glucose bolus “meal”), but not during the E-H clamp. A hyperinsulinaemic-hyperglycaemic clamp study involving dogs has demonstrated that high-fat feeding blunts NHGU in 136 comparison with chow-fed controls [285]. Thus, it could be argued that male AdPKCεKO mice may actually have improved hepatic and whole body insulin sensitivity following a one-week HFD through PKCε deletion in adipose tissue mediating improved NHGU via decreased plasma NEFA supply to the liver.

The absence of any noticeable changes in Rg’ between male AdPKCεKO and WT mice across all assessed tissues indicates that AdPKCεKO in male mice did not have any impact upon tissue-specific insulin sensitivity. This finding is consistent with the null change in GIR and Rd observed between male genotypes. Furthermore, the unaltered suppression of NEFA release between male genotypes coincides with absence of any improvements in the Rg’ of adipose tissues assessed in this study and contrasts with the further decreases in plasma NEFA excursions displayed by male AdPKCεKO mice during the one-week HFD GTT.

Overall, our E-H clamp studies have demonstrated that PKCε deletion in adipose tissue is associated with improved whole body insulin sensitivity in females, and to a lesser extent in males, following a one-week HFD. An increased suppression of HGO, potentially mediated by a decreased supply of FA from adipose tissue to the liver is proposed to underpin this improvement.

Mammals present many sex-based differences in fat deposition patterns, body fat percentages and in the metabolic and endocrine characteristics of various adipose tissue depots. These differences are thought to arise though differential actions of sex-specific hormones. In general, males preferentially deposit dietary NEFA in visceral adipose depots, which are implicated in metabolic dysfunction. Females on the other hand, preferentially deposit dietary NEFA into the lower-body SubQ adipose depot (which is considered to be metabolically protective). Subsequently, females display higher circulating levels of metabolically protective adipokines and a lower incidence of inflammation in comparison with males. These characteristics enable females to be generally more insulin sensitive than males [175, 181, 182, 286, 287]. PKCε may potentially be more prevalent and/or more activated in the SubQ adipose depot in comparison with visceral adipose depot. Future studies to investigate this have been mentioned earlier in this discussion. This potential increased prevalence and/or activation may diminish the ability of SubQ adipose to generate its beneficial metabolic 137 effects. As females have a greater proportion of SubQ adipose tissue in comparison with males, it is therefore proposed that the deletion of PKCε in adipose tissue may have facilitated a greater improvement in insulin sensitivity in female AdPKCεKO mice through restoring the beneficial characteristics of SubQ adipose tissue after one-week high-fat feeding.

It is generally accepted that larger adipocytes are more insulin resistant and have a greater potential to cause insulin resistance in peripheral tissues than smaller adipocytes (see Section 1.13 for further discussion). Furthermore, a study in overweight humans has reported a positive association between adipocyte size and ectopic lipid accumulation in liver and other visceral tissues [288]. Our results showed that a higher proportion of gonadal adipocytes from male and female AdPKCεKO mice after 16 weeks high fat feeding were smaller in area than WT controls. This important finding suggests that PKCε deletion in adipose tissue is associated with the improved function and decreased pathogenic potential of adipocytes during long-term high-fat feeding. This could be due to potential PKCε-mediated alterations towards adipokine and inflammatory cytokine production and release from adipose tissue. The contribution of adipokine and adipose-tissue specific cytokine production toward whole-body insulin resistance has been discussed in Section 1.12.

Our results provide compelling evidence to suggest PKCε acts indirectly via adipose tissue to increase hepatic glucose production. This is significant for a number of reasons. Firstly, it overturns the long-standing dogma which postulates that PKCε acts directly at the liver to achieve this outcome. Secondly, our findings provide further support for the emerging theme that PKCε can be a controller of lipid metabolism, in addition to being an effector of this metabolic process. It has long been known that dysfunctional lipid metabolism contributes to the generation of insulin resistance. The mechanisms underpinning this however, remain unclear. The findings presented in this chapter form a basis for gaining greater understanding of the mechanisms behind dysfunctional FA flux in adipose tissue and its associated impact on the generation and maintenance of hepatic and whole body insulin resistance.

We have extensively characterised the phenotype of PKCε deletion in adipose tissue within the context of glucose tolerance, insulin sensitivity and adipocyte size. An 138 important step is the identification of the genes, proteins and pathways in adipose tissue that are modulated by the deletion of PKCε in adipose tissue to mediate its protective phenotypes. Such delineation will be key to the development of targeted therapeutics designed to mimic the ablation of PKCε in adipose tissue and subsequently alleviate the development and/or continuation of insulin resistance in multiple tissues. These investigations are discussed in the next chapter.

139 CHAPTER 5 EFFECTS OF PROTEIN KINASE C EPSILON DELETION IN ADIPOSE TISSUE UPON LIPID METABOLISM AND CYTOKINE RELEASE

140 5.1 Introduction Our in vivo PKCε characterisation studies in the previous chapter demonstrated that PKCε ablation in adipose tissue is associated with improved glucose tolerance, suppressed NEFA release (under acute stimulatory conditions), enhanced whole-body insulin sensitivity and reduced adipocyte size during short and long term high-fat feeding. However, the metabolic pathways that are modulated upon the deletion PKCε in adipose tissue to achieve these outcomes were not addressed.

Most of the literature to date proposes that chronic PKCε activation via dietary lipid oversupply causes insulin resistance through direct interference of the canonical insulin signalling pathway in liver [136, 137, 143, 144]. In contrast, our in vivo studies suggest that the improvements in glucose tolerance and insulin sensitivity displayed by AdPKCεKO mice during a short term HFD may be due to associated decreases in FA flux from adipose tissue to the liver. PKCε has been strongly detected in lipid droplets, in addition to its localization at the plasma membrane [243]. Furthermore, PKCε activation is strongly proportional to DAG content at the lipid droplet [243]. This suggests that PKCε has the potential to impose actions upon lipid metabolism that occur within the lipid droplet. Studies from our laboratory involving global PKCε knockout mice under different nutritional states indicate that PKCε is involved in the modulation of lipid partitioning towards TAG storage in the pancreas [240], liver [140, 244], and adipose tissue [244]. Furthermore, we have also shown that cultured primary adipocytes obtained from global PKCεKO mice incorporated a higher amount of radiolabelled fatty acid tracer into TAG in comparison with WT-controls (unpublished work). This supports the suggestion that PKCε ablation in adipose tissue during a short-term HFD caused direct modulations in lipid metabolic pathways in adipose tissue, reducing FA flux toward the liver and subsequently protecting AdPKCεKO mice against glucose intolerance and insulin resistance.

The nature of the findings in the previous chapter also suggest that the improvements in glucose tolerance displayed by AdPKCεKO mice during a long term HFD may be associated with decreased inflammation. There is evidence in the literature to suggest that PKCε plays a pro-inflammatory role in myotubes [289] and astrocytes [290]. In

141 addition, PKCε has been shown to induce the expression of the pro-inflammatory cytokine IL-6 in adipocytes [291].

The objective of this chapter was to determine the mechanisms underpinning the link between PKCε ablation in adipose tissue and improved whole-body glucose tolerance and insulin sensitivity. To achieve this we aimed to (i) determine if PKCε ablation in adipose tissue altered the expression of genes encoding proteins involved in lipid metabolism, inflammation and adipocyte differentiation (ii) examine if PKCε ablation in adipose tissue altered total protein levels of lipid metabolism enzymes, and (iii) investigate whether PKCε knockout in adipose tissue mediated a decrease in inflammation via altered cytokine release.

142 5.2 Results

5.2.1 PKCε ablation in adipose tissue does not alter the expression of several genes involved in lipid metabolism, inflammation and adipocyte differentiation We first examined the mRNA levels of genes involved in lipid metabolism, inflammation and adipocyte differentiation to determine whether any of these physiological processes are altered in response to PKCε ablation in adipose tissue. mRNA was harvested from the gonadal adipose depot of AdPKCεKO and WT-floxed mice (from our ipGTT cohorts) fed a 16-week HFD, following a six-hour fast. This was converted into cDNA and quantified by RT-PCR. Procedures were conducted as per section 2.5.1.

Overall, there were no trending or significant changes in gene expression of most genes assessed. Notable exceptions included a significant increase in tnfα mRNA in female AdPKCεKO adipose tissue (Figure 5-1(c)) as well as trending increases in phlda1 and tnfα mRNA in male AdPKCεKO adipose tissue (Figure 5-2(b) and (c)). A trending decrease in leptin mRNA was also noticed in male AdPKCεKO adipose tissue (Figure 5-2(b)).

143 a 2.0 WT-Flox 2.0 Ad-PKCεKO

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hsl atglatgl HSLhsl gpam Gpamgpam lipin-1lipin-1Dgat-1dgat-1dgat-1Dgat-2dgat-2dgat-2ATGL Agpat-2agpat-2agpat-2Lipin-1 Perilipin-1perilipin-1perilipin-1Perilipin-2perilipin-2perilipin-2 2.0 b 2.0

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α α β β β β tnf il-1 inos saa CD68cd68 IL-1 iNOS F4/80f4/80SAA tgf- MCP-1mcp-1 TNF TGF- Figure 5-1 mRNA levels in female murine adipose tissue. mRNA was harvested from the gonadal adipose depots of female AdPKCεKO and WT- floxed mice fed a 16-week HFD, following a six-hour fast. mRNA was converted into cDNA and quantified by RT-PCR. Results expressed as fold change compared to WT- floxed control. a) Lipid metabolism genes b) Transcription factor, adipocyte-specific hormone and differentiation genes c) Inflammatory genes. WT-Floxed n=12 (black bars), AdPKCεKO n=14 (orange bars). Student’s T-test: *p < 0.05. 144 a 2.0 WT-Flox Ad-PKCεKO 1.5

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hsl atgl HSL Gpamgpam lipin-1Dgat-1dgat-1Dgat-2dgat-2 ATGL Agpat-2agpat-2Lipin-1 Perilipin-1perilipin-1Perilipin-2perilipin-2

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0.00.0 αα ββ ββ tnf il-1 inos saa CD68cd68 IL-1 iNOS F4/80f4/80 SAA tgf- MCP-1mcp-1 TNF TGF-

Figure 5-2 mRNA levels in male murine adipose tissue. Experimental details as per Figure 5-1. a) Lipid metabolism genes b) Transcription factor, adipocyte-specific hormone and differentiation genes c) Inflammatory genes. WT-Floxed n=12 (black bars), AdPKCεKO n=10 (orange bars).

145 5.2.2 PKCε ablation in adipose tissue is associated with increased protein expression of lipid esterification enzymes The knockout of PKCε in adipose tissue may mediate beneficial effects on glucose homeostasis during a long term HFD through the modulation of lipid metabolism pathways via altering the expression of proteins in a post transcriptional manner. To assess this possibility, proteins were extracted from the gonadal adipose tissue of AdPKCεKO and WT-floxed mice fed a 16-week HFD, following a six-hour fast. The infranatants (aqueous) and lipid fractions from tissue homogenates were isolated and separately immunoblotted for proteins involved in lipid esterification and lipolysis. Protein levels were quantified via densitometry. Further details on these procedures are outlined in section 2.5.2. DGAT-2 protein bands from the lipid fraction of male and female mice, as well as LIPIN-1 protein bands from the lipid fraction of male mice were not quantified as they below the level of detection.

Overall, there were no trending and/or significant differences in the levels of most proteins assessed between AdPKCεKO and WT-floxed mice, for both genders (Figure 5-3, Figure 5-4, Figure 5-5, Figure 5-6). Exceptions to this included a significant increase in DGAT-2 protein levels and a trending increase in LIPIN-1 protein levels (p=0.06) from the infranatant-fraction protein extract of male AdPKCεKO mice in comparison to WT-Floxed mice (Figure 5-6). Trending increases in GPAM protein levels were also observed within lipid-fraction protein extracts of male and female AdPKCεKO mice (Figure 5-4 and Figure 5-6). All observed changes occurred with proteins involved in the FA esterification pathway. Therefore, it is suggested that PKCε ablation in adipose tissue is associated with increased protein expression of lipid esterification enzymes.

146 Total-HSL - Infranatant Total-HSL - Fat 250 250

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Figure 5-3 Lipolysis protein levels in female murine adipose tissue.

Protein was harvested from gonadal adipose tissue of female AdPKCεKO and WT- floxed mice fed a 16-week HFD, after a six-hour fast. The infranatant (aqueous) and lipid fractions from the total protein extract were isolated and subjected to immunoblotting for lipolysis proteins. Protein levels were quantified via densitometry and normalised to protein levels of non-muscle myosin (housekeeping protein). Normalised protein amounts are expressed as percentage relative to WT-Floxed control. Representative images of each examined protein in female AdPKCεKO and WT-floxed mice are shown below the corresponding histogram. WT-Floxed n=12, AdPKCεKO n=14.

147

GpamGPAM - -Infranatant Infranatant GpamGPAM - - Fat Fat 250 250 200 200

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Figure 5-4 Lipid esterification protein levels in female murine adipose tissue.

Details as outlined in Figure 5-3. WT-Floxed n=12, AdPKCεKO n=14. 148 Total-HSL - Infranatant Total-HSL - Fat 200 200

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Figure 5-5 Lipolysis protein levels in male murine adipose tissue.

Protein was harvested from gonadal adipose tissue of male AdPKCεKO and WT-floxed mice fed a 16-week HFD, after a six-hour fast. The infranatant (aqueous) and lipid fractions from the total protein extract were isolated and subjected to immunoblotting for lipolysis proteins. Protein levels were quantified via densitometry and normalised to non-muscle myosin (housekeeping protein). Normalised protein amounts are expressed as percentage relative to WT-Floxed control. Representative images of each examined protein are shown below each corresponding histogram. WT-Floxed n=12, AdPKCεKO n = 10.

149 GPAMGpam -- InfranatantInfranatant GPAMGpam - Fat GPAM - Infranatant GPAM - Fat 200 200

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Figure 5-6 Lipid esterification protein levels in male murine adipose tissue.

Details as per Figure 5-5. Student’s T-test: * p<0.05 AdPKCεKO vs WT-Floxed. 150 5.2.3 PKCε ablation in adipose tissue is associated with increased IL-6 and decreased TNFα plasma concentrations It was suggested in the previous chapter that the enhanced glucose tolerance exhibited by AdPKCεKO mice during 16 weeks of HFD could be due to decreased inflammatory cytokine release from adipose tissue. To investigate this, we quantified the concentrations of inflammatory (TNFα, IL-1β, IL-6) and anti-inflammatory (IL-10) cytokines from plasma extracted from blood samples collected from 16-week high-fat fed WT-Floxed and AdPKCεKO mice following a six-hour fast. Details of the employed cytokine quantification procedure are listed in section 2.4.3.

IL-1β plasma concentrations were below the limit of detection in both male and female samples (Figure 5-7). Female mice displayed no difference in IL-10 plasma concentrations between genotypes. Female AdPKCεKO mice however, presented an increase in IL-6, as well as a decrease in TNFα plasma concentration as compared to WT controls (Figure 5-7 (a)).

IL-10 was below the limit of detection in all male mice assessed. IL-6 concentrations however, were near significantly increased in male AdPKCεKO mice (p=0.07). TNFα plasma concentrations did not change between male genotypes (Figure 5-7 (b)).

151 a Females WT-Flox 35 Ad-PKCεKO

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15 p = 0.07

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0 IL-10 IL-1β IL-6 TNFα

Figure 5-7 Plasma cytokine concentrations in AdPKCεKO and WT-floxed mice.

Concentrations of inflammatory (TNFα, IL-1β, IL-6) and anti-inflammatory cytokines (IL-10) were quantified from plasma extracted from blood samples collected from 16- week high-fat fed WT-Floxed (black bars) and AdPKCεKO (orange bars) mice after a six-hour fast. a) Female mice: WT-Floxed n=6, AdPKCεKO n=7 (b) Male mice: WT- Floxed n=5 AdPKCεKO n=8.

152 5.3 Discussion The results presented in this chapter have allowed us to gain a preliminary insight into how PKCε ablation in adipose tissue may modulate lipid metabolism, inflammation and adipocyte differentiation in response to a long term HFD. The deletion of PKCε in adipose tissue did not alter the expression of most genes encoding proteins involved in lipid metabolism and inflammation during extended high-fat feeding periods. Phlda1, a gene involved in adipocyte differentiation however, showed a tendency to be upregulated in our knockout model. The ablation of PKCε in the adipose tissue of 16- week fat-fed mice was linked to elevated protein expression of several lipid esterification proteins. In addition, increased plasma IL-6 and decreased TNFα cytokine concentrations were observed in AdPKCεKO mice during long-term high fat feeding.

The experiments in this chapter were only performed in 16-week HFD-fed mice. However, parallel experiments with age-matched chow-fed mice showing expected, diet-induced changes would help confirm that the lack of change in the expression of genes and proteins between WT-Floxed and AdPKCεKO mice was not due to inadequate sensitivity of our measurements. Furthermore, parallel studies in one-week fat fed mice will be required to assess whether greater alterations in the expression of lipid metabolism genes and proteins occur, in agreement with the general consensus that altered FA flux from adipose tissue to the liver is the predominant mediator of adipose- tissue induced insulin resistance during short-term high-fat feeding.

Overall, there were minimal differences between WT-Floxed and AdPKCεKO genotypes with respect to the expression of most genes assessed in this study. There were however, a few notable observations. A trending decrease in leptin mRNA expression was observed in the adipose tissue of male AdPKCεKO mice. Leptin is an adipokine which suppresses appetite and increases energy expenditure to maintain healthy energy balance. A decrease in leptin mRNA expression in male AdPKCεKO mice may therefore suggest that these mice had a higher body weight than WT-Floxed mice. This however, was not the case as there was no difference in body weights between male genotypes immediately prior to tissue harvest (data not shown).

153 An increase in adipose tissue tnfα gene expression was seen in both male and female AdPKCεKO mice. Though this may lead to the interpretation that inflammation is up regulated following PKCε deletion in adipose tissue, this should be considered together with other data. Inflammatory cytokines such as TNFα, IL-6 and MCP-1 are primarily produced and secreted from macrophages within adipose tissue rather than adipose tissue per se [277, 292, 293]. However, there were no observed increases in the expression of various macrophage markers, such as cd68 and f4/80, in the adipose tissue of AdPKCεKO mice. This suggests that macrophage generation, and therefore subsequent increases in inflammatory cytokine production, was not increased in AdPKCεKO mice (this will need to be confirmed via histological analysis of adipose tissue for macrophage accumulation and cytokine appearance). Also, no other genes encoding proteins associated with inflammation displayed increased mRNA expression. Furthermore, we had shown in the previous chapter that glucose tolerance was improved in male and female AdPKCεKO mice following 16 weeks of high-fat feeding. This result suggests that inflammation was down-regulated in these mice as inflammation is proposed to be the predominant mechanism underlying glucose intolerance during long term HFD [26, 194, 205]. These considerations lead to the suggestion that the observed increase in tnfα mRNA expression in the adipose tissue of AdPKCεKO mice was an anomaly and therefore not a true reflection of the metabolic environment within the adipose tissue of AdPKCεKO mice after a 16-week HFD.

The final notable change observed during our RT-PCR studies was the near-significant increase in phlda1 expression in the adipose tissue of male AdPKCεKO mice as compared to WT controls. The phlda1 gene encodes the protein T-Cell death-associated gene 51 (TDAG51), which has recently been found to play an important role in mediating preadipocyte differentiation into mature adipocytes - this is conducive to improved adipose tissue function and insulin sensitivity. Furthermore, the expression of phlda1 is inversely correlated to hepatic steatosis in mice [294]. Hence, it is suggested that the increase in phlda1 expression in this study may increase the formation of new adipocytes which may subsequently contribute toward the protective effect of PKCε deletion in male adipose tissue against insulin resistance during long term high- fat feeding. Phlda1 expression only increases during a short, specific time point of adipogenesis [294, 295]. In contrast, a separate study has found that PKCε protein 154 expression in fact increases in cultured pre-adipocytes during this timepoint [296]. To confirm that Phlda1 expression increases in AdPKCεKO male adipocytes (as well as female AdPKCεKO adipocytes) we will need to assess phlda1 expression within preadipocytes from WT and AdPKCεKO mice ex vivo over a time course following the induction of preadipocyte differentiation. Furthermore, we must assess the gene expression of other mediators and/or indicators of adipocyte differentiation, such as CCAAT/enhancer binding protein (C/EBPβ) and GLUT-4 [297] to confirm if the ablation of PKCε in adipose tissue is linked to increased adipocyte differentiation. During this study, we had attempted to measure the mRNA levels of preadipocyte factor 1 (pref-1), another factor involved in adipocyte differentiation [298]. However, the mRNA levels of this gene were below the limits of detection. The absence of changes in the expression of most genes in this study between genotypes, as well as the questionable impact of most observed changes, suggest that the improvement in insulin sensitivity displayed by AdPKCεKO mice was not associated with alterations in the expression of the genes encoding proteins involved in lipid metabolism and inflammation within gonadal adipose tissue.

Immunoblot analysis of the protein levels of lipid metabolism enzymes was conducted to assess whether PKCε ablation in adipose tissue during long term HFD altered the protein expression of these enzymes and hence, modulated the regulation of their associated lipid metabolism pathways. The observation of increased protein levels of multiple FA esterification enzymes (GPAM, LIPIN-1 and DGAT-2) within the gonadal adipose tissue of AdPKCeKO mice provides convincing early evidence to suggest that the FA esterification pathway is up-regulated in response to PKCε knockout in adipose tissue. Female AdPKCεKO mice did not display as many increases in enzyme levels as male AdPKCεKO mice. This may be due to the variability and semi-quantitative nature of immunoblots of subcellular fractions. The findings observed in these immunoblot experiments are in line with our previous published and unpublished findings (discussed earlier), which have shown an association between PKCε ablation and elevated triacylglycerol (TAG) levels in adipocytes, adipose tissue, pancreatic islets and liver [140, 240].

155 The unchanged mRNA expression of the gpam, lipin-1 and dgat-2 genes observed in AdPKCεKO mice when compared to WT-Floxed mice, together with their increased protein levels implies that PKCε, when present, may decrease protein levels through (i) conducting inhibitory post-transcriptional effects on the mRNA which encodes these FA esterification enzymes (ii) performing inhibitory effects on the protein machinery required for the specific translation of these genes, or (iii) activation of the cellular sensing mechanisms that may induce other potential inhibitory post-transcriptional modifications. PKCε, on account of its kinase activity, is most likely to generate these effects through phosphorylation.

A chronic up-regulation of the FA esterification pathway within adipose tissue would result in increased TAG levels and the subsequent expansion of this tissue. Paradoxically, we did not see any noticeable differences in gonadal adipose tissue weight between WT-Floxed and AdPKCεKO mice for both genders (Figure A-4). An explanation for this paradox may be found when the process of FA re-esterification is taken into account. Fifty percent of FA molecules released by lipolysis are taken back up into adipose tissue and re-esterified into TAG molecules under basal conditions. This percentage of re-esterified FAs increases with insulin stimulation because it increases FA uptake into adipose tissue, in turn providing more available FA for esterification [150, 161]. In this context, it may therefore be plausible that the increased protein levels of FA esterification enzymes in AdPKCεKO mice temporarily lead to a higher proportion of FAs being incorporated into TAG, in comparison to WT controls, during periods of acute systemic insulin increase (which occurs in response to a glucose challenge). This explanation may reconcile the potential up-regulation of the FA esterification pathway in AdPKCεKO mice with the minimal change in gonadal adipose tissue weight observed between genotypes (Figure A-4).

In adipose tissue, adipocytes are surrounded by a variety of immune cells, including macrophages and dendritic cells. FAs released from adipocytes are known to directly induce macrophage infiltration into adipose tissue, convert resident macrophages within adipose tissue from the M2 non-inflammatory phenotype to the M1 inflammatory phenotype and activate dendritic cells. This occurs mainly via FA-induced activation of TLR2/4 signalling on these immune cells, resulting in adipocyte dysfunction, local

156 cytokine accumulation and increased cytokine release from adipose tissue [83, 299, 300]. Upregulated esterification, even if temporary, may therefore reduce these immune cell processes through increasing the amount of FA within adipose tissue which becomes stored as metabolically inert TAG. This coincides well with the proposition that reduced inflammation may predominantly account for the dramatic increase in glucose tolerance shown by AdPKCεKO mice after 16 weeks of HFD.

Lipid esterification may not be the only route of lipid metabolism that is modulated by PKCε within adipocytes. Only two out of a multitude of proteins involved in lipolysis, total-HSL and PERILIPIN-1, were assessed in these immunoblot experiments. The effect of phosphorylation on esterification enzymes is poorly defined [156, 157]. However, it is widely known that phosphorylation is required for the activation and inhibition of many lipolytic proteins [154]. As this post-translational modification usually occurs during acute stimulatory conditions or under chronic starvation, we were unable to measure the protein levels of phosphorylated forms of HSL and PERILIPIN-1 in the current study. The absence of change in the levels of total HSL and PERILIPIN-1 therefore, does not essentially rule out that the amount of lipolysis occurring in adipocytes is unaltered after PKCε ablation in adipose tissue.

Basal plasma concentrations of inflammatory (TNFα, IL-1β, IL-6) and anti- inflammatory cytokines (IL-10) were measured in AdPKCεKO mice after 16 weeks of HFD to investigate whether associated improvements in glucose tolerance were due to PKCε−induced alterations in cytokine release from adipose tissue. The plasma concentration of each cytokine from our findings was comparable with those noted in the literature [301, 302] and from unpublished work by colleagues. Contrary to expectation, AdPKCεKO mice either did not display an increase in the plasma concentration of anti-inflammatory cytokine (IL-10), or did not present detectable levels of this cytokine. The trending decrease in TNFα plasma concentration shown by female AdPKCεKO mice and the unchanged TNFα plasma concentration between male genotypes is difficult to interpret alongside the increased tnfα mRNA expression observed in male and female AdPKCεKO mice and therefore requires further investigation. It may be that the trending decrease in TNFα shown by female

157 AdPKCεKO mice suggests that the deletion of PKCε in adipose tissue is associated with a decrease in inflammation.

Increased IL-6 plasma concentrations displayed by AdPKCεKO mice were unexpected. This finding contrasted with the work of Ohashi and colleagues, which demonstrated that PKCε overexpression in the 3T3-L1 adipocyte cell line increases IL-6 mRNA expression and IL-6 protein secretion into culture media [291]. Furthermore, the role of IL-6 in the inflammatory response remains unclear. It is widely acknowledged that IL-6 is a pro-inflammatory cytokine, which is chronically increased during obesity and associated with the pathophysiology of systemic and local insulin resistance [269, 303, 304]. Conversely, IL-6 has also been linked to improved insulin sensitivity and glucose homeostasis in a number of peripheral tissues including liver and skeletal muscle [305- 308]. A protective role of IL-6, rather than a pro-inflammatory role, within AdPKCεKO mice would lie in strong agreement with the findings from our characterisation studies from the previous chapter. Therefore, we propose that the observed increase in plasma IL-6 in male and female AdPKCεKO mice may potentially mediate a protective effect against insulin resistance during long-term high fat feeding. Further studies are required to confirm this.

Adiponectin, as discussed earlier, is an adipose tissue-specific hormone that exhibits insulin-sensitizing effects and is inversely correlated to obesity and insulin resistance [168, 171, 309]. Therefore, it is important to assess the whether PKCε ablation in adipose tissue affects adiponectin secretion after long-term high fat feeding. Based on the significant improvement in glucose tolerance in male and female AdPKCεKO mice during the GTT after 16 weeks of HFD, we hypothesise that an increase in plasma adiponectin concentrations may also occur in AdPKCεKO mice.

The findings in this chapter have highlighted of some of the metabolic processes which may be responsible for the enhanced glucose tolerance and insulin sensitivity displayed by AdPKCεKO mice. Firstly, we have demonstrated that these beneficial effects were not associated with PKCε−induced alterations in the expression of genes encoding proteins involved in lipid metabolism and inflammation during long-term high fat feeding. There was however, early evidence to suggest an association between PKCε in 158 adipose tissue and the mRNA expression of adipocyte differentiation genes under these conditions. Secondly, we have provided consistent evidence to suggest that the lipid esterification pathway is upregulated in the adipose tissue of AdPKCεKO mice fed a long-term HFD as indicated by increased protein levels of multiple enzymes in this pathway. Finally, we have shown that deletion of PKCε in the adipose tissue of 16- week fat-fed mice is associated with both a trending decrease in the secretion of the pro- inflammatory cytokine TNFα and a trending increase in secretion of the IL-6 cytokine which despite its long recognised role as a facilitator of insulin resistance, has been shown in recent years to mediate protection against this condition.

Our findings are compelling as they provide novel evidence supporting the emerging ideas that (i) PKCε is a controller of lipid metabolism, in addition to its role as an effector (ii) PKCε may decrease insulin sensitivity through routes other than the direct impairment of canonical insulin signalling and (iii) PKCε enhances the inflammatory response, which is proposed to be a primary mediator of whole-body insulin resistance during long-term high fat feeding, which is representative of the obese state. This knowledge, in conjunction with further studies to confirm current findings, elucidate further associated pathways and identify the direct targets of PKCε phosphorylation within these pathways, places us closer toward determining how PKCε in adipose tissue is implicated in the generation of dysfunctional lipid metabolism, enhanced inflammation and therefore, the subsequent generation of whole-body insulin resistance.

159 CHAPTER 6 SUMMARY AND FUTURE DIRECTIONS

160 Insulin resistance is a fundamental contributor toward the pathogenesis of T2D, which currently affects millions [3]. The ceramide accumulation in skeletal muscle and DAG- induced activation of PKCε in liver during dietary lipid excess are associated with insulin resistance in these tissues. However, there is a lack of knowledge pertaining to the mechanisms that link these factors. An acquisition of this understanding is important, as it will drive the design of improved therapeutics tailored toward targeting the exact metabolic pathways that specifically underpin insulin resistance.

The advent of lipidomics has unraveled a plethora of individual ceramide species which are hypothesised to display distinct physiological functions. The biological roles for many of these moieties however, have not been elucidated. Recent findings from our laboratory have for the first time suggested that ceramide species produced upon overexpression of specific CerS isoforms may not impede or may even improve insulin action in skeletal muscle cells [131]. In Chapter 3, we reinforced these findings and indicated the potential routes of ceramide synthesis that may produce these ceramide species in skeletal muscle.

A multitude of studies have established that enhanced FA flux from adipose tissue toward the liver is linked to elevated hepatic glucose production, a hallmark of hepatic insulin resistance [52, 53] [54] [55, 268]. However these studies are causative, providing minimal mechanistic insight into this important connection. We have previously failed to observe improved glucose tolerance in liver-specific PKCε knockout mice but obtained preliminary evidence suggesting a direct role for this enzyme in lipid metabolism. This led our laboratory to hypothesise that PKCε may be enhancing hepatic glucose production indirectly by promoting lipid efflux from adipose tissue. Subsequently, a full characterisation of the effects of PKCε in adipose tissue toward glucose tolerance, FA release and insulin sensitivity was conducted in Chapter 4. Furthermore, we have identified metabolic processes that may in part be responsible for these effects in Chapter 5. The main findings from this thesis are discussed below.

Our overall goal during our ceramide investigations was to further examine the novel protective effects upon skeletal muscle insulin action seen after the modulation of CerS isoforms. We determined that compensatory changes in the endogenous mRNA

161 expression of CerS isoforms and other ceramide synthesis enzymes overall did not occur after CerS overexpression. This eliminated compensatory changes in gene expression as a possible avenue for eliciting these novel effects upon insulin action in skeletal muscle. This investigation could be further extended to assess changes in protein expression and post-translational modification of CerS isoforms or other sphingolipid metabolism enzymes.

Despite this null result, we did highlight that enhanced flux through the salvage pathway of ceramide synthesis may potentially produce more metabolically beneficial ceramide species after the overexpression of multiple CerS isoforms. Though no changes in flux were seen through the de novo pathway of ceramide synthesis, we cannot completely rule out the potential of this pathway to elicit protective effects towards skeletal muscle insulin action. Repeating these flux assays using shorter periods of radiolabelled tracer exposure as well as in the absence of palmitate treatment may reveal flux changes through de novo pathway in response to CerS overexpression.

A final key finding observed during our ceramide investigations was the improvement and/or absence of notable decrease in insulin-stimulated GLUT4 translocation after the overexpression of all CerS isoforms. This result further confirmed the novel ability of certain ceramides to improve or at least not impede insulin action in skeletal muscle. It was important that this confirmation was achieved especially in light of recent publications, which have implicated CerS6 and its main ceramide product as mediators of insulin resistance. These publications however, do not demonstrate this specifically in skeletal muscle.

Overall the findings from Chapter 3 are significant as they establish that the physiological functions of ceramides in the context of skeletal muscle insulin resistance are more complex than suggested by the majority of current literature. Validation of these results may be achieved through parallel experiments involving the knockdown of individual CerS isoforms. Furthermore, the significance of our findings could be verified by extending the scope of our investigations to in vivo studies, which will evaluate and investigate the effects of skeletal muscle-specific overexpression and knockdown of CerS isoforms on glucose homeostasis in fat-fed mice.

162 The next step toward elucidating the metabolically beneficial effects of certain ceramides following CerS overexpression in skeletal muscle is to determine their intracellular site of action. Subsequently this will dictate the design of therapeutics that may promote the beneficial actions of ceramide species at these sites. One possible way of identifying the intracellular localisation of ceramide species is through the use of fluorescently labelled ceramide precursors in combination with fluorescence microscopy. Alternatively, ceramide species in different subcellular fractions could be determined. However, the extensive cytoskeletal network within skeletal muscle cells may present many challenges in this respect.

Characterisation of in vivo metabolic effects following the deletion of PKCε specifically in adipose tissue was successfully carried out in this thesis. This was the first time that such an investigation had been conducted. Firstly we observed that PKCε deletion in adipose tissue improved glucose tolerance during short and long term high fat feeding in association with greater suppression of plasma NEFA without compensatory increases in insulin secretion. Improved glucose tolerance during short- term high fat feeding was proposed to occur through increased suppression of NEFA release from adipose tissue. Alternatively, improvements in glucose tolerance during long term high-fat feeding were proposed to occur at least in part through decreases in inflammatory pathways and increases in beneficial adipokine release pathways in adipose tissue.

The suggestion that ablation of PKCε in adipose tissue mediated increases in whole- body insulin sensitivity was confirmed under conditions of short-term high-fat feeding by E-H clamp studies. During these investigations, female AdPKCεKO mice displayed improved whole-body insulin sensitivity. Increased hepatic insulin sensitivity, potentially mediated by further suppression of NEFA release, was proposed to drive this improvement. As such, it is suggested that the deletion of PKCε in adipose tissue diminishes the flux from adipose tissue to the liver to improve hepatic insulin sensitivity. Conducting the E-H clamp with simultaneous lipid infusion could confirm if increased suppression of NEFA release from adipose tissue is primarily responsible for the protection against glucose tolerance induced during short-term high-fat feeding. If the lipid infusion causes the GIR of KO mice to decrease to the same level as that of

163 lipid-infused WT mice, then this would provide strong evidence that diminished lipid flux from adipose tissue to liver (via PKCε deletion in adipose tissue) was responsible for the improvements in whole body insulin sensitivity observed after short-term high- fat feeding.

Though male AdPKCεΚΟ mice did not display significant improvements in insulin sensitivity in during the E-H clamp experiments, we cannot rule out that the deletion of PKCε in male adipose tissue does not improve glucose homeostasis and insulin sensitivity. Assessment of NHGU, which cannot be achieved during the E-H clamp, may be required to confirm whether this may be the case. In addition, it has been shown that pharmacologically decreasing FA release from adipose tissue enhances the ability of glucose to decrease hepatic glucose production independently of insulin - this capability is termed “glucose effectiveness” [310]. Investigating the effect of adipose tissue PKCε deletion upon glucose effectiveness therefore, may also be an alternative way to determine whether this deletion in males improves whole-body glucose homeostasis. Furthermore, the influence of FAs on glucose effectiveness underscores the importance of finding more effective therapeutic therapies directed toward lowering FA concentrations.

The varying Rg’ observed between different adipose tissue depots highlights the importance of examining adipose tissue depots from various regions of the body as separate entities. This is due to the fact that adipose tissue depots display differences in adipocyte size, lipid metabolism, vascularity, developmental gene expression, inflammatory susceptibility and adipokine production, which in turn may dictate the contribution of each adipose tissue depot toward hepatic insulin sensitivity [186, 311, 312]. In-depth comparisons of adipose tissue depots from WT mice employed in this study will enable these differences to be characterised.

A final key finding in our characterisation study was the association between PKCε deletion in the adipose tissue of 16-week fat-fed mice and smaller adipocyte size, a trait which indicates improved insulin sensitivity. To confirm whether this decrease in size parallels with decreased inflammation in these mice, we will analyse adipose tissue sections for the presence of macrophages and inflammatory cytokines.

164 Further mechanistic studies in Chapter 5 provided valuable insight into how PKCε deletion in adipose tissue mediated associated improvements in glucose tolerance and insulin sensitivity. PKCε deletion in adipose tissue from 16-week fat-fed mice did not alter the expression of several genes involved in lipid metabolism, inflammation and adipocyte differentiation. However it must be noted that only gonadal fat, a visceral adipose tissue depot, was employed to assess gene expression. Though it is generally accepted that most visceral fat depots (including gonadal fat) are less metabolically protective than lower-body subcutaneous fat [293, 311], the expression and/or activation of PKCε or its downstream mediators may also differ between visceral adipose tissue depots. PKCε may therefore regulate the expression of genes that underpin the particular metabolic characteristics of visceral adipose tissue to a greater degree in certain visceral adipose depots compared with others. In addition, the ablation of PKCε in lower-body subcutaneous tissue may also further improve its metabolically protective phenotype through altering the expression of these particular genes. Comparative gene expression analysis studies in other adipose tissue depots must be conducted to rule out that the deletion of PKCε in adipose tissue is not associated with modifying the expression of the genes assessed in this study.

Immunoblot analysis of adipose tissue from 16-week fat-fed mice demonstrated that PKCε deletion in adipose tissue was associated with increased protein expression of particular enzymes of lipid esterification. This suggests that observed decreases in NEFA release may be, in part, mediated by increased FA re-esterification. Assessment of lipid esterification in primary adipocytes from WT-Floxed and AdPKCεKO mice using radiolabelled palmitate tracer will help to confirm increased glycerolipid synthesis within the adipose tissue of AdPKCεKO mice. Evaluation of the effect of PKCε ablation in adipose tissue on the expression of proteins involved in fatty acid uptake and de novo lipogenesis will also be conducted in future experiments in combination with current efforts to optimise other procedures designed to measure lipolysis and FA re-esterification. In parallel, our laboratory will commence immunoblot analysis of proteins involved in mediating the inflammatory response.

A final result in this chapter was the trending increase in plasma concentrations of the IL-6 cytokine in AdPKCεKO mice. Whether IL-6 is a pro or anti-inflammatory 165 cytokine remains controversial. However, an increase in anti-inflammatory IL-6 within 16-week fat-fed AdPKCεKO mice would support the notion that decreased inflammation in adipose tissue in these mice was likely to be responsible in part for the observed improvements in glucose tolerance under these nutritional conditions.

Determination of the in vivo effects of PKCε ablation in adipose tissue on glucose and lipid homeostasis has provided convincing evidence to suggest that PKCε acts indirectly via adipose tissue to increase hepatic glucose production during various stages of high fat feeding. This is in contrast to the current consensus in the literature which proposes that PKCε acts directly at the liver to achieve this. Furthermore, these findings present PKCε specifically in adipose tissue as an important therapeutic target for treatment of insulin resistance and subsequent prevention of T2D. Identification of potential metabolic pathways which are altered by PKCε ablation in adipose tissue to achieve these effects further underscores the role of PKCε as a modulator of lipid metabolism and inflammation in adipose tissue. Overall, the results from chapters 4 and 5 generate a new understanding of the generation of dysfunctional FA flux from adipose tissue and its subsequent actions on hepatic and whole-body insulin resistance. From this knowledge, we are better placed to identify the protein targets of PKCε phosphorylation which are potentially implicated in these pathways. This may be achieved through the employment of phosphoprotemic techniques on isolated adipocytes from AdPKCεKO and WT mice. Subsequently, this will lead to the development of therapeutics designed to mimic the deletion of PKCε in adipose tissue, which will reduce the induction of insulin resistance during dietary lipid oversupply.

A link between lipid oversupply and insulin resistance has long been recognised [23]. As such, both dietary and exercise interventions have been proven to successfully treat insulin resistance [313] [314]. However, the long-term viability of these treatment strategies is low on account of patient incompliance, leading to a continued rise in the incidence of insulin resistance. This has highlighted the need for the development of pharmacological interventions which effectively target the pathways associated with the development and propagation of lipid-induced insulin resistance. In this thesis, we have identified and elucidated novel ways through which two common bioactive lipid metabolites, ceramide and DAG, exert their actions upon glucose and lipid metabolism

166 – processes which become dysregulated during insulin resistance. Through demonstrating the impact of these metabolites and associated enzymes upon the metabolic state of different organs involved in the development of insulin resistance, we have opened new avenues for therapeutic approaches.

167 CHAPTER 7 APPENDICES

168 Section A – Supplementary Data

LacZ CerS1 LacZ CerS2 LacZ CerS4

50 kDa

37 kDa

25 kDa

LacZ CerS5 LacZ CerS6

50 kDa 37 kDa 37 kDa

25 kDa

Figure A-1 Expression of endogenous and recombinant CerS isoforms in L6 myotubes. L6 myotubes were infected with increasing doses of recombinant adenovirus (shown as volume of viral stock in µl) facilitating the expression of CerS1, 2, 4, 5 and 6 or β- galactosidase (LacZ) as control. Cells were lysed and harvested for protein, which was subsequently quantified and immunoblotted with antibodies targeted to haemagglutinin (HA) (for the detection of CerS1, 2, 4 and 5) or CerS6. Figure adapted from [131].

169

a 150

100

(% (% control) 50

0 Phosphorylated / Akt Total Akt 0 5 0 5 0 5 0 5 GW4869 Dose (µM) 20 20 20 20

10 10 10 10 - - + + Palmitate - + - + Insulin b 150

100

(% (% control) 50

0 AmitryptyllineAmitriptyline Dose Dose (µM) (µ M) Phosphorylated / Akt Total Akt 0 0 0 0 10 20 50 10 20 50 10 20 50 10 20 50 - - + + Palmitate - + - + Insulin

Figure A-2 The effect of acidic and neutral sphingomyelinase inhibitors on insulin signalling in L6 myotubes.

Cells were treated with vehicle (shown as 0µM dose. GW4869 vehicle = DMSO, amitriptyline vehicle = H2O) or increasing doses of a) neutral sphingomyelinase inhibitor (GW4869), b) acidic sphingomyelinase inhibitor (amitriptyline) in the presence or absence of palmitate as indicated. Cells were subsequently stimulated with insulin as indicated, lysed and harvested for protein. Protein extracts were immunoblotted with antibodies targeted to phosphorylated-Akt (Ser473), total Akt (all isoforms) and β-actin. Phosphorylated-Akt and total Akt densitometry measurements were each normalised to β-actin prior to normalisation of phosphorylated-Akt with total-Akt. Results are presented relative to control values (% vehicle-treated, non- palmitate treated, insulin-stimulated cells) and are the means from three independent experiments carried out in duplicate.

170

WT KO WT KO

PKCε

Myosin

Figure A-3 Protein expression of PKCε in WT-Floxed and AdPKCεKO mice fed a 16-week HFD. Protein lysates harvested from the adipose tissue of AdPKCεKO and WT-Floxed mice fed a 16-week HFD were immunoblotted with an antibody targeted to PKCε. Non- muscle myosin was employed as a loading control.

a 10 b 10 8 8

6 6

4 4 Fat Pad Mass Mass Pad Fat Fat Pad Mass Mass Pad Fat (% (% Body Weight) 2 (% Body Weight) 2

0 0 WT - Cre WT - Floxed AdPKCεKO WT - Cre WT - Floxed AdPKCεKO

Figure A-4 Gonadal adipose tissue weights from mice fed a 16-week HFD. AdPKCεKO, WT-Cre and WT-Floxed mice were fed a HFD for 16 weeks, fasted for six hours and harvested for gonadal adipose tissue. Gonadal adipose tissue weight was recorded as the combined weight of both gonadal adipose depots harvested and is expressed as percentage mouse body weight recorded immediately prior to tissue harvest. a) Females b) Males. AdPKCεKO n=13 (f) n=8 (m), WT-Cre n=5 (f) n=9(m), WT-Floxed n=12(f) n=12(m).

171 Section B – Buffer and Diet Compositions

Antibody Dilution Buffer (1X solution) Bovine serum albumin (BSA) 5% (w/v) Phenol red 0.05% (v/v) Sodium azide 0.02% (v/v) 1X TTBS up to 500mL

High Fat Diet Hugo’s Copha Diet 2X stock (see below) All Safflower Oil 68mL AIN Vitamin Mix 29.6g Allowrie Lard (Melted) 500g

Hugo’s Copha Diet 2X stock (prepared by Garvan Institute Media Preparation Facility) Caesin 522g Sucrose 460g Starch (corn flour) 386g Homemade mineral mix (see below) 102g Trace minerals 29.6g Bran 114g Methionine 6.8g Gelatine 46g Choline Bitartate 9.2g

Homemade Mineral Mix (prepared by Garvan Institute Media Preparation Facility) Mixture 1 Sodium Selenate 0.28g (0.14g if anhydrous) Potassium Iodate 0.1g Starch Up to 10g Mixture 2 Mixture 1 1g Chromic Potassium Sulphate (ground) 0.55g Homemade Mineral Mix Manganese Carbonate (ground) 0.63g Iron Sulphate Heptahydrate (ground) 4.98g Zinc Carbonate 1.6g Copper Carbonate 0.3g Mixture 2 All Starch 133.9g Mix thoroughly then add the following Calcium Carbonate 356.71g Potassium Dihydrogen Orthophosphate 402.09g Sodium Chloride 74g Magnesium Oxide 24g

172 Krebs Ringer Buffer (1X solution) NaCl 1.3M CaCl2 21mM MgSO4 12.5mM KH2PO4 40mM HEPES 100mM D-(+)-Glucose 1.8mg/ml BSA 2% (w/v) H2O (Baxter water) up to 100mL

MOPS ruuning buffer (20X stock) MOPS 1M Tris 1M SDS 70mM EDTA 20mM dH2O up to 1L

RIPA buffer (1X solution) Trizma base 65mM NaCl 150mM Add dH2O to 180mL Adjust pH to 7.4 EDTA 5mM Igepal CA630 1% (v/v) Sodium deoxycholate 0.5% (w/v) SDS 0.1% (v/v) Glycerol 10% (v/v) NaF 10mM Sodium pyrophosphate 10mM dH2O up to 250mL Immediately prior to use add: Sodium orthovanadate, 1mM PMSF 2mM Protease inhibitor cocktail 1X

Transfer buffer (1X solution) Tris 25mM Glycine 192mM Methanol 10% (v/v) SDS 0.025% (v/v) dH2O up to 20L

Tris Buffered Saline (1X solution) Tris 10mM NaCl 150mM dH2O up to 1L

173 Tris Buffered Saline with Tween (1X solution) Tris 10mM NaCl 150mM Tween-20 0.05% (v/v) dH2O up to 20L

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