ASSESSING THE ROLE OF GLYCERONEOGENESIS IN

TRIGLYCERIDE METABOLISM

By

COLLEEN KLOCEK NYE

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Thesis Advisors:

Richard W. Hanson, Ph.D.

Satish C. Kalhan, M.D.

Department of Biochemistry

CASE WESTERN RESERVE UNIVERSITY

August, 2008

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Colleen Klocek Nye

Candidate for the Doctor of Philosophy degree*.

(signed) William L. Merrick, Ph.D. (chair of the committee)

Satish C. Kalhan, M.D.

Richard W. Hanson, Ph.D.

Martin D. Snider, Ph.D.

Arthur J. McCullough, Ph.D.

(date) June 18, 2008

*We also certify that written approval has been obtained for any proprietary material contained therein.

ii TABLE OF CONTENTS

LIST OF FIGURES…………………………………………………………………...... vii

LIST OF TABLES…………………………………………………………………...... viii

ACKNOWLEDGEMENTS………………………………………………………….....ix

LIST OF ABBREVIATIONS…………………………………………………………..x

ABSTRACT………………………………………………………………………….....1

CHAPTER 1: Introduction…………………………………………………………..3

Obesity, insulin resistance, and Type II diabetes……………………………………3

Triglyceride synthesis and metabolic sources of triglyceride glycerol……………..4

Glycerol kinase……………………………………………………………………….....4

Glucose uptake by the liver and peripheral tissues……………………………………..7

Glyceroneogenesis and PEPCK-C……………………………………………………..8

Discovery and significance of glyceroneogenesis under physiological

conditions………………………………………………………………………9

Discovery…………………………………………………………………………….....9

TG-FA cycling……………………………………………………………………….....9

Postprandial state………………………………………………………………………13

Postabsorptive state………………………………………………………………….....17

Response to pharmaceutical intervention: treatment of Type 2 diabetes

with thiazolidinediones………………………………………………………...18

Physiological role of glyceroneogenesis in other species…………………………...21

Rainbow smelt (Osmerus mordax)…………………………………………………….21

Parasitic helminth (Schistosoma mansoni)…………………………………………….22

iii Drosophila...... 23

Nematode worm (Caenorhabditis elegans)……………………………………………23

Consequence of ablation and overexpression of PEPCK-C:

transgenic mouse models……………………………………………………..24

PEPCK-C knockout in the liver………………………………………………………..24

Role of PEPCK-C in cataplerosis……………………………………………………...24

PEPCK-C knockout in white adipose tissue…………………………………………...26

Overexpresssion of PEPCK-C in white adipose tissue………………………………...27

Overexpression of PEPCK-C in skeletal muscle………………………………………29

Glyceroneogenesis in vivo…………………………………………………………….31

Statement of purpose…………………………………………………………………35

Hypothesis…………………………………………………………………………….36

CHAPTER 2: Experimental Procedures…………………………………………..37

Animals………………………………………………………………………………...37

Tracer study protocol…………………………………………………………………..38

Rationale for selection of dietary groups………………………………………………38

Rationale for design of tracer studies employing infusion of epinephrine…………….43

Triglyceride extraction and isolation of glycerol and fatty acids……………………...43

Isolation of plasma metabolites………………………………………………………..44

Measurements of radioactivity………………………………………………………...44

Sources of carbon and hydrogen in triglyceride glycerol……………………………...45

Appearance of [14C] label in the triose phosphate pool………………………………..45

Source of [3H] label of C-1 and C-3 of the triose phosphate pool……………………..48

iv Ratio of [14C]/[3H] of C-1 and C-3 of triglyceride glycerol……………………………48

Radioactivity of C-1 and (C-1 + C-3)…………………………………………………48

Phosphorylation of glycerol…………………………………………………………...49

Activity of PEPCK-C………………………………………………………………….49

Calculations……………………………………………………………………………49

Statistical analysis……………………………………………………………………..52

CHAPTER 3: Results……………………………………………………………….53

Glucose kinetics, gluconeogenesis, and the source of plasma

triglyceride glycerol…………………………………………………………...53

Glyceroneogenesis and glycolysis as a source of triglyceride

glycerol in adipose tissue………………………………………………………56

Fatty acid synthesis in the adipose tissue……………………………………………...59

PEPCK-C activity in the adipose tissue……………………………………………….59

Response to enhanced TG-FA cycling by epinephrine………………………………..66

The relative contribution of glyceroneogenesis and glucose to

triglyceride glycerol synthesis in skeletal muscle……………………………..66

CHAPTER 4: Discussion……………………………………………………………72

Significance of current methodology………………………………………………….72

Glyceroneogenesis in the adipose tissue………………………………………………74

Response to epinephrine infusion……………………………………………………..78

A computational model for adipose tissue metabolism……………………………….79

Glucose uptake and glyceroneogenesis by the adipose tissue:

A two-cell hypothesis………………………………………………………….80

v PEPCK-C activity in adipose tissue…………………………………………………...84

Glyceroneogenesis in the skeletal muscle……………………………………………..86

Hepatic glyceroneogenesis…………………………………………………………….88

Summary and remaining questions……………………………………………………90

What is the source of carbon for glyceroneogenesis?...... 90

What is the source of reducing equivalents for glyceroneogenesis?...... 91

REFERENCES………………………………………………………………………...92

vi LIST OF FIGURES

Figure 1. The pathway of glyceroneogenesis………………………………………..5

Figure 2. The triglyceride-fatty acid cycle…………………………………………..10

Figure 3. Tracer study protocol……………………………………………………...39

Figure 4. The incorporation of [3H] into G-3-P from bodywater……………………41

Figure 5. Relative contribution of glyceroneogenesis, direct and

indirect (via lactate) pathways of glucose, and [14C]/[3H]

ratio of C-1 and C-3 of triglyceride glycerol ………………………………...46

Figure 6. The activity of PEPCK-C in adipose tissue………………………………64

Figure 7. The two compartment (or two cell-type) hypothesis……………………..81

vii LIST OF TABLES

Table 1. Glucose kinetics, gluconeogenesis, and source of plasma

triglyeride glycerol…………………………………………………………..54

Table 2. The contribution of glyceroneogenesis and glycolysis to

triglyceride glycerol synthesis in adipose tissue……………………………..57

Table 3. Relative [14C] and [3H] radioactivity and [14C]/[3H] ratio

on C-1 and C-1 + C-3 of triglyceride glycerol in adipose tissue…………….60

Table 4. Incorporation of glucose carbon into de novo synthesized

fatty acids in adipose tissue…………………………………………………..62

Table 5. The effect of epinephrine infusion on the contribution of

glyceroneogenesis and glycolysis to triglyceride glycerol

synthesis in adipose tissue……………………………………………………67

Table 6. The contribution of glyceroneogenesis and glycolysis to

triglyceride glycerol synthesis in skeletal muscle……………………………69

viii ACKNOWLEDGEMENTS

This thesis is lovingly dedicated to my husband, Shad, my children, Austin, Skye and Baby Nye, and most especially to my mother, without whom I could not have accomplished this goal.

I want to thank my mentor, Satish Kalhan, for what has undoubtedly been the greatest educational experience of my life. You are brilliant and my life is better for having known you.

I want to thank my thesis advisor, Richard Hanson, for being my inspiration since the very first day I attended his lecture series in metabolism. You are nothing less than a star amongst stars. Thank you for your patience, kindness, advice and profound scientific

knowledge.

I was so fortunate to have had the unique opportunity to work with world

reknown experts in metabolism and biochemistry. What I will miss most is meeting with

both of you to discuss science. However, I equally enjoyed listening to your dialogue of

diverse scientific topics, and of life in general.

I want to thank the members of the Kalhan Lab who became my good friends and my support network: Clarita, Lourdes, Ed and Jose. Thank you so much for all of your help, guidance, and most importantly your friendship.

I also want to thank all the members of the Hanson Lab: it was a joy to have worked with you. I especially want to thank Parvin for her hard work and dedication, as well as her friendship and advice. Thank you, Jianqi, for your kind words and your vast scientific expertise.

ix LIST OF ABBREVIATIONS

3 [ H2]O Tritium-Labeled Water

U-[14C]glucose Uniformly Labeled [14C]Glucose

FFA Free Fatty Acids

TG-FA cycling Triglyceride-Fatty Acid Cycling

G-3-P Glycerol-3-Phosphate

CO2 Carbon Dioxide

OAA Oxaloacetate

PEPCK-C Phosphoenolpyruvate Carboxykinase-Cytosolic Form

PEP Phosphoenolpyruvate

GAP Glyceraldehyde-3-Phosphate

DHAP Dihydroxyacetonephosphate

FA CoA Fatty Acyl CoA

IMGU Insulin Mediated Glucose Uptake

NIMGU Non-Insulin Mediated Glucose Uptake

Rd Rate of Disposal

GLUT4 Glucose Transporter 4 a-v Arterio-Venous

TCA cycle Tricarboxylic Acid Cycle

NAD Nicotinamide Adenine Dinucleotide

GTP Guanosine Triphosphate

VLDL Very Low-Density Lipoprotein

NADH Nicotinamide Adenine Dinucleotide-Reduced Form

x IMTG Intramuscular Triglyceride

HSL Hormone Sensitive Lipase

LPL Lipoprotein Lipase

Ra Rate of Appearance

TZDs Thiazolidinediones

PPARγ Peroxisome Proliferator Activated Receptor Gamma

RXR 9-cis Retinoic Acid-Activated Retinoid Receptor

PPRE Peroxisome Proliferator Activated Receptor Response Element

aP2 Adipose-Specific Fatty Acid Binding Protein

mRNA Messenger Ribonucleic Acid

dATF-2 Drosophila Activating Transcription Factor-2

AMP Adenosine Monophosphate

CREB Cyclic AMP Response Element Binding Protein

S. mansoni Schistosoma mansoni

C. elegans Caenorhabditis elegans

Cre/loxP Cre and LoxP Recombination Sites

redox Reduction-Oxidation

ATP Adenosine Triphosphate

cDNA Complimentary Deoxyribonucleic Acid

VO2max Maximal Oxygen Consumption

2 [ H2]O Deuterium-Labeled Water

HPLC High Performance Liquid Chromatography

C-1 Carbon-1

xi C-3 Carbon-3

I Rate of Infusion

SA Specific Activity

Glc Glucose

GNG Gluconeogenesis

TG Triglyceride dpm Disintegrations Per Minute

BMI Body Mass Index

HOMA Homeostatic Model Assessment cAMP Cyclic Adenosine Monophosphate

xii Assessing the Role of Glyceroneogenesis in

Triglyceride Metabolism

Abstract

By

COLLEEN KLOCEK NYE

The formation of triglyceride in mammalian tissues requires the provision of glycerol-3-phosphate as the source of triglyceride glycerol. We have quantified the

relative contribution of glyceroneogenesis and glycolysis to triglyceride glycerol

synthesis in vivo in adipose tissue, skeletal muscle, and liver of the rat in response to a

chow diet (controls), 48 h of fasting, and lipogenic (high sucrose) diet. The rate of

3 glyceroneogenesis was quantified using the tritium ([ H2]O) labeling of body water

method, and the contribution of glucose, via glycolysis, was determined using [U-

14C]glucose tracer. In epididymal and mesenteric fat of control rats, glyceroneogenesis

accounted for ~90% of triglyceride glycerol synthesis. Fasting for 48 h did not alter

glyceroneogenesis in adipose tissue, whereas the contribution of glucose was negligible.

In response to sucrose feeding, the synthesis of triglyceride glycerol via both

glyceroneogenesis and glycolysis nearly doubled (vs. controls); however,

glyceroneogenesis remained quantitatively higher as compared to the contribution of

glucose. Enhancement of triglyceride-fatty acid cycling by epinephrine infusion resulted

in a higher rate of glyceroneogenesis in adipose tissue as compared with controls, while

the contribution of glucose, via glycolysis, was not measurable. Glyceroneogenesis

1 provided the majority of triglyceride glycerol in the gastrocnemius and soleus muscle. In the liver, the fractional contribution of glyceroneogenesis remained constant (~60 %) under all conditions and was higher than that of glucose. Thus, glyceroneogenesis, in contrast to glucose, via glycolysis, is quantitatively the predominant source of triglyceride glycerol in adipose tissue, skeletal muscle, and liver of the rat under conditions ranging from extended fasting to high sucrose feeding.

2 CHAPTER 1: Introduction

Obesity, insulin resistance, and Type 2 diabetes

Obesity in the United States and abroad has reached epidemic proportions and will only continue to increase as many cultures continue to adopt a westernized lifestyle of overindulgence in food accompanied by a decrease in exercise. Obesity is the result of long-term positive energy balance, when more energy is consumed than is required to meet the physiological demands of the individual. Obesity is highly correlated with insulin resistance, which is the most prominent metabolic defect of Type 2 diabetes. This is especially true when the adiposity is present intra-abdominally or viscerally (1). In obese subjects with insulin resistance, there is a mass effect of the adipose tissue, with increased levels of free fatty acids (FFA) being released from the enlarged total fat depot compared to lean individuals (2). In addition, insulin resistance involves an impaired suppression of lipolysis within the fat cells, in particular with upper-body obese subjects

(2). Furthermore, the ability of adipose tissue to recapture FFA by re-esterification is decreased in insulin resistant conditions (2). Elevated levels of FFA in the blood impair insulin-mediated glucose uptake by target tissues (i.e., skeletal muscle, adipose tissue) thus contributing to peripheral insulin resistance. In addition, the increased metabolic activity of visceral adipose tissue results in an increase in FFA delivered directly to the portal circulation contributing to hepatic insulin resistance, as evidenced by an increased rate of production of glucose, due to gluconeogenesis and glycogenolysis (2). The high portal FFA concentration further adds to the dyslipidemia observed with insulin resistance by providing substrate to the liver for the increased synthesis of triglycerides.

As accretion of fat leads to obesity, which may give rise to perturbations of lipid

3 metabolism; it is important to understand the role of triglyceride synthesis under different

physiological settings.

Triglyceride synthesis and metabolic sources of triglyceride glycerol

Triglyceride synthesis is critical for the accretion of fat in vivo and for the transport of lipids in the blood. In addition, triglyceride synthesis is an essential component of triglyceride-fatty acid (TG-FA) cycling (discussed below), in which fatty acids released from the adipose tissue following lipolysis, are re-esterified back to triglyceride in what is termed a “futile cycle.” Glycerol-3-phosphate (G-3-P), along with fatty acyl CoA, are the substrates required for the synthesis of triglycerides. G-3-P can be synthesized by phosphorylation of glycerol via glycerol kinase or by the reduction of dihydroxyacetone phosphate (DHAP) catalyzed by cytosolic glycerol-3-phosphate dehydrogenase. The metabolic sources of DHAP are glucose via glycolysis or pyruvate via glyceroneogenesis. Glyceroneogenesis may be defined as the de novo synthesis of G-

3-P from precursors other than glucose or glycerol (Fig. 1).

Glycerol kinase Glycerol kinase is present in significant quantities in the liver of all mammals. The activity of glycerol kinase in rat liver is ~2 units/g in fed animals compared to 0.62 units/g in humans (3;4). A decrease in glycerol kinase activity has been observed in response to fasting, as evidenced by values of 1.43 units/g and 1.26 units/g in animals fasted overnight and for 24 h, respectively (5-7). In contrast, glycerol kinase activity in skeletal muscle of rats and humans is extremely low, with values of

0.007 units/g and 0.009 units/g being reported for rat and human, respectively (8).

4 GAP Glycerol-3-PGlycerol-3-P + 3 FA CoA Triglyceride

Lactate

Alanine

PEPCK-C 5 Figure 1. The pathway of glyceroneogenesis. Glyceroneogenesis is defined as the de novo synthesis of G-3-P from

precursors other than glucose or glycerol, namely pyruvate, lactate, alanine, or any other (non-ketogenic) amino acid that

enters into the TCA cycle. Carbon flow of this pathway begins with pyruvate entering the mitochondria and proceeding to

oxaloacetate (OAA) via pyruvate carboxylase, OAA goes to malate which is transported to the cytosol where it is converted

back to OAA. The key regulatory enzyme in this pathway, the cytosolic form of phosphoenolpyruvate carboxykinase (GTP)

(PEPCK-C) (EC 4.1.1.32), catalyzes the GTP-dependent decarboxylation of OAA to form phosphoenolpyruvate (PEP). PEP is

metabolized to the level of the triose phosphate pool (glyceraldehye-3-phosphate and DHAP) where DHAP is reduced to G-3-

P via glycerol-3-phosphate dehydrogenase and NADH. 6 Glycerol kinase activity in adipose tissue of rats and humans is negligable (~0.002

units/g) (5;9;10).

Glucose uptake by the liver and peripheral tissues Intravenously administered

glucose is taken up by the splanchnic bed to a very limited extent, even under conditions

of hyperinsulinemia and hyperglycemia (11). Furthermore, in response to oral glucose,

Katz et al. (12) showed that 75% of the oral glucose escaped splanchnic removal in healthy human volunteers, and the peripheral tissues quantitatively play the dominant role in glucose disposal. Glucose uptake can occur through insulin mediated glucose uptake

(IMGU) or non-insulin mediated glucose uptake (NIMGU). With respect to IMGU, in the presence of ~70 µU/ml of insulin under euglycemic conditions, ~75% of whole body rate of disposal (Rd) of glucose is in skeletal muscle (12;13). In the presence of basal insulin levels (~6 µU/ml), ~20% of whole body Rd of glucose occurs in skeletal muscle;

~10% is IMGU, while ~10% is NIMGU (13). The majority of glucose disposal under basal conditions is NIMGU which occurs in the central nervous system (13).

In response to an oral glucose load, adipose tissue is responsible for ~5% of whole body Rd of glucose (14;15). Similar to what is observed in the skeletal muscle, insulin mediated GLUT4 translocation and glucose uptake into adipose tissue increases in response to glucose (16). In contrast, glucose uptake by the adipose tissue from rats decreases from 115 nmol/g tissue/min (fed) to 60 nmol/g tissue/min in response to fasting for 48 h, as determined by arterio-venous (a-v) difference techniques (17). Furthermore, there is a lower expression of GLUT4 (18;19) and a decreased uptake and phosphorylation of 2-deoxyglucose by isolated adipocytes from fasted rats (20).

7 Glyceroneogenesis and PEPCK-C Glyceroneogenesis is defined as the de novo

synthesis of G-3-P from precursors other than glucose and glycerol, namely pyruvate,

lactate, alanine, or any other (non-ketogenic) amino acid that can be converted to an

intermediate of the tricarboxylic acid (TCA) cycle. Pyruvate enters the mitochondria and proceeds to oxaloacetate (OAA) via pyruvate carboxylase, OAA is reduced to malate which is transported to the cytosol where it is converted back to OAA via NAD malate dehydrogenase. The key enzyme in this pathway, the cytosolic form of phosphoenolpyruvate carboxykinase (GTP) (PEPCK-C) (EC 4.1.1.32), catalyzes the

GTP-dependent decarboxylation of OAA to form phosphoenolpyruvate (PEP). PEP is metabolized to the level of triose phosphate (glyceraldehye-3-phosphate and DHAP) where DHAP is converted to G-3-P (Fig. 1).

PEPCK-C is present in significant quantities in the liver. In response to feeding,

PEPCK-C activity is ~2.1-2.5 units/g and increases to ~6.7-7.4 units/g after a 24 h fast

(21;22). In contrast, PEPCK-C is present in limited quantities in the skeletal muscle, where the activity in mice is reported to be ~0.08 units/g (23). Although Newsholme et al. (24) have reported an increase in PEPCK-C activity in skeletal muscle after a 48 h

fast, the regulation of PEPCK-C in skeletal muscle in response to hormone and nutrient

treatment has not been well established. In adipose tissue of the rat, PEPCK-C activity

has been reported to be ~0.040 units/g under postprandial conditions (21;22;25) and

~0.020 units/g in response to a sucrose supplemented diet followed by intravenous

glucose infusion (26). PEPCK-C activity increases in adipose tissue in response to

fasting, and was reported to be ~0.18-0.28 after a 24 h fast (21;22;25).

8 Discovery and significance of glyceroneogenesis under physiological conditions

Discovery Glyceroneogenesis was first described in vitro in white adipose tissue

40 years ago (27;28). The discovery of the key gluconeogenic enzyme, PEPCK-C, in this non-gluconeogenic tissue led investigators to examine its function in the adipose tissue.

Initial studies by Reshef, Ballard, Hanson and Leveille, demonstrated that the addition of pyruvate to rat epididymal adipose tissue, incubated in vitro, reduced FFA release by

65%, while not altering lipolysis (as determined by the amount of glycerol released into the medium) because of increased FFA re-esterification (21;29;30). These findings suggested that the pathway of glyceroneogenesis, named by Gorin and Shafrir in 1970

(31) played a physiological role in the TG-FA cycle (30).

TG-FA cycling TG-FA cycling is the re-esterification of FFA, which are released in excess, back to triglyceride during enhanced states of lipolysis such as fasting or exercise (Fig. 2). The hydrolysis of triglyceride in adipose tissue results in the production of glycerol and FFA. The FFA released upon lipolysis are either re-esterified within adipocytes, without leaving the adipose tissue (local/ intracellular TG-FA cycling), or are

released into the blood and re-esterified back to triglyceride in other tissues

(systemic/extracellular TG-FA cycling); these include, peripheral tissues (i.e., skeletal

muscle and adipose tissue) and the liver (112). In response to an overnight fast, local

TG-FA cycling in adipose tissue represents approximately 20-30% of the total, whereas

systemic recycling accounts for ~50% of TG-FA cycling (113). Since adipose tissue

lacks appreciable levels of glycerol kinase, the glycerol released upon hydrolysis cannot

be utilized for local TG-FA cycling, and therefore enters into the blood. In addition,

9 Ketone bodies VLDL G-3-P

PEPCK-C 10 Figure 2. The triglyceride-fatty acid cycle. During fasting, FFA released upon lipolysis are re-esterified within adipocytes,

without leaving the adipose tissue (local/intracellular recycling), or are released into the blood. Since adipose tissue lacks

glycerol kinase, and glucose uptake during fasting is significantly reduced, glyceroneogenesis may provide the majority of G-

3-P for local recycling. FFA which are released into the blood may be completely oxidized to CO2 by peripheral tissues,

converted to ketone bodies in the liver, or recycled (re-esterified) back into triglyceride in the liver (systemic/extracellular

recycling), with subsequent release as VLDL. However, only a small fraction of FFA released from adipose tissue are actually

oxidized, with the majority being recycled back into triglyceride in various tissues, namely the liver. Although the liver

possesses significant glycerol kinase activity, recent studies suggest that hepatic glyceroneogenesis is the predominant pathway

for G-3-P synthesis.

11 glucose uptake by the adipose tissue is significantly decreased in response to fasting.

Thus, the glyceroneogenic pathway was suggested to provide the majority of G-3-P necessary for re-esterification in adipose tissue during fasting (21;29;30;32).

The majority of systemic TG-FA cycling occurs in the liver. Although the liver contains glycerol kinase and can therefore utilize glycerol as a source for G-3-P, Kalhan et al. (33) have demonstrated that 10-60% of the very low-density lipoprotein (VLDL) triglyceride glycerol synthesized for systemic cycling of FFA is provided by glyceroneogenesis, whereas plasma glycerol contributes only ~6% in overnight fasting humans. Since the flux of triosephosphate to glucose (~10 µmol/kg/min) in the liver exceeds that to triglyceride glycerol (<1 µmol/kg/min), more glycerol is ultimately converted to glucose than to triglyceride glycerol (33). Similarly, since the contribution of pyruvate to the triosephosphate pool is far greater than that of plasma glycerol, the conversion of pyruvate to triglyceride glycerol will be much greater than that of glycerol

(33). Research by Siler et al. (34) supports the vital role of cytosolic redox state in regulation of the diversion of triosephosphate from gluconeogenesis to glyceroneogenesis. The reduction of DHAP to G-3-P is a redox-regulated step which requires the oxidation of NADH. Ethanol greatly enhanced the rate of triglyceride glycerol synthesis, via glyceroneogenesis, due to a decrease in the cytosolic NAD/NADH ratio in the liver that was induced by the catabolism of ethanol.

Skeletal muscle has been suggested to be a (possible) site of systemic TG-FA cycling. However, only local TG-FA cycling has been documented in oxidative muscle of overnight fasted rats (35). In humans, the presence of local TG-FA cycling has been evidenced by nearly equimolar rates of glycerol and FFA release from skeletal muscle

12 after an overnight fast (36). Furthermore, in response to treatment with a ß-adrenonergic

receptor agonist (isoprenaline), a concentration-dependent stimulation of glycerol release

was observed, whereas FFA release was not affected (36). Local TG-FA cycling has

been estimated to be one-twentieth of that found in adipose tissue, since less triglyceride

is present in skeletal muscle as lipid droplets deposited within muscle fibers (35). The physiological role of the intramyocellular triglyceride (IMTG) pool may be to provide a substrate source during exercise since well-trained endurance athletes exhibit increased

levels of IMTG. Yet insulin resistant states, such as Type 2 diabetes, are also associated with IMTG accumulation. Skeletal muscle contains glycerol kinase and PEPCK-C, although the activity of PEPCK-C in this tissue is approximately 10-fold higher than

glycerol kinase. Overexpression of PEPCK-C in skeletal muscle of the mouse resulted in

a marked increase in IMTG levels (23). Furthermore, studies by Jenson et al. (37)

suggest that even though glucose and glycerol contribute equally to triglyceride glycerol

synthesis in gastrocnemius of 24 h fasted rats, the majority of G-3-P is derived from what

they termed “indirect glycerogenesis,” (i.e. glyceroneogenesis) which originated from

steps further down the glycolytic pathway.

Postprandial state In response to a meal containing carbohydrate, the

concentration of blood glucose, and subsequently insulin, increases, resulting in enhanced

glucose uptake into insulin sensitive tissues. The activity of hormone sensitive lipase

(HSL), responsible for hydrolysis of triglyceride stored within adipocytes and subsequent

release of glycerol and FFA into the blood is decreased dramatically, whereas the activity

of lipoprotein lipase (LPL), which is stimulated by insulin, increases after a meal. LPL

hydrolyzes triglyceride contained within circulating chylomicrons. The FFA liberated

13 from this dietary lipid can then be esterifed back to triglyceride and deposited in

peripheral tissues. The portion of FFA released upon hydrolysis, which exceeds tissue

uptake, enters the plasma FFA pool and is cleared by the liver in a manner similar to FFA

derived from adipose tissue lipolysis (38). Thus, the dietary fatty acids that escape

deposition may be packaged into lipoprotein particles and secreted from the liver as

VLDL. Furthermore, insulin responsive pathways such as glycolysis, pentose phosphate

shunt, glycogen synthesis and lipogenesis are up-regulated after carbohydrate feeding.

Consequently, fatty acids synthesized de novo from glucose in the liver are exported as

VLDL and also deposited as triglyceride in the periphery. Thus, in the postprandial state, a source of G-3-P is required for esterification of dietary and de novo synthesized fatty acids, resulting in net deposition of triglyceride. Although glucose, via glycolysis, has historically been considered to be the major carbon source for synthesis of G-3-P in the white adipose tissue, skeletal muscle and liver (39-41), the pathway of glyceroneogenesis

must also be considered.

The majority of de novo fatty acid synthesis occurs in the liver of healthy humans

in response to refeeding with intravenous glucose or to a high carbohydrate mixed-meal

breakfast after an overnight fast (42). However, the estimated total fatty acid synthesis is

less than 500 mg/day as determined in vivo by stable isotope techniques (42). Although

the fractional contribution of hepatic de novo lipogenesis increases 6- to 10-fold in

response to excess carbohydrate feeding, the absolute rate remains quantitatively low in

healthy individuals (43). Unlike humans, de novo lipogenesis has been documented to

occur at high rates in adipose tissue of young rodents in response to normal feeding

(44;45). However, when Aarsland et al. (46) investigated whole-body net fat synthesis in

14 vivo in humans in response to 4 d of hyper-energetic carbohydrate feeding, they demonstrated that adipose tissue was the predominant site of de novo lipogenesis, even though the hepatic secretion of fat synthesized de novo increased ~35-fold during the study. While de novo lipogenesis in adipose tissue was shown to be active in humans, quantitative estimates by Diraison et al. (47) suggest that de novo lipogenesis by adipose tissue in vivo is minimal (<1 g) after acute or prolonged carbohydrate feeding. Thus, de novo lipogenesis in adipose tissue of humans can be induced under specific dietary conditions although it remains quantitatively negligible in terms of absolute lipogenesis.

Since glucose uptake and glucose responsive pathways are up-regulated in response to a meal containing carbohydrate, it only seems reasonable that glucose, via glycolysis, would provide the G-3-P required for triglyceride formation. This is in contrast to the in vivo studies performed by the Migliorini laboratory (48;49) which clearly demonstrate that in the adipose tissue, G-3-P is synthesized via the glyceroneogenic pathway in response to a high protein, carbohydrate free diet. This suggests that amino acids provide the triglyceride glycerol necessary for esterification of diet derived fatty acids under these conditions (48;49).

Although dietary and de novo synthesized fatty acids account for the majority of the fatty acids that contribute to net triglyceride synthesis in the postprandial state, FFA released during TG-FA cycling must also be considered. Since the importance of TG-FA cycling is emphasized during states of enhanced lipolysis, the occurrence of TG-FA cycling in the postprandial state is seldom highlighted. TG-FA cycling persists in the postprandial state, however, the contribution of systemic TG-FA cycling is decreased while intracellular TG-FA cycling is enhanced (50;51). In response to a meal containing

15 carbohydrate, increased levels of insulin are responsible for decreased lipolysis and

systemic TG-FA cycling, and an increase in intracellular fatty acid re-esterification, requiring a local source of G-3-P. Studies with humans have demonstrated a significant decrease in the rate of appearance (Ra) of glycerol, compared to the basal state, in response to infusion of glucose at 4 mg/kg/min, thus indicating suppression of lipolysis

(50). However, lipolysis is not completely inhibited, as evidenced by the persistent Ra

(release) of glycerol. Furthermore, the Ra of FFA decreased to a greater extent than the

Ra of glycerol, indicating re-esterification (or intracellular TG-FA cycling) was

stimulated in response to glucose infusion (50). In the same study, when exogenous

glucose infusion was increased to 8 mg/kg/min, the Ra of glycerol remained unchanged,

while the Ra of FFA decreased even more. The further increase in re-esterification in

response to higher levels of glucose infusion was attributed to the observed increase in

the concentration of insulin, which presumably increased glucose uptake into the adipocyte for the formation of G-3-P (50). The effect of insulin on FFA esterification in adipose tissue has often been attributed to the stimulation of glucose uptake and

consequent G-3-P production. However, Frayn et al. (52) showed that insulin infusion

(40 mU/ml/min) administered for 4.5 h after a meal high in carbohydrate and fat, resulted

in no further increase in glucose uptake (Frayn et al.). These studies also demonstrated that additional insulin significantly decreased plasma concentrations of FFA and had no

effect on glycerol release from the adipose tissue in the postprandial state (insulin clamp

versus unclamped) (52). Taking into account that exogenous insulin promoted no further

change in glucose uptake or lipolysis, while re-esterification was enhanced, this suggests

16 that the source of triglyceride glycerol for intracellular TG-FA cycling may be provided by the glyceroneogenic pathway.

Although the majority of triglyceride synthesis and subsequent storage takes place in the adipose tissue, skeletal muscle is also a site of net triglyceride deposition. In response to a meal containing carbohydrate, fatty acid oxidation by the skeletal muscle is decreased while glucose oxidation is increased. Similar to the assumption made about adipose tissue, it only seems logical that the source of G-3-P or triglyceride synthesis would be derived from glucose under these conditions. In contrast to what was observed in the adipose tissue, insulin does not appear to impact lipolysis in skeletal muscle, as evidenced by the lack of change in glycerol release in response to an oral glucose load, as well as to glucose and insulin (53). This suggests that triglyceride synthesis in the skeletal muscle supports the maintenance of basal triglyceride levels (54). In addition, the ectopic storage of triglyceride in tissues not designed for such high levels of accumulation (i.e., liver, skeletal muscle) may contribute to the disruption of lipid homeostasis and subsequently cause insulin resistance. Thus, during insulin resistant states, when glucose uptake by the skeletal muscle is decreased and net triglyceride deposition is enhanced, glyceroneogenesis may be responsible for the provision of triglyceride glycerol.

Postabsorptive state In response to an overnight fast, triglyceride formation in adipose tissue does not result in net triglyceride deposition; the majority of triglyceride formation under these circumstances occurs in response to higher levels of TG-FA cycling. Since glucose uptake is decreased is the postabsorptive state, the synthesis of G-

3-P via glyceroneogenesis for re-esterification of FFA becomes increasingly important.

17 Arterio-venous (a-v) differences across human subcutaneous abdominal adipose

tissue after an overnight fast have demonstrated that adipose tissue is strongly lipolytic

(releases FFA and glycerol), clears circulating triglyceride, glucose, ketone bodies and acetate, and produces lactate (55). Adipose tissue after an overnight fast is a net exporter of carbon, mainly as FFA (55).

Reimer et al. (56) used isolated perfused rat hind quarters to determine the uptake and utilization of substrates by skeletal muscle in response to fasting and showed that

skeletal muscle from fasting rats takes up glucose and fatty acids and releases lactate,

alanine, fatty acids, and glycerol. Furthermore, infusion of [1-14C]oleate had no impact on glucose uptake, whether insulin was present or not (56). In response to oleate infusion, 75% of the fatty acids were found in muscle lipids and 10% were oxidized to

CO2 (56).

Later in vivo studies measuring a-v differences across human forearm muscle

demonstrated uptake of FFA when fasting but not after 75 g of glucose ingestion (57).

Fatty acids were shown to be the main source of carbon atoms entering the resting

forearm (55). Similar to the studies performed using isolated perfused rodent muscle,

these data also indicated there was no significant a-v difference for glycerol during

fasting or after consumption of glucose.

Response to pharmaceutical intervention: treatment of Type 2 diabetes with

thiazolidinediones The thiazolidinediones (TZDs) are a class of anti-hyperglycemic

agent used to treat Type 2 diabetes. Two of these TZDs, rosiglitazone and pioglitazone,

have been approved for clinical use in the United States (58). TZDs do not directly affect

insulin secretion, but instead function by improving glycemic control. Insulin resistance

18 in the liver, skeletal muscle, and adipose tissue of rodents and humans is reduced in response to TZD treatment. The primary target of TZDs is the nuclear receptor peroxisome proliferator activated receptor-γ (PPARγ). Although PPARγ is expressed at

low levels in multiple cell types, a high level of expression is observed in adipose-derived

cells (59). PPARγ functions by dimerizing with another nuclear receptor, 9-cis retinoic

acid-activated retinoid receptor, RXR. The heterodimer then binds to a recognition

sequence (the peroxisome proliferator activated receptor response element or PPRE)

located in the promoter region of specific genes. Most genes directly regulated by

PPARγ are involved in lipid metabolism. These genes include: lipoprotein lipase, adipose-specific fatty acid binding protein (aP2), fatty acid synthase, PEPCK-C, and acyl

CoA oxidase (59;60;61).

Improvements of insulin action in liver and skeletal muscle caused by the administration of TZDs may be attributed to two possibilities. There may be a direct activation of low levels of PPARγ expressed in those tissues, or an indirect effect caused

by activating the highly expressed PPARγ present in adipose tissue (58). Until the

effects of TZDs are assessed in liver and muscle of PPARγ knockout mice, there will not

be definitive evidence of a direct insulin-sensitizing action in either of these tissues (58).

The response of adipocytes to insulin differs between the various fat depots.

Adipocytes in intra-abdominal or visceral fat are less responsive to insulin and exhibit increased rates of FFA release. Furthermore, differences also exist in the adipocyte population within a given fat depot. Some adipocytes may be small, newly differentiated and highly responsive to insulin; others may be large, lipid-filled, and insulin resistant.

In insulin resistant states, such as obesity and Type 2 diabetes, fat depots contain a high

19 degree of large, lipid-filled adipocytes (58). In vitro studies show that TZD activation of

PPARγ in fat depots stimulates the insulin-mediated differentiation program which

results in increased numbers of small, insulin-sensitive adipocytes (58). While in large

insulin-resistant fat cells, TZDs oppose inflammatory cytokine production and promote

apoptosis (58), they increase peripheral adiposity, but have no effect on, or significantly

reduce, visceral fat mass (62). Sequestering triglyceride in adipose depots, away from

liver and skeletal muscle is most likely an important insulin-sensitizing action of TZDs

(58).

Recent studies have found that the regulation of glyceroneogenesis occurs mainly

in visceral fat; the same fat depot implicated in the progression of obesity to Type 2 diabetes (63). PEPCK-C is among the several genes involved in fat metabolism that is directly stimulated by TZDs. When rosiglitazone was administered to diabetic patients for 12 wk, PEPCK-C mRNA was increased 9-fold and glyceroneogenesis was induced by

2.5-fold (64). Rosiglitazone also enhanced glyceroneogenesis in adipose tissue explants from human subjects, as determined by the incorporation of [1-14C]pyruvate into

triglyceride glycerol after 3 d of exposure to the drug (65). It is of interest that

rosiglitazone also induces the expression of the gene for glycerol kinase (66), so that both

the direct phosphorylation of glycerol via glycerol kinase, and the generation of G-3-P by

glyceroneogenesis, contribute to triglyceride synthesis in adipose tissue. Although

glycerol kinase could also contribute to the process of re-esterification, analysis of the

respective contributions of glycerol kinase and PEPCK-C in cultured adipocytes suggests

glyceroneogenesis accounts for at least 75% of the TZD effect (63). Furthermore, over-

20 expression of glycerol kinase was less than half as efficient as rosiglitazone treatment in decreasing FFA release from cultured adipocytes (66).

Many lines of investigation have pointed to glyceroneogenesis as one etiological factor in Type 2 diabetes (67). The implication of PEPCK-C in an anti-diabetic action of

TZDs is perhaps the reason why “fatless” mice (lipodystrophic model) that develop Type

2 diabetes are not responsive to TZD treatment (63). Future studies aimed at employing adipose tissue specific PEPCK-C knockout mice will prove to be invaluable in testing the hypothesis that PEPCK-C is an important target for the anti-diabetic actions of TZDs

(63).

Physiological role of glyceroneogenesis in other species

Rainbow smelt (Osmerus mordax) Since the discovery of glyceroneogenesis in the adipose tissue of the rat, this pathway has been shown to occur in adipose tissue

(63;67-70) and liver (33;68) from a variety of animal species. For example, this pathway plays a critical role in the rainbow smelt (Osmerus mordax), where the concentration of glycerol in the blood can reach 0.4 M during the winter months, when the ambient temperature of the north Atlantic water is below 1° C (71). Interestingly, these fish have a glycerol-3-phosphatase that allows them to make glycerol directly without going through triglyceride, as is required in mammals (71). Expression of the gene for PEPCK-

C is induced in the liver by the cold in parallel with the synthesis of G-3-P from amino acids via glyceroneogenesis (71). Studies with rainbow smelt held at low temperature (-

1° C) and injected with L-[2,3-13C]alanine or D-[U-13C]glucose demonstrated the conversion of both alanine and glucose to glycerol. However, these fish eat krill and

21 other small fish, so they have only negligible amounts of carbohydrate in their diets from which to make the glycerol via glycolysis. Therefore, glycerol is most likely synthesized via glyceroneogenesis, using amino acids as a carbon source (65).

Parasitic helminth (Schistosoma mansoni) The pathway of glyceroneogenesis has also been reported in the invertebrate Schistosoma mansoni, the parasitic helminth transmitted by fresh water mollusks and responsible for the major human endemic disease, schistosomiasis (72). For adult worms, which live in the bloodstream of vertebrates, glucose is largely available and represents the major nutrient source (73).

However, glucose is less abundant to the mollusk and cannot provide all the energy for

sporocyst (larval stage) differentiation. In addition, it has been demonstrated that

sporocysts stimulate the production of glutamine in mollusks, a substrate for both

gluconeogenesis and glyceroneogenesis (72). The incorporation of [14C]glutamine

indicated that the de novo synthesis of both glucose and glycerol occurred in sporocysts,

with the predominant production of glycerol. As suggested by Khayath et al. (72), the

increased expression of G-3-P dehydrogenase in sporocysts, compared with adult worms, was concordant with these data and with the need of G-3-P and further glycerol production by developing sporocysts. First, G-3-P is critical for membrane phospholipid synthesis which is a very important process for sporocyst multiplication (74). Glycerol derived from G-3-P could also represent an alternative source of energy and may represent an osmolyte in S. mansoni critical for adaptation of newly transformed sporocysts from fresh water to the hypertonic environment (72). Finally, the inhibition of

PEPCK activity, via 3-mercaptopicolinate, decreased the radioactivity found in glycerol

22 isolated from S. mansoni by more than 75%, supporting a key role for glyceroneogenesis

in these organisms.

Drosophila Glyceroneogenesis has also been shown to be active in the fat body

(the fly equivalent of the mammalian liver and adipose tissue) of Drosophila during its

larval stage (75). In order to analyze the physiological role of Drosophila ATF-2 (dATF-

2), Okamura et al. (75) generated dATF-2 knockdown flies using RNA interference.

ATF-2 is a member of the ATF/CREB family of transcription factors that is activated by

stress-activated protein kinases. Reduced dATF-2 in the fat body resulted in decreased

survival under starvation conditions due to smaller triglyceride reserves compared with

controls. Expression of the PEPCK gene was significantly reduced in the dATF-2

knockdown flies. Although the blood glucose level was unchanged in response to

starvation compared with controls, glyceroneogenic flux, measured by incorporation of

[1-14C]pyruvate into triglyceride glycerol, was reduced in lipids isolated from fat bodies

of knockdown flies. In addition, these studies provide evidence that glyceroneogenesis

occurs under conditions of normal feeding, since fat bodies of dATF-2 knockdown flies

fed under normal conditions exhibited reduced activity of glyceroneogenesis as compared

with control flies. Thus, PEPCK and glyceroneogenesis support triglyceride synthesis

under fasted conditions and in response to feeding in the Drosophila (75).

Nematode worm (Caenorhabditis elegans) Most recently, glyceroneogenesis has been implicated to be an important pathway in the nematode worm, Caenorhabditis elegans (76). Dietary restriction imposed on C. elegans by growth in axenic medium induced the down-regulation of lipid degradation and up-regulation of glyceroneogenesis

(76). The activity of PEPCK followed a similar trend. Thus, an increase in PEPCK

23 activity and the glyceroneogenic pathway may be contributing factors to the increase in

life span observed in C. elegans, as well as a wide range of species, in response to dietary

restriction.

Consequence of ablation and overexpression of PEPCK-C: transgenic mouse models

PEPCK-C knockout in the liver She et al. (77) used an allelogenic Cre/loxP gene

targeting strategy to create mice with a liver-specific ablation of the PEPCK-C gene.

Mice with a whole body ablation of PEPCK-C die within three days of birth. However,

mice with a liver-specific deletion in the gene for PEPCK-C, are viable but develop

profound fatty livers and hepatic steatosis after fasting, despite the up-regulation of genes

encoding FFA oxidizing enzymes. In addition, these transgenic mice remain euglycemic

after a 24 h fast; most likely due to an increase in the contribution of renal glucose

production. Thus, hepatic PEPCK-C may also integrate hepatic energy metabolism (77).

Role of PEPCK-C in cataplerosis In order to identify the abnormalities of hepatic

energy metabolism that lead to steatosis during fasting, Burgess et al. (78) studied

metabolic fluxes in isolated, perfused livers of mice lacking hepatic PEPCK-C by using

combined [2H] and [13C]NMR isotopomer analysis. After short-term fasting (4 h), the

production of glucose from glycogenolysis and glycerol remained normal, whereas gluconeogenesis from TCA cycle intermediates was nearly absent. Upon extended fasting (24 h), hepatic glucose production was derived entirely from glycerol. In addition, the mitochondrial reduction-oxidation (redox) state, as measured by the

NADH/NAD ratio, was 5-fold higher, hepatic TCA cycle intermediate concentrations were dramatically increased, and oxygen consumption was decreased. Flux through the

24 TCA cycle and pyruvate cycling was 10- and 40-fold lower, respectively. These findings suggested a broader role of PEPCK-C in energy homeostasis via cataplerosis, which might include contributions to both gluconeogenesis and glyceroneogenesis.

Cataplerosis involves reactions that dispose of TCA cycle intermediates generated by the entry of compounds into the cycle during the breakdown of amino acids and other metabolites (79). Just as anaplerosis involves the replenishment of TCA cycle intermediates during periods of biosynthesis (when intermediates leave the cycle), cataplerosis must provide for output of intermediates to maintain a delicate balance of the cycle (79). A high level of mitochondrial NADH, as observed in these knockout mice, is known to inhibit TCA cycle flux (80). Overproduction of NADH may be linked to the absence of PEPCK-C in three possible ways (78): 1) the NADH-consuming glyceraldehyde-3-phosphate dehydrogenase reaction is not actively contributing triose phosphate units into gluconeogenesis; 2) the demand for GTP/ATP required to convert

OAA to PEP is absent, so there is less demand to drive ATP synthesis; 3) excess cytosolic NADH is generated from gluconeogenesis via glycerol, further exacerbating the reduced energy needs of the PEPCK-C null liver. Thus, all NADH-producing steps in the

TCA cycle are inhibited. This is reflected by the dramatically reduced TCA cycle flux.

In addition to the NAD/NADH ratio, the TCA cycle is also controlled by the concentration of cycle intermediates.

The impact of hepatic PEPCK-C deletion on the concentration of TCA cycle intermediates is a direct reflection of its importance in cataplerosis. Under steady-state conditions, anaplerosis and cataplerosis must be equivalent in order to sustain constant levels of TCA cycle intermediates (79). With the loss of PEPCK-C, the most important

25 cataplerotic pathway in the liver, cycle intermediates accumulate because export of OAA

is blocked. The resultant increase in the concentration of TCA cycle intermediates causes

product and allosteric inhibition of cycle activity and fatty acid oxidation. Furthermore, since PEPCK-C converts the OAA formed in the TCA cycle to PEP, not only is the subsequent formation of glucose via gluconeogenesis hampered, the synthesis of G-3-P via glyceroneogenesis may also impaired (79).

The liver-specific PEPCK-C null mice exhibit dramatic increases in hepatic triglycerides (100%), plasma FFA (60%), and plasma triglyceride levels (34%) after a 24 h fast (78). Since glyceroneogenesis plays an important role in systemic TG-FA cycling, it has been suggested that the alteration in lipid metabolism in hepatic PEPCK-C null mice may stem from a loss of the glyceroneogenic pathway and subsequent interruption of TG-FA cycling (81;82). Although the data in this study offered little to support or challenge the role of glyceroneogenesis and TG-FA cycling in the development of hepatic steatosis, this study suggested a substantial alteration in hepatic energy homeostasis may be responsible. Although increased triglyceride accumulation in the presence of PEPCK-C ablation is opposite to observations in adipocytes (81), the number of pathways that interact with anaplerosis and cataplerosis is much greater in the liver compared with the adipocyte (79). The elucidation of the specific role of PEPCK-C in fat metabolism has yet to be deciphered. However, the resulting impaired lipid metabolism of the liver knockout mice indicated that PEPCK-C may play a vital role in the integration of multiple pathways of energy metabolism (77).

PEPCK-C knockout in white adipose tissue Olswang et al. (67) genetically engineered mice, abalting PEPCK-C in adipose tissue, by mutating the PPRE in the

26 PEPCK-C gene promoter. The PPRE site is essential for PEPCK-C gene expression in

adipose tissue of mice (67). The resulting homozygous mutant mice (PPRE-/-) did not express the gene for PEPCK-C in white adipose tissue and had lowered levels of

expression in brown adipose tissue. The PPRE-/- mice had a reduction in epididymal

white adipose tissue compared to controls (PPRE+/+), with ~25% of the mice displaying

lipodystrophy. Furthermore, there was an elevated release of FFA when epididymal

adipsoe tissue from 18 h fasted PPRE-/- mice was incubated with 25mM pyruvate at

37°C for 3 h. In contrast, heterozygous mice (PPRE+/-) showed significant inhibition of

FFA release in the presence of pyruvate. Therefore, ablation of PEPCK-C specifically in

adipose tissue, and thus the glyceroneogenic pathway, resulted in decreased synthesis of

triglyceride and was responsible for the reduced weight of the epididymal fat pads (67).

Overexpression of PEPCK-C in white adipose tissue Franckhauser et al. (70) used a chimeric gene to generate transgenic mice that overexpressed PEPCK-C in adipose tissue. PEPCK-C gene transcription was placed under control of a constitutive adipose-specific adipocyte lipid-binding protein gene (aP2) promoter. When compared to control mice, PEPCK-C activity was 4-fold higher in heterozygotes, and 13-fold higher in homozygotes. Glyceroneogenesis was observed in vitro in epididymal white adipose tissue by measuring the conversion of [14C]pyruvate to triglyceride glycerol. In

heterozygous and homozygous transgenic mice, the conversion was increased 2-fold and

2.5-fold, respectively, compared to control mice. Furthermore, re-esterification of FFA

was approximately 2-fold higher in transgenic mice as compared to controls. The

increased synthesis of G-3-P via the glyceroneogenic pathway, enabled the transgenic

mice to re-esterify a greater amount of FFA (70). Although food intake for control and

27 transgenic mice were nearly equal, the body weight of the homozygous transgenic mice was increased by 33% as compared to control animals. The weight of epididymal white adipose tissue in the transgenic mice increased by ~180% for the heterozygous mice and

~340% for the homozygous mice. Furthermore, total body fat was increased ~125% in heterozygous transgenic mice and ~200% in homozygous transgenic mice. Histological analysis of adipose tissue revealed hypertrophy in white adipocytes of transgenic mice.

Thus, the physical consequence of increased glyceroneogenesis, via overexpression of

PEPCK-C, was an accumulation of fat, which lead to obesity, not observed in controls.

These results introduced the possibility that the dysregulation of glyceroneogenesis may affect lipid deposition and contribute to obesity (70).

In response to intraperitoneal glucose administration, Franckhauser et al. (70) found that obese transgenic mice displayed normal glucose tolerance. Furthermore, whole-body insulin sensitivity measured with an insulin tolerance test, showed similar decreases in blood glucose concentrations in transgenic and control mice after insulin injection. In addition, a significant increase in basal [14C]2-deoxyglucose uptake was detected in white adipose tissue. Thus, adipose tissue was not insulin resistant in transgenic mice. Skeletal muscle glucose uptake was similar in transgenic homozygous mice and controls, and serum triglyceride levels under fed and starved conditions were the same for transgenic and control mice. Therefore, the overexpression of PEPCK-C only affected adipose tissue metabolism and did not alter skeletal muscle or liver metabolism. It is of great interest that obese transgenic mice showed no increase in plasma FFA, or any other signs of Type 2 diabetes, due to increased fatty acid re- esterification through glyceroneogenesis. This suggests an important role for

28 glyceroneogenesis in maintenance of lipid homeostasis and presents this pathway as a

potential site for pharmacological intervention.

However, when Franckhauser et al. (83) examined whether these transgenic mice

were protected from diet-induced insulin resistance by challenging them with a high fat

diet for six weeks, the mice developed severe obesity, were more hyperinsulinemic,

glucose intolerant, and insulin resistant than controls. Furthermore, serum triglyceride

levels were increased and hepatic fat deposition was observed. Thus, the triglyceride

accumulation in the adipose tissue (both white and brown) resulted in fat storage

saturation and prevented this tissue from buffering the flux of lipids in circulation, which

subsequently altered liver metabolism. This implies the existence of a threshold in the

increase of adipose tissue mass, which is most likely crucial for the appearance of ectopic

fat deposition (83). PEPCK-C overexpression and enhanced glyceroneogenesis resulted

in insulin sensitivity on a normal diet and paradoxically in insulin resistance in mice fed

a high fat diet (83).

Overexpression of PEPCK-C in skeletal muscle Most recently, Hakimi et al. (23)

generated transgenic mice which overexpress the gene for PEPCK-C in skeletal muscle.

This was accomplished by introducing a chimeric gene into mice which contained the

cDNA for PEPCK-C from the mouse linked to the human α-skeletal actin gene promoter

(84). The activity of PEPCK-C measured in gastrocnemius, soleus, and diaphragm of

transgenic mice was 10-fold higher than the controls. Overexpression of PEPCK-C

resulted in a profound alteration of the phenotype of the mouse. Most notable was an

extremely high level of physical activity accompanied by a 40% increase in VO2max. The increased oxidative capacity may be attributed to the high daily activity of the mice

29 which lead to dramatic increases in both mitochondrial biogenesis and IMTG

concentrations (up to 10X that of control animals). The PEPCK-Cmus mice did not accumulate lactate in their blood during maximal exercise. Furthermore, the mice generated little lactate and were able to use fatty acids as an energy source over the entire period of strenuous exercise (RER of 0.91 at exhaustion). These results indicated that the enhanced oxidative capacity of the muscle was supported by more complete oxidation of glucose/glycogen and increased IMTG levels.

It has been hypothesized that the rate of TCA cycle flux, and thus ATP generation via the respiratory chain, might be limited by the concentrations of intermediates in the cycle (85-88). Since the concentration of TCA cycle intermediates in the mitochondria of skeletal muscle greatly increases in response to exercise, the presence of PEPCK-C in muscle may provide a cataplerotic mechanism for the removal of intermediates during or after exercise (23). In this regard, ablation of PEPCK-C activity in the liver greatly decreased TCA cycle flux (78;89). Thus, it was speculated that an increase in the activity of the enzyme would have the opposite effect (23). Another possible role of PEPCK-C in skeletal muscle was suggested to be glyceroneogenesis (23;26). A considerable amount of triglyceride is synthesized and deposited in skeletal muscle in support of energy metabolism. Furthermore, in vivo studies demonstrate that exercise increases TG-FA cycling in skeletal muscle of the rat (35). In response to 48 h of fasting or in rats maintained on a high carbohydrate diet, the predominant pathway for triglyceride glycerol synthesis in gastrocnemius and soleus muscle has been demonstrated to be glyceroneogenesis, not glycolysis (26). Therefore, increased glyceroneogenesis may be

30 responsible for provision of G-3-P necessary for triglyceride glycerol synthesis in TG-FA cycling as well as the enhanced triglyceride deposition observed in PEPCK-Cmus mice.

Glyceroneogenesis in vivo

The elucidation of the pathway of glyceroneogenesis began in the 1960s, with early studies investigating this pathway using in vitro techniques (21;29). Subsequent

studies of glyceroneogenesis over the next thirty years were sparse, and

glyceroneogenesis remained largely ignored in the scientific literature. This situation has

slowly changed over the past decade, due in part to the availability of genetically

modified mice in which the gene for PEPCK-C has been altered, very few studies have

investigated glyceroneogenesis in vivo. To date, only nine studies (from three different

laboratories) have been conducted which include measurements of glyceroneogenesis in

vivo (33;48;49;69;90-94). Botion et al. (48) investigated this pathway in rats adapted to a

high protein, carbohydrate free diet by determining the relative contribution of

glyceroneogenesis versus glycolysis to triglyceride glycerol synthesis using the rate of

3 3 14 14 incorporation of [ H] from tritiated water ([ H2]O) and of [ C] from [U- C]glucose,

respectively, into the carcass, liver, and adipose tissue. Botion and colleagues (68)

pursued this study when results from previous experiments performed in vitro cast doubt

upon the importance of glyceroneogenesis. They had demonstrated that epididymal

adipose tissue from fed rats maintained on a high-protein diet had an increased

glyceroneogenic capacity, as evidenced by enhanced rates of incorporation of [14C] from

[14C]pyruvate into triglyceride glycerol, in the absence of glucose (68). Yet in the

presence of physiological concentrations of glucose, the contribution of pyruvate became

31 negligible in both high protein and control rats, with almost all of the triglyceride

glycerol derived from glucose (68). However, the metabolic significance of glyceroneogenesis was confirmed in an in vivo study (48) which showed higher rates of

glyceroneogenesis in both retroperitoneal and epididymal adipose tissue from rats

adapted to a high protein diet, as compared with controls. Furthermore, this was the first study to demonstrate the existence of hepatic glyceroneogenesis; prior to this publication,

glyceroneogenesis was considered to be an important pathway only in adipose tissue.

Glyceroneogenesis was first studied in vivo in humans by Kalhan et al. (33) who

measured hepatic glyceroneogenesis in pregnant and non-pregnant women after an

overnight fast. In this study, the fractional contribution of pyruvate and glycerol to the

synthesis of triglyceride glycerol obtained from the plasma was determined. The

contribution of pyruvate was quantified by employing the deuterium labeling of body

2 water ([ H2]O) technique, whereas the contribution of glycerol was measured using [3-

13C]glycerol tracer. Since the liver synthesizes a significant amount of triglyceride

during fasting and the glycerol Ra is increased as a result of enhanced adipose tissue

lipolysis, it was assumed that this glycerol would be the predominant source of hepatic

triglyceride glycerol synthesis (33). However, the results showed triglyceride glycerol

derived from pyruvate ranged from 10 to 60% in both pregnant and non-pregnant women,

whereas only ~6% of triglyceride glycerol was derived from plasma glycerol. This

suggested that glyceroneogenesis from pyruvate was quantitatively a major contributor to

plasma triglyceride glycerol synthesis in humans.

A more recent study by Kalhan et al. (90) focused on quantifying the contribution

of glyceroneogenesis versus glycolysis to triglyceride found in the VLDL of humans

32 with Type 2 diabetes. These subjects were studied before and after a 6-month behavioral

intervention therapy, during fasting and during a hyperinsulinemic/normoglycemic

clamp. The contribution of pyruvate, via glyceroneogenesis, to triglyceride glycerol

synthesis was measured using the deuterium labeling of body water technique, whereas

2 the contribution of glucose, via glycolysis, was measured using [6,6- H2]glucose. The

results showed that the estimated contribution of glyceroneogenesis to VLDL triglyceride

glycerol synthesis was ~53% and remained unchanged in response to clinical intervention

and during the hyperinsulinemic/normoglycemic clamp. Furthermore, glycolysis

contributed only 16% to triglyceride glycerol synthesis and this level of contribution did not change as a result of intervention therapy or in response to insulin plus glucose infusion, even though intervention resulted in an improvement in insulin sensitivity as evidenced by higher glucose uptake during the clamp. Thus, glyceroneogenesis, glycolysis, and plasma glycerol each contribute ~50%, ~15%, and ~5%, respectively, to the synthesis of hepatic triglyceride glycerol. The discrepant ~30% represents the

unlabeled triglyceride pool in the circulation. Nevertheless, glyceroneogenesis is quantitatively the major pathway for the synthesis of triglyceride glycerol in both studies, under all conditions examined.

The role that glyceroneogenesis may play in vivo in adipose tissue triglyceride synthesis under altered physiologic states or after pharmacologic treatment was investigated by Chen et al. (91). Glyceroneogenesis was measured in vivo in four different adipose depots of rodents maintained on a low carbohydrate diet, a high carbohydrate diet, or a high carbohydrate diet in conjunction with rosiglitazone treatment.

The methodological approach used in this study was unique in that it estimated the

33 fractional contribution of glyceroneogenesis to triglyceride glycerol synthesis based upon

2 long-term [ H2]O administration, while animals were maintained on their respective diets for 75 days. Glyceroneogenesis was found to be higher in adipose tissue from mice fed a low carbohydrate diet (50%) as compared to a high carbohydrate diet (17%). Thus, on a standard, high carbohydrate chow diet, the majority (83%) of triglyceride glycerol present in adipose tissue was derived from glycolytic metabolism of unlabeled glucose.

The administration of rosiglitazone to mice fed a high carbohydrate diet increased glyceroneogenesis in adipose tissue from 17% to 53%. This provided the first in vivo evidence that the upregulation of this pathway, presumably via increased PEPCK-C gene expression, occurred in response to thiazolidinedione treatment.

More recently, Brito et al. (49) has carried out in vivo analysis in rats which supported a role for dietary fatty acids in promoting glyceroneogenesis in adipose tissue under high protein, carbohydrate free conditions. In addition, Varady et al. (92) sought to identify a dietary regimen that could alter adipose tissue metabolism without causing loss of body weight or fat. In vivo triglyceride kinetic parameters revealed glyceroneogenesis was increased in response to a modified alternate-day fasting regimen (at least 50% caloric restriction on ‘fasting’ day) in adipose tissue of mice. Thus, when considering the studies of glyceroneogenesis in vivo, they are not only limited in number and progressively become more narrow, focusing on a particular tissue type and a specific dietary condition. What is lacking in the literature is a comprehensive study of glyceroneogenesis in vivo that provides simultaneous flux rates over this pathway in several tissue types of an intact animal. Furthermore, such measurements need to be established under multiple physiological conditions. Although the role of

34 glyceroneogenesis for esterification of fatty acids has been emphasized in adipose tissue

(and more recently the liver), the potential importance of this pathway in skeletal muscle

has been largely ignored, despite the fact that this tissue contains triglyceride stores, a low level of PEPCK-C activity, and is predominantly responsible for the insulin resistant state observed in Type 2 diabetics.

Statement of purpose

The extent to which glyceroneogenesis occurs in peripheral tissues under specific

dietary conditions or metabolic perturbations has yet to be determined. It was the goal of

this research to carry out the first comprehensive, whole-body analysis of

glyceroneogenesis in vivo. More specifically, we have determined the relative

contribution of glyceroneogenesis and glucose via glycolysis to the synthesis of

triglyceride glycerol in white adipose tissue (mesenteric and epididymal), skeletal muscle

(gastrocnemius and soleus), and the liver of intact rats. The rate of flux over the

glyceroneogenic and glycolytic pathways was investigated over a range of physiological

perturbations, i.e. chow-fed controls, 48 h fasted animals, and rats that were fed a

sucrose-supplemented, lipogenic diet. We also examined glyceroneogenesis in adipose

tissue in response to enhanced TG-FA cycling caused by infusion of epinephrine.

Establishing quantitative values for glyceroneogenesis in vivo will help define the extent

and distribution of this pathway in vivo and provide physiological insight into lipid

homeostasis.

35 Hypothesis

Due to its important role in TG-FA cycling and for controlling the level of FFA in the blood, we hypothesize that the pathway of glyceroneogenesis is critical for the regulation of triglyceride cycling and for the re-esterificaation of FFA in mammalian tissues. It is also a key element in the etiology of Type 2 diabetes in humans.

36 CHAPTER 2: Experimental Procedures

Animals Male Sprague-Dawley rats weighing ~250-275 g with indwelling carotid artery and jugular vein catheters, were obtained from Zivic-Miller Laboratories

(Zelienople, PA). The animals were housed in the Animal Resource Facility of Case

Western Reserve University under controlled temperature (22 ± 1°C) and light (on at

0600 h, off at 1800 h). All procedures involving the rats used in this study were reviewed

and approved by the Institutional Animal Care and Use Committee (IUCAC) and

conformed to American Association for Accreditation of Laboratory Animal Care

Guidelines. The animals were fed rat chow (Purina Mills Prolab RMH 1800; 21%

protein, 14% fat, 65% carbohydrate) and water ad libitum, and were provided with

environmental enrichment (chew squares). After entry into the facility, the rats were

allowed a minimum of four days to acclimate to the new surroundings prior to the study.

Glyceroneogenesis was quantified in four groups of animals: 1) Controls: food was removed at 7 AM on the morning of the tracer isotope study; water was provided ad

libitum. 2) Fasting: food was removed 48 h prior to the tracer study; water was provided ad libitum. 3) High carbohydrate diet: in addition to rat chow, the animals were given sucrose water (20% w/v) for 5 d. Food was removed at 7 AM, sucrose water was discontinued, and animals received glucose intravenously (15 mg/kg/min, as 10% solution) throughout the tracer study. 4) Response to epinephrine infusion: food was removed at 7 AM on the morning of the tracer study; water was provided ad libitum, and epinephrine (Hospira, Inc.; Lake Forest, IL) was infused (500 ng/kg/min) via the jugular vein catheter for 3 h.

37 Tracer study protocol (Fig. 3) Rats were given an intraperitoneal injection of 0.5

3 14 mCi of [ H2]O (1.0 mCi/ml) and [U- C]glucose, in isotonic saline solution, which was administered as a prime constant rate infusion (9.0 µCi/kg/h) via the jugular vein catheter

3 for 7 h. The rate of glyceroneogenesis was quantified using the tritium ([ H2]O) labeling

of body water method (Fig. 4), and the contribution of glucose, via glycolysis, was

determined using [U-14C]glucose tracer. The animals were unrestrained, awake, and

moved freely in the cage during the experiment. Blood samples (0.2 ml), were drawn

from the carotid artery catheter at 6, 6.5, and 7 h and plasma was separated. At 7 h, the rats were given an anesthetic dose of sodium pentobarbital (Abbott Laboratories) via the carotid artery catheter and the tissues of interest (epididymal and mesenteric fat, gastrocnemius and soleus muscles) were harvested while the tracer infusion was continued. Plasma and tissues were stored at -80°C until further analyses.

Rationale for selection of dietary groups Food was removed immediately prior to the tracer study protocol for the control animals, and thus, was representative of ~8 h of fasting; conditions in which PEPCK-C activity and glyceroneogenic flux was expected to be significant and quantifiable. We sought to investigate glyceroneogenesis under condtions where PEPCK-C activity and glyceroneogenic flux was expected to be high relative to controls; therefore, we selected conditions of 48 h of fasting. Furthermore, we wanted to investigate glyceroneogenesis under conditions where PEPCK-C activity and flux over this pathway was expected to be low relative to controls; accordingly, we selected lipogenic conditions of high carbohydrate feeding (20% sucrose supplementation) with continued glucose infusion. The infusion of exogenous glucose was carried out in order to maintain lipogenic/fed conditions throughout the course of the

38 Prime constant-rate infusion14 C]glucose[U-

3 [ H2]O Blood Samples Tissues

076 h 39 3 Figure 3. Tracer study protocol. Rats were given [ H2]O at time zero, followed by a prime constant-rate infusion of [U-

14C]glucose for 7 h. Animals moved freely about their cages during the study. Blood samples were collected during the final

hour of the infusion and tissues of interest were harvested while the tracer infusion was continued (see “Experimental

Procedures” for details).

40 G-3-P DHAP Glyceraldehyde-3-P NAD + NADH+H + CH 2OH CH 2OH CHO HO-CH C=O HC-OH CH2-OP CH2-OP CH 2-OP

+ 3 FA CoA

2-Phosphoglycerate COO - Triglyceride HC-OP CH 2OH Pyruvate H O Oxaloacetate 2 PEP - - - COO GTP GDP COO COO

C=O = C-OP C=O CH 2 PEPCK-C CH CH 3 COO - 2 Oxaloacetate CO 2 COO - NADH+H + C=O CH 2 + - COO + NAD NADH+H NAD + Malate Malate COO - COO - HO-CH HO-CH Fumarate COO - CH 2 CH 2 - - CH COO COO O H2 = HC - 41 COO Figure 4. The incorporation of [3H] into G-3-P from bodywater. The hydrogens of C-3 of pyruvate become labeled

through exchange with body water during transamination with alanine and keto-enol tautomerization, as well as during the

equilibrium of malate with fumarate, so that both hydrogens of C-3 of PEP will have the same SA as that of bodywater. Thus,

the [3H] of C-3 of G-3-P are derived entirely from pyruvate. Additional labeling occurs in the triose phosphate pool when C-1

and C-2 acquire [3H] directly from the bodywater. Thus, the [3H] of C-1 and C-2 are derived from the triose phosphate pool.

Hydrogens labeled with [3H] are highlighted in the red font.

42 tracer study. In addition, we examined glyceroneogenesis under conditions of enhanced

TG-FA cycling; thus, we induced cycling by infusing epinephrine intravenously (95).

Rationale for the design of tracer studies employing infusion of epinephrine We selected epinephrine, versus norepinephrine, to induce TG-FA cycling since infusion of epinephrine reflects a physiological state more likely to be observed. More specifically, epinephrine, the main stress-related hormone of the sympathetic nervous system, is released into the general circulation from the adrenal medulla and will present a uniform plasma and overall tissue concentrations (96). Whereas norepinephrine, the major neurotransmitter of the sympathetic nervous system, is released from axon terminals of sympathetic post-ganglionic neurons and exhibits steep concentration gradients between the synaptic clefts and plasma (97). Since basal epinephrine levels in rats (98;99) are approximately 10-fold higher than humans (97;100;101) and epinephrine infusion rates of

50 ng/kg/min in humans result in epinephrine concentrations observed during moderate exercise (102), we infused our rats at a rate of 500 ng/kg/min. The duration of the infusion study was significantly reduced to 3 h for epinephrine experiments due to the tachyphylactic response observed in infusion studies carried out for extended periods of time (101).

Triglyceride extraction and isolation of glycerol and fatty acids Triglycerides were extracted from tissues and plasma using a modification of the method of Folch et al.

(103). Briefly, adipose tissue was homogenized in 2:1(vol/vol) chloroform-methanol, while the muscle was homogenized in saline, using an IKA-ULTRA-TURRAX T 25 basic tissue blender. Plasma samples and tissue homogenates were incubated in

2:1(vol/vol) chloroform-methanol for 48 h at 4°C; then 4 mM MgCl2 was added, the

43 solution was mixed thoroughly, and centrifuged at 1000 X g for 1 h. The aqueous

supernatant was removed, taken to dryness, and further processed as described below.

The organic fraction containing the triglycerides was dried and saponified by the addition

of ethanolic KOH (0.5 N), followed by incubation at 70ºC for 1 h. To convert the

carboxylate salts to the free acids, the samples were acidified by the addition of 6 N HCl.

Fatty acids were extracted with hexane (Fluka) and dried, while the aqueous layer

containing glycerol was dried, reconstituted in a fixed volume and the concentration of

glycerol was determined fluorometrically (using a Cary Eclipse Fluorescence

Spectrophotometer) by the method of Wieland (104).

Isolation of plasma metabolites Plasma glucose, glycerol, and lactate were

separated by ion-exchange chromatography (BioRad AG 1-X8 hydrogen resin; AG 50W-

X8 formate resin). Glucose and glycerol, eluted in the neutral fraction, were further separated by liquid chromatography (Hewlett Packard Agilent 1100 Series HPLC with

Agilent ChemStation software; BioRad Aminex HPX-87P Carbohydrate Analysis

Column, 300 x 7.8 mm; mobile phase 100% water, flow rate 0.6 ml/min, column temperature 80ºC). The eluates were dried and reconstituted in a fixed volume. The

glucose concentration was determined enzymatically using the Beckman Glucose

Analyzer (glucose oxidase method), while the concentrations of glycerol and lactate were

determined enzymatically using a fluorometric assay (104;105).

Measurement of radioactivity The radioactivity of [3H] in a 20 μl aliquot of

plasma was measured to determine the specific activity of body water. An aliquot of

glucose, glycerol and lactate, isolated from plasma samples (above), was used to measure the total [3H] and [14C] radioactivity of each metabolite. The total [3H] and [14C]

44 radioactivity present in fatty acids and glycerol (from triglyceride) isolated from plasma

and tissues was also measured.

Sources of carbon and hydrogen in triglyceride glycerol Inherent to our

calculations for glyceroneogenesis is the fact that we determine the radioactivity of all

[3H] present on isolated triglyceride glycerol, assume complete and equal labeling of all

five hydrogens, and use the known labeling pattern of [3H] on C-3 of pyruvate. However, one may suggest that all [3H] radioactivity measured on triglyceride glycerol may be derived from the triose phosphate pool, not from glyceroneogenesis. Therefore, in order to lend support to our data which demonstrates the predominance of glyceroneogenesis in vivo, we measured the [14C] and [3H] radioactivity of C-1 and C-3 isolated from triglyceride glycerol via dimedon derivatization. We specifically examined mesenteric adipose tissue depots of glucose infused rats since values of glyceroneogenesis were highest in the depot of this group; conditions which were hypothesized to result in low glyceroneogenic flux rates.

Appearance of [14C] label in the triose phosphate pool Trioses from [U-

14C]glucose will result in complete labeling of carbons with [14C] (Fig. 5, left side of

diagram). On a stoichiometric equivalent basis, trioses from two molecules of pyruvate will result in no labeling of carbons (Fig. 5, center of diagram). Since glucose can be

metabolized to lactate in peripheral tissues, and lactate subsequently released and taken

up by other tissues, recycled [14C]lactate becomes available (Fig. 5, right side of

diagram). Note that recycled [14C]lactate proceeding to the level of the triose phosphate

pool will contain less label on C-1. Formation of PEP from pyruvate, via OAA, will

45 [14C]Glucose

H- O C = HC-OH HO-CH 14C/3H ratio of C-1 = 2C/8H = 0.25 HC-OH HC-OH 14 3 HCH C/ H ratio of C-3 = 2C/4H = 0.50 - OH

CH2OH CH2OH CH2OH CH2OH CH2OH 1 HO-CH HO-CH HO-CH HO-CH HO-CH 2

CH2-OP CH2-OP CH2-OP CH2-OP CH2-OP 3

COO- COO- COO- = C-OP= C-OP= PEP C-OP PEP

CH2 CH2 CH2

TCA COO- COO- COO- C=O C=O C-OH

CH3 CH3 CH3 46 Pyruvate [14C]Lactate Figure 5. Relative contribution of glyceroneogenesis, direct and indirect (via lactate) pathways of glucose, and

[14C]/[3H] ratio of C-1 and C-3 of triglyceride glycerol. The rectangle in the figure represents G-3-P in the triose phosphate

pool as derived from various precursors. When [14C]glucose is the precursor for triglyceride glycerol, C-1, C-2 and C-3 of G-

3-P will be labeled with [14C], however, only C-1 and C-2 will be labeled with [3H] in the triose phosphate pool. On a

stoichiometric equivalent basis, two molecules of pyruvate proceeding to the level of the triose phosphate pool will result in no

[14C] labeling of G-3-P, whereas the hydrogens on C-1, C-2 and C-3 of G-3-P will be completely labeled; hydrogens on C-3 of

pyruvate are completely labeled prior to reaching the triose phosphate pool (at PEP), whereas hydrogens on C-1 and C-2

become labeled at the level of the triose phosphate pool. [U-14C]glucose is metabolized to [U-14C]lactate, and [14C]

incorporation into G-3-P can occur indirectly via [14C]lactate. However, [14C]lactate proceeding to the level of triose

phosphate contains less label on C-1. Formation of PEP from pyruvate or lactate, via OAA, will result in the incorporation of

unlabeled carbon from the bicarbonate pool (pyruvate carboxylase and CO2 fixation) onto the C-1 position of PEP. As the

contribution of recycled glucose to G-3-P formation increases, this will result in an increase of the [14C]/[3H] ratio on C-3 and a

decrease of the ratio on C-1. Thus, a predominance of glyceroneogenic flux will result in a high ratio on C-3 (or C-1 + C-3)

versus C-1. Carbons labeled with [14C] are highlighted in blue font; hydrogens labeled with [3H] are highlighted in red font.

47 result in the incorporation of unlabeled carbon from the bicarbonate pool (pyruvate

carboxylase and CO2 fixation) onto the C-1 position of PEP.

Source of [3H] label of C-1 and C-3 of the triose phosphate pool Trioses from

glucose will result in complete labeling of hydrogens of C-1 upon entering the triose

phosphate pool (hydrogens of C-3 are unlabeled) (Fig. 5, left side of diagram). On a

stoichiometric equivalent basis, two molecules of pyruvate proceeding to the level of the

triose phosphate pool will result in two trioses with completely labeled hydrogens on C-1

and C-3 (Fig. 5, center of diagram). Hydrogens of C-3 of pyruvate are completely

labeled prior to reaching the level of the triose phosphate pool, whereas hydrogens of C-1

of pyruvate become completely labeled upon entering the triose phosphate pool.

Ratio of [14C]/[3H] of C-1 and C-3 of triglyceride glycerol As the contribution of

pyruvate to trioses increases (and the contribution of recycled [14C]lactate increases), the

ratio of [14C]/[3H] of C-1 will decrease while the ratio of [14C]/[3H] of C-3 will increase.

The ratio of C-1 is expected to decrease, since more unlabeled carbon will become

incorporated (from unlabeled C-1 of recycled [14C]lactate) while the [3H] will remain the

same, once [3H] incorporation into the triose phosphate pool plateaus (numerator will

decrease, denominator will not change). The ratio of C-3 is expected to increase since

the [14C] label of C-3 will continually increase as the contribution of recycled [14C]lactate increases while the [3H] of C-3 plateaus (numerator increases, denominator remains

unchanged).

Radioactivity of C-1 and (C-1 + C-3) The [14C] and [3H] radioactivity of the C-1

and C-3 triglyceride glycerol carbons was determined after dimedon derivatization,

according to the method of Reeves (106). Briefly, glycerol samples were subjected to

48 periodate cleavage and the formaldehyde formed was precipitated in a dimedon

derivative. Each glycerol sample was divided in half: one aliquot was taken directly

through the dimedon derivatization procedure; the other aliquot was phosphorylated

using glycerol kinase (below) to form G-3-P prior to carrying out the derivatization procedure. Incubation of glycerol (or G-3-P) with periodic acid will result in cleavage of vicinal diols. Phosphorylation of glycerol to form G-3-P will prevent the cleavage of C-3 by periodate. Thus, radioactivity of (C-1 + C-3), and C-1 only, can be determined from the same sample of triglyceride glycerol.

Phosphorylation of glycerol Triglyceride glycerol (~150 µmol) was enzymatically converted to G-3-P by incubating it overnight at 25˚C with 0.2 M ATP, 0.2

M MgCl2, and 10 U glycerokinase (Cellulomonas species, Sigma-Aldrich) in a final volume of 2.0 ml of 0.1 M phosphate buffer (pH 9.15). G-3-P was isolated by ion exchange chromatography using AG 1-X8 resin (BioRad) in 1N formic acid. The column was sequentially eluted with water, 1N formic acid, and 4N formic acid. G-3-P appeared in the 4N formic acid fraction. The recovery was >95%.

Activity of PEPCK-C The activity of PEPCK-C was determined by the method of

Ballard and Hanson (107), using adipose tissue isolated from 48 h fasted and glucose infused animals. The tissues were homogenized in 0.25 M sucrose, containing 5 mM

Tris-HCl at pH 7.4, and 1 mM dithiothreitol. A cytosolic fraction, prepared by centrifuging the homogenate at 30,000 X g for 30 min at 4ºC, was used to assay for

PEPCK-C activity.

Calculations The rate of appearance (Ra) of glucose in the blood was calculated during the isotopic steady state using the tracer dilution technique (108):

49 Ra (µmol/kg/min) = I/SA, where I is the rate of infusion of [U-14C]glucose tracer

(dpm/kg/min), and SA is the specific activity of glucose (dpm/µmol).

The SA of pyruvate was calculated from that of plasma water, assuming complete equilibration between the hydrogens on plasma water and the hydrogens on C-3 of pyruvate (Fig. 2). Therefore, the SA of pyruvate = 3 x SA of body water, assuming all three hydrogens on C-3 of pyruvate are in complete equilibrium with body water (109).

SA of body water = 1.11 x SA of plasma. The measured SA of plasma was multiplied by 1.11 since water constitutes 90% of plasma.

The rate of gluconeogenesis was calculated by dividing the [3H] SA of glucose with

the estimated SA of the triose phosphate pool. The SA of the triose phosphate pool = 4 x

SA of body water, since there are four hydrogens on each molecule of triose phosphate

(DHAP and glyceraldehyde-3-P) in equilibrium with body water (Fig. 2). It was assumed

that all the hydrogens on the triose phosphate molecules will be labeled to the same

extent as body water.

The rate of glyceroneogenesis in the adipose tissue and skeletal muscle was

calculated as follows:

Glyceroneogenesis (nmol/g/h) = ([3H] in triglyceride glycerol (dpm/g/h) x 2/5) /

([3H] SA of pyruvate (dpm/µmol) x 2/3).

The [3H] radioactivity of triglyceride glycerol was multiplied by 2/5 since only two

hydrogens in the glycerol moiety are derived from pyruvate (Fig. 2). The SA of pyruvate

was multiplied by 2/3 since only two hydrogens on C-3 of pyruvate are incorporated into

triglyceride glycerol. The SA of [3H] on C-1 and C-2 of triglyceride glycerol will be the

50 same as that of body water because of complete equilibrium in the triose phosphate pool.

In contrast, [3H] on C-3 of triglyceride glycerol are those derived from pyruvate.

The “total” (“direct” plus “indirect” via lactate) rate of the glycolytic contribution to triglyceride glycerol in adipose tissue and skeletal muscle was calculated as follows:

“Total” (direct plus indirect) incorporation of glucose into triglyceride glycerol

(nmol/g/h) = 2 x ([14C] radioactivity in triglyceride glycerol (dpm/g/h)) / ([14C] SA of glucose (dpm/µmol)).

The data are expressed as three-carbon equivalents by multiplying the results by 2.

The “direct” contribution of glucose carbon to triglyceride glycerol was calculated by subtracting the contribution of lactate (glyceroneogenesis) from the “total” glucose carbon incorporated into triglyceride glycerol. The contribution of lactate was calculated by multiplying the [14C] SA of lactate with the estimated glyceroneogenic flux from

pyruvate (corrected for the loss of [14C] in the citric acid cycle: dilution factor 2.2) (110).

Additional studies using four animals showed that the SA of the [14C]lactate pool reached

steady state within 1 h of the start of the [14C]glucose infusion (data not shown).

Plasma triglyceride glycerol from glyceroneogenesis (%) = ([3H] SA plasma

triglyceride glycerol (dpm/µmol) x 2/5) / ([3H] SA of pyruvate (dpm/µmol) x 2/3).

Both the SA of triglyceride glycerol and pyruvate were adjusted, since only two

hydrogens on C-3 of pyruvate are incorporated into the glycerol moiety.

Plasma triglyceride glycerol from glucose (%) = 100 x ([14C] SA of plasma

triglyceride glycerol (dpm/µmol) x 2) / ([14C] SA of glucose (dpm/µmol)).

The SA of triglyceride glycerol was multiplied by 2 since two three-carbon equivalents are formed from one molecule of glucose.

51 The rate of lipogenesis was calculated as described by Katz and Rognstad (111).

Statistical analysis Values are expressed as mean ± S.E. Analysis of variance

(ANOVA) was used to examine the differences between the three dietary groups. Two- tailed Student’s t-test (assuming unequal variance) was used to assess differences between two dietary groups for each tissue, and within the same dietary group between two adipose tissue depots (or two types of skeletal muscle).

52 CHAPTER 3: Results

Glucose kinetics, gluconeogenesis, and the source of plasma triglyceride glycerol

The mean plasma glucose concentration of control and 48 h fasted animals was 8.6 mM

and 6.3 mM, respectively (Table 1). The mean plasma glucose concentration (9.8 mM)

of the rats supplemented with sucrose was not significantly different when compared with

controls. The glucose Ra was significantly lower in 48 h fasted animals (17.7

µmol/kg/min) when compared with controls (35.6 µmol/kg/min) (Table 1). In the

sucrose supplemented group, intravenous glucose infusion resulted in complete suppression of endogenous glucose production. The fractional contribution of

gluconeogenesis, estimated from the incorporation of [3H] of body water into glucose, was significantly higher in 48 h fasted animals (58.6 %) compared with the control group

(28.5%) (Table 1). The fractional contribution of gluconeogenesis in the sucrose supplemented group could not be calculated, since endogenous glucose production was completely suppressed.

The plasma concentration of triglycerides was not different in the three groups

(Table 1). The fraction of plasma triglyceride glycerol derived from glyceroneogenesis was ~60%, and not different among the three groups (Table 1). Approximately 15% of

triglyceride glycerol was derived from glucose in the control group (Table 1). The contribution of glucose was lower in 48 h fasted rats (~11%), and was higher in sucrose supplemented animals (~28%). Although sucrose supplementation resulted in a higher contribution of glucose to plasma triglyceride glycerol when compared with controls, the contribution of glucose was less than that of glyceroneogenesis (Table 1). However, the estimated contribution of glucose includes both the direct and indirect (via lactate)

53 Glucose kinetics, gluconeogenesis, and source of plasma triglyceride glycerol

[Glc] Ra GNG [TG] Glyceroneogenesis Glycolysis

(mM) (µmol/kg/min) (%) (mM) (%) Control 8.6 ± 1.0 35.6 ± 4.3 28.5 ± 2.9 0.841 ± 0.07 57.6 ± 2.2 14.8 ± 1.9 (n=5) 48 h Fast 6.3 ± 0.6 17.7 ± 0.6* 58.6 ± 1.6** 0.844 ± 0.06 56.1 ± 6.7 10.6 ± 0.7 (n=5) 0.803 ± 0.06 59.3 ± 2.7 27.8 ± 5.6 Suc Sup + Glc Inf 9.8 ± 0.9 0 0 (n=5) 54 Table 1. Glucose kinetics, gluconeogenesis, and source of plasma triglyceride glycerol. Values are the means ± S.E. Data

of control rats, 48 h fasted rats, and those supplemented with sucrose. The plasma [Glc] (glucose concentration), Ra (rate of

appearance of glucose), estimated contribution of GNG (gluconeogenesis), [TG] (triglyceride concentration), and fractional

contribution of glyceroneogenesis and glycolysis to the formation of triglyceride glycerol isolated from plasma were

determined as described under "Experimental Procedures."

*, p < 0.05; **, p < 0.01 vs. control.

55 pathways. Therefore, the reported contribution of glucose to plasma triglyceride glycerol

also includes the [14C] incorporated into plasma triglycerides via glyceroneogenesis (from

[14C]lactate).

Glyceroneogenesis and glycolysis as a source of triglyceride glycerol in adipose

tissue Fasting for 48 h resulted in a significantly lower triglyceride concentration in three

rats, however the data in the entire group (n=5) were not different from controls. In the

sucrose supplemented animals, the concentration of triglyceride was significantly higher than controls (Table 2).

Glyceroneogenesis was quantified using [3H] incorporation into triglyceride glycerol and is expressed as incorporation of pyruvate equivalents. The rate of glyceroneogenesis in the control animals was ~600 nmol/g/h in the epididymal adipose depot and ~800 nmol/g/h in the mesenteric depot (Table 2). Glyceroneogenesis did not

change in response to fasting for 48 h. In contrast, sucrose supplementation resulted in a

significant increase in this pathway in both adipose tissue depots. Glyceroneogenesis

was higher in mesenteric fat compared with epididymal fat in all groups; however, a

statistically significant difference was only observed in the sucrose supplemented group

(Table 2).

“Total” and “direct” glucose carbon incorporated into triglyceride glycerol are

displayed in Table 2. 48 h fast caused a significantly lower incorporation of total glucose

carbon into triglyceride glycerol, as compared with controls. In contrast, sucrose

supplementation resulted in a higher total contribution of glucose carbon to triglyceride

glycerol (Table 2). The direct contribution of glucose to triglyceride glycerol was ~80

nmol/g/h in control animals in both adipose depots (Table 2). In the 48 h fasted

56 The contribution of glyceroneogenesis and glycolysis to triglyceride glycerol synthesis in adipose tissue

Epididymal Fat Mesenteric Fat

[TG] GlyceroneogenesisGlycolysis [TG] GlyceroneogenesisGlycolysis

Total Direct Total Direct (µmol/g) (nmol/g/h) (nmol/g/h) (µmol/g) (nmol/g/h) (nmol/g/h)

Control 362 ± 45.0 400 ± 21.6 294 ± 37.9 (n=5) 604 ± 56.6 82.3 ± 12.5 414 ± 17.7 789 ± 70.5 86.3 ± 13.7 94.8 ± 6.63* 66.3 ± 5.6* (-24.5 - 11.5)** 816 ± 96.6 (-42.0 - 10.5)* 48 h Fast 352 ± 34.4 511 ± 65.0 359 ± 45.9 (n=5) 149 ± 19.3 1410 ± 85.2**1111 ± 133* 277 ± 34.1*

Suc Sup + Glc Inf 723 ± 19.2** 1022 ± 57.1** 671 ± 67.9* 695 ± 11.7* (n=5) 57 Table 2. The contribution of glyceroneogenesis and glycolysis to triglyceride glycerol synthesis in adipose tissue. Values

3 are the means ± S.E.; range is indicated in parentheses. Glyceroneogenesis was quantified using tritium, [ H2]O, incorporation

into triglyceride glycerol and is expressed as nmol/g/h pyruvate incorporation. The contribution of glycolysis was quantified

using incorporation of [14C] into triglyceride glycerol, during a prime constant-rate infusion of [U-14C]glucose. “Total”

contribution of glycolysis includes three-carbon equivalents coming directly from glucose plus those coming indirectly from

[14C]lactate. “Direct” contribution represents three-carbon equivalents coming directly from glucose only; the [14C] label

arriving from [14C]lactate has been subtracted out. The [TG] (triglyceride concentration) and the relative contribution of

glyceroneogenesis and glycolysis to the formation of triglyceride glycerol isolated from epididymal and mesenteric adipose

tissue depots were determined after 7 h of tracer incorporation as described under "Experimental Procedures." *, p < 0.05; **,

p < 0.01 vs. control.

58 animals, the direct contribution of glucose, via glycolysis, was negligible. In contrast, sucrose supplementation resulted in a doubling of the direct contribution of glucose to triglyceride glycerol in both adipose tissue depots.

We confirmed the predominance of glyceroneogenesis, as compared to glycolysis, by examining the [14C]/[3H] ratio on C-1 and (C-1 + C-3) of triglyceride glycerol. C-1 and (C-1 + C-3) of glycerol were cleaved by periodate oxidation and the radioactivity of the dimedon derivative measured. We examined the mesenteric adipose tissue of sucrose supplemented rats, since glyceroneogenesis was highest in the adipose tissue of this group. As shown in Table 3, the [14C]/[3H] ratio was high on (C-1 + C-3) as compared with C-1 of triglyceride glycerol. A high [14C]/[3H] ratio on C-3 (or C-1 + C-3) suggests a greater contribution of glyceroneogenesis, relative to the direct contribution of glucose, via glycolysis, to triglyceride glycerol synthesis (Fig. 5).

Fatty acid synthesis in the adipose tissue The incorporation of [14C] of glucose into fatty acids was negligible in 48 h fasted animals and high in controls in both the epididymal and mesenteric adipose tissue (Table 4). Furthermore, fatty acid synthesis in the sucrose supplemented group was significantly higher, as compared to control animals in both adipose tissue depots examined.

PEPCK-C activity in the adipose tissue Maximal PEPCK-C activity (0.33 units/g tissue) was noted in mesenteric fat of 48 h fasted rats, which was significantly higher than that of the sucrose supplemented group (0.025 units/g tissue) (Fig. 6). Similar qualitative changes were observed in the epididymal adipose depot (0.14 units/g tissue in

48 h fasted rats and 0.020 units/g tissue in the sucrose supplemented group). PEPCK-C

59 Relative [14C] and [3H] radioactivity and [ 14C]/[3H] ratio on C-1 and C-1 + C-3 of triglyceride glycerol in adipose tissue

C-1 + C-3 dpmratio C-1 dpm ratio Animal [3H] [14C] [14C]/[3H] [ 3H] [14C] [14C]/[3H] 1 1574 2948 1.87 1220 189 0.15 2 1893 2482 1.31 1255 133 0.11 3 1812 2593 1.43 1145 139 0.12 4 1186 1925 1.62 496 174 0.35 5 1099 1731 1.58 423 173 0.41

1.56 ± 0.095 0.23 ± 0.063** 60 Table 3. Relative [14C] and [3H] radioactivity and [14C]/[3H] ratio on C-1 and C-1 + C-3 of triglyceride glycerol in

adipose tissue. C-1 and C-1 + C-3 (along with their corresponding hydrogens) were cleaved from triglyceride glycerol using

periodic oxidation and radioactivity of the dimedon derivative was measured as described under "Experimental Procedures."

The [14C] and [3H] radioactivity was measured and the [14C]/[3H] ratio was calculated. Each row represents the values

obtained from an individual rat of the sucrose supplemented group (mesenteric adipose depot). The mean ± S.E. is reported

for the [14C]/[3H] ratio of C-1 and C-1 + C-3. *, p < 0.05; **, p < 0.01 vs. C-1 + C-3.

61 Incorporation of glucose carbon into de novo synthesized fatty acids in adipose tissue

Epididymis Mesenteric Glucose Incorporation (nmol/g/h) Control 520 ± 160 360 ± 120 (n=5) 48 h Fast 0.84 ± 0.51 N.D. (n=5) Suc Sup + Glc Inf 5984 ± 934* 11753 ± 1882* (n=5) 62 Table 4. Incorporation of glucose carbon into de novo synthesized fatty acids in adipose tissue. Values are the means ±

S.E. The contribution of glucose carbon (N.D., none detected) to the de novo synthesis of fatty acids was quantified using

incorporation of [14C] into triglyceride fatty acids during a prime constant-rate infusion of [U-14C]glucose. Fatty acids were

obtained after hydrolysis of triglyceride isolated from epididymal and mesenteric adipose tissue depots as described under

"Experimental Procedures." *, p < 0.05 vs. control. 63 0.4 0.33 0.35 0.3 0.25 0.2 0.14 0.15 0.1 * ** 0.020 0.025 0.05 PEPCK Activity (U/g tissue) (U/g PEPCK Activity 0 Epididymal Mesenteric 64 Figure 6. The activity of PEPCK-C in adipose tissue. Epididymal and mesenteric adipose tissue was collected from 48 h

fasted animals (grey bars) and from animals maintained on a sucrose supplemented diet (black bars). The activity of PEPCK

was determined in these tissues as described under "Experimental Procedures" and is expressed as the mean ± S.E. for 3 rats.

The unit of activity is defined as one µmole of substrate converted to product/min at 37°C. *, p < 0.05; **, p < 0.01 vs. 48 h

fast.

65 activity was significantly higher in mesenteric fat versus epidymal fat of 48 h fasted animals.

Response to enhanced TG-FA cycling by epinephrine Epinephrine infusion resulted

in an elevated concentration of plasma glucose and an enhanced rate of lipolysis, as

indicated by the increase in the concentration of free glycerol in the plasma (data not

shown). In the control rats infused with saline, there was no change in the plasma

glucose and glycerol concentration.

Infusion of epinephrine caused a significant increase in glyceroneogenesis in

epididymal adipose tissue (Table 5). Similarly, glyceroneogenesis in mesenteric adipose

tissue was significantly higher in the epinephrine infused rats (978 nmol/g/h) as

compared to the control animals (479 nmol/g/h). Total glucose carbon incorporated into

adipose tissue triglyceride glycerol was higher in rats infused with epinephrine, when

compared with controls (Table 5). However, after accounting for the contribution via

lactate to triglyceride glycerol, the direct contribution of glucose via glycolysis was

negligible in both depots (Table 5).

The relative contribution of glyceroneogenesis and glucose to triglyceride glycerol

synthesis in skeletal muscle The concentration of triglyceride in the gastrocnemius and

soleus muscles were not different among the three groups (Table 6). Glyceroneogenesis

was quantifiable in both gastrocnemius and soleus in vivo and was not different in the

two muscle types. The rate of glyceroneogenesis in the control animals was ~75

nmol/g/h (Table 6). Sucrose supplementation resulted in a significant increase in

glyceroneogenesis in the gastrocnemius muscle only.

66 The effect of epinephrine infusion on the contribution of glyceroneogenesis and glucose to triglyceride glycerol synthesis in adipose tissue

Epididymal Fat Mesenteric Fat

GlyceroneogenesisGlycolysis Glyceroneogenesis Glycolysis

Total Direct Total Direct (nmol/g/h)(nmol/g/h) (nmol/g/h) (nmol/g/h)

Control 367 ± 21.6 25.1 ± 1.6 (-21.6 - -7.4) 479 ± 38.2 31.7 ± 16.0 (-27.2 - -3.1) (n=4)

Epi Inf (500 ng/kg/min) 672 ± 10.3* 102 ± 17.7* (-44.3 - 5.0) 978 ± 182*144 ± 38.2 (-103 - 9.3) (n=4) 67 Table 5. The effect of epinephrine infusion on the contribution of glyceroneogenesis and glycolysis to triglyceride

glycerol synthesis in adipose tissue. Values are the means ± S.E.; range is indicated in parentheses. Glyceroneogenesis was

3 quantified using tritium, [ H2]O, incorporation into triglyceride glycerol and is expressed as nmol/g/h pyruvate incorporation.

The contribution of glycolysis was quantified using incorporation of [14C] into triglyceride glycerol, during a prime constant-

rate infusion of [U-14C]glucose. “Total” [14C] contribution of glycolysis includes three-carbon equivalents coming directly

from glucose plus those coming indirectly from [14C]lactate. “Direct” contribution represents three-carbon equivalents coming

directly from glucose only; the [14C] label arriving from [14C]lactate has been subtracted out. The [TG] (triglyceride

concentration) and the relative contribution of glyceroneogenesis and glycolysis to the formation of triglyceride glycerol

isolated from epididymal and mesenteric adipose tissue depots were determined after 3 h of tracer incorporation as described

under "Experimental Procedures." *, p < 0.05 vs. control. 68 The contribution of glyceroneogenesis and glycolysis to triglyceride glycerol synthesis in skeletal muscle

Gastrocnemius Soleus

[TG] GlyceroneogenesisGlycolysis [TG] Glyceroneogenesis Glycolysis

Total Direct Total Direct (µmol/g) (nmol/g/h) (µmol/g) (nmol/g/h) (nmol/g/h) (nmol/g/h) Control 18.3 ± 2.3 5.7 ± 0.375.1 ± 9.3 19.5 ± 2.8 (-3.5 - 2.0) 6.4 ± 0.3 78.2 ± 16.3 (-3.5 - 4.0) (n=5) 48 h Fast (-10.5 - -2.0) 6.3 ± 0.4 10.9 ± 1.8 5.3 ± 0.6 118 ± 18.1 127 ± 21.0 (-10.5 - -0.5) (n=5) 12.0 ± 0.5 Suc Sup + Glc Inf 25.8 ± 1.5 (-3.0 - 2.5) 5.1 ± 0.3 31.7 ± 3.1* 5.4 ± 0.2 109 ± 4.0* 104 ± 3.2 (-3.0 - 1.5) (n=5) 69 Table 6. The contribution of glyceroneogenesis and glycolysis to triglyceride glycerol synthesis in skeletal muscle.

3 Values are the means ± S.E.; range is indicated in parentheses. Glyceroneogenesis was quantified using tritium, [ H2]O,

incorporation into triglyceride glycerol and is expressed as nmol/g/h pyruvate incorporation. The contribution of glycolysis

was quantified using incorporation of [14C] into triglyceride glycerol, during a prime constant-rate infusion of [U-14C]glucose.

“Total” contribution of glycolysis includes three-carbon equivalents coming directly from glucose plus those coming indirectly

from [14C]lactate. “Direct” contribution represents three-carbon equivalents coming directly from glucose only; the [14C] label

arriving from [14C]lactate has been subtracted out. The [TG] (triglyceride concentration) and the relative contribution of

glyceroneogenesis and glycolysis to the formation of triglyceride glycerol isolated from gastrocnemius and soleus muscles

were determined after 7 h of tracer incorporation as described under "Experimental Procedures." *, p < 0.05. 70 Total glucose carbon incorporated into triglyceride glycerol was ~20 nmol/g/h in

the gastrocnemius and soleus muscles of control rats (Table 6). Fasting for 48 h did not change the total glucose carbon incorporated into triglyceride glycerol in the skeletal muscle. In response to sucrose supplementation, a significant increase in total glucose incorporated into triglyceride glycerol was seen only in the soleus. After accounting for the contribution of lactate to triglyceride glycerol, the direct synthesis of triglyceride

glycerol from glucose via glycolysis was negligible in both the gastrocnemius and soleus

from all three groups of animals (Table 6).

71 CHAPTER 4: Discussion

I have examined the relative contribution of glyceroneogenesis and glucose, via

glycolysis, to triglyceride glycerol synthesis in the rat. My data show that

glyceroneogenesis is quantitatively the predominant pathway for triglyceride glycerol

synthesis in white adipose tissue, skeletal muscle, and liver during extended fasting, as

well as during periods of glucose availability. Surprisingly, the highest rates of

glyceroneogenesis in adipose tissue were observed in sucrose supplemented animals,

when fatty acid synthesis and triglyceride deposition were high.

Significance of current methodology Measurements of glyceroneogenesis in vivo,

have been carried out in both rodents and humans (33;48;49;69;90-94). The difficulty in

accurately quantifying this pathway lies in the fact that glucose can be metabolized to

lactate/pyruvate in peripheral tissues and subsequently taken up by adipose tissue (and

skeletal muscle) as a substrate for glyceroneogenesis. Botion and colleagues (48) were

the first group to measure glyceroneogenesis in vivo. They estimated glyceroneogenesis

by subtracting the triglyceride glycerol synthesis rate obtained with [U-14C]glucose

3 3 (glycolytic flux) from the rate obtained with [ H2]O (which results in [ H] labeling of the

triose phosphate pool and estimates triglyceride glycerol synthesis from all carbon sources). As pointed out by Botion et al. (48), this approach does not account for the

[14C] label incorporated into triglyceride glycerol via glyceroneogenesis, as it was

assumed that no appreciable turnover of [3H] or [14C]-labeled product occurred during the

experimental time period employed (1 h), and thus results in a minimal estimation of glyceroneogenesis.

72 In the present study, we accounted for [14C]lactate incorporation into triglyceride

glycerol via glyceroneogenesis. More specifically, by multiplying the rate of

3 14 glyceroneogenesis estimated by [ H2]O with the specific activity of [ C]lactate, and

subtracting [14C] incorporated via glyceroneogenesis from total [14C] incorporation into

triglyceride glycerol, we estimated triglyceride glycerol synthesis from glucose.

Although our estimates are true representations of glyceroneogenesis in vivo, we found

the contribution of glucose to be negligible. This was further confirmed when we

examined the ratio of [14C]/[3H] on C-1 alone versus the ratio obtained for (C-1 + C-3).

We anticipated that the ratio of [14C]/[3H] on C-1 of triglyceride glycerol from pyruvate

would be low relative to the triosephosphate formed from glucose. Formation of PEP

from pyruvate, via OAA, will result in the incorporation of unlabeled carbon from the bicarbonate pool (pyruvate carboxylase and CO2 fixation) onto the C-1 position of PEP.

This is what we observed, as the ratio of [14C]/[3H] on C-1 alone was a fraction (< 1) of

the ratio obtained for (C-1 + C-3), indicating that the carbon present in triglyceride

glycerol arrived indirectly via pyruvate.

These data suggest that the major sources of carbon in triglyceride glycerol are

pyruvate, lactate, alanine (or the carbon skeletons of any of other gluconeogenic amino

acid) via glyceroneogenesis. The negative values for the contribution of glucose to

triglyceride glycerol synthesis under certain dietary conditions could be due to the

following factors: First, we determined the [14C]lactate SA after its isolation from the

plasma. Although we are confident in the accuracy of our methods, a slight

underestimation of the concentration of lactate could result in an overestimation of

[14C]lactate SA. A higher [14C]lactate SA will result in an overestimation of the

73 glyceroneogenic flux from [14C]lactate. Second, the dilution factor (2.2) for the loss of

label in the citric acid cycle was based on literature values for the liver (110); we could

not find a similar estimate for the loss of label in adipose tissue. Therefore, the value we

have used may underestimate the dilution in the adipose tissue, since glyceroneogenic

flux relative to citric acid cycle flux in this tissue is far less than gluconeogenic flux.

relative to citric acid cycle flux in the liver. The smaller dilution factor will result in

overestimation of glyceroneogenesis from [14C]lactate. Third, we assume that all five

hydrogens of triglyceride glycerol are equally labeled and that both labeled hydrogens of

C-3 are derived from pyruvate. Since the magnitude of [3H] labeling of hydrogens on C-

3 will depend upon the contribution of pyruvate to triglyceride glycerol, this would result in an overestimation of [3H] on C-3, and thus, the contribution of glyceroneogenesis.

Glyceroneogenesis in the adipose tissue We have quantified glyceroneogenesis in adipose tissue in response to fasting, sucrose supplementation, and intravenous infusion of epinephrine. Our data show that during fasting there was no change in the rate of glyceroneogenesis, as compared to control values and the direct contribution of glucose to triglyceride glycerol was negligible. Glyceroneogenesis is thus likely to be critical for

TG-FA cycling, which requires the generation of G-3-P for triglyceride synthesis, particularly under conditions where the TG-FA cycling is increased (112). TG-FA cycling has been shown to be high in humans fasted for 96 h (113); we could not find any corresponding data on TG-FA cycling in the rat. However, there was a 50% lower rate of

glucose uptake by adipose tissue, as measured by a-v difference (17) in rats that were

fasted for 48 h. Fasting is associated with a lower expression of GLUT4 (18;19) and a

decreased uptake and phosphorylation of 2-deoxyglucose by adipocytes in vitro (20).

74 Since PEPCK-C activity increases in the adipose tissue of fasted rats (25), we anticipated

that glyceroneogenesis would be increased following 48 h of starvation, in order to

provide G-3-P for TG-FA cycling. Although not measured, the TG-FA cycling may

already be high in control animals (~8 h fast), so that starvation for 48 h did not cause a

further increase in TG-FA cycling and therefore no increase in glyceroneogenesis.

Both high carbohydrate feeding and intravenous glucose administration cause an

increase in the concentration of plasma insulin, increase in glucose uptake by the adipose

2 tissue, and promote lipogenesis (16;43;50;52). [ H2]O tracer studies with mice, showed

that glucose was the predominant source of triglyceride glycerol in both visceral and epididymal adipose tissue in animals that were fed a high carbohydrate diet for 75 days

(91). We thus expected glyceroneogenesis would be lower, and the contribution of glucose to triglyceride glycerol higher, in the rats maintained on a sucrose supplemented diet. The direct contribution of glucose to triglyceride glycerol synthesis was higher than controls in response to a sucrose supplemented diet, although glyceroneogenesis also increased and remained the predominant pathway for triglyceride glycerol synthesis

(Table 2). In addition, a 90% increase in lipogenesis was observed in the adipose tissue.

Wolfe and Peters (50) demonstrated that lipolysis continued to occur in humans when glucose was infused at 8 mg/kg/min and that the rate of lipolysis was significantly reduced due to enhanced intracellular TG-FA cycling (50). Therefore, the observed increase in glyceroneogenesis in our study may support triglyceride glycerol synthesis for enhanced intracellular TG-FA cycling, as well as the esterification of de novo synthesized fatty acids.

75 It is possible that a considerable fraction of the labeled fatty acids came from lipids that were synthesized in the liver as a consequence of sucrose feeding (43) and transported to the adipose tissue. The rate of lipogenesis has been shown to decrease markedly in the adipose tissue of rats of similar age as those used in our study (44;45).

Since we detected [14C] radioactivity in fatty acids isolated from plasma triglyceride of sucrose supplemented animals, such a possibility cannot be excluded. Despite the source of the newly synthesized fatty acids, the question remains as why glucose-derived carbon does not appear in significant quantities in triglyceride glycerol, since glucose is most likely being utilized for lipogenesis by the adipose tissue.

The results of the research presented in this thesis are the first to show that glyceroneogenesis is a major contributor to triglyceride glycerol synthesis in response to high carbohydrate feeding; during net triglyceride deposition and decreased systemic recycling. The rate of high glyceroneogenesis in sucrose fed animals noted in our study is at variance with the findings of Chen et al. (91) who found a lower fractional contribution of glyceroneogenesis and a higher contribution of glucose in mice fed a high carbohydrate diet for 75 days. These differences may be related to the short duration of our study, the specific difference in dietary regimen (i.e., high carbohydrate and low fat diet versus sucrose supplementation followed by glucose infusion in our study), and the tracer method employed. Chen et al. (91) developed a novel technique for determining the relative glyceroneogenic contribution to triglyceride glycerol using a probability parameter (n). In contrast to our approach, they do not assume equal labeling of hydrogens of the triose pool; their calculations are based upon the unique labeling pattern of triglyceride glycerol, which reflects the source of G-3-P: 3.5 hydrogens will be

76 labeled if G-3-P is from glycolysis (n=3.5); 5 hydrogens will be labeled if G-3-P is from glyceroneogenesis (n=5.0). Since their calculations are based upon the theoretical plateau for fully labeled glycerol, they use long-term label incorporation (15 or 64 days).

No difference was observed in n values when comparing the high carbohydrate vs. low carbohydrate diets (in both epididymal and mesesteric adipose tissue) after 26

2 days. By 75 days on the diet (64 days on [ H2]O), n was significantly higher

(glyceroneogenesis) in all adipose depots on the low carbohydrate diet. However, as time progresses, there is increased label incorporation and all molecular species become labeled. This makes identification of the original source of triglyceride glycerol more difficult to determine. Chen et al. (91) point out that due to the ability of n to change, experimental measurement of n by this technique is optimum when calculating triglyceride glycerol fractional synthesis, particularly under new experimental conditions.

It must be pointed out that the approach of Chen et al. (91) account for unlabeled glucose acquiring a labeled hydrogen on C-1 from the body water upon entry of glucose carbon into the triose phosphate pool (phosphoglucose isomerase step). This will result in labeling of one of the two hydrogens present on C-3 of DHAP (and thus G-3-P).

Therefore, our calculations would overestimate glyceroneogenesis by 25%. If we adjust our quantitative flux rates of glyceroneogenesis to account for such an overestimation, under conditions of glucose infusion, glyceroneogenesis would account for 53%

(epididymal) and 49% (mesenteric), with glucose via glycolysis (uncorrected values) contributing the remaining fraction. Thus, glyceroneogenesis still remains a significant contributor to triglyceride glycerol (~50%) in adipose tissue of rats fed a high carbohydrate diet.

77 In contrast, Chen et al. (91) shows that glyceroneogenesis accounts for 20%

(epididymal) and 16% (mesenteric) of triglyceride glycerol synthesis under high carbohydrate conditions. They site that the majority (83%) of triglyceride glycerol is derived from glycolysis and emphasize that glycolytic cycling of glucose to the level of pyruvate or OAA and back to G-3-P will contribute to triglyceride glycerol, thus complicating estimates of the proportion derived from glycolysis. The methodology of

Chen et al. (91) cannot correct for glycolytic cycling. However, we address this issue by accounting for glucose carbon that arrived in triglyceride glycerol indirectly via lactate.

Yet even if the majority of our glyceroneogenic flux rates are due to glycolytic cycling, it is of great interest that glucose proceeding down the glycolytic pathway would not directly go to G-3-P at the level of the triose phosphate pool, but instead proceed to the level of pyruvate/lactate, then, via glyceroneogenesis, back to G-3-P.

Response to epinephrine infusion The effect of enhanced TG-FA cycling on glyceroneogenesis was examined by determining the response of adipose tissue to epinephrine infusion. Epinephrine, a β-adrenegric agonist, increases TG-FA cycling in humans (95) and decreases glucose uptake by the adipose tissue (114). In addition, infusion of epinephrine causes an elevation in the concentration of lactate in the blood

(115). We anticipated that glyceroneogenesis would increase in response to epinephrine, since glucose uptake by adipose tissue is decreased. Epinephrine infusion of 500 ng/kg/min for 3 h resulted in enhanced glyceroneogenesis and no measurable contribution of glucose to triglyceride glycerol formation. This suggests that glyceroneogenesis is operative under conditions of enhanced TG-FA cycling and that lactate may provide a source of carbon for glyceroneogenesis.

78 A computational model for adipose tissue metabolism The increase in

glyceroneogenesis we observed in response to epinephrine infusion supports a

computational model of Kim et al. (116) to analyze key components of adipose tissue

metabolism. To better understand the factors that control the relative contribution of

glucose and pyruvate to G-3-P synthesis with the increased rate of TG-FA cycling, Kim et al. (116) developed a computational model of adipose tissue metabolism in humans during intravenous infusion of epinephrine. This model examined the effect of varying contribution of glucose and pyruvate in the basal state. While the rate of FFA production via lipolysis increased by 4-fold, the rate of intracellular re-esterification increased by up to 13% during epinephrine infusion. In the absence of changes in other hormone levels during epinephrine infusion (116), the re-esterification rate was primarily regulated by the availability of substrates. Model simulation showed that the increased rate of G-3-P synthesis occurred, with a greater contribution of glyceroneogenesis, regardless of its relative contribution in the basal state. It was also shown that the increased production of

FFA, due to the stimulated rate of lipolysis, increased the levels FA CoA by up to ~80%.

The subsequent ß-oxidation of FA CoA resulted in the increased ratio of acetyl CoA to free CoA, thus inhibiting the oxidation of pyruvate. Therefore, more pyruvate would be available for the synthesis of G-3-P, thereby increasing the flux through

glyceroneogenesis.

Since their model also indicated a lack of increase in the rate of glucose uptake

during epinephrine infusion, it raised a question about the effect of stimulated glucose

uptake on glyceroneogenesis. Therefore, the model of adipose tissue metabolism was

modified to simulate the physiological responses during euglycemic-hyperinsulinemic

79 clamp in humans (unpublished data, Kim, Saidel and Kalhan) by incorporating additional

metabolic pathways and their regulation by insulin. The model showed good agreement

with the physiological data from in vivo human studies (117;118). When the rate of

glucose uptake by the adipose tissue increased ~6-fold, there was ~20% increase in the

rate of G-3-P synthesis from both glucose and pyruvate. The model simulations indicated

that the decrease in redox state (i.e. NAD/NADH ratio) provided the major driving force

for increasing the synthesis of G-3-P, in addition to the higher availability of precursors.

Since the rate of glyceroneogenesis showed a similar increase, even with the stimulated

rate of glycolysis, this model suggests that glyceroneogenesis plays a significant role in

the regulation of re-esterification of FFA in response to physiological perturbations.

Glucose uptake and glyceroneogenesis by the adipose tissue: A two-cell

hypothesis Our data show that glucose uptake and glyceroneogenesis occur

simultaneously in the adipose tissue of control animals and was enhanced in sucrose

supplemented animals. The key issues of this study are as follows: 1) despite its uptake

by adipose tissue, glucose carbon is not going directly to G-3-P for triglyceride glycerol synthesis; and 2) glucose uptake is occurring despite the high rates of glyceroneogenesis in the tissue. The simultaneous occurrence of glucose uptake and glyceroneogenesis suggests either intracellular compartmentation of G-3-P in two different non-mixing pools, or the existence of two different cell types in the adipose tissue, a predominantly glycolytic cell and a predominantly triglyceride storing cell (Fig. 7). Currently, there are no data available that would support compartmentation of the pathways of triglyceride synthesis in a single cell type in adipose tissue that could readily explain our results. In

80 Glycolytic Cell Mature Adipocyte

Glucose

G-6-P TG

+ 3 FA CoA Triose G-3-P Triose

Pyruvate Alanine Pyruvate Pyruvate Lactate Lactate 81 Figure 7. The two compartmental (or two cell-type) hypothesis. Two different cell types exist within an adipose tissue

depot: 1) undifferentiated adipocytes which are not depositing triglyceride and are predominantly glycolytic (left side); 2)

differentiated adipocytes which display the phenotype characteristic of mature adipocytes (namely a large triglyceride

reservoir) and are depositing triglyceride (right side). The differentiated adipocytes may rely predominantly on

glyceroneogenesis for the deposition of triglyceride. 82 contrast, evidence of multiple cell types in the adipose tissue is well documented, thus lending support to a two-cell hypothesis.

Adipose tissue is a dynamic organ that includes a mixture of cell types. In addition to adipocytes, it contains endothelial cells and macrophages that primarily metabolize glucose to lactate (119;120). Furthermore, undifferentiated pre-adipocytes do not deposit triglycerides and may be primarily glycolytic; recent evidence indicates that there are least two different types of pre-adipocytes (121). In addition, PEPCK-C expression is a late event, occurring during terminal differentiation of the adipocyte (122)

Mature adipocytes are fully differentiated cells that exhibit an adipocyte-like phenotype, namely a large reservoir of triglyceride. Smith (58) has proposed that mature adipocytes range from young to old, and are insulin sensitive to insulin resistant, respectively. Each of the cell types may exhibit different patterns of substrate utilization, depending on the activity of the respective metabolic pathway predominating in the cell type relative to time, hormonal environment, and levels of nutrients.

Our data are a composite of the contribution of glyceroneogenesis and glucose metabolism in all of the cells types present in the adipose tissue. It is possible that the glycolytic cells metabolize glucose to lactate, which is then released into the interstitium where it is available as a glyceroneogenic substrate for the triglyceride-storing cell type.

The differentiated adipocytes, which express PEPCK-C, may rely predominantly on glyceroneogenesis for the deposition of triglyceride. After an overnight fast, the interstitial concentration of lactate is significantly higher in abdominal and femoral subcutaneous adipose tissue than in plasma of humans, indicating a local release of lactate (123). In response to an oral glucose load, both plasma and interstitial lactate

83 levels increased in parallel; however the interstitial lactate concentration was much greater than that in the plasma (123). In addition, adipose tissue expresses the gene for the monocarboxylate transporter-1, which may be responsible for both lactate release as well as uptake from the interstitium of adipose tissue (124). Our data showing the minor contribution of glucose to triglyceride glycerol synthesis are consistent with studies where glucose uptake is compromised by a GLUT-4 knockout in adipose tissue (125).

Mice with an adipose tissue specific-depletion of GLUT-4 maintain their adipose tissue triglyceride mass, suggesting that reduced glucose transport did not impair the overall esterification of fatty acids to triglyceride. In contrast, increased glucose metabolism, induced by over-expression of hepatic glucokinase in the adipose tissue of mice, resulted in higher glucose uptake and higher lactate production, without an increase in triglyceride synthesis (126). These studies are consistent with the two-cell hypothesis and with the compartmentation of glyceroneogenesis in the adipose tissue.

PEPCK-C activity in adipose tissue Chronic stimulation by transcriptional activators such as glucagon and fatty acids results in the up-regulation of PEPCK-C, whereas long term exposure to insulin or glucose serves to down-regulate PEPCK-C (19).

Therefore, under the chronic conditions employed in this study, where animals were fasted for 48 h or maintained on a sucrose supplemented diet for five days followed by a

7 h glucose infusion, it is expected that PEPCK-C protein levels, and thus activity, would be dramatically up-regulated or down-regulated, respectively. Accordingly, our data demonstrate that PEPCK-C activity was significantly lower in both epididymal and mesenteric tissue of rats exposed to chronic glucose availability compared to 48 h fasted animals (Figure 6).

84 There is ample evidence that the level of PEPCK-C activity influences the overall rate of glyceroneogenesis. Over-expression of PEPCK-C in adipose tissue caused an increased triglyceride deposition and obesity in the mice (70). The over-expression of

PEPCK-C in skeletal muscle of the mouse results in animals which have up to 10 times the triglyceride in the muscle than do controls (23). This indicates that high levels of

PEPCK-C in tissues can accelerate triglyceride synthesis (presumably via glyceroneogenesis). The total rate of glyceroneogenesis in adipose tissue from humans was markedly decreased as the BMI of the individual increased, suggesting that obesity causes a dysfunction of glyceroneogenesis (65). The activity of PEPCK-C in human adipose tissue explants correlated with the rate of glyceroneogenesis measured by

[14C]pyruvate incorporation into triglyceride glycerol in vitro. Recently, Chang et al.

(127) reported that there is a direct relationship between the body mass index of non-

diabetic women, but not with insulin resistance as measured by HOMA, and the level of

PEPCK-C mRNA in subcutaneous adipose tissue. This suggests that triglyceride

synthesis in adipose tissue is related to the level of expression of the gene for PEPCK-C

(the authors did not measure PEPCK-C activity in the tissues under investigation).

In contrast, we did not note a direct correlation between the activity of PEPCK-C

in the adipose tissue of the rats and the rate of glyceroneogenesis as determined by our

isotopic labeling techniques. However, the activity of PEPCK-C under all dietary

conditions studied was greater than the rate of glyceroneogenesis, so that carbon flux was

not limited by the activity of the enzyme. The most crucial aspect of glyceroneogenic

flux may be the provision of substrate (both pyruvate and fatty acids). In support of this

concept, upregulation of PEPCK-C activity in adipose tissue (high protein diet) results in

85 higher values of glyceroneogenesis only under conditions where there is a significant

increase in both glyceroneogenic and fatty acid substrates available (48;68). Since

PEPCK-C is never completely suppressed, and triglyceride synthesis is necessary under all physiological conditions (i.e., recycling of FFA to triglyceride during fasting, for deposition of dietary fat, and for esterification of de novo synthesized FA) glyceroneogenic flux will remain quantitatively important.

PEPCK-C may play additional metabolic roles other than supporting glyceroneogenesis in the adipose tissue and other tissues in which this pathway is active.

For example, deletion of the gene for PEPCK-C in the liver results in a marked reduction of citric acid cycle flux (78), presumably due to an altered rate of cataplerosis in the tissue. Furthermore, studies investigating rat adipose tissue amino acid metabolism in vivo demonstrate that adipose tissue is a site of glutamine synthesis in both fed and starved animals (17). These authors suggest that the principal substrates are amino acids derived from intracellular proteolysis, with glutamate and aspartate providing nearly all of the carbon required for the glutamine release observed (17). Thus, aspartate amino transferase may support an anaplerotic role of providing OAA to the TCA cycle for subsequent glutamine synthesis, while PEPCK-C may support a cataplerotic role for

removal of OAA to keep the cycle balanced. These areas will require further study.

Glyceroneogenesis in the skeletal muscle This is the first demonstration that

glyceroneogenesis occurs in skeletal muscles. In response to fasting, as well as sucrose

feeding, glyceroneogenesis was the main contributor to triglyceride glycerol formation,

whereas the direct contribution of glucose was not measurable. The low incorporation

noted in this study is, in part, due to the relatively small triglyceride pool in skeletal

86 muscle; the IMTG pool is typically a few micromoles/g of muscle (128). During fasting,

glyceroneogenesis would be expected to provide the G-3-P for triglyceride glycerol synthesis, since insulin-mediated GLUT-4 translocation and glucose uptake by the skeletal muscle is decreased (129). Furthermore, TG-FA cycling has been documented to be active in oxidative muscle of overnight fasted rats (35). Although we anticipated that glyceroneogenesis would be a functional pathway, due to the presence of PEPCK-C

activity in skeletal muscle (23), the predominance of this pathway was unexpected. Even

more surprising was the lack of direct contribution of glucose to G-3-P synthesis, given

that in response to an oral glucose load, skeletal muscle is responsible for the majority

(~85%) of insulin-mediated glucose uptake (12). Our data demonstrating a marginal

contribution of glucose to triglyceride glycerol in skeletal muscle are consistent with the

report of Guo and Jensen (37). These researchers originally attempted to quantify the

contribution of glucose to triglyceride glycerol using [6-3H]glucose tracer. However,

they found almost no incorporation of [3H] from glucose into triglyceride glycerol. The

[3H] label on [6-3H]glucose is lost when this tracer proceeds to the level of

pyruvate/lactate since the methyl hydrogens on C-3 are in rapid equilibrium with the

surrounding unlabeled pool of body water. This unlabeled molecule may then be utilized

as a substrate for the glyceroneogenic pathway, which would result in no [3H] label detection in isolated triglyceride glycerol. Using [6-14C]glucose, they were able to detect

[14C] incorporation into triglyceride glycerol. They found ~20% of triglyceride glycerol

was derived from glucose. The [14C] label could have arrived directly from [6-

14C]glucose or indirectly via [14C]lactate. Since no glucose contributed to triglyceride

glycerol formation using [6-3H]glucose, the [14C] label arrived from [14C]lactate.

87 Skeletal muscle includes both fast-twitch, glycolytic fibers and slow-twitch,

oxidative fibers. Higher levels of IMTG are present in skeletal muscle containing a higher proportion of oxidative fibers, such as the soleus. In contrast, the gastrocnemius contains a predominance of fast-twitch, glycolytic fibers (130), so that triglyceride glycerol synthesis from glucose in the gastrocnemius would be expected. The majority of glucose from an oral glucose load is taken up by the muscle, metabolized to lactate and released into the plasma, where it is used as fuel by other tissues or is converted to

glucose by the liver (see reference (131) for a review). Similar to what was observed in

the adipose tissue, the interstitial concentration of lactate measured in skeletal muscle in

both the fed and fasted states remains elevated as compared to the plasma (123;132). In

addition, lactate and pyruvate are exchanged across the sarcolemmal membranes by

facilitated transport involving specific monocarboxylate transporters (133;134). Thus, a

two-cell hypothesis could apply to the skeletal muscle where fast-twitch fibers

predominantly metabolize glucose to lactate, thereby providing a glyceroneogenic

substrate for adjacent slow-twitch fibers, containing higher IMTG levels (135).

Hepatic glyceroneogenesis The fractional contribution of gluconeogenesis to the glucose Ra responds as expected to the different dietary conditions (lower in controls

(~30 %) and increased to ~60 % after a 48 h fast), while glyceroneogenesis remained constant at about ~60 % under all conditions studied. Furthermore, glyceroneogenesis and not glucose metabolism via glycolysis was the predominant pathway of triglyceride glycerol synthesis in all dietary conditions studied. Glycolysis, like gluconeogenesis, was responsive to the different physiological conditions; changes in the fractional contribution of glucose to triglyceride glycerol synthesis were noted, but the overall range was from

88 10 – 30 % (48 h fasted vs. sucrose supplementation followed by glucose infusion). This is quite surprising, since exogenous glucose infusion would be expected to increase hepatic glucose uptake and glucose should then become the major source of G-3-P.

These results may indicate that levels of exogenous glucose infusion were not high enough to increase hepatic glucose uptake to a point where glucose,via glycolysis, would become the major source of G-3-P. If the rate of infusion of glucose were increased, a corresponding increase in glucose uptake and its subsequent conversion to triglyceride glycerol might have been observed. However, in response to sucrose supplementation followed by glucose infusion, hepatic glucose production was halted and gluconeogenesis was negligible. Furthermore, Kalhan et al. (90) have demonstrated that a normoglycemic-hyperinsulinemic clamp did not alter the relative contribution of pyruvate or glucose to triglyceride glycerol.

Since hepatic gluconeogenesis and glyceroneogenesis share many of the same enzymes, our data suggest that gluconeogenesis is regulated at a step which is not in common with glyceroneogenesis, most likely at a point above the triose phosphate level, potentially at fructose-1,6-bisphosphatase (90). This enzyme is allosterically regulated by fructose-2,6-bisphosphate and AMP (negative regulators) and ATP (a positive regulator); this regulation prevents futile cycling (136;137). During fasting, the concentration of fructose-2,6-bisphosphate decreases due to the activation of the phosphatase function of the bi-functional enzyme, 6-phosphofructo-2-kinase/fructose-

2,6-bisphosphatase, by cAMP (138). Increased levels of glucagon, and thus hepatic cAMP and ATP, would be expected to occur in 48 h fasted animals, resulting in negative allosteric regulation of the glycolytic enzyme phosphofructokinase, which synthesizes

89 fructose-1,6-bisphosphate, thereby increasing gluconeogenesis (139). Metabolic flux over the two pathways reflects allosteric regulation of key enzymes, such as phosphofructokinase and fructose-2,6-bisphosphatase, as well as the availability of substrate.

Summary and remaining questions The present study clearly demonstrates that high rates of glyceroneogenesis occur in three major tissues of the rat: the adipose tissue, skeletal muscle, and liver. As mentioned above, this pathway may play a significant role in the control of triglyceride turnover and be a potential target for the treatment of obesity and diabetes (140), emphasizing the importance of PEPCK-C in both carbohydrate and lipid metabolism. However, there are fundamental metabolic questions concerning the pathway of glyceroneogenesis that need further study.

What is the source of carbon for glyceroneogenesis? First, the carbon source for the G-3-P in adipose tissue and skeletal muscle is not well understood. Our research demonstrates that pyruvate is clearly the predominant source of carbon for glyceroneogenesis under normal physiological conditions ranging from extended fasting to chronic feeding. However, the relative contribution of molecules which enter the glyceroneogenic pathway, via pyruvate, has yet to be examined. Lactate is clearly a major precursor of triglyceride glycerol via glyceroneogenesis, but other intermediates, such as alanine and the carbon skeletons of gluconeogenic amino acids that enter the citric acid cycle are also potential candidates (79). We may speculate that during fasting, alanine, via pyruvate, is the major source of carbon for G-3-P synthesis. Whereas in response to an oral glucose load, lactate (derived from glucose) enters the glyceroneogenic pathway by way of pyruvate, and is the predominant carbon source.

90 What is the source of reducing equivalents for glyceroneogenesis? Another major

metabolic question concerning glyceroneogenesis is the source of the NADH for the

synthesis of G-3-P. Glyceroneogenesis requires two molecules of NADH in the cytosol

for every molecule of G-3-P synthesized; one is required at the glyceraldehyde-3-

phosphate dehydrogenase step and the other at glycerol-3-phosphate dehydrogenase.

One molecule of NADH is produced via cytosolic NAD-malate dehydrogenase when

malate is oxidized to OAA. The source of the other molecule is not clear for any

intermediate other than lactate, which generates an additional reducing equivalent in the

cytosol when it is oxidized to pyruvate via lactate dehydrogenase. It is possible that the

malate-aspartate shuttle produces net cytosolic NADH, essentially bringing reducing

equivalents from the mitochondria to the cytosol to support glyceroneogenesis or that

DHAP is reduced to G-3-P via the flavoprotein dehydrogenase in the inner mitochondrial membrane (i.e. a reversal of the G-3-P shuttle). In the latter case, the electrons for the synthesis of NADH would be generated in the mitochondria by fatty acid oxidation.

Alternatively, triglyceride may be synthesized using DHAP and not G-3-P as a source of triglyceride glycerol. Dihydroxyacetone phosphate acyltransferase (EC 2.3.1.42) is present in a broad variety of organisms (141;142), including humans (143) and may be an active participant in glyceroneogenesis. If so, it would preclude the need for the second molecule of NADH. Further research in this area will be required to resolve many of the important metabolic questions outlined above.

91 REFERENCES

1. Arner,P. 2003. The adipocyte in insulin resistance: key molecules and the impact of the thiazolidinediones. Trends Endocrinol. Metab 14:137-145.

2. Arner,P. 2002. Insulin resistance in type 2 diabetes: role of fatty acids. Diabetes Metab Res. Rev. 18 Suppl 2:S5-S9.

3. Robinson,J., and Newsholme,E.A. 1969. Some properties of hepatic glycerol kinase and their relation to the control of glycerol utilization. Biochem. J. 112:455-464.

4. Heinz,F., Lamprecht,W., and Kirsch,J. 1968. Enzymes of fructose metabolism in human liver. J. Clin. Invest 47:1826-1832.

5. Lin,E.C. 1977. Glycerol utilization and its regulation in mammals. Annu. Rev. Biochem. 46:765-795.

6. Knight,B.L., and Myant,N.B. 1970. A comparison between the effects of cold exposure in vivo and of noradrenaline in vitro on the metabolism of the brown fat of new-born rabbits. Biochem. J. 119:103-111.

7. Burch,H.B., Lowry,O.H., Meinhardt,L., Max,P., Jr., and Chyu,K. 1970. Effect of fructose, dihydroxyacetone, glycerol, and glucose on metabolites and related compounds in liver and kidney. J. Biol. Chem. 245:2092-2102.

8. Newsholme,E.A., and Taylor,K. 1969. Glycerol kinase activities in muscles from vertebrates and invertebrates. Biochem. J. 112:465-474.

9. Robinson,J., and Newsholme,E.A. 1967. Glycerol kinase activities in rat heart and adipose tissue. Biochem. J. 104:2C-4C.

10. Jungas,R.L., and Ball,E.G. 1963. Studies on the metabolism of adipose tissue. XII. The effects of insulin and epinephrine on free fatty acid and glycerol production in the presence and absence of glucose. Biochemistry 2:383-388.

11. DeFronzo,R.A., Ferrannini,E., Hendler,R., Wahren,J., and Felig,P. 1978. Influence of hyperinsulinemia, hyperglycemia, and the route of glucose administration on splanchnic glucose exchange. Proc. Natl. Acad. Sci. U. S. A 75:5173-5177.

12. DeFronzo,R.A., Jacot,E., Jequier,E., Maeder,E., Wahren,J., and Felber,J.P. 1981. The effect of insulin on the disposal of intravenous glucose. Results from indirect calorimetry and hepatic and femoral venous catheterization. Diabetes 30:1000- 1007.

92 13. Baron,A.D., Brechtel,G., Wallace,P., and Edelman,S.V. 1988. Rates and tissue sites of non-insulin- and insulin-mediated glucose uptake in humans. Am. J. Physiol 255:E769-E774.

14. Frayn,K.N., Coppack,S.W., Humphreys,S.M., and Whyte,P.L. 1989. Metabolic characteristics of human adipose tissue in vivo. Clin. Sci. (Lond) 76:509-516.

15. Ferre,P., Leturque,A., Burnol,A.F., Penicaud,L., and Girard,J. 1985. A method to quantify glucose utilization in vivo in skeletal muscle and white adipose tissue of the anaesthetized rat. Biochem. J. 228:103-110.

16. Kotani,K., Peroni,O.D., Minokoshi,Y., Boss,O., and Kahn,B.B. 2004. GLUT4 glucose transporter deficiency increases hepatic lipid production and peripheral lipid utilization. J. Clin. Invest 114:1666-1675.

17. Kowalski,T.J., Wu,G., and Watford,M. 1997. Rat adipose tissue amino acid metabolism in vivo as assessed by microdialysis and arteriovenous techniques. Am. J. Physiol 273:E613-E622.

18. Berger,J., Biswas,C., Vicario,P.P., Strout,H.V., Saperstein,R., and Pilch,P.F. 1989. Decreased expression of the insulin-responsive glucose transporter in diabetes and fasting. Nature 340:70-72.

19. Ezaki,O. 1997. Regulatory elements in the insulin-responsive glucose transporter (GLUT4) gene. Biochem. Biophys. Res. Commun. 241:1-6.

20. Bay,U., and Froesch,E.R. 1986. Glucose uptake and phosphorylation in fat cells of fasted and hypophysectomized rats. Mol. Cell Endocrinol. 47:217-224.

21. Reshef,L., Hanson,R.W., and Ballard,F.J. 1969. Glyceride-glycerol synthesis from pyruvate. Adaptive changes in phosphoenolpyruvate carboxykinase and pyruvate carboxylase in adipose tissue and liver. J. Biol. Chem. 244:1994-2001.

22. Reshef,L., and Hanson,R.W. 1972. The interaction of catecholamines and adrenal corticosteroids in the induction of phosphopyruvate carboxylase in rat liver and adipose tissue. Biochem. J. 127:809-818.

23. Hakimi,P., Yang,J., Casadesus,G., Massillon,D., Tolentino-Silva,F., Nye,C.K., Cabrera,M.E., Hagen,D.R., Utter,C.B., Baghdy,Y. et al 2007. Overexpression of the cytosolic form of phosphoenolpyruvate carboxykinase (GTP) in skeletal muscle repatterns energy metabolism in the mouse. J. Biol. Chem. 282:32844- 32855.

24. Newsholme,E.A., and Williams,T. 1978. The role of phosphoenolpyruvate carboxykinase in amino acid metabolism in muscle. Biochem. J. 176:623-626.

93 25. Reshef,L., Meyuhas,O., Boshwitz,C., Hanson,R.W., and Ballard,F.J. 1972. Physiological role and regulation of glyceroneogenesis in rat adipose tissue. Isr. J. Med. Sci. 8:372-381.

26. Nye,C.K., Hanson,R.W., and Kalhan,S.C. 2008. Glyceroneogenesis is the predominant pathway for triglyceride glycerol synthesis in vivo in the rat.

27. Ballard,F.J., Hanson,R.W., and Leveille,G.A. 1967. Phosphoenolpyruvate carboxykinase and the synthesis of glyceride-glycerol from pyruvate in adipose tissue. J. Biol. Chem. 242:2746-2750.

28. Reshef,L., Niv,J., and Shapiro,B. 1967. Effect of propionate on pyruvate metabolism in adipose tissue. J. Lipid Res. 8:688-691.

29. Reshef,L., Ballard,F.J., and Hanson,R.W. 1969. The role of the adrenals in the regulation of phosphoenolpyruvate carboxykinase of rat adipose tissue. J. Biol. Chem. 244:5577-5581.

30. Reshef,L., Hanson,R.W., and Ballard,F.J. 1970. A possible physiological role for glyceroneogenesis in rat adipose tissue. J. Biol. Chem. 245:5979-5984.

31. Shafrir,E., Gutman,A., Gorin,E., and Orevi,M. 1970. Regulatory aspects in carbohydrate metabolism of adipose tissue: glycolysis, glycogen synthesis, and glyceroneogenesis. Horm. Metab Res. 2:Suppl-5.

32. Hanson,R.W., Patel,M., Reshef,L., and Ballard,F.J. 1970. The role of pyruvate carboxylase and phosphopyruvate carboxylase (P-enolpyruvate carboxykinase) in adipose tissue. Hoppe Seylers. Z. Physiol Chem. 351:293.

33. Kalhan,S.C., Mahajan,S., Burkett,E., Reshef,L., and Hanson,R.W. 2001. Glyceroneogenesis and the source of glycerol for hepatic triacylglycerol synthesis in humans. J. Biol. Chem. 276:12928-12931.

34. Siler,S.Q., Neese,R.A., Parks,E.J., and Hellerstein,M.K. 1998. VLDL-triglyceride production after alcohol ingestion, studied using [2-13C] glycerol. J. Lipid Res. 39:2319-2328.

35. Tagliaferro,A.R., Dobbin,S., Curi,R., Leighton,B., Meeker,L.D., and Newsholme,E.A. 1990. Effects of diet and exercise on the in vivo rates of the triglyceride-fatty acid cycle in adipose tissue and muscle of the rat. Int. J. Obes. 14:957-971.

36. Enoksson,S., Hagstrom-Toft,E., Nordahl,J., Hultenby,K., Pettersson,N., Isaksson,B., Permert,J., Wibom,R., Holm,C., Bolinder,J. et al 2005. Marked reutilization of free fatty acids during activated lipolysis in human skeletal muscle. J. Clin. Endocrinol. Metab 90:1189-1195.

94 37. Guo,Z., and Jensen,M.D. 1999. Blood glycerol is an important precursor for intramuscular triacylglycerol synthesis. J. Biol. Chem. 274:23702-23706.

38. Parks,E.J., and Hellerstein,M.K. 2006. Thematic review series: patient-oriented research. Recent advances in liver triacylglycerol and fatty acid metabolism using stable isotope labeling techniques. J. Lipid Res. 47:1651-1660.

39. Margolis,S., and Vaughan,M. 1962. Alpha-Glycerophosphate synthesis and breakdown in homogenates of adipose tissue. J. Biol. Chem. 237:44-48.

40. Bally,P.R., Cahill,G.F., Jr., LeBoeuf,B., and Renold,A.E. 1960. Studies on rat adipose tissue in vitro. V. Effects of glucose and insulin on the metabolism of palmitate-1-14C. J. Biol. Chem. 235:333-336.

41. Watford,M. 2000. Functional glycerol kinase activity and the possibility of a major role for glyceroneogenesis in mammalian skeletal muscle. Nutr. Rev. 58:145-148.

42. Hellerstein,M.K., Christiansen,M., Kaempfer,S., Kletke,C., Wu,K., Reid,J.S., Mulligan,K., Hellerstein,N.S., and Shackleton,C.H. 1991. Measurement of de novo hepatic lipogenesis in humans using stable isotopes. J. Clin. Invest 87:1841- 1852.

43. Schwarz,J.M., Neese,R.A., Turner,S., Dare,D., and Hellerstein,M.K. 1995. Short- term alterations in carbohydrate energy intake in humans. Striking effects on hepatic glucose production, de novo lipogenesis, lipolysis, and whole-body fuel selection. J. Clin. Invest 96:2735-2743.

44. Karbowska,J., Kochan,Z., and Swierczynski,J. 2001. Increase of lipogenic enzyme mRNA levels in rat white adipose tissue after multiple cycles of starvation-refeeding. Metabolism 50:734-738.

45. Shrago,E., Glennon,J.A., and Gordon,E.S. 1971. Comparative aspects of lipogenesis in mammalian tissues. Metabolism 20:54-62.

46. Aarsland,A., Chinkes,D., and Wolfe,R.R. 1997. Hepatic and whole-body fat synthesis in humans during carbohydrate overfeeding. Am. J. Clin. Nutr. 65:1774- 1782.

47. Diraison,F., Yankah,V., Letexier,D., Dusserre,E., Jones,P., and Beylot,M. 2003. Differences in the regulation of adipose tissue and liver lipogenesis by carbohydrates in humans. J. Lipid Res. 44:846-853.

48. Botion,L.M., Brito,M.N., Brito,N.A., Brito,S.R., Kettelhut,I.C., and Migliorini,R.H. 1998. Glucose contribution to in vivo synthesis of glyceride- glycerol and fatty acids in rats adapted to a high-protein, carbohydrate-free diet. Metabolism 47:1217-1221.

95 49. Brito,S.C., Festuccia,W.L., Kawashita,N.H., Moura,M.F., Xavier,A.R., Garofalo,M.A., Kettelhut,I.C., and Migliorini,R.H. 2006. Increased glyceroneogenesis in adipose tissue from rats adapted to a high-protein, carbohydrate-free diet: role of dietary fatty acids. Metabolism 55:84-89.

50. Wolfe,R.R., and Peters,E.J. 1987. Lipolytic response to glucose infusion in human subjects. Am. J. Physiol 252:E218-E223.

51. Hammond,V.A., and Johnston,D.G. 1987. Substrate cycling between triglyceride and fatty acid in human adipocytes. Metabolism 36:308-313.

52. Frayn,K.N., Shadid,S., Hamlani,R., Humphreys,S.M., Clark,M.L., Fielding,B.A., Boland,O., and Coppack,S.W. 1994. Regulation of fatty acid movement in human adipose tissue in the postabsorptive-to-postprandial transition. Am. J. Physiol 266:E308-E317.

53. Bolinder,J., Kerckhoffs,D.A., Moberg,E., Hagstrom-Toft,E., and Arner,P. 2000. Rates of skeletal muscle and adipose tissue glycerol release in nonobese and obese subjects. Diabetes 49:797-802.

54. Hagstrom-Toft,E., Qvisth,V., Nennesmo,I., Ryden,M., Bolinder,H., Enoksson,S., Bolinder,J., and Arner,P. 2002. Marked heterogeneity of human skeletal muscle lipolysis at rest. Diabetes 51:3376-3383.

55. Coppack,S.W., Frayn,K.N., Humphreys,S.M., Whyte,P.L., and Hockaday,T.D. 1990. Arteriovenous differences across human adipose and forearm tissues after overnight fast. Metabolism 39:384-390.

56. Reimer,F., Loffler,G., Hennig,G., and Wieland,O.H. 1975. The influence of insulin on glucose and fatty acid metabolism in the isolated perfused rat hind quarter. Hoppe Seylers. Z. Physiol Chem. 356:1055-1066.

57. Coppack,S.W., Frayn,K.N., Humphreys,S.M., Dhar,H., and Hockaday,T.D. 1989. Effects of insulin on human adipose tissue metabolism in vivo. Clin. Sci. (Lond) 77:663-670.

58. Smith,S.A. 2003. Central role of the adipocyte in the insulin-sensitising and cardiovascular risk modifying actions of the thiazolidinediones. Biochimie 85:1219-1230.

59. Jowsey,I.R., Murdock,P.R., Moore,G.B., Murphy,G.J., Smith,S.A., and Hayes,J.D. 2003. Prostaglandin D2 synthase enzymes and PPARgamma are co- expressed in mouse 3T3-L1 adipocytes and human tissues. Prostaglandins Other Lipid Mediat. 70:267-284.

60. Debril,M.B., Renaud,J.P., Fajas,L., and Auwerx,J. 2001. The pleiotropic functions of peroxisome proliferator-activated receptor gamma. J. Mol. Med. 79:30-47.

96 61. Frohnert,B.I., Hui,T.Y., and Bernlohr,D.A. 1999. Identification of a functional peroxisome proliferator-responsive element in the murine fatty acid transport protein gene. J. Biol. Chem. 274:3970-3977.

62. Virtanen,K.A., Hallsten,K., Parkkola,R., Janatuinen,T., Lonnqvist,F., Viljanen,T., Ronnemaa,T., Knuuti,J., Huupponen,R., Lonnroth,P. et al 2003. Differential effects of rosiglitazone and metformin on adipose tissue distribution and glucose uptake in type 2 diabetic subjects. Diabetes 52:283-290.

63. Tordjman,J., Chauvet,G., Quette,J., Beale,E.G., Forest,C., and Antoine,B. 2003. Thiazolidinediones block fatty acid release by inducing glyceroneogenesis in fat cells. J. Biol. Chem. 278:18785-18790.

64. Cadoudal,T., Blouin,J.M., Collinet,M., Fouque,F., Tan,G.D., Loizon,E., Beale,E.G., Frayn,K.N., Karpe,F., Vidal,H. et al 2007. Acute and selective regulation of glyceroneogenesis and cytosolic phosphoenolpyruvate carboxykinase in adipose tissue by thiazolidinediones in type 2 diabetes. Diabetologia 50:666-675.

65. Leroyer,S.N., Tordjman,J., Chauvet,G., Quette,J., Chapron,C., Forest,C., and Antoine,B. 2006. Rosiglitazone controls fatty acid cycling in human adipose tissue by means of glyceroneogenesis and glycerol phosphorylation. J. Biol. Chem. 281:13141-13149.

66. Guan,H.P., Li,Y., Jensen,M.V., Newgard,C.B., Steppan,C.M., and Lazar,M.A. 2002. A futile metabolic cycle activated in adipocytes by antidiabetic agents. Nat. Med. 8:1122-1128.

67. Olswang,Y., Cohen,H., Papo,O., Cassuto,H., Croniger,C.M., Hakimi,P., Tilghman,S.M., Hanson,R.W., and Reshef,L. 2002. A mutation in the peroxisome proliferator-activated receptor gamma-binding site in the gene for the cytosolic form of phosphoenolpyruvate carboxykinase reduces adipose tissue size and fat content in mice. Proc. Natl. Acad. Sci. U. S. A 99:625-630.

68. Botion,L.M., Kettelhut,I.C., and Migliorini,R.H. 1995. Increased adipose tissue glyceroneogenesis in rats adapted to a high protein, carbohydrate-free diet. Horm. Metab Res. 27:310-313.

69. Brito,M.N., Brito,N.A., Brito,S.R., Moura,M.A., Kawashita,N.H., Kettelhut,I.C., and Migliorini,R.H. 1999. Brown adipose tissue triacylglycerol synthesis in rats adapted to a high-protein, carbohydrate-free diet. Am. J. Physiol 276:R1003- R1009.

70. Franckhauser,S., Munoz,S., Pujol,A., Casellas,A., Riu,E., Otaegui,P., Su,B., and Bosch,F. 2002. Increased fatty acid re-esterification by PEPCK overexpression in adipose tissue leads to obesity without insulin resistance. Diabetes 51:624-630.

97 71. Walter,J.A., Ewart,K.V., Short,C.E., Burton,I.W., and Driedzic,W.R. 2006. Accelerated hepatic glycerol synthesis in rainbow smelt (Osmerus mordax) is fuelled directly by glucose and alanine: a 1H and 13C nuclear magnetic resonance study. J. Exp. Zoolog. A Comp Exp. Biol. 305:480-488.

72. Khayath,N., Mithieux,G., Zitoun,C., Coustau,C., Vicogne,J., Tielens,A.G., and Dissous,C. 2006. Glyceroneogenesis: an unexpected metabolic pathway for glutamine in Schistosoma mansoni sporocysts. Mol. Biochem. Parasitol. 147:145- 153.

73. Skelly,P.J., and Shoemaker,C.B. 1996. Rapid appearance and asymmetric distribution of glucose transporter SGTP4 at the apical surface of intramammalian-stage Schistosoma mansoni. Proc. Natl. Acad. Sci. U. S. A 93:3642-3646.

74. Furlong,S.T., and Caulfield,J.P. 1991. Head group precursors modify phospholipid synthesis in Schistosoma mansoni. J. Lipid Res. 32:703-712.

75. Okamura,T., Shimizu,H., Nagao,T., Ueda,R., and Ishii,S. 2007. ATF-2 regulates fat metabolism in Drosophila. Mol. Biol. Cell 18:1519-1529.

76. Castelein,N., Hoogewijs,D., De,V.A., Braeckman,B.P., and Vanfleteren,J.R. 2008. Dietary restriction by growth in axenic medium induces discrete changes in the transcriptional output of genes involved in energy metabolism in Caenorhabditis elegans. Biotechnol. J. 6:803-812.

77. She,P., Shiota,M., Shelton,K.D., Chalkley,R., Postic,C., and Magnuson,M.A. 2000. Phosphoenolpyruvate carboxykinase is necessary for the integration of hepatic energy metabolism. Mol. Cell Biol. 20:6508-6517.

78. Burgess,S.C., Hausler,N., Merritt,M., Jeffrey,F.M., Storey,C., Milde,A., Koshy,S., Lindner,J., Magnuson,M.A., Malloy,C.R. et al 2004. Impaired tricarboxylic acid cycle activity in mouse livers lacking cytosolic phosphoenolpyruvate carboxykinase. J. Biol. Chem. 279:48941-48949.

79. Owen,O.E., Kalhan,S.C., and Hanson,R.W. 2002. The key role of anaplerosis and cataplerosis for citric acid cycle function. J. Biol. Chem. 277:30409-30412.

80. Bremer,J., and Wojtczak,A.B. 1972. Factors controlling the rate of fatty acid - oxidation in rat liver mitochondria. Biochim. Biophys. Acta 280:515-530.

81. Beale,E.G., Hammer,R.E., Antoine,B., and Forest,C. 2004. Disregulated glyceroneogenesis: PCK1 as a candidate diabetes and obesity gene. Trends Endocrinol. Metab 15:129-135.

82. Beale,E.G., Hammer,R.E., Antoine,B., and Forest,C. 2002. Glyceroneogenesis comes of age. FASEB J. 16:1695-1696.

98 83. Franckhauser,S., Munoz,S., Elias,I., Ferre,T., and Bosch,F. 2006. Adipose overexpression of phosphoenolpyruvate carboxykinase leads to high susceptibility to diet-induced insulin resistance and obesity. Diabetes 55:273-280.

84. Brennan,K.J., and Hardeman,E.C. 1993. Quantitative analysis of the human alpha-skeletal actin gene in transgenic mice. J. Biol. Chem. 268:719-725.

85. Sahlin,K., Katz,A., and Broberg,S. 1990. Tricarboxylic acid cycle intermediates in human muscle during prolonged exercise. Am. J. Physiol 259:C834-C841.

86. Gibala,M.J., MacLean,D.A., Graham,T.E., and Saltin,B. 1997. Anaplerotic processes in human skeletal muscle during brief dynamic exercise. J. Physiol 502 ( Pt 3):703-713.

87. Aragon,J.J., and Lowenstein,J.M. 1980. The purine-nucleotide cycle. Comparison of the levels of citric acid cycle intermediates with the operation of the purine nucleotide cycle in rat skeletal muscle during exercise and recovery from exercise. Eur. J. Biochem. 110:371-377.

88. Lee,S.H., and Davis,E.J. 1979. Carboxylation and decarboxylation reactions. Anaplerotic flux and removal of citrate cycle intermediates in skeletal muscle. J. Biol. Chem. 254:420-430.

89. Hakimi,P., Johnson,M.T., Yang,J., Lepage,D.F., Conlon,R.A., Kalhan,S.C., Reshef,L., Tilghman,S.M., and Hanson,R.W. 2005. Phosphoenolpyruvate carboxykinase and the critical role of cataplerosis in the control of hepatic metabolism. Nutr. Metab (Lond) 2:33.

90. Kalhan,S.C., Bugianesi,E., McCullough,A.J., Hanson,R.W., and Kelley,D.E. 2008. Estimates of hepatic glyceroneogenesis in type 2 diabetes mellitus in humans. Metabolism 57:305-312.

91. Chen,J.L., Peacock,E., Samady,W., Turner,S.M., Neese,R.A., Hellerstein,M.K., and Murphy,E.J. 2005. Physiologic and pharmacologic factors influencing glyceroneogenic contribution to triacylglyceride glycerol measured by mass isotopomer distribution analysis. J. Biol. Chem. 280:25396-25402.

92. Varady,K.A., Roohk,D.J., Loe,Y.C., Evoy-Hein,B.K., and Hellerstein,M.K. 2007. Effects of modified alternate-day fasting regimens on adipocyte size, triglyceride metabolism, and plasma adiponectin levels in mice. J. Lipid Res. 48:2212-2219.

93. Moura,M.A., Festuccia,W.T., Kawashita,N.H., Garofalo,M.A., Brito,S.R., Kettelhut,I.C., and Migliorini,R.H. 2005. Brown adipose tissue glyceroneogenesis is activated in rats exposed to cold. Pflugers Arch. 449:463-469.

94. Festuccia,W.T., Kawashita,N.H., Garofalo,M.A., Moura,M.A., Brito,S.R., Kettelhut,I.C., and Migliorini,R.H. 2003. Control of glyceroneogenic activity in

99 rat brown adipose tissue. Am. J. Physiol Regul. Integr. Comp Physiol 285:R177- R182.

95. Wolfe,R.R., Peters,E.J., Klein,S., Holland,O.B., Rosenblatt,J., and Gary,H., Jr. 1987. Effect of short-term fasting on lipolytic responsiveness in normal and obese human subjects. Am. J. Physiol 252:E189-E196.

96. Silverberg,A.B., Shah,S.D., Haymond,M.W., and Cryer,P.E. 1978. Norepinephrine: hormone and neurotransmitter in man. Am. J. Physiol 234:E252- E256.

97. Marangou,A.G., Alford,F.P., Ward,G., Liskaser,F., Aitken,P.M., Weber,K.M., Boston,R.C., and Best,J.D. 1988. Hormonal effects of norepinephrine on acute glucose disposal in humans: a minimal model analysis. Metabolism 37:885-891.

98. Lang,C.H., Molina,P.E., Skrepnick,N., Bagby,G.J., and Spitzer,J.J. 1994. Epinephrine-induced changes in hepatic glucose production after ethanol. Am. J. Physiol 266:E863-E869.

99. Jensen,J., Ruzzin,J., Jebens,E., Brennesvik,E.O., and Knardahl,S. 2005. Improved insulin-stimulated glucose uptake and glycogen synthase activation in rat skeletal muscles after adrenaline infusion: role of glycogen content and PKB phosphorylation. Acta Physiol Scand. 184:121-130.

100. Samra,J.S., Simpson,E.J., Clark,M.L., Forster,C.D., Humphreys,S.M., Macdonald,I.A., and Frayn,K.N. 1996. Effects of epinephrine infusion on adipose tissue: interactions between blood flow and lipid metabolism. Am. J. Physiol 271:E834-E839.

101. Marion-Latard,F., de,G., I, Crampes,F., Berlan,M., Galitzky,J., Suljkovicova,H., Riviere,D., and Stich,V. 2001. A single bout of exercise induces beta-adrenergic desensitization in human adipose tissue. Am. J. Physiol Regul. Integr. Comp Physiol 280:R166-R173.

102. Fellows,I.W., Bennett,T., and Macdonald,I.A. 1985. The effect of adrenaline upon cardiovascular and metabolic functions in man. Clin. Sci. (Lond) 69:215-222.

103. Folch,J., Lees,M., and Sloane Stanley,G.H. 1957. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226:497-509.

104. Wieland,O. 1974. Methods of Ezymatic Analysis. Academic Press. London. 1404- 1409 pp.

105. Gutmann,I., and Wahlefeld,A.W. 1974. Methods of Enzymatic Analysis. Academic Press. London. 1464-1472 pp.

100 106. Reeves,R.E. 1941. The Estimation of Primary Carbinol Groups in Carbohydrates. J. Am. Chem. Soc. 63:1476-1477.

107. Ballard,F.J., and Hanson,R.W. 1967. Phosphoenolpyruvate carboxykinase and pyruvate carboxylase in developing rat liver. Biochem. J. 104:866-871.

108. Steele,R., Wall,J.S., De Bodo,R.C., and Altszuler,N. 1956. Measurement of size and turnover rate of body glucose pool by the isotope dilution method. Am. J. Physiol 187:15-24.

109. Kalhan,S., and Parimi,P. 2000. Gluconeogenesis in the fetus and neonate. Semin. Perinatol. 24:94-106.

110. Hetenyi,G., Jr. 1982. Correction for the metabolic exchange of 14C for 12C atoms in the pathway of gluconeogenesis in vivo. Fed. Proc. 41:104-109.

111. Katz,J., and Rognstad,R. 1966. The metabolism of tritiated glucose by rat adipose tissue. J. Biol. Chem. 241:3600-3610.

112. Reshef,L., Olswang,Y., Cassuto,H., Blum,B., Croniger,C.M., Kalhan,S.C., Tilghman,S.M., and Hanson,R.W. 2003. Glyceroneogenesis and the triglyceride/fatty acid cycle. J. Biol. Chem. 278:30413-30416.

113. Elia,M., Zed,C., Neale,G., and Livesey,G. 1987. The energy cost of triglyceride- fatty acid recycling in nonobese subjects after an overnight fast and four days of starvation. Metabolism 36:251-255.

114. Kashiwagi,A., Huecksteadt,T.P., and Foley,J.E. 1983. The regulation of glucose transport by cAMP stimulators via three different mechanisms in rat and human adipocytes. J. Biol. Chem. 258:13685-13692.

115. Stainsby,W.N., Sumners,C., and Andrew,G.M. 1984. Plasma catecholamines and their effect on blood lactate and muscle lactate output. J. Appl. Physiol 57:321- 325.

116. Kim,J., Saidel,G.M., and Kalhan,S.C. 2008. A computational model of adipose tissue metabolism: evidence for intracellular compartmentation and differential activation of lipases. J. Theor. Biol. 251:523-540.

117. Coppack,S.W., Frayn,K.N., and Humphreys,S.M. 1989. Plasma triacylglycerol extraction in human adipose tissue in vivo: effects of glucose ingestion and insulin infusion. Eur. J. Clin. Nutr. 43:493-496.

118. Coppack,S.W., Frayn,K.N., Humphreys,S.M., Dhar,H., and Hockaday,T.D. 1989. Effects of insulin on human adipose tissue metabolism in vivo. Clin. Sci. (Lond) 77:663-670.

101 119. Newsholme,P., Curi,R., Gordon,S., and Newsholme,E.A. 1986. Metabolism of glucose, glutamine, long-chain fatty acids and ketone bodies by murine macrophages. Biochem. J. 239:121-125.

120. Newsholme,P., Gordon,S., and Newsholme,E.A. 1987. Rates of utilization and fates of glucose, glutamine, pyruvate, fatty acids and ketone bodies by mouse macrophages. Biochem. J. 242:631-636.

121. Tchkonia,T., Tchoukalova,Y.D., Giorgadze,N., Pirtskhalava,T., Karagiannides,I., Forse,R.A., Koo,A., Stevenson,M., Chinnappan,D., Cartwright,A. et al. 2005. Abundance of two human preadipocyte subtypes with distinct capacities for replication, adipogenesis, and apoptosis varies among fat depots. Am. J. Physiol Endocrinol. Metab 288:E267-E277.

122. Tontonoz,P., Hu,E., Devine,J., Beale,E.G., and Spiegelman,B.M. 1995. PPAR gamma 2 regulates adipose expression of the phosphoenolpyruvate carboxykinase gene. Mol. Cell Biol. 15:351-357.

123. Jansson,P.A., Larsson,A., Smith,U., and Lonnroth,P. 1994. Lactate release from the subcutaneous tissue in lean and obese men. J. Clin. Invest 93:240-246.

124. Bonen,A., Heynen,M., and Hatta,H. 2006. Distribution of monocarboxylate transporters MCT1-MCT8 in rat tissues and human skeletal muscle. Appl. Physiol Nutr. Metab 31:31-39.

125. Abel,E.D., Peroni,O., Kim,J.K., Kim,Y.B., Boss,O., Hadro,E., Minnemann,T., Shulman,G.I., and Kahn,B.B. 2001. Adipose-selective targeting of the GLUT4 gene impairs insulin action in muscle and liver. Nature 409:729-733.

126. Franckhauser,S., Munoz,S., Hidalgo,A., Ferre,T., Mas,A., Elias,I., Cerdan,S., Pujol,A., and Bosch,F. 2007. Increased Glucose Uptake by Adipose Tissue Expression of Glucokinase in Transgenic Mice leads to Lactate Production rather than Fat Accumulation. American Diabetes Association (Abstr.)

127. Chang,T.J., Lee,W.J., Chang,H.M., Lee,K.C., and Chuang,L.M. 2008. Expression of subcutaneous adipose tissue phosphoenolpyruvate carboxykinase correlates with body mass index in nondiabetic women. Metabolism 57:367-372.

128. Guo,Z., and Jensen,M.D. 1998. Intramuscular fatty acid metabolism evaluated with stable isotopic tracers. J. Appl. Physiol 84:1674-1679.

129. James,D.E., Brown,R., Navarro,J., and Pilch,P.F. 1988. Insulin-regulatable tissues express a unique insulin-sensitive glucose transport protein. Nature 333:183-185.

130. Maltin,C.A., Delday,M.I., Baillie,A.G., Grubb,D.A., and Garlick,P.J. 1989. Fiber- type composition of nine rat muscles. I. Changes during the first year of life. Am. J. Physiol 257:E823-E827.

102 131. Brooks,G.A. 2002. Lactate shuttles in nature. Biochem. Soc. Trans. 30:258-264.

132. Simonsen,L., Bulow,J., and Madsen,J. 1994. Adipose tissue metabolism in humans determined by vein catheterization and microdialysis techniques. Am. J. Physiol 266:E357-E365.

133. Roth,D.A., and Brooks,G.A. 1990. Lactate and pyruvate transport is dominated by a pH gradient-sensitive carrier in rat skeletal muscle sarcolemmal vesicles. Arch. Biochem. Biophys. 279:386-394.

134. Roth,D.A., and Brooks,G.A. 1990. Lactate transport is mediated by a membrane- bound carrier in rat skeletal muscle sarcolemmal vesicles. Arch. Biochem. Biophys. 279:377-385.

135. van Loon,L.J., Thomason-Hughes,M., Constantin-Teodosiu,D., Koopman,R., Greenhaff,P.L., Hardie,D.G., Keizer,H.A., Saris,W.H., and Wagenmakers,A.J. 2005. Inhibition of adipose tissue lipolysis increases intramuscular lipid and glycogen use in vivo in humans. Am. J. Physiol Endocrinol. Metab 289:E482- E493.

136. Pilkis,S.J., Chrisman,T.D., El-Maghrabi,M.R., Colosia,A., Fox,E., Pilkis,J., and Claus,T.H. 1983. The action of insulin on hepatic fructose 2,6-bisphosphate metabolism. J. Biol. Chem. 258:1495-1503.

137. Wu,C., Khan,S.A., Peng,L.J., Li,H., Carmella,S.G., and Lange,A.J. 2006. Perturbation of glucose flux in the liver by decreasing F26P2 levels causes hepatic insulin resistance and hyperglycemia. Am. J. Physiol Endocrinol. Metab 291:E536-E543.

138. Pilkis,S.J., Claus,T.H., and El-Maghrabi,M.R. 1988. The role of cyclic AMP in rapid and long-term regulation of gluconeogenesis and glycolysis. Adv. Second Messenger Phosphoprotein Res. 22:175-191.

139. Pilkis,S.J., El-Maghrabi,M.R., McGrane,M., Pilkis,J., and Claus,T.H. 1982. Regulation by glucagon of hepatic pyruvate kinase, 6-phosphofructo 1-kinase, and fructose-1,6-bisphosphatase. Fed. Proc. 41:2623-2628.

140. Cadoudal,T., Fouque,F., Benelli,C., and Forest,C. 2008. [Glyceroneogenesis and PEPCK-C: pharmacological targets in type 2 diabetes.]. Med. Sci. (Paris) 24:407- 414.

141. Sooranna,S.R., and Saggerson,E.D. 1979. Effects of starvation and adrenaline on glycerophosphate acyltransferase and dihydroxy acetone phosphate acyltransferase activities in rat adipocytes. FEBS Lett. 99:67-69.

142. Schlossman,D.M., and Bell,R.M. 1978. Glycerolipid biosynthesis in Saccharomyces cerevisiae: sn-glycerol-3-phosphate and dihydroxyacetone phosphate acyltransferase activities. J. Bacteriol. 133:1368-1376.

103 143. Datta,N.S., Wilson,G.N., and Hajra,A.K. 1984. Deficiency of enzymes catalyzing the biosynthesis of glycerol-ether lipids in Zellweger syndrome. A new category of metabolic disease involving the absence of peroxisomes. N. Engl. J. Med. 311:1080-1083.

104